6 Wood Discoloration

Olaf Schmidt 

Wood and Tree Fungi 

Biology, Damage, Protection, and Use

Olaf Schmidt 

Wood and Tree Fungi 

Biology, Damage, 

Protection, and Use 

With 74 Figures, 12 in Colors, and 49 Tables 

123

Professor Dr. Olaf Schmidt 

Universität Hamburg 

Zentrum Holzwirtschaft 

Abteilung Holzbiologie 

Leuschnerstraße 91 

21031 Hamburg 

Germany 

o.schmidt@holz.uni-hamburg.de 

Cover: Fruit body of serpula lacrymans and electrophoresis gel demontstrating species-specific priming PCR. 

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ISBN-10 3-540-32138-1 Springer Berlin Heidelberg New York 

ISBN-13 978-3-540-32138-5 Springer Berlin Heidelberg New York 

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Foreword 

Wood, as a raw material and a renewable biomass, has had great importance 

for thousands of years. Under suitable conditions, however, it is also easily 

degradable as part of the biological cycle. The processes of decomposition 

by fungi, and measures for protection against them, have been studied for 

quite a long time. The resulting knowledge on the causes and effects of wood 

degradation can hardly be overlooked. 

For more than 30 years, Olaf Schmidt has investigated the causes and effects 

of wood degradation by fungi and bacteria. Pioneering contributions have also 

been made in several fields, such as wood-inhabiting bacteria and molecular 

methods for fungal identification. Laboratory work is accompanied by teaching 

the field of wood deterioration by microorganisms, thus contributing to the 

broad spectrum of information accumulated. 

“WoodandTreeFungi”byOlafSchmidtpresentsthemostcomprehensive 

treatise on the fundamentals, causes, and consequences of decomposition of 

wood as well as measures for its prevention. The 1,400 references give an 

overlook of the vast amount of information evaluated. For a long time to come 

this book will be the competent source of knowledge about the fascinating 

interactions between wood and microorganisms. 

Walter Liese

Preface 

This book is the updated revision of the German edition “Holz- und Baumpilze” 

from 1994. Errors were corrected and new results were included. Particularly 

the chapter “Identification” was supplemented by molecular techniques. I realize 

that a one-author book on a relatively broad topic must include errors 

and also may have ignored recent literature. Strictly speaking, one should only 

write about things that they have experienced themselves, in the case of point 

this only concerns some aspects of bacteria and those fungi which inhabit the 

xylem of dead wood. Thus, current “secondary literature” was used for those 

chapters that are “on the edge” of my own research interest. 

For better readability, the authors of fungal names are not mentioned in 

the text, but summarized in an appendix. Fungal synonyms are also not given. 

These are available from Index Fungorum (www.indexfungorum.org/names/ 

names.asp). Fungal names cited from (older) publications were transferred to 

the current version. 

Thanks for general advice go to Prof. Dr. Dr. h.c. mult. Walter Liese, for critical 

reading to Prof. Dr. Dirk Dujesiefken (Chap. 8.2), Dr. Hubert Willeitner, 

and Dr. Peter Jüngel (Chap. 7.4), to Mrs. Ute Moreth for providing experimental 

data, Dr. Tobias Huckfeldt for several photographs, many colleagues for 

permission to use their photographs, Mrs. Christina Waitkus for transferring 

many pictures in an electronic version, to Springer-Verlag, particularly Mrs. 

UrsulaGrammandDr.D.Czeschlik,forco-operation,toMr.JardiMullinaxfor 

making my English understandable, and to Mrs. Cornelia Gründer for careful 

printing. 

Hamburg, December 2005 Olaf Schmidt

Contents 

1 Introduction 1 

2 Biology 3 

2.1 Cytology and Morphology.................................................... 3 

2.2 Growth and Spreading ......................................................... 10 

2.2.1 Vegetative Growth ..................................................... 10 

2.2.2 Reproduction of Deuteromycetes ................................. 16 

2.2.3 Sexual Reproduction.................................................. 18 

2.2.4 Fruit Body Formation ................................................ 24 

2.2.5 Production, Dispersal and Germination of Spores .......... 25 

2.3 Sexuality............................................................................ 26 

2.4 Identification...................................................................... 31 

2.4.1 Traditional Methods .................................................. 31 

2.4.2 Molecular Methods.................................................... 33 

2.5 Classification...................................................................... 47 

3 Physiology 53 

3.1 Nutrients ........................................................................... 53 

3.2 Air .................................................................................... 58 

3.3 Wood Moisture Content ....................................................... 60 

3.4 Temperature....................................................................... 67 

3.5 pH Value and Acid Production by Fungi ................................. 70 

3.6 Light and Force of Gravity .................................................... 74 

3.7 Restrictions of Physiological Data.......................................... 77 

3.8 Competition and Interactions Between Organisms................... 79 

3.8.1 Antagonisms, Synergisms, and Succession .................... 79 

3.8.2 Mycorrhiza and Lichens ............................................. 82 

4 Wood Cell Wall Degradation 87 

4.1 Enzymes and Low Molecular Agents ...................................... 87 

4.2 Pectin Degradation.............................................................. 92 

4.3 Degradation of Hemicelluloses.............................................. 93 

4.4 Cellulose Degradation.......................................................... 95 

4.5 Lignin Degradation ............................................................. 99

X Contents 

5 Damages by Viruses and Bacteria 109 

5.1 Viruses .............................................................................. 109 

5.2 Bacteria ............................................................................. 109 

6 Wood Discoloration 119 

6.1 Molding............................................................................. 121 

6.2 Blue Stain........................................................................... 125 

6.3 Red Streaking ..................................................................... 129 

6.4 Protection.......................................................................... 131 

7 Wood Rot 135 

7.1 Brown Rot.......................................................................... 135 

7.2 White Rot........................................................................... 138 

7.3 Soft Rot ............................................................................. 142 

7.4 Protection.......................................................................... 146 

8 Habitat of Wood Fungi 161 

8.1 Fungal Damage to Living Trees.............................................. 161 

8.1.1 Bark Diseases............................................................ 163 

8.1.2 Wilt Diseases ............................................................ 168 

8.2 Tree Wounds and Tree Care .................................................. 173 

8.2.1 Wounds and Defense Against Discoloration and Decay ... 173 

8.2.2 Pruning.................................................................... 177 

8.2.3 Wound Treatment...................................................... 178 

8.2.4 Detection of Tree and Wood Damages .......................... 179 

8.3 Tree Rots by Macrofungi ...................................................... 183 

8.3.1 Armillaria Species ..................................................... 186 

8.3.2 Heterobasidion annosum s.l......................................... 189 

8.3.3 Stereum sanguinolentum ............................................ 195 

8.3.4 Fomes fomentarius ..................................................... 195 

8.3.5 Laetiporus sulphureus ................................................ 197 

8.3.6 Meripilus giganteus .................................................... 197 

8.3.7 Phaeolus schweinitzii.................................................. 198 

8.3.8 Phellinus pini ............................................................ 198 

8.3.9 Piptoporus betulinus .................................................. 199 

8.3.10 Polyporus squamosus ................................................. 199 

8.3.11 Sparassis crispa ......................................................... 200 

8.4 Damage to Stored Wood and Structural Timber Outdoors......... 200 

8.4.1 Daedalea quercina ..................................................... 202 

8.4.2 Gloeophyllum Species................................................. 202 

8.4.3 Lentinus lepideus ....................................................... 205 

8.4.4 Paxillus panuoides ..................................................... 205 

8.4.5 Schizophyllum commune............................................. 206

Contents XI 

8.4.6 Trametes versicolor .................................................... 206 

8.5 Damage to Structural Timber Indoors.................................... 207 

8.5.1 General and Identification .......................................... 207 

8.5.2 Lesser Common Basidiomycetes in Buildings................. 212 

8.5.3 Common House-Rot Fungi ......................................... 214 

8.5.4 Prevention of Indoor Decay Fungi and Refurbishment 

of Buildings .............................................................. 233 

9 Positive Effects of Wood-Inhabiting Microorganisms 237 

9.1 “Myco-Wood”..................................................................... 238 

9.2 Cultivation of Edible Mushrooms .......................................... 239 

9.3 Biological Pulping ............................................................... 244 

9.4 “Palo Podrido” and “Myco-Fodder”....................................... 246 

9.5 Wood Saccharification and Sulphite Pulping ........................... 247 

9.6 Grinding and Steam Explosion.............................................. 248 

9.7 Recent Biotechnological Processes and Outlook....................... 249 

Appendix 1 

Identification Key for Strand-Forming House-Rot Fungi 253 

Appendix 2 

Fungi Mentioned in this Book 261 

References 267 

Subject Index 329

1 Introduction 

Wood is damaged by various agents (Table 1.1). 

This book addresses wood damage caused by microorganisms (fungi and 

bacteria). Wood damage by fungi has also been called “wood diseases” (“Holzkrankheiten”) 

and “wood pathology” (“Holzpathologie”). Because it concerns 

the substrate “tree” in the majority of dead cells, because all parenchyma cells 

in the wood of felled trees are dead after a few weeks, and, thus, because a dead 

tissue cannot fall ill, distance was taken to these terms. With regard to the 

microbial decomposition of biomass, in the English language there is a welldescribing 

differentiation between “biodeterioration”, which means unwanted 

biological destruction, and “biodegradation”, which means controlled degradation 

by microorganisms or their enzymes and degrading agents. Biodeterioration 

corresponds to the German “Holzzerstörung” and “Holzzersetzung”, 

and the latter positive aspect of wood biodegradation (“Holzabbau”) belongs 

to the area of “biotechnology of lignocelluloses” (Bruce and Palfreyman 1998; 

Chap. 9). 

Inthefollowingtext,themicrobialdamagetothexylem(wood)ofthetreeis 

mainly addressed. Since leaves, bark, and roots are entrance gates for parasites 

into the living tree that can lead to reduced tree growth and to lesser wood 

quality, some aspects of the area of “forest pathology” are included (Butin 

1995; Chaps. 5 and 8.1–8.3). The mechanisms of the decomposition of solid 

Table 1.1. Agents for wood damages 

– mankind: e.g., paper production, fire for cooking 

– conflagrations for agriculture 

– weathering, UV light 

– acids, bases, corrosion by salts, gases, discoloration by metals 

– wood insects: xylophagous beetles, termites, wasps, breeding ambrosia beetles, 

wood-colonizing ants 

– marine borers 

– bacteria: wetwood, discoloration, pit degradation, decay by erosion, tunneling, 

cavity bacteria 

– fungi: 

wood discoloration by molds, blue-stain fungi, red-streaking fungi 

wood decay by brown, soft, and white-rot fungi 

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2 1 Introduction 

wood apply essentially also to wood-based composites (plywood, fiberboard, 

particleboard, orientated strand board) (e.g., Chung et al. 1999) and to woodplastic 

composites (Simonson et al. 2004). Sutter (2003) and Unger et al. (2001) 

report on damages, conservation, and restoration of wood artifacts. Bacterial 

and soft-rot attack of archaeological wood is described by Blanchette (1995), 

Nelson et al. (1995), and Singh et al. (2003). 

The decomposition of biomass, which concerns wood and other lignocelluloses 

(annual plants), is a necessary part of the natural material cycle: during 

photosynthesis, wood and O2 are formed from CO2 and H2Obymeansoflight. 

In counterpart, the wood becomes degraded by fungi and bacteria to CO2,H2O 

and energy for microbial metabolism. 

In the forests of the earth, about 400 billion t of CO2 are bound. Without 

microbial degradation (or burning) of the biomass, the CO2 supply of the 

atmosphere necessary for photosynthesis would be used up in 20–30 years 

(Schlegel 1992), photosynthesis would grind to a halt, and the earth would be 

overfilled with non-decaying biomass. 

Humans retard wood degradation by microorganisms for economic reasons 

by wood protection measures (Willeitner and Liese 1992; Goodell et al. 2003; 

Müller 2005; Chap. 7.4) in order to prolong the use of the raw material wood. 

Thus, for example, the service life of a beech sleeper, which would amount to 

about 3 years without any protection, extends to about 45 years after impregnation 

with coal tar oil. 

Until around 1800, rot was considered punishment from God, and fruit 

bodies as eczemas. Still, in 1850, v. Liebig attributed decay to a “slow burning”. 

In 1874, Robert Hartig recognized the causality between pest and damage and 

is thus considered the father of forest and wood pathology (Merrill et al. 1975). 

The first pure culture of a wood-degrading fungus was succeeded to Brefeld 

(1881). 

Research on wood deterioration is done worldwide. The global network for 

cooperation in forest and forest products research is the International Union of 

Forest Research Organizations (IUFRO), which was created in Eberswalde, Germany, 

in 1892, and has 15,000 scientists in almost 700 member organizations 

in over 110 countries. Current research results on wood damages, protection, 

and investigation methods are introduced at the annual symposia of the International 

Research Group on Wood Preservation (IRG). Edible mushrooms 

cultured on wood are discussed at the meetings of the International Mycological 

Society. A recent comprehensive treatise on the various aspects of fungi is 

“The Mycota” (Esser 1994 et seq.). 

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2 Biology 

2.1 

Cytology and Morphology 

“Wood fungi” are eukaryotic and carbon-heterotrophic (free from chlorophyll) 

organisms with chitin in the cell wall, reproduce asexually and/or sexually 

by non-flagellate spores, filamentous, immovable and mostly land inhabiting. 

Damage to wood in water by fungi is described by Jones and Irvine (1971), Jones 

(1982) and Kim and Singh (2000). Soft-rot fungi belonging to the Ascomycetes 

and Deuteromycetes (Chap. 7.3) destroy wood with high moisture content in 

water or soil (e.g., Findlay and Savory 1954; Liese 1955). Fungi associated with 

leaf litter in a woodland stream were treated by Suberkropp (1997). 

In this book, a fungal cell, the hypha, is defined as one individual cell 

of mostly tubular shape that consists of a cell wall, contains a protoplasm 

with a nucleus and other organelles, and is in the “higher fungi” separated 

from its one or two neighbors by a transverse wall, the septum (Fig. 2.1). In 

analogy to the “higher plants”, where nearly every living cell is connected 

to its neighbors by cytoplasmic channels, the plasmodesmata, which pass 

through the intervening cell walls, also the protoplasms of neighbored hyphae 

areconnectedwitheachotherthroughtheporeordoliporesystem(Fig.2.2). 

This basic hypha is termed “vegetative hypha” in this book. This definition 

contrasts to others where one hypha, also termed generative hypha, is a more 

or less long filament consisting of several hyphal “compartments”, a definition 

that is hazy because the transition to the next higher unit, the mycelium, is 

flowing. The mycelium is thus the filamentous lining up of hyphae, consisting 

in young mycelia of only a few vegetative hyphae and in older ones of several 

and branched hyphae. Figure 2.1 shows septate hyphae as they occur in the 

wood-inhabiting Deuteromycetes, Ascomycetes, and Basidiomycetes. 

The diameter of hyphae reaches from 0.1–0.4µm for the microhyphae of 

Phellinus pini (Liese and Schmid 1966) to 60µm for the vessel hyphae in the 

mycelial strand (cord) of the True dry rot fungus, Serpula lacrymans,withan 

average for vegetative hyphae of about 2–7 µm (S. lacrymans: 3µm: Seehann 

and v. Riebesell 1988). Their length reaches from about 5µm for round/oval 

cells (spores) up to several micrometers. The size of many bacteria is between 

0.4 and 5µm. 

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4 2Biology 

Fig.2.1. Vegetative hyphae. C coenocytic hyphae, S septate hyphae 

Due to the smallness of the individual hypha and the use of microscopic and 

microbiological methods, fungi are microorganisms. This attachment does not 

contrast to the fact that fungi can form large and firm structures such as fruit 

bodies of decimeters in size like in the Tinder fungus, Fomes fomentarius (see 

Fig. 8.15). Those fruit bodies are, however, also composed of single hyphae. The 

main argument is, however, that the “actual fungus” is the vegetative hyphal 

system that can grow unlimited by simple mitotic reproduction without ever 

fruiting if fresh nutrients (wood, soil, agar) are available, and if growth in 

a certain biotope is not inhibited by the own or foreign metabolic products. 

Fungi are scientifically examined in microbiological or medical institutes 

(predominantly Deuteromycetes and Ascomycetes) and often in botanical institutes. 

They do, however, no longer rank among the plants. In multi-kingdom 

systems (Whittaker 1969), the “higher fungi” (Ascomycetes, Basidiomycetes) 

form the distinct group of fungi beside the Prokaryotes (Bacteria), Protista 

(eukaryotic single-celled organisms: slime fungi and “lower fungi”), plants, 

and animals (Müller and Loeffler 1992). Based on rDNA sequences, Woese and 

Fox (1977) divided the Prokaryotes into the kingdoms Eubacteria and Archaebacteria 

and later emphasized three domains, which were renamed Bacteria, 

Archaea, and Eucarya (see Fig. 5.1). 

The hyphal wall defines the shape of the hypha and provides the mechanical 

strength to resist the internal turgor pressure. The wall consists of various 

carbohydrates. Some yeast has mannan-β-glucans, while Ascomycetes, 

Deuteromycetes, and Basidiomycetes possess chitin-β-glucans, never cellulose. 

Chitin [poly–β(1-4)-N-acetoamido-2-deoxy-D-glucopyranose], which occurs 

except in fungi also in the exoskeleton of arthropods and crustaceans, and 

in some mollusks, is a macromolecule made of β-1,4-glycosidically linked 

N-acetylglucosamine units. Chitin synthases (CHS; EC 2.4.1.16) catalyze the 

formation of chitin from the precursor UDP-N-acetylglucosamine. In the yeast 

Saccharomyces cerevisiae, CHS I acts as a repair enzyme and is involved in the 

chitin synthesis at the point where the daughter and mother cells separate. CHS 

II participates in septa formation and CHS III in chitin synthesis of the cell 

wall (Robson 1999). Ascomycetes have two-layered cell walls, while walls of Basidiomycetes 

are multilamellar. The entire structure of the cell wall including 

extracellularlayers is complex(Toft1992;Robson1999):The wall offilamentous 

fungi may consist for example of an inner wall of about 10–20 nm composed 

of chitin microfibrils and an outer wall composed of a protein layer (about 

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2.1 Cytology and Morphology 5 

10 nm), a layer of glycoprotein (about 50 nm), and a slime layer, also termed 

mucilage layer, sheath, extracellular matrix or mycofibrils (about 75–100 nm). 

Slimelayersarecommontofungiandhavebeenfoundinbluestain,white, 

brown, and soft-rot fungi. They are composed of protein, lipid and carbohydrate 

containing material (α-glucan, β-1,3 and β-1,6-glucan) or of crystalline 

to membranous and fibrillar structures (Liese and Schmid 1963; Schmid and 

Liese 1965; Schmid and Baldermann 1967; Holdenrieder 1982; Green et al. 

1989). Various functions have been suggested for the slime layer (Schmid and 

Liese 1966; Sutter et al. 1984; Green et al. 1991b; Kim 1991; Abu Ali et al. 1997; 

Messner et al. 2003; Table 2.1). In Phanerochaete chrysosporium, the slime layer 

is composed of equal amounts of carbohydrates, lipids, and proteins, including 

five fractions with molecular weights between 30 and 200 kDa (cf. Messner 

et al. 2003). Production of the slime layer was influenced by iron, manganese 

and nitrogen concentration, temperature, and pH value (Jellison et al. 1997). 

Hyphae may be encrusted and covered with resinous material, oil drops, 

and calcium oxalate crystals (e.g., Holdenrieder 1982). 

The hyphal wall encloses the cytoplasm with its outer boundary, the plasmalemma. 

In the majority of fungi, ergosterol is the chief sterol in the plasma 

membrane and is used for fungal quantification (Chap. 2.4). Some antifungals 

like polyene and triazole act on this ergosterol (Robson 1999). The cytoplasm 

principally resembles that one of plants. There is one too many relatively 

small nuclei. Plastides are absent. Growing hyphae of Ascomycetes and Basidiomycetes 

show at the hyphal apex a mass of small vesicles, the “Spitzenkörper”. 

The tonoplast encloses a vacuolar system. Carbon is stored in glycogen 

vesicles and lipid vacuoles. Nitrogen is deposited as amino acids in the vacuolarsystemorasprotein.Phosphorusiscondensedaspolyphosphateinvolutin 

grana, often in vacuoles. Some yeast contains starch. 

Table 2.1. Possible functions for fungal slime layers 

– substrate recognition 

– adhesion to and establishing contact 

– covering the S3 layer of the wood cell wall during decay process 

– conditioning of the substrate for decay 

– modification of the extracellular ionic environment and pH-value 

– transport vector for low-molecular decay agents and enzymes to the wood (see Fig. 7.3) 

– transport vector for degradation products to the hypha 

– storage, concentration and retention of decay agents 

– regulation of the decay process, e.g., by controlling the glucose level 

– microenvironment for H2O2 maintenance needed for lignin degradation 

– storage of nutrients 

– permitting a film of liquid water to surround the wood cell wall 

– protection of the mycelium against dehydration and adverse environmental conditions 

– increase of surface area for aerobic respiration 

– storage of copper or CCA from attack of impregnated wood 

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6 2Biology 

The solutes in the cytoplasm and vacuolar system have a certain osmotic 

potential. If the potential is lower than that of the substrate, water is adsorbed 

across the membranes, increasing the volume of the cytoplasm (Jennings and 

Lysek 1999). An internal turgor needs to be generated for the elongation of the 

hyphal apex that is that water uptake and wall growth are in balance. 

Mycelium is the filamentous, partly branched, and in the wood-inhabiting 

Basidiomycetes usually whitish network made from some to numerous, in the 

light microscope hyaline to light yellow and in the case of blue-stain fungi 

brownish (melanin) hyphae. In Deuteromycetes, the pigmentation of the culture 

is manifold due to the various pigments of the conidia, whose color depends 

on the species. Mycelium forms the macroscopically visible thallus, the 

undifferentiated form of vegetative growth of fungi (thallobionts), which is not 

differentiated as it is the kormus of the “higher plants” into the organs, shoot 

axis, leaf, and root. Mycelium is the actual fungus with nourishing function and 

thus with wood decay ability. Under sufficient nutrient offer, mycelium is theoretically 

growable for an unlimited period. Sexuality with fruit body formation 

is not necessary for survival. For example, mycelium of an isolate of the Asian 

edible mushroom Shiitake, Lentinula edodes, is now maintained since about 

1940 exclusively on agar in the refrigerator without ever fructifying, but would 

immediately develop fruit bodies (Fig. 9.1) when favorable environmental conditions 

are provided (Schmidt 1990). The largest and longest-living wood fungi 

are Armillaria species. A clone of A. gallica in a Michigan forest covered 15 ha 

and was estimated at an age of about 1,500 years and a total biomass of 1,000 t 

(Smith et al. 1992). A clone of A. ostoyae estimated at 400–1,000 years covered 

an area of 6 km 2 in the Rocky Mountains (Anonymous 1992a). In Oregon, an 

A. ostoyae clone of 2,400 years stretched over an area of about 9 km 2 of forest 

soil (Schwarze and Ferner 2003). 

Deutero- and Ascomycetes have in the septum a central simple pore. Woodinhabiting 

Basidiomycetes (Homobasidiomycetes) contain the more complicateddoliporeseptumwithaparenthesomeonbothsides(Fig.2.2). 

The protoplasts of neighboring hyphae are connected through pores in the 

septa for the longitudinal migration of organelles and even nuclei, and for 

the transport of solutes (translocation; Chap. 3.1). Woronin bodies, which are 

composed of protein, block the pore when a hypha becomes injured. 

Fig.2.2. Septa (S) ofAscomycetes(a) 

and Basidiomycetes (b). P simple pore 

septum, D dolipore septum, H hyphae 

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The hyphal system expands by extension of individual hyphae that exhibit 

apical growth and by branching (Fig. 2.3). 

Different zones occur in the growing hypha (Fig. 2.4), which correspond to 

different ages and developmental stages (Huckfeldt 2003). 

Fig.2.3. Apical growth and hyphal branching system. One branch is sectioned to show 

the septum and some features of the protoplasm. N nucleus, ER endoplasmic reticulum, 

D dictyosome, V vacuole, M mitochondrion, Woronin bodies (dark) [reproduction with 

permission, from Jennings DH, Lysek G (1999) Fungal Biology, 2nd edn. Bios, Oxford, 

Fig. 1.1. page 6] 

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8 2Biology 

Fig.2.4. Ultrastructural features and zonation of growing hyphae of a house-rot fungus in 

woody tissue (MP, S1, S2andT wood cell wall layers). G glycogen, N nucleus, L lipid drops, 

Mi mitochondrion, MS multivesicles structure, V vacuole, Ve vesicle (from Huckfeldt 2003) 

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Due to the apical growth, the hyphal tip is the most sensitive part of the 

mycelium and gets the first contact with the substrate wood. Wood preservatives, 

unfavorable temperatures, shortage of nutrients, and moisture affect the 

tip. The tip contains different vesicles and membranous structures for cell wall 

synthesis and transport processes as well as enzymes for nutrient metabolism 

(Robson 1999). Like in other Basidiomycetes, the tip in Serpula lacrymans 

(Fig. 2.4) consists of three zones: 

In the apical zone, material for the structure of the cell wall, slime layer, 

and plasmalemma is concentrated and incorporated in the growing mycelium. 

Vesicles from the Spitzenkörper merge with the plasmalemma and deliver 

cell wall components. In the subapical zone, compartments and ribosomes 

are involved in the synthesis of cell wall material and secretion products. 

The basal zone contains the nucleus, or in the case of dikaryons, two nuclei. 

Vacuoles are involved in internal recycling processes, detoxification, storage, 

upkeep of turgor pressure, control of ionic strength as well as metabolization 

of compartments and macromolecules. The cytoskeleton, which consists of 

actin filaments and microtubuli, serves together with motor proteins to the 

upkeep of the zonation of the hyphal tip by changing the position of the 

compartments. Thus, the compartments continuously follow the growing tip 

to maintain the density of organelles in the subapical zone. Jennings and Lysek 

(1999) differentiated the apical growth zone with the extending hyphal tip, the 

absorption zone where there is uptake of nutrients, the storage zone in which 

nutrients are stored as reserve substances, and the senescence zone where dark 

pigments and lysis may occur. 

The hyphal system produces a loose network of filaments (aerial mycelium 

on the wood surface, substrate mycelium within wood and soil) or solid, 

morphologically differentiated units such as the strands of house-rot fungi 

and the rhizomorphs of Armillaria species (Chap. 2.2.1), and the fruit bodies. 

The mycelia of wood fungi differ considerably in their growth rate. Table 2.2 

shows the growth rate of some house-rot fungi. 

The growth rate serves as a characteristic for species identification in keys. 

Growth rate is also used as a hint of the age of the fungal infestation time of 

a building, e.g., in the case of damage by Serpula lacrymans (Chap. 8.5.3.4). 

However, mycelial extension depends on environmental conditions like temperature 

and nutrients, which differ between stable and favorable laboratory 

conditions and fluctuations in buildings. Furthermore, different isolates of 

a species commonly differ in growth rate (“strain variation”). In addition, 

dikaryons and monokaryons may differ in growth. For example, dikaryons of 

Lentinula edodes (Schmidt and Kebernik 1987), Serpula lacrymans (Schmidt 

and Moreth-Kebernik 1991a), and Stereum hirsutum (Rayner and Boddy 1988) 

grew faster than the monokaryons. Nevertheless, there are so-called “fastgrowing” 

wood fungi like the Cellar fungus, Coniophora puteana, withupto 

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10 2 Biology 

Table 2.2. Growth rate of house-rot fungi at optimum temperature (from Schmidt and 

Huckfeldt 2005) 

Group Species Number of Maximum radial 

investigated increase on agar 

isolates per day (mm) 

Dry-rot fungi Serpula lacrymans 2 4.0–5.1 

S. himantioides 2 7.0–11.0 

Leucogyrophana mollusca 6 1.0–3.3 

L. pinastri 4 2.4–4.2 

Meruliporia incrassata 2 2.8–3.2 

Cellar fungi Coniophora puteana 27 2.5–11.3 

C. marmorata 2 9.7–12.3 

C. arida 1 4.7 

C. olivacea 5 3.7–9.0 

White polypores Antrodia vaillantii 12 4.3–7.7 

A. sinuosa 4 4.0–8.0 

A. xantha 3 5.5–8.2 

A. serialis 3 3.5–3.9 

Oligoporus placenta 4 4.2–9.8 

Gill polypores Gloeophyllum abietinum 5 3.8–5.5 

G. sepiarium 4 6.8–8.3 

G. trabeum 5 7.1–9.1 

Oak polypore Donkioporia expansa 1 5.1 

11 mm radial increment per day on 2% malt extract agar at 23 ◦ C and “slowgrowing” 

species like S. lacrymans with up to 5mm at 19 ◦ C. 

Mycelium of wood-decay fungi predominantly grows as substrate mycelium 

insideofthesubstrateswood(orsoil)andisoftennotvisiblyontheoutside, 

thus, wood rot, particularly at incipient decay, is frequently not recognizable 

outwardly. By means of surface mycelium, growth additionally or predominantly 

occurs on the substrate surface, e.g., on nutrient agar or in the case of 

molds that grow superficially on timber and masonry. Aerial mycelium, e.g., in 

the white polypores in buildings (Antrodia spp.), is an intensively developed 

surface mycelium. The texture of the mycelial mat is manifold, e.g., flat on the 

substrate, crusty, woolly, felty, or zonate (Stalpers 1978). 

2.2 

Growth and Spreading 

2.2.1 

Vegetative Growth 

Simplistically, wood fungi live through two functionally different phases: the 

vegetative stage for mycelial spread and the reproductive stage for the elaboration 

of spore-producing structures. Rayner et al. (1985) extended the de- 

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2.2 Growth and Spreading 11 

velopment of a fungus in arrival, establishment, exploitation, and exit. The 

vegetative, asexual stage consists in wood fungi of vegetative hyphae with 

some specialized forms. The reproductive stage can both occur asexually or 

sexually (Schwantes 1996; Table 2.3). 

Functional specialization of the mycelium occurs during the vegetative stage: 

germination, infection, spread, and survival. These functions are correlated 

with different “fungal organs”. Spores (conidia, chlamydospores, also the sexually 

derived asco- and basidiospores) germinate under suitable conditions 

(moisture, temperature). The young germ hypha first shows some nuclei before 

the young mycelium grows with septation in the monokaryotic condition. 

Mycelial growth takes place via mitoses and synthesis of hyphal biomass. Infection 

and colonization of new substrates occurs by spores, hyphae, mycelium, 

and special forms like bore-hyphae, transpressoria, strands, and rhizomorphs. 

Prerequisites for the colonization of a substrate are suitable humidity and nutrient 

availability in the substrate or, like in Serpula lacrymans, theability 

of a fungus to transport nutrients and water and last, whether and by which 

organisms the substrate is already occupied (Rayner and Boddy 1988). Boring 

microhyphae of 0.1–0.4µm diameter develop e.g., in Phellinus pini at the 

hyphal tip without recognizable septum and produce boreholes of 0.3–3.3µm 

diameter probably by enzyme action (Schmid and Liese 1966). The appressorium 

is a hypha for the mechanical fixation to the substrate (Fig. 2.5a). The 

transpressorium (Fig. 2.5b) of the blue-stain fungi (Chap. 6.2) is a specialized 

boring hypha (Liese 1970); it is still unknown whether the penetration 

of the woody cell wall is by mechanical and/or enzymatic action. Transpressoriahavealsobeenfoundinthewhite-rotfungusPhellinus 

pini (Liese and 

Schmid 1966). 

Table 2.3. Functional and morphological differentiation of wood fungi (modified after 

Müller and Loeffler 1992) 

Developmental stage Function “Organ” 

Vegetative/asexual Germination Germ hypha 

Infection, Hypha, mycelium, 

spread boring hypha, 

appressorium, transpressorium, 

strand, rhizomorph 

Survival Chlamydospore, arthrospore, 

mycelia with resistance to 

dryness and heat 

Reproductive/asexual Anamorphic Fruit body, conidiophore, 

reproduction conidium 

Reproductive/sexual Teleomorphic Fruit body, 

reproduction ascus, basidium 

ascospore, basidiospore 

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12 2 Biology 

Fig.2.5. Appressorium and transpressoria of blue-stain fungi in wood. a Hyphae (H) of 

Ophiostoma piceae in the luminina (L) of a pine tracheid. A appressorium, T boring canal 

through the activity of a transpressorium, C wood cell wall. (LM, from Liese and Schmid 

1962); b Two transpressoria (EM, from Liese and Schmid 1966) 

Strands (cords) (Fig. 2.6) develop in a number of house-rot fungi and usually 

consist of three hyphal types, vegetative hyphae, thin fiber (skeletal) hyphae 

with mostly thick walls for strengthening, and broad vessel hyphae for nutrient 

transport (Nuss et al. 1991). These hyphae form a distinct mycelium in the longitudinal 

direction, which is, however, not so well organized like the tissue-like 

structure of the rhizomorphs. Also in contrast to rhizomorphs, strands develop 

behind the mycelial growth front. Particularly Serpula lacrymans overgrows 

larger distances of non-woody substrates and penetrates through masonry 

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Fig.2.6. Hyphae within a strand of Serpula lacrymans. H vegetative hyphae, V vessel hypha, 

F fiber hyphae (dark-field photo W. Liese) 

(only through the joints) between bricks or through old, crumbly bricks, and 

insulation materials. In the laboratory, some house-rot fungi overgrew by 

means of strands agar that contained wood preservatives (Liese and Schmidt 

1976) as well the fungal partner in dual culture. 

In the literature, there is however not always a uniform use of the terms 

“strands (= cords)” and “rhizomorphs”. For example, the strands of the American 

dry rot fungus, Meruliporia incrassata, have been termed rhizomorphs and 

were described as consisting of more or less parallel hyphae, outer (cortical) 

hyphae thick-walled and uniform in size (author: = fibers), inner (medullary) 

thin-walled hyphae, variable in size, and some differentiated into large conducting 

tubes (author: = vessels) (Palmer and Eslyn 1980). According to Burdsall 

(1991) “these two (S. lacrymans, M. incrassata) being similar and unique 

in forming large water-conducting rhizomorphs”. 

By means of his strand diagnosis, Falck (1912) was able to differentiate some 

house-rot fungi like S. lacrymans, Coniophora puteana, and Antrodia vaillantii 

macroscopically and microscopically. Table 2.4 shows an updated version for 

the above tree species based on recent measurements of mycelia in buildings 

andonagarculturesofgeneticallyverifiedisolates. 

As strand morphology is, after fruit body structure, a main feature to identify 

fungigrowingindoorsoranconstructionwood,anidentificationkeyforabout 

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14 2 Biology 

Table 2.4. Strand diagnosis for some common house-rot fungi (modified from Huckfeldt 

and Schmidt 2004, 2006) 

Serpula lacrymans 

Strands white, silver-grey to brown, more than 5 mm to 3 cm diameter, separable, with 

flabby mycelium in between, thick strands when dry breaking with clearly audible 

cracking (strands with mold contamination often not cracking any more), often in 

masonry; (S. himantioides: strands thinner than 2 mm and fibers 2–3.5 µmin 

diameter) 

Vegetative hyphae hyaline, partly yellowish, with large clamp connections, 2–4 µmin 

diameter 

Vessels at least partly numerous (in groups), 5–60 µmindiameter,notorrarely 

branched, with bar thickening up to 13 µmhigh 

Fibers refractive, 3–5 µm diameter, straight-lined, stiff, septa not visible, no clamps, 

lumens often visible 

Coniophora puteana, C. marmorata 

Strands first bright, then brown to black, up to 2 mm wide, to 1 mm thick, root-like, 

hardly removable (not so in C. marmorata), when removed usually fragile, partly 

with brighter center, underlying wood becoming black, also in masonry 

Vegetative hyphae usually without clamps, rarely multiple clamps (often indistinct 

when branched), 2–6 µmindiameter 

Vessels surrounded and interwoven by many fine hyphae (0.5–1.5 µm in diameter), 

difficult to isolate (preparation with KOH solution); drop-shaped, hyaline to 

brownish secretions (1–5 µm in diameter) often on hyphae; vessels due to 

preparation irregularly formed or distorted, up to 30 µm in diameter, thin-walled 

(slightly thick-walled with C. marmorata), without bars, with septa 

Fibers pale to dark brown, 2–4 µm in diameter, somewhat thick-walled, with relatively 

broad, usually visible lumen, some also branched, to be confused with generative 

hyphae 

Antrodia vaillantii, A. serialis, A. sinuosa, A. xantha 

Strands white to cream, partly somewhat yellowing or infected by molds, also ice 

flower-like, flexible also when dry, up to 7 mm in diameter, possibly also within 

masonry 

Vegetative hyphae with few clamps, 2–4 µm in diameter, often somewhat thick-walled; 

in KOH somewhat swelling 

Vessels not rare but in old strands difficult to isolate, up to 25 µm in diameter, thick-walled 

with middle lumen, without bars 

Fibers hyaline (in Antrodia xantha partly somewhat yellowish), numerous, 2–4 µmin 

diameter, hyphal tips with tapering ending cell walls, straight-lined, mostly 

unbranched, insoluble in KOH, but partly somewhat swelling, then with “blown up” 

hyphal segments 

20 strand-forming wood decay fungi based on Huckfeldt and Schmidt (2004, 

2006) is given in Appendix 1. 

The rhizomorphs of Armillaria mellea (Fig. 2.7) are tissue-like mycelial bundles 

with apical growth and consist of a black, gelatinous bark layer, followed 

by a pseudoparenchyma, and a central, loosely interwoven pith with vessel and 

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Fig.2.7. Rhizomorph of Armillaria mellea. Left: Apex with hair-like microhyphae (A), cortex 

(B) andpith(C). (from Hartig 1882); right: cross section (LM; from Schmid and Liese 

1970) 

fiber hyphae (Hartig 1882). By means of rhizomorphs, Armillaria species grow 

in the soil and infect the roots of living trees (Chap. 8.3.1). 

Under unfavorable conditions, resistance stages are formed. Spores are more 

resistant to heat, dryness, and wood preservatives than their mycelium. The hyphal 

cell water content is reduced, nutrients are concentrated, parts of the protoplasts 

or storage substances of neighboring cells are translocated in resting 

cells, and enzyme activity decreases (“latent life”). Chlamydospores (Fig. 2.8) 

are thick-walled spores with a brown cell wall, which occur in many blue-stain 

fungi. 

Formerly, it was believed that the vegetative mycelium of some wood-decay 

fungi is also resistant to dryness (Chap. 3.3) and heat (Chap. 3.4). Recent results 

show that this must not be true: When cultured on agar at about 28 ◦ C, 

the dikaryotic hyphae of Serpula lacrymans tend to revert to the monokaryotic 

condition, which regularly shows abundant arthrospores (Schmidt and 

Moreth-Kebernik 1990). In wood samples that were slowly dried or warmed, 

the substrate mycelium of S. lacrymans, C. puteana, Donkioporia expansa, 

and Gloeophyllum trabeum also formed arthrospores (Huckfeldt 2003). It was 

therefore assumed that these arthrospores are the agents for resistance against 

drying and heat. 

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16 2 Biology 

2.2.2 

Reproduction of Deuteromycetes 

Fungi that reproduce asexually (anamorphic fungi) are either yeasts or Deuteromycetes. 

The term “yeast” is descriptive and stands for any fungus that 

reproduces by budding. 

Deuteromycetes (Fungi imperfecti, colloquially: molds) is an artificial assemblage 

of fungi that reproduce asexually by conidia (conidiospores), either 

as the only form for propagation (imperfect fungi) or additionally (anamorph) 

to a sexual reproduction (teleomorph). When both the anamorph and the teleomorph 

are known, the fungus is called a holomorph (the whole fungus). The 

teleomorph may have one (mono-anamorphic) or many (pleo-anamorphic) 

asexual stages. In other words: Deuteromycetes are the conidia-producing 

forms of a fungus and may or may not be associated with a teleomorph. 

Many Deuteromycetes are supposed to have a teleomorph in the Ascomycetes, 

but they may also have basidiomycetous affinity. Also in the wood-inhabiting 

Deuteromycetes, the teleomorph often is of ascomycetous affinity as in the 

blue stain and soft-rot fungi, but some are anamorphs of Basidiomycetes 

like in the Root-rot fungus, Heterobasidion annosum [anamorph: Spiniger 

meineckellus (A.J. Olson) Stalp.; e.g., Holdenrieder 1989]. In the absence of 

a teleomorph, taxonomic affinity can be detected by the ultrastructure of 

the cell wall: Ascomycetes have two-layered walls, while the walls of Basidiomycetes 

are multilamellar. In terms of strict nomenclature, the teleomorph 

name takes precedence over the anamorph but in practice, a species is often 

identified according to the form in which it was found (Eaton and Hale 

1993), like in the case of the wood-inhabiting molds Aspergillus and Penicillium. 

The Deuteromycetes are usually divided in Coelomycetes and Hyphomycetes. 

Coelomycetes develop conidiophores within fruit bodies (conidiomata). 

In Hyphomycetes (or Moniliales), conidia develop on simple or aggregated 

hyphae. Conidium formation and conidiophore morphology are criteria to 

classify Deuteromycetes (Chap. 2.5). A simplified differentiation for woodinhabiting 

Deuteromycetes (Fig. 2.8) distinguishes between conidiospore (free 

cell fragmentation at the hyphal tip or a branch) and sporangiospore (development 

in a sporangium). 

Conidia of wood-inhabiting Deuteromycetes can be defined as mitotically 

developed (mitospores), immovable, mononuclear to more-nuclear, unicellular 

to more-celled, pigmentless (hyaline) to white, yellow, orange, red, green, 

brown, blue, or black colored (depending on the species) spores of different 

development, size, shape and surface (Fig. 2.9; Reiß 1997; Kiffer and Morelet 

2000). The variety of the spore pigments causes that molded substrates may be 

colorful. 

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Fig.2.8. Generalized view of conidia according to their development. C conidia, S sporangiospores, 

A arthrospores, Ch chlamydospores 

Fig.2.9. Conidia. Example of the manifold shapes and structures 

Fig.2.10. Developmental cycle of 

a deuteromycete. A conidium, B germ 

hypha, C development of conidiophore, 

D development of vesicle, E vesicle with 

conidia 

The series of spore germination, hyphal growth, and conidia production represents 

the asexual reproduction cycle of a deuteromycete fungus, illustrated 

in Fig. 2.10 by an Aspergillus species. 

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18 2 Biology 

The biological advantage of the conidia production to the Deuteromycetes 

(and anamorphs of Asco- and Basidiomycetes) is that these fungi can exit 

from an exploited substrate to arrive fresh nutrients by spores (mitospores) in 

huge numbers without the need of preceding sexuality. Distributed randomly 

by and through the air or by adhering to the surface of animals, spores are 

present everywhere. Disadvantageous is that without (para)sexuality clones of 

an original hypha are distributed. Conidia can develop independently from 

the karyotic stage of the hypha that is anamorphs can occur both on haploid 

and dikaryotic mycelium. 

2.2.3 

Sexual Reproduction 

A specific feature of the sexual reproduction of Ascomycetes and Basidiomycetes 

is that plasmogamy of haploid cells and karyogamy of two nuclei 

(n) to form a diploid nucleus (2n) are separated from each other temporally 

as well spatially by the dikaryophase (two-nuclei phase, dikaryon, n + n, ===) 

(Fig. 2.11). A dikaryotic hypha is one with two nuclei that derive from two 

haploid hyphae, but in which the nuclei are not yet fused by karyogamy. 

Particularly in Basidiomycetes, the dikaryotic phase is considerably extended. 

By conjugated division of the two nuclei (conjugated mitosis), by 

division of the dikaryotic hypha, and by means of a special nucleus migration 

connected with clamp formation both daughter cells become again dikaryotic. 

Fig.2.11. Generalized scheme of nuclear condition of haplo-dikaryotic Ascomycetes and 

Basidiomycetes. → haploid (n), ===> dikaryotic(n+n),=> diploid (2n), P plasmogamy, 

K karyogamy, M meiosis 

2.2.3.1 

Ascomycetes 

The life cycle of a typical ascomycete is shown in Fig. 2.12 (also Müller and 

Loeffler 1992; Eaton and Hale 1993; Schwantes 1996; Jennings and Lysek 1999). 

Haploid (n) spores (A, ascospores or conidia from an anamorph) germinate 

to haploid hyphae and after mitoses to haploid mycelium (B), which is 

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Fig.2.12. Generalized life cycle of an euascomycete. A ascospores or conidia, B germinated 

monokaryons, C plasmogamy of ascogonium (As)-trichogyne (T) and antheridium (An), 

D–G section of ascogonium after incorporation of “male” nuclei, D ascogenous hypha, E 

hook formation, F karyogamy in the tip hypha, G dikaryon and ascus after meiosis, H ascus 

after mitosis with eight ascospores, I anamorph with conidia 

the essential ascomycete with nutrition function and theoretically unlimited 

growth. Conidia may develop at the haploid mycelium as anamorph (I). 

Within the fruit body, hyphae develop to gametangia (“sexual organs”, C) 

connectedwithmitosis.Thetrichogyne(T,“copulationhypha”),whichderives 

from the ascogonium (As, “female gametangium”), fuses (plasmogamy, gametangiogamy) 

with the antheridium (An, “male gametangium”). The nuclei 

fromtheantheridiummigrate(therefore:male)throughthetrichogyneintothe 

ascogonium. There may be various modifications of the generalized scheme: 

Antheridia are absent, and mono-nuclear spermatia (from an anamorph) fuse 

with the trichogyne (deuterogamy). Somatogamy of “normal” hyphae takes 

place (see Chap. 2.2.3.2). One sex is missing or not functional, and fertilization 

occurs between two nuclei of the same sex (automixis). 

In the hymenial Ascomycetes (Ascohymeniales, wood-inhabiting Ascomycetes), 

the fruit bodies (ascocarps, ascomata) develop after the fertilization of 

the ascogonium from basal cells of the gametangia, and thus the fruit bodies 

predominantly consist of haploid hyphae (Fig. 2.13). 

From the “pollinated” ascogonium, ascogenous hyphae develop, into which 

migrates each one pair of two genetically different (compatible) nuclei. In 

Ascomycetes, the dikaryotic phase is limited and without nutrition function. 

By means of hook formation (Fig. 2.12E) the short-lived hook mycelium and 

the ascus (meiosporangium) develop, in which karyogamy and meiosis occur. 

Before ascospore formation, there is commonly an additional mitosis, which 

bringsthenumberofascospores(meiospores)intheascustoeight.Themature 

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20 2 Biology 

Fig.2.13. Structure of a fruit body (apothecium) of an ascomycete predominantly consisting 

of haploid hyphae (thin lines,onenucleus),somedikaryotichyphae(thick lines,twonuclei) 

and differently matured asci within the hymenium. As-, An ascogonium and antheridium 

before gametangiogamy, As+ fertilized ascogonium 

ascus is usually tube-shaped (“tube fungi”). The non-flagellate ascospores 

disper after disintegration of the ascus or via different opening mechanisms. 

The ascospores are mono-nuclear or after further mitosis multi-nuclear. They 

can be septate and show similar conidia characteristics of size, shape, color 

and wall sculpturing. 

The relatively small fruit bodies (less than 1 mm in diameter) of the woodinhabiting 

Ascohymeniales are the spherically closed cleistothecium, the pear- 

Fig.2.14. Fruit body types of Ascomycetes. P perithecium, A apothecium, C cleistothecium 

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shapedperithecium, e.g., inseveral blue-stainfungi, orthe disk-shapedapothecium 

(Fig. 2.14). 

2.2.3.2 

Basidiomycetes 

ThelifecycleofatypicalbasidiomyceteisschematicallyrepresentedinFig.2.15. 

The haploid basidiospore or conidium (A) germinates to the n-mycelium 

(B, monokaryon, primary mycelium). There are also asexual anamorphs in 

Basidiomycetes. According to Müller and Loeffler (1992), asexual anamorphs 

are supposed to occur almost just as frequently as in Ascomycetes: “they are 

named however only rarely with an own name, therefore hardly considered in 

the system of the Deuteromycetes and would be more frequent in the dikaryotic 

phase”. A known example among the wood-decay Basidiomycetes is Heterobasidion 

annosum with its anamorph Spiniger meineckellus. 

In the laboratory, monokaryons are capable of indefinite growth if they 

are regularly subcultured on fresh medium. In nature, characteristically the 

dikaryon or secondary mycelium develops. Basidiomycetes do not form sexual 

organs for plasmogamy, but monokaryotic hyphae come into contact one with 

another and fuse by somatogamy (C). If the nuclei are compatible, the dikaryon 

develops (D). This long-lived mycelium (Schwantes 1996) represents the essential 

basidiomycete that penetrates the substratum and absorbs nourishment, 

inthecaseofwoodfungiwithwood-decayfunction(D–G).Inabouthalfofthe 

Basidiomycetes, the dikaryon grows by clamp connections (clamp mycelium): 

Fig.2.15. Generalized life cycle of a homobasidiomycete. A basidiospores or conidia, B 

monokaryons after germination, C somatogamy, D dikaryon, E–G clamp formation, H–K 

basidium development, I karyogamy, J meiosis, K basidium with four basidiospores located 

in sterigmata 

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22 2 Biology 

A short branch arises on the side of the apical hypha and bends over. After 

synchronous (“conjugate”) division of the two nuclei (E), two daughter nuclei 

remain in the apical cell, one nucleus migrates into the branch (F), the branch 

end fuses with the subapical cell, and by septum formation, two dikaryotic 

hyphae have developed (G). Repeated conjugate divisions accompanied by 

septum formation result in an extensive dikaryotic mycelium (Jennings and 

Lysek 1999). Sometimes there are double or multiple (whorl) clamps (maximally 

eight) around one septum, e.g., in Coniophora puteana (four clamps). In 

a second method of dikaryotization, there is a division of the nuclei in the binucleate 

hypha followed by a migration of the daughter nuclei into the primary 

myceliumoftheoppositematingtype.Theforeignnucleusineachmycelium 

dividesanditsprogenymigratefromhyphatohyphathroughtheseptalpores 

until both parent mycelia have been dikaryotized (Alexopolus and Mims 1979). 

Depending on external factors, like season (temperature, air humidity), nutrients 

and light, large fruit bodies (tertiary mycelium, basidiocarp, basidioma) 

develop on the secondary mycelium (Fig. 2.16). 

In the fruit body of the hymenomycetes, the hymenium (fertile layer) develops 

(Fig. 2.16), in which the formation of basidia occurs (Fig. 2.15H–K). For 

Fig.2.16. Life cycle of a wood-decay basidiomycete. A haploid spores, hyphae, somatogamy 

and dikaryotic growth in the soil, B infection of the tree through a wound, C tree deterioration 

by the dikaryon, D fruit body formation (bracket); in the hymenium: E karyogamy, 

F, G meiosis, H mature basidium with four basidiospores 

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surface enlargement the hymenium may be e.g., net-like arranged (merulioid, 

S. lacrymans), warted (C. puteana), porous (A. vaillantii), or lamellate (Armillaria 

mellea). In the young basidium (Fig. 2.15H), karyogamy (I) and meiosis 

(J)occur.Fourhaploidnucleimigrateintooutgrowths(sterigmata)atthetop 

of the basidium (K) and are discharged as basidiospores. 

The spore size (5–20µm), shape (globose, cylindric, ellipsoid etc.), surface 

sculpturing (“ornamentation”: warted, crested, etc.), wall-thickness (thinwalled, 

double wall) (Ryvarden and Gilbertson 1993) and color (colorless or 

pigmented: white, yellow, orange, ochre, pink, brown, green, violet, black) 

are taxonomic characteristics. In the microscope, spores appear frequently 

bright to colorless (hyaline), e.g., in Daedalea quercina, Fomes fomentarius, 

H. annosum, Laetiporus sulphureus, Piptoporus betulinus and Trametes versicolor. 

Brownish spores separate e.g., the genus Serpula from other fungi with 

merulioid hymenium (Pegler 1991). Further characteristics are the violetstaining 

of amyloid spores (e.g., Stereum sanguinolentum)andthebrown-red 

staining of dextrinoid spores by JJK as well as the blue-staining of cyanophilic 

spores (C. puteana, H. annosum, Oligoporus placenta) by aniline blue (e.g., Erb 

and Matheis 1983). 

For the differentiation of the various fruit body types serve, e.g., the occurrence 

of sterile cells (cystidia) between the basidia (e.g., Antrodia spp. 

and Gloeophyllum spp.) and the construction of basidiocarps consisting of 

vegetative hyphae (monomitic), additionally of either skeletal or binding hyphae 

(dimitic) or of all three hyphal types (trimitic). Monomitic genera are 

Coniophora, Meripilus and Phaeolus, dimitic are Antrodia, Heterobasidion, 

Hirschioporus, Laetiporus and Phellinus, and trimitic are Daedalea, Fomes and 

Trametes. 

Most wood-inhabiting Basidiomycetes belong to the Homobasidiomycetes 

(formerly Holobasidiomycetes: single-celled basidium) and there to the Aphyllophorales 

with gymnocarpous (hymenium exposed while spores are still im- 

Fig.2.17. Common types of fruit bodies of wood-inhabiting Basidiomycetes. a Pileate with 

central stipe (Lentinula edodes cultured on wood by J. Liese in 1935). b Bracket-like (Piptoporus 

betulinus on a birch tree, photo T. Huckfeldt). c Resupinate (Serpula lacrymans in 

a building) 

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24 2 Biology 

mature) and non-lamellate fruit bodies. The Aphyllophorales show a number 

of different types of fruit bodies whose attachment to the substrate may also 

be rather distinctive: stalked, coral-like, club-like, bracket-like, resupinate, etc. 

(Ryvarden and Gilbertson 1993, 1994; Schwantes 1996). Simplistically, fruit 

bodies may be grouped into pileate with central stipe, bracket-like, and resupinate 

(Fig. 2.17). Fruit bodies may be annual (passing after spore discharge), 

biannual or perennial (every year new hymenial layers laid on the preceding 

ones). 

2.2.4 

Fruit Body Formation 

Fruit body initiation and development that occurs usually outside of the substrate 

are affected by various exogenous factors: humidity, temperature, light, 

nutrition, force of gravity, composition of air, and interactions with other 

organisms (Schwantes 1996). Endogenous factors cover the participation of 

phenol oxidases and other enzymes, cyclic adenosine monophosphate (AMP) 

and genes. Fruit body formation is often promoted by conditions, e.g., warmth 

in S. lacrymans, which are unfavorable for the vegetative development. 

In fungi that are not tolerant to dryness, like Pholiota and Pleurotus species, 

the fruit bodies frequently have a fleshy consistency and lose when drying 

their function irreversibly, so that in the northern hemisphere many forest 

fungi with annual fruit bodies preferentially fructify in damp-cool weather 

in the autumn. Dry-tolerant fruit bodies, like in Schizophyllum commune, 

continue spore production under humid conditions after dryness for many 

years. Others reduce the evaporation by hairy or “varnished” surfaces, like 

Inonotus and Ganoderma species. The concentric zonation of the pileus surface 

(rough and smooth in the change) of Trametes versicolor is influenced by 

humidity variation and the different colors of the individual zones by light and 

dark phases (Williams et al. 1981). In Coprinus comatus, fruit body primordia 

do not develop further without light (Jennings and Lysek 1999). Short-wave 

light (UV, blue) may influence fruit body development (Schwantes 1996). The 

Oyster fungus, Pleurotus ostreatus, only fruits below 16 ◦ C (Chap. 9.2), and 

the less tasty subspecies P. ostreatus ssp. florida at a higher temperature. Fruit 

bodies of the Winter fungus, Flammulina velutipes, appear also after snowfall. 

Serpula lacrymans fruits in the laboratory after a stimulating pretreatment of 

the mycelium for 3–4 weeks at the submaximal temperature of 25 ◦ C (Schmidt 

and Moreth-Kebernik 1991b; Chap. 3.4). Lentinula edodes is stimulated during 

its cultivation on wood in Asia by a cooling treatment. Schizophyllum commune 

fruits already on simple nutrient agar at room temperature. Gloeophyllum 

trabeum (Croan and Highley 1992a) and L. edodes (Leatham 1983) fructified 

on defined growth media. AMP was suitable for a Coprinus species (Uno 

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and Ishikawa 1973). Yeast extract, vitamin B1, traumatic influences through 

physical distortions to the mycelium, and the presence of another fungus or its 

mycelial extract or culture filtrate may be favorable (Stahl and Esser 1976; Leslie 

and Leonard 1979; Matsuo et al. 1992; Kawchuk et al. 1993). In S. commune,the 

development of a fruit-body-inducing substance (FIS) is genetically controlled 

(Leslie and Leonard 1979). In a Polyporus species, there are fi+ genes (fruiting 

initiation) (Stahl and Esser 1976; Esser 1989). The force of gravity determines 

that the yearly hymenial layers in the bracket-like, perennial fruit bodies of 

Fomes fomentarius also point to the earth center if the host tree is lying on the 

ground (Chap. 3.6). 

2.2.5 

Production, Dispersal and Germination of Spores 

Spores represent in the life cycle of a fungus a state of rest (low water content, 

high nutrient content; “latent life”) between the active phase of spore dispersal 

and start of new growth. 

Serpula lacrymans produces 300,000 (Falck 1912) to 360,000 (Rypáček 1966) 

and Piptoporus betulinus 31,000,000 (Kramer 1982) spores per hour and cm 2 of 

hymenium. Many forest mycorrhizal fungi fruit at higher air moisture content 

and lower temperature in the autumn. Among the tree parasites, Heterobasidion 

annosum disperse spores almost over the whole year, Laetiporus sulphureus 

in the autumn. 

Many Basidiomycetes disperse their spores actively for 0.1–0.2 mm (ballistospores) 

so that the spores more easily reach the open air (Schwantes 1996). 

In Schizophyllum commune, a liquid drop at the sterigma becomes larger and 

hurls the spore into the airflow (Müller and Loeffler 1992). Möykkynen (1997), 

using a wind tunnel, measured for the conidia of Heterobasidion annosum that 

a threshold speed of an airflow of 1.8 m/s liberates the spores. 

Falck (1912) calculated the mass of a spore of S. lacrymans as 171 × 10 −12 g. 

Fungal spores exhibit a density of 1.1 dp. In standing air, spores sink with 

sedimentation speeds of 0.03–0.55 cm/s (Reiß 1997). A continuously colonized 

area can expand 50 km over the year. In an appropriate air stream, spores 

can be transported up to 1,000 km (Burnett 1976). Furthermore, spores are 

spread by rain and snow. Animals distribute spores that are attached by the 

spore surface sculpturing (see Fig. 2.9) or remain indigested. Assumably by 

international trade, the causal agent of the Dutch Elm disease, Ophiostoma 

ulmi, was imported from Asia to Europe in 1918 (Chap. 8.1.2.1). 

The spore content in air is subject to characteristic rhythms. In Central 

Europe, it is higher in the summer at warm temperatures and low relative 

air humidity than in the winter. Basidiospores and ascospores are numerous 

in the air in spring and in autumn. Conidia have a maximum from June 

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26 2 Biology 

to September. In cities in temperate regions, the spore concentration of Cladosporium, 

mainly C. herbarum, often rises up to 10,000–15,000 spores/m 3 

air with peaks of more than 50,000 spores/m 3 (Nolard 2004). Air turbulence 

during stable areas of high pressure may result in daily rhythms, the concentration 

rising during the midday (Reiß 1997). Interiors with high dust content 

(e.g., the wood-processing industry) may exhibit increased spore contents. The 

lifespan of spores in free air is affected by temperature, air humidity, and sun 

exposure. As unpigmented spores are sensitive to UV light, pigmented spores 

predominate in the air. Exogenously dormant spores only germinate when the 

environmental conditions (nutrients, temperature, pH value) become favorable. 

Endogenously dormant spores fail to germinate even under favorable 

conditions, which is due to factors within the spore such as nutrient impermeability 

or the presence of endogenous inhibitors. Dormancy within these 

spores is broken by ageing when nutrients begin to enter or the inhibitors leach 

out (Robson 1999). 

Priortotheemergenceofoneormoregermtubes,sporesundergoaprocess 

of swelling during which they increase in diameter due to the uptake of water. 

The metabolic activity, production of protein, DNA and RNA all increase. 

The percentage of germinating spores depends on fungal species, spore age, 

temperature, available moisture, and substrate. In Serpula lacrymans, only 30% 

of sampled spores germinated in vitro (Hegarty et al. 1987). For the conidia of 

Heterobasidion annosum, the thermal cardinal points were 0 ◦ C minimum, between 

12 and 28 ◦ C optimum and 34 ◦ C maximum (Courtois 1972). Depending 

on the species, the duration of the germination ability reaches from a few days 

or weeks, like in Stereum species, to several years in Chaetomium globosum, 

and can reach up to about 20 years in S. lacrymans (Grosser et al. 2003). 

Germination of spores of wood fungi is favored by high air humidity, 

warmth, and pH values of 4–6. In Serpula lacrymans, citricacid(Hegarty 

et al. 1987) and vitamin B1 (Czaja and Pommer 1959) stimulated germination. 

Heartwood compounds may inhibit. 

2.3 

Sexuality 

The wood-inhabiting Ascomycetes and Basidiomycetes are either homothallic 

or heterothallic (Ryvarden and Gilbertson 1993). 

Homothallic fungi are self-fertile, that is no second mating type is required 

for sexual reproduction. Fertilization takes place at the same mycelium. Many 

Ascomycetes and about 10% of the Basidiomycetes belong to this type. 

Heterothallism includes both bipolar and tetrapolar fungi. In bipolar (unifactorial) 

species, incompatibility is controlled by a series of multiple alleles at 

one locus. Any dikaryon has two alleles that segregate at meiosis so that half 

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2.3 Sexuality 27 

the basidiospores have one allele and half the other. Compatible matings occur 

between monokaryons with different mating type factors. The inbreeding level 

is 50%. The outbreeding level in populations of bipolar polypores is over 90%. 

In tetrapolar (bifactorial) species, incompatibility is controlled by two series 

of multiple alleles at two loci on different chromosomes. The two pairs 

segregate independently at meiosis. Four different mating types rise from one 

dikaryon. In a fruit body of an isolate, basidiospores of the mating type AxBx, 

AxBy,AyBx and AyBy develop. These spores germinate to monokaryons. Fully 

compatible matings of monokaryons (+ in Table 2.5) occur when both factors 

are heterozygous (A#B#): AxBx and AyBy as well as AxBy and AyBx. In addition, 

there are hemicompatible matings, in which only one factor is different: AxBx 

and AxBy as well as AyBy and AxBy. 

The inbreeding level is 25%. The outbreeding level is very high. In Schizophyllum 

commune 450 A factors and 90 B factors can combine to over 40,000 

mating types (Raper and Miles 1958). Many Ascomycetes and about 25% of 

the examined Basidiomycetes are bipolar heterothallic (e.g., Oligoporus placenta). 

About 65% Basidiomycetes are tetrapolar (Raper 1966). Bipolar mating 

predominates among brown-rot fungi and tetrapolar mating among white-rot 

fungi (Rayner and Boddy 1988). Of 25 investigated brown-rot polypores, 17 

were bipolar, three were tetrapolar, three were heterothallic with type of mating 

system undetermined, one was homothallic, and one was reported by different 

authors as bipolar and tetrapolar (Ryvarden and Gilbertson 1993). The biological 

significance of heterothallism is that inbreeding is limited and outbreeding 

is enhanced, promoting gene flow between populations and decreasing the rate 

of speciation. 

Combination and recombination of the genetic material with plasmogamy, 

karyogamy, and haploidization, but without sexual organs, gamets and changes 

of generations, can take place by parasexuality, particularly in Deuteromycetes 

(Jennings and Lysek 1999). Nuclei of a hypha migrate by anastomosis into 

another hypha and multiply and spread there. In the case of a heterokaryon, 

Table 2.5. Mating scheme of tetrapolar heterothallic fungi 

AxBx AxBy AyBx AyBy 

AxBx − A B + 

AxBy A − + B 

AyBx B + − A 

AyBy + B A − 

− incompatible (A=B=), + compatible (A#B#) 

A common-A heterokaryon (A=B#): conjugate nuclear division and clamp formation 

blocked, variable nucleus content per hypha, 

B common-B heterokaryon (A#B=): nuclear migration and clamp cell fusion blocked (“false 

clamps”) 

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28 2 Biology 

some nuclear fusions and after haploidization new combinations occur. Usually, 

the diploid nuclei are unstable, and their ploidy number is regulated to the 

haploid stage by elimination of chromosomes or discharge of sections (Müller 

and Loeffler 1992). 

Illustrated by the tetrapolar heterothallic Serpula lacrymans,interstockmating 

of ten isolates is demonstrated in Table 2.6 (Schmidt and Moreth-Kebernik 

1991c). First, the four different mating type monokaryons of each isolate were 

obtained after fruit body stimulation (Schmidt and Moreth-Kebernik 1991b) 

and subsequent inbreeding according to Table 2.5. Then the 10×4 monokaryons 

were paired one with another in all possible combinations on agar. As in S. 

lacrymans, like in many Basidiomycetes, only the dikaryon forms clamps, it 

can be detected in the light microscope. In contrast, only the monokaryons of 

the fungus show abundant arthrospores. The heterokaryons of the type A=B# 

and the “false clamps” mycelia (A# B=) can be recognized from the mating 

diagram by calculation or by further pairings. The mating types of the isolate 

monokaryonsareshownintheuppertablepart. 

The mycelium of the F1-dikaryons of S. lacrymans grew faster at about 20 ◦ C 

than that of the two parental monokaryons (Schmidt and Moreth-Kebernik 

1991a), like this applies also to Lentinula edodes (Schmidt and Kebernik 1987) 

and Stereum hirsutum (Rayner and Boddy 1988). Thus, dikaryotic mycelium, 

which grows out from compatible monokaryons, looks like a bow tie (Fig. 2.18), 

that is, dikaryons can usually be detected macroscopically. 

In a sample of ten isolates, theoretically 20 different A and B factors can 

occur. In the S. lacrymans sample, there were however only four A and five 

B factors. This limited number of mating alleles contrasts with the regular 

observation of a high number of mating alleles in other Basidiomycetes (May 

et al. 1999) and indicated that S. lacrymans has a narrow genetic base. 

Fig.2.18. Bow tie-like outgrowth of the 

faster growing dikaryon (D) ofSerpula 

lacrymans from the slowly growing 

monokaryons (M) 

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2.3 Sexuality 29 

Table 2.6. Pairings among the four mating types of ten isolates of Serpula lacrymans (from 

Schmidt and Moreth 1991b) 

The matings in S. lacrymans have also shown some physiological differences 

between the mycelia of the different nuclear types. However, there was also consistency 

over the generations (parents as well as F1 and F2 generation), namely 

with regard to the growth rate, wood decay ability, as well as temperature and 

preservative tolerance (Chap. 8.5.3.4). 

Interfertility/intersterility tests are a useful criterion for the identification 

of unknown isolates and for separation of very similar species. Mating of 

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30 2 Biology 

a haploid mycelium with defined tester strains whose species affiliation is 

known only results in a dikaryotic/diploid mycelium if the isolate belongs to 

the same species. Mating is also possible between dikaryotic/diploid mycelium 

and monokaryotic/haploid mycelium (Buller phenomenon) in this way that 

one nucleus of the dikaryon enters a monokaryon of the same species. Complete 

absence of interfertility between monokaryons of the True dry rot fungus, S. 

lacrymans, and the Wild dry rot fungus, Serpula himantioides, (Harmsen 

et al. 1958) showed that both fungi are independent species. That is the True 

dry rot fungus should no more described as domestic variant of the wild 

species adapted to buildings, which was later confirmed by DNA techniques 

(Chap. 2.4.2.2). 

Intersterility must be approached cautiously, however, because intersterile 

populations (intersterility groups, ecotypes) occur that cannot be separated 

morphologically. For example, in Heterobasidion annosum (Chase and Ullrich 

1990), monokaryons isolated from fruit bodies sampled in pine forests 

(P-isolates) did not pair with isolates from spruce trees (S-isolates) (Korhonen 

1978a), and F-isolates were specialized for fir (Capretti et al. 1990). The different 

groups have been recently attributed to three distinct species (Niemelä and 

Korhonen 1998). Comparably, the five intersterility groups A, B, C, D, E within 

the annulate Armillaria mellea complex (Korhonen 1978b) were assigned to 

five biological species (Guillaumin et al. 1993). For edible mushrooms of the 

Pleurotus species, three North American, eight European, and five Asian intersterility 

groups have been found (Bao et al. 2004a). 

Another genetic system referred to as somatic or vegetative incompatibility 

restricts plasmogamy between genetically different heterokaryotic dikaryons. 

In 1929, Fomitopsis pinicola was the first basidiomycete to be studied by means 

of somatic incompatibility (cf. Högberg et al. 1999). The somatic incompatibility 

system can be defined as the rejection of nonself mycelia following hyphal 

anastomosis (Worrall 1997), thus assuring the isolation of unrelated individuals 

in nature. Cultures of the same genotype form a common mycelium, while 

cultures of different genotypes of a species or of different species separate themselves 

by a demarcation zone. Two isolates are incompatible if they carry different 

alleles at one or more vic loci. Self/nonself recognition is normally related 

to genetic uniqueness (Hansen and Hamelin 1999). Thus, there is a correspondence 

between the delimitation of genets by DNA fingerprints and vegetative 

compatibility tests. In some Basidiomycetes, however, vegetatively compatible 

isolates are not necessarily genetically identical or similar individuals, clones 

or genets, but closely related genets by chance may share similar vegetative 

compatibility alleles and do not recognize self from nonself. Kauserud (2004) 

grouped the European isolates of S. lacrymans into five widespread vegetative 

compatibility groups (VCGs). Due to low genetic variation of the fungus, the 

VCGs may not represent clones or inbred lineages, but rather different genets 

by chance share similar vic alleles (Kauserud et al. 2004a). 

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2.4 Identification 31 

In addition to the pairing of compatible monokaryons to insert genetic 

material in a fungus from another isolate, fusion of fungal protoplasts can be 

performed. Protoplast fusion can be used to make hybrids between cells of the 

same mating type, as well as of dikaryotic cells or even between species and genera. 

Spheroplasts (cell wall partially removed by lysing enzymes) or protoplasts 

(cell wall completely removed) fuse by electric influence or through osmotic 

active substances (polyethylene glycol) and some of them regenerate to new 

hyphae. Protoplast fusion is used for genetic studies as well as for isolate improvement. 

Experiments on wood fungi comprise, e.g., Auricularia polytricha, 

Gloeophyllum trabeum, Heterobasidion annosum, Lentinula edodes, Oligoporus 

placenta, Ophiostoma piceae, O. ulmi, Phanerochaete chrysosporium, Pleurotus 

ostreatus, Trametes versicolor, and Trichoderma spp. (Nutsubidze et al. 1990; 

Trojanowski and Hüttermann 1984; Royer et al. 1991; Sunagawa et al. 1992; 

Rui and Morrell 1993; Richards 1994; Tokimoto et al. 1998; Bartholomew et al. 

2001; Xiao and Morrell 2004). Interspecific fusions (Toyomasu and Mori 1989; 

Eguchi and Higaki 1992) and intergeneric fusions (Liang and Chang 1989) were 

reported. With increasing genetic distance of the fusion partners, however, the 

hybrids are instable, do no fruit, die, or the obtained fruit bodies correspond to 

one of the parents, that is, obviously one of the two nuclei has been eliminated 

before. 

Protoplasts have been produced from O. piceae with the aim of subsequently 

inserting genetic material capable of producing fluorescent proteins to allow visualization 

of hyphae of that species in wood by using fluorescence microscopy 

(Xiao and Morrell 2004). 

2.4 

Identification 

2.4.1 

Traditional Methods 

Determination keys and descriptions for Deuteromycetes are based on morphology, 

color, and development (conidiogenesis) of conidia and conidiogenous 

cells (Figs. 2.8–2.10) (Carmichael et al. 1980; Domsch et al. 1980; v. Arx 

1981; Wang 1990; Hoog and Guarro 1995; Schwantes 1996; Kiffer and Morelet 

2000; Samson et al. 2004). 

The fruit bodies of Ascomycetes and Basidiomycetes serve to identify species 

on the basis of macro- and microscopic characteristics using keys or illustrated 

books: Kreisel 1961; Domański 1972; Domański et al. 1973; Breitenbach and 

Kränzlin 1981, 1986, 1991, 1995; Moser 1983; Jülich 1984; Hanlin 1990; Jahn 

1990; Wang and Zabel 1990; Ryvarden and Gilbertson 1993, 1994; Huckfeldt 

and Schmidt 2005; yeasts: Barnett et al. 1990). There are identification kits 

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32 2 Biology 

for yeasts that employ assimilation tests of carbohydrates with a specifically 

adapted database, and also growth tests on carbon sources that are bound to 

a tetrazolium dye (Mikluscak and Dawson-Andoh 2005). An illustrated key for 

wood-decay fungi is in the Internet (Huckfeldt 2002). 

For wood-inhabiting Basidiomycetes, of which only mycelium is present, 

keys are based on microscopic characteristics of the hyphae and on growth parameters 

(Davidson et al. 1942; Nobles 1965; Stalpers 1978; Rayner and Boddy 

1988; Lombard and Chamuris 1990). Among the physiological characteristics, 

the Bavendamm test for the differentiation of brown- and white-rot fungi is 

based on the presence/absence of the phenol oxidase laccase (Bavendamm 

1928; Davidson et al. 1938; Käärik 1965; Niku Paavola et al. 1990; Tamai and 

Miura 1991; Chap. 4.5). Specific reactions to temperature (Chap. 3.4) provide 

further information. However, keys for mycelia are unable to differentiate 

closely related fungi such as the various Antrodia and Coniophora species. The 

strand diagnosis of Falck (1912; Table 2.4, Figs. 8.19–8.21) differentiates few 

indoor decay fungi like Serpula lacrymans, Coniophora puteana and Antrodia 

vaillantii. As house-rot fungi are the economically most important wood fungi 

by destroying wood during its final use within buildings and as not all indoor 

fungi fruit, a key including about 20 strand-forming indoor wood decay fungi 

(Huckfeldt and Schmidt 2004, 2005, 2006) is given in Appendix 1. 

In addition, there are monographs and descriptions of important tree 

pathogens (e.g., Ceratocystis and Ophiostoma species: Upadhyay 1981; Wingfield 

et al. 1999; Armillaria species: Shaw and Kile 1991; Heterobasidion annosum: 

Woodward et al. 1998) and of wood-degrading Basidiomycetes (Cockcroft 

1981; Ginns 1982) with data to taxonomy, morphology, ecology, growth behavior, 

and wood degradation in the laboratory and outside. A further possibility 

for identification is by national institutions against fee (Table 2.7). 

A list of collections and institutions with strain collections, compiled by 

German Collection of Microorganisms and Cell Cultures, is in the Internet 

(www.dsmz.de/species/abbrev.htm). Sixty-one culture collections in 22 European 

countries are united in the European Culture Collections’ Organisation 

(ECCO; www.eccosite.org). The World Federation of Culture Collections 

(WFCC; www.wfcc.info/index.html) is a worldwide database on culture resources 

comprising 499 culture collections from 65 countries. 

Table 2.7. Examples of institutions for identification, deposition, and purchasing of microorganisms 

German Collection of Microorganisms and Cell Cultures (DSMZ), Braunschweig 

Centraalbureau voor Schimmelcultures (CBS), Baarn, Netherlands 

International Mycological Institute (IMI), Kew, UK 

Belgian Coordinated Collections of Microorganisms (BCCM), Gent 

American Type Culture Collection (ATCC), Rockville 

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2.4.2 

Molecular Methods 

Molecular methods to characterize, identify, and classify organisms do not 

depend on the subjective judgment of a human being as it might occur using 

classical methods, but are based on the objective information (molecules) 

deriving from the target organism. Thus, molecular methods are increasingly 

used to identify organisms and for taxonomy research (molecular systematic). 

In the 1980s, molecular methods were established for wood-decay and staining 

fungi. Mainly, the fungal proteins (enzymes) and nucleic acids are used. It is 

outside the intention of this book to describe all molecular techniques that are 

currently used in the field of biology. The following overview comprises only 

some methods and results that are related to the characterization, identification, 

and phylogeny of wood-inhabiting fungi, particularly wood-decay fungi. 

Genome sequencing (meanwhile over 100 genomes are sequenced), molecular 

engineering, cloning, etc. are briefly addressed in other chapters. As an example 

of the latter, Lee et al. (2002) transformed the wild-type and the albino 

strain of the blue-stain fungus Ophiostoma piliferum with a green fluorescent 

protein (GFP) to microscopically differentiate the GPF-expressing fungi from 

other fungi in wood. 

2.4.2.1 

Protein-Based Techniques 

SDS polyacrylamide gel electrophoresis (SDS-PAGE) 

In SDS-PAGE, the whole cell protein is extracted from fungal tissue, denatured, 

and negatively charged with mercaptoethanol and sodium dodecyl sulfate 

(SDS). The proteins are separated according to size on acrylamide gels and 

visualized by Coomassie blue, amido black, fast green, imidazole-zinc or silver 

staining. The banding pattern obtained discriminates at the species level and 

slightly below. 

SDS-PAGE was used for wood-inhabiting Ascomycetes and Deuteromycetes 

like the Cancer stain disease fungus of plane, Ceratocystis fimbriata f. platani, 

(Granata et al. 1992) and Trichoderma species (Wallace et al. 1992). 

The technique also differentiated a number of wood-decay fungi (Schmidt 

and Kebernik 1989; Vigrow et al. 1989, 1991a; Schmidt and Moreth-Kebernik 

1991a, 1993; Palfreyman et al. 1991; McDowell et al. 1992; Schmidt and Moreth 

1995). For example, the closely related Serpula lacrymans, S. himantioides and 

the “American dry rot fungus”, Meruliporia incrassata, weredistinguished 

(Schmidt and Moreth-Kebernik 1989a). Figure 2.19 shows that the technique 

also detected a misnamed isolate of S. lacrymans. 

In addition, monokaryons and F1 dikaryons of S. lacrymans exhibited the 

typical species profile (Schmidt and Moreth-Kebernik 1990). There was no need 

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34 2 Biology 

Fig.2.19. Protein bands of Serpula lacrymans isolates (S) after SDS polyacrylamide gel 

electrophoresis. H false naming later identified as S. himantioides (from Schmidt 2000) 

to extrapolate on a possible influence of culture age or medium composition 

(Schmidt and Kebernik 1989). 

SDS-PAGE is fast when the sample originates from a pure culture and can 

be performed within 1 day. Reproducible homemade gels require accuracy 

and precautions, as acrylamide is carcinogenic in the unpolymerized form. 

Prefabricated gels are expensive. At least regarding wood-decay fungi, the 

method did not reach a practical application. 

Isozyme analyses 

Isozyme analyses have been used to distinguish similar and closely related 

species and forms, for investigations on the genetical variability and on the 

spread of pathogens (e.g., Blaich and Esser 1975; Prillinger and Molitoris 1981; 

Micales et al. 1992). Being functional proteins, isozymes are investigated by 

native electrophoresis or isoelectric focusing. There are a number of investigations 

on mycorrhizal fungi, e.g., Pisolithus and Scleroderma species (Sims 

et al. 1999) and on tree parasites, like Armillaria species (Bragaloni et al. 1997) 

and Heterobasidion annosum (Karlsson and Stenlid 1991). 

Two-dimensional gel electrophoresis, comprising isoelectric focusing and 

subsequent SDS-PAGE, is able to separate a sample of a large number of proteins. 

Immunological methods 

Wood fungi can be also detected and identified by immunological (serological) 

methods. Immunological assays use polyclonal antisera or monoclonal 

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antibodies. Antisera produced by animals like mice and rabbits as answer 

to the injection of mycelial fragments, extracts or culture filtrates are investigated 

by Western blotting, enzyme-linked immunosorbent assay (ELISA) 

or immunofluorescence (Clausen 2003). However, the experiments often exhibit 

cross-reactions with non-target organisms, even when monoclonal antibodies 

after fusion with myeloma cells (hybridomas) are used. Investigations 

were performed with e.g., Armillaria spp., Coniophora puteana, Gloeophyllum 

trabeum, Lentinula edodes, Lentinus lepideus, Oligoporus placenta, Phellinus 

pini, S. lacrymans, Trametes versicolor and with wood-stain fungi (Jellison and 

Goodell 1988; Palfreyman et al. 1988; Breuil et al. 1988; Glancy et al. 1990; 

Burdsall et al. 1990; Vigrow et al. 1991b, 1991c; Clausen et al. 1991, 1993; Kim 

et al. 1991a, 1991b, 1993; Toft 1992, 1993; McDowell et al. 1992; Clausen 1997a; 

Breuil and Seifert 1999; Hunt et al. 1999). 

The diagnostic potential lies in the identification of species without the 

need of preceding isolation and pure culturing and in the detection of fungi 

at early stages of decay (Clausen and Kartal 2003). The methods may become 

applicable when the producing techniques for hybridomas and diagnostic kits 

have been established. 

Immunological methods were also used to visualize the distribution of 

enzymes of wood-degrading fungi within and around the hypha and in woody 

tissue (e.g., Kim 1991; Kim et al. 1991a, 1993; Chap. 4). 

2.4.2.2 

DNA-Based Techniques 

Southern blotting of restriction fragments (RFLPs) 

In the RFLP technique, nuclear, mitochondrial or chloroplast DNA is treated 

with endonucleases, which each have a short nucleotide recognition site on 

the DNA target, and which cut the DNA into fragments. The fragments are 

separated on agarose gels and transferred by Southern blotting on nitrocellulose 

or nylon membranes. The addition of a special nucleotide probe, which 

hybridizes with a fragment, selects fragments from the present bulk (“smear”) 

of fragments. The probe may be radioactively labeled ( 32 Por 35 S) showing 

the hybridized fragment by autoradiography. Biotin, dioxigenin, or fluorescein 

probes visualize the fragment colorimetrically or as chemoluminescence. 

The different fragment pattern (restriction fragment length polymorphisms, 

RFLPs) differentiate species, intersterility groups and isolates, like as it was 

used e.g., for Armillaria spp. (Schulze et al. 1995, 1997). 

The technique is exact, but needs time and is methodically longwinded. 

Methods using the polymerase chain reaction (PCR) 

The procedure of PCR multiplies a part of DNA by a repeated (25–40 times) 

three-stage temperature cycle (amplification): the double strand is split into its 

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36 2 Biology 

single strands at about 94 ◦ C (denaturation), two nucleotide primers (15–30 

bases) attach to the complementary nucleic acid region at 35–60 ◦ C (annealing), 

and a thermostable polymerase synthesizes two new single strands at 

about 72 ◦ C (extension) by starting at the primers and using the four nucleotides 

present in the reaction mixture (Mullis 1990), that is the target DNA 

is doubled with each cycle. 

In real-time PCR techniques, the accumulation of PCR product is detected 

in each amplification cycle either by using a dye or a fluorescently labeled 

probe. Hietala et al. (2003) quantified Heterobasidion annosum colonization in 

different Norway spruce clones using multiplex real-time PCR. Eikenes et al. 

(2005) monitored Trametes versicolor colonization of birch wood samples. The 

technique of PCR-DGGE was used for arbuscular mycorrhizal fungi. A nested 

PCRofvariableregionsofthe18SrDNAwascombinedwithsubsequent 

separation of the amplicons using denaturing gradient gel electrophoresis 

(DGGE), and the method is intended to be used to discriminate closely related 

Glomus species (Vanhoutte et al. 2005). Vainio and Hantula (2000) performed 

DGGE analysis of fungal samples collected from spruce stumps. 

Randomly amplified polymorphic DNA (RAPD)-analysis 

RAPD analysis is based on PCR, but uses only one, short (about ten bases) 

and randomly chosen primer, which anneals as reverted repeats to the complementary 

sites in the genome. The DNA between the two opposite sites with 

the primers as starting and end points is amplified. The PCR products are 

separated on agarose gels, and the banding patterns distinguish organisms 

according to the presence/absence of bands (polymorphism). It is a peculiarity 

of RAPD analyses that they discriminate at different taxonomical level, viz. 

isolates and species, depending on the fungus investigated and the primer 

used (Annamalai et al. 1995). 

RAPD was used for tree parasites, such as Armillaria ostoyae (Schulze et al. 

1997) and H. annosum (Fabritius and Karjalainen 1993; Karjalainen 1996), 

mycorrhizal fungi (Jacobson et al. 1993; Tommerup et al. 1995) and edible 

mushrooms (Lentinula edodes: Sunagawa et al. 1995). Regarding wood decay 

fungi, Theodore et al. (1995) showed for S. lacrymans polymorphism among 

eight isolates. Another RAPD analysis exhibited similarity within S. lacrymans, 

which may be attributed to the low genetic variation of the species, but “normal” 

polymorphism in S. himantioides and Coniophora puteana (Schmidt and 

Moreth 1998). 

The German isolate Eberswalde 15 of C. puteana is obligatory test fungus for 

wood preservatives according to EN 113. The isolate is known for its variable 

behavior in wood decay tests. RAPD analysis was able to show that some alleged 

Ebw. 15 cultures held in different test laboratories are in reality subcultures 

from the British facultative test isolate FPRL 11e (Göller and Rudolph (2003), 

which explains the varying test results. 

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RAPD analysis does not require information of the target DNA and is fast 

when starting from pure cultures. However, at least four primers should be used 

to avoid spurious results, because the short primers imply a great sensitivity to 

contamination. In addition, the technique is unsuitable for the identification 

of unknown samples by comparison, because other not yet investigated fungi 

by chance may share a similar banding pattern. 

Use of ribosomal DNA 

The investigation of the ribosomal DNA (rDNA) has become popular, because 

the rRNA genes and spacers are assumed to evolve cohesively within a single 

species, to exhibit only very little sequence divergence between rDNA copies 

within single individuals, but to show normal levels of sequence divergence between 

species. The repetitive units of the nuclear rDNA of Eukaryotes consists 

of the conserved coding domains 18S and 28S rDNA. The conserved domains 

are interrupted by the non-coding variable internal transcribed spacer ITS I 

(between 18S and 5.8S) and ITS II (between 5.8S and 28S) which are informative 

for differentiation. The intergenic spacer IGS is located between the 28S 

andthe18SofthenextrDNAunit.Inthecaseofapresent5SrRNAgene, 

the IGS consists of two parts, IGS I and IGS II (Fig. 2.20). The conserved 

regions are preferentially used for phylogenetic analyses of genera, families, 

and orders. The rapidly evolving ITS spacers have become a popular choice 

for closely related species and at the subspecies level. After amplification by 

PCR, the amplicons are either restricted by endonucleases providing restriction 

fragments (RFLPs) which are subsequently separated according to size 

using agarose or polyacrylamide gel electrophoresis, or the DNA sequence is 

determined (“sequencing”). 

In addition to the nuclear rDNA, also mitochondrial rDNA was used for 

Basidiomycetes, e.g., by Bao et al. (2005a) in view of phylogenetic relationships 

among closely related Pleurotus species. 

Restriction fragment length polymorphism (RFLP) of rDNA 

RFLP analysis based on the rDNA was also called amplified ribosomal DNA 

restriction analysis (ARDRA). Depending on the intension, the RNA genes or 

the spacers are used. For RFLP analysis of the ITS, the ITS is first amplified, 

often using the “universal primers” ITS 1 and ITS 4 (White et al. 1990), which 

anneal to the evolutionary stable 18S and 28S rRNA genes. This attachment 

Fig.2.20. Schematic diagram of one rDNA unit. The number in the boxes is the size in base 

pairs for Serpula lacrymans (supplemented from Moreth and Schmidt 2005) 

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38 2 Biology 

to conserved rDNA regions allows the ITS amplification from fungi without 

prior knowledge of their ITS sequence. The PCR products are then digested 

either as single or as double digest by restriction endonucleases, of which some 

hundreds of different enzymes are known and each having an own recognition 

site on the DNA. 

Jonsson et al. (1999) identified mycorrhizal fungi in a spruce forest by ITS- 

RFLP comparison to reference material. Similarly, Johannesson and Stenlid 

(1998) identified tree parasites like Armillaria borealis and Heterobasidion annosum 

from bore core samples or mycelia isolated from wood. Regarding wooddecay 

Basidiomycetes, Zaremski et al. (1999) differentiated single isolates of 

C. puteana, Gloeophyllum trabeum and Oligoporus placenta by ITS-RFLPs. 

Various isolates of the closely related S. lacrymans and S. himantioides exhibited 

a distinct fragment profile for both fungi after digestion with HaeIII/TaqI 

(Schmidt and Moreth 1999). Restriction with TaqI differentiated S. lacrymans; 

S. himantioides, D. expansa, C. puteana, A. vaillantii, O. placenta and Gloeophyllum 

sepiarium by specific fragments (Fig. 2.21). 

Obviously, ITS-RFLP analysis is able to separate various wood decay fungi. 

It also detected misnamed isolates assumed to be S. lacrymans (Horisawa et al. 

2004; also Fig. 2.21) and identified unknown samples. The method is currently 

to be intended as a database for the identification of wood decay and associated 

fungi (Zaremski et al. 1999; Adair et al. 2002; Diehl et al. 2004; Råberg et al. 

2004). 

Advantageous is that the technique is fast and inexpensive. Limitations are: 

First, the use of universal primers implies sensitivity to contamination. Second, 

Fig.2.21. Species-specific ITS-RFLP pattern of isolates of Serpula lacrymans (L), S. himantioides 

(H), Coniophora puteana (C), Donkioporia expansa (D), Antrodia vaillantii (A), 

Oligoporus placenta (O), and Gloeophyllum sepiarium (G) generated by TaqI. M marker 

(50–1,000 bp). The culture X had been assumed to be C. puteana and was later identified 

by sequencing as C. olivacea 

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in view of a data collection to be used for identification by fragment length 

comparison, the limited ITS size of only 600–700 bases prevents a separation of 

all relevant fungi in a certain biotope by specific digest pattern. Third, the great 

number of possible fungi in a biotope is much greater than the few fragment 

patterns that would be present in data collections, that is, a not yet analyzed 

species may feign another fungus by exhibiting identical fragments. 

RFLP analysis of a 5.8S rDNA/ITS II/28S rDNA fragment was used to characterize 

five species of the Phellinus igniarius group (Fischer 1995) and 13 species 

of the Phellinus pini group (Fischer 1996). Corresponding DNA digestion of 

52 lignicolous European species with HpaII resulted in 44 distinct phenotypes 

and additional application of Hin6 IandHinf I in 48 species-specific and 

two genera-specific phenotypes (Fischer and Wagner 1999). RFLP patterns 

obtained from seven restriction enzymes assigned 34 Pleurotus strains to 11 

RFLP types, of which ten corresponded to biological species (Bao et al. 2004b). 

The intergenic spacer, either the IGS I alone or both IGS parts, has often 

been used for RFLP studies of Armillaria species (e.g., Harrington and Wingfield 

1995; Frontz et al. 1998; White et al. 1998; Terashima et al. 1998a; Kim 

et al. 2001). IGS I-RFLPs were also used to assign isolates of Heterobasidion 

annosum to intersterility groups (Kasuga and Mitchelson 2000) and to investigate 

the population structure of five Fennoscandian geographic populations 

of Phellinus nigrolimitatus (Kauserud and Schumacher 2002). 

rDNA Sequencing 

PCR-amplification and subsequent sequencing of parts of the ribosomal DNA 

avoid the main limitations of RFLPs because the whole information of hundreds 

of nucleotides of the target DNA is used. rDNA sequences may be used for diagnosis 

and for phylogenetic analyses (dendrograms) on relationships among 

fungi. Sequencing is nowadays the most important tool for molecular systematics 

and led to taxonomic rearrangements and changes in nomenclature. 

TheITSsequencesofagreatnumberofwoodfungiareknown,e.g.,from 

mycorrhizal fungi like Hebeloma velutipes (Aanen et al. 2001), from parasites 

like Armillaria species (Chillali et al. 1998) and Laetiporus sulphureus 

(Rogers et al. 1999), and from the red streaks producing Trichaptum abietinum 

(Kauserud and Schumacher 2003). Regarding wood decay fungi, a data set of 

rDNA-ITS sequences of 18 house-rot fungi is shown in Table 2.8 (Schmidt and 

Moreth 2002/2003) complemented by the 18S and 28S rDNA sequences of some 

important species (Moreth and Schmidt 2005). The ITS of some brown-rot and 

white-rot fungi was sequenced by Jellison et al. (2003). 

It is normal to deposit sequences in the international electronic databases 

for everyone’s use (European Molecular Biology Laboratory EMBL: www.ebi. 

ac.uk/embl; American GenBank: www.ncbi.nlm.nih.gov/genbank; DNA Data 

Bank of Japan DDBJ: www.ddbj.nig.ac.jp). 

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40 2 Biology 

Table 2.8. Sequenced and deposited rDNA regions of indoor wood decay fungi. Grey sequence 

known, 1–28 number of sequenced isolates, six-digit number EMBL database accession 

number (supplemented from Schmidt and Moreth 2002/2003 and Moreth and Schmidt 

2005) 

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Sequences of the ITS region (and the 18S and 28S rDNA) may be used to 

identify unknown fungal samples through sequence comparison by Basic local 

alignment search tool (BLAST) (e.g., www.ncbi.nlm.nih.gov/blast/bast.cgi). 

BLAST revealed ITS-sequence identity of a “wild” S. lacrymans isolate from 

the Himalayas with indoor isolates (White et al. 2001), identified misnamed 

isolates of S. lacrymans (Horisawa et al. 2004), identified Antrodia spp. and 

Serpula spp. isolations from fruit bodies and wood samples (Högberg and 

Land 2004), and confirmed Coniophora puteana isolates (Råberg et al. 2004). 

Kim et al. (2005) used a part of the 28S rDNA for identification of a number 

of basidiomycete fungi from playground wood products by BLAST. Partial 18S 

rDNA sequence of Sirococcus conigenus isolated from Norway spruce cankers 

was used by Lilja et al. (2005) to confirm the identification of the fungus. The 

whole IGS was sequenced to investigate intraspecific variation of mycorrhizal 

fungi like Laccaria bicolor (Martin et al. 1999). IGS I sequence analysis was 

used for Hebeloma cylindrosporum (Guidot et al. 1999) and Xerocomus pruinatus 

(Haese and Rothe 2003). IGS I analysis suggested that three different 

morphotypes/genotypes of an ectomycorrhizal fungus present in Kenya represent 

separate biological species (Martin et al. 1998). The IGS I region grouped 

isolates of Armillaria mellea s.s. in Asian, western North American, eastern 

North American and European populations (Coetzee et al. 2000). 

Sequences are used to construct phylogenetic trees (dendrograms) for phylogenetic 

analyses (molecular systematics). It is not unusual for those intentions 

to complement own data with sequences downloaded from the databases. 

For closely related fungi, like Armillaria species, IGS sequences were used for 

phylogenetic analysis (e.g., Terashima et al. 1998b). Also, ITS sequences may be 

applied to phylogenetic trees. An example of S. lacrymans and S. himantioides 

isshowninFig.2.22.ThetreeshowsthatisolatesofS. lacrymans collected in 

nature in Czechoslovakia, India, Pakistan and Russia group in the branch of 

indoor isolates (“Domesticus group”) but differ from wild Californian isolates 

(“Shastensis group”) (Kauserud et al. 2004b; also White et al. 2001; Palfreyman 

et al. 2003), suggesting a North American link between the anthropogenic 

isolates and the wild relative S. himantioides. Yao et al. (1999) applied ITS 

sequences to a phylogenetic study of Tyromyces s.l. 

For phylogenetic analyses of higher groups, genera, families and orders, 

often the conserved 18S and 28S rDNA are used. Bresinsky et al. (1999) and 

Jarosch and Besl (2001) sequenced 900 bases of the 28S rDNA of S. lacrymans, S. 

himantioides, Meruliporia incrassata and of Coniophora and Leucogyrophana 

species. Although it is not necessary to sequence the whole rRNA genes to construct 

trees, complete 18S and 28S rDNA sequences of a number of important 

wood-decay fungi are already known (Table 2.8). 

Nuclear and mitochondrial genes have different inheritance. Selosse et al. 

(1998) showed intraspecific polymorphism of the large subunit of mitochondrial 

rDNA in Laccaria bicolor. A sequence database of several ectomycorrhizal 

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42 2 Biology 

Fig.2.22. Phylogeny of Serpula 

lacrymans and Serpula himantioides 

ITS-rDNA sequences 

using maximum parsimony and 

Meruliporia incrassata as outgroup. 

Sequences are labeled 

with geographical origin of the 

isolate, followed by the isolate or 

collection name. Bootstrap support 

values (≥50%) are shown 

below the nodes. Stripped lines 

indicate nodes that collapsed in 

the strict consensus tree. Tree 

symbols indicate specimens derivedfromnature(otherwise 

from buildings), and star symbols 

pinpoint sequences derived 

from two newly discovered localities 

in Russia. Black squares 

indicate specimens having double 

character nucleotides in one 

or several positions, reflecting 

a heterozygous state (from 

Kauserud et al. 2004b) 

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basidiomycetes based on a portion of the large subunit of mitochondrial rDNA 

was assembled in view of identification (Bruns et al. 1998). 

rDNA sequencing yields a comprehensive pool of information, but is tedious 

and expensive. Further costs emerge if the PCR products are previously cloned. 

An automatic sequencer is too expensive for small laboratories. However, specialized 

sequencing services are meanwhile inexpensive, providing a sequence 

of about 800 bp length for about e10. 

Species-specific priming PCR (SSPP) 

As an advantage of the sequence divergence among fungi, oligonucleotide sequences 

may be used to design species-specific primers for PCR. At first sight, 

SSPP seems to be a powerful molecular identification tool for fungi. Subsequent 

restriction of the amplicon as well as the use of pure fungal cultures, axenically 

obtained samples, and precautions to exclude DNA from the laboratory or 

from contaminated field material are not required. Jasalavich et al. (1998) used 

primers that detect any basidiomycete fungus present, but not a particular 

species. Specific PCR primers were able to detect the aggressive biotypes 2 and 

4ofTrichoderma harzianum (T. aggressivum f. europaeum and f. aggressivum), 

which are strong parasites in the mushroom production of agarics, Shii-take, 

and Pleurotus species (Albert 2003). With regard to the tree-inhabiting Basidiomycetes, 

special ITS-primers were used for Heterobasidion annosum and 

Armillaria ostoyae (Garbelloto et al. 1996; Schulze and Bahnweg 1998). Specific 

primers distinguished A. mellea from the other four annulate European Armillaria 

species (Potyralska et al. 2002) and detected Phlebia brevispora (Suhara 

et al. 2005). 

To identify indoor wood decay fungi, specific oligonucleotide sequences that 

are located in the ITS II region of seven fungi and were previously tested for 

possible cross-reaction (Moreth and Schmidt 2000; Schmidt 2000) are suitable 

as primers for SSPP (Table 2.9). 

To make subsequent sample recognition easier, different distances of the 

DNA target region to the ITS 1 primer were considered, that is the amplified 

ITS regions exhibit a DNA fragment for each fungus of distinct and predictable 

length on the agarose gel, ranging from about 385 to 625 bp (Fig. 2.23). 

Oh et al. (2003) immobilized specific ITS oligonucleotides of some woodinhabiting 

fungi onto membrane filters for subsequent hybridization of DNA 

from field samples and detected e.g., Chaetomium globosum. 

A specific primer pair targeting the β-tubulin gene was able to distinguish 

between the mutant strain of Ophiostoma piliferum used for biocontrol of 

woodstain and the European and New Zealand wildtype isolates (Schröder 

et al. 2000). 

SSPP is precise and fast. The technique is already used in Germany for 

commercialfungaldiagnosis.However,SSPPdoesnotworkwithallfungi.The 

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44 2 Biology 

Table 2.9. Species-specific ITS-PCR primers (reverse) with target area (bp) in the ITS II if the 

ITS 1 primer of White et al. (1990) is used as forward primer (complemented from Moreth 

and Schmidt 2000) 

Species Specific primer (5 ′ → 3 ′ ) Target 

area (bp) 

Serpula lacrymans ATG TTT CTT GCG ACA ACG AC 567–587 

CAG AGG AGC CGA TGA ACA AG 459–478 

Serpula himantioides TCC CAC AAC CGA AAC AAA TC 410–429 

Coniophora puteana AGT AGC AAG TAA GGC ATA GA 614–633 

Antrodia vaillantii CAC CGA TAA GCC GAC TCA TT 498–517 

ACT GAC TAC AAA ATG GCG CG 445–464 

Oligoporus placenta TTA CAA GCC AGC ATA AAC CT 431–450 

Donkioporia expansa TCG CCA AAA CGC TTC ACG GT 525–544 

Gloeophyllum sepiarium GTT AAT AAA AAC CGG GTG AG 379–398 

Fig.2.23. Electrophoresis gel demonstrating species-specific priming PCR. L–A codes of 

specific primers which detected isolates of Serpula lacrymans (L), S. himantioides (H), 

Coniophora puteana (C), Donkioporia expansa (D), andAntrodia vaillantii (A). M marker 

(200–900 bp) (from Moreth and Schmidt 2000) 

closely related annulate European Armillaria species A. borealis, A. cepistipes, 

A. gallica, and A. ostoyae, exhibited rather similar ITS sequences and also 

intraspecific variation, that is a specific primer was only obtained for A. mellea 

(Potyralska et al. 2002; also Chillali et al. 1998). In addition, intraspecific 

variation may also occur with regard to the geographic origin of isolates 

(Kauserud et al. 2004b). The main limitation is, however, comparable to ITS- 

RFLPs, that the limited ITS size of only 600–700 nucleotides prevents the design 

of a specific primer for all relevant fungi of a certain biotope. In a practical 

view, also the technical effort becomes big on that score that a great number of 

specific primer has to be used for the diagnosis of an unknown sample. 

Microsatellites 

Microsatellites or simple sequence repeats (SSR) are hypervariable genomic 

regions characterized by short tandem repeat sequences of up to seven nu- 

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cleotide units that are distributed throughout the genomes of most Eukaryotes 

(Powell et al. 1996). The variability of the number of repeat units at a particular 

locus and the conservation of the sequences flanking the repeat make 

microsatellites valuable genetic markers. They provide information for identification 

and on genetic diversity and relationships among genotypes. For 

example, DNA fingerprinting with multilocus microsatellite probes suggested 

thatCapeTownisolatesofArmillaria mellea s.s. were introduced from Europe 

more than 300 years ago (Coetzee et al. 2001). 

Amplified fragment length polymorphism (AFLP) 

AFLP is a powerful tool for DNA fingerprinting and is based on (1) total genomic 

restriction, (2) ligation of primer adapters, and (3) unselective followed 

by selective PCR amplification of anonymous DNA fragments from the entire 

genome (Vos et al. 1995). AFLP markers are recognized as more reproducible 

compared to RAPD analyses and inter-simple sequence repeats (ISSRs), and are 

also able to give a higher resolution. AFLP analysis by Kauserud et al. (2004a) 

of European isolates of Serpula lacrymans belonging to five somatic incompatibility 

groups indicated that the species in Europe is genetically extremely 

homogenous by observing that only five out of 308 scored AFLP fragments 

were polymorphic. In contrast, S. himantioides as the closest relative to S. 

lacrymans possessed 31.3% polymorphic fragments. 

2.4.2.3 

Further Molecular Methods 

DNA-Arrays 

DNA-arrays (DNA-chips, microarrays) are tools in medical, pharmaceutical, 

and biological diagnosis of pathogens (genotyping, pathotyping) (Beier et al. 

2002; Wiehlmann et al. 2004). Basis is the increasing availability of sequence 

information of various viruses and bacteria. One chip can carry up to 10,000 

different DNA probes (e.g., oligonucleotides), which are raster-like bound on 

its surface. Nucleic acid molecules of the sample hybridize specifically with the 

corresponding DNA probe, and the hybridized chip areas are detected colorimetrically. 

Compared to PCR techniques, the sensitivity of the chip technology 

is lower than with species-specific PCR, and the chip techniques need experienced 

staff and expensive laboratory equipment. The great miniaturization and 

automation, however, allow the analyses of a great number of samples in a short 

time. Specific oligonucleotides to be used for arrays are already commercially 

available for several pathogenic bacteria and yeasts. A possible future use for 

wood fungi using specific oligonucleotides from rDNA sequences (Table 2.8) 

could be a new technique for fungal diagnosis. 

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46 2 Biology 

Fig.2.24. MALDI-TOF mass spectra of mycelia of each two closely related Serpula and 

Coniophora species (from Schmidt and Kallow 2005) 

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2.5 Classification 47 

Fatty acid profiles 

Microorganisms synthesize over 200 different fatty acids. The presence of 

specific acids and their relative amounts are constant for a particular species. 

Since the 1960s, bacteria and fungi are identified by gas chromatographic 

analysis of fatty acids, which were previously derivatized to methyl esters. The 

technique has also been used to identify wood-decay fungi like Phanerochaete 

chrysosporium, P. sordaria, Trametes versicolor, T. hirsuta,andT. pubescens 

(Diehl et al. 2003). 

MALDI-TOF mass spectrometry 

The technique of matrix-assisted laser desorption/ionization time-of-flight 

mass spectrometry (MALDI-TOF MS) was developed in the 1980s, and was 

used in many fields for peptide, protein, and nucleic acid analyses (Jürgens 

2004; Welker et al. 2004). The method was suitable to differentiate and identify 

viruses, bacteria, and fungi (yeasts and Deuteromycetes) (e.g., Fenselau and 

Demirev 2001). In MALDI-TOF MS, biomolecules and even whole cells are 

embedded in a crystal of matrix molecules, which absorb the energy of a laser. 

The sample is ionized by means of the matrix, and both the matrix and the 

analyte are transferred to the gas phase. The ions are accelerated in an electric 

field, and their time of flight is determined in a detector. After calibration of the 

instrument with molecules of known mass, the flight time of the analyte ions 

is converted to mass-to-charge ratios (m/z). Organism-specific signal patterns 

(“fingerprints”) in the mass range 2,000–20,000 Da were obtained. Figure 2.24 

shows the first MALDI-TOF MS fingerprints of Basidiomycetes, namely the 

closely related sister taxa Serpula lacrymans, S. himantioides and Coniophora 

puteana, C. marmorata (Schmidt and Kallow 2005). The obtained spectra may 

be used for subsequent diagnosis of unknown fungal samples by comparison. 

2.5 

Classification 

Approximately 120,000 fungal species are described. If the numerical ratio 

between vascular plants and fungi of 1:6 in botanically well-examined regions, 

like Great Britain, however, is transferred to a global scale of 270,000 vascular 

plants, 1.6 million fungi might exist. That is, so far only about 10% of the 

actual fungal species are described (Anonymous 1992b). Robson (1999) even 

estimated 3 million fungal species. 

Nomenclature regulates the constitution of names, their validity, legitimacy 

and priority or synonymy, and maintains a single correct name for each taxon 

(International Code of Botanical Nomenclature, St. Louis Code 2000). In view 

of the author names for fungi, these have to be only abbreviated when more 

than two letters are saved. Names are always abbreviated between a conso- 

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48 2 Biology 

nant and a vowel. The abbreviation should not cause confusion with other 

names. Contractions by omission of letters are avoided. Sanctioned names are 

indicated with “Fr.” or “Pers.” after the author of the first valid publication. 

An example might be shown by Trametes versicolor (L.: Fr.) Pilát (Table 2.10) 

(Jahn 1990). “(L.: Fr.) Pilát” means that Linné (L.) described the fungus with 

the name Boletus versicolor in “Species plantarum” in 1753. Fries (Fr.) included 

it as Polyporus versicolor in “Systema mycologicum” in 1821 that is the epithet 

“versicolor” was protected (sanctioned). Pilát placed it in the genus Trametes in 

1939. Particularly French mycologists prefer Coriolus versicolor (L.: Fr.) Quélet, 

because the French author included the fungus in this genus in 1886. In the 

various national colloquial languages and even within a state, different names 

are used. 

For the classification of fungi, there are different attempts of artificial and 

natural systems. The various groups of fungi have little in common, except 

the heterotrophy for carbon, that they are Eukaryotes, possess a slightly differentiated 

tissue, and exhibit in at least one period of life cell walls as well as 

spores as resting and distributing forms. Only for practical reasons they are 

nevertheless united. Multi-kingdom systems (Whittacker 1969) consider the 

polyphyletic origin of the fungi by attaching the slime fungi and “lower fungi” 

totheProtistaandthe“higherfungi”totheFungi,butbreaktherebythetraditional 

biological and ecological term fungus. A generally recognized fungal 

classification system does not exist, and it was ironically argued that there 

might be as many systems as there are systematists. Due to new knowledge, 

and depending on the priority, which is attached to a certain characteristic, 

taxonomic revisions occur in the classification system as well as changes of 

fungal naming (names of wood fungi: e.g., Larsen and Rentmeester 1992; Rune 

and Koch 1992). Current names are shown in Appendix 2. The coarse grouping 

in Table 2.11 is based on Müller and Loeffler (1992). 

About 2,000 described Protista group into six divisions that are independent 

from each other as well as from the “higher fungi”. The “higher fungi” 

Table 2.10. Naming of fungi, illustrated by Trametes versicolor (L.: Fr.) Pilát 

L.: Linné 1753 “Species Plantarum”: Boletus versicolor 

Fr.: Fries 1821 “Systema mycologicum”: Polyporus versicolor: 

→ sanctioning of the epithet “versicolor”, 

Pilát: Pilát 1939: placement in the genus Trametes 

Synonymous especially in France: 

Coriolus versicolor (L.: Fr.) Quélet (1886) 

Vernacular names: 

Germany: Schmetterlingsporling, Bunte Tramete, 

UK: Many-zoned polypore, 

France: Tramète chatoyant 

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with about 120,000 species may be grouped into three divisions and a form 

division: Zygomycota, Ascomycota with the classes Endomycetes (yeasts) and 

Ascomycetes, Basidiomycota including the Basidiomycetes, and Deuteromycota 

(Deuteromycetes). 

The important fungi that inhabit or destroy wood belong to the Ascomycetes, 

Basidiomycetes, or Deuteromycetes. Ascomycetes and Basidiomycetes have in 

common a dikaryotic phase and a haploid phase as mycelium, which does not 

sprout yeast-like. 

About 30,000 Ascomycetes (additionally about 16,000 lichen fungi) are characterized 

by the development of the meiospores in asci, the restriction of the 

dikaryon to the ascogenic hyphae in the fruit body, and the predominant 

gametangiogamy. In the Basidiomycetes (about 30,000 species), the mature 

meiospores are located in the sterigmata, and after somatogamy the dikaryotic 

phaseisextendedtothemycelium. 

As the third artificial group, the Deuteromycetes (30,000 species) are added 

whose vegetative characteristics correspond to the Ascomycetes or Basidiomycetes, 

in which, however, a teleomorph is not yet known or is either 

temporarily or generally not present. 

The term “microfungi” covers the Deuteromycetes and some Ascomycetes 

with microscopic structures. “Macrofungi (macromycetes)” means Basidiomycetes 

and Ascomycetes with large fruit bodies. 

There are different classifications of the Ascomycetes. A traditional way 

considers the appearance of the fruit bodies (ascomata): Hemiascomycetes 

(Protoascomycetes) do not form fruit bodies, Plectomycetes have protothecia 

or cleistothecia, Discomycetes show apothecia, Pyrenomycetes own perithecia, 

and Loculoascomycetes form pseudothecia (Schwantes 1996; Fig. 2.14). 

Another differentiation groups the Ascomycetes according to the time of development 

of the fruit bodies in the two groups, euascohymenial Euascomycetes 

and ascolocular Loculoascomycetes. In the first, the fruit body develops after 

the gametangiogamy, and in the latter, the primordia develop before the 

gametangiogamy. With priorization of the wall structure of the ascus, the 

Lecanoromycetidae show ascohymenial ascomata and a primitive archaeascus, 

the Euascomycetidae comprise ascohymenial fungi with a prototunicate 

Table 2.11. General classification of fungi 

Protista (2,000 species): Six divisions (e.g., slime fungi and “lower fungi”) 

Fungi (“higher fungi”), 120,000, three divisions and one form division: 

1. Zygomycota 

2. Ascomycota: yeasts and Ascomycetes, 46,000 (lichens included) 

3. Basidiomycota: rust fungi, smut fungi and Basidiomycetes, 30,000 

Deuteromycota: Deuteromycetes (imperfect fungi), 30,000 

with relevance to wood: Ascomycetes, Basidiomycetes, Deuteromycetes 

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50 2 Biology 

or unitunicate ascus wall, the Loculoascomycetidae have ascolocular ascomata 

development and mostly a bitunicate ascus wall, and the Laboulbeniomycetidae 

are ascohymenial fungi with prototunicate or unitunicate ascus wall. The 

separate group of the Taphrinomycetidae (Taphrinales) does not form ascomata, 

but the asci develop between the epidermis cells of the host plant. 

Classifications are shown in Kreisel (1969), Breitenbach and Kränzlin (1984), 

Müller and Loeffler (1992), Zabel and Morrell (1992), Schwantes (1996), and 

Hansen et al. (2000). However, a uniform and generally accepted classification 

does not exist. Thus, the Ascomycetes treated in this book are only classified 

by their orders (Tables 8.1–8.3). 

The traditional differentiation of the Basidiomycetes is based on two different 

principles. Practically to apply is the use of the morphology of the 

basidium. Holobasidiomycetes have unicellular basidia, and Phragmobasidiomycetes 

show septate basidia. To consider natural relationships better, a differentiation 

that is based on the kind of spore germination seems favorable 

(Müller and Loeffler 1992). The basidiospores of Homobasidiomycetes germinate 

by germ hyphae. Heterobasidiomycetes show repetitive germination. 

All Homobasidiomycetes possess a holobasidium. The Heterobasidiomycetes 

contain orders with phragmobasidia, but initial more primitive orders have 

holobasidia (Schwantes 1996). Based on the principle type of the fruit body, 

the Homobasidiomycetes may be grouped in Hymenomycetes, which have 

the hymenium exposed on basidiomata surface, and Gasteromycetes with the 

hymenium enclosed within basidiomata. 

Due to overlapping among the groups, lack of clarity, and different opinions 

among systematists, some authors (Müller and Löeffler 1992) abstain from 

uniting the orders into subclasses. 

The former subgrouping of the Homobasidiomycetes into Aphyllophorales, 

Agaricales, and Gasteromycetales did only consider the fruit body type. 

Schwantes (1996) differentiates four order groups: Apphyllophoranae (six orders), 

Agaricanae (three orders), Gasteromycetanae (nine orders), and Phallanae 

(one order). Apphyllophoranae and Agaricanae are almost in accordance 

with the term Hymenomycetes, and Gasteromycetanae and Phallanae with that 

one of Gasteromycetes. Numerous order and family-schemes especially for the 

polypores either use large and comprehensive groups like in Ryvarden and 

Gilbertson (1993, 1994) or numerous and small groups like in Hansen et al. 

(1992, 1997). 

Some common wood-inhabiting Basidiomycetes treated in this book are 

grouped in Table 2.12 according to Breitenbach and Kränzlin (1986, 1991, 

1995), except that the Coniophoraceae were placed in the Boletales. 

A great number of Deuteromycetes occur on wood, like molds (Aspergillus, 

Penicillium and Trichoderma species), blue-stain fungi (e.g., Aureobasidium 

pullulans, Cladosporium species, Discula pinicola), and soft-rot fungi (e.g., 

Paecilomyces variotii). 

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Table 2.12. Classification of some wood-inhabiting basidiomycetous genera 

Class Order Family Genus 

Heterobasidiomycetes Auriculariales Auriculariaceae Auricularia 

Darcymycetales Dacrymycetaceae Dacrymyces 

Homobasidiomycetes Aphyllophorales Sparassidaceae Sparassis 

Corticiaceae s. lato 

Phanerochaetaceae Phanerochaete 

Phlebiopsis 

Phlebiaceae Resinicium 

Stereaceae Amylostereum 

Chondrostereum 

Stereum 

Hymenochaetaceae Asterostroma 

Inonotus 

Phellinus 

Fistulinaceae Fistulina 

Ganodermataceae Ganoderma 

Polyporaceae s. lato 

Polyporaceae s. stricto Lentinus 

Pleurotus 

Polyporus 

Bjerkanderaceae Bjerkandera 

Oligoporus 

Tyromyces 

Coriolaceae Antrodia 

Diplomitoporus 

Donkioporia 

Trametes 

Trichaptum 

Daedaleaceae Daedalea 

Daedaleopsis 

Fomitaceae Fomes 

Fomitopsidaceae Fomitopsis 

Gloeophyllaceae Gloeophyllum 

Grifolaceae Grifola 

Heterobasidiaceae Heterobasidion 

Laetiporaceae Laetiporus 

Meripilaceae Meripilus 

Phaeolaceae Phaeolus 

Piptoporaceae Piptoporus 

Rigidoporaceae Physisporinus 

Schizophyllaceae Schizophyllum 

Agaricales Tricholomataceae Armillaria 

Flammulina 

Laccaria 

Coprinaceae Coprinus 

Strophariaceae Kuehneromyces 

Pholiota 

Boletales Boletaceae Boletus 

Coniophoraceae Coniophora 

Leucogyrophana 

Meruliporia 

Serpula 

Paxillaceae Paxillus 

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52 2 Biology 

The Deuteromycetes are usually divided in Coelomycetes and Hyphomycetes. 

Coelomycetes develop conidiophores within fruit bodies (conidiomata), 

which are either spherical with an apical opening (pycnidium), or flat, cupshaped 

(acervulus). Nearly all Coelomycetes are of ascomycetous affinity. In 

Hyphomycetes (Moniliales), fruit bodies are absent, and conidia develop on 

simple or aggregated hyphae. The “black yeasts” with melanized cell walls and 

nearly always with true mycelium (Chap. 6.2) are anamorphs of Dothideales 

and are therefore also included in the Hyphomycetes. 

The main criterion to classify Deuteromycetes is based on their mode of 

conidium formation. In addition, the conidiogenous cell is used to identify 

and classify Deuteromycetes. The conidiogenous cells can be borne directly in 

or from a vegetative hypha or on differentiated supporting structures. The entire 

system of fertile hyphae is called the conidiophore. Conidia can be formed 

in acropetal chains, or by basipetal succession, viz. the youngest conidium is 

formedatthebase,orbysympodialsuccession,whereeachnewlyformedconidium 

moves into terminal position so that a geniculate, elongate or condensed 

rachis develops. It is differentiated whether conidia result from fragmentation 

and demarcation of already existing hyphae (thalloconidia, arthroconidia) or 

by sprouting (blastoconidia), after the origin of their cell wall from the mother 

cell and whether only one conidium is formed (solitary) or several one behind 

the other in chains (catenulate) or as clusters (botryos). Criteria for the 

recognition of taxa are mostly different from the fundamental characters for 

biological classification. Instead, species are identified with artificial key features. 

Descriptions and classifications are by v. Arx (1981), Barnett and Hunter 

(1987), Wang (1990), Müller and Loeffler (1992), Hoog and Guarro (1995), 

Schwantes (1996), Reiß (1997), Jennings and Lysek (1999), Kiffer and Morelet 

(2000) and Samson et al. (2004). 

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3 Physiology 

The wood-inhabiting fungi as well as their colonization and damaging of 

wood are influenced by various physical/chemical and biological influences 

(Table 3.1). 

Physical/chemical factors comprise nutrients, water, air, temperature, pH 

value, light, and the force of gravity. Biological influences arise because of 

reciprocal effects between different organisms as antagonism, synergism, and 

symbiosis (e.g., Rypáček 1966; Käärik 1975; Rayner and Boddy 1988). When 

investigating the various factors, laboratory methods do not reflect the situation 

under natural conditions. Often it is difficult to vary a parameter without 

affecting the others. The individual factors in nature do not work isolated, but 

strengthen or weaken themselves mutually. 

Table 3.1. Influences on fungal activity 

physical/chemical: 

nutrients, water, air, temperature, pH-value, light, force of gravity 

biological: 

antagonism, synergism, symbiosis 

3.1 

Nutrients 

Fungi consist of about 90% water and 10% dry matter (chemical composition: 

Bötticher 1974). This dry matter has to be synthesized in the course of each 

hyphal division so that nutrients must be assimilated. Regarding the source of 

carbon, wood fungi are heterotrophic by using carbon from organic material, 

which derives from the autotrophic trees. In view of the biochemical way of 

nutrition, wood fungi are chemo-organotrophic. These fungi use organic compounds 

as hydrogen suppliers to produce energy from organic substances. This 

energy production is created by reduction-oxidation reactions (Schlegel 1992). 

Wood fungi are either parasites, which affect living tree tissue, or saprobes, 

which grow on dead wood. Both forms can be obligatory or facultative, as 

a saprobe may become a weakness or wound parasite with weakening or 

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54 3 Physiology 

wounding a tree. A parasite may remain active as a saprobe for some time 

after tree cutting. Schmiedeknecht (1991) differentiated five main groups of 

the heterotrophic way of life: parasites, nekrophytes, which affect living hosts 

either as weakness parasites or kill them by toxic effect, sarkophytes, which 

prepare freshly died tissue for saprobes, saprobes, and symbionts (also Rayner 

and Boddy 1988). 

In view of the use of wood nutrients (Table 3.2), wood-inhabiting microorganisms 

use carbon only from enzymatically easily accessible and digestible 

substrates, like simply constructed sugars, peptides, or fats, or from the storage 

material starch in the parenchyma cells. The wood decay fungi use carbon additionally 

from the complex, main components of the woody cell wall, cellulose, 

hemicelluloses, and lignin. 

The cell wall components can be degraded either directly within the wood 

cell wall or only as a pure component after isolation from the cell wall (Table 

4.3). In the laboratory, sugars such as glucose, maltose (in malt extract), and 

saccharose are suitable C-sources for most wood fungi. The wood-inhabiting 

fungi [yeasts (Chap. 9.5), molds, blue-stain fungi, red-streaking fungi in the 

early stage (Chap. 6)] and the wood-decay fungi during initial decay nourish 

predominantly of sugars and other components in the wood parenchyma cells. 

The quantity of these primary metabolites is usually below 10% related to the 

wood dry weight, and these metabolites occur usually only in living or just died 

sapwood parenchyma cells. For example, soluble nutrients in wood increased 

its susceptibility to soft-rot fungi and bacteria in ground contact (Terziew and 

Nilsson 1999). In Pinus contorta wood samples, triglycerides were consumed 

and mannose was the most depleted sugar by several blue-stain fungi (Fleet 

et al. 2001). The wood-degrading brown, white and soft-rot fungi (Chap. 7) use 

carbon additionally from the macromolecular cell wall components cellulose, 

hemicelluloses and lignin (the latter only with the white-rot fungi) (Chap. 4). 

Wood-inhabiting bacteria (Chap. 5.2) consume sugars and peptides of the 

parenchyma cells and affect non-lignified cell tissue (parenchyma cells, epithelial 

cells of the resin channels, sapwood bordered pits). Under natural 

Table 3.2. Grouping of wood microorganisms according to nourishment and damages 

wood inhabitants: 

bacteria, slime fungi, yeasts, 

staining fungi (molds, blue-stain fungi, red-streaking fungi at an early stage): 

growth on the surface and/or in the outer wood area, 

nutrition from the contents of parenchyma cells and sawwood capillary liquid 

wood decayers: 

brown-rot, white-rot, soft-rot fungi: 

wood rot as a result of nourishment from the polymeric components 

(cellulose, hemicelluloses, lignin) of the lignified cell wall 

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3.1 Nutrients 55 

conditions in the soil, in lakes, and marine environments, mixed bacterial 

populations of the erosion, cavitation and tunneling bacteria can degrade 

wood (Schmidt and Liese 1994; Daniel and Nilsson 1998; Kim and Singh 2000). 

Even a bacterial pure culture attacked woody cell walls (Schmidt et al. 1995) 

(Fig. 5.3c). 

Whereas the fungal cell wall with openings up to 10 nm hardly limits the 

uptake of water and small molecules, the plasma membrane is a selectively 

permeable barrier for the uptake and secretion of solutes. Water, non-polar 

and small uncharged polar molecules, like glycerol and CO2, can diffuse freely. 

Larger polar molecules and ions pass the membrane by means of diffusion 

or active transport (Rayner and Boddy 1988; Jennings and Lysek 1999). The 

uptake occurs mainly at the hyphal tips (Figs. 2.3, 2.4). Three main classes 

of nutrient uptake and transport occur in fungi, facilitated diffusion, active 

transport, and ion channels (Robson 1999). A constitutive low affinity transport 

system of facilitated diffusion allows the energy-independent accumulation of 

solutes like sugars and amino acids when present at a high concentration 

outside of the hypha, but not against a concentration gradient. When the 

solute concentration is low, carrier proteins are induced that have a higher 

affinity for the solute and mediate the energy-dependent uptake of solutes 

against a concentration gradient at the expense of ATP. During this process, 

fungi create an electrochemical proton gradient by pumping out hydrogen ions 

from the hyphae at the expense of ATP via proton pumping ATPases in the 

plasma membrane. The proton gradient provides the electrochemical gradient 

that drives nutrient uptake as hydrogen ions flow back down the gradient. 

A number of ion channels that are highly regulated pores in the membrane 

and allow influx of specific ions into the cell when open have been found 

in fungi. Ca 2+ stimulated K + channels carry an inward flux of K + ions and 

are thought to be involved in regulating the turgor pressure of the hypha. 

A mechanosensitive or stretch-activated Ca 2+ channel is opened when the 

membrane is under mechanical stress like during the generation of the high 

calcium gradient at the hyphal tip. 

During early growth, nutrients surrounding the young mycelium are in excess. 

As the mycelium develops further, nutrients in the center are increasingly 

utilized, nutrient depletion and accumulation of metabolic products occur 

beneath the colony center. Therefore, growth becomes restricted to the periphery. 

Different parts of the colony are at different physiological ages, with 

the youngest hyphae at the edge of the colony and the oldest, non-growing 

mycelium at the center (Robson 1999). 

The movement of the nutrient over short distances from a food source on 

to the regions devoid of the nutrient or nutrients required for growth can 

occur by diffusion within the aqueous phase of the cytoplasm (Jennings and 

Lysek 1999). As mycelial extension proceeds, nutrients are shifted from the 

site of absorption to another part of the mycelium by translocation (Jennings 

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1987, 1991). Translocation of nutrients is predominantly by water flow. Water 

flow is generated by the uptake of nutrients, particularly carbohydrates, by 

the mycelium such that the hyphae have a lower water potential than the 

substrate. In consequence, water flows into the hyphae and the hydrostatic 

pressure so generated drives a flow of solution towards the mycelial growth 

front.Thevolumeflowisdissipatedatthegrowthfrontbytheincreasein 

volume of the hyphae and the production of droplets at the hyphal apices. 

The droplets have a lower osmotic potential than the hyphae or that of the 

substrate from which the mycelium grows. This means that the water leaves 

the cytoplasm ultrafiltered by the plasmalemma of many of the nutrients 

in the translocation stream. Pressure-driven flow of solution has been studied 

particularly in Serpula lacrymans(Jennings 1991). It must occur in a wide range 

of fungi because droplets (guttation) are common among fungi. Guttation 

often occurs in white-rot fungi, like during growth of Donkioporia expansa 

in buildings and in the edible mushrooms Lentinula edodes and Pleurotus 

ostreatus when the colonization phase of the substrate is completed and the 

fungi start fruiting. In S. lacrymans, the droplets at the hyphal tips are slightly 

acidic (pH 3–4), which was related to the ability of the fungus to colonize 

alkaline substrates (Bech-Andersen 1987a). 

The dry weight of fungal mycelium consists of about 5% of nitrogen (% N 

of the Kjeldahl method ×4.4 corresponds to the protein content of fungi. Additional 

nitrogen is included, e.g., in the chitin). Wood typically has a very low 

nitrogen content. The average nitrogen for healthy hardwoods and softwoods 

was 0.09% of the dry weight of wood and reached to about 0.2% N (Rayner and 

Boddy 1988; Fengel and Wegener 1989; Reading et al. 2003) with an average 

carbon to nitrogen ration of 500 to 600:1. Nitrogen content changes over the 

wood cross section and is lower in wounded or decayed tissue. With regard to 

lignocelluloses, it has to be considered, however, that the majority of carbon 

is present as a cell wall component and thus enzymatically difficulty accessible, 

while the nitrogen compounds are more easily degradable. Altogether 

nitrogen, however, is a limiting factor. Fungi do not fix atmospheric nitrogen, 

how this some bacteria are able to do. Instead, fungi use nitrogen rationally, 

as nitrogen compounds are translocated to the growth front at the hyphal tips 

due to different turgor pressure in the mycelium (Watkinson et al. 1981; Jennings 

1987). Protein-rich woods, e.g., Pycnanthus angolensis, are colonized by 

bacteria after felling and during the drying process, which leads to undesirable 

discolorations (Chap. 5.2) (Bauch et al. 1985). For wood fungi, ammonium is 

a suitable inorganic source of nitrogen in vitro, while nitrate is usually not 

used. Organic nitrogen from amino acid mixtures in pepton or malt extract 

results in good growth on agar. 

There are several minerals in wood. The main inorganic components found 

in wood ash are K, Ca, Mg, Na, Fe, silica, phosphate, chloride, and carbonate 

(e.g., Fengel and Wegener 1989; also Wa˙zny and Wa˙zny 1964). By SEM-EDXA, 

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3.1 Nutrients 57 

Al, S, and Zn were detected (Rodriguez et al. 2003). Particle induced X-ray 

emission(PIXE)quantifiedP,S,K,Ca,Ti,Mn,Fe,Ni,Cu,Zn,Pb,Sr,Rb,Ba 

and F (Saarela et al. 2002). Inductively coupled plasma emission (ICP) showed 

a content of 50–100 ppm of manganese in Scots pine sapwood (Schmidt et al. 

1997a). Inorganic compounds comprise 0.1–0.5%, oxide basis, of total wood 

componentsintemperatezonesandupto4%intropicalwoods.Mineralelementsenterthelivingtreepredominantlythroughtheroot,whichisfrequently 

helped by mycorrhizae fungi. The wood-inhabiting fungi use metals present 

in wood for their growth and to degrade it (Chap. 4). Several metals are necessary 

to fungi, e.g., for wood degradation. Enzymes that participate in lignin 

degradation contain iron (lignin- and manganese peroxidases, cellobiose dehydrogenase) 

or copper (laccases) (Rodriguez et al. 2003). Iron, manganese, and 

copper are involved in the generation of hydroxy radicals or other oxidizing 

agents, which, in turn, attack wood (Henry 2003). 

Elements present in forest and other soils can also be a nutrient source for 

fungi, enhancing fungal capacity to degrade wood. The wood nitrogen content 

can be increased by ground contact or by means of translocation through the 

mycelium. Nitrogen can be taken up, e.g., by Serpula lacrymans mycelium 

from the soil under houses and transported in the strands to the place of wood 

degradation within buildings (Doi and Togashi 1989). 

Some wood-degrading Basidiomycetes are heterotrophic for vitamin B1 (thiamine). 

Heterobasidion annosum is auxoheterotrophic regarding the pyrimidine 

half of thiamine, can however synthesize the thiazole part of the vitamin 

(Schwantes et al. 1976). Some wood-decay fungi additionally need vitamin H 

(biotin). Suitable vitamin sources in vitro are yeast and malt extract. 

Thiamine is decomposed in hot alkaline medium. Therefore in the USA, 

poles had been treated with ammonium gas under high temperature (“dethiaminization”) 

to destroy the vitamin and, thus, to protect the wood against 

decay fungi. The poles, however, were for all that attacked by fungi, as thiamine 

from soil bacteria (Henningsson 1967) diffused into the poles during service 

(treatment of cut timber: Narayanamurti and Ananthanarayanan 1969). 

In addition to cell wall components, primary metabolites and storage material, 

wood contains a broad spectrum of extractable substances (extractives, 

accessory compounds, secondary metabolites) like waxes, fats, fatty acids and 

alcohols, steroids and resins (Fengel and Wegener 1989; Obst 1998). More than 

10,000 compounds were reported to occur in plants (Duchesne et al. 1992). 

Depending on the wood species, the type, quantity, and distribution of the extractives 

can vary considerably. They are particularly located in the heartwood, 

and after wounding and microbial infection also in the sapwood as wound reaction 

(Chap. 8.2.1). Heartwood is a dark-colored zone in the central part of 

the stems of most tree species and is physiologically formed from sapwood, 

followed by decreased moisture content, the death of parenchyma cells, and 

increased extractive content. Inhibiting extractives, which cause the natural 

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durability of many heartwood species develop during heartwood formation 

from starch and soluble carbohydrates (Magel 2000) and are mainly phenols, 

like terpenoids, flavonoids, stilbenes, and tannins (Fengel and Wegener 1989; 

Obst 1998; Roffael and Schäfer 1998; Imai et al. 2005). For example, pinosylvins 

inhibited brown-rot fungi (Celimene et al. 1999), flavonoids inhibited Gloeophyllum 

trabeum and Trametes versicolor (Reyes-Chilpa et al. 1998). While the 

extractives during the obligatory formation of a colored heartwood penetrate 

in the cell walls, those that develop by exogenous influences (facultatively colored 

heartwood), like wound reactions, do not impregnate cell walls (Koch 

2004). 

Omnivors are the only less specialized molds (Chap. 6.1), which can grow on 

wood, paper, wallpaper, books and leather, and dissolve even minerals from 

glass by acid production (Kerner-Gang and Schneider 1969). The “polyphage” 

H. annosum has a broad host spectrum of over 200 wood species (Heydeck 

2000). As a specialized parasite, Piptoporus betulinus attacks only birch trees 

(host spectrum: Jahn 1990; Ryvarden and Gilbertson 1993). 

Nutrient media to isolate, enrich, purify, and cultivate wood-inhabiting 

fungi are malt extract agar and potato dextrose agar of about pH 5.5. Bacterial 

isolates from wood grow on nutrient media like peptone/meat extract/yeast 

extract of about pH 7 (Schmidt and Liese 1994). For special microorganisms, 

selective media are commercially available, or standard agar is enriched with 

selecting compounds. If bacteria have to be eliminated during fungal isolations, 

the substrate can be acidified by lactic or malic acid or an antibiotic is 

added. Orthophenylphenol selects on white-rot fungi. Benomyl inhibits molds 

like Penicillium and Trichoderma species. 

3.2 

Air 

As aerobic organisms, wood fungi produce CO2, water, and energy by respiration 

and need therefore air oxygen (Table 3.3). 

The energy production from wood, if only cellulose is consumed, is shown 

in Table 3.4. Aerobes, however, do not respire carbohydrates totally, but use 

intermediates for their metabolism. 

Fungal activity is affected by the composition of the gaseous phase. Usually 

wood decay decreases at low O2 and high CO2 content, respectively. The O2 

Table 3.3. Aerobic degradation of wood to CO2, water and energy 

cellulose, hemicellulose, lignin from wood – (ectoenzymes) → 

sugars, lignin derivates – (uptake, intracellular enzymes) → CO2 +2(H) 

2(H) + 1/2O2 – (respiratory chain) → H2O + energy (ATP) 

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3.2 Air 59 

Table 3.4. Energy production from wood cellulose 

Assuming that 1 kg dry wood contains 48.6% cellulose: 

1 mol glucose (180 g) yields 2,835 kJ, 

180 g glucose correspond to 162 g cellulose 

[162 + 18; (1 mol H2O used for hydrolysis)], 

3 × 162 = 486, 

486 g cellulose yield 8,505 kJ (2,025 kcal) 

content in the wood of living oak trees varied season-dependently from 1–4% 

and the CO2-content from 15–20% (Jensen 1969). 

There are various reactions occurring in wood fungi that require oxygen, 

such as degradation of lignin, oxidative polymerization of phenols, and 

melanin synthesis in blue-stain fungi and other fungi. With the onset of differentiation, 

there is also an increased oxygen demand. When the reproduction 

is initiated, there is a high requirement for protein and nucleic acid synthesis, 

which energetically involves a higher demand on the fungal metabolism and, 

thus, increased oxygen utilization (Jennings and Lysek 1999). This reason as 

well as access to air currents for spore dispersal explain why most fungi form 

their fruit bodies at or near the substrate surface. 

A lack of oxygen can limit wood decay. Saprobes usually react more sensitively 

to O2 lack than parasites living within the heartwood: The saprobes 

Serpula lacrymans and Coniophora puteana survived without oxygen 2 and 

7 days, respectively (Bavendamm 1936), the parasitic heartwood destroyer 

Laetiporus sulphureus more than 2 years (Scheffer 1986). In Heterobasidion 

annosum, mycelial growth hardly decreased at 0.1% O2 content compared to 

20% (Lindberg 1992). The conidia of some blue-stain fungi still germinated 

at 0.25% O2 content, some Mucoraceae (molds) even in a pure N-atmosphere 

(Reiß 1997). 

The yeasts, which are able to get energy also facultatively anaerobically 

by fermentation, form an exception of the aerobic way of life among the 

fungi. During the alcoholic fermentation of the hexose sugars (Saddler and 

Gregg 1998) in coniferous wood sulphite spent liquors which was performed 

in former times e.g., in Switzerland, the produced hydrogen is not transferred 

to atmospheric oxygen, but to the organic H-acceptor acetaldehyde: 

2(H) + CH3CHO → CH3CH2OH (ethanol). At low oxygen content, anaerobic 

metabolites like ethanol, methanol, acetic acid, lactic acid, and propionic acid 

have been found also in Basidiomycetes (Hintikka 1982). 

Inthecourseofwooddegradation,theCO2 concentration may increase. 

Some wood-degrading Basidiomycetes, particularly heartwood destroyer, are 

tolerant of a high CO2 content, since they grew well at 70% CO2 and even at 

100% (Hintikka 1982), while forest-litter decomposing fungi were inhibited 

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at more than 20% CO2. Chaetomium globosum and Schizophyllum commune 

can fix CO2 into organic acids of the citric acid cycle (Müller and Loeffler 

1992). An increasing CO2 content inhibits the growth of many Deuteromycetes, 

which then partly change the metabolism to fermentation and also alter their 

filamentous growth manner to a yeast-like appearance (Reiß 1997; Jennings 

and Lysek 1999). 

The minimum air volume in wood for degradation by fungi is between 10 

and 20%: 10% in H. annosum, 20% in S. commune (Rypáček 1966). 

Reduction of the O2 content in wood effects a protection against fungal (and 

insect) decay. Such protection is performed by wet storage of wind-thrown 

wood by dipping and floating in water or sprinkling of piled wood. From 17.6 

million m 3 of windfalls after the storm in north Germany in 1972, 1.4 million 

were protected by watering and were sold until 1976 nearly without any quality 

loss (Liese and Peek 1987; Groß et al. 1991; Bues 1993). In 1990, 15 million m 3 

of round timber were stored by sprinkling in Germany. At the density of about 

0.5 g/cm 3 of spruce and pine wood, the 20% critical air volume is obtained 

through a wood moisture content of 120% u, so that alternating sprinkling is 

sufficient. With new methods, logs are wrapped by plastic foil and stored in an 

atmosphere of CO2 and/or N2 (Mahler 1992). 

The soft-rot fungi are an exception among the wood decay fungi. They exhibit 

lower a requirement for oxygen and can also live in water-filled wood 

tissue like in sprinkled cooling-tower wood with about 200% u moisture content, 

because the cooling-tower water is enriched with the necessary O2 by the 

spraying effect of the dripping water (Chap. 7.3). Among the Basidiomycetes, 

Armillaria mellea s.l. showed a strange behavior, as it caused in sprinkled 

Norway spruce logs tubes in the water-saturated sapwood, through which 

necessary oxygen for wood decay invaded the wood (Metzler 1994). 

In addition, (facultatively) anaerobic bacteria degrade the non-lignified sapwood 

bordered pits in sprinkled and ponded wood, so that wood permeability 

increases and the wood shows later the unwanted, because uneven, excessive 

uptake of wood preservatives or pigments (Willeitner 1971). 

3.3 

Wood Moisture Content 

As wood degradation by fungi involves enzymes, which are active in aqueous 

environment, and because hyphae consist of up to 90% of water, wood fungi 

need water. Water is also used for the uptake of nutrients, the transport within 

the mycelium and as solvent for metabolism. Without water, the metabolism 

rests. The resting phase occurs by means of spores, in wood fungi particularly 

by chlamydospores. Regarding the so-called dryness resistance of wood decay 

fungi (Theden 1972) it was however not proven if vegetative hyphae or spores 

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3.3 Wood Moisture Content 61 

survived. Water is taken up from the substrate wood, the soil, and from masonry 

etc. Altogether the moisture content of wood is the most important factor 

for wood degradation by fungi and thus also for wood protection. Moisture 

in wood exists in two different forms: Bound or hygroscopic water occurs 

within the cell wall by means of hydrogen bounds at the hydroxyl groups 

mainly in the cellulose and hemicelluloses and to smaller extent in the lignin. 

Freeorcapillarywaterinliquidformislocatedinthecelllumenaswell 

as in other holes and cavities of the wood tissue (e.g., Siau 1984; Smith and 

Shortle 1991). 

There are several methods of measuring wood moisture content (Vermaas 

1996): oven-drying method, microwave drying Danko (1994), distillation, Karl 

Fischer-titration, moisture meters based on electrical and dielectrical properties, 

continuous moisture meters, capacity admittance moisture meters, and 

hygrometric methods. Determination of the moisture content without destruction 

is done electrically by means of resistance measurement (Skaar 1988; Du 

et al. 1991a, 1991b; Böhner et al. 1993; Chap. 8.2.4). With increasing moisture 

content of wood from the oven-dry phase to the fiber saturation range (about 

30% u) the electrical resistance decreases approximately by the factor 1:10 6 . 

Moisture can be rapidly determined in practice using an indelible pencil that 

is the pencil line runs if the fiber saturation point is exceeded. 

The proportional wood moisture (% u) is determined gravimetrically by 

the wood mass before and after drying a wood sample at 103 ± 2 ◦ C: u (%) = 

[(MW − MD) : MD] × 100 (MW = mass of wet wood, MD = mass of dry wood). 

If heat-implied changes in the wood samples shall be excluded to take 

care of wood extractives and cell wall components for subsequent microbial/enzymatic 

degradation experiments or chemical analyses, drying of the 

wood specimens can be performed in an evacuated desiccator over silicagel 

or P2O5. Wood samples may be also conditioned to specific relative humidity 

conditions prior to and after decay, e.g., at 20 ± 2 ◦ Cand65±5%relativeair 

humidity. With the latter method, the theoretical dry weight (MDt) of a sample 

results from: MDt = (100 × MC): (100 + u) (MC = mass after conditioning, 

u = % wood moisture after air conditioning). However, weight loss methods 

using moisture-conditioned wood samples instead of oven-dry blocks are 

influenced by changes in hygroscopicity: For brown-rot, mass loss is slightly 

overestimated, for white rot, no difference occurs, while for soft rot, mass loss 

is slightly underestimated using the moisture-condition method (Anagnost 

and Smith 1997). 

To quantify the moisture content of fungal nutrient substrates, including 

wood, only the proportional water content of the substrate was considered in 

previous investigations. At the disposal to microorganisms, however, not the 

whole water content of the substrate is available, but only that part of the total 

water, which is not bound by solved substances (salts, sugars, etc.). The relative 

vapor pressure of a substrate (water activity aw, 0–1) results from the quotient 

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of the water vapor pressure in the substrate (p) and the pressure of pure water 

(p0)(aw=p/p 0 ) (Siau 1984; Rayner and Boddy 1988; Reiß 1997; Table 3.5). 

The minimum water activity (Table 3.5) is for most bacteria with 0.98 aw 

higher than for many molds, which grow still at 0.80 aw. The minimum for 

growth of wood-decay Basidiomycetes on agar is 0.97 aw. Xerotolerant and 

xerophilic molds like some Aspergillus species still grow at 0.62 aw. Those 

fungi grow in solutions of sodium chloride around 5–6 M (Jennings and Lysek 

1999) and tolerate an 80% saccharose solution (Schlegel 1992; Reiß 1997), 

generating the appropriate osmotic pressure within their protoplasm e.g., by 

the synthesis of glycerol. Below 0.6 aw usually no microbial growth occurs. 

The situation of high salt concentrations (sodium chloride) applies also 

to marine fungi. Various “lower fungi”, Deuteromycetes, Ascomycetes, and 

a few Basidiomycetes colonize wood in the sea (Kohlmeyer 1959; Volkmann- 

Kohlmeyer and Kohlmeyer 1993). As in marine fungi vacuoles constitute no 

more than about 20% of the volume of the protoplasm, there is no preferential 

accumulation of sodium chloride in the vacuoles. Marine fungi synthesize glycerol 

and other polyols (mannitol, arabitol) which contribute to their osmotic 

potential (Jennings and Lysek 1999). 

For growth and wood degradation by fungi, particularly at low water contents, 

the water potential (MPa) is the most important factor for water availability. 

It is defined as free energy of water in a system relative to pure water, 

and because in the relevant range all values are negative, it can be defined 

as that negative pressure (“subpressure”), which is necessary to extract water 

from the substrate (Griffin 1977). The water potential is affected by different 

factors (Siau 1984; Jennings 1991). These are particularly the size and form of 

the boundary surfaces both between water and firm matrix and between water 

and air (matrix potential), and the osmotic potential due to the occurrence 

of solved substances. The influence of the water potential on growth of wood 

fungi was first examined with simple substrates, like agar plates in Petri dishes, 

in controlled air humidity (Bavendamm and Reichelt 1938). The observed values 

of mycelial growth still at −14.5 MPa (aw 0.9), however, were later classified 

as too low. Instead, as lower limit about −4 MPa were determined (Griffin 1977; 

Griffith and Boddy 1991; Table 3.5). Serpula lacrymans did not grow on agar 

below −0.6 MPa (Clarke et al. 1980). 

Due to the occurrence of pores of different size (porosity of wood: Kollmann 

1987), the special significance of the matrix potential becomes obvious with 

increasing drying of wood tissue. In water-saturated wood, all cavities are 

filled, and a neglectably small pressure difference is sufficient for dehydration. 

With progressive drying, increasingly smaller openings become free from 

water (Table 3.5). Large openings in wood tissue with radii over 5µm like 

all cell lumens are free from water, if the matrix potential amounts to less 

than about −0.03 MPa. Between −0.03 and −14.5 MPa, pores from 5–0.01µm 

radius become empty (pits, boreholes by microhyphae). Below about −14 MPa, 

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Table 3.5. Correlations between water activity (aw, relative vapor pressure p/p0), water potential (MPa), maximum water-retaining pore radius 

(µm) within wood at 25 ◦C, wood emptiness class and microbial activity (compiled from Griffin 1977; Clarke et al. 1980; Siau 1984; Rayner and 

Boddy 1988; Viitanen and Ritschkoff 1991; Schlegel 1992; Reiß 1997) 

Water activity Water potential Pore radius Wood emptiness class Microbial activity 

(aw, p/p0) (MPa) (µm) 

1.0000 0 (free water) cell lumens, wood degradation and staining 

large openings after decay 

0.9999 −0.014 10.5 

0.9998 −0.028 5.2 

0.9993 −0.10 1.5 fiber saturation area, minimum for most wood fungi 

0.9990 −0.14 1.1 pits and small openings 

0.9975 −0.35 0.4 

0.9950 −0.69 0.2 

0.990 −1.4 0.1 half-maximum growth rate of 

wood-decay Basidiomycetes on agar 

0.980 −2.8 0.05 minimum for most bacteria 

0.970 −4.2 0.035 minimum for mycelial growth and wood decay of 

Serpula lacrymans 

0.960 −5.6 0.026 optimum for growth and sporulation of Aspergillus niger 

−6.0 no growth of S. lacrymans on agar 

0.920 −11.3 0.013 minimum for sporulation of A. niger 

0.900 −14.5 0.01 

0.880 < 0.01 temporary or intermolecular minimum for growth of A. niger 

0.840 openings in the cell wall minimum for germination of A. niger 

and growth of Paecilomyces variotii 

0.800 minimum for most molds 

0.750 minimum for halophilic bacteria 

0.650 minimum for A. repens 

0.600 lower limit for microbial growth 

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intermolecular cavities in the cell wall dry (liquid movement in wood: Siau 

1984; Skaar 1988). 

From the view of a hypha, a low water availability begins to become critical, 

if free water is no more located in the cell lumen void space, but liquid water 

exclusively within the cell wall and only water vapor in the lumen, or in other 

words, if the cell walls are fully hydrated yet with no water contained in the 

cellular spaces. This condition is defined as fiber saturation point or range 

(Babiak and Kúdela 1995) and lies at about −0.1 MPa (0.9993 aw), according to 

1.5µm pore radius (Table 3.5) and about 30% u wood moisture for woods of 

the temperate zones. The lower limit for wood degradation by Basidiomycetes 

is about −4 MPa (0.97 aw). 

Below fiber saturation, not only fungi are influenced by the moisture content, 

but also all technological properties of wood. With increasing moisture, e.g., 

elastic, strength, and insulation properties decrease. 

Relative air humidity (RH), which is in equilibrium with a substrate, and 

water activity of a substrate stand in the relationship: RH(%) = aw × 100. For 

example, 99.93% RH correspond to 0.9993 aw and thus to fiber saturation, so 

that the critical range for Basidiomycetes of 0.97 aw (Table 3.5) is exceeded by 

condensation in buildings. The S-shaped sorption isotherms, which indicate 

the dependence of the wood moisture on the relative air humidity of the environment, 

are shown by Siau (1984) and Kollmann (1987). Wood is dry at the 

relative vapor pressure of 0, and fiber saturation is reached at 1 (100% RH). 

Spruce sapwood samples placed over a saturated solution of K2SO4, which 

results in 97% RH and 26.5% u, showed 4.5% mass loss after 3 months of 

incubation with S. lacrymans (Viitanen and Ritschkoff 1991a). Wood samples 

in 93% RH according to 23–24% wood moisture content were overgrown by 

S. lacrymans and Coniophora puteana (Savory 1964). For the initial colonization, 

21% u was necessary (Huckfeldt et al. 2005; cf. Table 8.7). Coniophora 

puteana colonized wood samples of 18% moisture content when a moisture 

source was 20–30 cm away from the wood. Timber in buildings reached however 

till 45% humidity in the winter during night by condensation (Dirol and 

Vergnaud 1992). 

According to Skaar (1988), the wood moisture content of living trees 

amounted to 83% u in hardwoods in the sapwood and to 81% in the heartwood 

(average of 34 species) and in conifers to 149% in the sapwood and to 55% in 

the heartwood (average of 27 species). 

The moisture content in dead wood is determined by several factors: 

– fungal decay: For example, the wood moistures of dry heartwood samples 

of different wood species increased during decay by Trametes versicolor in 

84 days to 78–236% and by Oligoporus placenta to 108–286% (Smith and 

Shortle 1991). Regarding the sorptive capacity of wood (Cowling 1961; Anagnost 

and Smith 1997), Rawat et al. (1998) showed that the moisture content 

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of brown-rot decayed wood was more than that of undecayed samples at low 

relative humidities, but at higher humidities (about 37%) the situation was 

reversed. Absorptiveness also increased after pretreatment with blue-stain 

fungi (Fjutowski 2005). 

– moisture uptake, which can occur by five ways: rainfall, absorption from air, 

capillary penetration of water into wood in ground contact or in buildings 

by condensation on wood surfaces, water transport by the mycelium, and 

water formation by fungal metabolism, 

– loss of water: in wood with large pores by the force of gravity, furthermore 

by evaporation as a function of temperature, humidity, and matrix potential 

as well as by water transport via mycelium. 

The cardinal points of the wood moisture content for some decay fungi are 

shown in Table 3.6, whereby the data vary, however, depending on the fungal 

isolate, the wood species, and the testing method. Laboratory findings and 

practice observations may also yield different results (Ammer 1964; Savory 

1964; Cockcroft 1981; Thörnqvist et al. 1987; Viitanen and Ritschkoff 1991a; 

Huckfeldt et al. 2005). 

Generally, it applies to wood fungi: The minimum for wood decay is near 

the fiber saturation point of about 30% u, however, commonly slightly above 

this range because only then there is free water in the lumen void space. Some 

house-rot fungi, however, could colonize wood in laboratory culture, whose 

moisture was significantly below fiber saturation (minimum 18% u) before 

the mycelium contacts the woody substrate, because these fungi transported 

water from a moisture source by means of mycelium. The minimum for decay 

of pine wood samples by these house-rot fungi was between 22 and 37% u 

(Huckfeldt and Schmidt 2005; cf. Table 8.7). The optimum differs depending 

on the fungal species and affects the occurrence of different fungi in differently 

moist biotopes: For example, the optimum is at 50–100% for tree-inhabiting 

blue-stain fungi and below 50% for lumber blue-stain fungi (Bavendamm 

Table 3.6. Cardinal points of wood moisture content (% u) for some wood-decay fungi 

(literature data) 

Minimum Optimum Maximum 

Antrodia spp. 30 35–55 60–90 

Coniophora puteana 26–30 30–70 60–80 

Daedalea quercina 40 

Gloeophyllum spp. 30 40–60 80–210 

Heterobasidion annosum 45 

Lentinus lepideus 35–60 

Phlebiopsis gigantea 100–130 

Serpula lacrymans 26 30–60 55–225 

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1974). Wood fungi are inhibited as the cellular spaces of wood become fully 

saturated with water. The maximum wood moisture content allowing fungal 

growth is determined by the minimum air content within the wood cell. 

A certain water amount originates from fungal metabolism (Ammer 1964; 

Savory 1964). The assertion that S. lacrymans gets the total water, which is necessary 

to moisten dry wood, from the respiration of wood cellulose (Table 3.3), 

however, is wrong: It has to be considered that cellulose is not completely 

degraded to CO2 and 56% water. Intermediate metabolites for the synthesis 

of fungal biomass are necessary. According to Weigl and Ziegler (1960) about 

40% of the consumed cellulose is used for those metabolites. Furthermore, 

water production from carbohydrates is the rule for all breathing organisms. 

Nevertheless, some fungi, particularly S. lacrymans, show intensive guttation, 

that is excretion of water in drop form. 

In view of dry wood, in addition to spores also the mycelium of some fungi 

was said to be resistant to dryness (Table 3.7). 

The duration of this so-called dryness resistance depended on air humidity 

and temperature. Resistance lasted e.g., longer at 60% RH and low temperature 

than at 90% RH and high temperature. For S. lacrymans, the duration was 

8yearsat7.5 ◦ Cand1yearat20 ◦ C (Theden 1972). Dryness-resistant are also 

Coniophora species, indoor polypores, Gloeophyllum abietinum (on window 

timber), Lentinus lepideus (on sleepers), Paxillus panuoides, Schizophyllum 

commune, Stereum sanguinolentum, the soft-rot fungi and to smaller extent 

Heterobasidion annosum and Trichaptum abietinum. Towhatextentfungi, 

however, are qualified for dryness resistance, exclusively in the form of hyphae 

or as resistant spores was not examined in detail. 

Serpula lacrymans survived only in slowly drying wood samples. Own laboratory 

observations revealed that its dikaryons formed arthrospores in old, 

dry agar cultures, which points to monokaryotization. That is, the hyphae may 

have developed dryness-resistant resting stages if the substrate takes a long 

time to dry down, and the spores germinate again under sufficient moisture 

conditions. Thus, studies with wood samples that have been colonized 

by mycelium and subsequently slowly dried indicated that S. lacrymans, C. 

Table 3.7. Resistance to dryness (after Theden 1972) 

Years withstanding at ◦C 27 20 7.5 

Antrodia vaillantii ≥ 7 9 ≥ 6 

Coniophora puteana 0 2 4 

Coniophora marmorata 0 3 7 

Gloeophyllum abietinum 5 7 ≥ 7 

Gloeophyllum trabeum 11 ≥ 10 ≥ 8 

Lentinus lepideus 7 ≥ 9 ≥ 8 

Oligoporus placenta 9 ≥ 11 ≥ 5 

Serpula lacrymans 0.5 1 8 

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3.4 Temperature 67 

puteana, Gloeophyllum trabeum, and Donkioporia expansa may survive as 

arthrospores (Huckfeldt et al. 2005). 

3.4 

Temperature 

With respect to the temperature, Table 3.8 shows the cardinal points for some 

wood fungi. A comprehensive investigation was completed in 1933 grouping 

the species into low-temperature (optimum 24 ◦ Candbelow),intermediatetemperature 

(optimum between 24 and 32 ◦ C), and high-temperature group 

(optimum above 32 ◦ C) (Humphrey and Siggers 1933). For three species, e.g., 

Gloeophyllum sepiarium, minimum, and maximum temperatures were already 

determined (Lindgren 1933). It has to be considered, however, that considerable 

differences can exist between isolates of a species (Table 3.11). 

Generally, it applies to wood fungi: The minimum is usually at 0 ◦ C, because 

below the freezing point there is no liquid water available necessary for 

metabolism. Exceptions of growth below 0 ◦ C are possible, if the freezing point 

is decreased, e.g., by trehalose and glycerol or other polyhydric alcohols as 

anti-freeze agents which prevent ice-crystal formation within the hypha (Jennings 

and Lysek 1999). In some blue-stain and mold fungi, the lower limit for 

mycelial growth is at −7 to −8 ◦ C (Reiß 1997). Above the lower limit, the “reaction 

speed-temperature rule” begins to take effect, as in a certain temperature 

range, enzyme activity runs two to four times faster by increasing the temperature 

of about 10 ◦ C(Q10 value). Frequently, the optimum lies, depending on the 

species (and isolate) between 20 and 40 ◦ C. Psychrophilic fungi have their optimum 

below 20 ◦ C, mesophilic species between 20 and 40 ◦ C and thermophilic 

species over 40 ◦ C. Thermotolerant fungi, e.g., Phanerochaete chrysosporium 

and other fungi growing in wood chip piles, prefer the mesophilic range, tolerate 

however still 50 ◦ C. The maximum for mycelial growth and wood damage 

by most wood fungi is often at 40–50 ◦ C, because then the protein (enzyme) 

denaturing by heat takes effect. Fungi, however, may exhibit a change in gene 

expression, which leads to the synthesis of “heat-shock proteins (hsp)”. The 

hsps appear to prevent and repair general damage, denaturation and aggregation 

of other cellular proteins, as they are not only induced by heat, but also by 

heavy metals and oxidants (Jennings and Lysek 1999). 

Serpula lacrymans possesses a characteristic, which can be used for identification. 

With the optimum of about 20 ◦ C, slight growth still at 26–27 ◦ C, and 

growth stop at 27–28 ◦ C, the fungus differs from the other indoor wood decay 

fungi, like the Cellar fungus and the white polypores, as well as from other 

Serpula species, because, e.g., S. himantioides still grows at 31 ◦ C. There are, 

however wild Himalayan isolates of S. lacrymans that showed slight growth 

at 32 ◦ C (Palfreyman and Low 2002). In addition, in S. lacrymans also the 

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Table 3.8. Cardinal points of temperature ( ◦ C) for fungal growth and survival (mainly from 

Humphrey and Siggers 1933; Cartwright and Findlay 1958; Ammer 1964; Cockcroft 1981; 

Mirič and Willeitner 1984; Thörnqvist et al. 1987; Viitanen and Ritschkoff 1991; data of 

house-rot fungi from Schmidt and Huckfeldt 2005) 

Species lethal minimum optimum maximum lethal lethal lethal 

on agar on agar 4 h 

in 2 weeks (h) in wood 

Armillaria mellea 25–26 33 

Aspergillus niger 35–37 45–47 

Aureobasidium pullulans 25 35 

Daedalea quercina 5 23–30 30–44 

Fomes fomentarius 27–30 34–38 

Heterobasidion annosum 2–4 22–25 30–34 

Laetiporus sulphureus 28–30 36 

Lentinus lepideus 3–8 25–33 37–40 60 (0.5) 

Paxillus panuoides 5 22–25 29 

Phellinus pini 20–27 30–35 55 (0.5) 

Piptoporus betulinus 26–30 32–36 

Polyporus squamosus 24–25 30–38 60 (0.5) 

Schizophyllum commune 28–36 44 60 (0.5) 

Stereum sanguinolentum < 4 20–22 

Trametes versicolor 24–33 34–40 55 (0.5) 

Trichaptum abietinum 22–28 35–40 

Serpula lacrymans −6 0–5 20 26–27 30 55 (3) 50–70 

Serpula himantioides 25–27.5 32.5 > 35 65 

Leucogyrophana mollusca 25–27.5 32.5 30 ≥ 35 75 

Leucogyrophana pinastri 20–27.5 32.5 > 35 

Coniophora puteana −20/−30 0–5 22.5–25 27.5 ≥ 37.5 32.5 ≥ 37.5 60 (3) 70–75 

Coniophora marmorata 20–27.5 25 ≥ 37.5 35 ≥ 37.5 

Coniophora arida 25 27.5 35 

Coniophora olivacea 22.5–25 32.5–35 35 ≥ 37.5 

White polypores (old data) 3–5 25–31 35–38 80 (0.5) 

Antrodia vaillantii 27.5–31 35 37–40 65 (24) > 80 

Antrodia sinuosa 25–30 35 37–42.5 65 (3) 

Antrodia xantha 5 27.5–30 35 40–42.5 

Antrodia serialis 22.5–25 32.5–35 37.5–42.5 

Oligoporus placenta 3 25 35 40–45 65 (24) > 80 

Gloeophyllum abietinum 0–4 25–27.5 37.5–42.5 40–42.5 > 95 

Gloeophyllum sepiarium 5 27.5–32.5 ≥ 45 ≥ 45 60 (3) > 95 

Gloeophyllum trabeum 5 30–37.5 ≥ 45 ≥ 45 80 (1) > 95 

Donkioporia expansa 28 34 > 40 65 (24) > 95 

monokaryons tolerated 28 ◦ C (Schmidt and Moreth-Kebernik 1990), so that 

probably some data in the literature concerning growth of the fungus above 

27 ◦ C (Wälchli 1977) were based on monokaryons. Last, dikaryons of S. lacrymans 

(and some further fungi) can be reverted to the monokaryotic stage by 

cultivation at relatively high temperature and thus these cultures then also 

grew above 27 ◦ C. 

From the cultivation of edible mushrooms on wood (Chap. 9.2) it is known 

that the optimal temperature can be lower for fruit body formation than for 

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3.4 Temperature 69 

mycelial growth. Cultivation of Lentinula edodes in Asia involves a dipping 

of the colonized wood in cold water to stimulate fruit body development. On 

the other hand, S. lacrymans is stimulated to fruit in laboratory culture, if 

the mycelium is incubated first 3–4 weeks at 25 ◦ C and then 2 weeks at about 

20 ◦ C (Fig. 3.1; Schmidt and Moreth-Kebernik 1991b). In some fungi, spore 

germination is activated by high temperature, in nature for example after 

forest fires. 

The temperature curve of the mycelial growth rate must not correlate with 

that one of fungal activity. For example, the temperature range for growth 

may be broader than for wood degradation (Wälchli 1977). Furthermore, the 

temperature optima of enzymes isolated from fungi are often higher (50– 

60 ◦ C) than those of mycelial growth of the respective fungus. Some wood 

fungi tolerate extreme values beyond minimum and maximum by resistance 

to cold and heat, respectively. However, there are significant differences with 

regard to the test method used. Results from cultures on agar revealed that 

S. lacrymans survived 1 h at 55 ◦ C, Coniophora puteana 1hat60 ◦ C, Antrodia 

vaillantii 3hat65 ◦ C (Schmidt 1995a), and Gloeophyllum trabeum 1hat80 ◦ C 

(Mirič and Willeitner 1984). In colonized wood samples that were slowly dried 

before heating, S. lacrymans survived 4 h at 65 ◦ C, C. puteana 4h at 70 ◦ C, A. 

vaillantii 4h at 80 ◦ C and G. trabeum 4h at 95 ◦ C, assumably by developing 

resistant arthrospores (Huckfeldt et al. 2005). This great resistance of the fungi 

to heat challenges the use of a heat treatment procedure for the eradication 

of fungi in houses. In Denmark, whole houses are subjected to heat treatment 

against S. lacrymans (Koch 1991) (Chap. 8.5.4). 

Vegetative cells (bacteria and fungal hyphae) are destroyed by heating at 

80 ◦ C (pasteurization). Exceptions with growth of up to 113 ◦ Carebacteria 

(Archaea) in volcanic biotopes (geysers, black smokers). Spores are frequently 

Fig.3.1. Fruit body formation of Serpula 

lacrymans in laboratory culture stimulatedbyawarmthtreatment;25mycelial 

growth at 25 ◦ C, 20 growth increase at 

20 ◦ C, F fruit body 

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more thermotolerant than the corresponding mycelium. The basidiospores of 

S. lacrymans were killed by 32 h at 60 ◦ Cor1hat100 ◦ C (Hegarty et al. 1986). 

However, 4 h at 65 ◦ C reduced the germination rate from 30 to 8% (Hegarty 

et al. 1987). The heat resistance of basidiospores has also to be considered in 

view of eradication of indoor wood-decay fungi by heat treatment. 

As spore forming bacteria may survive 100 ◦ C, nutrient media for laboratory 

experiments are sterilized at 121 ◦ C and 210 kPa pressure in autoclaves. 

Alternatively, fractionated sterilization at 100 ◦ C (tyndallization) may be used 

(heating at 100 ◦ C on three successive days for 30 min to destroy vegetative 

cells; between the three heat phases incubation at room temperature to allow 

germination of survived spores). Heat-sensitive nutritives can by sterilized by 

membrane filtration using filter membranes with a pore size of 0.1–0.45µm. 

Insensitive laboratory equipment like glass material becomes sterile by 1 h of 

dry heat at 180 ◦ C. Wood samples for decay experiments may be sterilized by 

ethylene oxide in special devices. 

In many fungi, spores and also mycelia are resistant to cold. Thus, fungal 

cultures in international strain collections are conserved, except by lyophilization,alsoinliquidnitrogenat−196 

◦ C and not like it is usually done in small 

laboratories in the refrigerator on agar (or also on small wood pieces: Delatour 

1991). 

3.5 

pH Value and Acid Production by Fungi 

The pH value influences germination of spores, mycelial growth, enzyme activity 

(wood degradation), and fruit body formation. The optimum for wood 

fungi is often in slightly acid environment of pH 5–6 and for wood bacteria at 

pH 7. Basidiomycetes have an optimum range of pH 4–6 and a total span of 

about 2.5–9 (Thörnqvist et al. 1987). Ascomycetes, particularly soft-rot fungi, 

may tolerate more alkaline substrates to about pH 11. Thus, the pH values from 

3.3–6.4 in the wood capillary water of living trees and in aqueous extracts of 

wood and bark samples from trees of the temperate zones and from trading 

timbers (Sandermann and Rothkamm 1959; Rayner and Boddy 1988; Fengel 

and Wegener 1989; Landi and Staccioli 1992; Roffael et al. 1992a, 1992b) correspond 

with the pH demands of wood fungi. Over the tree cross section, pH 

differences can occur, that is for example the heartwood of oaks and Douglas 

fir is more acid than the sapwood. Furthermore, an initial pH value can be 

changed in the context of microbial succession, because bacteria may acidify 

or alkalize the substrate by their metabolites (fatty acid production in acid 

wetwood or methane or ammonia formation in alkaline wetwood; Chap. 5.2). 

Outside about pH 2 and 12, respectively, microbial activity is commonly prevented. 

The acid pH-extreme of Aspergillus niger is 1.5 (Reiß 1997). There 

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3.5 pH Value and Acid Production by Fungi 71 

are however fungi that even grow at about pH 0 like a Cephalosporium species. 

Among the bacteria, the Archaea Picrophilus oshimae and P. torridus have their 

pH-optimum at pH 0.7 and even grow at pH −0.06 (Anonymous 1996). 

Various wood fungi can change pH values near the extremes by means of pH 

regulation through their metabolic activity (Rypáček 1966; Humar et al. 2001). 

Alkaline substrates are acidified by the excretion of organic acids, particularly 

oxalic acid/oxalate (Jennings 1991). Oxalic acid is synthesized by oxaloacetase 

(EC 3.7.1.1) from oxalic acetate of the citric acid cycle (Micales 1992; Akamatsu 

et al. 1993a, 1993b) and can also derive from the glyoxylate cycle (Hayashi et al. 

2000; Munir et al. 2001). Table 3.9 shows the amount of oxalic acid produced 

by some house-rot fungi in vitro and the resulting pH value. 

Figure 3.2a shows the change of the pH value by Schizophyllum commune 

as an example of the pH-regulation curve of fungi. If there would not have 

been a pH-change caused by the fungus, the diagonal in Fig. 3.2a would have 

resulted. Nutrient liquids with acidic initial pH values become alkalized. For 

example, the initial pH of 4.2 changed stepwise to the final pH of 7.5. After 

3–4 weeks of culture, a nearly straight plateau of pH 7.5 derived from the initial 

pH values 4.2, 5.1, 6.0 and 7.5. In contrast, the alkaline initial pH value of 7.5 

was acidified in the first 2 weeks of culture (Schmidt and Liese 1978). 

Aerobic bacteria alkalize their substrates by ammonia release from proteins 

and amino acids (Schmidt 1986) and anaerobic bacteria alkalize the wetwood 

in trees by methane formation (Ward and Zeikus 1980; Schink and Ward 1984). 

Is less intensively examined by which metabolic pathways fungi alkalize acid 

media. This may occur by the consumption of anions or by the formation of 

ammonia from nitrogen compounds (Schwantes et al. 1976). 

While unbuffered laboratory nutrient media approach the natural habitat 

of wood fungi and show the physiologically produced pH value of a fungus, 

buffered media of different initial pH values results in that pH-range, within 

which a fungus can grow without adjusting the pH. The pH-optima received 

Table 3.9. Content of oxalic acid (g/L) and pH-value in nutrient liquid after 2 months of 

incubation (from Schmidt 1995; Schmidt and Moreth 2003) 

Species Isolate (g/L) pH 

Antrodia vaillantii FPRL14 1.85 2.4 

R112 0.63 2.8 

BAM 65 0.65 2.8 

DFP 2375 1.20 2.4 

Antrodia sinuosa MAD 2538 1.10 2.6 

Oligoporus placenta FPRL 280 0.25 2.2 

Coniophora puteana Ebw. 15 0.04 4.2 

Serpula lacrymans BAM 133 1.85 2.4 

Donkioporia expansa MUCL 29391 0.16 4.6 

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Fig.3.2. pH value regulation by Schizophyllum commune (a) and mycelial dry matter after 

growth with different initial pH values (b) for 7–28 days (from Schmidt and Liese 1978) 

in buffered and unbuffered media can differ. For example, Schizophyllum commune 

grewbestonbufferedagaratpH4.7–5.1,butreachedinunbuffered 

nutrient liquid the highest mycelial dry matter at pH 7.5 (Fig. 3.2b). Two pHoptima 

may occur (Fig. 3.2b). Frequently, the optimum pH value of enzyme 

activity of enzymes isolated from a fungus differs in vitro considerably from 

the pH value for the corresponding fungal growth. 

Most brown-rot fungi accumulate oxalic acid (oxalate) in rather large quantities 

and acidify their microenvironment usually to a greater extent than do 

the white-rot fungi (Table 3.9: Donkioporia expansa). pH-reduction by brownrot 

fungi was thought to favor the activity of some non-enzymatic systems 

hypothesized to be active in these fungi, as well as cellulolytic enzyme activity 

(Goodell 2003). In brown-rot fungi, oxalate serves as an acid catalyst for the 

hydrolytic breakdown of wood polysaccharides (Chap. 4). The acid attacked 

the hemicelluloses and the amorphous cellulose, thus increasing the porosity 

of the wood structure for hyphae, enzymes and low-molecular degrading substances 

(Green et al. 1991a; Shimada et al. 1991). The enzyme system to produce 

oxalate was also found in the white-rot fungi like Trametes versicolor (Mu et al. 

1996). White-rot fungi accumulate smaller amounts of oxalate and use it in 

connection with the enzymatic lignin degradation by lignin peroxidase and 

manganese peroxidase. Under extracellular condition, the mediators, veratryl 

alcohol cation radicals and Mn 3+ , produced by lignin and manganese peroxidase, 

respectively, catalyze the decomposition of oxalate to CO2 (Shimada 

et al. 1994). During intercellular metabolism, oxalate is formed by oxalate 

decarboxylase (EC 4.1.1.2) to formate and CO2, and the formate produced 

is converted to CO2 by formate dehydrogenase (EC 1.2.1.2), yielding NADH 

(Watanabe et al. 2003). Oxalate may be also metabolized by oxalate oxidase 

(EC 1.2.3.4) to CO2 and H2O2. There are, however, exceptions within both fun- 

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3.5 pH Value and Acid Production by Fungi 73 

gal decay groups. Gloeophyllum trabeum, for example, degraded 14 C-labelled 

oxalic acid to CO2 during cellulose degradation (Espoja and Agosin 1991), 

and only relatively slight acidification of nutrient liquids was observed for all 

indoor Gloeophyllum species (Schmidt et al. 2002a). 

The intensive production of oxalic acid by Serpula lacrymans, which is 

reflected by an acidification of the growth medium to pH 2.4 (Schmidt 1995b, 

Table 3.9), has been discussed in connection with the preferential occurrence 

of the fungus within buildings. Excess oxalic acid is neutralized to Ca-oxalate 

by calcium from brickwork or by chelating with iron from metals, stonewool 

and nails (Bech-Andersen 1987b; Paajanen and Ritschkoff 1991, 1992; Paajanen 

1993; Palfreyman et al. 1996). 

The indoor polypores, especially Antrodia vaillantii, are resistant to copper 

mainly due to the formation of Cu-oxalate (Da Costa 1959; Sutter et al. 1983; 

Collett 1992a, 1992b; Schmidt 1995b; Chap. 8.5.3.2). Humar et al. (2002) showed 

that A. vaillantii, Gloeophyllum trabeum and Trametes versicolor transformed 

copper(II) sulfate in wood into non-soluble and therefore non-toxic copper 

oxalate. Hastrup et al. (2005) evaluated wood decay of samples impregnated 

with copper citrate and found 11 out of 12 isolates of Serpula lacrymans to be 

tolerant towards copper citrate. Table 3.10 shows the ability of some house-rot 

fungi to grow on copper-containing agar. 

Table 3.10. Copper tolerance. Growth (±) of house-rot fungi on agar containing copper 

sulphate (from Schmidt 1995; Schmidt and Moreth 2003) 

Species Isolate Molarity of copper 

0.001 0.005 0.01 0.03 0.05 

Antrodia vaillantii FPRL 14 + + + + − 

FPRL 14a + + + − − 

UK 14 + + + (+) − 

BAM 65 + + + + (+) 

BAM 486 + + + − − 

DFPG 6911 + + + + − 

DFP 2375 + + + − − 

Sweden R112 + + + (+) − 

Sweden R113 + + + − − 

Antrodia sinuosa MAD 2538 + − − − − 

Oligoporus placenta Ebw. 125 + (+) − − − 

FPRL 280 + + (+) − − 

Findlay 304A + (+) − − − 

Coniophora puteana Ebw. 15 + − − − − 

Serpula lacrymans BAM 133 + − − − − 

Serpula himantioides MAD 213 + − − − − 

Donkioporia expansa MUCL 29391 + − − − − 

(+) one of two duplicates 

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Antrodia vaillantii decreased the life-time of timber impregnated with chromated 

copper arsenate and borate, respectively. Chromium, which plays a role 

in the fixation reactions of the elements (Bull 2001; Bao et al. 2005b), and arsenate 

as well as borate became soluble by oxalic acid and were washed out by 

rain (bioleaching) (Göttsche and Borck 1990; Cooper and Ung 1992a). Copper 

is precipitated into the insoluble form of the oxalate, rendering the copper 

inert. This leaching effect was used for biological remediation (recycling) of 

CC-treated wood waste (Leithoff et al. 1995; Stephan et al. 1996; Samuel et al. 

2003; Kartal and Imamura 2003). Arsenic and chromium free copper-organic, 

alternative preservatives which were recently developed in view of health and 

environmental aspects were also attacked (Humar et al. 2004; cf. Chap. 7.4). 

There are further possible candidates for bioremediation of CCA-treated wood 

such as Laetiporus sulphureus (Kartal et al. 2004). 

3.6 

Light and Force of Gravity 

At first sight, light might have no significance for fungi, because fungi are 

carbon-heterotrophic. The vegetative mycelium including the rhizomorphs 

of Armillaria species and the strands of house-rot fungi grow in nature in 

the absence of light, namely in the soil and within trees or timber (substrate 

mycelium), or in buildings hidden behind wall coverings and in the subfloor 

area. The growth within the substrate might be rather due to hygro-, hydro-, 

geo- and chemotropisms than to negative phototropism (Müller and Loeffler 

1992). Surface and aerial mycelia also grow in the dark like during the routine 

fungal culturing in the laboratory or at low light intensity like in the indoor 

polypores and Serpula lacrymans in buildings. 

A requirement for light occurs particularly with respect to the initiation of 

reproduction and the ripening of the fruit bodies. Light is the signal that the 

mycelium has reached the (irradiated) surface, where there the spores can be 

produced in an environment suitable for spore release (Jennings and Lysek 

1999). For fungi, light in the short wavelengths, blue light, is effective, while 

light with longer wavelengths is ineffective. The light acceptor of the photons 

hitting the mycelium is riboflavin, which then reduces a cytochrome. The 

required light quantities are low, below those of the full moonlight at a clear 

sky (0.23µWcm −2 ). 

During the cultivation of Lentinula edodes on wood (Chap. 9.2), the colonized 

wood substrate was exposed to light for 8–15 h/day (Schmidt 1990). 

In the dark, the primordia did not develop further or abnormal fruit bodies 

occurred. Particularly suitable are wavelengths from 370–420 nm and from 

620–680 nm. Daedalea quercina, Gloeophyllum abietinum, Lentinus lepideus, 

Paxillus panuoides and some other fungi develop abnormal and frequently 

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3.6 Light and Force of Gravity 75 

sterile“darkfruitbodies”onminetimber.Serpula lacrymans fruits in buildings 

in twilight. In the laboratory, the daily light-dark cycles are suitable. 

The Deuteromycetes Aspergillus niger and Paecilomyces variotii develop 

conidia both with light and in the dark, likewise the ascomycete Chaetomium 

globosum forms fertile cleistothecia. In other Ascomycetes, conidia formation 

is induced by light, while in darkness ascospores develop (Reiß 1997). LightdarkcyclesleadtoarhythmicchangeofgrowthandreproductionofPenicillium 

species and other Deuteromycetes. When the hyphae are irradiated, 

their growth rate is reduced to differentiation into conidia. Concentric rings 

develop on agar plates from the inoculum in periodically repeated distances 

(Schwantes 1996; Reiß 1997; Jennings and Lysek 1999). 

Some fungi can grow permanently on sites exposed to light, e.g., fungi 

growing on plant surfaces (leaves, phylloplane). Typical phylloplane fungi 

are Alternaria, Aureobasidium and Cladosporium species (Jennings and Lysek 

1999). Some of them are potential parasites, but also effect blue stain of timber 

as saprobionts. 

UV light, particularly 254 nm, has a lethal and mutagenic effect. Nucleic 

acids are damaged by UV-B of 260 nm by the photochemical induction of cyclobutan 

dimers, which prevents the correct transcription and reduplication 

of DNA (Panten et al. 1996). That is mycelium and colorless spores and bacteria 

can be damaged by sunlight. Microbial pigmentation, particularly black 

(conidia of Aspergillus niger) and yellow (e.g., bacterium Micrococcus luteus), 

is interpreted as a protection against the irradiance. UV is thus used in microbiological 

and molecular laboratories to reduce the amount of bacteria and 

fungi in the air, on laboratory surfaces and devices. 

Fungi may also use the direction from which the light is coming to orientate 

themselves (Jennings and Lysek 1999). During the primordium growth 

of Basidiomycetes, the stipe grows towards the light source. In the Pilobolus 

species (Mucoraceae), there is a ring of yellow-orange carotenoids in the sporangiophore 

below the subsporangial bladder, which is shaded by the spore 

mass in the sporangium. If the light received by the ring is not at a minimum, 

the sporangial stalk bends until it is, which gives the direction in which 

the sporangium will be shot off, up to a distance of 40 cm (Jennings and Lysek 

1999). The force of gravity takes effect immediately when the developing 

pileus shades the tip of the stipe (Schwantes 1996). This ensures that the pores 

and lamellae in the growing hymenium orientate to the earth’s center (positive 

gravitropism, Nultsch 2001) that is, the mature basidiospores can sink 

tothesoil.Aknownexampleofpositivegravitropismmayoccurinthefruit 

body of Fomes fomentarius. The perennial, bracket fruit bodies are located 

at the stem of beech trees. When the white-rotten tree is wind-thrown, the 

fungus lives for a certain time as saprobiont in the laying stem. The new hymenia 

developing on the “old” fruit body orientate with a 90-degree change 

of direction again towards the earth’s center. If there is by chance a further 

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Fig.3.3. Fruit body gravitropism of Flammulina velutipes growing a on wood chips in the 

laboratory, 5 days old, b during 1 × g conditions on a centrifuge in the orbit, 5 days old, 

c under micro-gravitation influence during the D2 Spacelab mission 1993, 7 days old. (from 

Kern and Hock 1996). d Fruit body of Schizophyllum commune grown during turning the 

dish upside down 

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3.7 Restrictions of Physiological Data 77 

change of the stem direction, the next hymenia again follow this change. 

An exception among the hymenia of following the gravity occurs in the resupinate 

fruit bodies of house-rot fungi. The hymenium points upwards in 

fruit bodies growing on the floor, and orientates to the side in fruit bodies 

growing on a wall. The gravity perception in fungi was investigated by 

fruiting experiments with Flammulina velutipes under micro-gravitation condition 

during the German D-2 spacelab mission 1993 in the US space shuttle 

Columbia (Kern et al. 1991; Fig. 3.3). A positively gravitropic reaction can be 

simply demonstrated in the laboratory if a Petri dish with grown mycelium 

of a well-fruiting fungus like Schizophyllum commune is upside down for 

fruiting. 

According to the statolithe theory, amyloplasts and the cytoskeleton in statocyte 

cells are involved in gravitropic reactions of plants. Fungi however do 

not possess statolithes. Gravity reaction of F. velutipes was hypothesized to 

occur as follows: In the case of correct negative gravitropic adjustment of 

the fruit body, a mycohormon that is produced in the lamellae is permanently 

transported into the upper pileus area. The hormone effects a length 

increase on all sides, mediated by the synthesis of vesicles and their following 

insert. Incorrect adjustment effects an unequal hormone distribution that influences 

vesicle formation and subsequent unilateral stretching growth (Kern 

1994). 

3.7 

Restrictions of Physiological Data 

Dataintheliteraturewithrespecttothephysiologyofwoodfungilikegrowth 

reactions to environmental factors should be valued with proviso. First, a fungus 

may be misnamed due to wrong identification. Thus, DNA-analyses of 

closely related house-rot fungi of the genera Antrodia and Coniophora, respectively, 

have shown that about 15% of all investigated isolates belonging 

to these genera and sampled from own and various other strain collections 

were wrongly identified. As extreme, an isolate named A. serialis revealed to 

be Donkioporia expansa (Schmidt and Moreth 2003). Second, due to changes 

in the taxonomy, there may be considerable confusion in older references, e.g., 

with respect to Antrodia vaillantii and Oligoporus placenta,becausebothhad 

been termed Poria vaporaria (Domański 1972). Third, generalizing statements, 

like a fungus is faster growing than others, have to be restricted, because there 

is considerable strain variation within a species. Table 3.11, based on isolates 

that had been verified by rDNA-ITS sequencing, shows as an example for variation 

that there are isolates of the so-called “fast-growing” Coniophora puteana 

exhibiting a lower growth rate than isolates of the “medium-growing” Antrodia 

vaillantii. Fourth, comparisons between different fungi/authors/publications 

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are only valuable if the test methods used are comparable and appropriate. It 

is for example senseless to compare both species in Table 3.11 with respect to 

the growth rate, if the experiment did not consider the different temperature 

optima of the species. 

Table 3.11. Examples for isolate variation within wood fungi (compiled from Schmidt et al. 

2002; Schmidt and Moreth 2003) 

Species Isolate with origin and year of isolation Temperature Maximum 

optimum daily radial 

growth at 

optimum 

temperature 

( ◦C) (mm) 

Antrodia FPRL 14, originally CBS 31 5.4 

vaillantii FPRL 14a, fruit body, UK 1936 28–31 4.3 

UK 14, via Denmark and BAM Berlin 28 5.5 

DFPG 6911, New Zealand 1953 28 5.4 

DFP 2375, BAM 28 5.8 

Sweden R112, greenhouse, Stockholm ≈ 1952 28 5.1 

Sweden R113 28–31 5.6 

Ottawa 11740, USA? 28 5.1 

BAM 65 25–28 4.9 

BAM 486 28 6.1 

Coniophora UK, FPRL 11e 22.5 2.5 

puteana BK-C-50, Uppsala 25 6.3 

74453-2, Uppsala 22.5 5.0 

FORINTEK 9 0, fruit body, Ontario 1973 25 4.8 

Eberswalde 15, ‘Normstamm I’ 1930 25 7.0 

BAM 260, building, Berlin 1940 22.5 4.5 

Zycha, München 1963 25 3.5 

outdoor fruit body, Hamburg 1997 22.5 7.0 

G 61, fruit body, cherry-tree, Karlsruhe 1985 22.5–25 4.8 

G 98, building, Karlsruhe 1990 25 7.5 

G 100, building, Karlsruhe 1990 22.5 9.3 






G 220, building, Karlsruhe 1996 22.5–25 10.0 

fruit body, Ludwigslust Castle 1998 22.5 10.5 

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3.8 Competition and Interactions Between Organisms 79 

3.8 

Competition and Interactions Between Organisms 

Except for axenic laboratory cultures, there are only a few cases in which 

a natural substrate remains occupied by only one species. A known case is 

Oudemansiella mucida in standing, but dead trunks of Fagus sylvatica due to 

the production of the antifungal compound, mucidin. Instead, nearly every 

substrate accessible to fungi can support more than one species (Rayner and 

Boddy 1988), that is, various fungi and bacteria compete for space, nutrients, 

water, and air. Each fungus has its own strategy to withstand competition. 

Competition may occur between species and between mycelia of the same 

species. As a result of the latter, wood colonized by Trametes versicolor shows 

that the individual colonies form black barrier (demarcation) lines, where the 

differentmyceliahaveinteractedwitheachothertoinhibitfurthermovementof 

each mycelium in the region of contact. Different parts of the same mycelium 

and even adjacent hyphae may compete. For example, reproducing hyphae 

might consume more nutrients and thereby affect the vegetatively growing 

hyphae. 

There are three main categories of the strategies or adaptations to ecological 

niches (Jennings and Lysek 1999). Through combative strategy, the fungus 

defends the substrate that has already been captured or attacks competitors 

occupying a substrate that is capable of capture (e.g., O. mucida). Through 

ruderal strategy, a substrate as yet unoccupied or only partly colonized is 

exploited. Those fungi do not attack potentially resistant substrates but degrade 

readily consumable or unusual compounds, like Pholiota carbonica (Europe, 

North America, Asia, North Africa) and P. highlandensis (USA), which both 

grow on former fire sites (Breitenbach and Kränzlin 1995). So these fungi 

occupy a substrate faster than possible competitors. Fungi concerned in the 

stress-tolerant strategy are adapted to environments that are too harsh for 

possible competitors. Examples for the latter are the soft-rot fungi growing in 

verywettimberoflowaircontent. 

3.8.1 

Antagonisms, Synergisms, and Succession 

Interactions (reciprocal effects) between wood fungi have been early investigated 

e.g., by Oppermann (1951) and Leslie et al. (1976), and were described 

in detail by Rayner and Boddy (1988). 

Antagonism (competitive reciprocal effect), the mutual inhibition and in 

abroadersensetheinhibitionofoneorganismbyothers,isbasedontheproduction 

of toxic metabolites, on mycoparasitism, and on nutrient competition. 

Antagonisms are investigated as alternative to the chemical protection against 

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tree fungi (“biological forest protection”) and against fungi on wood in service 

(“biological wood protection”) (Wälchli 1982; Bruce 1992; Holdenrieder and 

Greig 1998; Phillips-Laing et al. 2003). 

As early as 1934, Weindling showed the inhibiting effect of Trichoderma 

species on several fungi. Bjerkandera adusta and Ganoderma species were 

antagonistic against the causing agent of Plane canker stain disease (Grosclaude 

et al. 1990). Also, v. Aufseß (1976) examined mycelial interactions between 

Heterobasidion annosum and Stereum sanguinolentum and antagonistic fungi 

like Phlebiopsis gigantea and Trichoderma viride (also Holdenrieder 1984). 

Root rot by Heterobasidion annosum (Chap. 8.3.2) is the classical target for 

biological forest protection and the only example of a successful biological 

control of a fungal forest disease. Based on the work of Rishbeth, stump treatment 

with Phlebiopsis gigantea was developed and successfully used in several 

countries. Originally in England, the spread of root rot in pine sites was diminished 

by the immediate coating of the fresh stump surface with an aqueous 

spore (asexual arthrospores) suspension of P. gigantea (Meredith 1959; Rishbeth 

1963). The antagonist colonizes the stump, that is H. annosum cannot 

infect it by air-borne spores and thus an infection of neighboring trees via 

root grafts is prevented. The treatment of spruces yielded differently satisfactory 

results (Korhonen et al. 1994; Holdenrieder et al. 1997). Holdenrieder and 

Greig (1998) listed also several bacteria, which were antagonistic against H. annosum. 

Promising systems for the biological protection of growing trees have 

been studied against Armillaria luteobubalina, Chondrostereum purpureum, 

Phellinus tremulae, P. weirii, and Ophiostoma ulmi (Bruce 1998; also Palli and 

Retnakaran 1998). 

There were many attempts for biological wood protection (Bruce 1998). 

To date, the application of biological control to prevent wood decay and discoloration 

has been successful in the laboratory, but was often inconsistent 

in its performance in the field (Dawson-Andoh and Morrell 1997; Mikluscak 

and Dawson-Andoh 2004b). Much work has been done in the Forest Products 

Laboratory, Madison. In the laboratory, a blue stain fungus was inhibited by 

antibiotic substances from Coniophora puteana (Croan and Highley 1990) and 

Bjerkandera adusta (Croan and Highley 1993). Bacteria were examined for 

their suitability to prevent of blue stain (Bernier et al. 1986; Seifert et al. 1987; 

Benko 1989; Florence and Sharma 1990; Kreber and Morrell 1993; Bjurman 

et al. 1998; Payne et al. 2000; Bruce et al. 2004). A bacterial mixed culture decreased 

staining and molding of pine wood samples as well as decay by Trametes 

versicolor and Oligoporus placenta (Benko and Highley 1990). Streptomyces rimosus 

Sobin, Finlay & Kane (Croan and Highley 1992b) and its culture filtrate 

(Croan and Highley 1992c) prevented spore germination of Aspergillus niger, 

Penicillium sp. and Trichoderma sp.aswellasbluestainbyAureobasidium 

pullulans. Trichoderma species are extensively researched biological control 

agents for wood protection against decay fungi (Highley and Ricard 1988; 

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Murmanis et al. 1988; Morris et al. 1992; Doi and Yamada 1992; Bruce 1998; 

Phillips-Laing et al. 2003). Culture filtrates of Chaetomium globosum, Penicillium 

sp., Sporotrichum pulverulentum and Trichoderma viride decreased wood 

degradation by T. versicolor (Ananthapadmanabha et al. 1992). 

Current attempts for biological wood protection use a colorless mutant 

of the blue-stain fungus Ophiostoma piliferum. Round wood and cut timber 

is treated with a spore suspension of the mutant to reduce or even prevent 

subsequent natural colonization of the wood by blue-stain fungi (Blanchette 

et al. 1994; Behrendt et al. 1995; Schmidt and Müller 1996; White-McDougall 

et al. 1998; Ernst et al. 2004). Corresponding experiments used Gliocladium 

roseum to protect green lumber from molds, stain, and decay (Yang et al. 

2004a). Figure 3.4 demonstrates the inhibiting effect of O. piliferum against 

two blue-stain fungi in the laboratory. 

Synergism (mutualistic reciprocal effect) means the mutual promotion and 

inthebroadersensethepromotionofoneorganismbyothers.Topreparethe 

substrate, the pH value can be changed, vitamins can be excreted (Henningsson 

1967), and inhibiting heartwood compounds can be degraded. The nitrogen 

content may be increased by N-fixing soil bacteria (Baines and Millbank 1976), 

and nutrients can become more available (also Levy 1975a; Hulme and Shields 

1975). Neutralistic reciprocal effects, neither inhibition nor promotion, occur 

more rarely. 

Fig.3.4. Inhibition by a colorless mutant of Ophiostoma piliferum of blue-staining of wood 

samples by Phoma exigua and Aureobasidium pullulans. Wood samples A were previously 

dipped in a spore solution of O. piliferum and then all four samples were inoculated with 

the blue-stain fungi (from Müller and Schmidt 1995) 

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The various competition strategies and reciprocal effects influence the sequence 

(succession) of fungi and bacteria that are found at different stages in 

the degradation of a complex substrate like wood. Each species uses a different 

component of the substrate as it becomes available as a result of the degradation 

by the preceding species (Jennings and Lysek 1999). Primary colonists, 

bacteria and non-decay fungi (slime fungi, yeasts, molds), rely on relatively 

easy assimilable substrates such as simple sugars, starch and proteins and remain 

predominantly on the wood surface and within the outer wood parts, 

preparing the substrate for following organisms. There may occur a continued 

co-existence of non-decay organisms on the substrate. Or the primary colonists 

are followed by the decay fungi which are capable of degrading the relatively 

refractory wood cell wall components and which penetrate deeper into the 

wood such as staining fungi and the brown, soft and white-rot fungi (Levy 

1975a; Käärik 1975; Rayner and Boddy 1988). 

Schales (1992) found 15 wood-decay fungi on a wind-thrown beech tree 

and its stump. Chondrostereum purpureum and Stereum hirsutum occurred 

during the initial phase of 2 years. Bjerkandera adusta and Trametes versicolor 

were common in the following medium (optimum) phase of 5–7 years. 

Kuehneromyces mutabilis and Kretzschmaria deusta were observed in the final 

phase (also Jahn 1990; Röhrig 1991). Ten beech stumps showed within 4 years 

after tree felling 74 fungal species, 46 Basidiomycetes, 25 Ascomycetes and three 

Deuteromycetes (Andersson 1997a; also Willig and Schlechte 1995; Andersson 

1997b; Blaschke and Helfer 1999). Those surveys indicate that a substrate is 

colonized by more species than commonly described in literature and that 

some fungi occur earlier than expected. 

While most fungi colonizing wood use nutrients of the substrate, some are 

probably only passive occupants using the wood only as a support for fruit 

body formation. 

Interrelationships between trees and the fungi that inhabit them have been 

treated by Rayner (1993). 

3.8.2 

Mycorrhiza and Lichens 

Mycorrhiza (“fungal root”) is the association of mutual benefit (mutualistic interaction) 

between a fungus and the root of a higher plant (Agerer et al. to 1986; 

Willenborg 1990; Allen 1991; Schwantes 1996; Smith and Read 1997; Varma and 

Hock 1999; Egli and Brunner 2002; v.d. Heijden and Sanders 2002; Peterson 

et al. 2004). About 80–95% of the higher plants are capable of mycorrhization 

(e.g., Bothe and Hildebrandt 2003). 

Mycorrhizas are differently grouped. The grouping according to Hock and 

Bartunek (1984) in Fig. 3.5 distinguishes three major forms. 

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Fig.3.5. Major forms of mycorrhizas. 

Ek ectotrophic, En endotrophic, VA 

vesicular-arbuscular (modified from 

Hock and Bartunek 1984) 

The ectotrophic mycorrhiza (ectomycorrhiza) occurs predominantly on 

conifers and hardwoods of the boreal and temperate zone, particularly associated 

with Pinales, Fagales and Salicales. In many conifers and in beech 

and oak, the association is obligatory, and in other trees like elms it is facultative 

(Müller and Löffler 1992). The predominant part of the mycelium grows 

at the surface of side roots and forms a dense mycelial coat at the root tips. 

The hyphae penetrate between the cells of the outer root tissue by dissolving 

the middle lamellae, and coat the cells completely as “Hartig net” (Kottke 

and Oberwinkler 1986). The colonized roots do no longer possess root hairs; 

instead hyphae or rhizomorphs radiate into the soil. 

In the endotrophic mycorrhiza (endomycorrhiza) of the orchids, only a loose 

hyphal net is formed around the root, and the hyphae settle inside the cells 

in the root bark area. As an intermediate, the ectendotrophic mycorrhiza is 

particularly present on roots of 1 to 3-year-old conifers, whereby the hyphae 

penetrating into the bark cells degenerate with ageing. 

The most frequent form, the vesicular arbuscular mycorrhiza (VAM) occurs 

associated with over 200,000 wild and cultivated angiosperms, in addition, 

with Ginkgo biloba, Taxus baccata and Sequoia gigantea and S. sempervirens 

(Werner 1987), as well as predominant form in tropical forests. In the VAM, 

the unseptate hyphae extend inside the root cells bubble-shaped (vesicles) 

or branch out tree-shaped (arbuscules). The arbuscules develop by hyphal 

branching and become enclosed by the peri-arbuscular membranes from the 

plant (Bothe and Hildebrandt 2003). 

The benefit for the trees is the improved nutrient (amino acids) and mineral 

(N, P, K, Mg, Cu, Zn, Fe) support and the better water supply (Smith and 

Read 1997) due to the larger absorption area. Soils with frequently occurring 

ectomycorrhiza are commonly characterized by a lower nutrient content, the 

trees growing there would not be competitive without mycorrhizas (Schönhar 

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1989). The soil quality (ventilation, water permeability, stabilization of soil 

particles) is increased. The trees are more resistant to drying stress. In addition, 

mycorrhizal fungi play a role in tree defense against fungal pathogens 

(Strobel and Sinclair 1992). The fungi benefit from the supply of photosynthates 

(carbohydrates) from the trees and from supplements, e.g., thiamine. 

As much as 30–35% of the photosynthate by a beech forest is metabolized by 

the mycorrhizal fungi (Jennings and Lysek 1999). 

About one-third (2,000 species) of the “higher fungi” which grow in forests 

are mycorrhizal fungi (Egli and Brunner 2002). Among them there are many 

edible mushrooms (e.g., Boletus edulis, Cantharellus cibarius), but also poisonous 

species (e.g., Amanita species). Endotrophic mycorrhizal fungi are 

usually Ascomycetes. Ectotrophic fungi are usually Basidiomycetes such as 

Amanita species, B. edulis or the truffles (Ascomycetes). The about 150 VAM 

symbionts belong to the Zygomycetes, often to the genus Glomus. 

Many trees, like beech, oak, spruce, chestnut, pine, larch and willow, become 

stunted in sterile culture and previous mycorrhizal inoculation of seedlings 

improved tree growth (Ortega et al. 2004). Several obligatory mycorrhizal fungi, 

like B. edulis, only fruit in association with roots, partly host-specifically or 

with a narrow host spectrum, like Amanita caesarea predominantly associated 

with oaks, usually however hardly host-specifically, like A. muscaria at birch, 

eucalypts, spruce and Douglas fir (Werner 1987). The trees are usually less 

specific: Pinus sylvestris forms mycorrhizas with at least 155 fungal species and 

Picea abies with 118 fungi (Korotaev 1991). 

Artificial mycorrhization may de done in the tree nursery or during planting 

or by injection in the root area of old trees (Egli 2004; Evers and Pampe 

2005). About 500,000 l mycorrhizal inoculum was produced worldwide in 2003 

(Grotkass et al. 2004). 

With regard to the significance of the mycorrhizas in view of the forest 

dieback by pollution (Flick and Lelley 1985), there is a trend that young trees 

already show a fungal community, which is typical for old trees. The changed 

mycorrhiza was rated as signal for tree damage: “The fungi disappear before the 

trees” (Cherfas 1991). A negative correlation was found between the frequency 

of fungal occurrence and the content of nitrogen and sulfur compounds as 

well as ozone in the atmosphere: 71 species of fungi were observed in a certain 

area of the Netherlands from 1912–1954 and only 38 species between 1973 and 

1982. Also, the size of the fruit bodies decreased (Cherfas 1991). According 

to Schönhar (1989), the change of the mycorrhiza is particularly based on 

nitrogen imissions by fertilization. The possible role of mycorrhiza in forest 

ecosystems under CO2-enriched atmosphere in view of the global atmospheric 

change was discussed (Quoreshi et al. 2003). Experimental drought investigated 

in view of the expected reduction in water in Mediterranean regions showed 

that drought treatment did not delay mushroom appearance, but reduced 

mushroom production by 62% (Ogoya and Peñuelas 2005). 

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Investigations have been performed to regenerate the decreased mycorrhizal 

occurrence and the species change in forest damage sites by artificial 

inoculation and thus to improve the health of these trees and also of trees on 

other problematic sites (Römmelt et al. 1987; Marx 1991; Schmitz 1991; Lelley 

1992; Hilber and Wüstenhöfer 1992; Schmitz and Willenborg 1992; Göbl 1993; 

Kutscheidt and Dergham 1997). However, it has to be considered that thereby 

one intervenes only at the symptoms of the damage and not at its causes, that 

is, new inoculations without reduction of the emissions might be unsuccessful 

in the long run. To improve the isolate characteristics of mycorrhizal species, 

interstock matings have been done e.g., with Paxillus involutus (Strohmeyer 

1992). 

A further association of mutual benefit is lichens, a close and stable partnership 

between Ascomycetes (and rarely Basidiomycetes) with green algae 

or cyanobacteria (Kappen 1993). In the mutualistic form of lichens, the fungi 

receive organic nutrients and vitamins from the algae/bacteria and these get 

water and inorganic salts from the fungi. The association allows the pioneer 

settlement of inhospitable biotopes such as rocks with only traces of nutrients. 

In the antagonistic form, the fungi are parasitic to the algae, and the algae 

survive by increasing faster than they are destroyed by the fungi (Schubert 

1991). With respect to classification, the lichens are placed in the fungal system 

as lichenized fungi. 

Fungal associations with animals are the endosymbioses in the mycetomes 

of insects. Ectosymbioses occur in the “fungal gardens” of termites and in the 

cultivation of the ambrosia fungi in the drill ducts of bark beetles (Francke- 

Grosman 1958; Werner 1987). For example, Ips typographus is associated 

with ophiostomatoid fungi (Solheim 1999; Sallé et al. 2005). The fungi are 

transferred to the tree during the beetle attack and are considered important 

partners in beetle population establishment. In addition, fungi invade the 

host’s phloem and sapwood, where the hyphae can cause blue stain. Recently, 

a symbiosis between three partners was found: leaf cutter ants in Panama and 

Ecuador are associated with a basidiomycete fungus, but additionally with 

abacterium(Streptomyces sp.) which was shown to be antagonistic against 

a parasitic ascomycete that has a negative effect on the ant/basidiomycete interaction 

(Anonymous 1999). Aspects of the association of fungi and insects 

with the infected trees are described by Raffa and Klepzig (1992). 

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4 

Wood Cell Wall Degradation 

4.1 

Enzymes and Low Molecular Agents 

In view of the historical development of the research on wood degradation 

by fungi, this chapter starts with the enzymes involved in the decay of the 

woody cell wall, although it is now commonly accepted that non-enzymatic, 

low molecular weight metabolites are involved as precursors and/or co-agents 

with enzymatic cell wall degradation. 

Under the conditions within microbial cells, namely an aqueous environment 

with pH values around 6 and temperatures of 1–50 ◦ C, most reactions 

would run off only very slowly. Enzymes reduce the amount of the necessary 

activation energy as biocatalysts and control the reaction by substrate and 

effect specificity. More than 3,000 enzymes are described. 

Comparable with the lock/key principle, enzymes possess an active center, 

into which the substrate must fit, and which thus controls the conversion of 

the correct substrate (substrate specificity). The protein portion of the enzyme 

decides on the way of the reaction (effect specificity). Enzymes may consist 

only of protein or contain additional cofactors (e.g., Mg 2+ ,Mn 2+ ) or coenzymes 

(e.g., vitamin B1). Before the conversion of the substrate into a product, the 

enzyme substrate complex is formed: enzyme E + substrate S → enzyme 

substrate complex ES → enzyme E + product P. 

Studies on fungal polysaccharide hydrolyzing enzymes have shown a structural 

design composed of two functional domains, a catalytic core responsible 

for the actual hydrolysis and a conserved cellulose-binding terminus, with an 

intervening, glycosylated hinge region. A large number of genes encoding cellulases, 

hemicellulases, glucanases, amylolytic enzymes, and those hydrolyzing 

various oligosaccharides have been cloned from fungi. The best-studied organisms 

are Trichoderma reesei, Phanerochaete chrysosporium, andAgaricus 

bisporus in respect of cellulases and hemicellulases, and several Aspergillus 

species in respect of amylolytic enzymes, pectinases and hemicellulases (reviews 

by Penttilä and Saloheimo 1999; Kenealy and Jeffries 2003). For example, 

papain cleavage of cellobiohydrolase (CHB) from P. chrysosporium separated 

the catalytic domain from the hinge and binding domains. Restriction mapping 

and sequence analysis of cosmid clones showed a cluster of three structural 

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88 4 Wood Cell Wall Degradation 

related CHB genes. Within a conserved region, the deduced amino acid sequences 

of P. chrysosporium cbh1-1 and cbh1-2 were, respectively, 80 and 69% 

homologous to that of the Trichoderma reesei CBH I gene. Transcript levels of 

the three P. chrysosporium CHB genes varied, depending on culture conditions 

(review by Highley and Dashek 1998). Binding domains specific for xylan have 

also been identified (review by Kenealy and Jeffries 2003). 

Because of their valuable protein character, constitutive enzymes always 

present in the cell are the exception. Usually, the biosynthesis of the inducible 

enzymes is induced, if its presence is necessary, by the substrate or other 

molecules. Some work was done with regard to the regulation of extracellularly 

acting enzymes in fungi. For example with white-rot fungi, cellulase 

synthesis is induced in vitro by cellulose and repressed by glucose. As the wood 

cell-wall macromolecules are degraded outside the hypha, the most generally 

accepted view of the induction process is that the fungi produce a basic level 

of constitutive amount of enzyme that produces soluble degradation products 

that function as inducers. In Phanerochaete chrysosporium, which has served 

as a model organism for white-rot degradation studies, cellobiose concentration, 

a product of cellulase action, is controlled in at least four ways, by 

β-glucosidase, transglucosylation reactions, and two oxidative enzymes. As 

with cellulases, simple sugars repressed the production of most hemicellulosedegrading 

enzymes by white-rot fungi (review by Highley and Dashek 1998). 

For the naming of enzymes, particularly in former times “ase” was added 

to the name of the substrate (e.g., lignin, ligninase). Nowadays, the enzyme 

nomenclature indicates the enzymatic reaction. In accordance with the Nomenclature 

Committee of the International Union of Biochemistry and Molecular 

Biology (www.chem.qmul.ac.uk/iubmb/enzyme), enzymes are grouped 

according to their function into six classes and there into sub-groups: Oxidoreductases 

catalyze oxidation and reduction reactions by transferring hydrogen 

and/or electrons, transferases the transmission of different groups. 

Hydrolases hydrolyze glucosides, peptides etc., lyases catalyze non-hydrolytic 

cleavage, isomerases cause among other things reversible transformations of 

isomeric compounds, and ligases catalyze the covalent linkage of two molecules 

with simultaneous ATP cleavage. Each enzyme receives an EC number, which 

points out to its reaction. For daily use, the common, trivial names (ligninase, 

cellulase, xylanase), however, are still used. 

Some general characteristics of enzymes and of those enzymes involved in 

wood degradation are summarized in Table 4.1. 

The dry matter of wood consists of about 45% cellulose and, depending on 

wood species, of 20–30% hemicelluloses and 20–40% lignin. With exception 

of pectins in the middle lamella, which has significance to wood-inhabiting 

bacteria, further components such as the contents of parenchyma cells, resins, 

accessory compounds etc. are less considered in the following. Thus, relatively 

few enzymes are involved in the primary, extracellular enzymatic wood decay. 

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4.1 Enzymes and Low Molecular Agents 89 

Table 4.1. Some characteristics of enzymes for wood degradation 

altogether six classes of enzymes with subgroups: 

1. oxidoreductases, 2. transferases, 3. hydrolases, 

4. lyases, 5. isomerases, 6. ligases 

lignin degradation by oxidoreductases: 

lignin peroxidase EC 1.11.1.14 

manganese-peroxidase EC 1.11.1.13 

laccase EC 1.10.3.2 

hemicellulose degradation by hydrolases: 

endo-1,4-β-xylanase EC 3.2.1.8, xylan 1,4-β-xylosidase EC 3.2.1.37 

mannan endo-1,4-β-mannosidase EC 3.2.1.78, β-mannosidase EC 3.2.1.25 etc. 

cellulose degradation by hydrolases: 

cellulase EC 3.2.1.4 

β-glucosidase EC 3.2.1.21 

cellulose 1,4-β-cellobiosidase EC 3.2.1.91 etc. 

ectoenzymes: 

extracellular degradation of macromolecules pectin, hemicellulose, cellulose, lignin 

outside the hypha 

[uptake of degradation products (carbohydrate oligomers, dimers, monomers, 

lignin degradation products)] 

intracellular enzymes: 

metabolic transformation within the hypha to hyphal biomass, metabolites, energy 

endoenzyme: 

attack within the substrate, often randomly 

exoenzyme: 

attack at the non-reducing substrate end 

enzyme activity: 

in former times (but still used): 

international unit: 1 U = 1 µmol/min, (1 U = 16.67nkat) 

currently: 

kat (katal, catalytic activity): 1 kat = 1 mol/s 

The enzymes for the degradation of the cellulose and hemicelluloses within 

thewoodycellwallbelongpredominantlytothehydrolases,whichcleaveglucosidic 

bonds. Briefly (and thus not totally correctly), cellulose is hydrolyzed 

by cellulase, cellulose 1,4-β-cellobiosidase and β-glucosidase. The hemicelluloses 

xylan and mannan are degraded by endo-1,4-β-xylanase and mannan 

endo-1,4-β-mannosidase, respectively, followed by xylan 1,4-β-xylosidase and 

β-mannosidase and further enzymes for the side chains. Lignin is oxidatively 

degraded by the oxidoreductases lignin peroxidase and manganese peroxidase. 

Enzymatic wood degradation was summarized e.g., by Eriksson et al. 1990, 

Shimada 1993, Jennings and Lysek 1999, Goodell et al. 2003. 

Cellulose, hemicellulose, and lignin are as macromolecules too large to be 

taken up into the hypha. Therefore, the molecules are first degraded by extra- 

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cellular enzymes (ectoenzymes) into smaller fragments, which are taken up 

and then metabolized by intracellular enzymes to energy and fungal biomass. 

Independent of this place of action, an exoenzyme attacks at the end of a macromolecular 

substrate, while an endoenzyme splits within the molecule. These 

four terms are sometimes mixed up. 

Occurrence and distribution of enzymes and metabolites inside hyphae, in 

the hyphal slime layer, and within the attacked woody tissue were investigated 

by means of immunological methods and electron microscopy (Sprey 1988; 

Goodell et al. 1988; Srebotnik et al. 1988a; Blanchette et al. 1989, 1990; Daniel 

et al. 1989, 1990; Srebotnik and Messner 1990; Kim 1991; Green et al. 1991b; 

Kim et al. 1991a, 1991b, 1992, 1993; Lackner et al. 1991; Blanchette and Abad 

1992). TEM of immunogold-labeled hyphae of Trametes versicolor grown on 

carboxymethylcellulose localized β-glucosidase on the plasmalemma, in the 

hyphal cell wall, and in the hyphal sheath (review by Highley and Dashek 1998). 

Simple methods are used in screening tests to detect enzymes and to determine 

their activity. For example, a cellulose is added to a fungal culture, whose 

cellulolytic enzymes produce glucose. The glucose of the sampled culture filtrate 

reduces a test compound, which is added in oxidized form and changes 

its color by reduction. At a specific wavelength, the quantity of the converted 

test substance and thus of the developed glucose is measured and the activity 

of “cellulolytic enzymes” is calculated. Remazol brilliant blue, which binds 

to cellulose and hemicellulose by a microbially relatively inert linkage, may 

be mixed in agar. If cellulolytic or hemicellulolytic microorganisms or their 

enzymes are present, the still colored degradation products are released and 

clearing zones occur around the active colony, which can be also quantified 

(Schmidt and Kebernik 1988; Takahashi et al. 1992). For detailed investigations, 

various purification and enrichment steps may be used (chromatography, electrophoresis, 

etc.). 

The current unit of enzyme activity is katal (catalytic activity, kat), although 

in practice the old definition U is still used (Table 4.1). 

Microbial wood degradation is influenced by several major characteristics 

of the substrate wood (Table 4.2). 

Accessory compounds in the heartwood as well as resin excretion and wound 

reactions after wounding inhibit the colonization and spread of microorganisms 

within the tree (Chap. 3.1). 

The polymeric structure of the nutrients cellulose, hemicelluloses, and 

lignin requires that the degrading agents act outside the hypha. Cowling (1961) 

first stated that the known enzymes of the time were too large to penetrate 

intotheinteriorofthewoodcellwallandhypothesizedapossibleexistence 

of a small mass enzyme. The molecular weights of cellulases range from 13– 

61 kDa (Fengel and Wegener 1989). A cellulase of 40 kDa can exhibit a thickness 

of about 4 nm and a length of 18 nm (Messner and Srebotnik 1989). Frequently, 

about an 8 nm size was measured (Reese 1977; Messner and Stachelberger 

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4.1 Enzymes and Low Molecular Agents 91 

Table 4.2. Characteristics that make wood recalcitrant to fungi and bacteria 

– accessory compounds in the heartwood 

– resin excretion of softwoods, wound reactions of parenchyma cells in hardwoods 

– polymeric structure of the cell wall components 

– extracellular degradation of the nutrients 

– small pore sizes within the cell wall 

– complex structure of the woody cell wall 

– partially crystalline nature of cellulose 

– incrustation of the more easily degradable carbohydrates by lignin 

– structure and non-water-solubility of lignin 

1984; Murmanis et al. 1987). Thus, cellulases are too large for diffusing into 

the capillary areas of the cell wall from 0.5–4 nm pore size (average in spruce: 

1 nm: Reese 1977; Kollmann 1987) (Keilisch et al. 1970; Flournoy et al. 1991). 

This pore size excludes compounds with kDa mass greater than 6. Bailey et al. 

(1968) postulated a “pre-cellulolytic phase”. Meanwhile, so-called low molecular 

weight agents are known to be involved in the decay of the woody cell 

wall. As the different groups of wood decay fungi differ with regard to the 

participating low molecular agents, these aspects are treated separately in the 

chapters on cellulose and lignin degradation. 

The complex ultrastructure of the woody cell wall (e.g., Booker and Sell 

1998) affects its degradation (Liese 1970; Daniel 2003). A great part of the cellulose 

is bundled up by hydrogen bonds to larger, crystalline units (“crystalline 

cellulose”, Fig. 4.3), the elementary fibrils. The crystalline nature of the cellulose 

prevents an attack of many microorganisms. Several elementary fibrils 

result by linkage with hemicelluloses in the next larger unit, the microfibril. At 

the surface of the microfibrils, hemicelluloses form a bridge to the incrusting 

lignin, as chemical bonds exist between lignin and hemicelluloses (lignin carbohydrate 

complex, Koshijima and Watanabe 2003; Fig. 4.1). Several models 

depicting this molecular arrangement have developed (e.g., Kerr and Goring 

1975; Fengel and Wegener 1989) although there is no accepted model (Daniel 

2003). 

Principally, the carbohydrates cellulose and hemicelluloses are rather easily 

degradable, however, the lignin is resistant to most microorganisms due 

to its structure of phenylpropane units and the recalcitrant linkages between 

them. Thus, lignin incrustation of the carbohydrates inhibits the access to 

the consumable holocellulose. The hydrophobic nature of lignin further prevents 

a diffusion of the degrading enzymes inside the three-dimensional giant 

molecule. 

The composition of the microbial enzyme apparatus and its regulation affect 

the type of rot. Lignin (Fig. 4.4) is effectively degraded only by white-rot fungi 

and acts for other microorganisms as a barrier against wood decay. Table 4.3 

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Fig.4.1. Scheme of the association of cellulose (C), hemicelluloses (H) and lignin (L) within 

the woody cell wall 

Table 4.3. Lignified cell wall as carbon source for microorganisms 

Organism group Degradation of Degradation 

hemicellulose cellulose lignin of isolated within the 

components cell wall 

bacteria + + − + −a yeasts − − − 

molds + + − + − 

blue-stain fungi + − − + − 

soft-rot fungi + + − + + 

brown-rot fungi + + − −(+) + 

white-rot fungi + + + + + 

a cf. Chap. 5.2 

summarizes the behavior of the different groups of organisms against the 

nutritive “lignified cell wall”. It is differentiated if the cell wall component 

is degraded within the native woody substrate or only as sole nutritive after 

isolation from the wood. Within the bacteria, yeasts, and molds, only a few 

species are able to degrade isolated cell wall components. 

4.2 

Pectin Degradation 

Pectins comprise galacturans, galactans and arabinans as complex, branched 

polysaccharides of molecular weights of about 10 3 kDa. Galacturans are predominantly 

deposited in the middle lamella/primary wall (compound middle 

lamella) and in the tori of bordered pit membranes (Fengel and Wegener 1989). 

The content of galacturans in the wood is below 1%. They consist predominantly 

of α-1,4-linked galacturonic acid units and are split by hydrolases to 

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4.3 Degradation of Hemicelluloses 93 

galacturonic acid. Further enzymes are needed for the other pectins and for 

side chains. 

Various plant pathogenic fungi and bacteria and several soil (Bacillus spp.) 

and water bacteria (Clostridium spp.)arecapableofdegradingpectin(Schlegel 

1992). Wood-inhabiting bacteria lead to the “overuptake” of wood preservatives 

and paints (Willeitner 1971) after the degradation of the bordered sapwood 

pits (Chap. 5.2) during wet storage of wood. 

4.3 

Degradation of Hemicelluloses 

Hemicelluloses of wood are a complex combination of relatively short polymers 

made of xylose, arabinose, galactose, mannose, and glucose with acetyl and 

uronic side-groups. The major hemicellulose (polyose) of hardwoods is the O- 

Acetyl-(4-O-methylglucurono)-xylan, also named glucuronoxylan or briefly 

xylan. The xylan content in hardwoods ranges from 15 to 35%. For example, 

birch wood contains 22–30% xylan and 1–4% glucomannan, while pine 

wood contains 5–11% xylan and 14–20% glucomannan (Viikari et al. 1998). 

In monocotyledons, hemicelluloses may amount to 40% and exceed the cellulose 

portion. Beech wood xylan consists of about 200 β-1-4-linked xylose 

units (xylopyranose). About five to seven acetyl groups (linked to C-2 or 

C-3) and one 4-O-methylglucuronic acid residue (α-1-2) occur per ten xylose 

units (Timell 1967; Fengel and Wegener 1989; Eriksson et al. 1990; Puls 1992; 

Fig.4.2. Diagram of enzymatic xylan degradation. x xylose residue, Ac acetic acid residue, 

4-O-Me-GA 4-O-methylglucuronic acid residue 

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Saake et al. 2001). The enzymatic xylan degradation is shown as a diagram in 

Fig. 4.2. 

The xylan backbone is degraded (⇑) by the ectoenzyme endo-1,4-β-xylanase 

(“xylanase”, systematic name: 1,4-β-D-xylan xylanohydrolase, EC 3.2.1.8) 

within the xylose chain (endohydrolysis) to xylo-oligomers, xylobiose and 

xylose (Eriksson 1990; Eriksson et al. 1990). Intracellular and/or membranebound 

xylan 1,4-β-xylosidase (1,4-β-D-xylan xylohydrolase, EC 3.2.1.37) removes 

successively D-xylose residues from the non-reducing termini (exoenzyme) 

of small oligosaccharides. The side-groups are split by accessory enzymes: 

Acetylesterase (acetic-ester acetylhydrolase, EC 3.1.1.6) removes the 

acetyl groups. Xylan α-1,2-glucuronidase (xylan α-D-1,2-(4-O-methyl)glucuronohydrolase, 

EC 3.2.1.131) hydrolyzes the α-D-1,2-(4-O-methyl)glucuronosyl 

links (Puls 1992). Arabinose side-groups in arabinoxylans are removed 

by α-arabinosidase. The structure of different xylans and their enzymatic 

degradation is described by Bastawde (1992). 

The mannans (glucomannans, galactomannans, galactoglucomannans) of 

the conifers, consisting mainly of the hexose mannose, are similarly hydrolyzed 

by mannan endo-1,4-β-mannosidase (“mannanase”, 1,4-β-D-mannan mannanohydrolase, 

EC 3.2.1.78), β-mannosidase (β-D-mannoside mannohydrolase, 

EC 3.2.1.25) and accessory enzymes like β-galactosidase, α-glucosidase, 

and esterase (Eriksson et al. 1990; Takahashi et al. 1992; Viikari et al. 1998). 

Therearehemicellulosehydrogenbondstocellulosefibrilsandalsocovalent 

links with lignin. 

Oxalic acid of brown-rot fungi might be involved first in the degradation of 

the side chains of the hemicellulose, thus providing entrance to arabinose and 

galactose, and then depolymerize the main hemicellulose chain (and amorphous 

cellulose) (Green et al. 1991a; Bech-Andersen 1987b). 

Hemicellulose degradation is common in wood fungi, but rarer in bacteria. 

The soft-rot fungus Paecilomyces variotii produced plenty of xylanase (Schmidt 

et al. 1979), and glucuronidase was excreted, e.g., by Agaricus bisporus (Puls 

et al. 1987; Bastawde 1992). In Oligoporus placenta, xylanases were located in 

the hyphal sheath (Green et al. 1991b). 

Basidiomycetes, which prefer conifers in nature, degraded a spruce wood 

mannan more intensively than a birch xylan, and in reverse hardwood fungi 

showed greater activity against xylan (Lewis 1976). During incipient brownrot 

decay, the hemicellulose components are degraded first. In southern pine, 

earlystrengthlossupto40%wasassociatedwithlossofarabinanandgalactan 

components, and subsequent strength loss greater than 40% was associated 

with the loss of the mannan and xylan components. Since the cellulose microfibril 

is surrounded by a hemicellulose envelope, significant loss of cellulose 

was only detected at greater than 75% modulus of rupture loss (Curling 

et al. 2002). 

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4.4 Cellulose Degradation 95 

4.4 

Cellulose Degradation 

In the biosphere about 2.7 × 10 11 tofcarbonareboundinlivingorganisms. 

According to Schwarz (2003) about 4 × 10 10 t cellulose are produced per year. 

Cellulose occurs in all land plants, is always fibrillarly constructed and consists 

of β-1,4-linked glucose anhydride units (glucopyranose). The substrate for cellulose 

biosynthesis is UDP-glucose which is polymerized by cellulose synthase 

(UDP-glucose: 1,4-β-D-glucan 4-β-D-glucosyltransferase, EC 2.4.1.12) to β-1,4 

glucan chains. Depending on the wood species, the degree of polymerization 

(DP) of native cellulose ranges from 10,000 to 15,000 glucose anhydride units. 

In “native cellulose”, hydrogen bridges exist between the OH groups of neighboring 

glucose units and neighboring cellulose molecules. Tidy (crystalline cellulose) 

regions and areas of lower order (amorphous, paracrystalline cellulose) 

alternate (Fengel and Wegener 1989; Fig. 4.3). In Boehmeria nivea, cellulose 

crystals of about 300 glucose residues are interrupted vertically to the longitudinal 

axis by an amorphous region of 4–5 glucose residues (Schwarz 2004). There 

are different models for the arrangement of the cellulose molecules in the fibrils. 

Bacterial cellulose degradation including the cellulosome was treated by 

Schwarz (2003). There is still some uncertainty as how cellulose is degraded 

by fungi. Differences occur between the various groups of fungi, brown, white, 

and soft-rot fungi. 

Fig.4.3. Diagram of enzymatic cellulose degradation 

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Early workers investigating brown-rot fungi assumed that only cellulolytic 

enzymes were responsible for cellulose degradation. Cellulolytic activity was 

initially described using terminology of C1-Cx (Reese et al. 1950): native (crystalline) 

cellulose is prepared by C1 cellulase for the degradation by Cx cellulase, 

as C1 cellulase loosens the crystalline areas by cleaving the hydrogen bridges 

for the following attack by Cx cellulase. 

The C1-Cx model was later refined to refer to the action of general classes 

of exoglucanases and endoglucanases, respectively. As methods were further 

refined more specific functionalities were defined and newly isolated enzymes 

were found in brown-rot fungi. Brown-rot fungi produce several endo-1,4β-glucanases 

and β-glucosidases, but typically lack exo-1,4-β-glucanase activity. 

However, cellobiohydrolase and cellobiose dehydrogenase [cellobiose: 

(acceptor) 1-oxidoreductase, EC 1.1.99.18] have been isolated from Coniophora 

puteana. Brown-rot fungal wood degradation was recently reviewed by Goodell 

(2003). White-rot and soft-rot fungi produce the full cellulolytic enzyme 

system of endo- and-exoglucanases, and β-glucosidase. 

The enzymes produced are thought to act in concert with each other as 

well as with non-enzymatic systems. Attack occurs at the amorphous cellulose 

regions (Cx action) by cellulase (“endoclucanase”, systematic name: 

1,4-β-D-glucan 4-glucanohydrolase, EC 3.2.1.4), which endohydrolyzes 1,4-β- 

D-glucosidic linkages in cellulose and other β-D-glucans. A combined action 

takes place by cellulose 1,4-β-cellobiosidase (1,4-β-D-glucan cellobiohydrolase, 

EC 3.2.1.91), which hydrolyzes 1,4-β-D-glucosidic linkages in cellulose 

and cellotetraose, releasing cellobiose from the non-reducing ends (exoenzyme), 

and by glucan 1,4-β-glucosidase (1,4-β-D-glucan glucohydrolase, EC 

3.2.1.74), which acts on 1,4-β-D-glucans and related oligosaccharides and exohydrolyzes 

successive glucose units from the ends. The final hydrolysis of 

oligosaccharides is mediated by β-glucosidase (“cellobiase”,β-D-glucoside glucohydrolase, 

EC 3.2.1.21), which acts on terminal, non-reducing β-D-glucose 

residues with release of β-D-glucose. Cellobiose may be also attacked by cellobiose 

dehydrogenase [cellobiose:(acceptor) 1-oxidoreductase, EC 1.1.99.18] 

oxidizing cellobiose to cellobionolactone under reduction of O2 to H2O2,and 

Fe 3+ to Fe 2+ (Kruså et al. 2005). 

In the mold Trichoderma viride (T. reesei), three endoglucanases, two exoglucanases, 

and several β-glucosidases were found (Eriksson et al. 1990). In 

Sporotrichum pulverulentum Novobr. (anamorph of Phanerochaete chrysosporium), 

five endoglucanases, one exoglucanase and two β-gucosidases, which 

together with oxidizing enzymes (laccase and cellobiose: chinon oxidoreductase) 

caused a combined degradation of cellulose and lignin. Uemura et al. 

(1992) isolated six exoglucanases. In P. chrysosporium, cellulases have been 

classified into eight different families among the glycoside hydrolases (Samejima 

and Igarashi 2004). In addition, the importance of the cellobiose dehydrogenase 

(CDH) was shown, as this enzyme could participate in the extracellular 

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4.4 Cellulose Degradation 97 

metabolism of cellobiose instead of β-glucosidase.TheroleofCDHforcellulose 

degradation was discussed (Hyde and Wood 1997; Kruså et al. 2005). It 

was hypothesized that CDH act as link between cellulolytic and ligninolytic 

pathways (Temp and Eggert 1999). 

Insoluble, native cellulose is attacked comparatively slowly by a system of 

cellulolytic enzymes. A limited introduction of substituents into the cellulose 

molecule reduces the number of hydrogen bonds of cellulose chains in proportion 

to the degree of substitution (DS) and the pattern of occurrence along 

the cellulose chain. Depending on these features and the nature of the substituent, 

water solubility of cellulose derivatives may be obtained at DS values 

between 0.4 and 0.7, and cellulose loses its ordered structure and becomes 

enzymatically accessible. Cellulose acetates up to a DS of 1.4 were deacetylated 

by various enzyme preparations (Altaner et al. 2001). 

For in vitro-degradation tests, carboxymethylcelluloses (CMC) are often 

used as soluble cellulose substrate (e.g., Schmidt and Liese 1980). The molecular 

structure of CMCs was characterized e.g., by Saake et al. (2000). 

Pure crystalline cellulose substrates, like cotton or Avicel, are degraded by 

white and soft-rot fungi. Most brown-rot fungi hardly show enzyme activity 

against crystalline celluloses and attack only pre-treated cellulose derivatives 

(Highley 1988; Enoki et al. 1988), because brown-rot fungi do not possess the 

synergistic endo/exo glucanase system, but have only endoglucanases. Within 

the woody cell wall, brown-rot fungi, however, depolymerize cellulose rapidly. 

Thus, the presence of lignin, lignin breakdown products, hemicelluloses, or 

simple sugars was postulated. 

Due to the limitation of enzyme accessibility to the woody cell wall by its 

pore sizes, the conceptions on cellulose degradation within wood by brownrot 

fungi focused both on non-enzymatic procedures and enzymatic mechanisms 

(e.g., Eriksson et al. 1990; Highley and Illman 1991; Micales 1992; 

Ritschkoff et al. 1992; Goodell 2003). Bailey et al. (1968) postulated as preceeding 

non-enzymatic agent a “precellulolytic phase”. Koenigs (1974) and 

others showed that cellulose was oxidatively degraded by Fenton reagents 

[Fe(II) + H2O2 → Fe(III) + OH − +OH 0 ]. Since ferrous iron is required in Fenton 

reactions, which is, however, absent in oxygenated wood decay processes, 

asearchforamechanismtoreduceironwasmade.H2O2 can also react with 

copper ions and some chromium, vanadium and nickel species to generate 

OH 0 (Halliwell 2003). 

Numerous investigations stress the participation of oxalic acid (e.g., Green 

et al. 1991a, 1993; Micales 1992), as the acid reduces Fe 3+ to Fe 2+ , which forms 

from H2O2 the reactive hydoxylradical, which then depolymerizes the cellulose. 

In several brown-rot fungi, like Coniophora puteana, Serpula lacrymans 

and Oligoporus placenta, extracellular H2O2 was proven (Ritschkoff et al. 1990, 

1992; Ritschkoff and Viikari 1991; Backa et al. 1992; Tanaka et al. 1992). Serpula 

lacrymans dissolved by means of oxalic acid iron from stonewool, which 

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promoted fungal growth and wood decay together with H2O2 (Paajanen and 

Ritschkoff 1992). In wood samples impregnated with chrome copper arsenic, in 

contact with rusting iron, probably iron ions diffusing into the wood increased 

wood decay (Morris 1992). Iron-sulfate reducing soil bacteria increased the 

iron content in wood samples as well as the mycelial growth rate of Gloeophyllum 

trabeum and Oligoporus placenta (Ruddick and Kundzewicz 1991). In 

contrast, inorganic chelating agents and iron-binding siderophores decreased 

growth and wood decay by brown-rot fungi (Viikari and Ritschkoff 1992). The 

potential function of oxalate as reducing agent of Fe 3+ is, however, limited to 

theinsideofwoodysubstratesbecausethistypeofreactionshalloccuronly 

in the absence of light (Goodell 2003). The role of oxalate in brown-rot mechanisms 

may rather lie in a slow action on the hemicellulose matrix to help to 

open up the wood structure. 

Since the early work on Fenton systems for hydroxyl radical production, several 

hypotheses have been developed explaining the function of low molecular 

weight metabolites, metals, and radicals, which initiate cell wall degradation 

by brown-rot fungi. 

A compound, termed “glycopeptide”, isolated from Fomitopsis palustris, 

with a molecular weight of 7.2 to 12 kDa reduced O2 to OH 0 and catalyzed 

redox reaction between NADH as electron donor and O2 to produce H2O2 

andtoreduceH2O2 to OH 0 . The glycosylated peptide reduced Fe(III) to Fe(II) 

(Enoki et al. 2003). The glycopeptide may either diffuse as a deglycosylated 

“effector” form of lower molecular weight in the wood matrix or the shape of 

the glycopeptide is elongated allowing cell wall penetration or the glycopeptide 

generates longer-lived radicals such as superoxide which penetrate the wall 

microvoid spaces (Goodell 2003). 

A cellobiose dehydrogenase enzyme system was proposed to occur in Coniophora 

puteana and to bind and reduce iron in the presence of oxalate, which 

the fungus employs to generate and maintain the low pH environment at least 

in the vicinity of the hypha, which is required to avoid autoxidation of the 

reduced valence state of iron (Hyde and Wood 1997; Goodell 2003). 

“Low molecular weight fungal chelators” from Gloeophyllum trabeum (“Gt 

chelator”) mediated the production of hydroxyl radicals within the wood cell 

wall, immunolocalized in the S2 layer, and were termed as “chelator-mediated 

Fenton system” (CMFS). In CMFS, iron is reduced and then repeatedly “rereduced”, 

exceeding a 1:1 ratio for reduction of iron by catechol. Gt chelator 

in CMFS reactions reduced the cellulose crystallinity of wood and the molecular 

weight of Avicel crystalline cellulose (Goodell and Jellison 1998; Goodell 

2003). 

Shimokawa et al. (2004) provided evidence that Serpula lacrymans employs 

a Fenton reaction mediated by a quinone-type chelator, and preferentially degrades 

amorphous regions of cellulose in the non-enzymatic cellulose degradation. 

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4.5 Lignin Degradation 99 

The accessibility of the cellulose for cellulases can be improved by several 

pretreatments of wood (Chap. 9): for example, soaking increases the pore areas, 

and chemical pretreatments decrease the lignin content. 

4.5 

Lignin Degradation 

Next to cellulose lignin is the most abundant polymeric organic substance 

in plants. Of about 10 11 t of annual production of terrestrial biomass, about 

2×10 10 t are lignin (Jennings and Lysek 1999). Lignin is contrary to linear 

polysaccharides, like cellulose, a complex, stereoirregular, three-dimensional 

macromolecule (see Fig. 4.4, Nimz 1974; Higuchi 2002) in the range of 100 kDa 

(Abreu et al. 1999) and is highly hydrophobic reducing the hygroscopicity of 

wood. Lignin functions as a binding and encrusting material in the cell wall 

distributed with hemicelluloses in the spaces of inter-cellulose microfibrils in 

the cell wall. It acts as a cementing component to connect cells and harden 

Fig.4.4. Structural scheme of beech lignin (modified from Nimz 1974) 

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the cell walls of xylem tissues, which helps a smooth transportation of water 

through vessels and tracheids from roots to branches (Higuchi 2002). The incorporation 

of lignin into the cell wall gives trees with heights of 100 m the 

chance to remain upright. Lignin gives resistance against disease and wood 

decay by microorganisms. Lignin content amounts in softwoods to 26–39% 

(average 28%) (compression wood: 35–40%), in hardwoods of the temperatezoneto18–32%(average22%)(tensionwood:15–20%)andintropical 

hardwoods to 23–39% (Fengel and Wegener 1989). 

The monolignols (p-hydroxycinnamyl alcohols), p-coumaryl, coniferyl, and 

sinapyl alcohol, are the primary precursors and building units of all lignins 

(Fengel and Wegener 1989; Fig. 4.5). The biosynthetic pathway of monolignols 

starts from glucose via shikimic acid over phenylalanine and tyrosine, respectively, 

to p-coumaric acid which yields via intermediates p-coumaryl alcohol. 

p-coumaric acid is converted via caffeic acid and ferulic acid to coniferyl alcohol. 

Ferulic acid is transformed via 5-hydroxyferulic acid and sinapic acid to 

sinapyl alcohol (Higuchi 2002). 

For lignin polymerization (Li and Eriksson 2005), the monolignols are initially 

dehydrogenated by peroxidases and/or laccases to phenoxy radicals. The 

radicals then couple non-enzymatically to quinone methides as reactive intermediates. 

According to one proposal, dimeric quinone methides are converted 

into dilignols by water addition, or by intra-molecular nucleophilic attack by 

primary alcohol groups or quinone groups. Dilignols can also undergo enzymatic 

dehydrogenation to form the corresponding radicals, which in turn 

couple with phenoxy radicals to produce trilignols, etc. In a second mechanism, 

enzymatic dehydrogenation is restricted to monolignols. The polymerization 

evolves by successive non-radical addition of phenols to the quinone methides. 

In a third mechanism, the lignin polymer evolves from the polymerization of 

quinone methides. 

Most softwood lignins are as guaiacyl lignins (G-lignins) polymers which 

are predominantly made of coniferyl alcohol (spruce: C : S : p-C = 94 : 1 : 5). 

Hardwood lignins are guaiacyl-syringyl lignins (GS lignins) and consist predominantlyofCandS(beech:C:S:p-C 

= 56 : 40 : 4). Guaiacyl-syringylp-hydroxyphenyl 

lignins occur in grasses. Lignin quantity and composition 

Fig.4.5. Lignin building units. A pcoumaryl 

alcohol. B Coniferyl alcohol. 

C Sinapyl alcohol 

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vary also as with the tree age (Wadenbäck et al. 2004), between root and stem 

wood, heartwood and sapwood, xylem and bark, earlywood and latewood, and 

in different wood cells and cell wall layers. In the lignin molecule, the basic 

modules are linked with a variety of chemical bonds, ether and carbon-carbon 

linkages. Most bonds are covalent, of considerable variety and are equally in 

all three dimensions. The β-O-4 linkage (fat in Fig. 4.4) is the most frequent 

interunit linkage with about 50% (spruce) to 65% (beech) (Fengel and Wegener 

1989; Abreu et al. 1999; Higuchi 2002). 

Lignin forms an amorphic complex with hemicelluloses to encapsulate cellulose. 

As lignin represents a substance, which is hardly open to attack for most 

microorganisms, it protects within the woody cell wall the enzymatically more 

easily accessible carbohydrates against microbial degradation (Chap. 9, Table 

4.3). There are different model conceptions with regard to the arrangement 

of the three components (see Fig. 4.1). 

Causes for the resistance of lignin against microbial enzymes are: Aromatic 

rings are generally more difficulty degradable. The variety of the linkages 

between the building units and the hydrophobic nature require a breakdown 

system that is non-specific and, for the most part, nonhydrolytic as well as 

extracellular (Jennings and Lysek 1999; Reading et al. 2003). 

Overviews on lignin degradation are e.g., by Umezawa (1988), Higuchi 

(1990), Schoemaker et al. (1991), Jeffries (1994), Cullen and Kersten (1996), 

Yoshida (1997) and Koshijima and Watanabe (2003). 

An effective degradation of natural lignin (lignin within the woody cell wall) 

with respiration of the C-atoms from that aromatic ring exclusively occurs in 

white-rot fungi (Chap. 7.2). The residual lignin in wood degraded by brown-rot 

fungi is dealkylated, demethoxylated and demethylated, with some oxidation 

of the alkyl side chain. The aromatic ring is not attacked (Goodell 2003). Softrot 

fungi mainly cleave the methoxyl groups from the aromatic rings. Some 

bacteria demethylate or cleave within the alcoholic side chain, particularly 

in synthetic lignins with small molecular weights (dehydrogenation polymer, 

DHP) and in lignin model compounds (Fengel and Wegener 1989). For the 

“tunneling bacteria”, lignin degradation was postulated within the woody cell 

wall (Chap. 5.2). 

Many white-rot fungi produce extracellular phenol oxidases, which results 

in positive oxidase tests on nutrient agar with tannic and gallic acid. Only 

40% of the white-rot fungi studied produced the combination of lignin peroxidase 

and manganese peroxidase, whereas the combination of manganese 

peroxidase and laccase was more common. In an extreme case, Pycnoporus 

cinnabarinus produced only laccase, lacking both lignin and manganese peroxidase 

(Eggert et al. 1996; Li 2003). The test by Bavendamm (1928) is used 

since that time for the rapid differentiation of white and brown-rot fungi in the 

laboratory and is in identification keys for wood fungi among the first distinguishing 

characters (Stalpers 1978). Malt agar is supplemented with a lignin 

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model compound (Davidson et al. 1938; Lyr 1958; Käärik 1965; Rösch and 

Liese 1970; Tamai and Miura 1991) and inoculated with the unknown fungus. 

If the fungus excretes the phenol oxidase laccase (benzenediol:oxygen oxidoreductase, 

EC 1.10.3.2), tannic acid, etc. are oxidized (brownish discoloration), 

and it usually concerns a white-rot fungus. By definition, laccases catalyze the 

oxidation of p-diphenols and the concurrent reduction of dioxygen to water, 

although the actual substrate specifities of laccases are often broad (Eggert et al. 

1998). Most brown-rot fungi do not oxidize tannic acid, as they usually possess 

only intracellular tyrosinase (1,2-benzenediol:oxygen oxidoreductase EC 

1.10.3.1). However, misinterpretation may occur because tyrosinase can be set 

free through injuring the mycelium, e.g., by the inoculation procedure, which 

feigns then a white-rot fungus (Rösch 1972). Furthermore, also some intensive 

lignin decomposer, e.g., Phanerochaete chrysosporium,maycausenegativeor 

only weak Bavendamm reaction (Eriksson et al. 1990). Laccase is also present 

in several Deuteromycetes and Ascomycetes (Butin and Kowalski 1992; also 

Luterek et al. 1998). Phenol oxidase (laccase) and one-electron oxidation activity 

was shown for the soft-rot Deuteromycetes Cladorrhinum sp., Graphium sp., 

Scopulariopsis sp., and Sphaeropsis sp. (Tanaka et al. 2000). Niku-Paavola et al. 

(1990) used 2, 2 ′ -azino-di(3-ethylbenzothiazoline)-6-sulfonic acid as enzyme 

substrate, which is oxidized by laccase, while tyrosinase does not. 

Due to the effect of the laccase in vitro, in former times, lignin degradation 

was assumed to occur exclusively by phenol oxidases. The main significance 

of the laccase is, however, seen in the polymerization of phenols. Lignin polymerization 

by laccase occurs through the formation of phenoxy radicals by 

abstraction of hydrogen followed by a series of radical polymerization reactions. 

Thus, laccase has also been used to obtain wood composites like particle 

and MDF boards that were bound mainly or even solely by lignin when polymerized 

in situ by this enzyme (Kharazipour and Hüttermann 1998). On the 

other hand, laccases are involved in lignin degradation by fungi, which was 

confirmed by “synergistic cellulose lignin degradation models” (Ander and 

Eriksson 1976). In connection with the cell wall degradation, the significance 

of the phenol oxidases might be rather an adjusting function for the carbohydrate 

degrading enzymes (Eriksson et al. 1990). In fungi, laccases are also 

involved in pigmentation, fruit body formation, sporulation, and pathogenesis 

(Rättö et al. 2004). 

The isolation of the first ligninolytic enzyme was simultaneously obtained in 

two groups (Glenn et al. 1983; Tien and Kirk 1983) from culture filtrates of the 

white-rot fungus Phanerochaete chrysosporium. This fungus was well known 

as an intensive lignin decomposer, since it degraded 14 ClabeledligninstoCO2 

as well as dehydropolymers and model compounds (Kirk 1988). The enzyme, 

diarylpropane peroxidase (lignin peroxidase, LiP, “ligninase I”, diarylpropane: 

oxygen, hydrogen-peroxide oxidoreductase, EC 1.11.1.14) is a glucoprotein 

with a molecular weight of 42 kDa (also Srebotnik et al. 1988b), contains hem 

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(porphyrin with iron as central atom), needs extracellular H2O2, cleavesC-C 

bonds in a number of model compounds, and oxidizes benzyl alcohols to 

aldehydes or ketones. 

The key reaction of the LiP is a one-electron-oxidation of various nonphenolic 

compounds to generate instable aryl radical cations, as the enzyme 

delivers two electrons to hydrogen peroxide, which the enzyme then takes back 

from each one-phenyl propenoid unit (Kirk 1985; Higuchi 1990). Phenolic 

and non-phenolic lignin substructures are attacked, but not the intact lignin 

molecule. 

The radicals undergo subsequent non-enzymatic reactions to yield a variety 

of final products. The radical cations themselves act as oxidants. Thus, LiP 

initiates by means of different non-enzymatic reactions the cleavage of Cα-Cβ 

bond in the side chain, β-O-4 bond between side chain and next ring, as well 

the aromatic ring (Eriksson et al. 1990; Schoemaker et al. 1991; Fig. 4.6). Also, 

veratryl alcohol, which is produced independently of lignin degradation, can 

be oxidized by LiP to the radical cation, which itself can oxidize lignin (Jennings 

and Lysek 1999). 

The ligninolytic system of Phanerochaete chrysosporium is not induced by 

lignin but appears constitutively as cultures enter the secondary metabolism, 

that is, when primary growth ceases because of depletion of nutrients. Secondary 

metabolism was triggered by nitrogen, carbon, or sulphur limitation 

(review by Highley and Dashek 1998). Lignin cannot be used as only C source, 

but in cometabolism with cellulose or hemicellulose. A high O2 concentration 

(100% more suitable than 21%) was favorable (Kirk 1988). The enzyme was 

excreted by old, autolytic hyphae, but not by arthrospores and chlamydospores 

(Lackner et al. 1991). 

LiP has been found in several white-rot fungi, e.g., Trametes versicolor, 

Phlebia radiata (Dodson et al. 1987) and Bjerkandera adusta (Muheim et al. 

1990). There are numerous isomers of LiP with molecular weights of 40 to 

47 kDa, which differ in the carbohydrate portion of the protein (Evans 1991). 

The enzyme activity of LiP preparations is determined via Cα oxidation of 

Fig.4.6. Scheme of reactions initiated 

by lignin peroxidase. Cleavage of Cα-Cβ 

bond in the side chain (1), β-O-4 bond 

between side chain and next ring (2), 

and aromatic ring (3) 

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veratryl alcohol (with presence of H2O2) to the aldehyde, whose amount is 

measured at 310 nm (Faison and Kirk 1985; Schoemaker et al. 1991). 

The second enzyme involved in lignin degradation is manganese peroxidase 

(MnP) [Mn(II):hydrogen-peroxide oxidoreductase, EC 1.11.1.13], which needs 

free phenolic groups at the aromatic ring and does not oxidize veratryl alcohol. 

The hemoprotein enzyme was first detected in P. chrysosporium (Glenn and 

Gold 1985) and occurs e.g., in Armillaria species, Lentinula edodes, Pleurotus 

ostreatus and T. versicolor. It oxidizes in the presence of hydrogen peroxide 

Mn(II) to Mn(III), a strong oxidant, which oxidizes phenolic structures by 

single-electron-oxidation (Perez and Jeffries 1992; Kofugita et al. 1992; Robene 

Soustrade et al. 1992; Chatani et al. 1998; Kamitsuji et al. 1999). MnP polymerizes 

more extensively and depolymerizes less than lignin peroxidase (Tanaka 

et al. 1999). Treatment of water-insoluble 14 C-labeled milled wheat straw and 

milled straw lignin with MnP preparations from the white-rot fungus Nematoloma 

frowardii resulted in the direct release of 14 CO2 and in the formation 

of soluble 14 C-lignin fragments (Hofrichter et al. 1999). MnP also degraded 

polyethylene (Iiyoshi et al. 1998). 

For the degradation of native lignin, a fungus must have enzymes, which attack 

both phenolic and non-phenolic lignin components (Martinez-Inigo and 

Kurek 1997). The lignin peroxidase is most likely responsible for the degradation 

of the non-phenolic components and the laccase as well manganese 

peroxidases for the oxidation of the phenolic parts (Evans 1991). All together, 

there is a variety of oxidative enzymes that may be utilized by white-rot fungi for 

lignin degradation (Highley and Dashek 1998). Various enzymes, low molecular 

weight agents, free-radical reactions, and metals have been proposed to 

participate in lignin degradation (Messner et al. 2003; Reading et al. 2003): 

Lignin peroxidase (LiP), manganese peroxidase (MnP), cellobiose dehydrogenase 

(CDH), laccases, oxalate, hydrogen peroxide, small molecule mediators, 

methyl transferases, and the plasma membrane redox potential are involved 

in the degradation systems. There is, however, still some uncertainty on their 

accurate participation in lignin degradation. 

Progress has been made concerning the molecular genetics of lignin and 

cellulose biodegradation by white-rot fungi, primarily with Phanerochaete 

chrysosporium, but also with Bjerkandera adusta, Phlebia radiata, and Trametes 

versicolor (reviews by Highley and Dashek 1998 and Li 2003). Genes encoding 

Lip and MnP have been cloned and sequenced (e.g., Irie et al. 2000). The 

total genome sequence of P. chrysosporium has been disclosed by the DoE Joint 

Genome Institute in the USA, which has facilitated cDNA cloning of various cellulase 

genes from P. chrysosporium and successive production of recombinant 

proteins from them (Samejima and Igarashi 2004). The X-ray crystal structures 

of both LiP and MnP have also been elucidated. By means of recombinant 

DNA techniques, laccase catalysis has been studied, and the crystal structure 

of a T2-copper deleted laccase has been reported. In Pycnoporus cinnabarinus, 

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genes encoding two laccase isozymes have been cloned and sequenced (also 

Eggert et al. 1998). Glyoxal oxidase as a source of extracellular H2O2 was found 

to be encoded by a single gene. 

Occurrence and distribution of lignin peroxidase inside hyphae and whiterotten 

wood were examined by immuno gold labeling (Srebotnik et al. 1988a; 

Blanchette et al. 1989; Daniel et al. 1989, 1990; Blanchette and Abad 1992; Kim 

et al. 1993). The enzyme is particularly found in the hypha and the extracellular 

sheath, and less so in the wood cell wall and then near the hypha. In the cell 

wall, it is only considerably present in late degradation stages. It was concluded 

from this distribution that the lignin peroxidase attacks rather lignin fragments 

that had been set free from the cell wall, than that it binds at the polymeric 

lignin inside the intact wall. The primary degradation would have then taken 

place by low-molecular compounds like the cation radical of veratrylalcohol, 

which diffuses into the wall, produces there lignin fragments, which are then 

degraded by ligninase (Evans 1991). It may also assumed that the limited cell 

wall degradation starting from the cell lumen in close neighborhood of a hypha 

occurs directly by the enzyme towards closely neighboring lignin. This would 

agreewiththeearlyresultsoftheerosion-likecellwalldegradationbywhite-rot 

fungi (Schmid and Liese 1964; Liese 1970; Fig. 7.2b). 

There are many ways that a white-rot fungus could generate hydrogen peroxide 

required for LiP and MnP. Extracellular H2O2-producing enzymes are arylalcohol 

oxidase (EC 1.1.37), glyoxal oxidase, pyranose 2-oxidase (EC 1.1.3.10), 

and cellobiose dehydrogenase. Intracellular enzymes include glucose 1-oxidase 

(EC 1.1.3.4) (Leonowicz et al. 1999), pyranose 2-oxidase, and methanol oxidase 

(e.g., Daniel et al. 1994; Hyde and Wood 1997; Urzúa et al. 1998). OH 0 

may be also formed via hydroquinone redox cycling involving semiquinones 

produced by peroxidase or laccase which reduce both Fe(III) and O2 to provide 

the components for Fenton-type hydroxyl radical formation. It is not exactly 

known which enzyme plays the primary role in supplying H2O2 (Li 2003). 

From the only slow microbial decomposition of lignin results its significance 

for the formation of humic substances (e.g., Haider 1988; Schlegel 1992) and 

also for the lastingness of archaeological woods (Chap. 5.2). The suitability of 

lignins as fertilizer and for soil improvement was described by Faix (1992). 

Mikulášová and Košíkowá (2002) indicated a potential application of lignin 

biopolymers as antimutagenic agents in chemoprevention. 

There are some general prerequisites for the action of the degradative systems. 

As lignin is a highly oxidized polymer, reductive as well as oxidative 

reactions are required to effectively degrade it, both of which must occur aerobically. 

These reactions must be balanced or controlled to prevent redox cycling 

and free-radical-based polymerization of the degradation products. The oxidizing 

and reducing equivalents must be unique and continuously produced 

since extracellular regeneration would be improbable. Common biological 

compounds for reducing or oxidizing equivalents, such as NADH, which would 

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be difficult to regenerate once released extracellularly are precluded. In vitro, 

reduction of manganese dioxide was demonstrated for a ferrireductase system 

that includes NADPH-dependent ferrireductase and the iron-binding compound 

from Phanerochaete sordida (Hirai et al. 2003). Extracellularly formed 

free-radical species are able to diffuse away from their origin and mediate reactions 

with the insoluble lignin. The small, diffusible radicals and low-molecular 

agents achieve a greater area of reactivity than could be obtained by reactions 

catalyzed by enzymes or the fungi directly. The distance of the action from the 

hyphae also prevents self-inflicted damage to the fungus (Reading et al. 2003). 

The following description of systems to generate low molecular agents is 

according to Messner et al. (2003). 

In the “manganese peroxidase/Mn(II)/oxalate system”, there are two oneelectron 

reducing steps by Mn(II). The Mn(III) formed is chelated and released 

from the enzyme by the fungal metabolite oxalate. The relatively stable Mn(III) 

oxalate oxidizes phenolic lignin compounds and has been proposed to diffuse 

in the wood cell wall. 

In the “manganese peroxidase/Mn(II)/oxalate/cellobiose dehydrogenase 

system”, CDH is oxidized by O2 and metal ions such as Fe(III) and Cu(II) 

yielding H2O2, and Fe(II) or Cu(I) which react with H2O2 to generate hydroxy 

radicals which in turn demethoxylate and hydroxylate non-phenolic lignin. 

The phenolic lignin formed is then attacked by MnP-generated Mn(III). 

In the “manganese peroxidase/Mn(II)/oxalate/lipids system”, lipids extend 

the oxidative potential of MnP. Mn(III) promotes peroxidation of unsaturated 

fatty acids resulting in the formation of peroxyl radicals which are diffusible, 

potentially ligninolytic agents. Mn(III) also abstracts hydrogen from fatty 

acids to form acyl radicals. The system depolymerized both phenolic and 

non-phenolic lignin (Katayama et al. 2000). 

In the “lignin peroxidase/veratryl alcohol system”, the veratryl alcohol radical, 

generated during turnover of LiP, was proposed to act as a charge transfer 

system in wood. However, its short lifetime may prevent a diffusion into deeper 

cell wall areas. 

In the “laccase/mediator system”, laccases are combined with low molecular 

weight charge transfer agents, so-called mediators. The system is used to bleach 

pulp and depolymerized non-phenolic guaiacyl lignin. 

In the “glycopeptide system” (Enoki et al. 2003), low-molecular weight 

glycosylated peptides produce hydroxy radicals which modify lignin, resulting 

in new phenolic, benzyl radical, and cation radical substructures which are 

then attacked by LiP, MnP or laccase. The system also depolymerizes the wood 

carbohydrates (see Chap. 4.4). 

In the “coordinated Cu/peroxide system” (Messner et al. 2003), either hydrogen 

peroxide or organic peroxides, is the agent involved at least in the 

initial lignin degradation. Cu(II) is reduced to Cu(I) by either H2O2 or reducinggroupsinwood.Cu(I)formswithH2O2 

a reactive one-electron oxidant 

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that oxidizes phenolic and non-phenolic lignin. Cu(I) is reoxidized by lipid 

hydroperoxide. 

For the preferential white-rot type without the intense damage of cellulose, 

Teranishi et al. (2003) showed that Ceriporiopsis subvermispora produced 

ceriporic acid, which strongly inhibited the Fenton reaction to suppress the 

formation of OH 0 . 

In summary, meanwhile many details on the degradation of the various 

components of the woody cell wall are known. It may be considered, however, 

that in view of lignin and cellulose degradation, many results derive from only 

one fungal species, Phanerochaete chrysosporium (anamorph: Sporotrichum 

pulverulentum), and that this fungus has nearly no relevance for wood, neither 

for trees nor for construction timber, only for chip piles. 

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5 

5.1 

Viruses 

Damages by Viruses and Bacteria 

Viruses are small particles (10–2,000 nm in size) that infect Eukaryotes as obligate 

intracellular parasites. They reproduce by invading and taking over other 

cells as they lack own metabolism and the machinery for self-reproduction 

(Nienhaus 1985a). Typically, they carry either DNA or RNA surrounded by 

a coat of protein or protein and lipid. Plant viruses penetrate the shoot, leaf 

tissue and root via wounds or they are transferred by vectors [aphids, cicadas, 

nematodes, among fungi: Sphaerotheca lanestris (Erysiphales) on oak]. 

Partial bleaching of chlorophyll results in angular, circular (mosaic) or diffuse 

chloroses. Leaf damage, dwarfing or growth inhibition, distorted growth, 

and necrotic areas or lesions can occur, that is, virus infection can reduce 

the tree growth. Over 1,000 virus diseases of plants are described for Europe. 

Virus diseases in forest trees have been summarized e.g., by Nienhaus and 

Castello (1989) and Cooper and Edwards (1996). Viruses occur in several gymnosperms 

(Chamaecyparis, Cupressus, Larix, Picea and Pinus), angiosperms 

(Acer, Aesculus, Betula, Carpinus, Cormus, Corylus, Fagus, Fraxinus, Juglans, 

Populus, Prunus, Quercus, Rhamnus, Robinia, Salix, Sambucus, Sorbus and 

Ulmus) (Nienhaus 1989; Brandte et al. 2002), in bamboos and palms. Twig 

increase in horse chestnut (Butin 1995), and witches’-broom on beech and 

robinia are probably likewise due to the participation of viruses. Viruses have 

been detected several times in forest dieback sites (Parameswaran and Liese 

1988; Winter and Nienhaus 1989; Gasch et al. 1991). 

Viroids are infectious agents that consist of a single-stranded RNA. Viroids 

are smaller than viruses, lack a protein cover and are the smallest causal 

agents of plant diseases, like discolorations, chloroses and distorted growth, 

e.g., in coconut, cucumber, hop, potato and tomato (Schlegel 1992; Butin 1995; 

Nienhaus and Kiewnick 1998). About 33 species of viroids have been identified. 

5.2 

Bacteria 

“The Prokaryotes” (Dworkin et al. 2005) is a comprehensive reference on 

bacterial biology. 

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110 5 Damages by Viruses and Bacteria 

The term bacteria had been used for all Prokaryotes or for a major group 

of them. Based on the 16S rDNA sequence the Prokaryotes were divided into 

the kingdoms Eubacteria and Archaebacteria (Woese and Fox 1977). Later 

three domains, Bacteria, Archaea and Eucarya were renamed (Fig. 5.1) and 

confirmed by sequencing (Gray 1996). 

Archaea differ from other Prokaryotes in their membrane composition, 

flagella development and the similarity of their transcription and translation 

to that one of Eukarya. Many Archaea are extremophiles and live in geysers 

and black smokers at 80–110 ◦ C(Pyrodictium spp.), or in acid (about pH 0), 

alkaline, or saline (till 30% salt content) water like Halobacterium species. 

Methanogenic Archaea inhabit the digestive tracts of ruminants, humans, and 

termites, or soil, marshland and sewage etc. In trees, methanogenic Archaea 

are involved in the development of the alkaline wetwood (see below). 

Bacteria cover a major group of Prokaryotes and are ubiquitous in soil, 

water, as symbionts, or pathogens. They lack cell nucleus, cytoskeleton, mitochondria, 

and chloroplasts. The genetic information is located on a circular 

DNA strand, which is not covered by a nuclear membrane. Many bacteria contain 

plasmids with extrachromosomal DNA. Ribosomes are made of the 70S 

type (Eukaryotes: 80S). Reproduction is asexual by cell division. Exchange of 

genetic material can occur by transformation, transduction, and conjugation. 

About 10,000 species are identified, characterized, and deposited in culture 

collections, which might, however, represent only 10% of the actually existing 

species. Many bacteria are rod-shaped, sphere-shaped (cocci), helix-shaped 

(spirillum), or comma-shaped (vibrios). Common bacteria are minute, measuring 

0.4–5µm in size. They occur single, or double, or in chains or clusters. 

Fig.5.1. Phylogenetic tree of life based on rDNA data (from www:en.wikipedia.org) 

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5.2 Bacteria 111 

Gram staining divides Gram-positive and negative species according to their 

wall structure and the occurrence of pepitoglycan or lipids. With regard to 

oxygen, aerobes grow only in the presence of oxygen and anaerobes in its 

absence. The latter behavior may be either facultative or obligate. Many bacteria 

are motile, using either flagella, axial filaments, gliding, or changes in 

buoyancy. In some genera (Bacillus, Clostridium), the mother cell develops 

an endospore, which is rather resistant against heat, radiation and chemicals 

(Schlegel 1992). 

Actinobacteria are bacteria, which often live in the soil. They play important 

roles in plant decomposition, humus formation, and degradation (Filip 

et al. 1998) and are found on timber in soil contact. Some form branching filaments, 

which resemble the fungal mycelium (in former times: Actinomycetes), 

whereby the cell diameter of about 1µm is however smaller than that of most 

fungal hyphae. Some actinobacteria (e.g., Streptomyces) developaplentyof 

air-borne spores. 

There are various interactions between bacteria and plants, like increase of 

soil fertility by nutrient release, nitrogen fixation (Azotobacter), root symbioses 

(Rhizobium), decomposition and humification, and parasitism as causal agents 

of diseases. 

The pathogenic bacteria of woody plants belong to the genera Agrobacterium, 

Erwinia, Pseudomonas, andXanthomonas (Nienhaus and Kiewnick 

1998). Bacteria cause the fire blight [Erwinia amylovora (Burrill) Winslow 

et al.] of many species of the rose family (Tattar 1978), canker of poplar [Xanthomonas 

populi ssp. populi (Ridé) Ridé & Ridé], willow (X. populi ssp. salicis 

de Kam) and ash (Pseudomonas syringae ssp. savastanoi pv. fraxini Janse) 

(Butin 1995). Agrobacterium species infect the roots of a wide range of dicotyledonous 

plants and some gymnosperms causing crown gall and hairy 

root diseases. 

Since the late 1970s, Agrobacterium-mediated gene transfer is an important 

tool in genetic transformation of forest trees. During the disease process, 

a DNA segment of the bacterium (T-DNA) is integrated into the host plant 

genome. The T-DNA originates from a 200-kb plasmid (Ti plasmid) and foreign 

genes can be inserted into this DNA for transfer into the plant (Palli 

and Retnakaran 1998; Häggman and Aronen 1996), e.g., for gene-manipulated 

poplars and white spruce (Séguin et al. 1998). Kajita et al. (2004) transferred the 

gene for the enzyme feruloyl-CoA hydratase/lyase, which is involved in lignin 

(hydroxycinnamates) metabolism, from a bacterium into aspen by Agrobacterium 

tumefaciens (E.F. Smith & Townsend) Conn in view of producing trees 

with novel characteristics. 

Rickettsia and Rickettsia-like organisms (RLOs) (Proteobacteria) (100– 

800 nm) are obligate intracellular, Gram-negative bacteria with reduced metabolic 

activity. They cannot be cultured in nutrient medium. RLOs in plants are 

transferred by arthropodes, particularly cicadas, and multiply in the vector 

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and in the phloem or xylem. They cause e.g., leaf necrosis in oak and planes 

and distorted growth of larch (Nienhaus 1985b; Linn 1990; Butin 1995). 

Mycoplasmas (genus Spiroplasma) and phytoplasmas (in former times: 

MLOs; genus Phytoplasma) are the smallest (100–750 nm) independently 

growing bacteria. They are pleomorphic, sporeless, immovable, and filterable. 

Spiroplasma grows on nutrient medium, Phytoplasma does not. Plant 

pathogens are transferred by grafting, root grafts, vegetative propagation of 

infected material, Cuscuta species, and sucking insects, in which they multiply, 

into the phloem (Nienhaus and Kiewnick 1998). They cause a great 

number of yellow-type diseases, necroses, growth disturbances, or dying of 

rice, maize and sesame, vegetables, sugar cane, fruit trees, coconut palm, 

whitethorn, alder and ash, witches’-brooms on poplar, and sandal spike (Tattar 

1978; Nienhaus 1985a, 1985b; Linn 1990; Sinclair et al. 1990; Lindner 1991; 

Lederer and Seemüller 1991; Raychaudhuri and Mitra 1993; Raychaudhuri and 

Maramorosch 1996). 

Bacteria appear in trees and wood as both primary and secondary colonizers 

often in the context of succession together with fungi. They live on easily 

accessible nutrients and may prepare the substrate for fungi (Shigo 1967; 

Cosenza et al. 1970; Shigo and Hillis 1973; Shortle and Cowling 1978; Rayner 

and Boddy 1988). Soil bacteria may increase vitamin content (thiamine) of 

wood in ground contact, which promotes subsequent decay Basidiomycetes 

(Cartwright and Findlay 1958; Henningsson 1967). 

Bacteria penetrate into the sapwood of a tree via wounds. In hardwood 

vessels that are not closed by tyloses or other wound reactions, they might 

spread with the capillary water over larger distances. In softwood samples, 

however, only a few tracheids were passed due to the small free spaces within 

the pit membrane (Liese and Schmidt 1986). 

The wet heartwood (wetwood) of several tree species, particularly fir, hemlock, 

poplar, elm, also beech and oak, means any water-saturated and dead 

wood in living trees. Characteristics are the unpleasant smell of butyric acid 

and other acids, dark discolorations and gas escape from the heartwood if an 

increment borer has been used. The exact cause of wetwood formation, whether 

being due to bacteria or necrotic changes in the parenchyma cells, is not clarified. 

Wetwood develops in connection with mechanical wounds, branch breaking, 

decay, stem cracks, and insect attack. So-called acid wetwood, predominantly 

in conifers, contains several organic acids (butyric, acetic, propionic 

acid) produced by (facultative) anaerobe bacteria. Alkaline wetwood, mostly 

in hardwoods, develops with participation of obligate anaerobe methanogenic 

Archaea. These Prokaryotes attack the pits or cause their incrustation, give rise 

to discolorations, their metabolites may stress the tree, and the unpermeable 

wetwood tends to crack during drying (Carter 1945; Hartley et al. 1961; Wilcox 

and Oldham 1972; Bauch 1973; Knutson 1973; Bauch et al. 1975; Tiedemann 

et al. 1977; Ward and Pong 1980; Ward and Zeikus 1980; Schink et al. 1981; Mur- 

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doch and Campana 1983; Zimmermann 1983; Schink and Ward 1984; Kučera 

1990; Klein 1991; Walter 1993; Xu et al. 2001). 

Bacteria may be also associated with the development of false frost cracks in 

oak, ash, elm, poplar, and Silver fir. These radial shakes develop progressively 

from the stem interior, being initiated either from old cambial injuries or from 

pockets of fungal heart rot (Shigo 1972; Butin and Volger 1982). Occurrence, 

distribution, and enzyme activities of the bacteria isolated from pedunculate 

oak trees supported the assumption that bacteria may be involved in the 

weakening of the woody tissue in the area of the ray parenchyma cells so that 

mechanical factors like frost subsequently push the shake in the predamaged 

tissue (Schmidt et al. 2001). 

Several bacteria isolated from wet-stored stem wood were able to degrade 

pectin, hemicelluloses, and cellulose when these cell wall components had been 

supplied as isolated compounds (Schmidt and Dietrichs 1976). With regard to 

lignin, lignin derivatives or DHPs up to 1 kDa were attacked (Vicuña 1988). 

In view of bacterial wood degradation, bacteria attacked within partially 

lignified plant organs, like a shoot or a needle, only non-lignified tissue. The 

cell walls of the phloem cells of the vascular bundles were degraded, but those 

of the xylem part resisted. Inside woody tissue, bacteria preferentially feed 

soluble sugars, the content of parenchyma cells and attack non-lignified pit 

membranes (Liese 1970). In tension wood fibers, bacteria only consumed the 

cellulosic G-layer (Schmidt 1980). After a mild delignifying pretreatment of 

wood samples with sodium chlorite, however, bacteria caused mass loss up 

to 70% (Schmidt 1978), as the carbohydrates were now accessible. Figure 5.2 

Fig.5.2. Beech wood microtome sections with slightly reduced lignin content without (a) 

and after culture with Cellulomonas flavigena (b) 

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shows beech wood microtome sections whose lignin content have been reduced 

from naturally (21%) to about 19% and which subsequently had been used as 

the only carbon source for bacteria in liquid cultures. The microtome section in 

Fig. 5.2a represents the non-inoculated control. The section in Fig. 5.2b shows 

that only the highly lignified middle lamella primary wall region resisted to 

the bacterium Cellulomonas flavigena Kellerman and McBeth. However, the 

bacteria only consumed the carbohydrates of the pretreated wood. The lignin, 

which was dissociated from the pretreated woody cell wall by the bacteria, was 

not respired but was refound in the nutrient liquid, suggesting that lignin is 

a “ballast” to these bacteria that inhibits the dissimilation of the wood carbohydrates. 

The action of the chlorite pretreatment was assumed to result from 

the “opening” of the close association between carbohydrates and lignin in the 

woody cell wall so that the carbohydrates became accessible to the bacteria. 

Decay may have not been due to the reduction of the lignin content, because 

bacteria did not attack natural beech wood with 21% lignin content, but degraded 

pretreated Scots pine samples with a higher lignin content of about 

23% (Schmidt and Bauch 1980). 

Several bacteria were isolated from sawn Liriodendron tulipifera lumber already 

after 2 months of stacking (Mikluscak and Dawson-Andoh 2004a). After 

longer wood exposition under natural conditions, like in soil, or lakes and 

marine environment, the lignified cell wall was degraded by mixed populations 

and obviously the hurdle of the lignin barrier was cleared (Liese 1950; 

Liese and Karnop 1968; Schmidt et al. 1987; Fig. 5.3a). Dependent on the decay 

type within the wood cell wall, cavity, erosion, and tunneling bacteria 

were distinguished (Singh and Butcher 1991; Nilsson et al. 1992; Singh et al. 

1992; Daniel 2003). The two first types resemble the soft-rot types 1 and 2 

(Chap. 7.3). The tunneling bacteria are qualified by means of slime sheats to 

a gliding movement inside cell wall concavities created by themselves. The 

aggregates of the tunneling bacteria subcultured from the woody samples consisted 

of different bacterial species (Nilsson and Daniel 1992; Nilsson et al. 

1992). 

Aureobacterium luteolum Yokota et al. isolated from pond water caused erosions 

in the secondary wall in microtome sections of pine sapwood as substrate 

in 1 month of incubation, that is, bacterial wood degradation occurred obviously 

also by a pure culture under laboratory conditions (Schmidt et al. 1995; 

Fig. 5.3c). The result was however not reproducible using another strain of A. 

luteolum (Nilsson pers. comm.). 

In contrast to the xylem of healthy trees, which was rather “sterile”, wood 

samples from forest dieback sites contained several bacteria (Schmidt 1985; 

Schmidt et al. 1986). In view of the forest damage by pollution, bacteria (including 

RLOs and MLOs) were however assumed to be no causal agents, but 

rather, apart from other influences (emissions, climate, location), predisposing 

factors, or secondary parasites of the weakened trees. 

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Fig.5.3. Bacterial wood decay. a First photo (1944 by J. Liese) of wood cell walls degraded by 

bacteria showing destroyed pine wood tracheids within a foundation pile (from Liese 1950). 

b Degradation of a pine sapwood tracheid cell wall during 3 months of ponding in a lake 

(TEM, from Schmidt et al. 1987). c Degradation of a tracheid cell wall in the laboratory 

by Aureobacterium luteolum (TEM, from Schmidt et al. 1995). B bacterium, E erosion, 

S secondary wall, MP middle lamella/primary walls, R cell wall residues 

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In logs, which were stored for protection from decay fungi, staining and 

insect attack in the open (v. Aufseß 1986; Schmidt et al. 1986) or were sprinkled 

(sprayed) or water-stored (ponded) (Karnop 1972a, 1972b; Berndt and Liese 

1973; Schmidt and Wahl 1987), bacteria degraded in situ within a few weeks 

the non-lignified margo fibrils of the sapwood bordered pits (Fig. 5.4). Several 

bacterial isolates were obtained (Schmidt and Dietrichs 1976). The increased 

wood porosity may cause wood cracks during artificial drying and an irregular 

over-uptake of preservative solutions, varnishes, stains, or paints resulting in 

uneven finishes (Willeitner 1971). Wood spots due to increased permeability 

and bad smell of bacterial metabolites are current problems when wet-stored 

wood is used for indoor wood paneling. 

Timber in service is colonized by bacteria, if the wood is very wet and thus 

less suitable for fungi due to reduced oxygen content. Early reports (Liese 

1950; see Fig. 5.3a) on bacterial degradation refer as to wood in long-lasting 

ground contact (Levy 1975b), as in foundation piles, sleepers, or to wood 

in water, like in cooling towers, harbor constructions and boats (Liese 1955). 

Cell wall degradation even occurred in chromium-copper-arsenic-treated piles 

and poles (Willoughby and Leightley 1984; Singh and Wakeling 1993). The 

bacteria dissolved the toxic components and thus favored wood degradation 

by soft-rot fungi (Daniel and Nilsson 1985). Wood samples impregnated with 

chromium-copper-arsenate and incubated with bacterial pure cultures showed 

increased wood mass loss during subsequent incubation with Coniophora 

puteana (Willeitner et al. 1977). 

Bacteria are often found in archaeological woods from buried and waterlogged 

environments (Blanchette 1995; Björdal et al. 1999; Kim and Singh 1999, 

2000; Singh et al. 2003; Björdal et al. 2005; Schmitt et al. 2005). In those wet 

conditions, bacterial wood degradation is often associated with soft-rot fungi 

(Willoughby and Leightley 1984; Singh et al. 1991; Singh and Wakeling 1993). 

Fig.5.4. Destruction of a pine sapwood 

bordered pit showing the detachment of 

the torus(T) by bacterial (B)degradation 

of the margo fibrils. (REM, from Peek 

and Liese 1979) 

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Woods from Tertiary fossil forests after 20–60 million years of burial showed 

indications that nonbiological degradation was responsible for the changes in 

the cell walls (Blanchette et al. 1991). 

Bacteria are also involved in wood discoloration. The wood of the light 

African Ilomba, Pycnanthus angolensis, is colonized after felling during storage 

and shipment of the round timber. The bacteria spread in the stem interior 

and cause reddish-brown discoloration. Further discoloration develops during 

air-drying of the boards in the area of the stacked wood (sticker stain). As 

causal bacteria e.g., Pseudomonas fragi (Eichholz) Gruber was isolated, which 

remained active in the damp wood parts (contact with stacked woods) and 

increased there the pH value from about 5.5 to 7.5–8.5 by producing ammonia 

from the protein of the protein-rich wood species. This alkalinity results in 

chemical reactions (phenol oxidation and polymerization) of accessory components 

in the parenchyma cells, which cause the brown discoloration (Bauch 

et al. 1985). The bacterium also discolored wood samples in vitro (Fig. 5.5). 

Bacterial discoloration of Ilomba wood during air-drying could be almost 

completely prevented by previous dipping the fresh boards in a solution of 

each 5% formic acid and propionic acid. 

Several bacteria were isolated from beech trees that possessed an irregular 

stellar-shaped red heart (splash-heart). The bacteria caused also in vitro 

brown discoloration of light beech wood samples and wood capillary liquids 

by raising the pH value to over 7.3 (Schmidt and Mehringer 1989; also Mahler 

et al. 1986; Walter 1993). 

Pseudomonas aeruginosa (Schroeter) Migula discolored Obeche, Triplochiton 

scleroxylon (Hansen 1988). In water-stored pine stems, bacteria produced 

flavonoids from flavone glycosides, which diffused to the wood surface during 

drying the sawn timber and caused there brown discolorations (Hedley and 

Meder 1992). 

Bacteria were inhibited by chromium-copper wood preservatives and further 

preservation salts used against fungi. Concentrations used for fungi were 

mostly sufficient to prevent bacterial activity (Schmidt and Liese 1974, 1976; 

Liese and Schmidt 1975; Schmidt et al. 1975). Archaeological woods, like 

the Bremen Cog of 1380, are stabilized against further deterioration using 

polyethylene glycol (Hoffmann et al. 2004). 

Fig.5.5.Bacterial discoloration of Ilomba 

wood within 1 day by a pure culture of 

Pseudomonas fragi inoculated as a line 

on the light wood sample 

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Reviews on “bacteria and wood” are by Liese (1992), Schmidt and Liese 

(1994), Daniel and Nilsson (1998) and Kim and Singh (2000). 

Several bacteria, like Bacillus spp., Pseudomonas spp. and Streptomyces spp., 

were investigated in view of antagonism (Chap. 3.8.1) against fungal parasites 

(Armillaria spp.: Dumas 1992), wood staining fungi (Bernier et al. 1986; Seifert 

et al. 1987; Benko 1989; Florence and Sharma 1990) and decay fungi (Benko 

and Highley 1990). 

Bacteria are currently discussed in connection with the hygiene status of 

wood used for packing, transport, and preparation of foodstuffs. A study, which 

compared wooden and plastic boards used in kitchens, revealed that especially 

pine boards possess hygienic advantages due to its extractives compared to 

other woods and plastic (Milling et al. 2005). 

Pretreatment of spruce wood chips with the actinobacterium Streptomyces 

cyaneus (Krasil’nikov) Waksman for mechanical pulping decreased the energy 

consumption during fiberizing of 24% and increased some strength properties 

of handsheets (Hernández et al. 2005). 

To identify bacteria, predominantly on the basis of morphological and biochemical 

characteristics, “Bergey’s Manual of Determinative Bacteriology” 

(Buchanan and Gibbons 1974) is suitable. “Bergey’s Manual of Systematic Bacteriology” 

(Garrity 2001 et seq.) is the classic reference on bacterial taxonomy 

considering numerous rearrangements and changes in nomenclature, which 

are mainly due to molecular techniques notably sequencing of 16S rDNA and 

analysis of fatty acids. 

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6 

Wood Discoloration 

The damage of wood by fungi is essentially caused by the degradation of 

the cell wall by fungi, which decreases the mechanical wood properties and 

substantially reduces wood use. However, wood quality is also influenced by 

bacterial, algal and fungal discolorations (e.g., Grosser 1985; Zabel and Morrell 

1992; Eaton and Hale 1993). 

Discolorations in the wood of living trees, in round wood, timber and wood 

in service are long-known problems and are based on different biotic and 

abiotic causes (Bauch 1984, 1986; Kreber and Byrne 1994; Koch et al. 2002; 

Koch 2004; Table 6.1). 

Discolorations in standing trees occur after wounding by wound reactions 

ofthetree(Chap.8.2)andbythecolonizationofthestemwoodwithbacteria 

and fungi as a result of microorganism-own pigments (e.g., melanin of bluestain 

fungi, Zink and Fengel 1989) or of their metabolism (brown, white, and 

soft rot in trees, chemical reactions of accessory compounds after pH-change 

by wetwood bacteria and in the splash-heart of beech trees). 

Algae like Chlorococcum sp. and Hormidium sp. soiled and discolored timber 

surfaces (Ohba and Tsujimoto 1996; also Krajewski and Wa˙zny 1992a), whereby 

the green algae Chlorhormidium flaccidum (Kützing) Fot. and Chlorococcum 

lobatum (Kortschikoff) Fritsch & John caused even slight cell wall erosion 

(Krajewski and Wa˙zny 1992b). 

Table 6.1. Biotic and abiotic wood discolorations (completed after Bauch 1984; Butin 1995) 

tree reactions on wounding 

microbial discolorations 

– staining by algae, molds, blue stain and red-streaking fungi 

– grey stain of poplar wood by Phialophora fastigiata 

– pink stain by Arthrographis cuboidea 

– black streaking of beech wood by Bispora monilioides 

– red spotting of beech wood by Melanomma sanguinarum 

– “green rot” by Chlorociboria spp. 

– wood rots 

physiological reaction of living parenchyma cells (“Ersticken” of beech and oak) 

biochemical reaction by wood-own enzymes (“Einlauf” of alder) 

chemical reactions (iron-tannic acid reaction of oak, discoloration of hemlock by zinc) 

combined reaction (brown discoloration of Ilomba by bacterial pH-increase and 

subsequent chemical reaction of phenols) 

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120 6 Wood Discoloration 

The wood-discoloring molds and staining fungi live on nutrients in the 

parenchyma cells of the sapwood. Conifers and hardwoods, round wood, lumber, 

finished wood and wood products can be colonized. Discoloring fungi 

do not cause any or only very little cell wall attack. Prioritization of the color 

damage depends on subsequent wood use. 

Several Deuteromycetes and Ascomycetes stain woody substrates. Phialophora 

fastigiata (Hyphomycetes) causes a grey stain of poplar wood. Arthrographis 

cuboides (Hyphomycetes) produces a pink stain in several hardwoods 

and softwoods, and a naphthalenedione has been isolated from such wood 

(Golinski et al. 1995). Red alder wood used in the USA for furniture is stained 

reddish purple by Ophiostoma piceae if not rapidly processed after harvesting 

(Morrell 1987). Black streaking of beech logs occurs by Bispora monilioides. 

Red spotting of beech wood is effected by Melanomma sanguinarum (Dothideales). 

Paecilomyces variotii produces a yellow discoloration of oak wood 

during drying through its pH-change, which causes chemical reactions of the 

hydrolyzable gallotannins (Bauch et al. 1991). 

So-called green rot is caused by species of the ascomycete Chlorociboria 

(Helotiales). Chlorociboria aeruginascens and C. aeruginosa discolor rotten 

and moist hardwood (and conifer) branches and other woody debris in the 

forest (Jahn 1990). The green wood has often been employed in marquetry 

and veneering and is a feature of the famous Tunbridge ware (Ellis 1976). 

The naphthoquinone pigment, xylindein, produced by the fungus is mainly 

deposited in the ray parenchyma cells as well as in vessels and fibers adjacent 

to the rays. The pigment is now since more than 500 years durable 

(Blanchette et al. 1992a; Michaelsen et al. 1992). In a recent reproduction of 

a violin from the 17th century, green stained wood was used for the ornaments 

(Fig. 6.1). 

Fig.6.1. “Green rot” caused by Chlorociboria species. a Green-rotten poplar wood. The 

missing section was used for the green intarsia b of the replica c, d by T. Schmitt in 1998 of 

a violin by J. Meyer from 1670 (photos b–d:T.Schmitt) 

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6.1 Molding 121 

6.1 

Molding 

Thetermmoldoriginatesfromdailylifeandisnotataxonomicnameofasingle 

systematic group (Reiß 1997; Kiffer and Morelet 2000). The Deuteromycetes 

(fungi imperfecti) constitute an artificial group and comprise a great variety 

of 20,000–30,000 species of 1,700 genera of Hyphomycetes and 700 genera of 

Coelomycetes. The different molds have a broad spectrum of physiological 

response with regard to temperature, water activity, pH value etc. and thus can 

colonize and damage very diverse materials (molding). Molds are significant 

in view of damages to foodstuffs, deterioration of natural materials (leather, 

books, textiles, wallpapers), with regard to human and animal health, and for 

biochemists and the manufacturers of antibiotics [772 of about 3,200 admitted 

antibiotics originate from fungi: Müller and Loeffler (1992)], organic acids 

[e.g., citronic acid, malic acid: Rehm (1980)], enzymes (e.g., amylase, protease, 

lipase, cellulase, pectinase), cheese (Penicillium camemberti, P. roqueforti), 

salami sausages (P. nalgiovense), and “country cured ham” (Aspergillus spp., 

Penicillium spp.) (Schwantes 1996; Reiß 1997). Botrytis cinerea causes the 

“noble rot” of sweet wines. Fusarium oxysporum ssp. cannabis is used as an 

herbicide for suppressing marijuana plants (Kiffer and Morelet 2000). Even 

synthetic floor coverings, airplane fuels, oils, glues, paints, optical glasses, and 

textiles can be overgrown with and damaged by molds. 

With regard to lignocelluloses, seeds, seedlings, young tree roots (Schönhar 

1989), standing trees (Schmidt 1985), stored and blocked wood (Wolf and 

Liese 1977; Bues 1993), piled wood chips (Feicht et al. 2002) of the pulp industry 

(Hajny 1966), stored annual plants, like sugarcane bagasse (Schmidt and Walter 

1978), and books (Kerner-Gang and Nirenberg 1980) can be colonized by 

molds. Paecilomyces variotii (mold and soft-rot activity) is involved in the 

yellow discoloration of oak wood during storage and drying (Bauch et al. 

1991). There are German and European standards and test methods to measure 

growth of molds on and resistance of substrates like electrotechnical products, 

plastics, textiles, optic apparatus, and timber (Kruse et al. 2004). 

Frequently, molds are recognizable by their fast growth on the surface of 

substrates, on which conidia develop rapidly (Fig. 6.2a). Due to the speciesspecific 

color of the conidia, wood colonized by several mold species can make 

a multicolored impression, or it outweighs e.g., black due to Aspergillus niger 

or green after Penicillium spp. or Trichoderma spp. colonization. 

Trichoderma species were the most frequent fungi on spruce roots from forest 

dieback sites (Schönhar 1992). Stored beech stems are frequently colonized 

by Bispora monilioides, which causes black, radially arranged, elliptical strips 

on the fresh trunk cross surface. 

Molds develop on fresh cuts after tree felling, particularly on the moist 

sapwood, on inappropriately stored lumber, insufficiently dried and airtight 

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Fig.6.2. Molding. a Substrate surface with various molds (REM, photo W. Kerner). b Molding 

of insufficiently dried and then plastic-covered paneling 

sealed wood, like plastic-coated paneling (Fig. 6.2b), during sea transport of 

round timber and wood products under deck and in chip piles of pulp industry. 

Among 427 isolates from stacked yellow-poplar lumber, Penicillium 

implicatum and Aspergillus versicolor accounted for 29.7 and 14.5%, respectively 

(Mikluscak and Dawson-Andoh 2004b). 

Thehyphaepenetratethewoodonlyafewmillimetersandliveonparenchyma 

cells (sugar, starch, protein). In the laboratory, some species degraded 

isolated pectin, hemicelluloses, and cellulose, but not lignified cell walls. Thus, 

wood strength properties remain unchanged. 

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6.1 Molding 123 

Molded wood is, however, unmarketable. For decorative purpose, e.g., wall 

paneling (Fig. 6.2b), molded wood is unsuitable, as the color spots are not mechanically 

removable, but can be only masked by colored paints. Infected wood 

is not suitable for various hygienic requirements, e.g., packaging material. In 

addition, technological characteristics, for example the gluing of plywood, can 

be affected by molds (Wolf and Liese 1977). 

Mold growth in buildings is increasingly becoming a problem. Molding 

in indoor environments (Thörnqvist et al. 1987) is favored by high substrate 

moisture (water activity 0.9–1.0), high air humidity around 95%, warmth and 

insufficient ventilation (Viitanen and Ritschkoff 1991b), like in cellars and 

bathrooms. According to the German standard DIN 4108 part 2, the relative air 

humidity on the indoor surfaces shall not amount to over 80% (Borsch-Laaks 

2005). Moisture with following mold contamination can arise from condensation, 

flood, and various types of leaks. Excessive insulation after the petroleum 

crisis has markedly favored condensation areas (cold bridges), from cellars 

to attics, which rapidly become sites of mold growth. Accompanying lifestyle 

changes (frequent showers, new cooking methods, inadequate airing of bedrooms) 

have led increasingly to the production and accumulation of moisture 

in the home. A study in Belgium of isolated molds in homes of patients with 

allergic problems showed that more than 90% of those houses were contaminated 

by molds of the genera Cladosporium, Penicillium and Aspergillus (Nolard 

2004). Cladosporium sphaerospermum infiltrated 60% of the homes and 

was responsible for high contaminations, particularly in bedrooms and bathrooms. 

Aspergillus versicolor, Penicillium chrysogenum, P. aurantiogriseum, 

P. spinulosum, P. brevicompactum, Chaetomium globosum, Stachybotrys chartarum, 

andAlternaria alternata are often found on the walls of bedrooms, 

living rooms, and kitchens. While Cladosporium herbarum, a phytopathogen, 

does not grow in houses, large numbers of spores enter through windows and 

doors mainly during the summer months. 

Molds may cause health problems. About 200 fungal species produce various 

mycotoxins (about 100), of which some are highly toxic to humans and 

animals (mycotoxicoses) (Müller and Loeffler 1992; Schwantes 1996; Reiß 1997; 

Kiffer and Morelet 2000; Samson et al. 2004). The cancerogenic aflatoxins from 

Aspergillus fumigatus and A. flavus in food (agricultural crops, cereals etc, 

Meister and Springer 2004) are well known. Human health damage can further 

develop by mycoallergies through direct contact with a fungus or inhaled 

spores (molds in the living space). Five to 15% of the population suffering 

from respiratory allergy has been sensitized to one or several molds. Exposure 

of young children to molds and their metabolites may have a “stimulating” 

effect on the onset of later allergies (Nolard 2004). Mold allergies also occur 

in work environments. Woodworkers inhale spores of Cryptostroma corticale 

and Alternaria species (woodworker’s lung). “Bagassosis” may develop during 

bagasse processing. “Suberosis” is due to Penicillium glabrum growing on cork 

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bark. Pulpwood handler’s disease is caused by Alternaria species growing on 

paper pulp. Farmers inhale spores of Aspergillus fumigatus when damp hay 

is worked. Dustmen and compost makers may be exposed to molds when 

kitchen waste is stocked in closed containers for too long. Mushroom growers 

may be exposed to huge quantities of spores released by the basidiomycete 

they cultivate, and the culture substrate is sometimes contaminated by molds 

(mushroom grower’s disease) (Nolard 2004). 

Superficial mycoses occur on mucous membranes (fingernail bed, lips) and 

profound mycoses after wounding the skin or inner body (ear, eye, lung, blood 

vessels). Deuteromycetes are also significant in view of immunodepression in 

cases of transplants and of diminished defense mechanisms of AIDS sufferers. 

With regard to indoor environments (Frössel 2003; Hankammer and Lorenz 

2003) only a few molds are considered as producers of important toxic compounds 

which can be released in the environment and which may cause severe 

health problems (Samson and Hoekstra 2004). These are Alternaria alternata, 

Aspergillus flavus, A. fumigatus, A. versicolor, Chaetomium globosum, Emericella 

nidulans, Memnoniella echinata, andStachybotrys chartarum, whereby 

the latter is considered the most important toxic fungus in buildings producing 

the cytotoxic satratoxins. A questionnaire study among U.S. homebuilders, 

new homeowners, and real estate agents indicated that overall, respondents 

did not have a strong understanding of how mold forms in new constructions. 

Ten percent of homeowners believed that mold was an issue in their neighborhoods 

while 35% of home builders and 19% of real estate agents believed 

that this was an issue in the homes they built (Vlosky and Shupe 2004). The 

aspect of molds on indoor piled chips was treated by Feicht et al. (2002). Air 

sampling is performed to quantify and identify contamination. Measurement 

of microbial volatile organic compounds (MVOCs) in houses serves as note 

for contamination, especially for hidden contaminations (Keller 2002). Stachybotrys 

chartarum and Chaetomium globosum emitted ketones and alcohols 

(Korpi et al. 1999). There are also dogs trained to detect molds by sniffing. 

For remediation, first of all the cause of the damage (dampness) has to be 

removed continuingly (Neubrand 2004). In view of allergies, the spores may 

be taken away. There are primers and paints with prophylactic anti-molding 

substances, like organic sulfur-nitrogen compounds (thiocarbamate) and organic 

tin compounds (tributyltin oxide). Yang et al. (2004b) proposed that 

incorporating tree bark (white spruce), which inhibited mold growth in vitro, 

into the production of composite boards may increase the resistance of panels 

to fungi. 

Particularly several Trichoderma species are antagonistic against other organisms 

and also destroy (mycoparasitisms) fungal parasites and saprobionts 

(v. Aufseß 1976; Highley and Ricard 1988; Murmanis et al. 1988; Giron and Morrell 

1989; Doi and Yamada 1991; Dumas and Boyonoski 1992; Phillips-Laing 

et al. 2003). 

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6.2 Blue Stain 125 

There are various textbooks and keys to identify molds (e.g., Wang 1990; 

Kiffer and Morelet 2000; Samson and Hoekstra 2004; Samson et al. 2004). 

The attachment of a species to the molds is not always strict. There are overlappings 

with blue-stain and soft-rot fungi since fungi traditionally implicated 

in wood discoloration can cause soft rot if the conditions are suitable (e.g., Alternaria 

alternata, Cladosporium herbarum, Aspergillus fumigatus) and many 

soft-rot fungi are highly melanized (e.g., Phialophora spp.). That is, a fungus 

all may show the typical superficial mold growth and is treated in textbooks 

on molds, but also effected blue stain, or produced weight loss in soft-rot tests 

(Seehann et al. 1975; Daniel 2003). 

6.2 

Blue Stain 

Blue stain (synonymous sap stain) is a blue, grey or black, radially striped 

wood discoloration of sapwood, which can be caused by about 100 to 250 

(Käärik 1980) fungi belonging to the Ascomycetes and Deuteromycetes. Seifert 

(1999) and others differentiated three groups of blue-stain fungi: – Ceratocystis, 

Ophiostoma and Ceratocystiopsis species (Upadhyay 1981; Perry 1991; 

Gibbs 1999), – black yeasts such as Hormonema dematioides, Aureobasidium 

pullulans, Rhinocladiella atrovirens, andPhialophora species, – dark molds 

such as Alternaria alternata, Cladosporium sphaerospermum,andC. cladosporioides. 

Yang (1999) differentiated dark staining fungi, such as Ophiostoma 

piliferum on jack pine, Ceratocystis minor on white pine, and C. coerulescens 

on white spruce, and light staining fungi, such as O. piceae, C. adiposa and 

Leptographium sp. Frequently, like in the Ophiostoma species, the teleomorph 

is a perithecium (Figs. 2.14, 6.3E). Blue stain occurs in conifers, particularly 

in pine, but also in spruce, fir, and larch, in hardwoods, like beech and birch, 

and in tropical woods. The stain may be superficial or penetrate deeply into 

the wood. In heartwood species, only the sapwood discolors, since blue-stain 

fungi live mainly on the content of the parenchyma cells. Figure 6.3 shows 

some details of blue stain. 

The hyphae are brown colored due to melanin (Zink and Fengel 1989) and 

relatively thick (Fig. 6.3C). Some species like A. pullulans develop dark-brown, 

thick-walled chlamydospores (Fig. 6.3D). The blue-black color of the wood 

develops as optical effect due to refraction of light. Hyphae penetrate into stem 

wood from cross sections or radially through bark fissures and move via the 

medullary rays. Easily accessible nutrients (sugars, carbohydrates, starch, proteins, 

fats, extractives) are taken up from the ray parenchyma cells. Xylanase, 

mannanase, pectinase and amylase have been detected in several blue-stain 

fungi (Schirp et al. 2003a). From the rays, the hyphae penetrate into the longitudinal 

tracheids with mechanical pressure through the torus of the bordered 

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Fig.6.3. Blue stain in wood. A Artificial bluing of pine boards by Phoma exigua. B Detail. 

C Thick, brown hyphae of P. exigua. D Chlamydospores (photo G. Koch). E Perithecia (A, 

B, C, E from Schmidt and Huckfeldt 2005), — 5 cm, --- 5 mm 

pits (thin hyphae through the margo) and grow there from cell to cell through 

the pits. Because fungi colonize the sapwood tracheids and fibers, components 

of the capillary liquid also might be used as nutrients. Although there are 

special microhyphae, transpressoria (Fig. 2.5), which can break through the 

wood cell wall, probably by physical pressure and/or enzymatic action (Schmid 

and Liese 1966; Liese 1970), in most cases the strength properties of wood are 

hardly affected. Thus, the occasionally used term “blue rot” is wrong. Some 

species however caused some strength loss. Toughness was the property most 

seriously affected (Seifert 1999; Schirp et al. 2003b). In most cases, however, 

the damage to wood is mainly cosmetic. The damage however affects domestic 

and export earnings for the forest industries. For example, Pinus radiata in 

New Zealand is highly susceptible to blue stain with an estimated annual loss 

in revenue of NZ$ 100 million per year (Thwaites et al. 2004). 

Temperature minimum depends on the species, and is between 0 and −3 ◦ C; 

the optimum is between 18 and 29 ◦ C and the maximum is between 28 and 

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6.2 Blue Stain 127 

40 ◦ C. The moisture span reaches from fiber saturation close to umax.Inmany 

species, the optimum is between 30 and 120% (Käärik 1980; Schumacher and 

Schulz 1992). For log colonization, moisture loss in the felled tree of 10–15% 

is sufficient. Blue stain occurs during seasoning or transportation of green 

lumber before the wood is dried and is enhanced at relative humidities above 

90% (Seifert 1999). 

Blue-stain fungi were arranged into different ecological groups (Butin 1995): 

In blue stain of stems (primary blue stain), spores of Ophiostoma species (moisture 

optimum 50–130%), particularly Ophiostoma piceae (Harrington et al. 

2001) and also Discula pinicola are transferred by wind in bark wounds (forest 

work or wood transport) as well as by bark beetles particularly in un-debarked 

pine stems which are allowed to dry out slowly over weeks or months while lying 

in the forest (Neumüller and Brandstätter 1995). Hormonema dematioides, 

A. pullulans, and a Leptographium species were the most frequently isolated 

stain-fungi from bark and sapwood of living Pinus banksiana trees. There were 

indications that none of the well-known log-staining fungi was associated with 

healthy living jack pine trees, and it was deduced that prompt transportation 

of logs from forests to sawmills and sanitary treatment of log storage yards 

helps to reduce the severity of log staining before sawing (Yang 2004). The 

most aggressive sapstain species on fresh Pinus contorta logs was Ceratocystis 

coerulescens, followed consecutively by Leptographium spp., C. minor, O. piliferum, 

O. piceae, O. setosum, C. pluriannulata, andA. pullulans (Fleet et al. 

2001). Discula pinicola is the main cause of the so-called internal blue stain, 

which is characterized by a central wood discoloration without any external 

staining. A comparison of the growth of several blue-stain fungi in freshly cut 

pine billets has been performed by Uzunović and Webber (1998). The bluestain 

fungal composition on Pinus radiata logs harvested in New Zealand and 

shipped to Japan showed differences between summer and winter transport 

(Thwaites et al. 2004). 

Blue stain of sawn timber (secondary blue stain) is caused e.g., by Cladosporium 

species (moisture optimum 50–100%) and Strasseria geniculata (Butin 

1995) in sawn timber that is not completely dry or badly stacked in timber 

yards (Schumacher et al. 2003). 

The classical distinction in primary and secondary blue-stain fungi was not 

confirmed however by the frequent occurrence of D. pinicola both in stored 

pine stems and in boards (Schumacher and Schulz 1992). Battens of Sitka 

spruce were stained by O. piceae when the surface moisture content in a stack 

was 22% or more (Payne et al. 1999). 

Tertiary blue stain (moisture optimum 30–80%) results frequently from A. 

pullulans and Sclerophoma pithyophila on timber that has been converted into 

products, was painted and re-imbibe moisture while in service, like wooden 

façades, window frames, garage doors and garden furniture. Through damages 

of the coating in window wood e.g., by nails or due to inappropriate 

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window construction, water is taken up, distributes in the wood and cannot 

evaporate through the coat layer. Fungi start growing and their mycelia, spore 

masses or perithecia (Fig. 6.3E) cause the paint layer to flake off with further 

moisture increase (Sell 1968). Hyphae of A. pullulans were able to grow 

through alkyd paints (Sharpe and Dickinson 1992). Colonization of painted 

wood by blue-stain fungi was treated by Bardage (1997). Tertiary blue-stain 

fungi do not originate from infected stems or lumber, but are new infections. 

Colonized wood shows excessive uptake of solutions, so that spot-shaped 

color differences develop after painting, similarly like at the excessive uptake 

caused by bacteria. The isolate A. pullulans P 268 is test fungus in the standard 

EN 152. 

Air-borne blue stain means the spread of blue-stain fungi by wind or rain, 

insect blue stain is due to fungi, which are associated with bark beetles (Solheim 

1992). 

There are different results in view of blue staining of wood that derives from 

forest dieback sites. Practical observations and fungal isolations (Schmidt 

1985) showed that wood from polluted forest sites was more stained than 

that from healthy forests. Laboratory experiments however did not show these 

differences (Liese 1986; Saur et al. 1986). Klepzig et al. (1996) found different 

interactions of ecologically similar saprogenic fungi with healthy and 

abiotically stressed trees. Regarding the storage of spruce, pine and beech 

stems (v. Aufseß 1986; Göttsche-Kühn and Frühwald 1986; Schmidt et al. 1986; 

Schmidt and Wahl 1987; Nimmann and Knigge 1989) the wood from diseased 

trees first tended to faster discolorations due to fungal attack. However, after 

longer storage no relation was found between the state of health of the 

tree and the damage extent during storage. On the contrary, the stems of 

healthytreeswereevenmorestronglydiscolored,sincetheirlongerlasting 

drying period provided for the fungi a longer time favorable growth 

conditions. Stored planks from damaged pine trees were also slightly less 

stained than wood from healthy trees (Schumacher and Schulz 1992). Altogether, 

there are no results justifying the occasionally used term “damage 

wood”. 

Incubation of fresh Scots pine sapwood samples with blue-stain fungi increased 

wood absorptiveness and the wood may show a greater ability to 

impregnation with water-based preservatives (Fojutowski 2005). 

Stained wood is used due its color effects by Swedish woodworkers and 

was also used to produce attractive violins (Seifert 1999). Corresponding attempts 

to stain timber artificially did however not yield regular discoloration 

of the samples (Fig. 6.3A). It is possible to remove the stain from the wood 

using oxidizing agents such as sodium chlorite or hydrogen peroxide (Seifert 

1999). 

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6.3 Red Streaking 129 

6.3 

Red Streaking 

Red-streaking discoloration (known as “Rotstreifigkeit” in Germany) is one of 

the most common and important damage in seasoning logs and sawn lumber, 

occurring only in conifers (spruce, pine, fir) and recognized as a distinct condition 

in continental Europe. The stripe-shaped to spotted yellow to reddishbrowndiscolorationextendsinlogsfromboththeirbark-coveredfacesand 

from their cut ends (Butin 1995; Baum and Bariska 2002) (Fig. 6.4). Stems that 

are not debarked show a rather flat discoloration and debarked stems exhibit 

a streakier staining (v. Pechmann et al. 1967). 

Causal agents are several white-rot Basidiomycetes, in spruce particularly 

Stereum sanguinolentum (Kleist and Seehann 1997) and Amylostereum areolatum.InsouthGermany,Amylostereum 

chailettii is common (Zycha and Knopf 

1963; v. Pechmann et al. 1967). In pine, red streaking is mainly due to Trichaptum 

abietinum (Butin 1995). According to Kreisel (1961), S. sanguinolentum 

and T. abietinum occur often together in stored logs. 

Red streaking develops if the wood remains in a semi-moist state over 

a long period, especially in the warmer season (v. Pechmann et al. 1967). The 

fungi gain access to the wood through the exposed cut ends and bark fissures. 

The mycelium reaches its greatest density in the medullary rays, where the 

fungus uses the primary storage compounds in the ray parenchyma cells. 

From there, the discoloration spreads axially deeply in the wood, penetrating 

the bordered pits and also by thin bore hyphae that perforate the tracheids 

cell wall (Kleist and Seehann 1997; Kleist 2001). Logs may be stained during 

overseas shipment, and red streaks producing fungi become again active in 

rewetted boards due to their ability to dryness resistance. The staining is mainly 

an oxidative process (Butin 1995). Kleist (2001) stated that the fungi involved 

excrete the pigments. 

The moisture optimum of most species lies between 50 and 120% u. Redstreaking 

fungi are slowly growing white-rot fungi, so that initially no serious 

strength loss is connected with turning red. During longer colonization however 

an intensive white rot develops with substantial mass and strength loss, so 

that red streaking damage represents a transition from discoloration to decay 

(v. Pechmann et al. 1967; Peredo and Inzunza 1990). 

Secondary infections by brown-rot fungi may occur. Red-streaked wood 

samples were degraded in the lab test more strongly by brown-rot fungi than 

controls without pre-infection. From reddish discolored fir wood, 26 Basidiomycetes 

(white and brown rot) and numerous blue-stain and mold fungi 

were isolated (v. Pechmann et al. 1967). From Pinus radiata wood, different 

molds, blue-stain fungi, Stereum sp. and the white-rot fungi Ganoderma sp., 

Schizophyllum commune and Trametes versicolor were isolated (Peredo and 

Inzunza 1990). Spruce wood samples from forest dieback sites contained more 

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Fig.6.4. Red-streaking discoloration of spruce wood by Stereum sanguinolentum. a Fruit 

bodies of S. sanguinolentum on the crosscut stem surface. b Wood discoloration some 

centimeters beneath the surface (photos G. Kleist) 

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6.4 Protection 131 

often A. areolatum and S. sanguinolentum compared to samples from healthy 

forests (Schmidt et al. 1986). 

Stereum sanguinolentum Bleeding Stereum 

small, thin, resupinate to semipileate fruit body, soft-leathery-crusty, bowlshaped, 

upper surface: felty, concentrically zonate, yellow-brown, whitish-wavy 

margin (Fig. 6.4a); bright to grey-brown hymenium blood-red after injury; 

dimitic (Breitenbach and Kränzlin 1986); amphithallic (Calderoni et al. 2003); 

apart from the saprobic way of life also parasitic after penetration through 

wounds and thus the most important species of “wound rot of spruce” (Butin 

1995); stacked wood not attacked; genus Stereum with multiple clamps (Kreisel 

1969). 

Trichaptum abietinum Fir Polystictus 

fruit body: annual, resupinate to semipileate and pileate, singly and roofing 

tile-like; upper surface: white-grey-brown, thin, felty, hirsute, zonate, leathery; 

pore surface: young net-shaped to porous, old: labyrinthine; young hymenium 

reddish with angular violet pores, later brown-violet; dimitic (Breitenbach and 

Kränzlin 1986); tetrapolar heterothallic (Nobles 1965); saprobic on stumps, 

stored logs and finished wood; severe white rot at high wood moisture; rarely 

on living trees (Kreisel 1961). 

6.4 

Protection 

To avoid microbial wood discoloration, the generally suitable measures against 

fungi (e.g., Liese et al. 1973; Liese and Peek 1987; Groß et al. 1991; Yang and 

Beauregard 2001) are listed in Table 6.2. 

Felling in the cold season and fast processing of the stems through well 

coordination between forestry and wood industry reduces microbial activity 

during storage of the stems in the forest. Cool, shady, and ventilated storage 

without ground contact and with unhurt bark to maintain high wood moisture 

content and to prevent lateral infections are favorable. Lumber discoloration 

can be prevented by prompt air-drying in well-ventilated stacks protected 

against rain by a roof, or by kiln-drying. Wet storage of stemwood by sprinkling 

or ponding protects against fungi and insects. Currently, stem storage 

Table 6.2. Preventive measures to avoid microbial wood discolorations and decay 

– felling in the cold season 

– appropriate storage of fresh wood 

– coordination between forestry and wood industry 

– drying 

– wet storage 

– storage in N2/CO2 atmosphere 

– chemical preservation 

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is performed in a N2/CO2 atmosphere (Mahler 1992; Bues and Weber 1998; 

Maier et al. 1999). 

During wet storage, however, wood quality may become reduced by degradation 

of the pits by anaerobic bacteria (Willeitner 1971; Karnop 1972a, 1972b; 

Adolf et al. 1972; Fig. 5.4), by oxidative discolorations of phenolic compounds 

diffusing outward (Höster 1974), and by brown discoloration of the outer 

log parts through phenolics from the bark (Peek and Liese 1987; Bues 1993). 

Sprinkled stems were even colonized by Armillaria mellea, which “drilled” 

a borehole from the bark into the xylem to provide itself with air and subsequently 

decayed the wet wood (Metzler 1994). 

Discoloring fungi and molds may be rather tolerant towards several fungicides, 

which inhibit decay fungi. Numerous protective agents were investigated 

for their effectiveness against mold and blue-stain fungi: e.g., Karstedt 

et al. (1971), Wolf and Liese (1977), Nunes et al. (1991), Laks et al. (1993), 

Wakeling et al. (1993), and Suzuki et al. (1996). Sodium pentachlorophenate 

(PCP-Na) had been used for dipping and spraying procedures against discoloration 

and decay (Willeitner et al. 1986). In view of the negative impact on 

humans, animals, plants, and the environment, utilization of PCP and import 

of PCP-treated woods are however restricted in Germany due to contaminations 

of PCP with polychlorinated dibenzodioxines and dibenzofuranes as well 

as due to the development and release of these compounds during burning of 

PCP containing woods. Dependent of material and intended purpose, e.g., 

boron compounds, quaternary ammonium compounds or dithiocarbamates 

may be used (Chap. 7.4). Solid wood, wood composites (Gardner et al. 2003), 

and gypsum wallboard treated with borate were tested for mold performance 

(Fogel and Lloyd 2002). Boron compounds were used against blue-stain in 

Norway spruce (Babuder et al. 2004) and rubber wood (Akhter 2005). Against 

discolorations of drying oakwood by Paecilomyces variotii, treatment of the 

fresh wood with 5–10% propionic acid was recommended (Bauch et al. 1991). 

Growth of molds and bacteria during the outdoor storage of sugarcane bagasse 

on Trinidad that is used there for the production of fiberboards was reduced 

by organic sulfur compounds and propionic acid (Liese and Walter 1980). 

Although blue-stain fungi do not reduce wood quality significantly, discoloration 

is considered as substantial damage and is a perpetual problem of 

round wood and timber. Despite felling during the cold season as well as using 

ventilated stacking of the lumber, damage nevertheless occurs by blue-stain 

fungi. A two-year experiment with pine wood using different felling times and 

storage variations showed that damage of the round timber might be reduced 

and that rapid timber seasoning has the greatest influence (Schumacher and 

Schulz 1992). 

Un unsolved problem is the discoloration of bright tropical woods, like 

Pycnanthus, Virola, Aningeria and Pterygota (Bauch et al. 1985), after felling 

and during shipment and drying of the sawn timber (Karstedt et al. 1971; 

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Fougerousse 1985). Discolorations result from oxidative reactions of accessory 

compounds with atmospheric oxygen and phenol oxidases (e.g., Neger 1911; 

Oldham and Wilcox 1981), from chemical reactions of wood contents with 

metals [iron, zinc: e.g., Bauch (1984)], or from microorganisms, particularly 

blue-stain fungi, and in some woods, like Ilomba, from “combined influences” 

[bacterial pH-change and subsequent chemical reactions (Bauch 1986; see 

Fig. 5.5, Table 6.1)]. The practical processing of wood preservation in the 

tropics against discolorations and decay is summarized by Willeitner and 

Liese (1992) (also Findlay 1985). 

Comprehensive investigations on red streaks producing fungi, their reduction 

of wood quality and on suitable storage are described by v. Pechmann 

et al. (1967). Since fungal damage is usually only superficial in the first months, 

deeper discolorations can be limited to a practically insignificant extent, if 

the log does not remain in the forest in the warm season longer than some 

months. The wet to moist condition of the wood should rapidly run through either 

by suitable forest storage (no ground contact, ventilated, shady), or a high 

moisture content should be maintained in the sapwood by an unhurt bark. 

Attempts of a “biological wood protection” by antagonism are described in 

Chap. 3.8.1. 

To prevent enzyme-mediated, non-microbial sapwood discolorations such 

as sticker stain in ash or grey stain in oak, logs were treated with fumigants to 

kill living parenchyma cells (Amburgey et al. 1996; also Schmidt et al. 1997b; 

cf. Chap. 8.1.2.2). 

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7 

Wood Rot 

There are three types of fungal wood rot: brown, white, and soft rot (see 

Figs. 7.1–7.4). Further terms are either older names (e.g., destruction rot = 

brown rot), specifications (red rot = white rot by Heterobasidion annosum)or 

terms used in practice (marble rot = white rot with black demarcation lines) or 

false names (blue rot = blue stain). According to the classical school of thought 

a fungal species causes only one type of decay, and species causing different 

rots shall not be grouped in the same genus [e.g.: Lentinus lepideus: brown rot; 

Lentinula (in former times Lentinus) edodes:whiterot]. 

Regarding the delineation between the three decay types, there are, however, 

exceptions: The brown-rot fungus Coniophora puteana produced cavities to be 

typical of soft-rot fungi and erosion and thinning of the cell wall to be characteristic 

of white-rot fungi (Kleist and Schmitt 2001; Lee et al. 2004). Fistulina 

hepatica revealed the soft-rot mode in cell walls rich in syringyl lignin, whereas 

brown rot was associated with cells rich in guaiacyl lignin (Schwarze et al. 2000). 

Several white-rot Basidiomycetes like Phellinus pini (Liese and Schmid 1966) 

as well as Inonotus hispidus and Meripilus giganteus caused cavities (Schwarze 

and Fink 1998; Schwarze et al. 1995a), which differed between the host trees, 

cell type, and location in the annual ring. Cavities in the secondary wall of 

fibers and tracheids were also found to be caused by two Armillaria species as 

well as by Stereum sanguinolentum, Ganoderma applanatum, and Grifola frondosa 

(Schwarze and Engels 1998). It was hypothesized that soft-rotting activity 

of white-rot Basidiomycetes may commonly precede white rotting when the 

fungus invades previously uninfected zones in the xylem, in which moisture 

content is high. Delignification of Norway spruce tracheids by Stereum sanguinolentum 

was associated with the presence of radial and concentric clefts 

containing cell wall entities in the secondary wall (Schwarze and Fink 1999) 

supporting observations of a radial and concentric arrangement of cell wall 

constituents within the S2 (Sell and Zimmermann 1993). 

7.1 

Brown Rot 

Brown rot is caused by Basidiomycetes, which metabolize the carbohydrates 

cellulose and hemicelluloses of the woody cell wall by non-enzymatic and 

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136 7 Wood Rot 

enzymatic action and leave the lignin almost unaltered (Fig. 7.1A; Chap. 4), 

whereby the brown color develops. 

Brown-rot fungi do not produce lignin-degrading enzymes. There are however 

reports of lignin peroxidase and manganese peroxidase in some brown-rot 

fungi, and lignin loss or metabolization by brown-rot fungi have been reported. 

Particularly in later stages of decay, the highly lignified middle lamella/primary 

walls were observed to undergo attack. Also, the penetration of the wood cell 

wall by bore holes removes lignin in the process, all suggesting that low molecular 

weight lignin degrading agents and potentially even lignin degrading 

enzymes max occur in some brown-rot fungi, at least with localized activity 

(Goodell 2003). Laccase activity was also found in Coniophora puteana (Lee 

Fig.7.1. Brown rot. A Cubic crack. B Wood cell wall showing remaining lignin after carbohydrate 

degradation (TEM, photo W. Liese). C Brown cubical rot by Oligoporus amarus. MP 

middle lamella/primary walls, S secondary wall, L lumen 

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7.1 Brown Rot 137 

et al. 2004), and in Gloeophyllum trabeum and Oligoporus placenta (Goodell 

2003). Non-enzymatic, low molecular agents produced by the brown-rot fungi 

are responsible for initial stages of cell wall attack (Goodell 2003; Chap. 4). 

Of about 1,700 wood-degrading Basidiomycetes in North America, only 120 

species (7%) caused brown rot, and of these 79 (65%) were polypores (Eriksson 

et al. 1990; Ryvarden and Gilbertson 1993). White-rot fungi distribute broader 

over the different basidiomycetous groups and some belong to the Ascomycetes 

(Rayner and Boddy 1988). Most brown-rot fungi affect conifers (Ryvarden and 

Gilbertson 1993), while white-rot fungi occur more frequently on hardwoods. 

Brown rot occurs in standing trees, felled and processed wood as well as in 

sapwood and heartwood. In the northern hemisphere, the majority of timber 

used in construction is from conifers. Thus, a large part of wood in outdoor and 

indoor service is destructed due to the action of brown-rot fungi. Brown rot is 

usually uniformly distributed over the substrate. A brown cubical pocket rot is 

caused by Laurelia taxodii in cypress and by Oligoporus amarus (Fig. 7.1C) in 

incense cedar. Decay pockets are localized and surrounded by firm wood (Zabel 

and Morrell 1992). A woody substrate both may show brown rot and white rot; 

a standing tree of Picea engelmannii exhibited “white pocket rot” by Phellinus 

pini in the heartwood (Chap. 8.3.8), and after wind throw the healthy areas 

became brown-rotten (Blanchette 1983). Brown-rot wood debris is extremely 

stable due to its content of slightly modified lignin and has remained unaltered 

in the soil for centuries. In conifers forests, this humic material may comprise 

up to 30 vol% in the upper layers (Swift 1982; Ryvarden and Gilbertson 1993). 

Table 7.1 lists some important brown rot. 

Table 7.1. Some common brown-rot fungi 

Fungus Predominant occurrence 

standing timber timber softwood hardwood 

tree outdoors indoors 

Laetiporus sulphureus × × 

Phaeolus schweinitzii × × 

Piptoporus betulinus × × 

Sparassis crispa × × 

Gloeophyllum spp. × × 

Daedalea quercina × × 

Lentinus lepideus × × 

Paxillus panuoides × × 

Antrodia spp. × × 

Coniophora spp. × × 

Serpula lacrymans × × 

Meruliporia incrassata × × × 

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Brown-rot fungi colonize the wood via the rays and spread in the longitudinal 

tissue through pits and by means of microhyphae. They grow inside 

the cell lumina (Fig. 7.1B) and there in close contact with the tertiary wall. 

The low-molecular agents and/or the cellulolytic enzymes penetrate through 

the relatively resistant tertiary wall (high lignin content) and diffuse into the 

secondary wall, where they degrade the carbohydrates completely (Fig. 7.1). 

Typically, brown-rot fungi do not cause lysis zones around their hyphae, while 

this is characteristic of many white-rot fungi. The hyphae are surrounded by 

slimelayers(Table2.1). 

In the early stages of decay, the carbohydrates are rapidly depolymerized. 

In Serpula lacrymans, the compression strength is decreased by 45% at only 

10% mass loss (Liese and Stamer 1934). Hemicellulose degradation runs up 

to about 20% mass loss faster than the respiration of the cleaving products. 

The relative lignin content increases parallel to carbohydrate degradation, the 

absolute lignin content slightly decreases. Due to the rapid cellulose depolymerization, 

the dimensional stability particularly decreases. The wood breaks 

up into rectangular blocks if it shrinks by drying (Fig. 7.1A), which led to the 

former term “destruction rot”. In some older literature, brown rot is falsely 

named as “red rot”, which however means the typical white-rot caused by Heterobasidion 

annosum.Inadvanceddecay,brown-rottenwoodcanbecrushed 

with one’s fingers to a brown powder (“lignin”). “House rot” means decay inside 

buildings, mostly by brown-rot fungi, particularly by Serpula lacrymans, 

Meruliporia incrassata, Coniophora species, Antrodia species, Donkioporia expansa 

(white rot) and Gloeophyllum species. There are further about 60 more 

rarely indoor occurring fungi (Table 8.6). 

7.2 

White Rot 

White-rot research has been reviewed by Ericksson et al. (1990) and Messner 

et al. (2003). White rot means the degradation of cellulose, hemicelluloses, 

and lignin usually by Basidiomycetes and rarely by Ascomycetes, e.g., 

Kretzschmaria deusta and Xylaria hypoxylon. White rot has been classified 

by macroscopic characteristics into white-pocket, white-mottled, and whitestringy, 

the different types being affected by the fungal species, wood species, 

and ecological conditions. From microscopic and ultrastructural investigations, 

two main types of white rot have been distinguished (Liese 1970). 

In the simultaneous white rot (“corrosion rot”), carbohydrates and lignin 

are almost uniformly degraded at the same time and at a similar rate during all 

decay stages. Typical fungi with simultaneous white rot are Fomes fomentarius, 

Phellinus igniarius, Phellinus robustus,andTrametes versicolor in standing 

trees and stored hardwoods (Blanchette 1984a). Wood decayed by F. fomen- 

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7.2 White Rot 139 

tarius, T. versicolor and some other fungi shows black demarcation lines (zone 

lines) (Fig. 7.2a), by which different species, or incompatible mycelia of the 

same species separate themselves from each other, or mycelia dissociate themselves 

from not yet colonized wood (“marble rot”, in German: “Marmorfäule”). 

The lines result from fungal phenol oxidases, whereby fungal compounds or 

also host-own substances are transformed to melanin (Li 1981; Butin 1995). 

As a function of the moisture distribution in wood, or between different fungal 

species or incompatible genotypes, a compartmentalization of individual 

decay centers can result from black pseudosclerotic layers of firmly structured 

mycelium (Rayner and Boddy 1988; Eriksson et al. 1990). 

Cell wall decay can start by microhyphae producing holes in the secondary 

wall (Schmid and Liese 1966), which flow together to larger wall openings with 

advancingdecay.Usually,however,thehyphaegrowinsidethelumenwithclose 

contact to the tertiary wall. The hypha surrounded by a slime layer (Table 2.1) 

excretes the degrading agents, which are active only in direct proximity of the 

hypha. Thus, a lysis zone develops under the hypha, and the hypha produces 

groovesinthewallwhichisgraduallyreducedinthickness,likearivererodes 

the ground (Schmid and Liese 1964; Liese 1970; Fig. 7.2b). 

Fig.7.2. White rot. a Simultaneous white rot by Trametes versicolor in beech wood with black 

demarcation lines. b Clamped hypha of T. versicolor digging into the cell wall (TEM, from 

Schmid and Liese 1964). c Successive white rot by Ganoderma adspersum in the Chilean 

“palo podrido” (photo J. Grinbergs). d White pocket rot (photo W. Liese) 

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In the successive (sequential) white rot, e.g., by Heterobasidion annosum 

(root rot in spruce), Xylobolus frustulatus (“Rebhuhnfäule” in standing and 

felled oaks: Otjen and Blanchette 1984, 1985), or in the Chilean “palo podrido” 

(Fig. 7.2c), lignin and hemicelluloses degradation run faster at least in 

early stages of attack, so that first cellulose relatively enriches. Further fungi 

showing successive white rot are e.g., Ceriporiopsis subvermispora, Dichomitus 

squalens, Inonotus dryophilus, andMerulius tremellosus. Frequently, e.g., 

by Phellinus pini (Liese 1970) in the heartwood of living conifers as well as 

by Bjerkandera adusta and some other fungi (Blanchette 1984a; Otjen et al. 

1987), there are small, elongated cavities within a wood tissue, where the 

lignin and also the hemicelluloses are “selectively” (preferentially) degraded 

(“selective white rot”, “selective delignification”, preferential white rot). The 

greatest part of the cellulose remains. These decayed regions are surrounded 

by tissue that appears sound (white pocket rot, honeycomb rot; Fig. 7.2d). 

With advancing decay, the wood becomes fibrous in texture by the decay of 

the more lignified middle lamella/primary wall area. Some Ganoderma species 

caused within a wood tissue as well white pocket rot as simultaneous rot, or, 

depending on the wood species, white pocket rot in birch and oak and simultaneous 

rot in poplar (Blanchette 1984a; Dill and Kraepelin 1986; Otjen and 

Blanchette 1986). 

The terms “selective white rot” and “selective delignification” have been 

propagated in the period of biopulping research (Chap. 9.3) as these terms 

promise more experimental success than would do names like successive white 

rot. As in most cases of “selective white rot” and particularly in late stages 

of attack, cellulose is also degraded to some extent, the term “preferential 

delignification” should be used. 

Many white-rot fungi, e.g., Heterobasidion annosum (Hartig 1874), Fomes 

fomentarius, Ganoderma species, and Trametes versicolor cause black spots of 

manganese dioxide deposits in the attacked wood (Blanchette 1984b; Erickson 

et al. 1990; Daniel and Bergman 1997). Manganese deposits may occur in 

connection with lignin degradation by manganese peroxidase. Physisporinus 

vitreus, isolated from cooling-tower wood (Schmidt et al. 1996) exhibited these 

manganese deposits predominantly in the slime layer and in the inner S2 

beneath a hypha shown by TEM/EDX spectra (Fig. 7.3B). 

White-rot fungi attack predominantly hardwoods, either as pioneer organisms 

or later in the context of a succession. As conifers are the main timbers 

used in the northern hemisphere for constructions, white-rot fungi occur there 

rarely in buildings. In Table 7.2, some important white-rot fungi are specified. 

In all white rot types, the wood strength properties are reduced to a lesser 

extent than in brown-rotten wood, since at the same mass loss, lesser cellulose 

is consumed, and it does not come to cracking or cubical rot. In a very late 

stage of attack, a wood mass loss of 97% has been measured. 

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7.2 White Rot 141 

Fig.7.3. Manganese deposits occurring during decay of a Scots pine sapwood block by 

Physisporinus vitreus. A Wood sample with black manganese deposits after culture (from 

Schmidt et al. 1996). B TEM-micrograph showing electron-dense material in the hyphal 

slime layer (c) and the secondary wall (b). C TEM/EDX spectra of manganese and other 

elements in different areas of the attacked wood (see B). a control from a healthy area within 

the S2, b spectrum from the S2 beneath a hypha, c spectrum from dense deposit material 

within the hyphal slime layer. The copper peaks result from the metal grids. (from Schmidt 

et al. 1997a) 

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Table 7.2. Some common white-rot fungi 

Fungus Predominant occurrence 

standing tree timber outdoors softwood hardwood 

Armillaria mellea × × × 

Donkioporia expansa indoor × × 

Fomes fomentarius × × 

Heterobasidion annosum × × 

Meripilus giganteus × × 

Phellinus pini × × 

Polyporus squamosus × × 

Schizophyllum commune × × 

Stereum sanguinolentum × × × 

Trametes versicolor × × 

7.3 

Soft Rot 

The term “soft rot” was originally used by Findlay and Savory (1954) to describe 

a specific type of wood decay caused by Ascomycetes and Deuteromycetes 

which typically produce chains of cavities within the S2 layer of soft- and 

hardwoods in terrestrial and aquatic environments (Liese 1955), for example 

when the wood-fill (Fig. 7.4a) in cooling towers became destroyed despite 

water saturation, and when poles broke, although they were protected against 

Basidiomycetes. About 300 species (Seehann et al. 1975) to some 1,600 examples 

of ascomycete and deuteromycete fungi (Eaton and Hale 1993) cause soft rot, 

e.g., Chaetomium globosum (Takahashi 1978), Humicola spp., Lecythophora 

hoffmannii, Monodictys putredinis, Paecilomyces spp., and Thielavia terrestris. 

Soft-rot fungi differ from brown-rot and white-rot Basidiomycetes by growing 

mainly inside the woody cell wall (Fig. 7.4b). The wood is colonized via 

the wood rays. In conifers, the fungi penetrate, starting from the tracheidal 

lumina, by means of thin perforation hyphae of less than 0.5µm thickness into 

the tertiary wall and re-orientate then as thin hyphae after L-bending in one 

direction or after T-branching in both directions along the microfibrils in the 

secondary wall (soft rot type 1, Nilsson 1976). 

In longitudinal wood sections, hyphal activity is recognizable in the polarized 

light by rhombus-shaped cavities in the secondary wall of different size and 

arrangement (Levy 1966; Butcher 1975), which may be lined up like a string 

of pearls (Fig. 7.4c): The thin hypha stops its growth and the cavity is then 

developed around the hypha by the release of enzymes (putatively endoglucanases) 

along what is described as the proboscis hypha. Within the cavity, 

hyphal thickness increases to about 5µm. From the tip of the cavity, the next 

fine hypha starts its growth, which results in the next cavity, and continuous 

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7.3 Soft Rot 143 

Fig.7.4. Soft rot. a Woodfill from a cooling tower. b Hole-shaped decay of the secondary 

walls in latewood tracheids of pine sapwood (LM, photo M. Rütze). c Cavities inside a fiber 

(LM, photo by W. Liese) 

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enlargement of existing cavities and the formation of new cavities lead to total 

destruction of the S2 layer (Eriksson et al. 1990; Daniel 2003). SEM and TEM 

showed that the hyphae are normally associated with a variety of granular and 

fibrillar materials including extracellular slime (Table 2.1), melanin and lignin 

breakdown products. In Lecythophora mutabilis, CCA was concentrated in the 

granular material (Daniel 2003). Several causes are discussed for oscillating 

hyphal growth and cavity formation (Table 7.3). 

In cross sections, the cavities appear hole-shaped (“initial stage”) and increase 

with advancing decay to larger wall openings (Fig. 7.4b). Finally, it 

comes to circular detaching of the tertiary wall (“advanced stage”). Because of 

their high lignin content, the tertiary and primary walls are attacked in the end 

(“late stage”). It remains an incomplete skeleton of middle lamella/primary 

walls (“destruction stage”). 

In the soft rot type 2, which particularly occurs in hardwood (Zabel et al. 

1991), the hyphae erode particularly from the lumen the tertiary wall and 

penetrate till the middle lamella/primary wall. As rare variant, diffuse and 

irregular cavities in the secondary wall were described (Anagnost et al. 1994). 

Soft rot develops also in monocotyledons (bamboos: Liese 1959; Sulaiman 

and Murphy 1992). In a broader definition for soft rot, each significant fungal 

decay of the woody cell wall by non-basidiomycete fungi was suggested, which 

however contrasts to the white-rot causing Ascomycetes. 

Since the tertiary and middle lamella/primary wall are resistant over longer 

time against fungal attack due to stronger lignification (Fig. 7.4b), wood with 

soft rot frequently first will not be recognized with the naked eye. Also with 

the “hammer test” it does not result in the hollow sound of decayed wood 

(Liese 1959), so that in former times during repair work of poles accidents 

arose several times by pole breaks due to the unawareness of the officials. Soft 

rot penetrates slowly from the outside to the wood center. Moist wood is dark 

colored and the surface is soft. Although softening of wet wood is typical, 

attacked CCA treated timber has shown degraded wood to be hard. The dry 

wood shows cubical rot with a fine-cracked, charcoal-like surface (Fig. 7.4a). 

Table 7.3. Possible factors involved in cavity formation by soft-rot fungi 

– chemical and morphological structure of the wood cell wall (Liese 1964) 

– accumulation of toxic phenolic substances from lignin degradation (Liese 1970) 

– oscillating cellulase activity as reaction to the produced sugars (Nilsson 1974) 

– unequally distributed chemical factor of the carbohydrates in the cell wall 

(Nilsson 1982) 

– composition and distribution of lignin in the cell wall 

– nutrients obtained by cavity formation allowing only limited growth of a hypha 

– small channel between two cavities due to intense enzyme production at the 

hyphal apex and less enzyme production at the hyphal basis (Eriksson et al. 1990) 

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7.3 Soft Rot 145 

Further infection symptoms are the blunt fracture and short-fibrous breaking 

out of splinters when puncturing. 

Within the cell wall, soft-rot fungi degrade cellulose and hemicelluloses. 

Compared to the brown-rot fungi the cellulolytic agents diffuse, however, not 

sodeepintothecellwall,butremainindirectproximityofthehyphae(Liese 

1964). Lignin is not (or little) attacked at least in the initial stage, mainly 

by demethylation, so that soft rot with regard to the decay type resembles 

brownrot.IsolatedligninsandDHP’sarenotdemethylated.Inligninmodel 

compounds, the β-O4 linkage and the aromatic ring were cleaved (Eriksson 

et al. 1990; also Bauch et al. 1976). 

The inhibiting effect of lignin was demonstrated by the result that a delignifying 

pretreatment promoted the carbohydrate degradation (Zainal 1976). 

Wood decay by soft-rot fungi is further affected by the quantity and type of the 

lignin: Lignin-rich softwood with lignin predominantly made of coniferyl units 

is more resistant than the lignin-poorer hardwood made of sinapyl-coniferyl 

units (Nilsson et al. 1988; Eriksson et al. 1990). In conifers, wood decay occurs 

preferentially in the late wood (Fig. 7.4b) with its relative low lignin and high 

cellulose content. 

Due to the intensive carbohydrate degradation, soft-rot fungi, just like 

brown-rot fungi, already cause about 50% decrease of impact bending at only 

5% mass loss, and cracks occur by the reduction of the dimensional stability. 

Soft rot develops in trees, stored wood, and in outside used wood. Soft-rot 

fungi can decay wood under extreme ecological conditions, which are unsuitable 

for Basidiomycetes: constantly wet wood till almost water saturation, like 

in harbor constructions and ships, but not permanently submerged, as well 

as wood in soil contact, like poles, piles, sleepers (Liese 1959). Several soft-rot 

fungi were found on rotting branches (Butin and Kowalski 1992). Soft-rot fungi 

(and Basidiomycetes) under marine conditions were described by Kohlmeyer 

(1977), Leightley and Eaton (1980) and Troya et al. (1991). The wood moisture 

tolerance of the fungi reaches from dryness resistance to decay at almost water 

saturation. For example, Chaetomium globosum and Paecilomyces spp. did not 

show any inhibition of their decay ability in beech wood samples of 200% 

wood moisture content (Liese and Ammer 1964). With altogether relatively 

low oxygen demand, soft-rot fungi receive the necessary oxygen for the decay 

of water-saturated wood in cooling towers by the sprinkling effect of the water, 

which brings oxygen in solution. Thermophilic species and those with the ability 

of heat resistance destroy wood in the inner of wood chip piles (Hajny 1966; 

Smith 1975). Chaetomium globosum canstartgrowinginnutrientsolutions 

with initial pH values from 3 to 11. Some soft-rot fungi decay woods with high 

natural durability, like Bongossi or Teak. After 21 years of outdoor exposure 

in soil, the heartwood of several hardwoods exhibited soft rot in about threequarters 

of all the samples, about one-quarter white rot and only 3% brown 

rot (Johnson and Thornton 1991). Soft-rot fungi are tolerant to chrome fluo- 

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rine salts, which inhibit brown and white-rot fungi, but are sensitive to copper 

(Chap. 7.4). Wood in soil contact must therefore be treated with a preservative 

that contains copper if coal tar oil is not applied. Large economic losses developed 

nevertheless in Australia when hundreds of thousands of eucalyptus 

poles, which were treated with chrome copper arsenic, prematurely failed by 

soft rot due to unequal preservative distribution in the wood (Dickinson et al. 

1976; Liese and Peters 1977; Greaves and Nilsson 1982). Several soft-rot fungi 

were isolated from CCA treated (Zabel et al. 1991; Wong et al. 1992) and coal 

tar oil-impregnated poles (Lopez et al. 1990; Dickinson et al. 1992). 

7.4 

Protection 

This chapter focuses on fundamentals upon prevention of wood damage by 

fungi, and protection and preservation of wood (e.g., Willeitner and Liese 1992; 

Eaton and Hale 1993; Palfreyman et al. 1996; Murphy and Dickinson 1997; 

Zujest 2003; Goodell et al. 2003; Müller 2005). Protection in the broader sense 

comprises non-chemical methods like organizational measures and measures 

by design, use of naturally durable woods, application of antagonisms, or wood 

modifications that do not affect the environment. Preservation predominantly 

stands for chemical measures. 

Table 7.4 shows the conditions for the development of wood fungi and 

protection principles that can be deduced from them. 

The principle of the wood protection consists of changing at least one of the 

three life prerequisites of fungi in wood in such a way that the development 

of fungi is impossible or at least inhibited. Fungal attack can be prevented 

Table 7.4. Prerequisites for the development of wood fungi and principles of protection 

deduced from them (supplemented from Willeitner and Schwab 1981) 

Prerequisite Preventive measure Protection principle 

Suitable moisture Reduce, keep away Timber drying, 

constructional wood protection, 

wood modification 

Suitable food Make inedible Use of durable wood, 

chemical wood preservation, 

wood modification, 

(use of antagonisms) 

Sufficient oxygen Keep away Drying, wet storage, 

storage in CO2/N2 atmosphere, 

use below the water level 

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(Willeitner and Schwab 1981; Erler 2002; Willeitner 2000, 2003; Goodell et al. 

2003; Böttcher 2005; Borsch-Laaks 2005; Schmidt 2005) by: 

– organizational protection (e.g., short and appropriate wood storage), 

– use of durable wood species (natural methods), 

– keeping away water by structural wood protection measures by design: 

appropriate surface and weather protection, use of vapor barriers, avoidance 

of condensation due to thermal insulations, salient roof to protect timber 

from rain, drawing off of rain, barrier to avoid direct contact between wood 

and adjacent material, or inside the wall against raise of moisture from the 

ground, 

– chemical wood preservation, 

– wood modifications that increase dimensional stability of wood, reduce 

uptake of moisture, or make it hard to digest, 

– use of antagonisms. 

The moisture conditions in wood are of decisive importance for the development 

of wood fungi (Chap. 3.3). Table 7.5 shows the hazard classes of wood 

[to be replaced by “use classes” according to prEN 335-1 (2004) respectively 

ISO] that depend on wood use and timber moisture according to the German 

standard DIN 68800, parts 2 and 3 (1990, 1996), the corresponding potential 

application of durable timber, and the minimum requirements of chemical 

preservation measures. 

Natural durability means the wood-own resistance against bacteria, wooddecay 

fungi, beetles, termites and marine borers, which will differ for a timber 

species against the various organisms. Wood durability is based on the presence 

of accessory compounds, whereby it concerns numerous compounds from 

different chemical classes (Fengel and Wegener 1989; Obst 1998). They are produced 

in the living tree during transition from the sapwood to the heartwood 

and are deposited in the heartwood (Taylor et al. 2002). Thus only the heartwood 

exhibits natural durability, while the sapwood of all wood species is only 

little or not durable. The European standard EN 350-2 (1994) uses a five-class 

system to group 128 timbers according to their durability against fungi. Wood 

with high durability against fungi (durability class 1, very durable) is e.g., 

greenheart (durable also against termites and marine organisms). European 

oak is durable (class 2), walnut is moderately durable (class 3), Norway spruce 

is slightly durable (class 4), and European beech not durable (class 5) (also Augusta 

and Rapp 2003, 2005; Willeitner 2005a). Natural durability of some bamboo 

species against four decay fungi was investigated by Remadevi et al. (2005). 

The influence of the felling time on resistance is controversially discussed. 

It has to be considered that fresh winter-felled wood is less susceptible to 

damage due to other moisture, drying, and climatic conditions than wood 

felled in the summer. There are however no differences if the wood is carefully 

dried (Willeitner 2005a). 

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Table 7.5. Hazard classes of timber, conditions for wood use, resistant wood species, and 

chemical preservation measure 

Hazard condition for wood use durable wood minimum 

class preservative measure 

0 indoors, if 

wood moisture ≤ 12%, 

timber open at 3 sides or 

coating against insects 

none 

1 indoors colored heartwoods prevention of insects 

air humidity ≤ 70%, sapwood proportion < 10% 

wood moisture < 20% 

2 indoors: colored heartwoods prevention of 

air humidity > 70% of durability class fungi and insects 

in wet areas: 1, 2 or 3 

water-repellent coating 

outdoors: without 

weathering 

3 outdoors: weathered colored heartwoods prevention of 

without permanent of durability class fungi and insects, 

ground or water contact 1 and 2 weatherproof 

indoors: wet rooms 

4 permanent ground colored heartwoods prevention of 

or fresh water contact, of durability class 1 fungi and insects, 

special prerequisites weatherproof, 

for cooling towers and prevention of 

marine timber soft-rot fungi 

There is still a worldwide spread superstition that wood properties like 

resistance against fungi depend on the moon. The wood of trees felled at 

a certain date related to the moon phase is thought not to swell nor shrink, 

to be incombustible, resistant to fungi and insects, and to become very hard. 

Those oscillating changes of the properties of the woody tissue, which mainly 

consists of dead fiber or tracheid cell walls, are biologically impossible. Thus, 

all specifications are in contradiction to scientifically based results (Wa˙zny and 

Krajewski 1984; Seeling 2000; bamboo: Yamamoto et al. 2005). The positive 

effects of a certain felling date observed in the practice may be due to other 

influences: People which believe in lunar influences select in the forest wellgrown 

trees, use appropriate drying, storage methods and wood design, that 

is, the wood, its processing and use are of high quality and thus the wood is 

more resistant to deterioration. 

There are several standards to determine the resistance of untreated wood 

and wood-based composites against fungi and also to test the efficacy of 

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preservatives. In Europe, the standards are ruled by the European Committee 

for Standardization (Table 7.6; Willeitner 2005b). 

Figure 7.5 shows a Kolle flask that is used according to the European standard 

EN 113 to determine the toxic values of wood preservatives against wooddestroying 

Basidiomycetes cultured on agar medium. The method can be also 

used to test the natural durability of timber species etc. 

Chemical wood preservation is used, if structural-constructive measures, or 

natural durability, or wood modifications alone are insufficient for an increased 

wood endangering to meet the requirement of long-term use of wood. Not 

durable wood species or those of insufficient durability and the sapwood 

of all wood species can be made resistant for a long time against damage 

by treatment with appropriate wood preservatives, provided that the wood 

shows permeability. Prerequisite is, corresponding to the wood use, to bring 

effective formulations in sufficient amount deeply into the wood (Schoknecht 

and Bergmann 2000) using appropriate methods (Willeitner and Schwab 1981; 

Table 7.6. European standards that deal with resistance and preservation of wood against 

fungi 

EN 335 (1992/95) Durability of wood and wood-based products; Definition of 

hazard classes of biological attack (3 parts) 

EN 350 (1994, 2 parts), EN 460 (1994) Durability of wood and wood-based products – 

Naturaldurabilityofsolidwood 

ENV 12038 (1996) Durability of wood and wood-based products – Wood-based panels 

ENV 12404 (1997) Durability of wood and wood-based products – Assessment of the 

effectiveness of a masonry fungicide to prevent growth into wood of Dry rot Serpula 

lacrymans (Schumacher ex Fries) F.S. Grey 

EN 113 (1996) Determination of toxic values of wood preservatives against wood 

destroying Basidiomycetes cultured on agar medium 

EN 152 (1989) Test methods for wood preservatives; Laboratory method for 

determining the protective effectiveness of a preservative treatment against blue 

stain in service (2 parts) 

EN 252 (1990) Field test method for determining the relative protective effectiveness of 

a wood preservative in ground contact 

EN 330 (1993) Wood preservatives; Field test for determining the relative protective 

effectiveness of a wood preservative for use under a coating and exposed out of 

ground contact: L-joint method 

ENV (prestandard) 807 (2001) Wood preservatives – Determination of the effectiveness 

against soft rotting micro-fungi and other soil-inhabiting micro-organisms 

ENV 839 (2002) Wood preservatives – Determination of the effectiveness against wood 

destroying Basidiomycetes – Application by surface treatment 

ENV 12037 (1996) Wood preservatives – Field test method for determining the relative 

protective effectiveness of a wood preservative exposed out of ground contact 

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Fig.7.5. Kolle flask according to EN 

113 to determine the toxic values of 

wood preservatives against wooddestroying 

Basidiomycetes cultured 

on agar medium. a Non-impregnated 

control. b Impregnated sample 

Tables 7.9, 7.10). It is distinguished between “preventive wood preservation” 

(Kleist 2005) and “controlling wood preservation” after a damage (Sallmann 

2005). 

There are different national regulations with regard to testing, approval, 

application and toxicological aspects of chemical wood preservatives. Thus, 

the following only describes the German situation (Fischer 2005; Reifenstein 

2005). 

The official approval of wood preservatives used for load-bearing construction 

takes place by the “Deutsches Institut für Bautechnik (DIBt)”, which evaluates 

the results of tests that had been performed in view of minimum requirements 

(efficacy, no unfavorable side effects). The Federal Institute for Risk Assessment 

(BfR) evaluates hygienic-toxicologic aspects of the preservative and 

the Federal Environmental Office (UBA) its ecotoxicologic behavior. Preservatives 

with approval obtain a general national approval by the DIBt. Important 

characteristics of a wood preservative are described by test ratings (Table 7.7). 

About 95% of the mainly professionally used preservation salts possess the 

DIBt approval (23% share of the market), while only 10% of the predominantly 

solvent-based preservatives that are used by do-it-yourselfers have been previously 

proven by neutral boards. 

Wood preservatives for non-load-bearing constructions can receive a quality 

mark by the RAL Quality Community of Wood Preservatives, including an 

evaluation by BfR and UBA. 

For blue stain-preventing preservatives for timber outdoors without ground 

contact including windows and outside joinery, a registration process concern- 

Table 7.7. Test ratings of wood preservatives in view of efficacy 

P prevention of fungi 

Iv prevention of insects 

Ib control of insects 

W for weathered wood without permanent soil or water contact 

E for wood in permanent soil or water contact or with dirt deposits in joints 

M preventionofSerpula lacrymans to grow through brickwork 

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ing efficacy and toxicology is possible by the German Association of Varnish 

Industry (VdL) on a voluntary basis. 

All wood preservatives with DIBt approval, RAL quality mark, and the 

VdL-blue stain preventing preservatives are listed and specified by the active 

components in the annual wood preservative register (Deutsches Institut für 

Bautechnik 2005; Table 7.8). There is also a consumer guide on wood preservatives 

by the German Federal Ministry of Consumer Protection, Food and 

Agriculture (2003). 

Water-based boron salts without chromate are only suitable for inside use 

due to their leachability (Peylo and Willeitner 1995, 2001). Wood preservatives 

based on protein borates greatly retarded the leaching of boron from 

treated timber (Thevenon et al. 1998). In chromate-containing salt mixtures, 

the biocides are fixed to the wood tissue (Bull 2001). By the fixation process, 

the hexavalent chromium (Cr VI ) is reduced by wood components to the trivalent 

less toxic Cr III . This helps to stabilize the other preservative components 

in the wood (Bao et al. 2005b), in different degrees, e.g., copper is almost 

completely fixed. Therefore, those mixtures are also suitable for outside use. 

Chrome fluorine boron salts (CFB) are suitable for inside and outdoor use 

without ground or permanent water contact. Chrome copper salts (CC) are 

Table 7.8. Major groups of wood preservatives for prevention and control of decay fungi and 

insects (based on Deutsches Institut für Bautechnik 2005) 

DIBt approval-preservatives: 

for prevention: 

water-based preservatives 

boron, CFB, CC, CCA, CCB, CCF salts 

quaternary ammonium compounds 

quaternary ammonium-boron compounds 

chromium-free copper compounds (Cu-HDO, Cu-quaternary ammonium, 

Cu-triazol) 

various other compounds 

solvent-based preservatives (e.g., Al-HDO, pyrethroides) 

solvent-based and water-soluble preservatives (only insects, carbamates) 

coal tar oil distillates (creosotes) 

special compounds for wood-based composites (only fungi, anorganic boron 

compounds, K-fluorides, K-HDO) 

for control: 

water-based and solvent-based preservatives to control insects 

boron compounds, quaternary ammonium compounds, carbamates 

to prevent growth of Serpula lacrymans through masonry 

RAL quality mark-preservatives to prevent blue stain, decay fungi, insects, and termites, 

to control insects, and to protect masonry against S. lacrymans 

Blue stain preventing primers according to VdL instructions 

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allowed for indoor and outdoor wood, especially if it is exposed to leaching, 

and also for wood in ground contact, e.g., poles, exterior structures, such as 

decks and fences, mine timber (Narayanappa 2005), and wood in permanent 

water contact, such as cooling towers and marine works. Chrome copper salts 

with the addition of either boron (CCB) or fluorine (CCF) may be used indoors 

and outdoors. 

Chrome copper arsenic salts (CCA) are restricted to outdoor use and certain 

application such as noise barriers (Commission Directive 2003/2/EC 2003). 

In the USA and Canada, industry registrants voluntary agreed to withdraw 

CCA treatment for use in such residential applications as decks, fences, and 

playground components effective as of 2004, although it is still registered for 

commercial/industrial products (Bao et al. 2005b). Component leaching from 

CCA-treated wood during above-ground exposure was affected by climatic 

variables like precipitation and temperature (Taylor and Cooper 2005). Bao 

et al. (2005b) showed for CCA, CCB and acid copper chromate (retention about 

7kg/m 3 ) fixation times of 8–32 days at 21 ◦ C and between 12 and 48 h at 50 ◦ C. 

Treatment of freshly impregnated wood with hot steam of 110 ◦ C for 1 h was 

also suitable for sufficient fixation (Peek and Willeitner 1981, 1984; also Cooper 

and Ung 1992). Timber treated with water-based fixing salts should thus be 

protected from rain, depending on the type of preservative, to avoid leaching of 

the not yet fixed components, which would decrease the protection and pollute 

the environment. Cookson et al. (1998) evaluated the fungicidal effectiveness 

of water-repellent CCAs. 

Molybdenum and tungsten have been studied as substitutes for arsenic in 

CC-salts (Cowan and Banerjee 2005). Schultz et al. (2005a) used a mixture of 

copper(II) and oxine copper for an outdoor ground-contact exposure. 

Toxicological aspects have lead to an increased use of chromium-free preservatives 

that are just as fixing. These preservatives are based e.g., on ACQ (alkaline 

copper quaternary ammonium salts), copper HDO [bis-(N-cyclohexyldiazeniumdioxy)-copper] 

and Cu-triazoles. Some of these products also include 

boron. Quaternary ammonium compounds are used as N-dimethylalkylbenzylammoniumchloride, 

didecylpoly(ethox)ethylammoniumborate (polymeric 

Betain), and N,N-didecyl-N-methyl-poly-(oxethyl)-ammoniumpropionate. 

Zabielska-Matejuk et al. (2004) showed antifungal activity of bis-quaternary 

ammonium and bis-imidazolium chlorides (also Pernak et al. 1998). 

Didecyldimethylammoniumtetrafluoroborate inhibited mold and stain (Kartal 

et al. 2005a) and decay fungi (Kartal et al. 2005b). Copper(II) octanoate/ 

ethanolamines were investigated by Humar et al. (2003). Mazela et al. (2005) 

used copper monoethanolamine complexes with quaternary ammonium compounds. 

The group of triazoles as wood preservatives was treated by Wüstenhöfer 

et al. (1993). 

Solvent-based preservatives contain e.g., Al-HDO [tris (N-cyclohexyl-diazeniumdioxy)-aluminum] 

or triazoles (e.g., tebuconazole, propiconazole) as 

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fungicides and pyrethroides as insecticide. Pentachlorophenol is totally banned 

and γ-hexachlorocyclohexane (lindane) is not used any more. The addition 

of emulsifiers enables the use of various organic substances as emulsions in 

water-based systems. 

Distillates of coal tar oil (creosote) are a complex mixture of some hundred 

compounds, mainly polycyclic aromatic hydrocarbons (Majcherczyk and 

Hüttermann 1998), and are allowed for outdoor use of timber in permanent 

ground and water contact that is not in contact with human beings. Preferably, 

creosotes are applied to wood that is exposed to leaching, e.g., railroad sleepers 

and telegraph poles. According to legislation, the content of benzo(a)pyren 

is limited to 50 ppm, classified by the Western-European Institute for Wood 

Preservation as type WEI B and C. 

Boron compounds, quaternary ammonium compounds, and carbamates are 

suitable to prevent growth of Serpula lacrymans through masonry. 

For the protective effect, the moisture content, the type of preservative (Table 

7.8), the treatment process (Tables 7.9, 7.10), and the duration of treatment 

have to be considered. In addition, the timber species and part of the timber 

determine the permeability for preservatives. The treatability (EN 350-2) 

varies between completely permeable (class 1: easy to treat like the sapwood of 

Quercus robur and Pinus sylvestris) to extremely refractory (class 4: extremely 

Table 7.9. Applicability of wood preservatives and treating processes depending upon the 

wood moisture content (modified from Willeitner and Liese 1992) 

Moisture Wood Treating process 

content preservative 

water- creosote organic 

based solventbased 

green 

much above 

yes no no sap-displacement (diffusion) 

fiber saturation 

markedly above 

yes no no diffusion, long-term soaking 


slightly above 

yes no no diffusion, long-term soaking, 

simple methods, OPM 


below 

yes no (yes) soaking, simple methods, 

(diffusion), (pressure processes) 

fiber saturation yes yes yes pressure processes (except OPM), 

soaking, simple methods 

underlined preferably recommended, in brackets possible but not recommended, 

OPM oscillating pressure method 

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Table 7.10. Major groups of application procedures of wood preservatives (after Willeitner 

and Schwab 1981; Willeitner and Liese 1992) 

Pressure processes, which use intervals of any difference of pressure until 0.8 N/mm 2 

(including or excluding vacuum) and which follow different treatment schedules, 

dependent on preservative and timber, yield deep penetration and high retention. 

In long-term procedures, the timber or the part of it to be treated is kept completely 

immersed (soaking) in the preservative, which slowly (some days) penetrates into 

the wood. 

Short-term procedures (dipping, spraying, deluging, brushing) yield only little 

penetration (surface treatment) and low retention. 

Special procedures preserve timber in use, like bored-holes in constructional timber, 

pilings or poles, and pastes or bandages used as a ground-line treatment for poles 

difficult to treat like the heartwood of Q. robur). The role of bordered pits to 

the refractory nature of softwoods has been reviewed by Usta (2005). 

The penetrability of refractory timbers like Picea abies (class 3–4) can 

be improved by using oscillating pressure methods (Breyne et al. 2000), by 

incising methods or by ponding. During the latter, bacteria attack the pits, 

but only irregularly and only in the sapwood. There were attempts to pretreat 

wood with chemicals (Militz and Homan 1992) and with enzymes to improve 

the permeability of conifers (Adolf 1975; Militz 1993) and hardwoods (Knigge 

1985). Jagadeesh et al. (2005) improved penetration and retention of CCA in 

bamboo by using shockwaves. 

Therearedifferentpreconditionsforchemical treatmentofwood(Willeitner 

and Liese 1992): All wood must be debarked and free from phloem remainings 

(“white-peeled”) before treatment. An exception is sap-displacement treatments 

(Boucherie process), where a water-based preservative is introduced 

under low pressure at the butt end of freshly felled trees replacing the sap of 

the sapwood by the preservative. Sap-displacement is out of use now for wood, 

but is applied to bamboo culms, whereby boron-salts are most commonly and 

successfully used (Liese and Kumar 2003). When the water content of wood is 

much above fiber saturation, a water-based preservative either of a high concentration 

or by long-term soaking in its solution can be used to distribute the 

chemicals into the timber by diffusion. Chemical treatment of seasoned wood 

with moisture content below fiber saturation requires penetration of a liquid 

into the capillary structure of the wood and subsequent distribution into the 

wooden tissue. Movement is effected either by externally applied pressure or 

by internal capillary forces. 

Before treatment, working the timber like final cross cuttings, sawing, borings 

and shapings should be completed. Otherwise, the newly exposed part of 

the timber has to be treated once more, e.g., by brushing it several times. This 

applies also to later developing drying shakes if the wood was insufficiently 

dried before treatment. 

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Table 7.9 shows the applicability of wood preservatives and treating processes 

depending upon the wood moisture content. 

For pressure treatments [full-cell process, empty-cell process (Lowryprocess, 

Rüping-process), vacuum-process] and principally for creosote, the 

moisture content should be below fiber saturation. For short-term procedures 

(superficial treatments) and also for water-based preservatives, at least the 

wood surface must begin to dry. To bring the active substances into the wood, 

the procedures may be arranged into four major groups (Table 7.10). 

In Germany, there are 234 pressure plants and 2,115 plants that use soaking 

(Quitt 2005). The necessary retention of a preservative depends on the endangerment 

of the wood, on the efficacy of the active ingredient, and the treatment 

procedure. The minimum quantities are shown in the respective DIBt approval. 

The preventing chemical preservation of wood-based composites is regulated 

in DIN 68800 part 5 (1978). 

Wood plastic composites (WPCs) are a new material with plastic as a matrix 

and embedded wood particles and fibers as well as distinctive additives 

(Teischinger et al. 2005). The material is produced. e.g., by an extrusion process 

or injection molding process. Wood is added for better technical properties 

and for cost reduction. WPCs are increasingly used as a substitute for wooden 

decks especially in North America. Marketing of WPC products as “maintenance 

free” has been a key factor contributing to their success with the 

consumer. WPCs are nevertheless susceptible to fungal degradation despite 

the close association of wood with the plastic. Wood particles close to the surface 

of WPC products can attain moisture levels high enough to facilitate the 

onset of decay. Borates markedly reduced mass loss of WPC by Gloeophyllum 

trabeum in a soil block test (Simonsen et al. 2004). Mankowski et al. (2005) 

showed almost no mass loss by G. trabeum and Trametes versicolor in samples 

that had been treated with zinc borate. 

The non-chemical protection and chemical preservation of bamboo are 

described by Liese (2002) and Liese and Kumar (2003). 

Methods to determine the amount of active substances in the wood and 

to measure penetration depth are described by Petrowitz and Kottlors (1992), 

Schoknecht et al. (1998) and Schoknecht and Bergmann (2000). An overview 

is in the Internet (www.holzfragen.de/seiten/hsm reagenzien.html). N-cyclohexyl-diazeniumdioxide 

in impregnated pine wood was measured by direct 

thermal desorption-gas chromatography-mass spectrometry (Jüngel et al. 

2002). 

Since about 1975 critical reports increase with regard of possible environmental 

impacts by chemical wood preservation, like by pentachlorophenol and 

chromate-containing preparations, the pollution of the soil by leached chemicals 

(Willeitner 1973; Willeitner et al. 1991; Leiße 1992; Hartford 1993) and 

due to problems arising from the disposal of treated timber (Marutzky 1990; 

Voß and Willeitner 1992). Pentachlorophenol (PCP) has protected wood since 

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1935 from staining and from decay by fungi and insects (Prewitt et al. 2003). 

The life expectancy of utility poles increased from approximately 7 years in 

an untreated pole to about 35 years in a treated pole, thereby saving utility 

companies millions of dollars in replacement costs. In the USA, 36 million 

PCP-treated poles have been estimated to be in service in 1990. In view of the 

negative impact on humans, animals, plants, and the environment, utilization 

of PCP and import of PCP treated woods are however restricted in Germany. 

Disposal of spent treated wood has increasingly become a major concern. 

Popular methods, such as burning (incineration, combustion) and land filling, 

are costly or even impractical because of increasingly strict regulatory requirements. 

Recycling of the preserved wood and removal of the toxic preservatives 

from the treated wood is of great importance. Research in this area (Lin and 

Hse 2005) focus on direct recycling of preserved wood into composite manufacturing, 

CCA removal from spent CCA-treated wood performed by lowtemperature 

pyrolysis, solvent extraction, hydrogen peroxide extraction (Kim 

et al. 2004), electrodialytic remediation (Christensen et al. 2005), biological 

remediation, and dual treatment processes involving biological remediation 

and chemical extraction. Li and Hse (2005) liquefied CCA-treated wood in 

polyethylene glycol and removed more than 90% of the metals by precipitation 

from aqueous solvents. Kartal and Imamura (2005) used chitin and chitosan 

for remediation of CCA-treated wood. Studies on bioremediation, particularly 

creosote, DDT, lindane and PCP, used several bacteria and fungi (review by 

Majcherczyk and Hüttermann 1998). Fungi which excrete high amounts of oxalic 

acid and are copper tolerant like Antrodia vaillantii (Collett 1992a, 1992b; 

Schmidt 1995b) have been used to bio-recycle CCA and CCB treated wood 

(Leithoff et al. 1995; Stephan et al. 1996; Kartal and Imamura 2003; Samuel 

et al. 2003; Humar et al. 2004; Kartal et al. 2004). Clausen (1997b) enhanced 

CCA removal from treated wood by Bacillus licheniformis (Weigmann) Chester. 

There is a great bulk of investigations on new, alternative wood protection 

procedures that deal with the chemical and/or physical modification of wood 

(e.g., Militz and Krause 2003). Rapp and Müller (2005) grouped the recent wood 

protection procedures that are already used or are expected to be used into 

wood modification, wood hydrophobization, and supercritical fluid treatment. 

Wood modification comprises various treatments that decrease the swelling 

of the woody cell wall and thus its accessibility for the fungal degradation 

agents. 

Reactive organic compounds like acetic anhydride (“acetylation”) are introduced 

in the wood (Hill et al. 1998), which react with the hydroxyl groups of 

the cell wall polymers and thus increase the dimensional stability of the wood 

as well as its resistance against decay and discoloring fungi. Acetylation with 

acetic anhydride results in covalently bonded acetyl groups (“plugging of hydroxyl 

groups”) in the wood and acetic acid as a by-product. Acetylated wood 

is non-toxic and has no harmful impact on the environment, but may have an 

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unpleasant smell. Stake tests according to EN 252 with acetylated pine wood 

samples showed that the resistance of samples with an acetyl content of about 

20% equals that of CCA treated wood with 10 kg/m 3 retention (Larsson Brelid 

et al. 2000). Brown-rot decay became zero at a weight percent gain (WPG) of 

about 20% due to acetylation, and white-rot was prevented at a WPG of about 

12% (Ohkoshi et al. 1999). As other anhydrides, propionic, butyric and hexanoic 

anhydrides were tested against brown, white and soft-rot fungi (Suttie 

et al. 1999; Papadopoulus 2004). Several carboxylic acid anhydrides were used 

for pine sapwood (Dawson et al. 1999). 

Impregnation with melamine resins leads to a deposition of the resin in the 

cell wall (Rapp et al. 1999) and there to the “blockade of hydroxyl groups” 

without chemical linkage, which also improves the mechanical properties and 

durability of wood (Rapp and Peek 1996; Lukowsky et al. 1999). 

Impregnation with 1,3-dimethylol-4,5-dihydroxyethylen urea (DMDHEU) 

effects the “linking-up of neighbored hydroxyl groups” by etherification with 

the N-methylol groups (Rapp and Müller 2005). There was no significant 

weight loss by Trametes versicolor of beech wood samples with 25% WPG of 

DMDHEU (Verma et al. 2005). 

There are various methods to produce thermally modified timber (“thermal 

modification of wood”) which leads to improved dimensional stability 

(Tjeerdsma et al. 1998) and biological resistance, but also partial wood degradation 

and discoloration. The processes have in common that the wood is 

subjected to temperatures between 160 and 260 ◦ Cinanatmospherewithlow 

oxygen content (Leithoff and Peek 1998; Rapp 2001; Ewert and Scheiding 2005). 

Potentially toxic byproducts have been considered by Kamdem et al. (2000). In 

Europe, about 45,000 m 3 of thermally modified timber were produced in 2004. 

Four basic technologies have been established: the Finnish “Thermo wood”, 

the Dutch “Plato wood”, the French “Retification”. Heat is transferred to the 

wood in the gas phase of air, exhaust fumes of combustion gases or nitrogen. 

The German “oil heat treatment” uses a vegetable oil (rape) for heat transfer, 

which additionally affects hydrophobization (Sailer et al. 2000; Bächle et al. 

2004). The wood is used outdoors, e.g., for façade covering, noise barriers, 

and in gardens for decks, and indoors, e.g., for floorings. Four years lasting 

field tests of wood samples from the four European industrial heat treatment 

processes indicated that heat treated wood appears to be not suitable for in 

ground application, since only durability classes in the range from 2 (durable) 

to 4 (slightly durable) were achieved (Welzbacher and Rapp 2005). Thermalhygro-mechanically 

densified wood showed reduced hygroscopy and improved 

mechanical performance, and resistance to fungal degradation (Schwarze and 

Spycher 2005). 

Wood hydrophobization occurs by oils, waxes, paraffins, and silicons. Sailer 

(2001) and Rapp et al. (2005) used vegetable oils. The oil, which is deposited in 

the cell lumina, reduces water uptake without inhibiting vapor release. A wax- 

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type end coating of logs considerably reduced stain and checking (Linars- 

Hernandez and Wengert 1997). Hill et al. (2004) impregnated the wood cell 

wall with silane monomers, which polymerize in situ. Furuno and Imamura 

(1998) used sodium silicate-boron. The use of organic silicon compounds 

was reviewed by Mai and Militz (2004). Both hydrophobization and increased 

wood density were obtained in doubly modified wood samples when the wood 

was treated with reagents bearing isocyanate, carboxylic anhydride or oxirane 

functionstoinducereactionswiththeOHgroupsandwhenthereagentsalso 

carried a polymerisable function by incorporating a monomer (styrene or 

methyl methacrylate) into the wood (Bach et al. 2005). 

Supercritical fluid treatments use the principle that the preservative carrier 

e.g., CO2 possesses at a certain pressure and temperature at the same time the 

properties of a gas and a liquid. The effective substance is similarly well soluble 

as in an organic solvent, but the penetration into the wood is deeper due to the 

minimal surface tension. At the end of the treatment, the carrier regains the 

gas phase by falling below the supercritical point that is the carrier loses the 

dissolving ability for the preservative, which remains deposited in the wood, 

and leaves the wood. Due to the minimal swelling of the wood, supercritical 

fluid treatment is particularly suitable for size-constant components like windows 

and doors (Rapp and Müller 2005). Morrell et al. (2005) impregnated 

wood-based composites with tebuconazole using supercritical carbon dioxide. 

Chitosan, a linear copolymer of β(1-4)-linked 2-amino-2-deoxy-D-glucopyranose 

and 2-acetoamido-2-deoxy-D-glucopyranose residues, is produced 

commercially by alkaline deacetylation of chitin. Most chitosan is produced in 

India, Japan, Poland, Norway, and Australia, mainly based on crab and shrimps 

shells discarded by the canning industries in the USA and Japan. In contrast to 

chitin, which is highly crystalline and thus insoluble in water and most organic 

solvents, chitosan is soluble in diluted acids (Eikenes et al. 2005). It was tested 

against a brown-rot fungus (Lee et al. 1993), and blue-stain and mold fungi 

(Chittenden et al. 2003, Torr et al. 2005). Wood decay tests according to EN 113 

showed that Coniophora puteana and Gloeophyllum trabeum were inhibited by 

about 6 kg chitosan/m 3 , but Oligoporus placenta may be stimulated (Schmidt 

et al. 1995). Militz et al. (2005) showed a protecting effect against all three 

fungi. On the other hand, chitin and chitosan act as chelators for metal ions 

and enhanced removal of CCA components from treated sawdust in view of 

remediation of CCA-treated wood (Kartal and Imamura 2005). 

Vanillin polymerized by laccase reduced the weight loss by C. puteana from 

25 to 5% (Rättö et al. 2004). Proteinase inhibitors like hen egg white inhibited 

growth of Ophiostoma piceae in pine sapwood samples (Abraham et al. 1997). 

Antioxidants enhanced efficacy of organic biocides in decay tests (Schultz et al. 

2005b). Chelators create metal limited conditions (Viikari and Ritschkoff 1992) 

or interact with enzymatic systems, like 2-hydroxypyridine-N-oxide (Mabicka 

et al. 2004). Cashew (Anacardium occidentale) nut shell liquid (CNSL), which 

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is a mixture of phenolics extracted from the shells of the cashew nut, reduced 

growth of some decay fungi (Pelayo et al. 2000). Venmalar and Nagaveni (2005) 

tested copperised CNSL and neem (Azadirachta indica) seed oil, containing 

azadirachtin, as preservatives. Alcoholic neem leaves extracts decreased wood 

mass loss by Oligoporus placenta and Trametes versicolor (Dhyani et al. 2005). 

Recent research on the various aspects of modified wood was compiled at 

the Second European Conference on Wood Modification (2005). 

There were (and are) many attempts at biological wood protection. To 

date, the application of microbiological control to prevent wood decay and 

discoloration has been successful in the laboratory, but inconsistent in its performance 

in the field (reviews by Bruce 1998; Bjurman et al. 1998; Chap. 3.8.1). 

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8 

Habitat of Wood Fungi 

Microbial damages to trees and wood can be differentiated into damage to the 

living tree, to felled and stored wood and in outside use, and to wood in indoor 

use. 

Such grouping is however rather for didactical reasons. There are many 

overlappings: For example Daedalea quercina is occasionally found as wound 

parasite on living oaks, frequently on stumps, more rarely on timber in outdoor 

use, like sleepers or bridge timber, and sometimes also on buildings (halftimbering 

and windows). Stereum sanguinolentum causes as well the “wound 

rot” of spruce trees (Butin 1995) as the red streaking of stored coniferous wood 

(v. Pechmann et al. 1967). 

8.1 

Fungal Damage to Living Trees 

This chapter belongs to the field of “forest pathology” and only gives an 

overview. For further reading see Tattar (1978), Schwerdtfeger (1981), Sinclair 

et al. (1987), Hartmann et al. (1988), Schönhar (1989), Butin (1995), Schwarze 

et al. (1997), and Nienhaus and Kiewnik (1998). Defense mechanisms of the 

trees are described by Blanchette and Biggs (1992) (also Chap. 8.2.1). 

The tree can be already damaged on its flowers, seeds, and seedlings by 

fungi that belong to the Oomycetes, Deuteromycetes, or Ascomycetes. Among 

the more frequently occurring fungi on flowers or inflorescences are host 

specific Taphrina species that affect alder catkins, or female flowers of poplar, 

and Thekopsora areolata damaging spruce inflorescence (Butin 1995). 

Seeds can be damaged by non-specific molds of the genera Alternaria, 

Fusarium, Penicillium,andTrichothecium. Among the specialists that can cause 

internal rotting of seeds are Rhizoctonia solani on beechnuts and Ciboria 

batschiana on acorns. Conedera et al. (2004) list several parasitic fungi that 

colonize chestnuts. 

Heat damage in seedlings is often followed by secondary infections by Alternaria, 

Fusarium, and Pestalotia species. Thelephora terrestris, Helicobasidium 

brebissonii, Rosellinia minor and R. aquila can smother seedlings or 

young plants. Seedling rots are among the most common diseases in the forest 

nursery. Important fungi on conifer seedlings are Phytium debaryanum, Phy- 

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162 8 Habitat of Wood Fungi 

tophthora species, Fusarium species, Rhizoctonia solani, andMacrophomina 

phaseolina. The Shoot tip disease of conifer seedlings is caused by Strasseria 

geniculata, Botrytis cinerea, and Sphaeropsis sapinea. Sirococcus shoot dieback 

of spruce is caused by Sirococcus strobilinus, particularly on Picea pungens and 

Pinus contorta. Meria laricis causes the Meria needle-cast of young larch. The 

Table 8.1. Some leaf diseases caused by fungi (compiled from Butin 1995) 

Disease Causal fungus Classification 

Needle-cast of Douglas fir Rhabdocline pseudotsugae Sydow Rhytismatales (A) 

Phaeocryptopus gauemannii (Rohde) Petrak Dothideales (A) 

Lophodermium needle blight Lirula macrospora (R. Hartig) Darker Rhytismatales (A) 

of spruce 

Spruce needle reddening Lophodermium piceae (Fuckel) Höhn. Rhytismatales (A) 

Spruce needle rust Chrysomyxa species Uredinales (B) 

Rhizosphaera needle browning Rhizosphaera kalkhoffii Bubák Coelomycetes (D) 

of spruce 

Lophodermium needle-cast Lophodermium seditiosum Minter, Rhytismatales (A) 

of pine Staley & Millar 

Lophodermella pine needle-cast Lophodermella sulcigena (E. Rostrup) Höhn. Rhytismatales (A) 

Naemacyclus needle-cast of pine Cyclaneusma minus (Butin) DiCosmo, Rhytismatales (A) 

Peredo & Minter 

Dothistroma needle blight of pine Mycosphaerella pini E. Rostrup ap. Munk Dothideales (A) 

Pine needle rust Coleosporium species Uredinales (B) 

Larch needle-cast Mycosphaerella laricina (R. Hartig) Neger Dothideales (A) 

Herpotrichia needle browning Herpotrichia parasitica (R. Hartig) Dothideales (A) 

of Silver fir E. Rostrup 

Silver fir needle blight Hypodermella nervisequia (DC.) Lagerb. Rhytismatales (A) 

Silver fir needle rust Pucciniastrum epilobii (Pers.) Otth Uredinales (B) 

Black snow mold Herpotrichia juniperi (Duby) Petrak Dothideales (A) 

White snow mold Phacidium infestans P. Karsten s.l. Helotiales (A) 

Keithia disease of Thuja Didymascella thujina Rhytismatales (A) 

(E. Durand) Maire 

Giant leaf-blotch of sycamore Pleuroceras pseudoplatani (Tubeuf) Monod Diaporthales (A) 

Powdery mildew of maple Uncinula tulasnei Fuckel, Erysiphales (A) 

Uncinula bicornis (Wallr.) Lév. 

Tarspotofmaple Rhytisma acerinum (Pers. ) Fr. Rhytismatales (A) 

Cristulariella leaf spot of maple Cristulariella depraedans (Cooke) Höhn. Hyphomycetes (D) 

Birch leaf rust Melampsoridium betulinum (Pers.) Kleb. Uredinales (B) 

Beech leaf anthracnose Apiognomonia errabunda (Roberge) Höhn. Diaporthales (A) 

Oak leaf browning Apiognomonia quercina (Kleb.) Höhn. Diaporthales (A) 

Oak mildew Microsphaera alphitoides Grif. & Maubl. Erysiphales (A) 

Taphrina gall of alder Taphrina tosquinetii (Westend.) Magnus Taphrinales (A) 

Leaf browning of hornbeam Gnomoniella carpinea (Fr.) Monod Diaporthales (A) 

Asteroma carpini (Lib.) Sutton Coelomycetes (D) 

Apiognomonia leaf browning Apiognomonia tiliae (Rehm) Höhn. Diaporthales (A) 

of lime 

Poplar leaf blister Taphrina populina Fr. Taphrinales (A) 

Marssonia leaf-spot of poplar Drepanopeziza punctiformis Gremmen Helotiales (A) 

Septotinia leaf blotch of poplar Septotinia populiperda Helotiales (A) 

Waterman & Cash ex Sutton 

Poplar and willow leaf rust Melampsora species Uredinales (B) 

Anthracnose of plane Apiognomonia veneta (Sacc. & Speg.) Höhn. Diaporthales (A) 

Leaf blotch of Horse chestnut Guignardia aesculi (Peck) Stew. Dothideales (A) 

A ascomycete, B basidiomycete, D deuteromycete 

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8.1 Fungal Damage to Living Trees 163 

Table 8.2. Some fungal damages to buds, shoots, and branches (compiled from Butin 1995) 


Cucurbitaria bud blight of spruce Gemmamyces piceae (Borthw.) Cassagrande Dothideales (A) 

Grey mold Botryotinia fuckeliana (de Bary) Whetzel Helotiales (A) 

Sphaeropsis shoot-killing of pine Sphaeropsis sapinea (Fr.) Dyko & Sutton Coelomycetes (D) 

Pine twisting rust Melampsora pinitorqua E. Rostrup Uredinales (B) 

Brunchorstia dieback of conifers Gremmeniella abietina (Lagerb.) Morelet Coelomycetes (D) 

Shoot shedding of pine Cenangium ferruginosum Fr. Helotiales (A) 

Juniper rust Gymnosporangium sabinae (Dickson) Winter Uredinales (B) 

Kabatina shoot killing Kabatina thujae Schneider & Arx Coelomycetes (D) 

of Cupressaceae 

Pollaccia shoot blight of poplar Venturia macularis (Fr.) E. Müller & Arx Dothideales (A) 

Myxosporium twig blight of birch Myxosporium devastans E. Rostrup Coelomycetes (D) 

Marssonina leaf and shoot blight Drepanopeziza sphaerioides (Pers.) Höhn. Helotiales (A) 

of willow 

A ascomycete, B basidiomycete, D deuteromycete 

Beech seedling disease is due to Phytophthora cactorum. OtherPhytophthora 

species attack chestnuts. Rosellinia quercina, Cylindrocarpon destructans and 

Fusarium oxysporum lead to root damage in young oaks. 

Forest canopy fungi were investigated by Stone et al. (1996). A total of 344 

different morphotaxa of endophytic fungi were isolated from leaves of Theobromae 

cacao. Most common were Colletotrichum sp., Xylaria sp. and Nectria 

sp. Inoculation of sterile leaves of young cocoa trees with these endophytes 

reduced subsequent damage by a parasitic Phytophthora sp. (Kull 2004). 

Many species of fungi are capable of causing leaf diseases. Hardwood leaf 

diseases showing superficial fungal growth, or swollen, raised, or dead leaf areas, 

may be grouped simplistically into leaf spot, blotch, anthracnose, powdery 

mildew, leaf-blister, and shot-hole. Conifers may show needle spot, cast, blight, 

and rust (Tattar 1978; Stephan 1981; Butin and Kowalski 1989; Stephan et al. 

1991). Table 8.1 only lists some fungi causing leaf diseases. Details on a specific 

disease may be read in Butin (1995). 

Some fungal damages to buds, shoots, and branches are listed in Table 8.2. 

8.1.1 

Bark Diseases 

Some bark diseases caused by fungi are listed in Table 8.3. 

Three bark diseases are described in detail. 

8.1.1.1 

Beech Bark Disease 

Beech bark disease (Fig. 8.1) has been known in Europe since about 1849 

and was imported to North America (Shigo 1964; Parker 1974; Schütt and 

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Table 8.3. Some bark diseases (compiled from Butin 1995, supplemented from Jung and 

Blaschke 2005) 


Phacidium disease of conifers Phacidium coniferarum (Hahn) DiCosmo Helotiales (A) 

Spruce bark disease Nectria fuckeliana Booth Hypocreales (A) 

Crumenulopsis stem canker Crumenulopsis soraria (P. Karsten) Groves Helotiales (A) 

of pine 

Pine stem rust Cronartium flaccidum (Alb. & Schwein.) Uredinales (B) 

(Resin-top disease) Winter 

Endocronartium pini (Pers.) Hiratsuka Uredinales (B) 

White pine blister rust Cronartium ribicola J.C. Fischer Uredinales (B) 

Larch canker Lachnellula willkommii (R. Hartig) Dennis Helotiales (A) 

Beech canker Nectria ditissima Tul. Hypocreales (A) 

Beech bark disease Nectria species Hypocreales (A) 

Black bark scab of beech Ascodichaena rugosa Butin Rhytismatales (A) 

Fusicoccum bark canker of oak Fusicoccum quercus Oudem. Coelomycetes (D) 

Chestnut blight Cryphonectria parasitica (Murrill) Barr Diaporthales (A) 

Dothichiza bark necrosis and Cryptodiaporthe populea (Sacc.) Butin Diaporthales (A) 

dieback of poplar 

Canker stain of plane Ceratocystis fimbriata (Ellis & Halstead) 

Davidson f. platani Walter Ophiostomatales (A) 

Stereum canker rot of Red oak Stereum rugosum (Pers.) Fr. Aphyllophorales (B) 

Pezicula canker of Red oak Pezicula cinnamomea (DC.) Sacc. Helotiales (A) 

Coral spot Nectria cinnabarina (Tode) Fr. Hypocreales (A) 

Sooty bark disease of sycamore Cryptostroma corticale (Ell. & Ev.) Hyphomycetes (D) 

Gregory & Waller 

Sudden oak death Phytophthora ramarum (Werres, Pythiales (O) 

De Cock & Man in’t Veld) 

A ascomycete, B basidiomycete, D deuteromycete, O oomycete 

Lang 1980; Eisenbarth et al. 2001). It develops particularly on trees older 

than 60 years of European Fagus sylvatica and American beech F. grandifolia 

by a disturbance of the water regime due to a abiotic/biotic factor complex: 

moist site, dry summer, participation of the Beech scale, Cryptococcus fagisuga 

(Lunderstädt 2002) and either one of two bark-killing Ascomycetes, in Europe 

Nectria galligena and in North America N. coccinea var. faginata (Mahoney 

et al. 1999), and possibly of mycoplasmas. Classical pathogenesis is an often 

short-lived mass reproduction of the Beech scale, which causes subcortical 

changes and subsequent infestation with Nectria. Xylem breeding Trypodendron 

domesticum and Hylecoetus dermestoides may follow. The larval galleries 

may be subsequently colonized by white-rot fungi. The susceptibility to the 

disease is biotically effected, whereby the physiological condition of the tree 

and its genetic potential determine the efficacy of the damaging agents (Beech 

scale, Nectria spp., beetles, white-rot fungi). The outbreak and/or healing can 

be controlled by the site conditions (Braun 1977; Lunderstädt 1992). 

The fungus invades the bark that was previously altered by the feeding 

activity of the Beech scale. A red-brown to blackish (bark tannic substances) 

slimy liquid may ooze from the bark tissue (Wudtke 1991). Under the bark 

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Fig.8.1. Beech bark disease. a tarry spots on the bark, b occluded bark lesions, c determination 

of extent of necrosis by scoring the bark with a timber scribe, d early stage of necrosis, 

e late stage with incipient white rot (from Butin 1995, by permission of Oxford University 

Press) 

develop dark regions with dead cambium to over 1 m in extension. Small 

necroses with exposed wood may be closed by callus formation, which leads 

to a T-shaped fault in the xylem. Tylosis formation causes wilting. Massive 

invasions can result in tree dieback. Larger necroses form infestation gates for 

white-rot fungi (Bjerkandera adusta, Fomes fomentarius, Fomitopsis pinicola, 

Hypoxylon species, Stereum hirsutum) (Eisenbarth et al. 2001). 

8.1.1.2 

Chestnut Blight 

The Chestnut blight (chestnut bark canker) (Fig. 8.2) is caused by the ascomycete 

Cryphonectria parasitica (Halmschlager 1966; Heiniger 1999). The 

pathogen was imported on Asian rootstock to New York in 1904 and caused 

lethal cankers on more than 3.5 billion susceptible American chestnut trees, 

Castanea dentata, across 9 million acres of the eastern US, being there at that 

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Fig.8.2. Pathogenesis of Chestnut blight by Cryphonectria parasitica (translated from 

Heiniger 1999, with permission of Swiss Federal Institute for Forest, Snow and Landscape 

Research) 

time the most important hardwood species. The disease appeared in Europe 

first in 1938 in Genoa in the European chestnut sites (Castanea sativa)ofItaly, 

then in southern France, Spain, Switzerland (1948), Germany (1992), and Eastern 

Europe. The fungus penetrates as a spore by means of wind, rain, insects, 

or birds through wounds into the bark until the cambium. Then reddish-brown 

bark spots that break to longitudinal fissures, branch-surrounding necroses, 

wilt, and death of the affected branch or crown region occur. One- to 2mm-large, 

orange-yellow-ochre pustules (conidiomata, ascomata) develop on 

the bark. 

The disease in Europe does not run however as intensively as in the USA 

probably due to lesser aggressive fungal strains. The reduced pathogenicity is 

caused by Cryphonectria-hypovirus 1 that infests the fungus, that is, it becomes 

lesser virulent and only produces superficial cankers, which soon heal up. The 

virusisalsofoundinthenaturalC. parasitica populations in Japan and China, 

but not in the North American populations. To limit the distribution of the 

fungus in non-infested countries, there are official regulations (European and 

Mediterranean Plant Protection Organization) (Heiniger 2003). 

Breeding experiments are performed between C. dentata and resistant Asian 

species. There are also attempts on a biological control based on vegetative 

pairing of hypo-virulent fungal isolates with virulent strains. Infested sites 

are inoculated with hypo-virulent isolates that can transfer the virus in the 

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pathogen if both belong to the same vegetative compatibility group (Haller- 

Brem 2001). There are biotechnological attempts for transgenic chestnut trees 

(Gartland and Gartland 2004). 

8.1.1.3 

Plane Canker Stain Disease 

The Plane canker stain disease (plane tree canker) (Fig. 8.3) is caused by the 

ascomycete Ceratocystis fimbriata f. sp. platani (Wulf 1995). The disease was 

for the first time observed on Platanus species in 1926 in the eastern USA 

and occurred in the 1940s in Europe [France, Italy, Spain, Switzerland, Turkey; 

Clerivet and El Modafar (1994)]. About 80% of the city-trees along motorways 

became destroyed until 1950 in the USA. Marseille lost over 1,500 100-year-old 

plane trees in 12 years. The fungus penetrates through wounds predominantly 

after pruning, more rarely by insects, into the bark of the stem and the branches 

and leads to cambium dying and elliptical bark necroses. Later, it colonizes 

the outer sapwood with bluish-brown discoloration. Excretion of toxins by 

the fungus and tyloses effect wilting of individual crown portions. Thus, the 

fungus both produces a bark and a wilt disease (Butin 1995). The trees die 

usually within 3–6 years. Reproduction organs are predominantly found on 

Fig.8.3. Plane canker stain disease. a Symptoms on plane, b stem cross section showing 

stained wood, c tangential stem section showing the stain as streaks, d phialide with conidia 

of the Chalara anamorph, e conidiophore with chlamydospores, f perithecium, g ascospores 

(from Butin 1995, by permission of Oxford University Press) 

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cut sections of debranched or felled trees: perithecia with ascospores and three 

different anamorphs, e.g., Chalara. Necroses can by secondarily colonized by 

other fungi (Chondrostereum purpureum, Schizophyllum commune). 

Transfer may be reduced by hygienic measures like removal of infested trees. 

National and EC regulations have to be considered. 

8.1.2 

Wilt Diseases 

Diseases that affect the vascular system of a plant are called wilt diseases. 

A fungus causes moisture stress that leads to wilting, killing of large branches 

and even entire trees (Tattar 1978). Two important wilt diseases caused by 

fungi are Dutch elm disease and Oak wilt. 

8.1.2.1 

Dutch Elm Disease 

Dutch elm disease (“elm dying”) (Fig. 8.4) is caused by the ascomycetous 

fungus Ophiostoma ulmi s.l. (Gibbs 1974; Rütze and Heybroek 1987; Sinclair 

et al. 1987; Ouellette and Rioux 1992; Butin 1995; Harrington et al. 2001; 

Kirisits et al. 2001; Nierhaus-Wunderwald and Engesser 2003). The pathogen 

is composed of two separate species or three subgroups: the non-aggressive 

(NAG) subgroup, referred to as O. ulmi, and the two races, Eurasian (EAN) and 

North American (NAN), of the aggressive subgroup, referred to as O. novoulmi 

(Brasier 1999). The disease was probably imported from East Asia around 

1917 over France into the Netherlands in 1919 (NAG), where 1920/21 the first 

comprehensive investigations took place, so that the disease was called Dutch 

elm disease. In 1923, it had arisen for the first time in England, 1930 via veneer 

wood in the USA, 1934 in almost all European countries and 1939 in the former 

Soviet Union (Heybroek 1982). Between 1940 and 1960 it receded, but again 

a new aggressive eastern race (EAN), probably from Romania/Ukraine, spread 

westward over the whole of Europe and eastward to middle Asia. Assumably 

after the import to North America, there the aggressive western race (NAN) 

shall have developed and imported to Great Britain (Röhrig 1996). 

The wood loss in an economical view is very great. Altogether, hundreds of 

millions of decade- to centuries-old elm trees in Europe, North America, and 

in parts of Asia were destroyed. About 25 million elms died since the 1970s in 

England (Wörner 2005), and in Utrecht and Amsterdam, half of all the elms 

died. 

Scolytid bark beetles are the principal agents of the long-distance transmission 

introducing the pathogen into healthy trees during adult feeding. In 

Europe, the principal vectors are Scolytus scolytus and S. multistriatus, but also 

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Fig.8.4. Pathogenesis of Dutch elm disease (translated from Nierhaus-Wunderwald and 

Engesser 2003, with permission of Swiss Federal Institute for Forest, Snow and Landscape 

Research) 

other vector species are recognized (Wingfield et al. 1999). In North America, 

vectors are the imported S. multistriatus and the American elm bark beetle 

Hylurgopinus rufipes. The females select almost exclusively diseased, dying, or 

already dead elms for their breeding galleries. The larvae take up the pathogen, 

which is passed on alive via the pupa to the young beetle. The young beetles 

contaminated with spores (conidia or ascospores) infect healthy trees in twig 

crotches of small branches during maturation feeding. Since this bark is too 

thin for oviposition, the beetles leave the healthy tree and use the thick-barked 

parts of diseased elms for their breeding galleries. The change between the stem 

of infected elms and the branches of healthy trees makes the Scolytus beetles 

effective vectors (v. Keyserlingk 1982). Root graft transmission via connections 

from adjacent trees is the major cause of elm death in urban areas. 

The principal reaction compounds developing in elms following invasion 

by the fungus are cadinane sesquiterpenoids [mansonones, elm phytoalex- 

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ins; e.g., Meier and Remphrey (1997)]. Barrier zones containing starch-filled 

parenchyma and swollen ray parenchyma have also been observed. During 

pathogenesis, the fungus develops within the xylem vessels with associated 

tyloses and vessel plugging, ultimately resulting in the wilt syndrome 

(Smalley et al. 1999), promoted by fungal toxins (cerato-ulmin) (Brasier et al. 

1990). A branch cross section shows dark spots in the earlywood, which form 

brownish longitudinal strips in the tangential plane. An infection with a nonaggressive 

strain can be buried by new annual rings (chronic form); an aggressive 

strain grows through the annual ring borders (acute form), and the tree 

can die within 2 years. 

The use of pheromones as an attractant does not cover all beetles. Fungal 

inhibitors such as benomyl only result in a dilatory effect. There were attempts 

of a biological control with the bacterium Pseudomonas syringae van Hall and 

with Trichoderma species (Aziz et al. 1993). Mansonones inhibited the growth 

of O. ulmi in vitro. Control measures are felling of infected or weakened trees as 

well as debarking and burning the bark and thicker branches in order to reduce 

the beetle population. In view of resistance to the pathogen, the major sources 

of genes for resistance are possessed by Asiatic elms. The responses of the European 

and North American elms vary depending on the individual subgroups 

of the pathogen. Classical breeding for resistance by selection of individuals 

from native populations have been made since the 20s, and hybrid elms have 

been bred, incorporating natural resistance from Asian elms. There are indications, 

which are based on DNA techniques, that most of the English elms, 

Ulmus minor var. vulgaris, are clones deriving from an Italian tree exported 

by the Roman agronomist Columella from Latium via Spain to England. That 

would explain the observed small genetic diversity within the English elms 

and thus their high susceptibility to the pathogen (Wörner 2005). Currently, 

two biotechnological approaches are pursued: Double-stranded RNA viruses, 

known as d-factors, may have the potential to reduce aggressivity if introduced 

into a fungal population at large in sufficient quantities to become established 

and spread through fungal populations. Transgenic Ulmus procera trees have 

been produced using Agrobacterium rhizogenes (Riker et al.) Conn and A. 

tumefaciens as mediator, demonstrating that a variety of exogenous genes can 

be expressed in regenerant elms (Gartland and Gartland 2004). 

8.1.2.2 

Oak Wilt Disease 

The North American oak wilt (Fig. 8.5; Rütze and Liese 1980, 1985a; Sinclair 

et al. 1987) is a vascular disease that is endemic among oaks in the USA and 

caused by the ascomycete Ceratocystis fagacearum. It was recorded for the first 

time in Minnesota in 1928, Wisconsin in 1942, already in 1979 in 21 US states 

east of the Great Plains and is now also found in Texas and Tennessee. The 

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fungus can both infect red oaks (Quercus falcata var. pagodaefolia, Q. rubra, 

Q. shumardii)andwhiteoaks(Q. alba, Q. bicolor, Q. macrocarpa, Q. michauxii, 

Q. muehlenbergii). Red oaks become systematically infected and die quickly, 

mostly within the year of first wilting symptoms and sometimes within a few 

weeks after infection. The economically more important white oaks are more 

resistant and show the damage often being restricted to just a few branches. 

The lower susceptibility of the white oak is attributed to smaller earlywood 

vessel diameter, more intensive tylosis formation resulting in a slower spread 

of the fungus in the tree and the ability to “bury” infected tissue by a new 

annual ring. 

The infection usually occurs via root graft transmissions between the diseased 

and healthy trees (Fig. 8.5a), so that the distribution is low with 1 to 2 m 

(maximum 8 m) per year. Local spreading via root grafts can be inhibited by 

ditches. The fungus invades the vessels of the youngest two annual rings and 

stimulates the adjacent parenchyma cells to tylosis formation. Thus, wilt and 

defoliation occur in the undersupplied crown regions. Additionally, wilt toxins 

are produced. The leaves become flabby and discolor, are light green from the 

edge, and later bronze-brown in red oak and pale-light brown in white oak. 

After tree death, the hyphae grow inward in the sapwood as well as outward 

Fig.8.5. Transmission of the Oak wilt 

fungus, Ceratocystis fagacearum, via 

root-grafts (a), during maturation feeding 

of bark boring beetles (b), and from 

sporulating mat by sap feeding nitidulids 

(c) (from Rütze and Parameswaran 1984) 

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through the cambium under the bark. In the cambial layer particularly in red 

oak, 5 to 8-cm-large sporulation mats (usually conidia) develop from May 

to October, which cause bark detachment and fissure by means of pressure 

cushions. 

There are two different ways for long-distance transmission by insects (about 

100 m/year): First, oak bark beetles (Pseudopityophthorus spp.) breed in dying 

or dead oaks, and the young beetles transfer the pathogen during the maturation 

feeding on shoots and twigs of healthy oaks (Fig. 8.5b). Since asexual 

spores do not develop in the larval galleries, this transmission way has only 

less significance. Second, sap beetles, particularly Nitidulidae, are attracted by 

the smell of the sporulating mats and transmit infectious material to healthy 

trees into fresh wounds, attracted by their smell (Fig. 8.5c) (Appel et al. 1990). 

The nitidulids effect that the bipolar heterothallic fungus is dikaryotized and 

develops ascospores, if conidia with contrary mating factor were introduced 

from other sporulation mats. Since wounds are infectious only a few days in 

healthy oaks, this infection way has also less significance. Furthermore, the 

subcortical mats of C. fagacearum were observed to be rapidly overgrown by 

Graphium pyrinum Goid. (anamorph of Ophiostoma piceae). This colonization 

reduces the chance of contamination of the insect vectors with spores of the 

pathogen and is likely to contribute to the low efficacy of insect transmission 

(Rütze and Parameswaran 1984). 

Since 1951, the import of unpeeled oak logs from North America to Germany 

was allowed according to a plant protection order, if the wood derives 

from healthy areas (“white counties”), in accordance with the plant protection 

departmentoftheUSDepartmentofAgriculture.Ithadhowevertobeconsidered 

that also the European oaks, although usually white oaks (Quercus petraea 

and Q. robur), are more susceptible from nature and that the European oak 

bark beetle Scolytus intricatus is more aggressive in its transmission behavior 

than the North American species. In order to prevent the import of the fungus 

(Gibbs et al. 1984), oak wood became subject to specific treatment requirements 

under Council Directive 77/93/EC including bark removal, kiln drying, 

etc.Sincesuchwoodcannotbeconvertedtoveneers,thosemeasureswould 

have equaled practically an import stop for oak logs and the endangerment of 

the European veneer industry. Thus, experiments were performed in a cooperative 

venture between the Federal Research Center for Forestry and Forest 

Products Hamburg and the Universities of Minnesota and West Virginia on log 

fumigation with bromomethane (methyl bromide) as a means of ensuring that 

thelogswerefreefromC. fagacearum (Liese et al. 1981; Schmidt 1988). The 

European community permitted by EEC Plant protection guidelines of 1978 

the import of unpeeled oak logs if they were disinfected before export with 

240 g bromomethane per m 3 of wood for 3 days at a minimum temperature 

of 3 ◦ C in plastic tents (Rütze and Liese 1983). The use of bromomethane has 

fallen off considerably since the Montreal Conference of 1997 because of its 

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8.2 Tree Wounds and Tree Care 173 

destruction to the ozone layer. Log fumigation needs an exemption. In Europe, 

to monitor that sufficient bromomethane fumigation of the oaks has been carried 

out, the TTC test (Brunner and Ruf 2003) is suitable. The test is based on 

the fact that the gas kills not only the oak wilt fungus but also the living cells 

of oak sapwood. These cells would survive for several months in logs that are 

not treated with gas. Increment cores of the whole sapwood are treated with 

a 1% solution of 2,3,5-triphenyl-2H-tetrazolium chloride (colorless), which 

discolors dark red to triphenylformazan in contact with living cells by their 

dehydrogenase activity (Rütze and Liese 1985b). 

Fumigation with bromomethane has also been applied to four pathogenic 

fungi in larch heartwood (Rhatigan et al. 1998). Due to the pending restrictions 

of bromomethane for phytosanitation in general, the potential substitution by 

sulfuryl fluoride and iodomethane was investigated (Schmidt et al. 1997c, 

Unger et al. 2001). 

There are privileges of the import regulations for the fewer endangered white 

oak: no fumigation during winter months, however immediate debarking and 

burning of the bark as well as immediate wood processing. Since the wood of 

both oak groups is hardly or not at all to differentiate by appearance, a color 

test is suitable: When sprayed on the heartwood of any species of white oak 

a sodium nitrite solution produces a blue-black color within a few minutes, 

whereas the color is a reddish brown in red oak (Willeitner et al. 1982). 

The possible oak wilt transmission to Europe was discussed several times 

in connection with the increasing illness of European oaks (Siwecki and Liese 

1991). These damages develop however due to a complex effect of abiotic factors 

(dryness and pollutants as predisposing factors, severe winter frost as acute 

stressor) and biotic influences (leaf-eating insects, nematodes, phytoplasmas, 

and Armillaria spp. as weakness parasites, other Ceratocystis species, other 

fungi.) The literature on the role of pathogens in the present oak decline in 

Europe has been compiled by Donaubauer (1998). 

8.2 

Tree Wounds and Tree Care 

8.2.1 

Wounds and Defense Against Discoloration and Decay 

Initiation for discolorations and decay are predominantly wounds that are 

frequently caused by animals chewing, branch breaking, pruning, mechanized 

wood harvest, construction injury, and motor traffic (Tattar 1978). 

Rots in living trees might occur fast or result from processes of many years, 

which frequently remain hidden for a long time, until fruit bodies appear, or 

the tree is broken, thrown by the wind, or felled. 

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After wounding, tree-own discolorations (deposition of heartwood substances) 

develop by living cells, followed by microbial stain and finally by 

wood rot (Shigo and Hillis 1973; Hillis 1977; Shortle and Cowling 1978; Bauch 

1984; Rayner and Boddy 1988; Fig. 8.6). 

Depending on the fungus and tree species, brown, white, or even soft-rot 

decay can develop in the tree. Sapwood and/or heartwood can be colonized. 

Fungi may be saprobionts of parasites. Development and spread of decay are 

influenced by the tree species, which can be susceptible, like birch or poplar, 

or exhibits natural durability in its heartwood due to inhibiting accessory 

compounds. 

It has to be distinguished between passive mechanisms, which are already 

present before damage, and active defense mechanisms, which trees trained in 

the course of their phylogenetic development to limit wounds, infections, and 

senile damages (Blanchette 1992; Duchesne et al. 1992; Rayner 1993). 

After the xylem is wounded, two defense functions have to be differentiated: 

First, the tree must avoid an interruption of the transpiration stream by air 

embolism, and second, limit the spread of invaded microorganisms (Liese and 

Dujesiefken 1996). 

When a softwood tracheid is injured, its lumen is filled with air at ambient 

pressure. Thus, a pressure drop exists across the pit membranes of the watercontaining 

neighboring tracheids. Their tori are therefore pulled against their 

pit borders, and the air-blocked tracheid is thus sealed off from the waterconducting 

tracheids (Zimmermann 1983). Conifers protect themselves from 

Fig.8.6. Model of successive changes in 

the stem wood after prior injury to the 

bark. w wound, c callus margin, f fruit 

body, b barrier zone, r rot, m microbial 

wood discoloration, t tree-own wood 

discoloration (after Shigo 1979) 

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wounds and penetrating microorganisms by phenolic compounds, terpenoids, 

and resin (Tippett and Shigo 1981). 

In hardwoods, the defense reactions depend on physiologically active parenchyma 

cells. The water-conducting system is protected against damage by 

tyloses, plugs or membranes, and phenolic substances or suberin are deposited 

on the cell wall or in the lumen (Schmitt and Liese 1993). 

For the graphic understanding of the spatial cut off within a tree, Shigo 

developed the CODIT model (Fig. 8.7; Shigo and Marx 1977; Shigo 1984), 

which stands for “compartmentalization of decay in trees”. The model means 

that the tree protects itself from penetrating microorganisms by four inhibiting 

walls and that the spatial expansion of discoloration and decay is determined 

by the anatomical structure of the wood. The axial “walls 1” with the weakest 

partitioning effect are formed by the closure of the vessels and pits above and 

underneath a wound by gums and tyloses. The tangential “wall 2” stem-inward 

occurs by the annual ring borders and by the sapwood/hardwood boundary. 

The radial “walls 3” are caused by the lateral wood rays. The most effective 

compartmentalization is by “wall 4”, also termed barrier zone, formed by the 

cambiumaftertheinjurywithincreasedparenchymacontent. 

The CODIT model interprets the tree-own reactions as compartment formation 

against microorganisms. It seems, however, more biological that the 

tree protects itself first from penetrating air, particularly since wood fungi 

can only settle the tissue if air is present. With changed definition, the term 

CODIT has been expanded by Liese and Dujesiefken (1989, 1996): the D does 

not only stand for decay, but also for damage and covering desiccation as well 

as dysfunction. 

The histological changes that occur in wood and bark as wound reactions 

in hardwoods are schematically shown in Fig. 8.8. 

The parenchyma cells die at the surface of the damaged wood area. The 

tissue beneath the wound plane also dies, without mobilizing reserve materials, 

Fig.8.7. CODIT model with walls 1 to 4 

(after Shigo 1979) 

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Fig.8.8. Changes in the xylem and phloem of hardwoods after wounding (after Liese and 

Dujesiefken 1989) 

since the defense reactions in the wood begin temporally retarded. In this 

bright zone of about 1 cm, the vessels remain open, and the lumina do not 

contain inclusions. With increasing distance to the wound, reserve material 

is mobilized, and the vessels are closed. In beech, the degeneration of the 

parenchyma is limited, as parenchyma cells in the wounded area are divided 

by transverse walls and limit the damage by suberization of the wound-near 

compartments (Schmitt and Liese 1993). 

A closure by tyloses (Schmitt and Liese 1994) only takes place in tree species, 

which possess pit sizes of at least 8µm. Trees without tyloses, like lime and 

maple, can prevent air embolism by blocking the vessels with plugs. In birch, 

the ladder-shaped vessel openings are closed on one side by membranes, and 

parenchyma cells excrete fibrillar material in neighboring vessels and fibers 

(Schmitt and Liese 1992a). 

The tissue behind the wound area, which is discolored by means of accessory 

compounds and which contains died parenchyma cells and vessels 

out of function, had been termed protection wood. As it is colonized however 

frequently by fungi, it obviously does not possess increased durability. 

The healthy wood outside this area shows microscopically in an area of a few 

millimeters mobilization of reserve material and vessel closure, but no fungi, 

so that the actual protective layer obviously lies outside of the visible discoloration. 

Also in the phloem the parenchyma dies at the wound surface and the 

tissue beneath is set out of function. A wavy-shaped wound periderm, which 

attaches the periderm of the young callus bark to the outer bark, develops in 

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the transition of the discolored to the functional phloem (Trockenbrodt and 

Liese 1991). 

The cambium reacts to the damage at the wound margin with intensified 

cell formation (callus) to overwall the opened wood body (Stobbe et al. 2002a). 

The wound wood, which is later formed outside the callus, effectively limits 

discoloration and decay outward. 

8.2.2 

Pruning 

Forest tress are pruned to produce high-class timber, trees in urban areas are 

pruned for safety reasons and along motorways and power-lines for clearance. 

Each cut causes a wound, which leads in the exposed wood to discoloration 

and decay (Fig. 8.9). 

Until the 80s in Germany, the flush cut had been regarded as the correct 

method when removing a branch back at the stem. Studies on the pruning 

of hardwoods carried out by Shigo and staff (Shigo 1989) caused confusion. 

Comparing the effects of different cut locations of a total of 750 pruning 

wounds on 115 street and park trees led to the Hamburg Tree Pruning System 

(Dujesiefken and Stobbe 2002), which is integrated since 1992 into the 

German rules and regulations for tree care methods. The recommendations 

Fig.8.9. Discoloration reaching far into 

thestemofhorsechestnut9years 

after flush cut pruning (a); reduced 

discoloration after a branch collar cut 

(b) (from Dujesiefken and Stobbe 2002) 

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for branches without branch collar are also part of the European Tree Pruning 

Guide. According to the branch attachment (branches with or without a collar), 

the cut has to be done outside the stem so that the branch bark ridge is not 

damaged. Flush cuts have to be avoided. When pruning dead branches, the 

distinctive swelling at branch base must remain at the stem. Regardless of the 

time of year and the tree species, radical tree pruning, e.g., a drastic removal 

of crown parts, should not be done. If possible, branches greater than 5 cm 

in diameter of weak compartmentalizing trees (e.g., Aesculus, Betula, Malus, 

Populus, Prunus, Salix), and greater than 10 cm of strong compartmentalizing 

trees (e.g., Carpinus, Fagus, Quercus, Tilia) should only be reduced partially 

rather than completely. 

For organizational reasons and due to nature protection laws, pruning is 

usually done during the dormant season. However, wounding of maple, birch, 

beech, oak, ash, lime tree and spruce showed on the basis the intensity of 

the wood discolorations that injuries should be avoided in hardwoods during 

the dormant stage and in spruce from late summer to winter due to different 

wound reactions (Lenz and Oswald 1971; Armstrong et al. 1981; Dujesiefken 

et al. 1991; Schmitt and Liese 1992b). 

8.2.3 

Wound Treatment 

In the 50s and 60s, large stem wounds were shaped out and filled with concrete. 

Since concrete and wood shrink and extend differently under weather 

influence, shakes develop, water penetrates and leads to rot. Since the 70s, the 

cleaned wounds were treated with wound dressings or with wood preservatives. 

Disinfection of the opened wood body with ethanol or alcoholic iodine 

solutionbeforewoundtreatmentdidnotledtoapreventionofdiscoloration 

and decay in beech and ash (Dujesiefken and Seehann 1995). The use of wood 

preservatives was disputed for tree care measures, as they are not developed for 

theprotectionoftreewounds.Thetreatmentofartificialwoundswithwood 

preservatives resulted in beech in more intensive discolorations behind the 

wound area than at wounds, which were only treated with wound dressings. 

Wound dressings belong to the plant preservatives. In Germany, they must 

be tested according to efficacy and environmental compatibility to become 

licensed (Balder 1995). 

Alternatively, cavities can be foamed with polyurethane (Dujesiefken and 

Kowol 1991). Figure 8.10 shows reduced discoloration in a beech tree after 

filling the wound with polyurethane. 

Currently, traffic wounds on street trees are covered by black plastic wraps, 

which promote the development of a surface callus overgrowing the wound 

area (Fig. 8.11). 

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8.2.4 

Detection of Tree and Wood Damages 

Fig.8.10. Discoloration of beech at the 

control wound protected with wound 

dressing (a) and at the cavity filled with 

polyurethane (b) (from Dujesiefken and 

Kowol 1991) 

To investigate trees with regard to microbial damage, particularly to detect decay, 

discolorations, cavities, shakes and generally pathological changes, as well 

as to determine wood quality in felled timber, construction wood and woodbased 

composites, numerous methods are available. Inspection methods were 

described by McCarthy (1988, 1989), Zabel and Morrell (1992), Niemz et al. 

(1998), Londsdale (1999), Harris et al. (1999), Unger et al. (2001). Methods 

can be classified as destructive, nondestructive, or near-nondestructive. They 

reach from technically simple procedures like using increment bore tools to 

expensive equipment like computer tomography (Habermehl and Ridder 1993; 

Habermehl 1994) as well from subjective visual inspection to objective molecular 

techniques. In view of tree care, noninvasive or less destructive methods are 

preferable (Niemz et al. 1999; Kaestner and Niemz 2004). Modern techniques 

for nondestructive characterization and imaging of wood were reviewed by 

Bucur (2003) and comprise ionizing radiation computed tomography, thermal 

imaging, microwave imaging, ultrasonic imaging, nuclear magnetic resonance 

and neutron imaging. Some methods are preferentially used for trees, others 

for lumber, some may be used on the spot, others are pure laboratory tech- 

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Fig.8.11. Use of a plastic wrap to improve 

the development of a surface callus. 

a Fresh, cleaned wound. b Wound 

covered with a black plastic wrap. 

c After9weeksnearlyhalfofthewound 

coveredwithabrightsurfacecallus 

tissue (from Stobbe et al. 2002b) 

niques, and some of the latter are capable to identify the causal agent. Due 

to some overlapping in their use, the methods are listed together in Table 8.4. 

Limits of ultrasonic evaluation of wood defects have been shown by v. Dyk and 

Rice (2005). There is a great bulk of references on the various techniques; thus, 

only examples are given in Table 8.4. 

The earliest nondestructive evaluation of trees is the visual inspection of 

the tree condition (growth, foliation, wilt) and occurrence of wounds, resin 

excretion, necrosis, canker, or fruit bodies. Visual inspection is also applied 

for lumber, poles, and wood in indoor use. Fruit bodies might serve to identify 

the causal agent. This visual inspection is by definition neither objective nor 

sure. Olfactory detection is done by the use of sniffer dogs that detect dry rot 

(Koch 1991), molds, or termites (Zabel and Morrell 1992). 

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Table 8.4. Inspection methods for fungal activity and wood quality in trees and timber 

Method Procedure Advantage, disadvantage Reference 

Optical Visual Non-destructive, in situ, subjective 

Endoscopy Hidden spaces in buildings, bore holes may be required Janotta (1995) 

Rhizoscopy Root system Seufert et al. (1986) 

Light microscopy Simple, destructive Schwarze et al. (1997), Anagnost (1998) 

Electron microscopy Accurate, photographic record, destructive Liese (1970), Daniel (2003) 

IR/NIR/FTIR spectroscopy Laboratory method, printed record Körner et al. (1992), Schwanninger et al. (2004) 

UV microspectrophotometry Laboratory method, 3D wood topochemistry Koch and Kleist (2001) 

Raman spectroscopy Laboratory method, wood topochemistry Röder et al. (2004) 

Spectrometrical GC-MS Laboratory method, MVOC’s, mold and rot detection Keller (2002), Blei et al. (2005) 

MALDI-TOF MS Laboratory method, fungal identification Schmidt and Kallow (2005) 

Acoustic Speed of ultrasound (Impulse hammer) Non-destructive, in situ, density of sound wood must be known Rust (2001), Niemz et al. (2002) 

Acoustic emission Non-destructive, in situ Noguchi et al. (1992) 

Electrical Electrical resistance, conductivity Shigo et al. (1977), Kučera (1986) 

(Shigometer, Vitamat) Less destructive, in situ, handy devices Larsson et al. (2004) 

Nuclear magnetic resonance Non-destructive, not transportable, expensive Müller et al. (2002) 

Radar Ground penetrating radar for root investigation, in situ Barton and Montagu (2004) 

Microwaves Non-destructive Takemura and Taniguchi (2004) 

Mechanical Increment cores Handy instruments, low cost, destructive Niemz et al. (1998) 

Needle penetration (Pilodyn) Handy instruments, low cost, nearly non-destructive Niemz and Kučera (1999) 

Drill resistance (Resistograph) Portable instruments, printed data plots, destructive Rinn (1994), Isik and Li (2003) 

Thermographic Heat radiation Non-destructive, handy instruments, resolution insufficient Niemz et al. (1998) 

Radiographic X-ray, γ-ray computed tomography Non-destructive, in situ, expensive Habermehl (1994) 

Calorimetric Isothermal microcalorimetry Laboratory method, non-destructive Xie et al. (1997) 

Microbiological Culturing to pure culture Laboratory method, fungal identification 

Biochemical CO2 Laboratory method, fungal activity Kirk and Tien (1986) 

ATP Laboratory method, fungal activity McCarthy (1983), Bjurman (1992a) 

Chitin Laboratory method, fungal quantification Nilsson and Bjurman (1998) 

Ergosterol Laboratory method, fungal quantification Pasanen et al. (1999), Dawson-Andoh (2002) 

pH-value Non-destructive, fungal activity, brown/white rot differentiation Peek et al. (1980) 

Sniffer dogs Non-destructive, detection of dry rot, molds Koch (1991), Keller et al. (2004) 

Molecular Protein gel electrophoresis Laboratory method, fungal identification Schmidt and Kebernik (1989), Vigrow et al. (1989) 

Immunology Laboratory method, early decay, fungal identification Vigrow et al. (1991a,b), Clausen (1997a) 

DNA-based methods Laboratory method, fungal identification, objective White et al. (2001), Schmidt (2000) 

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The type and intensity of a biological attack can be recognized by different 

macromorphologic changes of the wood tissue. Typical discolorations occur 

on and inside wood that is colonized by molds, blue stain, and red-streaking 

fungi. Brown- and soft-rotten woods differ in color and shape of the brown 

and soft-rotten cubes, and white-rotten wood between simultaneous and white 

pocket rot. 

Various mechanical and physical wood changes occur when wood-inhabiting 

microorganisms colonize wood. Wood mass (weight) loss is a commonly 

used measure of decay capability. The basic calculation is: [(weight before − 

weight after): weight before] ×100%. The extent of decay in a specimen that 

was sampled from attacked wood can be determined the same way, if its dry 

weight is compared to that of a comparable healthy control: mass loss ML 

(%) = [(DW1 −DW2) :DW1] × 100 (DW1 = dry weight of control, DW2 =dry 

weight of decayed sample). 

Mass loss of wood samples exposed to fungi is likewise used to determine the 

efficacy of wood preservatives and to examine the natural durability of wood 

species. There is a permanent discussion if fungal pure cultures or artificial 

mixed cultures should be used in laboratory tests (Kolle flask method, soilblock 

test, vermiculite method) or if soil contact decay tests are preferable. 

Laboratory tests are reproducible as they are based on defined test fungi. 

Field stake tests result in a severe exposure condition as the natural microbial 

composition may contain microorganisms that degrade wood, biodegrade 

organic wood preservatives or modify inorganic preservatives making them 

more susceptible to leaching (Nicholas and Crawford 2003). In Europe, the 

Kolle flasks method with malt extract agar and defined wood blocks of 5 × 

2.5 × 1.5 cm in size from Scots pine sapwood and European beech is used for 

Basidiomycetes according to the standard EN 113 (Fig. 7.5; Table 7.6). In this 

method, specified isolates of certain fungal species, e.g., Coniophora puteana 

Ebw. 15, have to be used. The wood decay capacity of the test organisms is, 

however, erroneously named “virulence”, although it concerns fungi and not 

viruses.Soft-rotfungi tests are performedinplasticcontainers with vermiculite 

(grainy substance of aluminum iron magnesium silicate) as moisture and 

nutrient depot. The standard soil block test AWPA E10 uses either 14-mm or 

19-mm wood cubes that are exposed to white- and brown-rot fungi that were 

previously inoculated onto wood wafers on top of a sterile moist soil bed in 

a bottle. Soil bed testing based on the methodology described in the European 

Pre-standard ENV 807 uses 100 mmlong ×10mmrad ×5mmtang specimens that 

are exposed to the naturally soil-inhabiting microorganisms (v. Acker et al. 

2003). Field stake tests use stakes or posts of the selected wood species that 

are half buried vertically in soil and inspected for decay at intervals. Wood 

assembly above-ground tests (post-rail, L-joint, lap-joint), all including some 

type of joint that effectively traps rainwater, simulate decking, door frames or 

joinery (Zabel and Morrell 1992; Nicholas and Crawford 2003). 

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8.3 Tree Rots by Macrofungi 183 

The degree of wood decay can be quantified by changes in wood strength 

properties, modulus of rupture, work to maximal load in bending, maximal 

crushing strength, compression perpendicular to the grain, impact bending, 

tensile strength parallel to the grain, toughness, hardness, and shear strength 

(Wilcox 1978; Zabel and Morrell 1992; Nicholas and Crawford 2003). 

Isothermal microcalorimetry has been used to determine the activity of 

fungi after exposure to high and low temperature, oxygen depletion, and drying 

(Xie et al. 1997). 

Different stainings detect fungal hyphae and spores in woody tissue (Erb 

and Matheis 1983; Krahmer et al. 1986; Weiß et al. 2000). Treatment with 

safranine and astra blue stains lignified wood areas red and lignin-free parts 

blue, and thus differences between sound and decayed wood may become visible. 

Light-microscopic degradation patterns have been summarized (Schwarze 

et al. 1997). There is a key to identify wood decays based on light microscopic 

features (Anagnost 1998). 

Transmission (TEM) and raster electron microscopy (REM) result in detailed 

views of the cell wall decay by the various groups of fungi (Liese 1970; 

Daniel 1994). UV microspectrophotometry (UMSP) characterizes lignin and 

phenolic compounds in situ, determines their content semiquantitatively in 

the various layers of the wood cell wall (Koch and Kleist 2001), and has also 

been applied to measure lignin content after microbial wood attack (Bauch 

et al. 1976; Schmidt and Bauch 1980; Kleist and Seehann 1997; Kleist and 

Schmitt 2001). General wood quality, microbial activity in wood, and composition 

in fossil specimens may be quantified by chemical analyses of the wood 

cell wall components, by UV and IR spectroscopy, and by gas chromatography/mass 

spectrometry of lignin components (Faix et al. 1990, 1991; Nicholas 

and Crawford 2003; Schwanninger et al. 2004; Uçar et al. 2005). 

Biochemical methods to quantify microbial activity comprise assay of chitin 

as component of the fungal cell wall (Braid and Line 1981; Vignon et al. 

1986; Jones and Worrall 1995; Nilsson and Bjurman 1998) and ergosterol as 

fungal membrane component (Nilsson and Bjurman 1990; Pasanen et al. 1999; 

Dawson-Andoh 2002). 

Molecular methods to detect and identify fungi, like protein gel electrophoresis, 

immunology, and DNA-based techniques, are described in 

Chap. 2.4.2. 

8.3 

Tree Rots by Macrofungi 

There is a broad spectrum of macrofungi (macromycetes) affecting trees. Most 

fungi belong to the Homobasidiomycetes (Table 2.12). About 20 species have 

greater economic importance. Among them, the Agaricales are represented 

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by Armillaria. The other important fungi belong to the Aphyllophorales and 

there predominantly to the Polyporaceae sensu lato (“polypores”: Ryvarden 

and Gilbertson 1993, 1994). These polypores are summarized by the practical 

forester as “tree polypores” (Table 8.5; Seehann 1971). Fungi occurring on 

park and urban trees have been compiled e.g., by Seehann (1979), Wohlers et 

al. (2001), Wulf (2004) and Dujesiefken et al. (2005). Fungi affect predominantly 

older hardwoods and conifers of all climate zones. Infection occurs through 

wounds (wound parasites). Weakened trees may be more susceptible to fungi 

(weakness parasites). However, samples of dead wood from weakened spruces 

of different damage classes from forest dieback sites did not show differences in 

decayexperimentswith Heterobasidion annosum,Trametes versicolor (Schmidt 

et al. 1986), Coniophora puteana, Gloeophyllum abietinum and Oligoporus 

placenta (Liese 1986), compared to healthy trees. 

Fungi either penetrate via the roots (root rots) or the stem (stem rots). 

Root-decay Basidiomycetes are e.g., Armillaria species, Heterobasidion annosum, 

Meripilus giganteus, Phaeolus schweinitzii, andSparassis crispa. Among 

the Ascomycetes, Rhizina undulata (Pezizales) attacks the roots of spruce, 

pine and larch, and Kretzschmaria deusta (Xylariales) invades injured roots 

of beech, horse chestnut, elm, lime tree, maple, and plane causing white rot 

in the root and the stem (Butin 1995; Schwarze et al. 1995b; Baum 2001). 

Some common stem-decay Basidiomycetes in Europe (Butin 1995) and the 

USA (Zabel and Morrell 1992) are listed in Table 8.5. Most English names derive 

from Käärik (1978), Larsen and Rentmeester (1992) and Rune and Koch 

(1992). 

Fungi may attack the heartwood (heart rots) and effect thus a considerable 

strength and volume reduction of the tree xylem. They cause either brown or 

white rot in a several years of development, whereby all combinations between 

hardwoods and conifers as well as brown rot and white rot occur. However, 

also a soft-rot decay pattern may develop in the standing tree. Tree decay fungi 

have great economical importance, since a great part of the wood body can 

be devaluated, and felling of infected trees may be necessary. After felling, 

windthrow, or death of the tree, some fungi continue growth as saprobes in 

the wood for several years, then however usually die, that is, typically they 

do not endanger structural timber. The variously sized fruit bodies (basidiocarps, 

basidiomata) are either pileate, shelf-shaped, bracket-like, coral-like, 

or resupinate (see Fig. 2.17). Shape and size of the pores are distinguishing 

features (Breitenbach and Kränzlin 1986; Ryvarden and Gilbertson 1993, 1994; 

Krieglsteiner 2000). Beside fungi with annual fruit bodies, species with perennial 

basidiomes produce new hymenial layers each year and may become very 

large, hard and woody (see Fig. 8.15a). 

Daedalea quercina, Fomes fomentarius, Phellinus igniarius, Laetiporus sulphureus, 

Piptoporus betulinus, Polyporus squamosus, andMeripilus giganteus 

occur predominantly on hardwoods. Heterobasidion annosum, Phaeolus 

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Table 8.5. Some stem-decay Basidiomycetes 

Amylostereum areolatum (Chaill.: Fr.) Boidin white 

Armillaria mellea (Vahl: Fr.) Kummer, Honey fungus, and further Armillaria white 

species 

Bjerkandera adusta (Willd:Fr.)P.Karsten,Smokeypolypore white 

Chondrostereum purpureum (Pers.: Fr.) Pouzar, Silver-leaf fungus white 

Climacocystis borealis (Fr.: Fr.) Kotl. & Pouzar white 

Coniophora arida (Fr.: Fr.) P. Karsten brown 

Coniophora olivacea (Fr. Fr. ) P. Karsten brown 

Daedalea quercina (L.: Fr.) Fr., Maze-gill brown 

Daedaleopsis confragosa (Bolton: Fr.) J. Schröter white 

Fistulina hepatica (Schaeffer: Fr.) Fr., Beef-steak fungus brown 

Fomes fomentarius (L.: Fr.) Kickx, Tinder fungus white 

Fomitopsis pinicola (Sw.: Fr.) P. Karsten, Red-belted polypore brown 

Ganoderma adspersum (S. Schulzer) Donk, white 

Ganoderma applanatum (Pers.) Pat. white 

Ganoderma lipsiense (Batsch) G.F. Atk., Artist’s conk white 

Ganoderma lucidum (Curtis: Fr.) P. Karsten white 

Grifola frondosa (Dicks.: Fr.) S.F. Gray white 

Heterobasidion annosum (Fr.: Fr.) Bref., Root rot fungus white 

Inonotus dryadeus (Pers.: Fr.) Murr. white 

Inonotus hispidus (Bull.: Fr.) P. Karsten white 

Laetiporus sulphureus (Bull.: Fr.) Murr., Sulphur polypore brown 

Meripilus giganteus (Pers.: Fr.) P. Karsten, Giant polypore white 

Oligoporus stipticus (Pers.: Fr.) Kotl. & Pouzar brown 

Onnia tomentosa (Fr.: Fr.) P. Karsten white 

Phaeolus schweinitzii (Fr.: Fr.) Pat., Dye polypore brown 

Phellinus chrysoloma (Fr.) Donk white 

Phellinus hartigii (Allesch. & Schnabl) Pat. white 

Phellinus igniarius (L.: Fr.) Quélet, False tinder fungus white 

Phellinus pini (Brot.: Fr.) A. Ames, Ochre-orange hoof polypore white 

Phellinus pomaceus (Pers.: Fr.) Maire white 

Phellinus robustus (P. Karsten) Bourdot & Galzin white 

Pholiota squarrosa (Pers.: Fr.) Kummer white 

Piptoporus betulinus (Bull.: Fr.) P. Karsten, Birch polypore brown 

Pleurotus ostreatus (Jacq.) Kummer, Oyster mushroom white 

Polyporus squamosus (Hudson: Fr.) Fr., Scaly polypore white 

Resinicium bicolor (Alb. & Schwein.: Fr.) Parm. white 

Schizophyllum commune Fr.: Fr., Split-gill white 

Sparassis crispa Wulfen: Fr. brown 

Stereum rugosum (Pers: Fr.) Fr. white 

Stereum sanguinolentum (Alb. & Schwein.: Fr.) Fr., Bleeding Stereum white 

Trametes hirsuta (Wulfen: Fr.) Pilát white 

Tyromyces caesius (Schrader: Fr.) Murr., Blue cheese polypore brown 

Tyromyces stipticus (Pers.: Fr.) Kotl. & Pouzar brown 

Xylobolus frustulatus (Pers.: Fr.) Boidin, Ceramic parchment white 

Rot 

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schweinitzii, Phellinus pini, and Sparassis crispa inhabit softwoods. Species of 

Armillaria attack both tree groups. 

Inthefollowing,somecommontreefungiaredescribed,mostlyinnote 

form. For details see Seehann (1971, 1979) and textbooks e.g., by Butin (1995), 

Breitenbach and Kränzlin (1986, 1991), Rayner and Boddy (1988), Jahn (1990), 

Ryvarden and Gilbertson (1993, 1994), Krieglsteiner (2000), and Schwarze et al. 

(2004). 

8.3.1 

Armillaria Species, Honey Fungi 

The genus Armillaria (Fr.: Fr) Staude comprises worldwide about 40 species. 

The rather similar fungi form rhizomorphs in the soil and beneath the tree 

bark, the mycelium shines in the dark, the secondary mycelium is diploid 

and normally clampless (Marxmüller and Holdenrieder 2000). There are exannulate 

and annulate species (Shaw and Kile 1991; Guillamin et al. 1993). 

In Europe, five intersterility groups that had been referred to as A, B, C, D, 

E (Korhonen 1978b) within the annulate Armillaria mellea complex were assumed 

until the 1980s to be polymorphic members of the species Armillaria 

mellea (“Armillaria mellea complex”). In the 90s, the groups were assigned 

to five biological species (Guillaumin et al. 1993; Nierhaus-Wunderwald 1994; 

Holdenrieder 1996): 

A=Armillaria borealis Nordic honey fungus, 

B = Armillaria cepistipes, 

C = Armillaria ostoyae Dark honey fungus, 

D=Armillaria mellea s.s. Honey fungus, 

E = Armillaria gallica Marxm. & Romagn. 

Based on the verification of isolates by mating tests between monospore cultures, 

between diplonts and haplonts (Buller phenomenon), and by somatic 

compatibility tests, morphological variation of the fruit bodies of the five 

annulate European species was recently shown in color plates with suitable 

characters for species identification (Marxmüller and Holdenrieder 2000). In 

North America, nine annulate species are known (Anderson and Ullrich 1979; 

Anderson et al. 1980; Bruhn et al. 2000). The six species in Australasia (Kile and 

Watling 1983) are incompatible with European and North American species. 

In Africa, a subspecies of A. mellea was found (Agustian et al. 1994). 

Occurrence: The Armillaria species differ in host preference, pathogenicity 

(primary parasite, opportunist attacking weakened plants, destructive agent of 

non living tissue resulting in heart wood rot), geographical distribution, type 

and frequency of rhizomorphs, and in cultural characteristics such as mat 

morphology and optimum temperature (Rishbeth 1985, 1991; Shaw and Kile 

1991; Guillaumin et al. 1993; Marxmüller and Holdenrieder 2000; Schwarze 

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and Ferner 2003; Prospero et al. 2003). The damage, Armillaria root disease 

(Hartig 1874, 1882), occurs in conifers and hardwoods, particularly spruce, 

pine, maple, poplar, oak, in plantations of fruit, vine, flowers, ornamentals, 

and tropical cash crops (Seehann 1969; Schönhar 2002a; Schwarze and Ferner 

2003). The fungi occur also on stumps, piles, etc., and even in sprinkled wood 

(Metzler 1994). 

Physiology: Parasite, saprobe, white rot; slow growth in the laboratory; 

Characteristics: in pine and spruce, resin excretion; white, fan-like mycelial 

mats and brown-black, inside white rhizomorphs (0.25–4 mm; Schmid and 

Liese 1970; see Fig. 2.7) between bark and wood (Hartig 1874; Fig. 8.12a); 

wood colonized by living mycelium shining in the dark; clampless; 

Fig.8.12. Armillaria mellea. a Fruit 

bodies and rhizomorphs (translated 

from Hartig 1874); b White-rotten 

stump with rhizomorphs after removing 

the bark. c Fruit bodies and white 

mycelial sheet beneath the bark (photo 

W. Liese) 

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Fruit body (Fig. 8.12c): central stipe (to 15 cm), cap 5–15 cm in diameter; 

annual, in groups on stumps and at the root collar in late autumn; upper 

surface (A. mellea): small, yellow-brown scales on honey-yellow ground (Honey 

fungus); gill surface: cream-white to brownish-red gills; monomitic; clamps 

only at the basidium basis; pileus with white ring; young edible, danger of 

sickness when insufficiently cooked or overmatured; 

Significance: The Armillaria fungi, which are feared by the foresters, belong 

to the most important and cosmopolitan pathogens inside and outside 

the forest. They can attack almost all species of hardwoods and conifers of 

all ages (Hartig 1874; Schönhar 1989; Livingston 1990; Klein-Gebbinck and 

Blenis 1991; Gibbs et al. 2002). They live as saprobes in the soil on dead wood 

remainders and on stumps. The transition to the parasitic phase occurs, if the 

tree is weakened by stress (other parasites, wetness, dryness, pollution), so that 

forest damage sites showed increased occurrence of Armillaria. The infection 

occurs by rhizomorphs (Fig. 8.13). Solla et al. (2002) showed that A. mellea 

Fig.8.13. Development and transmission of Armillaria root disease (translated from 

Nierhaus-Wunderwald 1994, with permission of Swiss Federal Institute for Forest, Snow 

and Landscape Research) 

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and A. ostoyae penetrated Picea sitchensis root bark without prior wounding, 

but neither species formed rhizomorphs. The rhizomorphs grow in the soil 

from tree to tree and serve for nutrient translocation and infection. If the tree 

does not succeed in defending the fungus by histological or chemical barriers 

(Woodward 1992a; Wahlström and Johansson 1992), the fungus spreads between 

bark and xylem in the cambial region. The sap stream is interrupted, and 

toxic metabolites are excreted by the fungus. If the whole cambium is colonized 

around the stem, the tree dies rapidly (“cabium killer”). Beside the parasitic 

way of life, the fungus can spread via the wood rays in the heartwood of the 

root and stem basis (butt rot). Armillaria species and Heterobasidion annosum 

showed an increased occurrence in forest dieback sites (Kehr and Wulf 1993). 

AdirectcontrolofArmillaria spp. (e.g., Fox 1990) is practically impossible, 

particularly since the fungus occurs almost everywhere in the soil. In Oregon, 

the upper ground layer was colonized over an area of about 9 km 2 by only 

one mycelial clone of A. ostoyae, whose age was supposed to be 2,400 years. 

In England, a clone of A. gallica of about 500 years of age covered an area of 

9 ha. In France and Germany, clone diameter reached about 200 m in diameter 

(Marxmüller and Holdenrieder 2000). 

Armillaria is more frequent on soils with balanced microclimate and high 

air humidity at ground level as well as on nutrient-rich soils of about pH 5. 

Since young conifers are particularly susceptible on former hardwood soils, 

oldstumpsandrootsshouldberootedoutbeforeplantingconiferstolimit 

the vitality of the fungus, which, during its saprobic phase, depends on easily 

degradable nutrients (Butin 1995). Isolation of infected tree groups by 30 to 50cm-deep 

ditches is usually unsuccessful. Armillaria-infected plants in gardens 

and parks should be promptly removed. The resistance of the plant hosts can 

be increased by suitable soil preparation, good planting, and tree care. Douglas 

fir, Sitka spruce, fir and larch are lesser susceptible species. The application of 

chemicals within the root range is strenuous and therefore only suitable for 

valuable garden and park trees (Schönhar 1989). 

Pinosylvin from Pinus strobus inhibited mycelial growth of A. ostoyae 

(Mwangi et al. 1990). Growth rate, spread and survival of rhizomorphs decreased 

by several bacteria, particularly Pseudomonas fluorescens Migula (Dumas 

1992), Trichoderma species (Dumas and Boyonoski 1992), wood-inhabiting 

Basidiomycetes (Pearce 1990) and mycorrhizal fungi (Kutscheidt 1992). 

8.3.2 

Heterobasidion annosum s.l. Root Rot Fungus, Fomes Butt Rot 

From the Root rot fungus, several intersterility groups have been distinguished, 

which differ in relation to distribution, fruit body morphology and host tree 

(Korhonen 1978a). In Europe, three groups have been referred to as P-group 

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(pine), S-group (spruce), and F-group (fir) (Holdenrieder 1989; Siepmann 

1989; Capretti et al. 1990; Stenlid and Karlsson 1991; Korhonen et al. 1992). In 

North America occur the P- and S-type. The Asian forms are lesser characterized 

(e.g., Dai and Korhonen 1999). The three European forms show significant 

differences in their distribution and host preference and have been attributed 

to three distinct species (Niemelä and Korhonen 1998; Korhonen and Holdenrieder 

2005): 

Heterobasidion annosum s.s. corresponds to the European P-type of H. 

annosum s.l. and may named pine root rot fungus, as it typically occurs 

in pine forests. In addition, the fungus attacks Juniperus communis, Picea 

abies, P. sitchensis, Pseudotsuga menziesii, Larix decidua, L. x eurolepsis, and 

L. kaempferi. The distribution area covers the whole of Europe except for the 

most northern forests and possibly the great parts of Siberia. 

Heterobasidion parviporum (European S-type of H. annosum s.l.; Spruce 

root rot fungus) occurs in Europe nearly exclusively on Picea abies, but as it 

seems, it is not found in Western Europe. In Russia, it attacks also Abies sibirica 

and in East Asia further Picea and Abies species. 

Heterobasidion abietinum (European F-type of H. annosum s.l.; Fir root 

rot fungus) occurs in fir forests from the Pyrenees to South Polonia and the 

Caucasus, particularly on Abies alba, but also on A. borisii-regis, A. cephalonica 

and A. nordmanniana. 

The three closely related species can be differentiated by cultural studies, 

mating tests and DNA techniques. The hymenium of H. parviporum has small 

pores (up to 5 pores/mm) and the upper side shows short hairs, while H. annosum 

s.s. has bigger pores and a bald upper side. The features of H. abietinum 

often overlap with those of the two former species, but its occurrence on firs 

is a suitable clue (Korhonen and Holdenrieder 2005). Hybridization of the 

species occurs in the laboratory. A natural hybrid between S- and P-type has 

been found in North America, but generally, hybrids occur more easily between 

forms from different continents. Regarding the evolution of H. annosum 

s.l., the origin of H. parviporum and H. abietinum seems to be East Asia, as 

there occurs a form that showed high compatibility with all three species. Assumably, 

H. annosum s.l. spread from the eastern Himalayas and has thereby 

increasingly differentiated via different routes: H. abietinum arrived in Europe 

via the South Asian conifer forests, H. parviporum via northern Asia, and the 

American S-type reached North America over the Bering Strait. Not much is 

known on the P-types (Korhonen and Holdenrieder 2005). Molecular analyses 

have shown a close relation of the genus Heterobasidion to the Russulales. 

The following description concerns H. annosum s.l. 

Occurrence: common in Europe, North America; predominantly conifers; in 

heartwood and rootwood of spruce, larch and Douglas fir; in pine restricted to 

the root area due to greater resin content; broad host range of over 200 woody 

plants (Heydeck 2000); largest diameter of a genet smaller than 30 m, only in 

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single cases up to 55 m; maximum age of an individual genet around 200 years 

(Queloz and Holdenrieder 2005); 

Physiology: white rot, root rot, butt rot, so-called red rot due to reddish 

discoloration of the wood; at initial decay preferential lignin degradation, later 

simultaneous white rot (Peek and Liese 1976); parasite and saprobe; 

Characteristics: anamorph Spiniger meineckellus (A.J. Olson) Stalp. 

(Fig. 8.14C) on agar and fresh wood samples at high relative humidity: clubshaped 

thickened conidiophore after spore dispersal like a morning star 

(“Brefeld conidia” as identification feature: Brefeld 1889); flask-shaped increase 

of the stem basis of spruce by cambial irritation; resin excretion; 

Fruit body (Fig. 8.14A): annual to enduring crusty brackets in autumn, 

often resupinate (1 cm thick, 3–20 cm wide) in rows and roofing tile-similar, 

usually fused, at the stem basis and on flat-running roots, frequently covered 

by needle litter; yearly a new pore layer; fresh: tough, old: hard and woody; 

upper surface: bumpy-wrinkled, brown, often zonate, leathery-crusty, whiteyellowish 

margin; pore surface: white-cream with circular-angular pores (4– 

5/mm); dimitic; bipolar. 

Significance: The fungus is one of the most important pathogens in coniferous 

forests of temperate regions (Hartig 1874, 1878; Rishbeth 1950, 1951; 

Zycha et al. 1976; Hallaksela 1984; Tamminen 1985; Benizry et al. 1988; Schönhar 

1990; Woodward 1992a, 1992b; LaFlamme 1994; Woodward et al. 1998; 

Heydeck 2000; Greig et al. 2001; Gibbs et al. 2002; Korhonen and Holdenrieder 

2005), which causes substantial damage particularly in older forests. The infection 

occurs by germinating spores or by mycelium that is already present 

in roots or soil. Several infection ways are possible: by basidiospores (also 

conidia) via stump infection (Redfern et al. 1997), by mycelial growth through 

root graft transmission from diseased to healthy roots (Hartig 1878; Schönhar 

2001), or via spores [germinable about 1 year: Brefeld (1889)], which are 

washed into the soil by rain and germinate on the roots. The fungus penetrates 

into older roots through wounds and into young uninjured roots through the 

thin bark (Rishbeth 1951; Peek et al. 1972a, 1972b; Lindberg and Johansson 

1991; Lindberg 1992; Solla et al. 2002). The hyphae penetrate into sound spruce 

roots via the pit channels of the thick-walled stone cork cells. The walls of the 

following thin-walled stone cork cells and the sponge cork cells are degraded. 

The fungus colonizes the tracheids from the bark rays via the wood rays. The 

tracheids are degraded by enzymes and perforated by microhyphae (Peek and 

Liese 1976). Embryos of Pinus spp. showed three days after artificial inoculation 

intercellular penetration of hyphae through the epidermis and into the cortex 

(Nsolomo and Woodward 1997). Infection of spruce seeds of 4–7 days of age 

showed that infective structures on the root surfaces were evident 24 h after 

inoculation. Internal colonization of cortical tissues started after 24–48 h and 

reached the endodermis within 72 h. Severe destruction of stelar cells occurred 

12–15 days postinfection (Asiegbu et al. 1993). Infection of nonsuberized and 

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Fig.8.14. Fomes root rot by Heterobasidion annosum. A Fruit bodies at the stem basis. B 

Sequent sections of a stem showing the different color and decay zones (photos W. Liese); 

C Pathogenesis, a longitudinal section through a spruce with heart rot, with stem cross 

sections, b cross section through a stem at an early stage of disease, c a late stage in the 

wood decay, d fruit bodies, e Brefeld conidiophores with conidia, f a heart rot caused by 

Armillaria sp. shown for comparison (from Butin 1995, by permission of Oxford University 

Press) 

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young suberized roots of spruce seedlings showed host reaction to delimit the 

infection by the formation of a necrotic ring barrier in the outer cortex. In cases 

where the inner cortex became infected, hyphae accumulated just before the 

endodermis, which acted as a new barrier. Only in nonsuberized roots, the stele 

was almost completely digested within 3 days after inoculation (Heneen et al. 

1994a). In woody roots 2–4 mm in diameter, fungal infection was restricted 

to the remnant cortex cells and the rhytidome after an incubation period of 

20 days; accumulation of granular materials prevailed in the infected periderm 

cells, which enclosed degenerated hyphae, both leading to the conclusion that 

the rhytidome acts as a successful barrier to infection of the inner parts of 

the root for at least 20 days following inoculation (Heneen et al. 1994b). Stem 

infections are rare and limited to wounds at the root collar (Schönhar 1990). 

Main infection is by airborne basidiospores that germinate on fresh stump 

surfaces. Infection of neighboring trees occurs by vegetative mycelia via root 

contacts. Once established in the root system, the fungus can remain active 

for about 60 years. The fungus spreads into trees of the next generation from 

infected stumps (Vasiliauskas and Stenlid 1998). 

The significance of the fungus is not only based on its parasitic capability 

to kill living roots, but it is at the same time causal agent of “red rot”, which 

ascends in the heartwood (heart rot) of the stem and is economically usually 

more serious. In Europe, on average, a 10% stem wood devaluation is counted 

for spruce by “red rot”. In Scotland, the fungus is responsible for 90% of 

losses due to rot (Blanchette and Biggs 1992). The yearly damage in Germany 

amounts to e56 million (Dimitri and Tomiczek 1998) and in the EU countries 

to about e500 million (Woodward et al. 1998). “Red rot” increased in forest 

dieback sites. 

The parasitic phase of the fungus develops first as root rot. In pine, the 

fungus predominantly grows stemwards in the root cambium area, until it is 

stopped by resin formation and a bark wound periderm. Large root parts die 

off. In the less resinous spruce, fir, larch and Douglas fir, fungal activity shifts, 

as soon as the mycelium reaches roots of more than 2 cm in diameter, into the 

root interior, that is, side roots and thus also the infected tree remain alive. 

Only if all roots are colonized, the mycelium also grows into the cambium and 

kills the tree. 

The saprobic phase begins with penetration in the heartwood. Sapwood 

colonization occurs only after felling due to reduction of moisture content and 

particularly due to inhibition by the living sapwood (Shain and Hillis 1971). 

The effects of heartwood colonization depend on the tree species. In pine, the 

fungus spreads usually only insignificantly in the stem, but the tree dies due to 

the root damage. In larch, the mycelium grows in the heartwood/sapwood area 

and reaches likewise only low stem height. In spruce, the fungus climbs up in 

the stem 25–40 cm/year (Stenlid and Redfern 1998). Likewise, the Douglas fir 

stem can be colonized. The infected wood shows first a “1. color zone” (grey- 

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violet striping), then a “bright hard rot” (light brownish, wood still firm), later 

a “dark hard rot” (brownish-red, only wood structure remaining) and finally 

a “soft rot” (Fig. 8.14B; Zycha 1964), where the wood is fibrously dissolved 

and interspersed with small, white spindle-like nests with a black center of 

manganese deposits (Fig. 8.14C) (Hartig 1978; see Chap. 7.2). 

Imperiled for H. annosum are first plantings on formerly agriculturally 

used pasture soils and arable lands (“field-dying”, German: “Ackersterbe”). 

Conifers on base-rich and compacted ground, and on sites with very variable 

moisture content suffer more from the disease than those on acidic, more open 

soils with a more uniform water supply (Butin 1995; Schönhar 1997; Heinsdorf 

and Heydeck 1998). The inhibition of acidophilic, antagonistic mycorrhizas 

may play a role. A direct control is difficult, and only preventing measures are 

used (Schönhar 1990, 2002b). Rooting out and removing the infected stumps 

as well as isolating the infected sites by ditches are difficult and not always 

successful (Schönhar 1989). The most effective measure is to perform thinnings 

during the wintertime, as spore infection decreases during frost (Korhonen 

and Holdenrieder 2005). In not-yet-infected first plantings, the stumps which 

are the starting point for a propagation of the fungus via root grafts, have 

been coated on the fresh surface with carbolineum, which however delays the 

stumpdecomposition.Immediatetreatmentofthefreshsurfacewithasodium 

nitrite solution prevented spore germination of H. annosum.Aschemical,also 

urea (Schönhar 2002b) and boron compounds are used (Pratt 1996). Originally 

in the U.K and later in Scandinavia and further European countries, a spore 

solution of the antagonistic fungus Phlebiopsis gigantea is immediately applied 

to the fresh stump surface of pines (Meredith 1959; Rishbeth 1963; Schwantes 

et al. 1976; Lipponen 1991; Gibbs et al. 2002) and spruce (Korhonen et al. 1994; 

Holdenrieder et al. 1997). There are spore preparations, which are specifically 

suited for spruce, but generally, P. gigantea is more suitable for pines. The 

wood can be automatically inoculated with spores through holes in the saw 

blade of the harvester (Metzler et al. 2005). The antagonist overgrows the 

stump cross surface, so that H. annosum cannot colonize it by spores. Thus, an 

infection of neighboring trees over root grafts is prevented. Further antagonists 

to H. annosum are treated by Holdenrieder and Greig (1998) and compiled by 

Woodward et al. (1998). 

Root graft transmission can be reduced by far planting faces and admixture 

of hardwoods. Lesser sensitive hardwoods as well as fir or larch should be 

selected for particularly endangered sites instead of spruce and pine. In vitro, 

mycelial growth was inhibited by stilbenes, flavonoids and lignans (Zycha et al. 

1976; Shain and Hillis 1971; Yamada 1992). Breeding attempts with the aim of 

red-rot resistant tree clones were performed, but did yet not reach a practical 

use. Recent resistance research mainly deals with the genetic mechanisms of resistance 

and the physiology of defense reactions (Korhonen and Holdenrieder 

2005). Viruses in the root rot fungus, which are morphologically similar to the 

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Cryphonectria-hypovirus (Chap. 8.1.1.2), only reduced spore germination of 

the fungus. 

8.3.3 

Stereum sanguinolentum, Bleeding Stereum, 

Bleeding Conifer Parchment 

Occurrence: conifers, particularly spruce; as saprobe causing red streaking 

discoloration (see Fig. 6.4a); 

Significance: white rot, most important fungus involved in “Wound rot of 

spruce” (Butin 1995); 2/3 of about 20% of annual harvest of fir wood with fungal 

damage affected by wound rots, particularly by S. sanguinolentum (Schönhar 

1989); wounds often due to mechanized wood harvest or bark damage by game; 

infection of the opened wood body by spores; also transmission of mycelial 

fragments by woodwasps (Sirex spp.); small and superficial wounds often 

closed by resin excretion; extension of white rot in the outer stem wood with 

reddish discoloration; fast rot extension (20 cm/year) in the first years after 

infection; rot spreads more rapidly after injuries at the root collar than after 

wounding the stem or small roots; injured roots of less than 2 cm in diameter 

and wounds in more than 1-m distance of the stem foot hardly lead to stem rot. 

To prevent wound rot by S. sanguinolentum, tree harvest should be done 

carefully and injuries treated with a wound dressing. Amylostereum species 

maybealsoinvolvedinwounddecayofspruceandotherconifers,A. areolatum 

and A. chailletii, both also being associated with woodwasps (Vasiliauskas 

1999). 

8.3.4 

Fomes fomentarius, Tinder Fungus, Hoof Fungus 

Occurrence: common, circumboreal, south to North Africa, through Asia to 

eastern North America; mostly hardwoods, common on birches in the north 

and on beeches in the south, also on oak, lime tree, maple, poplar, and willow, 

rarely on alder and hornbeam, exceptionally on softwoods (Schwarze 1994, 

2001); 

Fruit body (Fig. 8.15a): perennial (over 30 years, increase in early summer 

to autumn), thick, large (to 50 cm in diameter), hard brackets, mostly solitary; 

often high at the stem; firmly attached to the bark; upper surface: light brown to 

blackish-grey, bulging-zonate; pore surface: flat, cream-brownish hymenium 

with white margin; circular pores (4–5/mm); trimitic; soft-tough trama beneath 

a 1 to 2-mm-thick hard crust; 1–3 new hymenial layers per year; up to 

240 million spores per cm 2 hymenium and hour; tetrapolar. In former times 

(e.g., in Haitabu), the trama was soaked with salpetre for tinder production. 

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Fig.8.15. Fruit bodies of tree decay fungi. a Fomes fomentarius. b Laetiporus sulphureus. 

c Meripilus giganteus. d Phaeolus schweinitzii. e Phellinus pini. f Piptoporus betulinus. 

g Polyporus squamosus. h Sparassis crispa (photos T. Huckfeldt) 

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A part of a fruit body was found at the “Ötzi” mummy. Still, around 1890, about 

50 t of trama per year were sampled in the Bavarian, Bohemian and Thuringian 

forests for fire igniting, as styptic, and for the production of hoods, gloves and 

trousers (Hübsch 1991; Scholian 1996). 

Significance: one of the most remarkable “large polypores”; infection of 

weakened and old trees via bark wounds or branch breakings; natural member 

in the biocoenosis of birch and beech forests; simultaneous white rot with 

black demarcation lines; at final stage, danger of windthrow; involved in the 

final decay of beech bark-diseased trees; saprobe on thrown or felled trees for 

several years (“Verstocken”). 

8.3.5 

Laetiporus sulphureus, Sulphur Polypore, Giant Sulphur Clump 

Occurrence: cosmopolitan, Europe, western North America, northeast Asia; 

preferentially on hardwoods with colored heartwood, like oak and robinia, 

also apple, beech, cherry, elm, lime tree, maple, pear, plum, poplar, willow, 

rarely on conifers; common on park and urban trees (Schwarze 2002); 

Fruit body (Fig. 8.15b): annual (summer to autumn), conspicuous (upper 

surface: sulfur-yellow to reddish) wavy-velvety brackets (15–40 cm); pore surface: 

sulfur-yellow with angular pores (3–4/mm); single or in clusters; fresh: 

succulent-soft, later: inflexible, chalk-like, straw-colored to grey; dimitic; eaten 

in North America; 

Significance: infection of the stemwood usually via wounds; brown rot in 

the heartwood; yellowish mycelium in broad, bind-like strips along the tears 

and shakes that develop in the wood; sapwood usually not attacked; infected 

trees alive for many years till broken or thrown by storm; rarely saprobic, e.g., 

on wooden boats. 

8.3.6 

Meripilus giganteus, Giant Polypore 

Occurrence: circumboreal in the northern hemisphere, but nowhere common; 

usually hardwoods, particularly horse chestnut, beech, lime and oak; often on 

park and urban trees (Seehann 1979; Schwarze 2003); 

Fruit body (Fig. 8.15c): annual (summer to autumn) on stumps of freshly 

felled trees and at the basis of standing trees; often apparently growing from 

the ground, but always in contact to wood; large and pileate with fan-shaped 

to spatulate pilei from a common base; aggregates to 1 m in diameter and 70 kg 

fresh weight; upper surface: cream-white to yellow-brown zonate; pore surface: 

cream to yellow-orange-brown pores (3–5/mm), rapidly blackish when 

bruised or cut; monomitic; eaten in Japan; 

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Significance: white rot in damaged roots of usually older trees, weakened 

due to compressed soil, asphalting, salting, and by wounds due to building 

operations or road traffic; fruit bodies indicate a heavily destroyed root system 

leaving only little time to the trees for surviving; stem hardly infected. Tree 

care in the urban area reduces the damage. 

8.3.7 

Phaeolus schweinitzii, Dye Polypore, Velvet Top Fungus 

Occurrence: circumglobal, in European conifer forests north to 70 °N in Finnmark, 

Norway, particularly pine, Douglas fir, also spruce and larch, rarely on 

hardwoods (Ryvarden and Gilbertson 1993); 

Fruit body (Fig. 8.15d): annual (late summer), easy-passing; at the stem 

basis or on the soil on hidden roots; stipitate, short, central, upward more 

thick, cylindrical to knotty stipe, first with spinning-top-like, later with several 

tile-like caps (to 40 cm); on the cross cuts of felled trees with lateral stipe; 

frequently including plant residues or small branches during ripening; upper 

surface: when young orange, later yellowish-brown, old often black; yellowbrown 

margin; woolly; pore surface: angular pore (1–2/mm) layer at first 

orange,latergreenishtorustybrown,discolorswithpressurered-brownish; 

monomitic; 

Significance: brown rot, major cause of butt rot in the heartwood of old 

pine and Douglas fir; frequently in conifers forests on former hardwood soils 

(Schönhar1989);firstonroots andinstemwounds,laterinthe stemheartwood, 

less ascending the stem (butt rot); decayed wood and laboratory cultures with 

turpentine smell; saprobe on dead trees, stumps and logs for several years. 

8.3.8 

Phellinus pini, Ochre-Orange Hoof Polypore 

Occurrence: circumglobal, widespread in northeast Europe on pine, in North 

America and Asia on other conifers as well (Heydeck 1997; Frommhold and 

Heydeck 1988); 

Fruit body (Fig. 8.15e): perennial (to 50 years), brackets only 5–20 years 

after infection near branch holes and stubs; often high at the stem of old 

trees (Naumann 1995), 5–12 cm; upper surface: zonate, rough, cracked, at 

first rust-brown, hirsute, later dark-brown-blackish, glabrous and encrusted; 

pore surface: yellow to grey-brown with round to angular/daedaleoid pores 

(1–3/mm); dimitic, bipolar; 

Significance: infection of old (30–50 years) pine and larch at exposed heartwood 

(branch stubs, wounds); living sapwood usually not penetrated; often 

high at the stem (Hartig 1874; Liese and Schmid 1966); from deep-reaching 

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dead branches decay upwards and downwards in the stem; white-pocket rot, 

preference for latewood of Pinus and Larix (Liese and Schmid 1966; Blanchette 

1980), pockets in some hosts concentrated in the earlywood bands (“laminated 

rot”, ring shake); occurrence of transpressoria and formation of cavity-shaped 

decay pattern (Liese and Schmid 1966); local bark deepenings, outer sapwood 

resin-infiltrated (in former times wood used as resinous wood); in spruce, 

infection also via sapwood; wood still relatively firm at early decay; dying after 

tree felling. 

8.3.9 

Piptoporus betulinus, Birch Polypore, Birch Conk Fungus 

Occurrence: circumboreal, north to Norwegian North Cape at 71 °N (Ryvarden 

and Gilbertson 1994); only birch; also in gardens and parks; 

Fruit body (Fig. 8.15f): annual (summer to late autumn), but enduring; 

solitary and in groups; shell-shaped, fan-like brackets (8–30cm); pilei pendent, 

dimidiate, or reniform; often several meters high on the stem; upper surface: 

dull-smooth, unzonate, young cream-white, later ochre-brown to grey-brown, 

old usually cracked; pore surface: white to cream-brownish circular to angular 

pores (3–5/mm); dimitic; some isolates bipolar (Stalpers 1978); fruit body 

previously used in Fennoscandia as a cushion for knives, which do not rust 

while standing in the fruit body; 

Significance: weakness-parasite, host-specific on older and weakened (e.g., 

lack of light) birch; infection via wounds (branch breakage); brown rot; danger 

of windthrow. 

8.3.10 

Polyporus squamosus, Scaly Polypore 

Occurrence: circumpolar in Europe (north to Finnmark at 70 °N), Australia, 

Asia, and America; hardwoods such as ash, beech, elm, horse chestnut, lime, 

maple, planetree, poplar, and willow (Schwarze 2005); frequently on urban and 

park trees; 

Fruit body (Fig. 8.15g): annual (early summer); solitary or in groups from 

a branched base; usually laterally stipitate, with circle to fan-like cap (to 80 cm 

wide and 2 kg fresh weight); upper surface: yellow-ochre with concentrically 

arranged light to dark-brown, scale-like patches, smooth and sticky; pore 

surface: cream-yellowish with angular-oval pores (1–2/mm); whitish stipe (up 

to 10 cm) at the basis dark-brown to black-felty; dimitic; tetrapolar (Stalpers 

1978); young edible; 

Significance: white rot in the heartwood of living and dead hardwoods with 

black demarcation lines after penetration through wounds. 

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8.3.11 

Sparassis crispa 

Occurrence: rare in Europe; particularly pine, also Douglas fir, spruce and fir; 

Fruit body (Fig. 8.15h): annual (summer to late autumn); solitary at the root 

area of living pines, lateral and at cross surface of stumps and fallen stems; 

hemispherical to cushion-shaped; resembling a large (up to 70 cm and 6 kg 

fresh weight) sponge, cauliflower, or coral (German: “Krause Glucke”); consisting 

of numerous, wavy, narrow upright-standing branches deriving from 

a fleshy stalk; frizzy, leaf-like branch-ends partly growing together, similar to 

Icelandic moss; surface: smooth, cream, later ochre, when old with brown margin, 

finally completely brown; hymenium on the outside, downward arranged 

side of the branches; monomitic; when young well edible mushroom (in Germany 

certified as market fungus) with whitish meat, spicy morel-similar smell 

and nut-like taste; fruit bodies also on agar cultures; some isolates tetrapolar 

(Stalpers 1978); 

Significance: parasitically in roots of older pines, ascending to 3 m high 

with brown rot in the stem heartwood; decayed wood with turpentine smell; 

economically important wood losses in pine and Douglas fir (Heydeck 1994). 

8.4 

Damage to Stored Wood and Structural Timber Outdoors 

After felling or falling of a tree, the living cells die some time later. The active 

defensesystemsdonotfunctionanylonger.Somefungithatarealreadypresent 

in the stem can continue degradation by their now saprobic way of life, e.g., 

Fomes fomentarius. The exposed wood surfaces however rapidly dry, and new 

ecological conditions develop. Thus, the stem usually provides a new energyrich 

substrate for rapid colonization by several saprobic organisms (Zabel and 

Morrell 1992). 

Colonization and discolorations of the stem in the forest occur frequently 

within short time by bacteria, algae, slime fungi, molds, and blue-stain and 

red-streaking fungi. After longer exposure wood decays by brown, white 

and soft-rot fungi develop, which may be summarized as “decay of stored 

wood”, or “colonization of fallen and cut wood” (Rayner and Boddy 1988). 

Among the Basidiomycetes are e.g., Armillaria gallica, Bjerkandera adusta, 

Chondrostereum purpureum, Fomes fomentarius, Stereum spp., Schizophyllum 

commune and Trametes versicolor. Several fungi are involved in the decomposition 

of the stumps remaining in the soil e.g., Armillaria spp., B. adusta, 

C. purpureum, Daedalea quercina, Fistulina hepatica, Ganoderma spp., Gloeophyllum 

spp., Grifola frondosa, Heterobasidion annosum, Meripilus giganteus, 

Phaeolus schweinitzii, Phlebiopsis gigantea, Pleurotus ostreatus, Stereum spp., 

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8.4 Damage to Stored Wood and Structural Timber Outdoors 201 

S. commune and T. versicolor. On tree residues remaining in the forest (top, 

branches) grow e.g., B. adusta, C. purpureum, Coniophora puteana, Gloeophyllum 

sepiarium, Stereum sanguinolentum and T. versicolor. Forest-litter 

degrading Basidiomycetes were described by Frankland et al. (1982). 

Damages on roundwood (logs, poles) and boards may occur during transport 

and inappropriate storage e.g., by C. puteana, Fomitopsis pinicola, Gloeophyllum 

trabeum, Paxillus panuoides, Phlebiopsis gigantea, S. sanguinolentum 

and Trichaptum abietinum. Wood chips are damaged by B. adusta, Gloeophyllum 

spp., Phanerochaete chrysosporium, T. versicolor, and by several Deuteroand 

Ascomycetes (molds, blue-stain and soft rot fungi). Several bacteria, yeasts, 

Deuteromycetes and Ascomycetes were found in stored annual plant residues, 

like sugarcane bagasse (Schmidt and Walter 1978). 

Yeasts commonly colonize twigs, leaves, litter, and humus, are however also 

found on freshly sawn lumber (Mikluscak et al. 2005). 

Structural timber that is used outdoors in ground contact, like sleepers, 

poles, posts, fences, bridges and garden furniture, is attacked by soft-rot fungi if 

it is insufficiently treated with wood preservatives. Among the Basidiomycetes 

occur e.g., Antrodia vaillantii, H. annosum, Lentinus lepideus, Leucogyrophana 

pinastri, Oligoporus placenta, Phanerochaete sordaria, Phlebiopsis gigantea, 

Serpula himantioides, Sistotrema brinkmanni, Trametes versicolor and Trichaptum 

abietinum (e.g., Lombard and Chamuris 1990; Morrell et al. 1996). 

Mine timber was decayed by A. vaillantii and C. puteana as well as by 

Armillaria spp., G. sepiarium, H. annosum, L. lepideus, L. pinastri, O. placenta, 

Paxillus panuoides, Schizophyllum commune, Serpula lacrymans, Stereum spp. 

and T. versicolor (Eslyn and Lombard 1983). Earliella scrabosa, Loweporus 

lividus, Rigidoporus lineatus, and R. vinctus were isolated from gold mine 

poles in India (Narayanappa 2005). 

Wood in fresh water, like in cooling towers, is often destroyed by soft-rot 

fungi. Among the Basidiomycetes, e.g., Donkioporia expansa and Physisporinus 

vitreus have been isolated from cooling-tower woods (v. Acker and Stevens 

1996). The latter fungus degraded pine sapwood samples that showed a final 

moisture content of up to 320% u (Schmidt et al. 1996). Schwarze and 

Landmesser (2000) hypothesized that the preferential degradation of tracheidal 

pit membranes is associated with the adaptation of this fungus to very wet 

substrates. Wood in salt water below (not permanent) the sea level, as in harbor 

constructions, is predominantly attacked by Deuteromycetes and Ascomycetes 

and rarely by Basidiomycetes (Jones et al. 1976; Kohlmeyer 1977; Leightley 

and Eaton 1980). Basidiomycetes, like Antrodia xantha, Daedalea quercina, 

Gloeophyllum sepiarium, Laetiporus sulphureus, Lentinus lepideus, Phlebiopsis 

gigantea, Schizophyllum commune and Xylobolus frustulatus dominate in wood 

above the water level, like in docks, stakes or boats (Rayner and Boddy 1988). 

Damages on stored and structural timber in outside use can be reduced or 

even avoided by means of protection measures against fungal activity described 

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inChap.6.4:winterfelling,shortandadequatestorageofthefreshroundwood, 

wet storage, rapid drying, storage in a gas atmosphere (N2/CO2), and storage 

of cut timber in well-ventilated piles with protection against rain as well as 

chemical protection. 

In the following, some common Basidiomycetes on wood in outside use are 

described, mostly in note form. For details see also Grosser (1985), Breitenbach 

and Kränzlin (1986, 1991), Zabel and Morrell (1992), Eaton and Hale (1993), 

Ryvarden and Gilbertson (1993, 1994), Bech-Andersen (1995), Butin (1995), 

Kempe (2003), Krieglsteiner (2000), and Weiß et al. (2000). 

8.4.1 

Daedalea quercina, Maze-Gill, Thick-Maze Oak Polypore 

Occurrence: circumglobal and throughout Europe, North America, North and 

Central Asia, North Africa; in northern Europe only on oaks, in central and 

southern Europe also on Acer, Carpinus, Castanea, Chamaecyparis, Corylus, 

Eucalyptus, Fagus, Fraxinus, Juglans, Juniperus, Populus, Picea, Prunus, 

Robinia, Sorbus, Tilia, and Ulmus (Wa˙zny and Brodziak 1981); 

Fruit body (Fig. 8.16h): perennial, single or fused, broadly sessile, dimidiate, 

flat or ungulate, sometimes imbricate, sometimes nodular or deformed, large 

brackets (up to 30 cm wide and 8 cm thick) often high at the stem; hard and 

corky to woody; upper surface: grooved, uneven, covered with nodes, glabrous 

or somewhat pubescent, cream, ochraceous grey to brown; pore surface: sinuous, 

or daedaleoid to labyrinthine, or almost lamellate, pores 1–4 mm wide 

measured tangentially, walls up to 3 mm thick; monstrous fructification in the 

dark; trimitic; bipolar; 

Significance: brown rot in the durable heartwood of oaks and other hardwoods; 

on wounded standing trees via exposed heartwood, dead branches, 

on stumps, fallen stems, on sleepers, poles, stakes, wooden bridges, mine 

timber; occasionally in buildings on weathered timber, like window sills and 

half-timbering. 

8.4.2 

Gloeophyllum Species, Gill Polypores 

Three Gloeophyllum species are relevant to wood. The fungi have similar fruit 

bodies and life conditions (Hof 1981a, 1981b, 1981c; Grosser 1985; also Bavendamm 

1952a), and are thus usually united as “wood gill polypores”. They 

are widespread in Europe, North America, North Africa, and Asia on conifers 

and hardwoods. Gloeophyllum abietinum is a somewhat southern species, G. 

trabeum a southern species. 

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Fig.8.16. Fruit bodies of decay fungi on stored wood and on timber in outdoor use. Gloeophyllum 

abietinum. a Upper side. b Lower side. c Darkness fruit bodies; Gloeophyllum 

sepiarium d Upper side. e Lower side; Gloeophyllum trabeum f Upper side. g Lower side. 

h Daedalea quercina; i Lentinus lepideus; j Paxillus panuoides; Schizophyllum commune 

k Upper side. l Lower side. m Trametes versicolor (photos T. Huckfeldt) 

Gloeophyllum abietinum, Fir Gill Polypore 

Fruit body (Fig. 8.16a,b): perennial, pileate (2–8 cm wide), broadly attached, 

ofteninrowsortile-like,ontimberlowersideresupinate;uppersurfacehirsute 

to velutinate, in age zonate, scrupose to warted or smooth, rusty yellow, 

reddish-brown to dark grey and black when old, when young whitish-yellowbrown, 

wavy, sharp margin; hymenophore ochre-grey brown, wavy lamellae 

(8–13/cm, behind the margin) with anastomosing, serrate, mixed with poroid 

areas; monstrous fruit bodies in the dark (Fig. 8.16c); trimitic; bipolar; 

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Strands: only rarely on timber in laboratory culture, cream-ochre-dark 

brown; fibers to dark brown; no vessels. 

Gloeophyllum sepiarium, Yellow-Red Gill Polypore 

Fruit body: (Fig. 8.16d, e) annual to perennial, pileate, broadly sessile, dimidiate, 

rosette shaped, often imbricate in clusters from a common base or fused 

laterally, to 7 cm wide, 12 cm long and 6–8 mm thick, margin slightly wavy; 

upper surface when young yellowish brown, then reddish brown and grey to 

black when old; scrupose, warted to hispid, finally zonate often differently 

colored; hymenophore with straight lamellae (15–20/cm, behind the margin), 

edges of lamellae golden brown in active growth, later umber brown, side surface 

of lamellae ochre-brown; usually mixed with daedaleoid to sinuous pore 

areas (1–2/mm); monstrous fruit bodies in the dark; trimitic; bipolar; 

Strands: only rarely on timber in laboratory culture, white-cream; fibers 

yellow to brown, no vessels. 

Gloeophyllum trabeum, Timber Gill Polypore 

Fruit body (Fig. 8.16f, g): annual to perennial, pileate, sessile, imbricate with 

several basidiomes from a common base or elongated and fused along wood 

cracks, to 3 cm wide, 8 cm long, 8 mm thick; upper surface soft and smooth, 

hazelnut to umber brown to grayish when old, weakly zonate to almost azonate, 

lighter margin; hymenophore semi-lamellate or labyrinthine to partly poroid 

(2–4/mm), rarely lamellate specimens with up to four lamellae/mm along the 

margin, ochre to umber brown; monstrous fruit bodies in the dark; dimitic; 

bipolar; 

Strands: only on timber in laboratory culture, white-beige to yellow-orangegrey 

brown, below 1 mm thick; fibers yellow to brown; no vessels. 

Significance: predominantly saprobic, G. sepiarium and G. trabeum exceptionally 

on living trees; belonging to the strongest brown-rot fungi of coniferous 

structural timber; often on stumps; broad moisture optimum (about 40 to 

200% u; Table 8.7), on stored timber and on finished timber that is again 

moistened, like poles, posts, fences, sleepers and mining timber. The fungi are 

the most important destroyers of conifers windows (cf. Fig. 8.17) that had accumulated 

moisture due to inappropriate window construction and handling 

faults by the user (e.g., injuring of the lacquer layer by nails). For example, 

3.5 million (7%) of wooden windows were partly or completely destroyed by 

fungi, predominantly by G. abietinum, in Germany between 1955 and 1965 

(Seifert 1974). Fungi survive in the sun-warmed and dry window timber due 

to their heat and dryness resistance [G. abietinum: 5–7yearssurvivalindry 

timber: Theden (1972)]. Fungi cause (by means of substrate mycelium) decay 

first only in the wood interior (“interior rot”). The serious brown rot under 

the varnish layer is often only recognized if fruit bodies develop. Except on 

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window timber, the gill polypores occur in buildings after moisture damages 

or incorrect structure on roofing timbers, on façades, outside doors, balconies, 

andontimberinsaunasandmines. 

8.4.3 

Lentinus lepideus, Scaly Lentinus 

Occurrence: temperate zones, common in Europe, North America, former 

Soviet Union, India; conifers, particularly Pinus, alsoAbies, Cedrus, Larix, 

Picea, Pseudotsuga, Tsuga; 

Fruit body (Fig. 8.16i): mainly eccentric, stipe (up to 7 cm long), pileus 5– 

15 cm wide; fleshy-tough to hard in age, initially convex, later applanate; upper 

surface: pale to cream or purplish brown, with brownish scales (name!) in radial 

orientation; lower surface: whitish to yellow-ocher, serrate gills; monstrose, 

sterile fruit bodies in the dark (Seehann and Liese 1981); dimitic (Kreisel 

1969); 

Significance: brown rot of heartwood, via wounds and dead branches in 

standing trees, on stumps, felled logs, serious damage on structural timbers 

outdoors in ground contact (poles, sleepers, fence posts, stakes, wooden 

bridges, harbor timbers) (Bavendamm 1952b), on mine timber; particularly 

dangerous due to resistance to heat, desiccation and coal tar oil (test fungus 

in EN 113 for tar oil and comparable compounds); degradation of pine heartwood 

(interior rot) in improperly impregnated (drying shakes developed after 

treatment) poles and sleepers; rarely in buildings, particularly in the cellar 

and on damp timber on the ground floor, on joist heads in contact with wet 

masonry, door posts, roof timber; pleasant smell of the fresh mycelium of Peru 

balsam. 

8.4.4 

Paxillus panuoides, Stalkless Paxillus 

Occurrence: mostly conifers; 

Fruit body (Fig. 8.16j): annual, thin, small (2–12 cm), shell-shaped, bellshaped, 

small eccentric stipe or attached, solitary or in groups, also tile-like; 

upper surface: pale-yellow to olive brown; lower surface: saffron-orange gills; 

monomitic; normal fructification in the dark (Kreisel 1961); 

Significance: slowly growing, but serious brown rot; rarely at the basis of 

living pines, on stumps, stored wood, structural timber outdoors (sleepers, 

bridges, balconies), garden furniture, mine timber, rarely in buildings, associated 

with the Coniophora spp., on very moist places in cellars, cow-sheds, 

greenhouses. 

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8.4.5 

Schizophyllum commune, (Common) Split-Gill 

Occurrence: circumglobal, temperate to tropical, very common, predominantly 

hardwoods like Fagus, Quercus, Tilia, fruit woods, bamboos, straw, tea-leaves, 

coconut fibers; 

Fruit body (Fig. 8.16k, l): annual, but durable, thin, small, shell-shaped 

(1–5 cm), dimidiate; usually in groups, leathery-tough; upper surface: greybrown 

to flesh-colored becoming white with dryness, downy-woolly; lower 

surface: appearing as if gilled, hymenium covering fan-like arranged, at the 

beginning grey, later violet-brown pseudolamellae, which are lengthwise split 

and outwardly bent (Fig. 3.3d); hygroscopic movements of the split lamellae 

by being hard and rolled up in dry weather and being again flexible and 

sporulating after years of dryness when again moist; monomitic, tetrapolar 

(Raper and Miles 1958); formerly eaten in Assam, Congo, Peru and Thailand, 

and used as chewing gum in Hong Kong, Indonesia and Malaysia (Dirol and 

Fougerousse 1981); fructification also in culture; 

Significance: white rot; as wound parasite on living trees after bark fire 

damage, on stumps, stored stems, frequently on beech as first colonizer; on 

stored and structural timber outdoors surviving dryness and exposition to 

sun by dryness resistance; in the tropics serious wood destroyer, fruit bodies 

often on imported timber; in vitro only little wood decay (Schmidt and Liese 

1980). 

8.4.6 

Trametes versicolor, Many-Zoned Polypore 

Occurrence: circumglobal, very common throughout Europe, dead wood of 

almost all hardwoods, particularly Fagus, alsoBetula, no attack of Quercus, 

Castanea, and Robinia (Jacquiot 1981), rarely conifers, also fruit woods after 

pruning; 

Fruit body (Fig. 8.16m): annual, often reviviscent, hard-leathery, sessile 

or effused-reflexed, pilei dimidiate-substipitate, convex or imbricate, rarely 

resupinate, to 10 cm wide, often in large imbricate clusters, rarely solitary; 

upper surface: hirsute to tomentose, highly variable in color, with sharply 

contracted concentric zones of brown, buff, reddish or bluish colors (name!), 

often green by algae; lower surface: cream-white to ochraceus-yellow, angular 

to circular pores (4–5/mm); in the dark self-colored fruit bodies with totally 

white hirsute upper surface; trimitic; tetrapolar; 

Significance: white-rot, often with black demarcation lines (“marble rot”); 

on wounded or dead standing trees, on stored stems, common on 4–6 years 

old hardwood stumps; rarely on sleepers, fence posts, garden timber; on mine 

timber; dryness resistance; used after World War II in the former East Germany 

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8.5 Damage to Structural Timber Indoors 207 

for the production of “myco-wood” for pencils, rulers, etc. (Luthardt 1963); 

test fungus in EN 113 for hardwood samples. 

8.5 

Damage to Structural Timber Indoors 

8.5.1 

General and Identification 

The indoor wood decay fungi (“house-rot fungi”) cause considerable economical 

damage in buildings. They may be considered to be the most important 

“wood fungi” as they deteriorate wood at the end of the economical series 

“forestry” – “timber harvest” – “storage” – “structural timber” – “indoor use”. 

For Britain, it has been estimated that the cost of repairing fungal damage of 

timber in construction in 1977 amounted to £ 3 million per week (Rayner and 

Boddy 1988). An estimate for the former East Germany amounts to an avoidable 

damage in old houses of e1.5 billion (Huckfeldt 2003). In the northern 

hemisphere, mainly coniferous wood is used as interior structural timber, in 

Germany particularly Picea abies. The most important wood-degrading fungi 

within buildings in Europe and North America are therefore fungi that cause 

brown rot in conifers. White-rot fungi, which preferentially attack hardwoods, 

are less common in buildings. Depending on the state of knowledge, formerly 

often only three more well-known species (groups) were called house-rot 

fungiinEurope:theTruedryrotfungus,Serpula lacrymans, the cellar fungi 

Coniophora spp. (formerly only C. puteana) and the indoor polypores, formerly 

called “Poria group” (probably mainly Antrodia vaillantii). These three 

groups cause about 80% of the fungal wood damages in buildings. Recently, 

the Oak polypore, Donkioporia expansa, has also been accepted as important 

indoor rot fungus (Kleist and Seehann 1999). The Gill polypores (Falck 

1909) may be included to the indoor species as they are common destroyer of 

painted coniferous window timber (Fig. 8.17) and also occur on damp roofing 

timber. 

There are some evaluations on the frequencies of the various species involved 

in indoor wood decay. A survey of 1,500 buildings in New York State 

from 1947 to 1951 showed several fungi and Hyphodontia spathulata, G. sepiarium, 

A. xantha, andG. trabeum as most frequent isolations from decayed 

wood (Silverborg 1953). An investigation of 3,050 buildings in Poland showed 

53.8% S. lacrymans, 22.4% C. puteana and 11.3% A. vaillantii (Wa˙zny and 

Czajnik 1963). A survey of 1,200 biotic damages in buildings of the former 

East Germany over 21 years resulted in 34.8% S. lacrymans, 14.6% Coniophora 

spp., 13% soft rot and 8.7% “Poria” (Schultze-Dewitz 1985). An evaluation of 

749 damages in Belgium between 1985 and 1991 revealed 59.4% S. lacrymans, 

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Fig.8.17. Gloeophyllum sp. on window joinery. Fruit body and brown-rotten softwood 

10.1% C. puteana, C. marmorata, 9.5%Donkioporia expansa, 2.3%Antrodia 

vaillantii, A. sinuosa, A. xantha and some further species (Guillitte 1992). 

An evaluation of a total number of 3,434 decay fungi in Norwegian buildings 

from 2001 to 2003 found as the most frequent fungi 18.4% Antrodia 

species, 16.3% C. puteana, 16.0% S. lacrymans and 2.9% G. sepiarium (Alfredsen 

et al. 2005). A recent survey over 4 years in 63 buildings in North 

Germany yielded 36 basidiomycetous species (Table 8.6). Supplemented by 

literature research, altogether about 70 different house-rot fungi have been 

reported (Huckfeldt and Schmidt 2005). However, those literature compilations 

might be uncertain due to the use of synonyms and the change in fungal 

nomenclature. 

A survey of 5,000 cases of damage in multistorey houses revealed that all 

timbers without sufficient basic protection are endangered, but that there are 

different damage centers in a home: “Poria” and soft rot in the attic and upper 

floor, and S. lacrymans and Coniophora spp. on the ground and in the cellar 

(Schultze-Dewitz 1990). 

Some of the less common indoor Basidiomycetes are listed in Table 8.6. 

Among them, Lentinus lepideus is particularly found in damp cellars, on the 

ground floor and in beam-ends in contact with wet masonry (Bavendamm 

1952b). Paxillus panuoides occurs in cellars (Bavendamm 1953). Daedalea 

quercina affects structural oak-wood (windows, half-timbering). Falck (1927) 

mentioned for cellars Polyporus squamosus and Coggins (1980) also Laetiporus 

sulphureus, Phlebiopsis gigantea and Trametes versicolor. A description 

of the Dry rot fungus and other fungi in houses and on timber in exterior use 

has been compiled by Bech-Andersen (1995). Some of the more rare indoor 

species normally occur on trees or timber in outdoor use and are described 

in Chaps. 8.3 and 8.4. Further indoor damages are discolorations of window 

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Table 8.6. Species and frequency of house-rot fungi and accompanying fungi in buildings 

in northern Germany (from Huckfeldt and Schmidt 2005) 

Species Frequency 

Serpula lacrymans 53 

Coniophora puteana 7 

Antrodia sp. 6 

Antrodia xantha 5 

Coprinus spp., three species 5 

Donkioporia expansa 5 

Asterostroma cervicolor 4 

Antrodia sinuosa 3 

Antrodia vaillantii 2 

Coniophora marmorata 2 

Dacrymyces stillatus 2 

Diplomitoporus lindbladii a 2 

Gloeophyllum trabeum 2 

Lentinus lepideus 2 

Leucogyrophana pinastri 2 

Leucogyrophana pulverulenta 2 

Paxillus panuoides 2 

Trechispora farinacea 2 

Asterostroma laxum a 1 

Cerocorticium confluens a 1 

Cerinomyces pallidus a,b 1 

Gloeophyllum abietinum 1 

Gloeophyllum sepiarium 1 

Gloeophyllum sp. 1 

Grifola frondosa a 1 

Heterobasidion annosum 1 

Hyphoderma praetermissum 1 

Leucogyrophana mollusca 1 

Oligoporus placenta 1 

Oligoporus sp. 1 

Phellinus contiguus 1 

Phellinus pini 1 

Pluteus cervinus a 1 

Stereum rugosum 1 

Trametes multicolor 1 

Trichaptum abietinum 1 

Volvariella bombycina 1 

non-decay fungi: 

Peziza repanda 5 

Reticularia lycoperdon 3 

Cladosporium sp. 2 

Fuligo septica 1 

Ramariopsis kunzei 1 

Scutellinia scutellata a 1 

a For the first time proven to occur in houses 

b First proof in Germany (Huckfeldt and Hechler 2005) 

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timber and outside doors by blue-stain fungi and molding in damp rooms 

(Chap. 6) (Frössel 2003; Hankammer and Lorenz 2003). 

The common house-rot fungi are serious wood decayers. Among them, S. 

lacrymans is considered in Europe as most dangerous and most hardly controllable 

fungus due to its ability to transport nutrients and water. Traditionally, it 

is also supposed to possess some further specific features, which, however, do 

not all stand up to laboratory results. Nevertheless, in Germany, S. lacrymans 

has to be clearly differentiated from the other house-rot fungi in view of refurbishment.Morefar-reachingmeasureshavetobeperformedinthecaseofits 

presence. Thus species identity should be known. 

For identification, fruit bodies are preferentially used (Grosser 1985; Breitenbach 

and Kränzlin 1986; Jahn 1990; Ryvarden and Gilbertson 1993, 1994; 

Krieglsteiner 2000; Weiß et al. 2000; Kempe 2003; Bravery et al. 2003). A diagnostic 

key for fungi on structural timbers based on their fruit bodies is 

available in the internet and is to be completed in time (Huckfeldt 2002). 

Some species only rarely fructify in buildings, or after isolation in laboratory 

culture, or do it never. However, some house-rot fungi form mycelial strands 

(cords). The classical strand diagnosis from Falck (1912) is old and includes 

only a few species. A diagnostic key including color photographs based on 

measurements in infected buildings and on wood samples in laboratory culture 

comprises several species (Huckfeldt and Schmidt 2004, 2006). An updated 

version is shown in Appendix 1. A recent textbook comprises photographs and 

identification keys for fruit bodies and strands of fungi occurring on wood in 

indoor and exterior use (Huckfeldt and Schmidt 2005). 

If neither fruit bodies nor strands, but only vegetative mycelia are present, 

e.g., if only mycelium is found in buildings, or as it is the case for fungi cultured 

in the laboratory on agar, there are keys and books for mycelia (Nobles 

1965; Stalpers 1978; Lombard and Chamuris 1990). However, some genera 

among the house-rot fungi are hardly or not at all distinguishable into species, 

like Antrodia, Coniophora and Leucogyrophana. Thus, molecular methods 

may be used (Chap. 2.4.2). Among the DNA-based techniques, species-specific 

ITS-PCR differentiated seven indoor wood-decay Basidiomycetes (Fig. 2.23, 

Table 2.9; Moreth and Schmidt 2000). The technique is meanwhile used in 

Germany for commercial identification of house-rot fungi. Sequencing of the 

internal transcribed spacer (ITS) of the ribosomal DNA (rDNA) and subsequent 

sequence comparison by BLAST with ITS sequences from correctly 

identified fungi deposited in the nucleotide databases is to date the best 

molecular tool for diagnosis (Table 2.8 and Fig. 2.22; Schmidt and Moreth 

2002, 2003). 

There is a great number of investigations on the physiology of house-rot 

fungi in text books (e.g., Jennings and Bravery 1991), monographs (e.g., Cockcroft 

1981), and publications that may used in support of identification. Among 

the physiological parameters, growth rate and reaction to wood moisture con- 

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Table 8.7. Cardinal points of wood moisture content (% u) of some house-rot fungi for 

colonization and decay of wood (after Huckfeldt and Schmidt 2005) 

Species Minimum for Minimum Optimum Maximum 

colonization for decay for decay for decay 

(moisture source (mass loss (mass loss (mass loss 

20–30 cm away) above 2%) above 10%) above 2%) 

Serpula lacrymans 21 26 45–140 240 

Leucogyrophana pinastri 30 37 44–151 184 

Coniophora puteana 18 22 36–210 262 

Antrodia vaillantii 22 29 52–150 209 

Donkioporia expansa 21 26 34–126 256 

Gloeophyllum abietinum 20 22 40–208 256 

Gloeophyllum sepiarium 28 30 46–207 225 

Gloeophyllum trabeum 25 31 46–179 191 

tent and temperature are important features. However, some of the older data 

suffer in so far as they derive from only vague or incorrectly identified fungi. 

Data that are based on genetically verified fungi are shown in Tables 2.2, 3.8– 

3.11, and 8.7. 

Regarding the most important influence on wood decay, wood moisture, 

opinion has it that the indoor polypores need moisture above the fiber saturation 

range, which often occurs only after wetting with water, whereas the 

Coniophora spp. mostly attack wood, which was moisturized by vaporous water 

or by contact with damp material. The Dry rot fungus is halfway as it 

germinates on contact-wetted timber, but takes water from wet substrates by 

capillary mechanism and translocates water in its mycelium to timber for 

further growth (Schultze-Dewitz 1985). 

In piled Scots pine sapwood samples placed on agar in 2-L Erlenmeyer 

flasks, a continuous wood moisture gradient developed within 6 weeks by diffusion 

from the agar via the lowest sample, which was water-saturated to the 

uppermost air-dried sample (Huckfeldt 2003). Table 8.7 shows that all fungi 

subsequently inoculated on the agar near the bottom wood sample degraded 

very wet wood. For example, S. lacrymans showed more than 2% wood mass 

loss in a sample of 240% final moisture content. The optimum moisture for 

decay (mass loss above 10%) varied among the species from 36 to 210% u. The 

minimum moisture for decay (mass loss above 2%) was slightly below fiber 

saturation and for C. puteana and G. abietinum significantly low at 22% u. 

Minimum moisture for wood colonization was for some fungi around 20% u, 

whereby the wood sample was 20–30 cm away from the agar as the water source 

(Huckfeldt and Schmidt 2005). 

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8.5.2 

Lesser Common Basidiomycetes in Buildings 

The following species description starts with some lesser common fungi and 

ends with the most serious European fungus, the True dry rot fungus Serpula 

lacrymans,inorderofatransitiontotheremedialtreatments.Daedalea 

quercina, Gloeophyllum species, Lentinus lepideus and Paxillus panuoides, 

which also occur in buildings, have been already described in Chap. 8.4. The following 

data are based on observations and measurements in attacked buildings 

and on genetically verified pure cultures on wood samples in the laboratory 

(Huckfeldt 2003; Huckfeldt and Schmidt 2005; Huckfeldt et al. 2005; Schmidt 

and Huckfeldt 2005), and were supplemented mainly from Grosser (1985), 

Breitenbach and Kränzlin (1986), Ryvarden and Gilbertson (1993, 1994), and 

Bravery et al. (2003). 

8.5.2.1 

Diplomitoporus lindbladii 

Occurrence: circumpolar in the conifers zone, in Europe throughout the conifer 

forest regions, but rare in the Mediterranean region, North America, also on 

hardwoods; 

Fruit body (Fig. 8.18a): annual to biannual, resupinate, becoming widely 

effused (a few decimeters), up to 6 mm thick, biannual basidiomes thicker, 

frayed margin, easily separable; upper surface white-cream, grey when old; 

pore surface with 2–4 circular-angular pores/mm, to 3 mm deep; trimitic; 

allantoid to cylindrical, hyaline spores (5–7 × 1.5–2µm); bipolar; 

Strands (Fig. 8.18b): on timber in laboratory culture, white, yellowing when 

dry, root-like, iceflower-like, similar to A. vaillantii; fiberssimilartoA. vaillantii,butsolublein5%KOH; 

Significance: white rot, indoors. 

8.5.2.2 

Asterostroma cervicolor and A. laxum 

Fruit body (Fig. 8.18c): resupinate, sheet-like, thin, whitish to ochre or cinnamon, 

hardly distinguishable from mycelium; no pores; may be found on 

masonry; spores warty (A. cervicolor), without warts (A. laxum); monomitic; 

Strands and mycelium (Fig. 8.18d): cream-brown, up to 1-mm-wide strands 

with a rough appearance, flexible when dry, sometimes across and inside masonry 

over a long distance, brown strands often present next to fruit body, 

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Fig.8.18. Diplomitoporus lindbladii a Fruit body and detail. b Mycelium and strands on 

white-rotten wood; Asterostroma cervicolor c Fruit body on a ground ceiling. d Knotty 

mycelium and strands on a floorboard. e Stellar setae; Dacrymyces stillatus f Young fruit 

bodies. g Old fruit bodies on window joinery (photos T. Huckfeldt) — 5 cm, - - - 5 mm 

embedded in white mycelium or in fruit bodies (A. laxum); surface mycelium 

of A. cervicolor first white, then brown, partly only small mycelial plugs; 

Stellar setae (Fig. 8.18e): within basidiome, mycelium and strand (German: 

“Sternsetenpilz”); setae dichotomically branched, to 90 µm in diameter and 

partly rare in A. laxum, setae rarely branched and to 190µm indiameterin 

A. cervicolor; 

Significance: white-rot, softwoods, often on joinery, e.g., skirting boards, 

floor and ceiling boards, windows, fiber and gypsum boards, decay often 

limited in extent. 

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8.5.2.3 

Dacrymyces stillatus, Orange Jelly 

Fruit body (Fig. 8.18f, g): yellow-orange-red, also whitish, dark orange when 

dry, button-shaped, lenticular to mug- or plate-like, 1–15 mm wide, gelatinouselastic,slimy 

meltingwhenold,solitary andingroups,oftentwodifferentforms 

on the same place, a brighter form with basidiospores and a darker form with 

arthrospores, often appearing through paint; 

Significance: white rot, softwoods and hardwoods, wood darkens, decay 

commonly patchy with small pockets of rot, often restricted to interior of 

timber, on window and doorframes, common outdoors on windows, claddings 

and along the gable board of the roof (Alfredsen et al. 2005). 

8.5.3 

Common House-Rot Fungi 

There is a bulk of knowledge on the common indoor wood decay fungi due 

to their economic importance. Thus, these species and species groups are 

described in more detail in the following (also Findlay 1967; Bavendamm 

1969; Coggins 1980; Cockcroft 1981; Grosser 1985; Jennings and Bravery 1991; 

Ryvarden and Gilbertson 1993, 1994; Krieglsteiner 2000; Weiß et al. 2000; 

Kempe 2003; Sutter 2003; Huckfeldt and Schmidt 2005). 

8.5.3.1 

Donkioporia expansa, Oak Polypore 

This fungus is only recognized since the 1920s as relevant for practice and 

since about 1985 as important decay fungus in buildings (Kleist and Seehann 

1999; Erler 2005). Assumably, the species was often overlooked despite the less 

common decay type of a white rot in buildings and the large size of its fruit 

bodies. A reason it was overlooked may be that damage is often restricted to 

wood interior and not noticed until fruit bodies appear and furthermore that 

the fruit bodies are inconspicuously embedded in plentiful surface mycelium. 

Occurrence: fairly rare, Central Europe, North America, in Germany preferentially 

in the south, at least in Europe almost exclusively restricted to structural 

timber, preferably Quercus, but also Castanea, Fraxinus, Populus and 

Prunus,frequentlyalsoonindoortimberofPicea and Pinus; 

Fruit body (Fig. 8.19a, b): perennial, resupinate, first white, then ochre to 

reddish-tobacco-brown to grey with ageing, to 10 cm thick, becoming widely 

effused to a few square meters, firmly attached, an walls wavy to stairs-like, 

often multi-layered, tough-elastic with silvery surface when fresh, hard and 

brittle when dry, easily separable when old, mainly made up of long tubes, 4–5 

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Fig.8.19. Donkioporia expansa a Fruit body and mycelium. b Detail showing the long pores. 

c Old mycelium. d Strand-like structures grown on wood in laboratory culture; Antrodia 

vaillantii e Mycelium and strands. f Fruit body and detail. g Antrodia sinuosa fruit body 

and detail. h Antrodia xantha fruit body and detail. i Antrodia serialis fruit body and detail. 

j Oligoporus placenta fruit body and detail (photos b–j: T. Huckfeldt) — 5 cm, --- 5 mm 

circular to angular pores/mm, often amber guttation drops, which leave behind 

small black pits when dry; trimitic; ellipsoid spores 4.5−7 × 3.2−3.7µm; 

Mycelium (Fig. 8.19a,c): inside wood shakes and cavities, at high air humidity 

also on free wood surfaces with thin, skin-like mycelial flaps with bizarre 

seeds, later thick, brownish surface mycelium, guttation as on fruit bodies, 

black demarcation lines between mycelium and woody substrate; 

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Strands (Fig. 8.19d): not yet observed in buildings, strand-like structures 

on wood samples in laboratory culture, rare, cream, yellowish to grey-brown, 

root-like, hidden under mycelium; 

Significance: The Oak polypore inhabits damp areas in kitchens, bathrooms, 

WC, cellars, cow-sheds, occurs on beams, under floors, in mines, on 

bridge timber, and cooling tower wood [Azobé, Bangkirai; v. Acker et al. 

(1995); v. Acker and Stevens (1996)]. It produces a white-rot. Continuous high 

wood moisture promotes growth (defective sanitary facilities, cooling tower 

wood). The fungus is often found at beam-ends that are enclosed in damp 

walls. At initial attack of softwoods, the timber surface remains often nearly 

intact (“interior rot”). In laboratory culture, minimum wood moisture for 

wood colonization was 21% u and for wood decay 26%. Greatest wood mass 

losses occurred between 34 and 126% (Table 8.7). Moisture maximum was 

256%. Temperature optimum was 28 ◦ C, and maximum was 34 ◦ C(Table3.8). 

The fungus survived for 4 h in dry wood of 95 ◦ C (Huckfeldt 2003). Wood 

mass losses according to EN 113 were: oak sapwood 45%, oak heartwood 

23%, beech 50%, birch 60%, pine sapwood 40% (Kleist and Seehann 1999). 

Assumably,thereisnospreadbystrandsfrommoisttodrywoodandno 

growth through the masonry because strands were only found in vitro to date. 

Thus, refurbishment only needs drying and exchange of destroyed timber. In 

oaks, the fungus is often associated with the death-watch beetle, Xestobium 

rufovillosum. 

8.5.3.2 

Indoor Polypores: Antrodia Species and Oligoporus placenta 

Four Antrodia species and O. placenta may be assigned to the indoor polypore 

fungi. 

Occurrence: circumglobal in the coniferous forest zone, mostly on softwoods 

(Findlay 1967; Domański 1972; Coggins 1980; Lombard and Chamuris 

1990; Grosser 1985; Lombard 1990; Ryvarden and Gilbertson 1993, 1994; 

Krieglsteiner 2000; Sutter 2003); 

Antrodia vaillantii occurs circumglobal in the coniferous forest zone and in 

Europe widely distributed, but rather rare in Fennoscandia. It is the most frequent 

fungus in British mines (Coggins 1980). Antrodia sinuosa is circumpolar 

in the boreal conifer zone, widespread in Europe, North America, East Asia, 

North Africa, and Australia (Domański 1972). The species was in Sweden with 

1,045 damages between 1978 and 1988 with 13% portion the most common indoor 

polypore (Viitanen and Ritschkoff 1991a). Antrodia serialis attacks logs 

and piles, causes heart rot in standing trees and occurs widespread, also in 

Himalaya and Africa (Seehann 1984; Breitenbach and Kränzlin 1986), rarely 

(1.4%) in buildings (Viitanen and Ritschkoff 1991a; Coggins 1980), within 

the roof area, in cellars and under corridors (Domański 1972). Antrodia xan- 

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tha (Domański 1972) occurs in Europe and North America on fallen stems, 

branches, stumps, in greenhouses (Findlay 1967), at windows (Thörnqvist et al. 

1987), on timber in swimming pools and in flat roofs (Coggins 1980). Oligoporus 

placenta is rare, but widespread in Europe except for the Mediterranean. 

In North America, the fungus is the most common wood decayer in ships 

(Findlay 1967) and was exported to Great Britain (Coggins 1980). In North 

America, O. placenta and A. serialis are common on mine timber and poles 

(Gilbertson and Ryvarden 1986). 

Antrodia vaillantii, Mine polypore, Broad-spored white polypore 

Fruit body (Fig. 8.19f): annual, resupinate, first white, then light yellow to grey, 

drying, as corky layer (to 1 cm thick) on the wood underside or above as pad; 

2–4 circular-angular pores/mm hymenium, to 12 mm long; dimitic; hyaline 

spores 5–7 × 3–4µm; 

Strands (Fig. 8.19e): pure white, felty, 0.5–7 mm in diameter, ice flower-like, 

flexiblealsoifdry;fibersnumerous,white,flexible,2–4µm thick, unsoluble 

in 5% KOH; vessels not rare, to 25µm in diameter, partly with thick walls and 

reduced lumen, no wall thickenings; vegetative hyphae with clamps, 2–6µm 

in diameter, often also thick-walled. 

Antrodia sinuosa, White polypore, Small-spored white polypore 

Fruit body (Fig. 8.19g): similar to A. vaillantii, annual,resupinate,to5mm 

thick; 1–3 circular-sinuous pores/mm, to 3 mm long; dimitic; hyaline spores 

4−6×1−2µm; 

Strands: similar to A. vaillantii. 

Antrodia xantha, Yellow polypore 

Fruit body (Fig. 8.19h): annual, resupinate, first yellowish, then pale, whitecream, 

crusty to bracket-shaped, to 10 mm thick, 1 m wide; 3–7 circularangular 

pores/mm, to 5 mm long; margin without pores; on vertical substrates 

small knots, to 8 mm large, partly grown together; dimitic; hyaline spores 

4−5×1−1.5µm; 

Strands: similar to A. vaillantii, but partly yellow discolored, later often pale 

and then undistinguishable from A. vaillantii. 

Antrodia serialis, Effused tramete, Row polypore 

Fruit body (Fig. 8.19i): annual to biennial, resupinate to pileate, first white to 

cream-ochre, then pink-spotted, to 6 mm thick, to a few decimeters wide; 2–4 

circular, partly slitted pores/mm, to 5 mm long; distinct, wavy margin; also in 

rows; dimitic; hyaline spores 4−7 × 3−5µm; 

Strands: not yet found. 

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Oligoporus placenta, (Reddish) Sap polypore 

Fruit body (Fig. 8.19j): annual, resupinate, either white to grey-brown (form 

monticola) or later pink to salmon-violet (reddish form placenta)(Domański 

1972), easily passing, to 1 cm thick; 2–4 circular-angular-slitted pores/mm, to 

15 mm long; monomitic; hyaline spores 4−6 × 2−2.5µm; 

Strands: on wood samples in laboratory culture, white, partly yellowing, 

easily refractable, to 1 mm in diameter; fibers and vessels rare or absent. 

Significance: The five “indoor polypores” form a group of brown-rot fungi that 

are associated with the decay of softwoods in buildings. In Central Europe, 

these fungi belong after the Dry rot fungus, Serpula lacrymans, andtogether 

with the Coniophora cellar fungi to the most common indoor decay fungi. They 

accounted for 14% of indoor decay fungi in Denmark (Koch 1985) and Finland 

(Viitanen and Ritschkoff 1991a). A survey in California ranked A. vaillantii, 

A. sinuosa, A. xantha and O. placenta with29%occurrenceasthemaingroup 

(Wilcox and Dietz 1997). 

The polypores have similar biology and distribution (Lombard and Gilbertson 

1965; Donk 1974; Breitenbach and Kränzlin 1986; Lombard and Chamuris 

1990; Bech-Andersen 1995; Schmidt and Moreth 1996, 2003). They differ in 

their fruit body, spore morphology (Jülich 1984; Ryvarden and Gilbertson 

1993, 1994) and sexuality. Some species also fruit in laboratory culture, which 

supports identification of mycelia and tests for sexuality. Antrodia vaillantii 

is tetrapolar heterothallic (Lombard 1990), A. serialis, A. sinuosa and O. 

placenta are bipolar (Domański 1972; Stalpers 1978). Three Antrodia species 

develop strands (Falck 1912; Stalpers 1978; Jülich 1984), O. placenta only in 

vitro. However, the vegetative mycelium that has been isolated from decayed 

wood is hardly distinguishable (Nobles 1965). Due to the limited differentiating 

features, misinterpretations occur. 

Furthermore, the nomenclature has a confusing history and is still not always 

uniform (Cockcroft 1981). Fungi have been variously classified as Polyporus, 

Poria, Amyloporia, Fibroporia (Domański 1972). Misleading synonyms in the 

older literature such as Polyporus vaporarius and Poria vaporaria have been 

used for different species, viz. A. vaillantii (Bavendamm 1952c), A. sinuosa, 

and O. placenta. According to Ryvarden and Gilbertson (1994), the Reddish 

sap polypore, formerly Tyromyces placenta (Fr.) Ryv., was placed in Oligoporus, 

since the genus Tyromyces is restricted to fungi causing a white rot. Older 

synonyms are Postia placenta (Fr.) M.J. Larsen & Lomb., Poria placenta (Fr.) 

Cooke sensu J. Eriksson, Poria monticola Murr., and the haploid standard 

strain Poria vaporaria (Pers.) Fr. sensu J. Liese (Domański 1972). Postia is 

a nomen provisorium/nudum in the sense of Fries and illegitimate in the sense 

of Karsten. Isolate MAD 698 of Postia placenta was thoroughly investigated in 

view of brown-rot decay mechanisms (e.g., Clausen et al. 1993; Highley and 

Dashek 1998). Difficulties may increase because O. placenta separates into the 

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forms placenta with salmon-pink fruit bodies (“Reddish sap polypore”) and 

monticola, never with reddish stain (Domański 1972). Monokaryotic isolates 

of O. placenta were used for testing wood preservatives in Germany (Poria 

vaporaria “standard strain II”) and are obligatory in the recent European 

standard EN 113 (see Table 3.9, 3.10, named “Poria placenta” FPRL 280). Even 

literature from 2005 uses the names Postia placenta and Poria placenta. 

For species identification in the case that only vegetative mycelium is present, 

rDNA-ITS sequencing separates the five species (Schmidt and Moreth 2003; 

Chap. 2.4.2.2). 

For an easier understanding during a practical valuation of a fungal damage, 

the different fungi are often summarized as “indoor polypores” or as “Vaillantii 

group”, particularly because they differ from the Cellar fungus and Dry rot 

fungus by their mycelia, strands, and fruit bodies. The polypores, particularly 

A. vaillantii, form a well-developed white and cottony surface mycelium without 

“inhibition colors”, which, thus, can be confused with the young mycelium 

of the Dry rot fungus. Polypore mycelium spreads ice flower-like over the substrate, 

that of the Dry rot fungus is converted with ageing into silvery-grey 

skins, and that of the cellar fungi is dominated by fine black strands. White 

(A. vaillantii), to string-thick, smooth and flexible strands develop within 

the mycelium and grow over non-woody substrates and also through porous 

masonry (Grosser 1985), the latter, however, less intensive than by the Dry 

rot fungus. The white to yellow (A. xantha) orred(O. placenta f. placenta) 

fruit bodies show pores that are visible with the naked eye (Fig. 8.19). The 

dry wood shows the typical brown-cubical rot. It is often said that the cubes 

caused by the polypores and the cellar fungi are smaller than those by the 

Dry rot fungus. The cube size varies however also as a function of the wood 

moisture content (Grosser et al. 2003). After advanced decay, the dried substrate 

of most brown-rot fungi can be ground with the fingers to a brown 

powder (“lignin”). 

The polypores attack predominantly coniferous woods in damp new and 

old buildings, particularly in the upper floor, furthermore mine timber, stored 

timber as well as timber in outside use, particularly in the soil/air zone, such 

as poles and sleepers. They also attack trees as wound parasites and live 

on stumps and fallen trees (Krieglsteiner 2000). Antrodia serialis was found 

in over-mature Sitka spruce trees (Seehann 1984). “Dry” wood should not 

become infected. In the laboratory, however, wood of 22% moisture content 

was colonized (Table 8.7). As so-called “wet-rot fungi” (Coggins 1980; Bravery 

et al. 2003), they need wet wood with moisture contents from 30 to 90% u for 

a long time. According to literature, the optimum is around 45% (Table 3.6). 

Laboratory experiments revealed that minimum moisture for wood decay by 

A. vaillantii was 29% and the optimum 52 to 150% (Table 8.7). With timber 

drying, Antrodia species were supposed to die (Bavendamm 1952c; Coggins 

1980). However, more convincing seems that they only stop growth (Grosser 

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1985). In the laboratory, over 11 years were survived by “dryness resistance” 

(Theden 1972), so that fungi may come to life again. There is also resistance to 

high temperature: Antrodia vaillantii, A. sinuosa and O. placenta survived on 

agar 3 h at 65 ◦ C. Antrodia vaillantii and O. placenta withstood heat of 80 ◦ C for 

4 h in slowly dried wood samples (Huckfeldt 2003), which has to be considered 

in view of a possible treatment of infected homes with hot air. 

Some species destroy timber in soil contact, like poles and palisades, even 

if it is properly impregnated with chrome-copper salts (Stephan et al. 1996). 

Especially A. vaillantii but also A. xantha and O. placenta are known for 

copper tolerance (Da Costa and Kerruish 1964) due to the production of oxalic 

acid (Rabanus 1939; Da Costa 1959; Sutter et al. 1983, 1984; Jordan et al. 

1996). Strain variation occurred (Da Costa and Kerruish 1964; Collett 1992a, 

1992b), and monokaryons were more tolerant than their parental strains (Da 

Costa and Kerruish 1965). In vitro, A. vaillantii was the most copper-tolerant 

fungus among the five species (Table 3.10) and produced most oxalic acid 

(Table 3.9; Schmidt and Moreth 2003). Antrodia vaillantii is also tolerant to 

arsenic (Göttsche and Borck 1990; Stephan and Peek 1992). 

8.5.3.3 

Cellar fungi: Coniophora species 

Occurrence: The genus Coniophora comprises about 20 species occurring 

worldwide with a broad host range primarily on conifers (Ginns 1982). Seven 

species occur in Europe (Jülich 1984) and five in Western Germany (Krieglsteiner 

1991). Coniophora puteana is frequently associated with brown-rot 

decay in European buildings. The fungus was estimated to be twice as common 

as the Dry rot fungus in the UK (Eaton and Hale 1993). It comprised over 

50% of the inquiries at the Danish Technological Institute (Koch 1985), 16.3% 

in Norway (Alfredsen et al. 2005), and 13% at the Finnish Forest Products 

Laboratory (Viitanen and Ritschkoff 1991a). The fungus has been used for 

nearly 70 years as a test fungus for wood preservatives in Europe. It also occurs 

in the USA, Canada, South America, Africa, India, Japan, Australia, and New 

Zealand. Further “cellar fungi” that attack indoor timber in Europe are especially 

C. marmorata,andalsoC. arida and Colivacea(Fig. 8.20). In Europe, the 

cellar fungi cause with about 10% frequency the two to third most common 

fungal indoor wood decay after S. lacrymans. In Australia and New Zealand, C. 

arida and C. olivacea are common. Some further Coniophora species also occur 

in buildings, mines and glass houses, but predominantly in warm climatic 

zones (Ginns 1982). The species can be differentiated by their fruit bodies 

(Jülich and Stalpers 1980; Breitenbach and Kränzlin 1986; Krieglsteiner 2000). 

However, the species concept within Coniophora is difficult because there are 

only a few, and unstable characteristics, which complicates species identification 

in infected buildings. With regard to isolates in culture, Coniophora cannot 

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Fig.8.20. Cellar fungi. Coniophora puteana a Fruit body. b Fruit body margin. c Fruit body 

detail with warts. d Strands in a false ceiling. e Strands on a steel girder. f Coniophora arida 

fruit body. g Coniophora olivacea fruit body (photos T. Huckfeldt) — 5 cm, --- 5 mm 

be differentiated at the species level by morphological and cultural characteristics 

(Stalpers 1978). Thus, isolations from buildings were summarized as C. 

puteana/ C. marmorata (Guillitte 1992). Sequencing of the rDNA-ITS separated 

the species (Schmidt et al. 2002b). Based on fruit-body identification, C. marmorata 

is rather common in southern Germany. The following description is 

based mainly on Huckfeldt (2003), Huckfeldt and Schmidt (2005) and Schmidt 

and Huckfeldt (2005). 

Coniophora puteana, (Brown) Cellar fungus 

Fruit body (Fig. 8.20a–c): annual, resupinate, light to dark brown, first whiteyellow, 

then brownish; indistinct, fibrous margin; to 4 mm thick, to a few 

decimeters wide, firmly attached, fragile when dry; warty knots up to 5 mm 

thick; monomitic; yellow-brown spores 9−16 × 6−9µm; 

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Strands (Fig. 8.20d, e): first white, soon brown-black, to 2 mm thick, rootlike, 

fragile, black wood beneath the strands; fibers brown, 2–5µm thick, 

lumina visible; vessels 10–30µm thick, often deformed, no bars; vegetative 

hyphae mostly clampless, rarely with multiple clamps, with brown drops (1– 

5µm) holding the hyphal net together. 

Coniophora marmorata, Marmoreus cellar fungus 

Fruit body: annual, resupinate, pale to olive-brown, grey margin, to 0.4 mm 

thick, to 15 cm wide, separable, felty; dimitic; no picture available because not 

yet found in buildings in northern Germany; 

Strands: brownish, to 1 mm thick, easily separable, no drops. 

Coniophora arida,Aridcellarfungus 

Fruit body (Fig. 8.20f): annual, resupinate, white-ochre to yellow-brown, light 

margin, to 0.3 mm thick, to 10 cm wide, firmly attached, smooth to felty, finefrayed 

margin; monomitic; 

Strands: rare, white to brown, 0.1 mm thick. 

Coniophora olivacea, Olive cellar fungus 

Fruit body (Fig. 8.20g): annual, resupinate, olive-brown, margin lighter, fraying 

with strands, to 0.6 mm thick, to 6 cm wide, firmly attached, smooth to warty, 

fibrous-cottony, septate cystidia, monomitic, partly merging fruit bodies; 

Strands: brown, thin. 

Significance: The older European literature on occurrence, biology and significance 

of the cellar fungi summarizes the several fungi to C. puteana. This 

fungus was said to be the most common species in new buildings. It however 

occurs also in damp old buildings, on stored wood, timber in soil contact like 

poles, piles, sleepers and on bridge timber as well as rarely on stumps and 

as wound or a weakness parasite on living trees (Bavendamm 1951a; Grosser 

1985; Breitenbach and Kränzlin 1986; Sutter 2003). Of 177 Basidiomycetes 

on American mine timbers, 83 isolates were C. puteana (Eslyn and Lombard 

1983). In buildings it does not occur, like the name misleadingly suggests, 

only in cellars, but it can ascend everywhere on damp timber up to the roof 

(Schultze-Dewitz 1985, 1990). Beside softwoods, it attacks also several hardwoods 

(Wälchli 1976). As a so-called wet rot fungus (Bravery et al. 2003) with 

relatively high requirement for moisture from 30 to about 70% u and the optimum 

around 50% (Table 3.6), all timber in the area of damp walls (beam 

ends and wall slats), damp floors and ceilings in kitchens, bathrooms and toilets 

as well as all timber in areas with water vapor development (swimming 

pools, launderettes) is endangered. In vitro, minimum moisture of C. puteana 

for wood colonization was 18% u and for decay 22%. The optimum moisture 

was broad, from 36 to 210% (Table 8.7). Damage by the cellar fungi is quite 

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comparable with that one of the Dry rot fungus and can even exceed it. A fresh 

floorboard can be completely destroyed in 1 year, so the danger exists that 

furniture or persons can fall through. These types of damages occurred in 

Germany frequently during the building boom in the postwar years, if insufficiently 

dried wood were used, or the homes had not sufficiently dried before 

they were moved into and drying was prevented by humidity-impermeable 

painting, linoleum, or carpet. 

The cellar fungi belong to the fast-growing house-rot fungi and reached on 

agar at 23 ◦ C up to 11 mm radial increase per day (Table 3.11). The optimum 

temperature (Table 3.8) was between 20 and 27.5 ◦ C, whereby C. marmorata 

preferred the warmer range, and the maximum was between 25 and about 

37.5 ◦ C. Isolate Ebw. 1 of C. puteana survived 15 min. at 60 ◦ C(Miričand 

Willeitner 1984) and 3 h at 55 ◦ C (Table 3.8). In slowly dried wood samples, 

even 4 h at about 70 ◦ C were withstood (Huckfeldt 2003). The data concerning 

a possible dryness resistance of the fungus vary: after observations from practice, 

it dies when drying; up to 7 years were however survived in dry wood in 

the laboratory (Theden 1972). There was isolate variation with regard to the 

sensitivity to wood preservatives (Gersonde 1958). 

Recognition characteristics (Fig. 8.20): The diagnosis is not always easy, 

since fruit bodies are rare and colonized wood shows frequently no or only meager 

surface mycelium (Käärik 1981). The few centimeters to several decimeters 

wide, resupinate, brownish fruit bodies resemble those of the Dry rot fungus, 

are however thinner. The species C. puteana is easy to recognize of the warty 

knots on the hymenophore (name: “carrying cones”). Characteristic on agar 

are double and multiple clamps. The initial stages of the rot are frequently 

ignored, since hardly infection signs become visible on exposed wood exterior 

surfaces, e.g., on baseboards, while the wood at the backside is already 

completely rotten and overgrown by thread-thin, radiate to root-like, brown 

to black strands (Fig. 8.20d,e). Early signs of rot are often dark discolorations 

under the paints. 

8.5.3.4 

Dry-rot fungi: Serpula species, Leucogyrophana species, Meruliporia incrassata 

This chapter deals with the brown-rot causing dry-rot fungi, namely Serpula 

lacrymans and S. himantioides,andtheLeucogyrophana species, L. mollusca, 

L. pinastri and L. pulverulenta (Fig. 8.21). Due to its economic relevance in 

Europe, emphasis is laid on S. lacrymans, however, the American pendant, the 

American dry rot fungus, Meruliporia incrassata,isconsidered. 

The way of spelling of the epithet “lacrimans”, which can be attributed to 

Fries (1821), is linguistically correct, however illegal, since the original spelling 

by Wulfen in 1781 was with “y” (Pegler 1991). 

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Occurrence and significance: The True dry rot fungus, S. lacrymans, isthe 

most dangerous house-rot fungus in central, eastern, and northern Europe, 

northwards to the Hebrides. It grows however also in cooler areas of Japan 

(Doi 1991), Korea, India, Pakistan and Siberia (Krieglsteiner 2000), in New 

Zealand and southern Australia (Thornton 1991), in Mexico, Canada and in the 

northern USA (Rayner and Boddy 1988). The data concerning its involvement 

in fungal indoor damage reach from 16% in Norway (Alfredsen et al. 2005) 

over 22% in Denmark (Koch 1991), 54% in Poland (Wa˙zny and Czajnik 1963) 

and North Germany (Schmidt and Huckfeldt 2005) to 59% in Sweden (Viitanen 

and Ritschkoff 1991a). For example, the annual repair costs of dry rot damage 

amount to at least 150 million £ in Great Britain (Jennings and Bravery 1991). 

Since the fundamental work by Hartig (1885), Mez (1908), Falck (1912: 

cf. Hüttermann 1991) and Wehmer (1915) S. lacrymans belongs to the bestinvestigated 

fungi. The older observations and results are described by Liese 

(1950), Bavendamm (1951b), Cartwright and Findlay (1958), Harmsen (1960), 

Savory (1964), Wagenführ and Steiger (1966), Findlay (1967), Bavendamm 

(1969), Coggins (1980) and Segmüller and Wälchli (1981). A literature search 

from 1988 lists 1200 publications (Seehann and Hegarty 1988). Informative 

photographsfordiagnosisonthebasisfruitbodies(Fig.8.21a,b)areby 

Grosser (1985) and on the Internet (www.hausschwamminfo.de). Younger reviews 

and laboratory findings to the biology and physiology are by Jennings 

and Bravery (1991), Viitanen and Ritschkoff (1991a), Schmidt and Moreth- 

Kebernik (1991a), Eaton and Hale (1993), Huckfeldt (2003), Schmidt (2003), 

Huckfeldt and Schmidt (2005), Huckfeldt et al. (2005), Schmidt and Huckfeldt 

2005). There is a German instruction leaflet with experiences from the practice 

on life conditions and refurbishment (Grosser et al. 2003). 

As cause of the special danger of the fungus the following features were 

specified: Its “omnipresent” spores germinate on damp wood or other cellulosic 

materials (paper, cardboard), and the mycelium can reach wood by 

growing over and through substrates that do not serve as a nutrient. For initial 

colonization, it only needs low wood moisture content. The conventional 

wisdom is that it is the only fungus that can infect so-called “dry” timber 

(min. 21% u) and masonry (min. 0.6% water content) and widely spread by 

mycelium (Fig. 8.21c) and its highly developed strands (Fig. 8.21d; name: “small 

serpent”), thereby growing over and through wood and several other 

materials, like porous or ruptured masonry or its wall joints, supplying channels 

for electricity, and water pipes (Coggins 1991; Jennings 1991). However, 

recent laboratory experiments showed that S. lacrymans is not unequalled as is 

also other indoor fungi colonized dry wood (Table 8.7). Coggins (1980, 1991) 

stressed that the initial colonization of a substrate, as for example the growth 

throughwalljoints,occursbytheyoungesthyphaeofthevegetativemycelium, 

in contrast to the infection way of Armillaria species that do this by means of 

rhizomorphs. In contrast, the strands develop as a secondary mycelium behind 

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Fig.8.21. Dry-rot fungi. Serpula lacrymans a Fruit body. b Detail. c Mycelium. d Strands. 

e Serpula himantioides fruit body; Leucogyrophana pinastri f Old fruit body. g Detail. h Old 

strands and sclerotia, iMyceliumand sclerotia.jYoungsclerotia.kLeucogyrophana mollusca 

fruit body. l Hair-like strands and sclerotia. m Mycelium and sclerotia; Leucogyrophana 

pulverulenta n Old fruit body, o Mycelium and strands (photos b–o:T.Huckfeldt)—5cm, 

---5mm 

the growth front and serve rather to transport nutrients to the hyphal margin. 

Alkaline materials to pH 10 can be overgrown, and alkalinity is decreased by 

excretion of liquid (pH 3–4) at the hyphal tip. An acute infection is often for 

a longer time not recognized due to the “hidden way of life”. Spores and still 

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alive mycelia can lead to re-infections in the case of careless or inappropriate 

remedial treatments (Bravery et al. 2003). Thick mats of surface mycelium may 

cover the attacked timber assumably preventing the wood from drying. 

Serpula lacrymans occurs predominantly in older buildings and in the cellar 

and ground floor area (Schultze-Dewitz 1985, 1990; Koch 1990). Uninhabited 

and poorly ventilated houses and all buildings with high relative air humidity in 

connection with damages to the structural fabric are particularly endangered. 

Importantcausesofdryrotinfectionsarebuildingdefectsthataffectincreased 

wood moisture content (e.g., Paajanen and Viitanen 1989). The mycelium reacts 

sensitively to draught and humidity removal, generally to climatic changes, so 

that it often develops in false ceilings and false soil areas under floors and 

behind wall coverings, from where it spreads. Because of this hidden way of 

life, often only fruit bodies on masonry, baseboards, doorframes or stairway 

steps show that the higher floors are already infected. In extreme cases, e.g., 

during the refurbishment of listed buildings, all timbers as well as large parts 

of the masonry have to be removed. A survey of houses in northern Germany 

indicated that old buildings are particularly at risk, which had insulating 

windows as the only measure of heat insulation. Now, the moisture in the 

building condenses on other weak spots like empty spaces of the brickwork at 

the back of heaters (Huckfeldt et al. 2005). 

Except in homes, the fungus occurs on mine timber and rarely in the open 

(poles, sleepers), but in the boreal climate not in the forest. However, according 

to Pegler (1991), the species occurs outdoors in Central Europe and North 

America, and according to Bech-Andersen (1995), in the Himalayas in conifers 

forests. Phylogenetic trees based on the rDNA-ITS sequence showed that the 

outdoor isolates from the Himalaya and from California belong to the species 

S. lacrymans (Chap. 2.4.2.2). Phylogenetic analyses indicate that the indoor 

isolates of S. lacrymans may have originated from an ancient lineage closely 

related to the Californian outdoor isolates (Kauserud et al. 2004b). 

In the open, the Wild merulius S. himantioides (Fig. 8.21e) is common, in 

Europe frequently on spruce wood, stumps, structural timber in outdoor use, 

and rarely on living trees. Occasionally, it is also found in buildings (Falck 

1927; Harmsen 1978; Grosser 1985; Seehann 1986; Pegler 1991). 

As further dry-rot fungi occur three Leucogyrophana species (Fig. 8.21f–o) 

in the forest on fallen stems and branches, and on wood in indoor use: L. mollusca, 

L. pinastri (Schulze and Theden 1948; Siepmann 1970) and L. pulverulenta 

(Harmsen 1953). They differ from Serpula by smaller spores (Ginns 1978; 

Pegler 1991; Breitenbach and Kränzlin 1986). Leucogyrophana pulverulenta is 

rather common in Denmark. The three fungi need a higher wood moisture 

content than S. lacrymans (cf. Table 8.7). 

Whereas S. lacrymans is restricted in North America to the northern parts 

of the USA and Canada, the American dry rot fungus Meruliporia incrassata 

(first reported in the USA in 1913) occurs particularly in the southern states 

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and the Pacific northwest of the USA (Verrall 1968; Palmer and Eslyn 1980; 

Gilbertson and Ryvarden 1987; Burdsall 1991; Zabel and Morrell 1992; Eaton 

and Hale 1993; Jellison et al. 2004). Being a warm-temperature fungus, two 

isolates from the USA and Canada grew best between 22.5 and 25 ◦ Canddied 

after3weeksofculturingatabout35 ◦ C (Schmidt 2003). Burdsall (1991) named 

24–30 ◦ C as the optimal temperature range for growth and above 36 ◦ Casthe 

lethal temperature. Jellison et al. (2004) quoted 28–30 ◦ C as the optimal range 

for growth, and 3–30 h at 40 ◦ C for lethal. Sapwood and heartwood of many 

gymnosperms and angiosperms are attacked. It was rarely found on standing 

trees, infrequently on felled logs and stumps, on structural timber outdoors 

such as in mills, lumber yards, on shingles, on bridge timber, posts, but is common 

on moist wood or wood located near a permanent or intermittent water 

supply if the wood is untreated (Palmer and Eslyn 1980). Some characteristics 

ofwooddecaybythisfungusaresimilartothoseofS. lacrymans, notably 

its sensitivity to dryness by mostly dying in pure culture tests with southern 

pine blocks of 30% wood moisture at 90% RH at 27 ◦ C(PalmerandEslyn 

1980), and its ability to transport nutrients and water from a feeding source to 

the advancing mycelial front spreading over non-wooden mortar and bricks. 

Pictures of mycelium and strands are by Zabel and Morrell (1992). 

The Serpula and Leucogyrophana speciesaswellasM. incrassata can be 

differentiated by their fruit bodies and strands (Appendix 1). Molecular techniques 

separate the vegetative mycelia (Chap. 2.4.2). The following description 

is based on observations and measurements in buildings and on results from 

wood samples in laboratory tests (Huckfeldt and Schmidt 2005; Schmidt and 

Huckfeldt 2005) and is supplemented especially for M. incrassata from Palmer 

and Eslyn (1980), Gilbertson and Ryvarden (1987), and Burdsall (1991). 

Serpula lacrymans, (True) Dry rot fungus 

Fruit body (Fig. 8.21a, b): annual to perenniell, resupinate to effused-reflexed 

and imbricate, sometimes stalactite-like, rust-brown, old: black; bulging, 

white-yellowish, sharp margin; fleshy-thick (to 12 mm), to 2 m wide, hymenophore 

merulioid; first monomitic, later dimitic containing fibers; yellowbrown, 

thick-walled spores 9−12 × 4.5−6µm; tetrapolar; 

Strands (Fig. 8.21d): young: white; old: grey-brown; to 3 cm in diameter, 

audibly breaking when dry, embedded in flabby mycelium; fibers 3–5 µm 

thick, hardly septate, without buckles, straight, rigidly, refractive; vessels to 

60µm thick, with bar-likes or warty wall thickenings, not or rarely branched. 

Serpula himantioides, Wild merulius 

Fruit body (Fig. 8.21e): annual, resupinate, sometimes membrane-like, rustbrown; 

white, sharp, not bulging margin, < 2 mm thick, hymenophore smooth 

to merulioid; yellow-brown, thick-walled spores 9−12 × 5−6µm; tetrapolar; 

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Strands: white to grey-brown, about 1 mm in diameter, microscopic characteristics 

similar to S. lacrymans. 

Leucogyrophana mollusca, Soft dry rot fungus 

Fruit body (Fig. 8.21k): resupinate, orange to yellow-brown; old: grey-blackish; 

white, cottony-frayed margin; 1–2 mm thick, to a few decimeters wide, easily 

separable; hymenophore merulioid, tooth-like elevations; uneven, brownviolet 

to grey-black sclerotia (Fig. 8.21m), 1–6 mm, often in groups; yellowishbrown 

spores 6−7.5 × 4−6µm; 

Strands (Fig. 8.21l): hair-like, first cream-yellow, soon brown-black, below 

1 mm thick, separated from mycelium (“barked”), flexible when dry, fragile 

when old; no fibers; vessels up to 25µm thick, numerous, in groups, with 

bar-thickenings. 

Leucogyrophana pinastri, Mine dry rot fungus, Yellow-margin dry rot fungus 

Fruit body (Fig. 8.21f, g): resupinate, first yellow-orange, then olive-yellow to 

brown, grey-black when old, to 1 m wide, hymenophore merulioid to irpicoid 

to hydnoid; round-oval, brown-black sclerotia to 2–3 mm thick; hyaline to 

yellowspores5−6×3.5−4.5µm; 

Strands: first yellowish, then grey-brown (Fig. 8.21h), hair-thin, separated 

from mycelium; no fibers; vessels to 15µm thick, numerous, in groups, with 

bar-thickenings. 

Leucogyrophana pulverulenta, Small dry rot fungus 

Fruit body: resupinate, first sulphur-canary yellow, then (Fig. 8.21n) oliveyellow 

to cinnamon-brown, also grey-black when old, white, indistinct margin, 

to 20 cm wide; hymenophore smooth to merulioid, no sclerotia; hyaline to 

yellow, thick-walled spores 5−6 × 3.5−4.5µm; 

Strands (Fig. 8.21o): white, to 2 mm thick, not clearly separated; no fibers; 

vessels to 20µm thick, numerous, in groups, bar-thickenings indistinct or 

absent. 

Meruliporia incrassata, American dry rot fungus 

Fruit body: similar to S. lacrymans, annual, resupinate to effused, 20 cm or 

more in length, thin, easily separable, whitish to buff margin, grey center, 

becoming darker as it matures; 1 to 12 mm thick, fleshy, brittle when dried; 

first appearing as a felted pad of mycelium with formation of pores beginning at 

the center, subsequent fertile to the margin; hymenophore poroid, occasionally 

merulioid; whitish to buff or ochre-grey when fresh, grey-brown to black when 

drying, unequally circular to angular pores, 1–3/mm; monomitic; thick-walled 

oblong to ellipsoid spores, variable in size, 8−16 × 4−8µm; 

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Strands: first as vein-like structures in the mycelium, often extending into 

soil or masonry, appearing whitish when young, browny-black with age (Eaton 

and Hale 1993), 0.3–5.1 cm in diameter, length up to 9 m (Palmer and Eslyn 

1980). 

Recognition characteristics of S. lacrymans (Fig. 8.21) 

Wood: The relatively large cubes of the brown-cubical rot (Fig. 7.1a) are no 

reliable characteristic. Painted doorframes or baseboards first show blisters 

and fine tears in the lacquer and after longer infestation, wavy surfaces. 

Fruit body: The brownish, to 12 mm thick and 2 m size, mostly resupinate 

fruit body growing on wood or masonry (Fig. 8.21a) is conspicuous. From 

shakes and vertical planes grow pad and bracket-like fruit bodies. The gyrosoreticulate 

hymenophore is traditionally named “merulioid” (Fig. 8.21b), which 

derives from the former generic name Merulius. The margin is whitish, often 

bulging and always with a sharply limited front. Particularly at the margin, 

as also with the mycelium, arise liquid drops of neutral pH value due to 

guttation, which led to the naming lacrymans (watering). Fresh fruit bodies 

have a pleasant smell like fungi, but putrefy after sporulation and then easily 

stink (from the ammonia). The old, dry, then black-brown fruit bodies hardly 

show the merulioid structure. Fruit bodies develop over the whole year, with 

an amassment in the late summer until winter (Nuß et al. 1991). 

Affected areas are often widely covered with brown, elliptical, yellow-brown 

spores with small, pointed extension at an end and partly with up to five intracellular 

oil droplets (Hegarty and Schmitt 1988; Pegler 1991; Nuß et al. 1991). 

Falck (1912) calculated the spore release by a 1-m 2 fruitbodyto3×10 9 spores 

per hour. 

First, however, inconstant fructification in the laboratory culture was obtained 

by Falck (1912), Cymorek and Hegarty (1986b) stimulated fructification 

by 12 ◦ C incubation and by natural temperature change in the open (cool) 

(Hegarty and Seehann 1987; Hegarty 1991). Fruit bodies relatively often developed 

in pure cultures, if the mycelium was first incubated for about 4 weeks at 

25 ◦ Conmaltagarandthenatabout20 ◦ C and natural daylight (Schmidt and 

Moreth-Kebernik 1991b; Fig. 3.1). 

Mycelium (Fig. 8.21c) and biology: During initial growth, with sufficient humidity 

and standing air, often a white, woolly thick aerial mycelium develops, 

which is rapidly interspersed by the typical strands. Yellow to wine-red (also 

violet) discolorations (“inhibition colors”) by restraining influences [light, 

accumulation of toxic metabolites, increased temperature: Zoberst (1952), 

Cartwright and Findlay (1958)] are characteristic and led to the former generic 

name Merulius, going back to the yellow beak of the male blackbird Turdus 

merula (Coggins 1980). Older mycelium collapses to removable, dirty grey to 

silvery skins, in which the branched strand system is embedded. The match- 

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to pencil-thick, up to 2 to 4-m-long, grey-brown and on their surface fibrously 

roughened strands (Fig. 8.21d, Table 2.4; Falck 1912) break when being dry with 

audible cracking. Strands are formed only in aerial mycelium, and there as well 

by dikaryotic as by monokaryotic mycelium, and not in substrate mycelium 

and reach (at 20 ◦ C) 5 mm length increase per day (Nuß et al. 1991). 

The fungus is tetrapolar heterothallic. Only dikaryons show clamps (Harmsen 

et al. 1958), while only monokaryons form plentifully arthrospores 

(Schmidt and Moreth-Kebernik 1991c). Contrary to Antrodia sinuosa and 

Coniophora puteana, the clamps are as large as the hyphal diameter (Nuß 

et al. 1991). Matings between different isolates of S. lacrymans revealed physiological 

differences between the different mycelial types, but also constancy 

of the characteristics over several generations (Schmidt and Moreth-Kebernik 

1989b, 1990, 1991a): The dikaryons (parents and F1 and F2 generation) grew 

significantly faster than the mycelia of the two appropriate monokaryons and 

the two heterokaryon types (A# B=, A= B#). Regarding wood decay, dikaryons 

and monokaryons showed greater activity than the heterokaryons (also Elliott 

et al. 1979). Monokaryons and heterokaryons however tolerated higher temperature 

than the dikaryons, by growing still at 28 ◦ C. Monokaryons also endured 

higher protective agent concentrations and this was also proven for Antrodia 

vaillantii and Gloeophyllum trabeum (Da Costa and Kerruish 1965). Related to 

practice, such physiological differences between the different mycelial types 

could become relevant, since dikaryons can revert under adverse conditions 

to the monokaryotic stage, as for example G. trabeum by arsenic (Kerruish 

and DaCosta 1963) and S. lacrymans by relatively high temperature (Schmidt 

and Moreth-Kebernik 1990). The more tolerant monokaryons would survive 

and can mate under again favorable conditions to dikaryons and thus have 

overcome the adverse environment. 

The vegetative hyphae in the aerial mycelium are thicker (about 6µm) than 

the hyphae within woody tissue, with about 2µm. Within wood, medallion 

clamps also occur. The distance between the two clamps is shorter than in 

aerial mycelium, and often almost right-angled hyphal branching occurs. Morphologic 

characteristics of mycelium, fruit body, and spores were described by 

Nuß et al. (1991). 

Conifers are preferred. Hardwoods with dark heart like oak and chestnut 

are more resistant than light species (Wälchli 1973). Beside wood and masonry, 

composite woods (chipboards, fiberboards), carpets, and textiles are attacked 

and insulating materials (Grinda and Kerner-Gang 1982) like mineral wool are 

through-grown and damaged (Bech-Andersen 1987b). 

Because of the relatively low optimal temperature range of 17 to 23 ◦ C, the 

mycelium grows preferentially in the cooler cellar and ground floor areas. The 

total span reaches from 0 to 26–27 ◦ C, and growth stops at 27–28 ◦ C, which 

differentiates the species from the similar S. himantioides. The mycelium died 

on agar at 55 ◦ C for 3 h (Table 3.8, also Mirič and Willeitner 1984). In dried 

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wood samples, however, only 70 ◦ C for 4 h were lethal (Huckfeldt et al. 2005). 

The spores were killed after 1 h at 100 ◦ C (Hegarty et al. 1986). Thus, hotair 

treatment procedures of attacked buildings (see below), as they are used 

in Denmark and also proposed for Germany, kill neither the spores nor the 

hyphae growing within large-dimensioned timbers and masonry. 

The minimum wood moisture for initial colonization is 21% u (Huckfeldt 

2003). The opinion has it that this infection of wood below the fiber saturation 

range of about 30% is possible, because the Dry rot fungus is particularly 

effective to transport nutrients and water by means of mycelium and strands, 

and here particularly by the vessel hyphae, from a moist nutrient source [wood 

over fiber saturation or wet masonry: Dickinson (1982)] to the infestation of 

“dry wood” (Wälchli 1980; Jennings 1987, 1991; Coggins 1991; Savory 1964). 

Not to stamp out, even in recent publications, is the erroneous opinion that 

S. lacrymans is extraordinary to colonize dry timber by the exclusive water 

production via its own enzymatic wood decay (Chap. 3.3). Also incorrect is 

that it takes up the necessary water from the air humidity. 

Compared to Cellar fungus and the indoor polypores, the Dry rot fungus 

was considered to be sensitive to high wood moisture content (Cartwright 

and Findlay 1958). There is an older reference that it even reduced high wood 

moistures by guttation in favor of higher air humidity (Miller 1932). The 

optimal wood moisture for initial decay is about 30–40% u and shifts with 

longer decomposition rather to 40–60% (Wälchli 1980). The maximum of 

about 90% (Wälchli 1980) was higher than the 55% moisture content often 

cited in the older literature. In piled wood samples (Table 8.7), the optimum 

wood moisture was between 45 and 140%, and even samples with initial values 

of 240% wood moisture were decayed with wood mass loss over 2% (Huckfeldt 

and Schmidt 2005), so that the total span reached from 21 to 240%. The common 

term in English “Dry rot fungus” (Savory 1964; Coggins 1980; Bravery et al. 

2003) and in German “Trockenfäule-Erreger” is paradoxical, since the Dry rot 

fungus also (like all other decay fungi) needs free water in the cell lumina 

for the enzymatic wood decay and is susceptible to desiccation. By means 

of mycelium (and strands), the fungus transports beside nutrients and water 

also minerals, e.g., the wood-decay limiting nitrogen (Watkinson et al. 1981) 

from the soil under a house to wood decay in the interior (Doi 1989; Doi 

and Togashi 1989; also Weigl and Ziegler 1960; Jennings 1991). After Savory 

(1964), the main significance of the strands lies in the nutrient translocation 

and not in the water transport (also Bravery and Grant 1985). Literature data 

to the requirements for temperature and humidity are also by Viitanen and 

Ritschkoff (1991a). 

The mycelium of S. lacrymans is said to show dryness resistance of many 

years. However, the few experiments available revealed that it can reach at least 

under laboratory the dryness resistance only by a slow moisture removal. Assumably,themyceliumneedstimetorevertfirstintothemonokaryoticstage 

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with its resistant arthrospores. Furthermore, the resistance at 20 ◦ Camounted 

only about 1 year. Only at low temperature (7.5 ◦ C), the fungus survived several 

years (Theden 1972; also Savory 1964). Nevertheless, the remaining infected 

areas form a danger potential for new growth. Infected timber parts can exhibit 

just so much moisture to enable a slight growth and thus a longer survival 

than by means of dryness resistance (Grosser 1985). Furthermore, the danger 

of re-infection may derive from the dryness-resistant spores, whose duration 

of germ ability was said to amount to 20 years. In infected buildings, S. 

lacrymans frequently produces basidiospores, and basidiospores seem to be 

the main agent of dispersal (Falck 1912; Langendorf 1961; Schultze-Dewitz 

1985). Vegetative spread by mycelium and strands seems to be restricted to 

within buildings or the soil in subfloor space (Doi 1991). However, according 

to Wälchli (1980) the infection occurs instead by mycelium that is brought in 

with timber from other remedial treatments and via wooden boxes or shoes. 

Beside the requirement for low temperature, the preferential indoor occurrence 

of S. lacrymans was attributed to the intensive synthesis and secretion of 

oxalic acid (Jennings 1991; cf. Table 3.9), whose excessive production was said 

to be neutralized as calcium oxalate by calcium from masonry or by chelating 

with iron from girders (Bech-Andersen 1985, 1987a, 1987b; cf. Palfreyman 

et al. 1996). Oxalic acid is also implicated in copper tolerance of fungi. Although 

a single isolate of S. lacrymans was only able to grow on agar at a low 

concentration of copper sulphate (Table 3.10), Haustrup et al. (2005) showed 

11outof12isolatestobetolerantagainstcoppercitrate.Theimplicationof 

calcium in oxalate precipitation was also shown for M. incrassata (Jellison et al. 

2004). Thus, dry rot attack in buildings is often found in the ends of beams, 

which are not separated from the masonry. 

During controversies, e.g., in the context of house buying, frequently the 

question of the infection date plays a role, for whose determination the daily 

average mycelial growth is often used. According to Jennings (1991), the linear 

mycelial extension on wood, masonry and insulants ranges from 0.65 to 

9mm/d. Assuming a 5-mm radial increase per day on malt agar at optimal 

temperature (Table 2.2), 15 cm follow per month. Due to the changing and 

not always optimal conditions in buildings and because different isolates of 

the fungus exhibited considerable differences in growth rate [1.5–7 mm/d: Cymorek 

and Hegarty (1986a); Seehann and v. Riebesell (1988)], an exact age 

determination on the basis of the mycelial extension is impossible. Similarly, 

the decay of pine sapwood samples varied among 25 isolates from 12 to 56% 

in 6 weeks of cultivation (Cymorek and Hegarty 1986a; Thornton 1991), and 

different isolates differed likewise in their sensitivity to wood preservatives 

(Abou Heilah and Hutchinson 1977; Cymorek and Hegarty 1986a; Wa˙zny and 

Thornton 1989a, 1989b, 1992; Wa˙zny et al. 1992). Important is also the decision 

if the mycelium in a building is alive or dead. Subculturing on malt agar is 

possible, but isolations from mycelium are often contaminated by molds. Vital 

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staining with fluorescein diacetate is suitable (Huckfeldt et al. 2000; also Koch 

et al. 1989; Bjurman 1994). 

The possibilities to identify S. lacrymans cover the classical methods of fruit 

body investigation (Grosser 1985; Pegler 1991), strand diagnosis (Falck 1912; 

Table 2.4, Appendix 1), and mycelium analysis by identification key (Stalpers 

1978). As modern techniques, protein polyacrylamide gel electrophoresis 

(Schmidt and Kebernik 1989; Vigrow et al. 1989; Palfreyman et al. 1991; 

Fig. 2.19) and immunological tests (Palfreyman et al. 1988; Vigrow et al. 1991c; 

Toft 1992, 1993; Glancy and Palfreyman 1993) were tested for suitability. DNA 

techniques have been established (Schmidt 2000) and are already used commercially. 

MALDI-TOF mass spectrometry was capable of differentiating the 

mycelium of the True dry rot fungus and its closest relative the Wild merulius 

(Schmidt and Kallow 2005; Fig. 2.24). Measurement of microbial volatile organic 

compounds (MVOCs) may identify wood-decay fungi (Bjurman 1992b). 

Pinenes, acrolein, and ketones were found in Serpula lacrymans, Coniophora 

puteana, and Oligoporus placenta (Korpi et al. 1999). Mono- and sesquiterpenes, 

aliphatic alcohols, aldehydes and ketones, and some aromatic compounds 

were emitted by Fomitopsis pinicola, Piptoporus betulinus, and further 

species (Rosecke et al. 2000). Blei et al. (2005) showed that MVOC analysis 

was able to distinguish pure cultures of Antrodia sinuosa, C. puteana, Donkioporia 

expansa, Gloeophyllum sepiarium, S. lacrymans, and S. himantioides. 

Field experiments, however, were influenced by the distance of sampling from 

the infested and/or destroyed wood and also by the rates of air changes. 

To improve the technique of MVOC analysis, Keller et al. (2005) measured 

volatile compounds in non-infested living and bedrooms as a background 

reference for infestation. Trained sniffer dogs can also detect S. lacrymans 

(Koch 1991). 

If S. lacrymans is proven, the fungus is (beside longhorn beetle and termites) 

the only biological damage causer for which there is the obligation in 

some German states (Hamburg, Hessen, Sachsen, Thüringen, and Saarland) 

to become registered. Since costs of refurbishment can be considerable (to 

e3,000 per m 2 living space), the determination of the extent of the damage and 

theremedialtreatmentsshouldbedonebyarenownedcompany.InGermany, 

refurbishment has to follow the standard DIN 68800 part 4. In the case of a lawsuit, 

§459 of the German Civil Code regarding “regress for material defects” 

takes effect. 

8.5.4 

Prevention of Indoor Decay Fungi and Refurbishment of Buildings 

All decaying fungi need water for wood decay. Elimination of the source of 

moisture and drying of wood and masonry after prolonged wetting are the 

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most important remedial treatments. Since S. lacrymans can transport water, 

it cannot be excluded that sources of dampness are overlooked during repair, 

and thus more-extensive measures are necessary for its control. 

The first remedial treatment of dry rot infestation is described in the Bible 

in Leviticus 14:33–48. Preventive measures against all house-rot fungi are 

avoidance of general building defects and of those during refurbishment of old 

buildings: moisture ascending in the masonry, seeping rain water, insufficient 

ventilation, installation of wet or infested timber and wet fillers, allside walled 

beam ends, lack of building drainage, condensation water by wrong thermal 

insulation and inappropriate vapor barriers, unsatisfactory underside blockage 

of buildings without cellars, wrong structure of floors, reuse of attacked 

building debris, leakages in bathrooms and insufficient wood protection. 

To the common causes belong also unrepaired building damage: leaky roofs, 

shattered windowpanes, leaky or sweating water and heater lines, clogged or 

defective rainwater and drainage facilities as well as water damage caused by 

burst piping, defective washing machines and dishwasher water pipelines, cellar 

floodings and fire-fighting water (Thornton 1989a; Paajanen and Viitanen 

1989; Bricknell 1991; Doi 1991; Wälchli 1991). 

Particularly regarding cellar fungi, flooring in new buildings should not 

done too early. Damp bulk goods in ceilings shall be avoided. 

The danger of infestation exists via spores and by infected timber and 

wooden boxes, which are stored as firewood in damp cellars, and by mycelium 

viatheshoesofworkers. 

If a fungus is found, it should be first determined whether it concerns S. 

lacrymans or another fungus, as this decision may require the obligation to 

register the fungus and influences the extent of remedial treatments. In cases of 

doubt, laboratory identification should be performed by appropriate institutes, 

national testing institutions, offices for plant protection or in the laboratories of 

wood preservative manufacturers. The German standard DIN 68800 demands 

that if an exact species identification is not possible, then refurbishment is to 

be proceeded in such a way, as if the True dry rot fungus were present. 

Then the extent of the damage has to be established. German guidelines for 

control measures are listed in Table 8.8 (Grosser et al. 2003). 

Table 8.8. German guidelines for control measures during refurbishment 

DIN 68800 Part 4: Wood preservation; control measures against wood-destroying fungi 

and insects, issue 1992 

Part 3: Wood preservation; protective chemical wood preservation, issue 1990 

Part 2: Wood preservation in building construction; protective structural measures, 

issue 1984 

DIN 52175: Wood preservation; term, fundamentals, issue 1975 

Concretization rule for building work (VOB part B) 

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Refurbishment methods are described by Grosser (1985), Blow (1987), 

Wälchli (1991), Bech-Andersen (1995), Gründlinger (1997), Sutter (2003), 

Bravery et al. (2003) and Grosser et al. (2003), briefly: Elimination of the 

source of moisture, removal of all infected timber 1 m beyond the last evidence 

of fungus or decay, disposal of the attacked timber and the other infected 

building materials, physical (heat) and chemical treatment (boron, quaternary 

ammonium compounds) of infested masonry with certified preservatives 

for those species that colonize brickwork, use of preservative-treated 

timbers for replacement following DIN 68800, and providing adequate ventilation. 

Eradication in the roof space with hot air as it is used against insects (Paul 

1990) is already done or is being considered to fight fungi in some European 

countries (Koch 1991; Sallmann 2005). However, first these treatments are 

technically wrong in view of a safe killing of mycelium and spores of house-rot 

fungi in wood and in masonry, since the necessary heat (Schmidt and Huckfeldt 

2005; Huckfeldt et al. 2005; Table 3.8) is not obtained, particularly not in the 

inside of thick timber. Second, heat treatment is economically doubtful due 

to the endangerment of the structural fabric and third, from an ecological 

viewpoint, enormous energy is needed. 

Microwaves are also used or being considered as an alternative method. 

Irradiation tests with microwaves from 1990 to 1992 in Denmark in about 100 

cases of fungal infestation killed the mycelium of S. lacrymans that previously 

had been inserted into the brickwork within 10 min (Bech-Andersen and Andersen 

1992; Kjerulf-Jensen and Koch 1992). However, microwave treatment 

is a fire risk if metal fastenings are present in the timber (Bravery et al. 2003) 

and there are general doubts on the suitability of the technique for buildings 

(Sallmann 2005). 

For registered historical buildings and wood artifacts, the suitability of fumigants 

was tested mainly for the control of insects, but also to control decay 

fungi. Against fungi, bromomethane and ethylene oxide have been used (Unger 

et al. 2001). Fumigants, however, do not provide protection against new infestations. 

In the laboratory, aminoisobutyric acid, which is analogous to the amino 

acid alanine, reduced the decay of wood samples by S. lacrymans from 22 to 

1% (Elliott and Watkinson 1989). An intervention in the trehalose metabolism 

of S. lacrymans was suggested to influence the internal translocation processes 

(Jennings 1991). The binding of iron by chelating agents inhibited mycelial 

growth, EDTA prevented decay of pine samples by Coniophora puteana, Gloeophyllum 

trabeum and Oligoporus placenta (Viikari and Ritschkoff 1992), and 

tellurium acid wood decay by C. puteana (Lloyd and Dickinson 1992). Polyoxin 

acted as inhibitor of the chitin synthase of several fungi (Johnson and 

Chen 1983). Particularly the Trichoderma species display a wide arsenal of 

antagonistic mechanisms that make these fungi attractive as biological control 

agents (Highley and Ricard 1988; Giron and Morrell 1989; Doi and Yamada 

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1991; Rattray et al. 1996; Bruce 2000). Bacteria decreased wood decay by O. 

placenta (Murmanis et al. 1988; Benko and Highley 1990). 

From a biological point of view, there is no reason that all indoor wood 

decay fungi should be a problem. The biological requirements of the common 

species are known. Control measures are straightforward. Even once a fungus 

is established, it is mainly only necessary to change the conditions in the 

building to a long-term removal of moisture. There was only slight wood decay 

by some house-rot fungi below the fiber saturation range of about 30% u. The 

lower limit for decay of pinewood samples (mass loss slightly over 2% within 

5 months) was 22% (Table 8.7). This also applies to the feared S. lacrymans. 

This fungus turned out in many laboratory tests on temperature and drying 

effects to behave rather sensitively when compared to the cellar fungi and 

the indoor polypores. The only biological specific features of S. lacrymans are 

its more highly developed strand system to transport nutrients from a moist 

feeding source over considerable distances and to colonize new substrate, its 

formation of thick surface mycelium that prevents the colonized wood from 

drying, and its ability to grow through masonry. 

The most important measure against all fungi in buildings is to detect and 

eliminate the cause of the increased moisture content of wood and masonry 

that is in contact with wood as well to exclude any re-moistening, including 

throughcondensationandfaultsbythehomeuser.Ifthedestroyedtimber 

has been replaced and lasting dryness of the wood can be guaranteed, there is 

no need for further provision, from the biological view, as there is no fungus 

known which destroys dry wood (below 22% u), not even S. lacrymans.Since 

practice, however, shows that in many cases a lasting dryness cannot be ensured 

in buildings, there are specific recommendations (and in Germany regulations) 

forthecaseofS. lacrymans infestation. 

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9 

Positive Effects of Wood-Inhabiting 

Microorganisms 

Particularly after the OPEC oil embargo of the 1970s, research turned towards 

the utilization of renewable resources like wood, yearly plants, and lignocellulosic 

waste from forestry and agriculture instead of oil as raw material for 

chemical and biological processes (“biotechnology of lignocelluloses”) (Eriksson 

et al. 1990; Dart and Betts 1991). 

Among the substantial causes that make the biological conversion of lignocelluloses 

difficult (Table 4.2), the most serious obstacle is the incrustation 

of the degradable carbohydrates cellulose and hemicelluloses by the lignin 

barrier, which is not surmountable by most microorganisms. Table 9.1 groups 

some bioconversions that have been done in the past or are recently investigated 

or already performed into those microbial processes, which go well 

directly with lignocelluloses, and into those, which need a pretreatment of the 

substrate. Only the wood-degrading white, brown, and soft-rot fungi, and the 

wood-degrading bacteria can degrade the native woody cell wall without any 

pretreatment of the substrate. Whereas brown and soft-rot fungi and assum- 

Table 9.1. Biotechnological procedures with lignocelluloses without and after substrate 

pretreatment 

conversion without substrate pretreatment 

– “myco-wood” 

– production of edible mushrooms 

– biological pulping 

pretreatment of the substrate and subsequent microbial conversion 

biological pretreatment 

– “palo podrido” and “myco-fodder” 

chemical pretreatment 

– hydrolysis of wood with acids and use of glucose for yeast production, ethanol 

fermentation and microbial transformations to amino acids, antibiotics, enzymes, 

vitamins 

– sulphite pulping process and use of hardwood pentoses in the spent liquor for yeast 

production and of softwood hexoses for ethanol fermentation 

– pulping and subsequent use of enzymes for deinking of waste paper 

physical pretreatment 

– grinding of lignocelluloses to improve accessibility to enzymes 

– steam explosion methods to open the wood structure for bioconversions 

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238 9 Positive Effects of Wood-Inhabiting Microorganisms 

ably also the wood-degrading bacteria only clear the hurdle of lignification, 

exclusively the white-rot fungi and their ligninolytic system additionally use 

the lignin as a carbon source and are therefore predestined for bioconversions 

(Table 4.3). All other microorganisms as well as their isolated enzymes need 

first a pretreatment of the substrate wood, which loosens the chemical/physical 

association of carbohydrates and lignin or reduce the lignin content or improve 

the physical accessibility of the degrading agents to the substrate. The various 

possibilities of a pretreatment can be grouped into biological, chemical, 

and physical methods (Dart and Betts 1991). Saddler and Gregg (1998) distinguished 

four main pretreatment methods currently being researched and 

commercialized to make lignocelluloses more easily digestible to hydrolytic 

enzymes while preserving the yield of the original carbohydrates for bioconversions: 

organosolv, steam explosion, dilute-acid prehydrolysis, and ammonia 

fiber explosion. Some of the bioconversions described below like “myco-wood” 

or “palo podrido” may occur a little strangely to some readers, but are examples 

that wood bioconversion can work. 

9.1 

“Myco-Wood” 

In Eberswalde, Germany, around 1930, J. Liese started to cultivate edible mushrooms 

on wood like Flammulina velutipes, Kuehneromyces mutabilis, Lentinula 

edodes (Fig. 2.17a) and Pleurotus ostreatus to improve the food situation 

of the population (Liese 1934). Due to the import stop of wood from overseas 

into the German Democratic Republic (GDR) at that time which was 

needed for pencils etc., his student, W. Luthardt thought about a possible 

use of the wood substrate remaining after mushroom production to produce 

pencils and other form-stable products. In 1956, Luthardt got the patent for 

“myco-wood” for the GDR and in 1957 under license for the Federal Republic 

of Germany: “Myco-wood is a wood that is loosened through the controlled 

action of certain wood-inhabiting fungi and which has changed its technological 

characteristics to a large extent or may obtain defined technical qualities” 

(Luthardt 1969). For myco-wood production, 50-cm-long stem sections of Fagus 

sylvatica were inoculated on the crosscut surface with a mycelium paste of 

Pleurotus ostreatus or Trametes versicolor, respectively, and were incubated in 

the constant climate of former air-raid shelters for different periods. Through 

the controlled white rot, a white and porous raw material free from tension 

was obtained that showed improved carving and sharpening ability to be used 

for form-constant products like pencils, rulers, and drawing boards. For example, 

after 3 months of incubation, the wood showed 30% mass loss, was 

completely colonized by mycelium, and was now suitable for rulers. One of 

these rulers is still used in our laboratory and looks like newly manufactured. 

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9.2 Cultivation of Edible Mushrooms 239 

About 120 million myco-wood pencils were produced in the GDR from 1958 

to 1961. The microbially modified wood also showed faster water absorption 

and desorption and was thus used for wood forms of the glass industry. Due 

to water-vapor film between wood and glass, it was possible to produce 12,000 

goblets using a myco-wood form instead of 800 glasses using normal wood 

(Luthardt 1963). Attempts to produce myco-wood also took place with tropical 

woods (Eusebio and Quimio 1975; Arenas et al. 1978) and bamboo (W. Liese, 

pers. comm.). 

9.2 

Cultivation of Edible Mushrooms 

Although actual data could not be obtained, the worldwide production of edible 

mushrooms cultivated on straw and wood may be in the range of 2 million t 

(fresh weight basis) per year (Table 9.2), so that the cultivation of mushrooms 

represents the economically most important microbial conversion of lignocelluloses 

(Chang and Hayes 1978). 

Without knowledge of the biological background, about 2,000 years ago, 

the Shii-take, Lentinula edodes, (Fig. 2.17a) was already cultivated on wood 

Table 9.2. Production of edible mushrooms (after various reports in the journal “Der 

Champignon”) 

Year (× 1,000 t) (%) 

Mushrooms worldwide 1991 4,273 100 

Agarics (Agaricus spp.) 1,590 37.2 

Oyster mushrooms (Pleurotus spp.) 917 21.5 

Auricularia spp., Tremella spp. 605 14.2 

Shii-take (Lentinula edodes) 526 12.3 

Enoki (Flammulina velutipes) 187 4.4 

Nameko (Pholiota nameko) 40 0.9 

Grifola frondosa 2005 35 

Mushrooms worldwide 1997 6,344 100 

China 4,000 63.1 

Japan, Taiwan, Korea, etc. 1,005 15.8 

EU 908 14.3 

North America 431 6.8 

Shii-take worldwide 1997 1,322 100 

China 1,125 85.1 

Japan 133 10.0 

Taiwan, Korea 44 3.4 

EU 0.995 

France 0.450 

Germany 0.150 

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in Asia. The name Shii-take means “Pasania-fungus”, because the mushroom 

was grown on the “Shii-tree” (Castaneopsis (Pasania) cuspidata, Japanese chinquapin). 

For the cultivation of this excellently tasting mushroom (compared 

to the Shii-take, the commercially produced agarics taste like nothing) as “natural 

log cultivation”, originally in China, and later in Japan, logs and branch 

sections were exposed to natural, passive inoculation by wind-borne spores 

and were stacked in the forest for fruit-body formation. About 300 years ago, 

the Shii-take was cultivated by farmers for extra income to be sold on local 

markets. The bark surface of logs, particularly from Quercus serrata or other 

fagaceous trees, was broken with an axe to improve the chances of inoculation. 

Since the 1920s, pure spawn culture was placed (“spawning”) into holes 

drilled into the logs. For the colonization phase of the substrate by mycelium, 

the inoculated logs were first placed as stacks in the forest or in greenhouses 

until the mycelium grew out. The colonized woods were then set up individually 

or stacked crosswise in the forest (“growing yard”) or in greenhouses for 

fruit body formation. Eight to 12 months after inoculation, there is the first 

flush of mushrooms, and cropping of logs occurs over about 5 years. Since 

the 1970s in Taiwan, Japan, and China, the Shii-take is produced commercially 

on chopped wood (chips) and wood waste like sawdust under controlled 

conditions such as defined substrate composition, temperature, light conditions, 

relative humidity, and wood moisture content. The big breakthrough 

for sawdust substrates was the use of plastic bags, in which the substrate can 

be compressed, sterilized, inoculated, and grown out (Fig. 9.1). The woody 

substrate is supplemented with amendments (bran, whole meal, urea etc.), 

watered for a suitable moisture content, and inoculated with special isolates. 

In this “bag” or “artificial log” culture, the mycelium knits the substrate into 

a solid block. The methods for Shii-take production have been recently summarized 

by Miller (1998). In the local experiments (Schmidt and Kebernik 

1986; Schmidt 1990), different wood wastes such as chips (Fig. 9.1), sawdust, 

Fig.9.1. Shii-take (Lentinula edodes) 

fruit-bodies grown on wood waste chips 

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leaves and needles from some hardwoods and also from spruce and pine were 

used. A short colonization phase of the substrate was done at about 24–28 ◦ C 

in closed plastic bags or similar containers. Readiness of the Shii-take to fruiting 

became visible in the closed bag, when brown-black wet spots occurred 

between the white mycelial mat along the outer surface of the artificial log 

and the bag. Then the substrate was a solid block, and the substrate containers 

were opened or removed for fruiting at lower temperature of about 

12–20 ◦ C. After bag removal, the outer mycelial surface becomes brown and 

leathery. Then the logs were sprayed with water once a day, and natural daylight 

in a greenhouse was used to stimulate primordia formation. The artificial 

light–dark cycle requires light in the 3,700 to 4,200-nm range and intensity 

of 400–500 lux (Miller 1998). Several flushes occur within 1 year. After each 

cropping, the dry substrate may be re-wetted e.g., by soaking in cold water. 

This soaking both replaces the water that has been lost by the growth of 

the fruit bodies and the cold stimulates the development of the next primordia. 

The yields amounted to about 100% biological efficiency (fungal fresh 

weight: dry weight wood; Royse 1985). In Taiwan, for example, 516 companies 

produced about 24,000 t of fresh fungi on chopped substrates in 1985, 

and a similar quantity was obtained, however, by over 5,000 farmers on wood 

sections. 

The Shii-take was for a long time the most common mushroom cultivated 

on wood worldwide. Altogether, the fungus was the second most frequent 

cultivated mushroom with its main production in Japan after the Agaricus 

species, which are traditionally cultivated on wheat straw that is composted 

with manure or some other nitrogen-rich additive. The Shii-take has been 

however overhauled through the increased production of Pleurotus species 

particularly in China. Worldwide 526,000 t Shii-take were harvested in 1991 

and 1.3 million t in 1997 (Table 9.2). Beside Asia, some Shii-take cultivation 

is performed in the USA, Canada, and Europe. In Germany, there is a handful 

of commercial Shii-take growers producing some hundreds of tons. A great 

part of Chinese and Japanese Shii-take is exported in dry condition to Taiwan, 

Singapore, USA, Canada, Australia, and Europe. In Germany, 100 g of dry, 

imported Shii-take cost about e10. The local market price varies for outdoorgrown 

fungi due to seasonal influences from e10 to 40 per kg fresh weight. 

Because of the slow growth of the Shii-take mycelium during the colonization 

phase, the cultivation on shopped substrates is endangered by contaminations, 

partly leading to parasitism, particularly by Trichoderma species like T. hamatum, 

T. harzianum, T. parceramosum,T. pseudokoningii, T. reesei and T. viride 

(Albert 2003). Thus, the colonization phase is commonly performed with 

pasteurized (60–100 ◦ C) ore autoclaved substrates in plastic bags (Schmidt 

1990). 

The fundamentals of Shii-take production are known outside of Asia. The 

first cultivations in Europe were performed by Mayr (1909) and Liese (1934; 

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Fig. 2.17a), and research on the biological and physical demands of the fungus 

were done in the USA (e.g., Leatham 1982; Royse 1985) and in Europe (e.g., 

Zadraˇzil and Grabbe 1983; Rohrbach 1986; Müller and Schmidt 1990; Lelley 

1991; Kalberer 1999). Basically, the procedure consists of four main steps as 

shown in Fig. 9.2: pre-culture of a certain isolate, propagation of the mycelium 

for inoculation by growth on sterile grains (spawn production), colonization 

phase of the sterilized substrate in plastic bags, and fruiting phase on the 

opened containers. 

The reasons why the Japanese and Chinese in particular have been so successful 

in Shii-take cultivation are not known. Generally, the cultivation of 

so-called “alternative or exotic mushrooms” has got to have the right feel for 

it. The Shii-take belongs to “demanding mushrooms” while the Oyster mushroom, 

Pleurotus ostreatus, is easily satisfied through its fast growth ability on 

several substrates such as lignocellulosic waste (Pettipher 1987) and is thus 

lesser sensitiveness to contamination. In North America and Europe, particularly 

in Italy and Hungary, frequently Pleurotus species such as P. ostreatus are 

grown on chopped wheat straw, but also stem sections (Fig. 9.3) or chopped 

waste is used by hobby breeders and commercially. The market price of this 

lesser-tasting fungus in Germany amounts to e5–10/kg fresh weight. Further 

fungi that are cultivated on lignocelluloses are e.g., Agrocybe aegerita, 

Auricularia auricula-judae, Flammulina velutipes, Grifola frondosa, Hericium 

erinaceus, Kuehneromyces mutabilis, andPholiota nameko (Miller 1998). Research 

results and practical tips for mushroom culturing occur in the German 

Fig.9.2. Main steps of Shii-take production: a Maintenance of a selected isolate on agar. 

b Mycelial growth on grains for inoculation. c Substrate colonization in closed plastic bags. 

d Fruiting phase after removal of plastic bag (from Schmidt 1990) 

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Fig.9.3. Pleurotus ostreatus cultivation 

on beech wood billets in Germany in 

1936 (photo J. Liese) 

journal “Der Champignon”. The international work is treated at the meetings 

of the International Mycological Society. 

Concerning the nutritional value of fungi, it may be considered that a fresh 

fruit body contains predominantly water and only about 10% dry matter. 

For 100 g of fresh Shii-take, 92.6 g water, 4.3 g carbohydrates, 1.9 g protein, 

0.3 g lipids, 0.7 g ballast material, and 0.5 g minerals have been measured, 

corresponding to 109 kJ. Minerals in decreasing order were K, P, chloride, Ca, 

Mg, Na, Zn, fluoride, Fe, and Cu. The vitamins comprised C, pantothenic acid, 

nicotine amide, E, B1, folic acid, and D (Schulz 2002; also Spiegel 2001). Thus, 

considering the high price of the tasty mushrooms species, their significance 

as food lies rather in culinary appeal. 

For thousands of years, mushrooms have been known as a source of medicine, 

particularly in Asia. Among these non-culinary mushrooms, e.g., Ganoderma 

species are grown on wood waste to obtain medically active compounds 

(Miller 1998). For example, he has shown that the methanol extract of the G. 

lucidum fruitbodyhasastronginhibitoryactivityofthe5α-reductase that 

is involved in the benign prostatic hyperplasia of older men (Liu et al. 2005). 

Those “medicinal mushrooms” are widely sold as a nutritional supplement 

and are touted as being beneficial to health. Asian people believe that the Shiitake 

has antivirus, antibactericidal, antitumour (e.g., Mori et al. 1989) and 

cholesterol-decreasing effects. In view of the possibly increased heavy metal 

content and radiation load that had been measured in some forest mushrooms, 

indoor-cultured fungi are harmless, but are usually lesser tasty than outdoorgrown 

fungi. In Asia, the quality of mushrooms grown in a bag or bottle culture 

is considered inferior. 

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9.3 

Biological Pulping 

Mechanical and chemical processes for pulp and paper production consume 

energy and chemicals. Their wastes have to be controlled in view of environmental 

aspects. Biotechnological processes have thus been successfully 

implemented in the pulp and paper industry during the last decade driven 

by the objective to reduce manufacturing costs using new delignification processes 

and by environmental considerations (Messner et al. 2003). The application 

of white-rot fungi, or their ligninolytic systems, was one option for 

this. The aim was termed as biological pulping or briefly biopulping. In its 

strict sense, biopulping was defined as the pretreatment of wood chips with 

selectively delignifying white-rot fungi prior to mechanical or chemical pulping 

(Messner 1998). In a broader sense, the term biopulping is also used for 

any biochemical assistance to the pulping process such as the application of 

blue-stain fungi for resin reduction or the use of enzymes for bleaching and 

deinking. 

Nilsson had found Sporotrichum pulverulentum (first termed Chrysosporium 

lignorum) in chip piles in Sweden, where it caused serious damages (Bergman 

and Nilsson 1966). In 1972, Henningsson et al. described the fungus as the 

thermophilic white-rot basidiomycete Phanerochaete chrysosporium (teleomorph 

of S. pulverulentum) causing defibration of wood. In the late 1960s, 

Eriksson in Stockholm had already started research to decrease the lignin 

content in the wood microbially by treatment of wood chips with white-rot 

fungi (Eriksson 1985; Eriksson et al. 1990). Mechanical pulp was produced 

from chips pretreated with P. chrysosporium by Ander and Ericksson (1975). 

Because white-rot fungi of the “selective delignification type” would also attack 

the carbohydrates sooner or later, cellulase-less mutants such as Cel 44 of S. 

pulverulentum have been produced by UV irradiation of conidia (Ander and 

Eriksson 1976) and later by crossing of Cel − -mutants with monokaryons of 

high ligninolytic activity (Johnsrud 1988). 

Phanerochaete chrysosporium has also been isolated in the USA in the Arizona 

desert (Burdsall and Eslyn 1974). Also in the late 1960s, Kirk in Madison 

began research on P. chrysosporium with the isolation of lignin peroxidase (Tien 

and Kirk 1983; see Chap. 4.5), and since the 1980s, biopulping is investigated 

in the USA (Kirk et al. 1993). 

There were masses of investigations and publications on various aspects 

of biopulping during the past four decades. They report on the successful reduction 

of chemicals and manufacturing and energy costs as well as on the 

application of further white-rot fungi such as Ceriporiopsis subvermispora, 

Dichomitus squalens, Merulius tremellosus, and Phlebia brevispora. For example, 

when biopulped chips are used to produce mechanical pulp, energy for 

refining was reduced from 25 to 35% and the sheet strength properties are 

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9.3 Biological Pulping 245 

typically improved 20 to 40% (Hunt et al. 2004). A 20% reduction was obtained 

in the total pulping time necessary for achieving pulp and paper properties 

comparable to those from controls (Chen et al. 1999). Körner et al. (2001) 

showed that non-sterile incubation of wood chips with Coniophora puteana 

yielded energy savings of about 40% during refining of wood chips, a three 

times higher bending strength and more than half reduced water absorption 

and swelling of fiber boards. The topic of biological treatment of chips of was 

reviewed by Messner (1998). 

Despitethemassiveamountofmoneyandworkdevotedtobiopulping, 

a sweeping success seems however vague. The difficulties involved are mainly 

microbiological problems: It is generally difficult to scale-up small-sized laboratory 

experiments with fungal pure cultures via medium-sized rotating fermentors 

with controlled aeration and temperature to the final aim of obtaining 

the same result in chip silos or even in large-sized chip piles under natural outdoor 

conditions. During controlled biopulping, the different white-rot fungi 

may be grown on wood chips for 10 to 15 days. In a wood chip pile, available 

nutrients, humidity, and temperature are, however, favorable to contamination 

by many fungi. Most common are Trichoderma species, of which some 

excrete antibiotics against other fungi. Uneven distribution of the inoculum, 

unsuitable or uneven oxygen and carbon dioxide amounts, unfavorable or uneven 

wood moisture content, and increase of the temperature to 50 ◦ Coreven 

to the incineration point are common problems of large-sized outdoor bioconversions 

in piled substrates. An example with respect to brown-rot fungi 

is the successful laboratory and pilot-scale experiments by Leithoff (1997) to 

bio-leach chromium, copper and other elements from treated waste wood by 

means of Antrodia vaillantii (Chap. 7.4) and the failure of the method using 

larger chip piles under practical conditions. Nevertheless, it has been stated 

that development of the biopulping process has reached the pilot scale as far as 

the use of white-rot fungi for mechanical and sulphite pulping is concerned, has 

already been tested on a commercial scale with Ophiostoma piliferum for craft 

pulping (Messner 1998) and that “biopulping ... is close to mill application” 

(Messner et al. 2003). 

As a “by-product”, the biotechnological attempts of using fungi or their 

enzymes in the pulp and paper industry in processes as biopulping, biobleaching, 

and fiber modification have spurred the understanding of the mechanisms 

of wood decay (Chap. 4). It may however be mentioned that the most often 

investigated fungus with respect to enzyme mechanisms, P. chrysosporium,has 

beside chip piles no relevance for wood, neither for trees nor for constructional 

timber. 

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9.4 

“Palo Podrido” and “Myco-Fodder” 

In the evergreen temperate rainforests of southern Chile, Philippi (1893) found 

in the heartwood of dying and fallen hardwoods (Eucryphia cordifolia, Nothofagus 

spp. and other trees) a white, spongy-wet wood tissue (Fig. 9.4a, also 

Fig. 7.2c), which may occupy the entire interior of logs. This white-rotted 

wood, called “palo podrido” (rotted wood) or “huempe”, develops by the action 

of Ganoderma species like Ganoderma adspersum (Martínez et al. 1991a, 

1991b; Barrasa et al. 1992; Bechtold et al. 1993) and other white-rot Basidiomycetes, 

associated yeasts and bacteria (González et al. 1986), in the moist 

forest climate during a long time. Environmental factors such as a lack of 

desiccation and frost during the year in tropical forests may have reduced 

the mechanical stress on the wood and maintained conditions that promote 

delignification (Eriksson et al. 1990). Low nitrogen content of the wood was 

considered to be a major factor that contributed to this selective delignification 

(Dill and Kraepelin 1986). Black manganese deposits indicating the correlation 

to manganese peroxidase have been found in palo podrido by Barrasa et al. 

(1992) and others. Rodriguez et al. (2003) detected several iron-chelating catechol 

compounds in palo podrido samples, whose relation to lignin or fungal 

metabolites remained however unclear. 

Palo podrido has been used by rural population as feed for foraging cattle. 

Healthywood,eveningrindedform,hasaverylowrumendigestibility.Thus, 

the development of palo podrido by the action of fungi may be termed as 

“biological wood pretreatment”. Due to the fungal delignification particularly 

in the area of the middle lamella/primary walls, the woody tissue is loosened 

and now edible by cattle. Figure 9.4b demonstrates that the Chilean cow prefers 

the pineapple-like palo podrido (Fig. 9.4a) to the surrounding grass. Mainly 

through the opening of the wood structure, now the anaerobic rumen bacteria 

cangetaccesstothedigestiblewoodcarbohydrates.Thereductionofthelignin 

content from 22% of healthyNothofagus wood to about 6%in the corresponding 

palo podrido sample (Dill and Kraepelin 1986) may have promoted bacterial 

activity, but is probably no premier factor, as it has also been stated for the 

bacterial degradation of chemically pretreated wood (Chap. 5.2). The rumen 

bacteria convert the wood carbohydrates in palo podrido to fatty acids like 

acetic, propionic, and butyric acid. This fermentation is the “biotechnological” 

part of palo podrido. Last, the cow uses the fatty acids and also the continually 

dying bacteria to produce meat and milk. 

Lignocelluloses which have been specifically treated with fungi to improve 

the digestibility and protein content for use as ruminant feed have been termed 

as “myco-fodder” (Heltay 1999; also Eriksson et al. 1990). For example, the 

digestibility of straw that was treated with Lentinula edodes for 2 months 

showed increased digestibility by 28% (Zadraˇzil 1985; Zadraˇzil and Brunnert 

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9.5 Wood Saccharification and Sulphite Pulping 247 

Fig.9.4. “Palo podrido” caused by Phlebia chrysocreas (a) andaChileancoweating“palo 

podrido” (b) (photos J. Grinbergs) 

1980). As a by-product, production of edible mushrooms may increase the 

economy of fungal straw treatment. 

9.5 

Wood Saccharification and Sulphite Pulping 

Both wood saccharification with acids and sulphite pulping may be termed 

“chemical wood pretreatment” when the obtained sugars are subsequently 

used for microbial or enzymatic conversions. 

The acid wood saccharification yields monosaccharides from the wood carbohydrates. 

Hydrolysis of lignocelluloses either with diluted or concentrated 

acids has been practiced on a large commercial scale for many years. This technique 

was used in the USA in the 1910s and in Germany and in Switzerland 

during the Second World War. About 10 million m 3 of wood were saccharified 

by acid hydrolysis with up to 48% sugar yield of the possible 70% yield in 

the former Soviet Union around 1983 (Wienhaus and Fischer 1983). The main 

product is glucose, which is the universal sugar for the majority of organisms. 

Glucose can by either converted by yeasts, e.g., Candida utilis, aerobically 

to fodder yeast (single cell protein, SCP; Dart and Betts 1991) or for human 

feed, or glucose is anaerobically fermented to ethanol to be used as chemical 

feedstock or as petrol substitution (Decker and Lindner 1979). Glucose fermentation 

to ethanol was one of the first complex biological processes mastered by 

man and became an important fuel and chemical feedstock in the mid-19th 

century. However, with the rapid growth of the petroleum and petrochemical 

industry following World War I, fermentation has been restricted primarily 

to the brewing and distilling industries (Saddler and Gregg 1998). Ethanol 

can be also obtained from xylose by the xylose-fermenting yeast Pachysolen 

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tannophilus. Other fungi, e.g., molds, as well as aerobic and/or anaerobic bacteria 

can produce amino acids, antibiotics, enzymes, organic acids, solvents, 

and vitamins from glucose or glucose-containing wastes. Technical problems 

of acid hydrolysis, such as corrosion of the reaction vessels and formation of 

noxious by-products, led to research on enzymatic hydrolysis processes, which 

promised, for example, higher sugar yields (Saddler and Gregg 1998). 

Spent sulphite liquors contain at about 50% of the employed wood as lignin 

sulphonic acids and simple sugars from the hemicelluloses. A number of applications 

for lignosulphonates or the entire spent sulphite liquors have been 

developed in the past (e.g., Faix 1992). Since the early past century, the hexoses 

in spent softwood liquors were converted by yeasts to alcohol and the pentoses 

in hardwood liquors to fodder or feeding yeast, respectively. For example, a mill 

in Switzerland produced (in 1980) in two tanks (320 m 3 ) 82,000 hL alcohol and 

7,000 t of yeast cells, respectively. In Sweden, 1.2 million hL of alcohol was produced 

in 33 plants in 1945 (Herrick and Hergert 1977). In the 1980s, the sugars 

in spent sulphite liquor were converted by means of the soft-rot deuteromycete 

Paecilomyces variotii for use as animal feed in Finland (“Pekilo-process”; Forss 

et al. 1986). Han et al. (1976) cultured Aureobasidium pullulans on straw hydrolysate 

for production of single cell protein. Ek and Eriksson (1980) used 

Sporotrichum pulverulentum for water purification and protein production. 

Anaerobic treatment of pulp mill effluents by bacteria was reviewed by Guiot 

and Frigon (1998). 

9.6 

Grinding and Steam Explosion 

Among the physical pretreatment methods, grinding of lignocelluloses increases 

the inner surfaces of the wood cell wall and thus improves the accessibility 

for enzymes to the cell wall components. The particle size must be 

reduced to 50µm to maximize the effect. The energy costs become prohibitive 

at particle sizes of 200µm (Dart and Betts 1991). 

Steam explosion methods saturate the lignocellulose with steam and then 

allow it to undergo explosive decompression. The treatment releases acids 

that contribute to the disruption of the cell wall (Dart and Betts 1991). In the 

steaming-extraction process, chopped wood was treated in watery or alkaline 

solution for a few minutes at 185–190 ◦ C (1,100–1,200 kPa). Subsequent washing 

with water or thin sodium hydroxide solution separated the wood into 

a solid component containing lignin and cellulose and a liquid phase of the 

hemicelluloses (Dietrichs et al. 1978). In vivo digestibility of wood in a test 

cow increased from about 5% of natural wood to 80% for steam-treated wood 

(Puls et al. 1983). The hemicellulose fraction was used to produce Paecilomyces 

variotii mycelium and enzymes (Schmidt et al. 1979). 

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9.7 Recent Biotechnological Processes and Outlook 249 

9.7 

Recent Biotechnological Processes and Outlook 

Several new applications of enzymes have reached, or are approaching, the 

stage of commercial use in the pulp and paper industry. These include e.g., 

enzyme-aided bleaching with xylanases, direct delignification with oxidative 

enzymes, energy-saving refining with cellulases, pitch removal with lipases, 

slime control (Klahre et al. 1996) in the paper machine, removing contaminants 

in the recycle stream, as well as deinking (Kenealy and Jeffries 2003; Messner 

et al. 2003). 

A colorless mutant of the blue-stain fungus Ophiostoma piliferum was used 

to control pitch problems (Blanchette et al. 1992b; Farrell et al. 1993; Brush et al. 

1994; also Fischer et al. 1994), and chip treatment with O. piliferum decreased 

energy consumption and increased strength properties in mechanical pulps 

(Forde Kohler et al. 1997). 

Enzymes used in pulping can increase the yield of fiber, decrease further 

refining energy requirements, or provide specific modifications to the fiber. 

Cellulases, hemicellulases, and pectinases allowed for better delignification of 

the pulp and savings in bleaching chemicals without altering the strength of 

the paper (Kenealy and Jeffries 2003). Laccase and protease reduced energy 

requirements in mechanical pulping. Cellulases and hemicellulases have been 

used in the refining of virgin fibers. Agricultural residues like wheat and rice 

straw have been mechanically pretreated followed by treatment with enzymatic 

cocktails from Lentinula edodes for pulp production (Giovannozzi-Sermanni 

et al. 1997). 

The initial studies on the use of enzymes in bleaching were performed with 

a goal of imitating the wood-decaying action of fungi in nature (Iimori et al. 

1998; Viikari et al. 1998). However, different mixtures of lignin and manganese 

peroxidases did not consistently delignify unbleached craft pulp. The use of 

xylanases in bleaching can improve lignin extraction, alter carbohydrate and 

lignin association, or cleave redeposited xylan. Recently, laccases or manganese 

peroxidases, either alone or combined with low molecular weight mediators, 

have been examined. In the laccase-mediator concept, laccase is combined with 

a low molecular weight redox mediator resulting in generation of a strongly 

oxidizing co-mediator, which then specifically degrades lignin (Jakob et al. 

1999; Sealey et al. 1999). “Novel xylanases” deriving from thermophilic and 

alkaline sources are of importance due to the prevailing conditions in pulp processing. 

Progress in the knowledge of the xylanase-encoding DNA sequences 

and the expression of xylanases in other microorganisms may lead to further 

development in this area (Kenealy and Jeffries 2003). 

Waste paper is the primary raw material of the European paper industry. 

For Germany, the amount of waste paper for paper production has been 

forecasted to about 14 million t in 2005. In 1995, the average composition of 

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waste paper in a deinking plant consisted of 41% newspapers, 39% magazines, 

9% wood-free paper, 6% unusable paper and board, 4% other bright papers, 

and 1% non-paper components (Hager 2003). More than 70% of mixed office 

waste paper consists of uncoated papers that are printed with copy or 

laser printer toners, which may be difficult to remove by conventional, alkaline 

deinking (Kenealy and Jeffries 2003). Fibers may be treated by hydrolyzing 

enzymes to remove print (deinking). Cellulases are particularly effective in 

the removal of toners from office waste papers. It was concluded that the primary 

role of cellulases in deinking involves separating ink-fiber agglomerates 

and dislodging or separating ink particles and fibrous material in response 

to mechanical action during disintegration (Kenealy and Jeffries 2003). Few 

experiments have used oxidative enzymes for deinking. The missing potential 

for the reduction of specks that derive from residual ink and the observed 

lignin modification rendered laccase either alone or combined with the mediator 

1-hydroybenzotriazole unsuitable for practical ink elimination of wood 

containing waste paper (Hager et al. 2002). Recycled paper sludge generated 

during repulping was simultaneously hydrolyzed with fungal cellulase and fermented 

with the yeast Kluyveromyces marxianus to convert cellulose fibers to 

ethanol (Lark et al. 1997). 

Papers made from secondary fibers often show a higher microbial load 

which is of disadvantage for some applications, e.g., as hygienic papers (Cerny 

and Betz 1999). 

Anaerobic treatment of pulp mill effluents was reviewed by Guiot and Frigon 

(1998).PhanerochaetechrysosporiumandTrametes versicolor havebeenusedto 

degrade the chlorolignins in the effluents produced during chlorine bleaching 

(Eriksson et al. 1990). The ligninolytic systems of white-rot fungi, particularly 

P. chrysosporium, was used to degrade several persistent environmental 

pollutants such as benzo(a)pyrene, DDT, and dioxin. Bacteria metabolized 

dibenzo-p-dioxin (Wittich et al. 1992). Aerobic bioremediation techniques 

for the cleanup of creosote and PCP-contaminated soils were reviewed by 

Borazjani and Diehl (1998) (also Prewitt et al. 2003). Aerobic PCP transformation 

initially produced small amounts of pentachloroanisole; however more 

than 75% of both chemicals disappeared in 30 days from the test soil. Under 

methanogenic conditions, PCP was reductively dechlorinated to tetra-, tri-, 

and dichlorophenols (D’Angelo and Reddy 2000). Bioremediation of wood 

treated with preservatives using white-rot fungi was treated by Majcherczyk 

and Hüttermann (1998). The peroxidases of white-rot fungi unspecifically oxidize 

aromatic compounds by generating such a high redox-potential that they 

“burn down” all available aromatics present in the proximity of the mycelia. 

Phanerochaete laevis transformed polycyclic aromatic hydrocarbons (Bogan 

and Lamar 1996). Tubular bio-filters filled with straw, which were previously 

colonized with Pleurotus ostreatus mycelium was used to filter out ammonia 

from the waste air of cattle sheds (Majcherczyk et al. 1990). Experiments on the 

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9.7 Recent Biotechnological Processes and Outlook 251 

biodegradability of coal tar oil (creosote) by 16 bacterial species and six fungi 

using IR-spectra as an indicator for attack have been unsuccessful, assumably 

due to the complex mixture of some hundred toxic compounds in the tar oil 

(Schmidt et al. 1991). 

Bark extracts of Acacia spp. (wattle or mimosa bark extract) and wood extracts 

of Schinopsis spp. (quebracho wood extract) are rich sources of tannins. 

Tannins are used for a long time in leather tanning and for the production of 

adhesives (Pizzi 2000; Roffael et al. 2002). Copper tannate was tested as a possible 

wood preservative (Pizzi 1998). In 1950, worldwide about 300,000 t of 

tannin extracts were produced (Herrick and Hergert 1977). Main commercial 

producers are Argentina, South Africa, Brazil, Paraguay, Zimbabwe, Indonesia, 

Kenya, and Chile. The hot water extract of spruce and larch bark contains a high 

amount of carbohydrates and was thus unsuitable for adhesives. The soft-rot 

fungus Paecilomyces variotii reduced the carbohydrate content, so that the tannins 

were suitable as adhesives (Schmidt et al. 1984; Schmidt and Weißmann 

1986). Wagenführ (1989) used a commercial pectinolytic enzyme preparation 

to reduce the carbohydrate content. 

Despite the massive amount of money and effort devoted over the past 

decades to the microbiological or enzymatic conversions/treatments of lignocelluloses, 

several of the projects started with enthusiasm have suffered success 

or practical utilization or even loss of interest. Oil has remained the premier 

raw material for chemicals of all types (Little 1991). However, the foreseeable 

limitation of oil resources and thus the probable increase in the cost of 

petroleum-derived feedstock will provide the necessary incentive to further 

research. But, the development and utilization of alternative processes also 

depend on political interests and geographical aspects. As an example for the 

latter, the use of biofuels (rapeseed oil methylester, RME) may be a possible 

substitute for fossil fuels, which also contribute substantially to the increase in 

CO2 in the atmosphere. In Germany, the share of RME on the whole consumption 

of diesel fuel however is 4.3% and cannot exceed 7% due to the limited 

arable acreage. In the end, economy and subsidization will decide on future 

research. 

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Appendix 1 

Identification Key for Strand-Forming House-Rot Fungi 

(According to Huckfeldt and Schmidt 2004) 

All key points must be considered before a decision. Numbers in parentheses refer to 

the preceding key point. The question mark points out that only few samples have been 

investigated. 

1 fungus causes (intensive) rot 2 

1∗ fungus does not cause (intensive) rot – possible error: rotten wood is 

overgrown or infection is in initial stage; no vessel hyphae (if vessels then 

usually within strands, forward to 3) 

34 

2(1) brown-cubicalrot;nosetae;sporesalwayseven(alsoinoil-immersion) 3 

2∗ white rot; vessels always less than 15 µm in diameter 24 

3(2) strands clearly recognizable, but often overgrown by mycelium 4 

3∗ strands indistinct (microscopic investigation necessary; start at (4) if 

vessels are present) 

16 

4(3) strands over 5 mm in diameter, removable from the substrate, frequently 

surrounded by thick mycelium or hidden in masonry, wood etc.; dry 

strands break with clearly audiblecracking; fiber (skeletal) hyphae refractive; 

vessels with internal wall thickenings (bars), to 60 µm indiameter; 

vegetative hyphae with clamps see (12) Serpula lacrymans 

4∗ strands under 5 mm in diameter or firmly attached to the substrate 5 

5(4,16) strands hair-like, often branched and clearly defined (with “bark”), below 

0.5mm in diameter and often below mycelium, removable; no fibers; or 

strands/mycelium with sclerotia 

6 

5∗ strands not hair-like, not clearly defined (without bark); no sclerotia; 

fibers present or absent 

7 

6(5) sclerotia large, to 6 mm in diameter, round, often somewhat irregular, 

sometimes absent; strands hair-like, with bark, cream to yellow, redbrown 

to black when old, under 0.5 mm in diameter, somewhat flexible 

when dry; no fibers; vessels to 25 µm in diameter, numerous, in groups, 

withbars,cellwallto1µm thick; some vegetative hyphae bubble-like 

swollen to 10–25 µm in diameter and according literature with medallion 

clamps, always with clamps; strands also in masonry; only on softwoods 

Leucogyrophana mollusca 

6∗ sclerotia small and oblong, to 2.5 mm long, brown to grey, sometimes 

absent; strands hair-like, with bark, yellowish, grey to brown, probably 

darker when old, covered by lighter mycelium or exposed, under 0.5 mm 

in diameter, somewhat flexible when dry; no fibers; vessels to 25 µm in 

diameter, but often partly thickened, numerous, in bundles, with bars; 

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254 Appendix 1 

vegetative hyphae with clamps, 2.5–4.5 µm in diameter; strands also in 

masonry; probably only on softwoods Leucogyrophana pinastri 

7(5) strands with vessels (sometimes rare; search; wood fibers may be mistaken 

for vessels) 

8 

7∗ strands without vessels 9 

8(7) strands without fibers, however usually with vessels and vegetative hyphae 

with clamps 

10 

8∗ strands with fibers, vessels and vegetative hyphae with clamps 11 

9(7) strands with fibers and vegetative hyphae 16 

9∗ juvenile strands only with vegetative hyphae (old strands sometimes with 

vessels and fibers) 

22 

10(8) vessels rare, often narrowed at the septa; fibers absent or indistinct; veg- 15 

etative hyphae with clamps 

10∗ vessels numerous, to 21 µm in diameter, often in bundles, with septa, bars 

indistinct or absent, cell wall to 1 µm thick; no fibers; vegetative hyphae 

with clamps, 1–4µm in diameter, small hyphae partly with thickened 

cell wall; strands indistinct, just as embedded as those of S. lacrymans, 

somewhat flexible when dry, white, cream-yellow to grey, always brittle, 

to 2 mm in diameter, also in masonry Leucogyrophana pulverulenta 

11(8) vessels with bars (sometimes absent in very young strands), to 60 µm 

in diameter, fibers straight-lined, not flexible (with aqueous or ethanol 

preparation, may be flexible in KOH) 

11∗ vessels without bars, but with clearly defined septa, rarely over 30 µmin 

diameter; mycelium not silver grey (if molds absent), fibers flexible or not 

12(11) fibers refractive, (2–) 3–5 (–6.5) µm in diameter, fibers within strands 

near fruit body to 12 µm in diameter, straight-lined, septa not visible, 

no clamps, thick-walled, lumina often visible; vessels at least partly numerous 

(in groups), 5–60 µm indiameter,notorrarelybranched;with 

bars, these up to 13 µm high; vegetative hyphae hyaline, partly yellowish, 

brown when old, with large clamps, 2–4 µm in diameter, near fruit body 

to 4 µm in diameter; strands white, silver-grey, greytobrown,to3cm 

wide, usually with flabby mycelium in between, dry strands breaking 

with clearly audible cracking (strands contaminated with molds often 

not cracking any more); aerial mycelium cotton-woolly, soft, white, lightgrey 

to silver-grey, with yellow, orange or violet spots (“inhibition color”), 

often several square meters on walls, ceilings and floors, in the draught 

collapsing fast; on hardwoods and softwoods; strands often in masonry; 

(S. himantioides can be excluded, if strands thicker than 2 mm and at least 

some fibers more than 4.5 µmindiameter) Serpula lacrymans 

12 ∗ see before, but fibers (1.5−) 2–3.5 (−4) µm (sometimes not clearly distinguishable 

from S. lacrymans); strands to 2 mm in diameter, root-like 

branched and not as surrounded by thick mycelium as S. lacrymans;fruit 

body to 2 mm thick Serpula himantioides 

13(11) vegetative hyphae partly swelling up to 5–10 (−20) µm, fibers up to 2.5 

(−3) µm, vessels up to 40 µm; mycelium white, sometimes going yellow 

(if vessels swelling up: see 15) 

13 ∗ vegetative hyphae not swelling, ± regular diameter, at septa sometimes 

smaller; strands and aerial mycelium predominantly consisting of fibers, 

12 

13 

22 

14 

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Appendix 1 255 

thesebrighttobrown;vesselssolitary;vegetativehyphaepresent,however 

partly rare (search) 

14(13) fibers light to dark-brown, flexible or not, older strands not snow-white; 

on hardwoods and softwoods 

14 ∗ fibers hyaline or pale yellow, flexible; strands whitish to cream, partly 

somewhat yellowing or rarely infected by molds, also ice flower-like, 

flexible when dry, up to 7 mm in diameter; fibers numerous, 2–4µm 

in diameter (in Antrodia xantha partly somewhat yellowish, hyphal tips 

with tapering ending cell walls), narrow lumina, straight-lined, mostly 

unbranched, insoluble in 3% KOH, [if dissolving, see Diplomitoporus 

lindbladii (31), check rot type, if fibers missing], but in KOH swelling, 

sometimes with ‘blown up’ hyphal segments; vessels not rare but in old 

strands difficult to isolate, up to 25 µm in diameter, thick-walled with 

middle lumen, without bars; vegetative hyphae with few clamps, 2–4 (–7) 

µm in diameter, sometimes medallion clamps, often somewhat thickwalled; 

surface mycelium white to cream, thin, aerial mycelium in nodraught 

or under-floor areas partly some square meters large, white 

to cream, later also stalactite-like growth from above; strands also in 

masonry(?);probablyonlyonsoftwoods;genusAntrodia (species not 

surely distinguishable on the basis of their strands/mycelia) 

Antrodia vaillantii, A. sinuosa, A. xantha, A. serialis 

15(10,14) vegetative hyphae with clamps; strands first cream to loam-yellow, then 

brownish to ochre, up to 3 mm wide, root-like branches, similar to those 

of Coniophora puteana, however not becoming black; surface mycelium 

first dirty-white to yellowish, then loam-yellow, brownish to ochre, near 

fruit body partly violet; vegetative hyphae refractive, (1.5−) 2.5–3–5 (−5) 

µm in diameter, partly thickened; fibers indistinct, 1.5–5 µmindiameter 

(often only in darker strands); vessels hyaline, sometimes with ‘blown up’ 

hyphal segments, up to 15 (−25) µm in diameter, without bars, but with 

septa, with clamps; on and within (?) masonry and wood, often in damp 

cellars; brown rot Paxillus panuoides 

15 ∗ vegetative hyphae without or rarely with clamps, rarely multiple clamps 

(more often at margin of fruit body, often indistinct, since branched), 

2–6 (–9) µm indiameter;strandsfirstbright,thenbrowntoblack,up 

to 2 mm wide, to 1 mm thick, root-like, hardly removable (not so with C. 

marmorata), when removed usually fragile, partly with brighter center, 

underlying wood becoming partly black; fibers pale to dark brown, 2–4 

(−5) µm in diameter, somewhat thick-walled, however with relatively 

broad, usually visible lumen, also branched, to be confused with vegetative 

hyphae; drop-shaped, hyaline to brownish secretions (1–5 µm in 

diameter) often to be found on hyphae; vessels in strands surrounded and 

interwoven by fine hyphae (0.5–1.5 µm in diameter), therefore preparation 

with H2SO4 and KOH solution, due to preparation irregularly formed 

or distorted, up to 30 µm in diameter, thin-walled (or slightly thick-walled 

with C. marmorata), without bars, but with septa; often also in masonry 

etc., genus Coniophora (species not surely distinguishable on the basis of 

their strands/mycelia) e.g., Coniophora puteana, C. marmorata 

16(3,9) mycelium on masonry, concrete etc.; vessels possibly not visible or missing, 

untypical or small; if star-shaped setae present see (25) 

15 

5 

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16∗ mycelium not on or in masonry 17 

17(16) fibers present; vessels absent; vegetative hyphae with clamps (in older 

parts rare); mycelium and strands only on wood 

18 

17∗ fibers missing or very rare; vegetative hyphae present (see (5) if vessels 

present, search for vessels, being rare in young strands) 

22 

18(17) fibers partly with swelling and partly with regular diameter, 2.5–4.5 µm 

in diameter (in fruit body sometimes larger), flexible,luminasmall,often 

visible, sometimes punctually larger; vegetative hyphae thin-walled, 1–2 

(−2.5) µm in diameter, with clamps, but no medallions; cystidia possible; 

myceliumcreamtocorky,firmandtough,oftenincavitiesandshakesin 

wood below fruit body; on oak, half-timbering; brown rot 

Daedalea quercina 

18∗ fibers not swelling, ± regular in diameter; hyphae in wood usually possess 

clamps of medallion type 

19 

19(18) mycelium rough-velvet; usually two-layered, at least two-colored: white 

mycelium close to wood and covered by yellow, reddish to brown aerial 

mycelium; fibers 1.5–5 µm in diameter, discolored at darker mycelial areas; 

vegetative hyphae with clamps; grey mycelia cannot be differentiated; 

often at windows 

20 

19∗ mycelium fine-velvet to silky; not distinctly two-layered; fibers 1.5–2 

(−2.5) µm in diameter, hyaline, straight, rarely branched; vegetative hyphae 

always with clamps, 1.5–2 µm in diameter; if hyphae wider see (14); 

mycelium firm and tough, first white, then with yellow, ochre to violet 

spots; covering cavities and shakes in wood, easy to remove; mycelia and 

strands so far only proven for wood; monstrous “dark fruit bodies”, sometimes 

with little caps; usually on softwoods; brown rot; genus Lentinus 

(species not surely distinguishable on the basis of their strands/mycelia) 

e.g., Lentinus lepideus 

20(19) fibers up to dark-brown (examine dark areas); aerial mycelium cream, 

ochre to dark-brown, underneath white to cream mycelium (not always 

clearly visible, use pocket-lens); colored fibers 1.5–3 (–4.5) µm in diameter; 

vegetative hyphae 2–4.5 µm in diameter, with clamps; strands rare, 

then forming structures of a few centimeters, these first bright, reddish, 

then red-brown to grey; in cavities dark, monstrous tap-, pin-, antlers- or 

cloud-like “dark fruit bodies”; only on softwoods 

Gloeophyllum abietinum 

20∗ colored fibers and surface mycelia not so dark, 2–5 µm indiameter; 

sometimes also “dark fruit bodies” 

21 

21(20) mycelium white, cream to light brown; rarely short strands of few centimeters 

of length, these first bright, then yellowish to ochre-brown and usually 

covered by mycelium; colored fibers light to dark yellow, light-brown to 

brown, 2–4.5 µm in diameter (partly broader); vegetative hyphae hyaline, 

2–4µm in diameter, with clamps; arthrospores rare, 3−4 × 10−15 µm, 

cylindrical; often in shakes; tap-, pin-, antlers- or cloud-like “dark fruit 

bodies”; only on softwoods Gloeophyllum sepiarium 

21∗ mycelium white, beige, yellow-orange to light grey-brown; strands under 

1 mm in diameter and not clearly defined; surface mycelium whiteyellow 

to grey and usually covered by mycelium; colored fibers very light 

yellow, gold-yellow to light-brown, 1–4 µm in diameter, septa clearly 

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22(13,17) 

recognizable; vegetative hyphae hyaline, 2–4 µm in diameter, thin-walled, 

with clamps; ‘dark fruit bodies’ also antlers-shaped, often with brighter 

tips; often in shakes; only on wood (softwoods and hardwoods) 

Gloeophyllum trabeum 

vegetative hyphae without clamps 23 

22∗ vegetative hyphae with clamps, partly with swellings, 1–2 µmindiameter; 

fibers and vessels only in older strands; fibers 0.5–2 (−3) µm, hyaline, 

straight-lined, thick-walled, septa not visible, no clamps, no reaction in 

KOH; vessels 6–40 µm in diameter, thin-walled or slightly thick-walled, 

hyaline, vessels in strands surrounded and interwoven by fine hyphae 

(0.5–1.5 µm in diameter); mycelium pure white or pink, if being undisturbed 

lasting so, easily removable, but sensitive; strands often sunk in 

mycelium; on softwoods, rarely on hardwoods; brown rot; genus Oligoporus 

and similar fungi (species indistinguishable by strands/mycelia) 

e.g., Oligoporus placenta 

23(22) arthrospores thin-walled, cylindrical 1.5−2.5 × 5−12 µm; vegetative hyphae 

hyaline, thin-walled or slightly thick-walled, 2–3 (−4) µmindiameter, 

without clamps, but with primordial clamps; vessels indifferent, septa 

present, thin-walled, to 12 µm in diameter; in older parts sometimes small 

fibers(comparewith12);myceliumwhitetoyellow,easilyremovable,but 

sensitive; strands often sunk in mycelium 

monokaryon of Serpula lacrymans 

23∗ arthrospores absent or different 34 

24(2) setae present, simple setae or stellar setae, within white to cream 25 

mycelium, partly only very small nests of setae (search) 

24 ∗ setae absent 28 

25(24) stellar setae present; vegetative hyphae without clamps 27 

25 ∗ setae not clearly stellar-shaped or simply branched, partly rooted 26 

26(25) simple, dark-brown, to 180 µm long setae within mycelium, strand and 

fruit body; fibers pale yellow, thin-walled, 2–3 µm in diameter, rarely 

branched; vegetative hyphae hyaline, 1.5 µm in diameter; mycelium 

downy, loam-yellow to brown, also white when young; strand-like structures 

up to 4 mm wide and 0.5 mm thick, firmly attached, often fingershaped 

branched; usually on hardwoods (often on framework), very rare 

on softwoods; so far proven for oak, ash, false acacia, elm, beech, fir and 

spruce; white rot Phellinus contiguus 

26 ∗ simple, dark-brown setae in fruit bodies and mycelium, under 100 µm; 

other species of the genus Phellinus known to occur in buildings (species 

not surely distinguishable on the basis of strands/mycelia) 

Phellinus nigrolimitatus, P. pini, P. robustus 

27(25) stellar setae dichotomously branched, to 90 µm in diameter, in fruit body, 

mycelium and strand, partly rare; vegetative hyphae with septa, 2–4 µmin 

diameter; strands cream to red-brown, fibrous surface; partly embedded 

in white mycelium or fruit body; spores subglobose, smooth; strands on 

and in masonry; white rot Asterostroma laxum 

27 ∗ stellar setae only rarely branched, up to 190 µm in diameter, in fruit body, 

mycelium and strand; vegetative hyphae with septa, 1.5–3 µm indiameter; 

strands cream-brown, up to 1 mm in diameter; surface mycelium 

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first white, then brown, partly small mycelial plugs; spores subglobose, 

tuberculate; strands on and in masonry; white rot 

Asterostroma cervicolor 

28(24) fibers absent (also in aqueous preparation); sometimes strands with vessels 

28∗ fibers present; strands without vessels; mycelium sometimes with vessel- 

like hyphae 

29(28) crystalline asterocystidia in fruit body and strand, up to 20 µm indiameter, 

cystidia stipe to 11 µmlong,2µmindiameter; vegetative hyphae 

with clamps, 1.5–3 µm; vessels thin-walled, to 15 µmindiameter,without 

bars, but with septa; no fibers; strands snow-white to cream, 0.2–1 mm 

in diameter (?), mostly short und near fruit body; fruit body smooth; to 

date only found on softwood; white rot Resinicium bicolor 

29∗ without asterocystidia 30 

30(29) vegetative hyphae with clamps, partly with bubble-like swellings, 1–2 

(−4) µm; no fibers (if fibers present, see 31); sometimes with vessels (then 

no swellings), small clamps, 4–9 (?) µm in diameter; strands snow-white 

to cream, 0.2–1(?) mm in diameter, fragile, often only short and near 

fruit body; fruit body resupinate, thin, poroid, grandinioid or smooth, 

fragile; spores warty, translucent and small, 4−5.5×3−4.5 µm; so far only 

found directly on damp wood; white rot; genus Trechispora (species not 

distinguishable on the basis of strands/mycelia); 

in buildings Trechispora farinacea, T. mollusca 

30∗ other characteristics 34 

31(28) fibers insoluble in 3% KOH, sometimes slightly swelling, partly under 32 

3 µm in diameter; mycelium partly with brown crust 

31 ∗ fibers completely soluble in 3% KOH, 2–4.5 (−8?) µm in diameter, thickwalled 

to solid (‘filled’), similar to A. vaillantii (14); no vessels; vegetative 

hyphae with few clamps, 1–2.5 µm indiameter;surfacemycelium 

without crust, usually meager, partly forming compact plates, white to 

light-brown; strands white, partly somewhat yellowing, root-like, richly 

branched, radiate or ice flower-like, fibrous, up to 2 mm in diameter; so 

far mycelium only proven to occur on wood; white rot 

Diplomitoporus lindbladii 

32(31) arthrospores often lemon-shaped, hyaline, thick-walled, 5−7×7−12(−?) 

µm, in surface mycelium, which lies close to the wood, and in substrate 

mycelium; in white mycelium: fibers hyaline to brown, to 2 µmindiameter, 

not very thick-walled and hardly separable from vegetative hyphae; vegetative 

hyphae hyaline with clamps, these often difficult to find, 1–2 µmin 

diameter; vessels not proven; in colored mycelium: fibers light-brown to 

brown, 1.5–3 (−4.5) µm in diameter; vegetative hyphae hyaline to brown, 

thick-walled, rarely clamps, 2–6 (−7)µm in diameter, branched; vessels to 

11 µm in diameter; strands usually absent or short and under mycelium; 

mycelium first white to cream, then yellowish, grey to brown, when old 

often luxuriant, firm and tough, frequently with paper-like, firm, brown 

crust, predominantly in shakes and cavities, usually with amber guttation 

drops or with brown to black spots (remainders of dried guttation), 

in constructions white to cream; surface mycelium partly with distinct 

29 

31 

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margin; sometimes with poroid fruit bodies within surface mycelium 

(1–90 mm thick), then also wider hyphae; white rot, preferential sapwood 

decay,hardwoodandsoftwood,nooronlysomegrowthonmasonry 

Donkioporia expansa 

32 ∗ arthrospores, strands or mycelia different 33 

33(32) strands black, very clear, with separate crust layer, often also hollow 

when old (32), clearly thicker than 1 mm, only on wood with bark rests 

or in wood in the area of in-growing roots, examine for in-growing roots; 

hardwood and softwood; white rot rhizomorphs of Armillaria spp. 

33 ∗ strands or mycelia different 34 

34(1,23, 

30,33) 

on masonry, rough-casting etc.; no or slightwood decay: e.g., species of the 

genera Coprinus, Peziza (white strands), Scutellinia, Pyronema, molds 

(e.g., Cladosporium) and slime fungi (Enteridium, Fuligo, Trichia) 

34 ∗ further species on wood which so far were rarely found in buildings: e.g., 

species of the genera Daldinia, Fomitopsis, Hyphodontia, Phanerochaete, 

Phlebiopsis, Pleurotus, Polygaster, Trametes; see also Table 8.6 

www.taq.ir

Appendix 2 

Fungi Mentioned in this Book 

(see also Tables 8.1–8.3; most English names according to Larsen and Rentmeester 1992, 

Rune and Koch 1992, some names suggested as new by the author) 

Scientific name, English name Significance in this book 

Agaricus bisporus (J.E. Lange) Pilát agaric mushroom 

Agrocybe aegerita (Brig.) Singer edible mushroom on wood 

Alternaria alternata (Fr.) Keissl. toxic mold, blue stain 

Amanita caesarea (Scop.: Fr.) Pers. mycorrhizete 

Amanita muscaria (L.: Fr.) Hook. mycorrhizete 

Amylostereum areolatum (Chaill.: Fr.) Boid. red streaking 

Amylostereum chailletii (Pers.: Fr.) Boid. red streaking 

Antrodia serialis (Fr.: Fr.) Donk, Effused tramete indoor wood 

Antrodia sinuosa (Fr.: Fr.) P. Karsten, White polypore indoor wood 

Antrodia vaillantii (DC: Fr.) Ryv., Mine polypore indoor wood 

Antrodia xantha (Fr.: Fr.) Ryv., Yellow polypore indoor wood 

Armillaria borealis Marxm. & K. Korh., Nordic honey fungus tree parasite 

Armillaria cepistipes Velen. tree parasite 

Armillaria gallica Marxm. & Romagn. tree parasite 

Armillaria luteobubalina Watling & Kile parasite 

Armillaria mellea (Vahl: Fr.) Kummer, Honey fungus tree parasite 

Armillaria ostoyae (Romagn.) Herink, Dark honey fungus tree parasite 

Arthrographis cuboides (Sacc. & Ellis) Sigler pink stain 

Aspergillus flavus Link cancerogenic mold 

Aspergillus fumigatus Fres. cancerogenic indoor mold 

Aspergillus niger van Tieghem, Black mold mold 

Aspergillus versicolor (Vuill.) Tiraboschi mold on poplar wood, 

indoor mold 

Asterostroma cervicolor (Berk. & Curtis) Massee indoor wood 

Asterostroma laxum Bres. indoor wood 

Aureobasidium pullulans (de Bary) Arn. blue stain 

Auricularia auricula-judae (Fr.) Quélet edible mushroom on wood 

Auricularia polytricha (Mont.) Sacc. protoplasts 

Bispora monilioides Corda black streaking of beech 

logs 

Bjerkandera adusta (Willd:Fr.)P.Karsten,Smokeypolyporetreerot Boletus edulis Bull.: Fr. mycorrhizete 

Botrytis cinerea Pers. noble rot of wines, seedling 

shoot tip disease 

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Candida utilis (Henneberger) Lodder & Kreger yeast, glucose conversion 

Cantharellus cibarius Fr. mycorrhizete 

Ceratocystis adiposa (E.J. Butler) C. Moreau blue stain 

Ceratocystis coerulescens (Münch.) B.K. Bakshi blue stain 

Ceratocystis fagacearum (Bretz) Hunt Oak wilt disease 

Ceratocystis fimbriata (Ellis & Halstead) Davidson f. platani 

Walter 

Plane canker stain disease 

Ceratocystis minor (Hedgc.) J. Hunt blue stain 

Ceratocystis pluriannulata (Hedgc.) C. Moreau blue stain 

Cerinomyces pallidus Martin indoor wood 

Ceriporiopsis subvermispora (Pilát) Gilb. & Ryv. lignin degradation, 

biopulping 

Cerocorticium confluens (Fr.: Fr.) Jül. & Stalp. indoor wood 

Chaetomium globosum Kunze: Fr. soft rot 

Chlorociboria aeruginascens (Nyl.) Kan. Small-spored green 

wood-cup 

‘green rot’ 

Chlorociboria aeruginosa (Pers.: Fr.) Seaver (large-spored) ‘green rot’ 

Chondrostereum purpureum (Pers.: Fr.) Pouzar, Silver-leaf tree rot, stored and exterior 

fungus 

wood 

Ciboria batschiana (Zopf) Buchwald acorn rot 

Cladosporium cladosporioides (Fres.) de Vries blue stain 

Cladosporium herbarum (Pers.) Link mold, blue-stain 

Cladosporium sphaerospermum Penz. blue stain, indoor mold 

Climacocystris borealis (Fr.)Kotl&Pouzar treerot 

Coniophora arida (Fr). P. Karsten, Arid cellar fungus indoor wood 

Coniophora marmorata Desm., Marmoreus cellar fungus indoor wood 

Coniophora olivacea (Fr.) P. Karsten, Olive cellar fungus indoor wood 

Coniophora puteana (Schum.: Fr.) P. Karsten, (Brown) Cellar 

fungus 

indoor wood 

Coprinus comatus (O.F. Müller: Fr.) S.F. Gray light influence 

Cryphonectria parasitica (Murr.) Barr Chestnut blight 

Cryptostroma corticale (Ellis & Everh.) P.H. Greg & S. Waller mold, woodworker’s lung 

Cylindrocarpon destructans (Zins.) Scholten oak root parasite 

Dacrymyces stillatus Nees: Fr., Orange jelly indoor wood 

Daedalea quercina (L.: Fr.) Fr., Maze-gill stored and exterior wood 

Daedaleopsis confragosa (Bolton: Fr.) J. Schröter white rot 

Dichomitus squalens (P. Karsten) D.A. Reid successive white rot, 

biopulping 

Diplomitoporus lindbladii (Berk.) Gilb. & Ryv. indoor white wood 

Discula pinicola (Naum.) Petrak blue stain 

Donkioporia expansa (Desm.) Kotl. & Pouzar, Oak polypore indoor white-wood 

Emericella nidulans (Eidam) Vuill. toxic mold 

Earliella scrabosa Gilb. & Ryv. mine timber 

Fistulina hepatica (Schaeffer: Fr.) Fr., Beef-steak fungus tree rot 

Flammulina velutipes (Curtis: Fr.) Singer edible mushroom on wood 

Fomes fomentarius (L.: Fr.) Kickx, Tinder fungus tree rot 

Fomitopsis palustris (Berk. & M.A. Curtis) Gilb. & Ryv. cellulose degradation 

Fomitopsis pinicola (Swartz: Fr.) P. Karsten, Red-belted 

polypore 

brown rot 

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Fuligo septica Gmelin indoor wood 

Fusarium oxysporum (Schlecht.: Fr.) ssp. cannabis herbicide, oak root parasite 

Ganoderma adspersum (Schulzer) Donk “palo podrido” 

Ganoderma applanatum (Pers.) Pat. white rot 

Ganoderma lipsiense (Batsch) G.F. Atk., Artist’s conk white rot 

Ganoderma lucidum (Curtis: Fr.) P. Karsten medicinal mushroom 

Gliocladium roseum Bainier antagonism 

Gloeophyllum abietinum (Bull.: Fr.) P. Karsten, Fir gill 

polypore 

stored and exterior wood 

Gloeophyllum sepiarium (Wulfen: Fr.) P. Karsten, Yellow-red 

gill polypore 

stored and exterior wood 

Gloeophyllum trabeum (Pers.: Fr.) Murr., Timber gill stored and exterior wood 

polypore 

Grifola frondosa (Dicks.: Fr.) S.F. Gray white rot, edible mushroom 

Hebeloma cylindrosporum Romagn. mycorrhizete 

Hebeloma velutipes Bruchet mycorrhizete 

Helicobasidium brebissonii (Desm.) Donk seedling smothering 

Hericium erinaceus (Bull.: Fr.) Pers. edible mushroom on wood 

Heterobasidion abietinum Niemelä & Korhonen, Fir root 

rot fungus 

tree parasite 

Heterobasidion annosum (Fr.: Fr.) Bref. s.s., Pine root 

rot fungus 

tree parasite 

Heterobasidion parviporum Niemelä & Korhonen, 

tree parasite 

Spruce root rot fungus 

Hormonema dematioides Melin & Nannf. blue stain 

Hyphoderma praetermissum (P. Karsten) J. Eriksson & Strid indoor wood 

Hyphodontia spathulata (Schrader) Parm. indoor wood 

Inonotus dryadeus (Pers.: Fr.) Murrill white rot 

Inonotus dryophilus (Berk.) Murrill successive white rot 

Inonotus hispidus (Bull.: Fr.) P. Karsten white rot 

Kluyveromyces marxianus (E.C. Hansen) Van der Walt yeast, ethanol production 

Kretzschmaria deusta (Hoffman) P.M.D. Martin white-rot ascomycete 

Kuehneromyces mutabilis (Schaeff.: Fr.) Singer & A.H. Sm. edible mushroom on wood 

Laccaria bicolor (Maire) Orton mycorrhizete 

Laetiporus sulphureus (Bull.: Fr.) Murrill, Sulphur polypore tree rot 

Laurelia taxodii (Lentz & H.H. McKay) Pouzar brown pocket rot 

Lecythophora hoffmannii (van Beyma) W. Gams soft rot 

Lecythophora mutabilis (J.F.H. Beyma) W. Gams & McGinnes soft rot 

Lentinula edodes (Berk.) Pegler, Shii-take edible mushroom on wood 

Lentinus lepideus (Fr.: Fr.) Fr., Scaly Lentinus stored and exterior wood 

Leucogyrophana mollusca (Fr.: Fr.) Pouzar, Soft dry 

rot fungus 

indoor wood 

Leucogyrophana pinastri (Fr.: Fr.) Ginns & Weresub, 

Mine dry rot fungus 

indoor wood 

Leucogyrophana pulverulenta (Sow.: Fr.) Ginns, Small indoor wood 

dry rot fungus 

Loweporus lividus (Kalchbr.: Cooke) J.E. Wright mine timber 

Macrophomina phaseolina (Tassi) Goid. conifer seedling parasite 

Melanomma sanguinarum (P. Karsten) Sacc. red spotting of beech wood 

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Memnoniella echinata (Rivolta) Galloway toxic mold 

Meria laricis Vuill. Meria needle-cast of larch 

Meripilus giganteus (Pers.: Fr.) P. Karsten, Giant polypore tree rot 

Meruliporia incrassata (Berk. & Curtis) Murr., American indoor wood 

dry rot fungus 

Merulius tremellosus Schrader successive white rot, 

biopulping 

Monodictys putredinis (Wallr.) Hughes soft rot 

Nectria coccinea var. faginata Lohmann, Watson & Ayers Beech bark disease 

Nectria galligena Bres. Beech bark disease 

Nematoloma frowardii (Speg.) E. Horak lignin degradation 

Oligoporus amarus (Hedgc.) Gilb. & Ryv. brown pocket rot 

Oligoporus placenta (Fr.) Gilb. & Ryv., (Reddish) 

indoor wood 

Sap polypore 

Oligoporus stipticus (Pers.: Fr.) Kotl. & Pouzar brown rot 

Ophiostoma novo-ulmi Brasier Dutch elm disease 

Ophiostoma piceae (Münch) H. and P. Sydow blue stain 

Ophiostoma piliferum (Fr.) H. and P. Sydow blue stain 

Ophiostoma setosum Uzunovic, Seifert, S.H. Kim & Breuil blue stain 

Ophiostoma ulmi (Buisman) Nannf. Dutch elm disease 

Oudemansiella mucida (Schrad.) Höhn competition 

Pachysolen tannophilus Boidin & Adzet yeast, xylose fermentation 

Paecilomyces variotii Bain. soft rot 

Paxillus involutus (Batsch: Fr.) Fr. mycorrhizete 

Paxillus panuoides (Fr.: Fr.) Fr., Stalkless Paxillus stored and exterior wood 

Penicillium aurantiogriseum Dierckx indoor mold 

Penicillium camemberti Thom cheese mold 

Penicillium brevicompactum Dierckx indoor mold 

Penicillium chrysogenum Thom indoor mold 

Penicillium glabrum (Wehmer) Westling mold, suberosis 

Penicillium implicatum Biourge mold on poplar wood 

Penicillium nalgiovense Laxa salami-sausages mold 

Penicillium roqueforti Thom cheese mold 

Penicillium spinulosum Thom indoor mold 

Peziza repanda Pers. indoor wood 

Phaeolus schweinitzii (Fr.:Fr.)Pat.,Dyepolypore treerot 

Phanerochaete chrysosporium Burds. ligninase, biopulping 

Phanerochaete laevis (Fr.) J. Eriksson & Ryv. detoxification 

Phanerochaete sordaria (P. Karsten) J. Eriksson & Ryv. fatty acid profiles, lignin 

degradation 

Phellinus chrysoloma (Fr.) Donk white rot 

Phellinus contiguus (Pers.) Pat. indoor white-rot 

Phellinus hartigii (Allesch. & Schnabl) Pat. white rot 

Phellinus igniarius (L.: Fr.) Quélet, False tinder fungus white rot 

Phellinus nigrolimitatus (Romell) Bourdot & Galzin, 

Black-edged polypore 

tree rot 

Phellinus pini (Brot.: Fr.) A. Ames, Ochre-orange 

hoof polypore 

tree rot 

Phellinus pomaceus (Pers.: Fr.) Maire white rot 

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Phellinus robustus (P. Karsten) Bourdot & Galzin white rot 

Phellinus tremulae (Bondartsev) Bondartsev & Borrisow parasite 

Phellinus weirii (Murrill) Bilb. parasite 

Phialophora fastigiata (Lagerb. & Melin) Conant grey stain of poplar 

Phlebia brevispora Nakasone specific PCR, biopulping 

Phlebia chrysocreas (Berk. & M.A. Curtis) Burds “palo podrido” 

Phlebia radiata Fr. lignin degradation 

Phlebiopsis gigantea (Fr.) Jül., Conifer parchment antagonism 

Pholiota carbonica A.H. Sm. competition 

Pholiota highlandensis (Peck) Hesler & A.H. Sm. competition 

Pholiota nameko (T.Itô)S.Ito&S.Imai ediblemushroom 

Pholiota squarrosa (Pers.: Fr.) Kummer white rot 

Phoma exigua Sacc. blue stain 

Physisporinus vitreus (Pers.: Fr.) P. Karsten, Pole fungus manganese deposits 

Phytophthora cactorum (Lebert & Cohn) Schröter Beech seedling disease 

Phytium debaryanum Hesse conifer seedling parasite 

Piptoporus betulinus (Bull.: Fr.) P. Karsten, Birch polypore tree rot 

Pleurotus ostreatus (Jacq.: Fr.) Kummer, Oyster fungus edible mushroom on wood 

Pleurotus ostreatus ssp. florida edible mushroom on wood 

Pluteus cervinus (Schaeffer) Kummer indoor wood 

Polyporus squamosus (Hudson: Fr.) Fr., Scaly polypore tree rot 

Pycnoporus cinnabarinus (Jacq.) Fr. lignin degradation 

Ramariopsis kunzei (Fr.) Corner indoor wood 

Resinicium bicolor (Alb. & Schwein.: Fr.) Parm. indoor white-rot 

Reticularia lycoperdon Bull. indoor wood 

Rhinocladiella atrovirens Nannf. blue stain 

Rhizina undulata Fr. root-decay ascomycete 

Rhizoctonia solani Kühn beech nut rotting 

Rigidoporus lineatus (Pers.) Ryv. mine timber 

Rigidoporus vinctus (Berk.) Ryv. mine timber 

Rosellinia aquila (Fr.) de Not. seedling smothering 

Rosellinia minor (Höhn) Francis seedling smothering 

Rosellinia quercina R. Hartig oak root parasite 

Saccharomyces cerevisiae Meyen: E.C. Hansen yeast 

Schizophyllum commune Fr.: Fr., Split-gill stored and exterior wood 

Sclerophoma pithyophila (Corda) v. Höhn. blue stain 

Scutellinia scutellata Lambotte indoor wood 

Serpula himantioides (Fr.: Fr.) P. Karsten, Wild merulius indoor wood 

Serpula lacrymans (Wulfen: Fr.) Schroeter apud Cohn, indoor wood 

(True) Dry rot fungus 

Sirococcus conigenus (DC.) P.F. Cannon & Minter tree parasite 

Sirococcus strobilinus Preuss Sirococcus shoot dieback 

Sistotrema brinkmanni (Bres.) J. Eriksson stored and exterior wood 

Sparassis crispa Wulfen: Fr. tree rot 

Sphaeropsis sapinea (Desm.) Dyko & Sutton Conifer seedling shoot tip 

disease 

Sphaerotheca lanestris Harkn. virus vector 

Stachybotrys chartarum (Ehrenb.) Hughes toxic mold 

Stereum hirsutum (Willd.: Fr.) S.F. Gray, Hairy Stereum stored and exterior wood 

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Stereum rugosum (Pers.: Fr.) Fr. white rot 

Stereum sanguinolentum (Alb. & Schwein.: Fr.) Fr., 

red streaking, Wound rot of 

Bleeding Stereum 

spruce 

Strasseria geniculata (Berk. & Broome) Höhn blue stain, Conifer seedling 

shot tip disease 

Thekopsora areolata (Fr.) Magnus spruce inflorescence 

damage 

Thelephora terrestris Erh. seedling smothering 

Thielavia terrestris (Apinis) Malloch & Cain. soft rot 

Trametes hirsuta (Wulfen: Fr.) Pilát white rot 

Trametes multicolor (Schaeffer) Jül. indoor wood 

Trametes pubescens (Schum.) Pilát fatty acid profile 

Trametes versicolor (L: Fr.) Pilát, Many-zoned polypore stored and exterior wood 

Trechispora farinacea (Pers.) Liberta indoor wood 

Trechispora mollusca (Pers.) Liberta indoor wood 

Trichaptum abietinum (Dicks.: Fr.) Ryv., Fir Polystictus red streaking 

Trichoderma hamatum (Bonord.) Bainier mushroom parasite 

Trichoderma harzianum Rifai mushroom parasite 

Trichoderma parceramosum Bissett mushroom parasite 

Trichoderma pseudokoningii Oudem. mushroom parasite 

Trichoderma reesei E.G. Simmons enzymes 

Trichoderma viride Pers.: Fr. mold, antagonism, 

enzymes 

Tyromyces caesius (Schrader: Fr.) Murrill, Blue cheese 

polypore 

brown rot 

Tyromyces stipticus (Pers.: Fr.) Kotl. & Pouzar brown rot 

Volvariella bombycina (Schaeffer: Fr.) Singer indoor wood 

Xerocomus pruinatus (Fr.) Quélet IGS sequence 

Xylaria hypoxylon (L.) Grev. white-rot ascomycete 

Xylobolus frustulatus (Pers.: Fr.) Boidin, Ceramic parchment tree rot 

www.taq.ir

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Subject Index 

Abiotic wood discolorations 119 

Accessory compounds 57, 81, 90–1, 147, 

175–6, 194 

Acetylation of wood 156 

Acid production by fungi 70–4, 94, 97–8, 

104, 220, 232 

Actinobacteria 111 

Air 58–60, 174–5 

Algae on wood 119, 200 

Allergies through fungi 123–4 

Alternative wood protection 79–81, 156–9 

Amplified fragment length polymorphism 

(AFLP) 45 

Antagonisms 79–81, 124, 170, 194, 235 

Antrodia species 13–4, 23, 32, 38, 41, 44, 69, 

73–4, 77, 138, 156, 201, 207–11, 216–7, 

218–20, 230, 233, 255 

Archaea 110 

Archaeological wood 105, 116–7 

Armillaria root disease 187 

Armillaria species 6, 9, 14–5, 23, 32, 34–6, 

38–9, 41, 43–5, 60, 74, 80, 104, 118, 132, 

135, 173, 184, 186–189, 200–1, 224, 259 

Arthrospores 15, 17, 66, 69, 80, 103 

Ascomycetes 16, 18–20, 27, 31, 49–50, 70, 

84, 102, 120, 125, 137–8, 142, 184, 201 

Ascospores 19–20, 75 

Asexual development 10–6 

Aspergillus species 16–7, 50, 62–3, 70, 80, 

87, 121–5 

Asterostroma species 212–3, 257–8 

Aureobasidium pullulans 50, 80, 125, 

127–8, 248 

Bacteria 4, 47, 54, 58, 60, 63, 69–71, 75, 80, 

93–5, 98, 109–18, 132, 156, 200, 236, 246, 

251 

Bacteria and forest damages 110–3, 170 

Bacteria and wood used in kitchens 118 

Bacteria as antagonists 80, 118, 170, 189 

Bacteria as pathogens to trees 111, 114 

Bacterial degradation of pits 60, 93, 112, 

116, 132 

Bacterial wood degradation 55, 113–4 

Bacterial wood discoloration 56, 117 

Bark damages 163–8, 175–6, 199 

Bark extracts 251 

Basidiomycetes 16, 18, 21, 25, 27, 31, 47, 

49–50, 59, 64, 70, 84, 112, 129, 135, 137–8, 

182, 208, 222, 224 

Basidiospores 21, 23, 70, 193, 226, 229, 232 

Bavendamm test 32, 101–2 

Beech bark disease 163–5, 197 

Biological forest protection 80 

Biological influences on fungi 53, 79–85 

Biological pulping 140, 244–5 

Biological wood protection 80, 159 

Bioremediation of preservatives 74, 156 

Biotechnology of lignocelluloses 237–51 

Biotic wood discolorations 119–33 

Blue stain 80, 125–8 

Blue stain fungi 11–2, 21, 50, 54, 59, 65, 67, 

81, 125–8, 132, 200, 210 

Boron wood preservatives 74, 132, 151, 153, 

155, 194 

Brown pocket rot 136–7 

Brown rot 135–8, 184, 198–200, 202, 205, 

219–20, 223 

Brown rot fungi 32, 72, 96–7, 101, 129, 

135–8, 204, 207, 218 

Butt rot 189–90, 198 

Carbon dioxide 58–60, 66, 84, 158, 245 

Carbon sources for fungi 53–5 

Cavity bacteria 114 

Cavity formation 114, 135, 142–4, 199 

Cellar fungi 220–3, 234, 236 

Cellulases 87, 95–7, 121, 249–50 

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330 Subject Index 

Cellulose 54, 59, 66, 88, 90–2, 95–9, 237 

Cellulose degradation 72, 87–9, 92, 95–9, 

104, 113–4, 122, 135, 138, 145 

Ceratocystis fagacearum 32, 170–2 

Ceratocystis fimbriata 32–3, 167–8 

Ceratocystis minor 32, 125, 127 

Chaetomium globosum 26, 43, 60, 75, 81, 

123–4, 142, 145 

Chemical wood discolorations 117, 119, 133 

Chemical wood preservation 149–53 

Chestnut blight 165–7 

Chitin 4, 56, 156, 158, 183, 235 

Chitosan for wood protection 158 

Chlamydospores 15, 17, 60, 103, 125–6 

Chlorociboria species 119–20 

Chromium wood preservatives 74, 116–7, 

144–5, 151–2, 156, 220 

Cladosporium species 26, 50, 123, 125, 127 

Clamp formation 21–2 

Classification of fungi 47–52, 162–4 

Coal tar oil wood preservatives 146, 151, 

153 

CODIT model 175 

Collections of fungi 32 

Competitions 79–85 

Conidia 16–8, 25, 31, 52, 59, 75, 121, 123, 

191–2 

Coniophora species 9, 13–5, 22–3, 32, 35–6, 

38, 41, 44, 46–7, 59, 64, 66, 69, 77, 80, 

96–8, 116, 135, 138, 158, 182, 201, 205, 

207–11, 218, 220–3, 230, 233, 235, 245, 

255 

Copper tolerance 73, 220, 232 

Copper wood preservatives 74, 146, 151 

Cryphonectria parasitica 165–6 

Cubical rot 138, 145, 219, 229 

Dacrymyces stillatus 214 

Daedalea quercina 23, 74, 161, 184, 200–1, 

202, 208, 256 

Damage to buds, shoots and branches 163 

Damage to seeds and seedlings 161–2 

Dark fruit body 75, 203–4 

Deinking 250 

Demarcation lines in wood 79, 139, 197, 

199, 206 

Detection methods of tree and wood 

damages 179–83 

Deuteromycetes 16–8, 31, 47, 49–50, 52, 60, 

102, 120–1, 124–5, 142, 201 

Development of Ascomycetes 18–20, 27 

Development of Basidiomycetes 21–3 

Development of Deuteromycetes 16–8 

Diplomitoporus lindbladii 212, 258 

Discula pinicola 50, 127 

Disposal of preservative-treated wood 74, 

155–6, 250 

DNA-arrays 45 

DNA-based techniques 35–46, 233 

DNA-restriction 35, 37–9 

DNA-sequencing 39–44, 77, 210, 219 

Donkioporia expansa 15, 38, 44, 56, 67, 138, 

201, 207–8, 214–6, 233, 259 

“Dry rot” 231 

Dry rot fungi 207–8, 210, 219–20, 223–33 

Dutch elm disease 168–70 

Edible mushrooms 36, 68, 84, 188, 197, 

199–200, 206, 239–43 

Environmental concerns of wood 

preservation 150–6 

Environmental pollutants 84–5, 132, 153, 

155, 250 

Enzymatic bleaching 249 

Enzymes 57, 60, 87–107, 238, 249 

Enzymes in pulp and paper production 

249–50 

Erosion bacteria 114–5 

Ethanol production 59, 247–8 

Excessive preservative uptake 60, 93, 116 

Fatty acid profiles of fungi 47 

Felling time of trees 131, 147–8 

Fenton reaction 97–8, 105–6 

Fermentation of spent sulphite liquors 59, 

248 

Fiber saturation point 63–5, 231, 236 

Fixation of wood preservatives 151 

Flammulina velutipes 24, 76–7, 238, 242 

Fomes butt (root) rot 189–95 

Fomes fomentarius 4, 23, 25, 75–6, 138, 140, 

165, 184, 195, 200 

Force of gravity 25, 74–7 

Forest diseases 84–5, 109, 112, 114, 121, 

161–73, 184, 186–200 

Formal genetics of fungi 26–31 

Frost cracks 113 

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Subject Index 331 

Fruit body formation 16, 19–25, 52, 74–7, 

102, 180, 229, 240–3 

Fruit body types of Ascomycetes 20–1, 31, 

49–50 

Fruit body types of Basidiomycetes 23–4, 

31, 50, 184 

Fumigation against fungi 172–3, 235 

Fungal cell wall 4, 16, 55 

Fungal dry matter 53, 56, 243 

Fungal protoplasm 5–9 

Fungi imperfecti 16, 121 

Fungi in cooling towers 60, 140–2, 201, 216 

Fungi in piled wood chips 107, 121, 201, 245 

Fungi in wood artifacts 235 

Fungi on hardwoods 137, 140, 142, 184, 

188, 195, 199, 202, 206, 212, 214 

Fungi on leaf litter 201 

Fungi on masonry 224, 227, 231–2, 235–6 

Fungi on mining timber 201, 204–6, 216–7, 

219–20, 222 

Fungi on monocotyledons 144, 206 

Fungi on poles 142, 201, 204–5, 217, 219, 

222 

Fungi on softwoods 137, 142, 184, 186, 188, 

190, 195, 198, 200, 202, 205, 207, 212–4, 

219–20, 230 

Fungi on stored wood 121, 128, 138, 145, 

200–7 

Fungi on structural timbers indoors 137, 

201–2, 204–5, 207–33 

Fungi on structural timbers outdoors 137, 

145, 200–7, 219, 226–7 

Fungi on stumps 80, 82, 187, 191, 200, 

204–6, 217, 219, 227 

Fungi on trees 65, 82, 121, 127–8, 137–8, 

145, 161–73, 183–200, 202, 206 

Fungi on urban tress 184, 189, 197, 199, 216 

Fungi on window joinery 202, 204, 207–8, 

214 

Fungi on wood in aquatic habitats 62, 142, 

145, 201, 205 

Ganoderma species 24, 80, 129, 135, 

139–40, 200, 246 

Gloeophyllum species 15, 23–4, 31, 35, 38, 

58, 66–7, 73–4, 98, 137–8, 155, 158, 200–1, 

202–05, 207–8, 211, 233, 235, 256–7 

“Green rot” 119–20 

Grouping of Bacteria 110–2 

Grouping of Deuteromycetes 16, 52 

Grouping of wood preservatives 150–1 

Growth rate of mycelium 9–10, 78, 98, 210, 

223, 232 

Guttation 56, 215, 229, 231 

Hazard classes of timber 148 

Heart rots 184, 193, 198–9, 202, 205, 216 

Heat treatment against fungi 70, 220, 231, 

235 

Hemicellulose degradation 89, 92–4, 122, 

135, 145 

Hemicelluloses 54, 87–8, 91–2, 237, 248 

Heterobasidion species 16, 21, 23, 25–6, 

30–4, 36, 38–9, 43, 57–9, 66, 80, 135, 138, 

140, 184, 189–95, 200–1 

Heterothallic (homothallic) fungi 26–7 

Hook formation 19 

House rot fungi 12–4, 32, 65, 68, 71, 73, 

77–8, 108, 138, 207–33 

Hypha 3–8, 11–2, 12, 55 

Hyphal zonation 8–9 

Identification keys for fungi 31–2, 101, 210, 

253–9 

Identification of Ascomycetes 31–2 

Identification of Bacteria 118 

Identification of Basidiomycetes 31–47, 

186, 210, 233 

Identification of Deuteromycetes 31, 52, 125 

Immunological methods for enzymes 35, 

98, 105 

Immunological methods for fungi 34–5, 

233 

Inspection methods for fungal activity 

179–83 

Interactions between microorganisms 

79–85 

Internal transcribed spacer of rDNA 37–44 

Intersterility groups 29–30, 35, 39, 186, 

189–90 

Isolate variation in fungi 9, 78, 220, 223, 232 

Isozyme analysis 34 

Laccase 32, 57, 101–2, 104, 106, 136, 158, 

249–50 

Laetiporus sulphureus 23, 25, 39, 59, 184, 

197, 201, 208 

Leaf diseases 109, 162–3 

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Lentinula edodes 6, 9, 24, 31, 35, 43, 56, 69, 

74, 104–5, 135, 238–42, 246, 249 

Lentinus lepideus 35, 66, 74, 135, 201, 205, 

208 

Leucogyrophana species 41, 201, 223, 

225–7, 228, 253–4 

Lichens 85 

Light 24, 26, 74–5, 199, 241 

Lignin 54, 88–9, 91, 99–107, 113–4, 138, 

145, 183, 219, 237, 246 

Lignin carbohydrate complex 91–2, 94, 99, 

237, 249 

Lignin degradation 72, 89, 91–2, 99–107, 

136, 138 

Lignin peroxidase 57, 72, 89, 102–6, 249 

“Lower fungi” 48–9 

Low-molecular agents 72, 98, 104–6, 137–8, 

249 

MALDI-TOF mass spectrometry 46–7, 233 

Manganese deposits 57, 140–1, 246 

Manganese peroxidase 57, 72, 89, 104, 106, 

140, 246, 249 

Mannan 89, 94 

Mating of mycelia 26–30 

Melanin 125 

Meripilus giganteus 23, 135, 184, 197, 200 

Meruliporia incrassata 13, 33, 41, 138, 223, 

226–7, 228–9, 232 

Microbial volatile organic compounds 

(MVOCs) 124, 233 

Microsatellites 44–5 

Microscopic methods for fungi 183 

Microwaves against fungi 235 

Minerals in wood 56–7, 83, 97–8 

Molding 58, 121–5, 210 

Molds 16, 54, 58–9, 67, 120–1, 132, 200, 232 

Molds as pathogens 123–4 

Molecular genetics of enzymes 104–5, 249 

Molecular techniques for fungi 33–47, 183, 

190, 210, 227 

“Moon wood” 148 

Mushroom production on wood 237, 

239–42 

Mycelium 3, 6, 9–11, 32, 55, 189, 215, 223 

Mycoallergies 123–4 

“Myco-fodder” 246 

Mycoplasmas 112, 164, 173 

Mycorrhiza 36, 38–9, 41, 82–5, 189 

Mycoses 124 

Mycotoxicoses 123 

“Myco-wood” 206, 237–9 

Naming of enzymes 88 

Naming of fungi 47–8 

National regulations for wood preservation 

149–50 

Natural durability 58, 145–7, 182 

Nectria species 164 

Needle diseases 162–3 

Nitrogen in wood 56, 81, 246 

Nutrient uptake 55 

Nutrients for fungi 53–8, 81, 146 

Oak disease in Europe 173 

Oak wilt disease 170–3 

Oligoporus placenta 23, 27, 31, 35, 38, 64, 

77, 80, 94, 97–8, 137, 158–9, 201, 218, 

219–20, 233, 235–6 

Ophiostoma piceae 31–2, 120, 125, 127, 158, 

172 

Ophiostoma piliferum 32–3, 43, 81, 125, 

127, 245, 249 

Ophiostoma ulmi 25, 31–2, 80, 168–70 

Oxalic acid (oxalate) 71–3, 94, 97–8, 104, 

220, 232 

Oxygen 58–9, 103, 111–2, 116, 133, 145–6, 

245 

Paecilomyces variotii 50, 63, 75, 94, 120–1, 

132, 248, 251 

“Palo podrido” 139–40, 246–7 

Parasits 53–4, 59, 186–200, 219 

Paxillus panuoides 66, 74, 201, 205, 208, 255 

Pectin 92–3, 125 

Penicillium species 16, 50, 58, 75, 80–1, 

121–3, 161 

Pentachlorophenol 153, 155–6, 250 

Phaeolus schweinitzii 23, 184, 198, 200 

Phanerochaete chrysosporium 5, 31, 47, 67, 

87–8, 96, 102–4, 107, 201, 244, 250 

Phellinus pini 11, 23, 35, 39, 135, 137, 140, 

186, 198, 257 

Phenol oxidases 32, 102, 133 

Phlebiopsis gigantea 80, 194, 200–1, 208 

pH-value 26, 58, 70–2, 98, 117, 119, 145, 225 

pH-value of wood 70, 112 

www.taq.ir

Subject Index 333 

Phylogenetic analysis 41–2 

Physical/chemical influences on fungi 

53–77 

Piptoporus betulinus 23, 25, 58, 184, 199, 

233 

Plane canker stain disease 167–8 

Pleurotus ostreatus 24, 30–1, 43, 56, 104, 

200, 238, 242, 250 

Polymerase chain reaction 35–45 

Polypore fungi 27, 66, 137, 184, 196–7, 219 

Polyporus squamosus 184, 199, 208 

Positive effects of wood microorganisms 

237–51 

Prerequisites for fungal activity 84, 90, 97, 

101, 105, 145–6 

Pretreatment of wood for fungi 99, 237–8, 

246–8 

Prokaryotes 4, 109–10, 112 

Protection measures against fungi 131–3, 

146–59, 166, 168, 170, 172–3, 178, 180, 

189, 194–5, 201–2, 216, 233–6 

Protein-based techniques for identification 

33 

Protoplast fusion 31 

Pruning 177–8 

Randomly amplified polymorphic DNA 

(RAPD) 36–7 

Red-streaking discoloration 129–31, 195 

Red-streaking fungi 54, 129–31, 200 

Relative air humidity 64, 68, 123, 226–7 

Relativity of experimental data 77–8 

Remedial measures of indoor rot 233–6 

Resistance to heat 15, 69–70, 111, 204–5, 

216, 220, 223, 231 

Resistance to dryness 15, 60, 66, 129, 145, 

204–6, 220, 223, 231–2 

Restriction fragment length 

polymorphism of DNA (RFLP) 35, 37–9 

Rhizomorphs 9, 12–5, 186–9 

Ribosomal DNA (rDNA) 37–44, 110, 118, 

210, 226 

Rickettsia 111 

Root graft transmission 169, 171, 188, 191, 

194 

Root rots 184–90, 197–8, 200 

Sap stain 125–8 

Saprobes 53, 59, 187, 195, 198 

Schizophyllum commune 24–5, 27, 60, 66, 

71–2, 77, 129, 168, 200–1, 206 

SDS polyacrylamide gelelectrophoresis 

(SDS-PAGE) 33–4, 233 

“Selective delignification” 140, 244 

“Selective white-rot” 107, 140 

Septum 6, 50 

Serpula species 3, 9, 11–5, 23–6, 28–30, 

32–8, 41–2, 44–7, 56–7, 59, 62–4, 66–9, 

73–5, 97–8, 138, 153, 201, 207–11, 218, 

223–8, 229–36, 253–4, 257 

Sexual development of fungi 16, 18, 26–9 

Simultaneous white rot 138–9, 191, 197 

Size of wood microorganisms 3 

Slime fungi 48–9, 200 

Slime layer around hyphae 5, 90, 105, 

138–40, 144 

Sniffer dogs 124, 180, 233 

Soft rot 135, 142–6, 184, 207 

Soft-rot fungi 50, 54, 60, 66, 70, 102, 116, 

142–6, 201 

Solvent-based preservatives 152 

Somatic incompatibility 30 

Sparassis crispa 184, 186, 200 

Species-specific priming PCR 43–4, 210 

Spore dispersal 25–6, 123–4 

Spore germination 26, 50, 232 

Spores 15–9, 21, 23–5, 81, 102, 123 

Sporotrichum pulverulentum 81, 96, 107, 

244, 248 

Stainings for fungi 183, 233 

Standards for wood preservation 148–9 

Steam explosion treatment 238, 248 

Stem decays 184–5, 193, 195, 199 

Stereum sanguinolentum 23, 66, 80, 129–30, 

131, 135, 161, 195 

Stereum species 9, 26, 28, 80, 82, 131, 168, 

195, 200–1 

Strand diagnosis 13–4, 210, 253–9 

Strands (cords) 12–3, 57, 204, 210, 212–3, 

216–9, 222–3, 227–31 

Succession 82 

Successive white rot 139–40 

Super critical fluid treatment 156, 168 

Surveys on occurrence of house-rot fungi 

207–9, 218, 220, 224 

Symbioses 82–5 

Synergisms 81 

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Temperature 6, 24–6, 32, 67–70, 78, 110, 

121, 126, 145, 186, 210–1, 216, 223, 227, 

229–30, 245 

Test ratings of wood preservatives 150 

Thermal wood modification 157 

Trametes versicolor 23–4, 31, 35–6, 47–8, 

58, 64, 72–3, 79–80, 82, 90, 103–4, 129, 

138–40, 155, 157, 159, 200–1, 206–7, 208, 

238, 250 

Translocation of nutrients 15, 55–7, 189, 

225, 227, 231, 235–6 

Transpressorium 11–2, 126, 199 

Treatmentprocessesoftimberwith 

preservatives 153–4 

Tree care 177–8 

Tree defense against microorganisms 

174–7, 189, 193, 195 

Tree polypores 184 

Tree rots 82, 183–200 

Trichaptum abietinum 39, 66, 129, 131, 

201 

Trichoderma species 31, 33, 43, 50, 58, 80–1, 

87–8, 96, 121, 124, 170, 189, 235, 245 

Tunneling bacteria 114 

Tyrosinase 102 

UV light 1, 26, 75, 183, 244 

Vegetative development of fungi 7–14, 19, 

21 

Viruses 47, 109, 166, 170, 194–5 

Vitamins 25–6, 57, 81, 84, 87, 112, 243 

Water 53, 60–1, 146 

Water activity 62–3, 121, 123 

Water formation by fungi 58, 66, 231 

Water potential 56, 62–3 

Water storage of wood 60, 113, 116, 131–2, 

187 

Water transport by fungi 6, 231, 234 

Water-based preservatives 151 

Wet heartwood 70–1, 110, 112 

“Wet rot” 219, 222 

White pocket rot 137–8, 140 

White rot 129, 135, 137–42, 184, 191, 195, 

198–9, 206, 212–4, 216 

White rot fungi 32, 72, 88, 101–2, 129, 

138–42, 244, 246 

Wilt diseases 168–73 

Wood cell wall 92 

Wood damaging agents 1 

Wood decays 135–46, 186–200 

Wood discolorations 117, 119–32 

Wood dry weight 61, 182 

Wood hydrophobization 156–8 

Wood modifications 156–7 

Wood moisture 57, 60–7, 123, 127, 129, 

145–7, 201, 204, 210–1, 216, 219, 222, 224, 

226–7, 230–1, 233–6, 245 

Wood parenchyma 117, 119, 122, 125, 129, 

133, 175 

Wood preservation 117, 124, 133, 146, 

149–53, 178, 182, 234 

Wood protection 131–3, 146–59 

Wood saccharification 247–8 

Wood strength properties 183 

Wood-based composites 149, 155, 230 

Wood-decay fungi 10, 32–3, 40, 43, 47, 54, 

57, 82, 185 

Wood-inhabiting fungi 23, 32, 54, 82, 121, 

125 

Wood-plastic composites 155 

Wound dressings 178, 195 

Wound parasites 53, 174, 184, 206, 222 

Wound reaction in trees 112, 174–8 

Wound rot of spruce 195 

Wound treatment 178–80 

Xylan 89, 93, 249 

Xylan degradation 89, 93–4, 125 

Yeasts 4, 32, 47, 59, 201, 246–8 

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6 Wood Discoloration

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