Frequency of Cyanogenesis in Tropical Rainforests of Far North ...

Frequency of Cyanogenesis in Tropical Rainforests of 

Far North Queensland, Australia 

REBECCA E. MILLER*, RIGEL JENSEN and IAN E. WOODROW 

School of Botany, The University of Melbourne, Victoria, 3010, Australia 

Background and Aims Plant cyanogenesis is the release of toxic cyanide from endogenous cyanide-containing 

compounds, typically cyanogenic glycosides. Despite a large body of phytochemical, taxonomic and ecological 

work on cyanogenic species, little is known of their frequency in natural plant communities. This study aimed to 

investigate the frequency of cyanogenesis in Australian tropical rainforests. Secondary aims were to quantify the 

cyanogenic glycoside content of tissues, to investigate intra-plant and intra-population variation in cyanogenic 

glycoside concentration and to appraise the potential chemotaxonomic significance of any findings in relation to the 

distribution of cyanogenesis in related taxa. 

Methods All species in six 200 m 2 plots at each of five sites across lowland, upland and highland tropical rainforest 

were screened for cyanogenesis using Feigl–Anger indicator papers. The concentrations of cyanogenic glycosides 

were accurately determined for all cyanogenic individuals. 

Key Results Over 400 species from 87 plant families were screened. Overall, 18 species (45 %) were cyanogenic, 

accounting for 73 % of total stem basal area. Cyanogenesis has not previously been reported for 17 of the 18 species, 

13 of which are endemic to Australia. Several species belong to plant families or orders in which cyanogenesis has 

been little reported, if at all (e.g. Elaeocarpaceae, Myrsinaceae, Araliaceae and Lamiaceae). A number of species 

contained concentrations of cyanogenic glycosides among the highest ever reported for mature leaves—up to 

52mgCNg 1 d. wt, for example, in leaves of Elaeocarpus sericopetalus. There was significant variation in cyanogenic 

glycoside concentration within individuals; young leaves and reproductive tissues typically had higher 

cyanogen content. In addition, there was substantial variation in cyanogenic glycoside content within populations 

of single species. 

Conclusions This study expands the limited knowledge of the frequency of cyanogenesis in natural plant 

communities, includes novel reports of cyanogenesis among a range of taxa and characterizes patterns in intra-plant 

and intra-population variation of cyanogensis. 

Key words: Australia, b-glycosidase, chemotaxonomy, cyanogenic glycoside, cyanogenesis, defence, hydrogen cyanide, 

polymorphism, Queensland, screening, secondary metabolite, tropical rainforest. 

INTRODUCTION 

Cyanogenesis is the ability to release toxic hydrogen 

cyanide (HCN) from endogenous cyanide-containing compounds. 

It has long been recognized in plants (Conn, 1991; 

Seigler, 1991), and has been recorded in ferns, fern-allies, 

gymnosperms, as well as monocotyledonous and dicotyledonous 

angiosperms from >550 genera and 130 plant 

families (Conn, 1981; Poulton, 1990; Jones, 1998). Cyanogenesis 

in plants requires the presence of either an unstable 

cyanohydrin, or of a stable cyanogen and its degradative 

enzymes (Seigler, 1991). While cyanolipids have been 

identified from a few taxa (Mikolajczak et al., 1970), cyanogenesis 

in plants most commonly results from the hydrolysis 

of cyanogenic glycosides (Conn, 1981). Autotoxicity in 

intact plants is prevented by the spatial separation— 

either at the subcellular or at the tissue level—of the cyanogenic 

glycoside and catabolic enzymes (Kojima et al., 1979; 

Selmar, 1993a; Poulton and Li, 1994; Zheng and Poulton, 

1995; Hickel et al., 1996). The catabolism of cyanogenic 

glycosides is therefore initiated upon tissue disruption, 

due to mechanical damage or ingestion by herbivores, for 

example, which enables mixing of enzymes and cyanogenic 

substrate (Wajant et al., 1994; Patton et al., 1997). 

Little is known about the frequency of cyanogenesis 

in natural plant communities. This is despite a large body 

of literature documenting cyanogenesis in >2650 species of 

angiosperms worldwide (Lechtenberg and Nahrstedt, 1999). 

Indeed, as many as 11 % of all plant species are predicted to 

be cyanogenic (Jones, 1998). Historically, much of the 

interest in cyanogenesis centred around recording toxic 

plants with the potential for stock and human poisoning, 

the high frequency of cyanogenesis among food plants 

(Jones, 1998), and the potential utility of cyanogenesis 

and the structure of specific cyanogens in elucidating phylogenetic 

relationships between taxa (e.g. Gibbs, 1974). As a 

consequence, much of what is known about the frequency 

of cyanogenesis comes from surveys of regional floras, or 

of specific taxonomic groups. There are a number of 

substantial chemotaxonomic works incorporating information 

on cyanogenesis (see Hegnauer, 1966, 1973, 1986, 

1989, 1990; Gibbs, 1974), several smaller and more specific 

chemotaxonomic works (e.g. Tjon Sie Fat, 1978, 1979b; 

Spencer and Seigler, 1985; van Wyk and Whitehead, 

1990; Seigler, 1994), including work on Australian Acacia 

spp. (Conn et al., 1985; Maslin et al., 1988), and numerous 

inventories of cyanogenic species (e.g. Rosenthaler, 1919; 

Seigler, 1976a, 1976b; Tjon Sie Fat, 1979a; Francisco and 

Pinotti, 2000). Several Australian researchers were active in 

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the field early last century, reporting a number of 

cyanogenic species and the specific cyanogenic constituents 

involved (e.g. Petrie, 1912, 1913, 1914, 1920; 

Finnemore and Cox, 1928, 1929; Finnemore and 

Cooper, 1936, 1938). More than 700 species in the 

Queensland flora were screened by Smith and White 

(1918). Further phytochemical screening of the Queensland 

flora was conducted by Webb (1948, 1949); however, there 

was negligible testing among tropical taxa. Importantly, 

many of these records are based on qualitative tests performed 

using herbarium specimens, and tests of this kind 

using dried material are by no means conclusive. One further 

limitation of these data in terms of extrapolating to 

natural plant communities is that negative results were 

often not reported. 

Tropical rainforests are of particular interest in the 

study of plant chemical defences. Elevated herbivore pressure 

in tropical environments is hypothesized to have 

favoured both a diverse array of defences and high levels 

of investment in chemical defence (Levin, 1976; Levin and 

York, 1978; Coley and Barone, 1996; Kursar and Coley, 

2003). Indeed, in the field of plant secondary chemistry, the 

tropical rainforest has been the focus of intense interest 

in co-evolutionary relationships between plants and 

herbivores, and the extraordinary inter-specific and intraplant 

variation in chemical defence strategies (Feeny, 

1976; Levin, 1976; Coley et al., 1985; Coley and 

Kursar, 1996). 

On the whole, community-level studies of the distribution 

of chemical defences have focused on a small set of Asian 

and African rainforests (McKey et al., 1978; Gartlan et al., 

1980; Davies et al., 1988; Waterman et al., 1988). In 

addition, these and other studies have tended to focus on 

C-based ‘quantitative’ defences (e.g. Coley, 1983, 1988). 

Among N-based defences, the greater awareness of the 

frequency of alkaloid-bearing plants in tropical and other 

ecosystems is a consequence of extensive phytochemical 

screening for bioactive compounds (e.g. Swanholm et al., 

1960; Hartley et al., 1973; Smolenski et al., 1974; Collins 

et al., 1990; Hadi and Bremmer, 2001) or ecological 

research into specific plant–herbivore interactions (e.g. 

Janzen and Waterman, 1984; Hartmann et al., 1997), rather 

than systematic studies within natural communities (but 

see Mali and Borges, 2003). 

There are some hypothesis-driven surveys for cyanogenesis. 

Two studies have investigated the frequency of cyanogenesis 

in floras, investigating evolutionary and ecological 

hypotheses about exposure to herbivory (see Kaplan et al., 

1983; Adsersen et al., 1988; Adsersen and Adsersen, 1993). 

However, only the survey of Thomsen and Brimer (1997) in 

Costa Rican rainforest has been conducted in a systematic 

fashion, well defined with respect to forest area and plant 

size. As with other N-based defences, work on cyanogenic 

tropical rainforest species worldwide is limited, and 

there has been no work on cyanogenesis in Australian tropical 

rainforests, or any other form of chemical defence. The 

lack of work on cyanogenesis in diverse tropical rainforests 

is further surprising, as it is a readily detectable and constitutive 

chemical defence (Gleadow and Woodrow, 

2000b). 

Here we report some of the findings of a large-scale 

quantitative survey for cyanogenesis in Australian tropical 

rainforests. First, we investigated the frequency of cyanogenesis 

and the contribution of cyanogenic species to 

biomass (basal area) in lowland, upland and highland tropical 

rainforest in north Queensland, Australia. Conducting 

the survey in a standardized fashion will enable comparison 

with other communities (e.g. Thomsen and Brimer, 1997). 

Secondly, intra-plant and intra-population variation in concentrations 

of cyanogenic glycosides was quantified. The 

high level of endemism among the Australian tropical rainforest 

flora (Webb and Tracey, 1981) and the number of 

rainforest taxa previously untested for cyanogenesis underscore 

the potential for novel reports of cyanogenesis within 

different taxonomic groups. Finally, therefore, this study 

aimed to investigate the potential taxonomic significance 

of cyanogenesis reported here. Overall, this study aimed 

to expand the limited knowledge of the frequency of 

cyanogenesis in the Australian flora and in natural plant 

communities. 

MATERIALS AND METHODS 

Field sites 

Field work was conducted between July 1999 and 

September 2002. Six 200 m 2 plots (20 · 10 m) were established 

at each of five sites (total area 1200 m 2 per site) in 

lowland and upland rainforest in the tropics of north east 

Queensland, Australia (Fig. 1). Six plots were selected for 

two reasons: first, the number of new species captured with 

each additional plot had reached a plateau at around five 

species and, secondly, it was a realistic sample size given 

the time and resources available. Sites were selected to 

capture maximum species diversity as on the Atherton 

Tablelands, forest type and species composition vary 

both with substrate and with altitude (Tracey, 1982). Further, 

to enable comparison of forests occurring on different 

soil types, two pairs of sites with similar altitude and rainfall 

were selected. The first pair comprised a site on soil derived 

from basalt at Lamins Hill and one on soil derived from 

granite at Mt Nomico (Fig. 1). The second pair comprised 

sites on soils derived from basalt and rhyolite at Longlands 

Gap (Fig. 1). A fifth site in lowland rainforest near 

Cape Tribulation and Myall Creek, on soil derived 

from metamorphic substrate, was also surveyed (Fig. 1). 

The distribution of cyanogenic species in relation to 

resource availability (soil nutrients) will be addressed 

elsewhere (see Miller, 2004). 

It was not possible to control strictly for logging history; 

all sites on the Atherton Tablelands had been selectively 

logged prior to the declaration of the Wet Tropics World 

Heritage Area in 1988. Detailed site descriptions are provided 

in Miller (2004). 

All individuals (palms, trees and vines) with a diameter at 

breast height (dbh) of >5 cm were tagged and identified. 

All additional species (dbh

Myall Creek 

Cape Tribulation 

Daintree River 

Port Douglas 

Western 

Australia 

Northern 

Territory 

South 

Australia 

Queensland 

Mareeba 

Atherton 

Tableland 

Atherton 

Cairns 

Mt Nomico 

Lamins Hill 

Malanda 

Longlands Gap 

0 50 km 

New South 

Wales 

F IG. 1. Location of study sites in tropical rainforest in far north east Queensland, Australia. There were two pairs of sites in upland rainforest on the Atherton 

Tableland: Lamins Hill (17 224 0 S, 145 425 0 E) and Mt Nomico (17 133 0 S, 145 404 0 E), and two sites at Longlands Gap (17 27 0 S, 145 28 0 E). A fifth site was 

in lowland rainforest near Myall Creek and Cape Tribulation (16 062 0 S, 145 269 0 E). 

Samples of all cyanogenic species, as well as species 

dominant at each site, and some rare species were pressed 

and lodged at the University of Melbourne (MELU) and 

Brisbane (BRI) Herbaria. Accession numbers, where 

assigned, are listed (Appendix). 

Climate 

The climate in far north Queensland is characterized 

by a marked wet season from December to April. Climate 

recording stations in the study area are scattered, therefore 

data specific to each site were not available. While located 

in the tropical latitudes, because of its higher altitude, the 

climate of the Atherton Tableland is semi-tropical. Mean 

annual temperature within the study area on the Tableland 

is 22 C (Nix, 1991), with a minimum of 10 C (Hall et al., 

1981; see Graham et al., 1995). All the upland and highland 

rainforests surveyed on the Atherton Tableland have 

high average annual rainfall, generally in the range 

2000–3000 mm plus cloud interception (Tracey, 1982). 

For example, the average annual rainfall at Lamins Hill 

is 3584 mm, based on 30 years of records from the Queensland 

Bureau of Meteorology (see Osunkoya et al., 1993). In 

contrast, the climate in the coastal lowland tropical rainforest 

near Cape Tribulation is characterized by higher 

temperatures. Mean daily temperatures range from 28 C 

in January to 22 C in July, and temperatures may reach 

the mid to high 30 s during the summer months. Average 

annual rainfall is also high, at 3928 mm recorded at Cape 

Tribulation (based on 65 years of records from the 

Queensland Bureau of Meteorology). 

Sites 

Upland and highland rainforest. The upland rainforest at 

Lamins Hill [17 224 0 S, 145 425 0 E; altitude 850 m above 

sea level (a.s.l.); Fig. 1] is classified as complex mesophyll 

vine forest on basalt (type 1b; Tracey, 1982; Tracey and 

Webb, 1975). This forest type typically occurs on upland 

sites (400–800 m), on high fertility kraznozem soils derived 

from basalt, with high rainfall (Tracey, 1982). It is characterized 

by a closed canopy with multiple tree layers, and 

an uneven canopy with height ranging from 35 to 45 m 

(Tracey 1982). 

The upland rainforest at Mt Nomico (17 133 0 S, 

145 404 0 E, 900 m a.s.l.; Fig. 1) is within the Gillies 

Range State Forest, and is on low nutrient soil derived 

from granite. The forest is classified as complex notophyll 

vine forest, typical of upland granitic soils (type 6; Tracey, 

1982; Tracey and Webb, 1975). 

In the highland rainforest of Longlands Gap State Forest 

(altitude 1100–1200 m a.s.l.), there is a sharp boundary 

in forest type defined by basalt (17 277 0 S; 145 285 0 E) 

and rhyolite (17 273 0 S, 145286 0 E) parent substrates.

On basalt, the forest is complex notophyll vine forest 

(type 5a; Tracey, 1982; Tracey and Webb, 1975), a forest 

type characterized by an uneven canopy (20–40 m high) and 

numerous tree layers, typical of basaltic cool wet uplands 

and highlands (Tracey, 1982). On rhyolite, the forest is 

simple microphyll vine forest (type 9; Tracey, 1982), 

which is common on upland granitic soils (800–1300 m), 

and is characterized by an uneven canopy 20–25 m high, 

with emergent Agathis atropurpurea (up to 35 m). 

Lowland rainforest. The fifth site was in lowland 

rainforest in the Daintree World Heritage Area, near 

Cape Tribulation (Fig. 1). The site was near to Thompson 

and Myall Creeks (16 062 0 S, 145 269 0 E; altitude 40 m 

a.s.l.). The forest is complex mesophyll vine forest (type 1a; 

Tracey, 1982), the canopy is irregular, from 25 to 33 m in 

height, and supports a great diversity of species and life 

forms, including many palms and lianas. The floristic 

composition is patchy, with considerable variation in 

canopy and understorey dominants over small distances 

(Webb et al., 1972; Tracey, 1982). The soil is relatively 

nutrient-poor red clay loam podsol derived from 

metamorphic substrate. 

Sampling for detection of cyanogenesis 

Whole leaf samples (1–2 g f. wt) were taken from individuals 

with a dbh >5 cm, and from all additional species 

in the lower strata, until at least three individuals of each 

species had been sampled. Where possible, the youngest 

fully expanded leaves without epiphyllous communities 

were selected; however, in the case of samples acquired 

by pruning shears attached to extension poles or by slingshot 

and rope from the canopy, it was not always possible 

to be selective. In instances where it was not possible to 

obtain samples from large canopy trees, samples were taken 

from nearby individuals of the same species. Fresh whole 

leaf samples used for cyanide testing were stored in air-tight 

bags on ice until tested for cyanide 2–6 h later using Feigl– 

Anger (FA) papers (Feigl and Anger, 1966). Individuals of 

rare species, and those with little foliage, were sampled only 

once for analysis in the laboratory where a quantitative 

assay was used to test for cyanogenesis using freezedried 

ground tissue. Because cyanogenesis is known to 

be a polymorphic trait (e.g. Hughes, 1991; Aikman et al., 

1996), a minimum of three individuals of all species was 

tested, more where species were common. Owing to the 

diverse and heterogeneous nature of the forests, multiple 

individuals of each species were not always represented in 

the plots. Therefore, for these rare species, testing individuals 

located outside the plots was required, and still there 

were several species for which only single samples were 

obtained. 

A range of other factors was taken into consideration 

when sampling. For example, given that young leaves 

of tropical species, in particular, are typically more highly 

defended than older leaves (Coley, 1983; Coley and Barone, 

1996), both young (soft expanding leaves) and old (recent 

fully expanded) leaves were tested for cyanogenesis, where 

possible. In addition, depending on availability, fruit and 

flowers from several species were tested. Owing to the large 

number of species and samples in the survey, it was not 

possible to examine seasonal trends in defence; however, 

individuals of the majority of species were tested in wet 

and dry seasons for qualitative changes in cyanogenic status, 

as there is some evidence for seasonal variation in 

cyanogenesis (Seigler, 1976b; Janzen et al., 1980; Kaplan 

et al., 1983). Finally, because edaphic factors have been 

reported to affect the expression of cyanogenesis (e.g. 

Urbanska, 1982), species found on more than one substrate 

type were tested at each site. 

Sample collection, handling and storage for quantitative 

analysis 

In addition to samples for fresh cyanide tests, whole leaf 

samples (again recent fully expanded leaves) were taken for 

quantification of cyanogenic glycosides. All individuals of 

species that produced a positive result and individuals of 

rare species were sampled. Depending on sampling conditions, 

samples were either placed immediately into liquid 

nitrogen, or placed in a sealed air-tight bag and kept on ice 

for 2–6 h until snap frozen in liquid nitrogen. Frozen 

samples were transported to the laboratory on dry ice, 

freeze-dried and stored on desiccant at 20 C for analysis. 

Freeze-dried samples were ground using either a cooled 

IKA Labortechnic A10 Analytical Mill (Janke and Kunkel, 

Stanfen, Germany) or, for smaller samples, an Ultramat 2 

Dental Grinder (Southern Dental Industries Ltd, Bayswater, 

Victoria, Australia). 

Chemical analyses 

Detection of cyanogenesis: Feigl–Anger papers. The 

presence of cyanogenic compounds in fresh field samples 

was determined using FA indicator papers (Feigl and 

Anger, 1966). FA papers were selected in preference to 

picrate papers because FA papers are more sensitive 

(Nahrstedt, 1980) and less prone to giving false-positive 

results (Brinker and Seigler, 1989). FA test papers were 

prepared according to Brinker and Seigler (1989). Because 

FA papers can be sensitive to moisture and light (Brinker 

and Seigler, 1989), they were stored in the dark and on 

desiccant until use. 

Fresh leaves (approx. 1–2 g f. wt) were crushed in 

duplicate screw-top vials. Old and young foliage samples 

were tested separately. To facilitate cyanogenesis, 05mLof 

water was added to one of the vials, and pectinase from 

Rhizopus spp. (Macerase Ò Pectinase, 441201 Calbiochem Ò , 

Calbiochem-Novabiochem Corp., La Jolla CA, USA) 

(04g L –1 )in01 M Tris–HCl (pH 68) was added to the 

other. Pectinase has been found to have non-specific b-glycosidase 

activity (Brimer et al., 1995) and, therefore, in 

the absence of sufficient endogenous b-glycosidase, enables 

tests for the presence of cyanogenic glycosides to be made. 

The indicator papers were suspended above the tissue by 

means of the screw-top lid, and vials were left at room 

temperature and checked after 12 and 24 h. This 24 h 

time period, used in other surveys (e.g. Dickenmann, 

1982; Thomsen and Brimer, 1997; Buhrmester et al., 

2000; Lewis and Zona, 2000), was selected to avoid spurious 

test results due to substantial bacterial contamination

which may occur beyond 24 h (Saupe et al., 1982). Tissue of 

known cyanogenic species, Prunus turneriana or Ryparosa 

javanica, was used as a positive control. Moderate to 

strongly cyanogenic samples gave a positive result within 

a few minutes or up to a few hours, while more weakly 

cyanogenic samples took several more hours. In accordance 

with the recommendations of Brinker and Seigler (1989), a 

new test was conducted for any samples producing a slow 

positive response (24 h) in case of interference by microbial 

cyanogenesis. In addition, any samples with inconclusive 

colour change were re-tested. An individual was considered 

cyanogenic if a positive, repeatable result was obtained, 

and a species was considered cyanogenic if at least 

one individual produced a consistent and repeatable 

positive result. A negative test result indicates the absence 

of a cyanogenic glycoside, or of the specific cyanogenic 

b-glycosidase, or both. 

In the few instances where insufficient tissue was available 

for both FA paper tests using fresh leaves and subsequent 

laboratory analysis, cyanogenesis was determined 

based on the quantitative assay of freeze-dried ground 

leaf tissue (as described below) by comparison with a 

negative tissue control (e.g. Alstonia scholaris or Aglaia 

meridionalis). 

In this study, tests for cyanogenesis used approx. 

1–2 g f. wt of leaves, which is larger than tissue samples 

tested in previous surveys (e.g. 50 mg f. wt by Lewis and 

Zona, 2000; and 200 mg f. wt by Thomsen and Brimer, 

1997; Buhrmester et al., 2000). According to Dickenmann 

(1982) who used 500 mg f. wt tissue, a weak positive reaction 

with FA papers, where part of the paper turns blue, 

indicated approx. 2–20 mg HCN kg –1 f. wt, while a strong 

reaction indicated >50 mg HCN kg –1 f. wt. These lower 

values equate to just over 6–60 mg HCN g –1 d. wt using a 

conversion based on the mean foliar water content of 

several species in this study, which was 70 %. In this 

study, based on fresh leaf tests and quantitative analysis 

of freeze-dried tissue from the same sample, the threshold 

sensitivity of FA papers was similar, within the range 

5–8 mg HCN g –1 d. wt. This threshold sensitivity of FA 

papers corresponds well to the criteria of Adsersen 

et al. (1988) for classifying individuals as cyanogenic, 

where individuals with

T ABLE 1. Summary of the concentration of cyanogenic glycosides in leaves and other plant tissues from cyanogenic species found in six plots in upland/highland (U) and 

lowland (L) tropical rainforest at five sites: high nutrient basalt sites at Lamins Hill (B1) and Longlands Gap (B2), and low nutrient sites on granite at Mt Nomico (G), on rhyolite 

at Longlands Gap (R) and on metamorphic substrate near Cape Tribulation (M) 

Concentration of cyanogenic glycosides in different plant parts (mg CN g 1 d. wt) 

Form Species Family Forest Site Leaf Stem Floral parts Fruit/seed 

H Alocasia brisbanensis 

(F.M.Bailey) Domin* 

Araceae U B1 – – – – 

T Beilschmiedia collina B.Hyland Lauraceae U B1, B2, G, R Old: 28.4–1263 (n = 36) – – – 

Yg: 1039–1391 

T Brombya platynema F.Muell. Rutaceae L M 156.4–1285 (n = 20) – – – 

0–10.5 (n = 27) 

T Cardwellia sublimis F.Muell. Proteaceae U, L B1, B1, G, R, M Old:11.5–69.8 (n = 21) – Bud: 779–851 Seed: approx.8 

Yg: 733.1, tips: 219.1 Flwr: 130–334 

T Cleistanthus myrianthus 

Euphorbiaceae L M Old: 1.1–17.3 (n = 41) – Flwr: 30.7 Mature: 160–377 

(Hassk.) Kurz 

Yg: 78.4–80.4 (n = 3) Immature: 821 

T Clerodendrum grayi Munir Verbenaceae U B1, B2, G Old: 1325–4800 (n = 6) – Bud: 733 – 

Flwr 440–942 

T Elaeocarpus sericopetalus F.Muell. Elaeocarpaceae U G, R 1100–5056 (n = 10) Sdlg: 1739–2679 – 1028 

Tree: 135 

V Embelia grayi S.T.Reynolds Myrsinaceae U B1, B2 10–81 (n = 3) – – – 

V Flagellaria indica L. Flagellariaceae U, L B1, G, M 11–177.3 (n = 6) – – – 

T Helicia australasica F.Muell. Proteaceae L M 42.2–155.2 (n = 8) † – – – 

T Helicia blakei Foreman Proteaceae U B1 Mean: 17.9 (n = 2) – – – 

T Helicia nortoniana 

(F.M.Bailey) F.M.Bailey* 

Proteaceae L M – – – – 

V Passiflora sp. (Kuranda BH12896) Passifloraceae L M 313–922 (n = 4) – – – 

T Mischocarpus exangulatus 

Sapindaceae U B1 Old: 133; yg: 222 (n = 1) – – – 

(F.Muell.) Radlk. 

T Mischocarpus grandissimus 

(F.Muell.) Radlk. 

Sapindaceae U, L G, M 49–680 (n = 2) 

– – – 

761–2006 (n = 4) z 

T Opisthiolepis heterophylla L.S.Sm. Proteaceae U B1, B2 Old: 10–208 (n = 7) – – – 

Yg: 2000 

V Parsonsia latifolia (Benth.) S.T.Blake 

Apocynaceae U B1, G, R 765–4835 (n = 3) – – – 

T Polyscias australiana 

(F.Muell.) Philipson 

T Prunus turneriana 

(F.M.Bailey) Kalkman 

T Ryparosa javanica (Blume) 

Kurz ex Koord. & Valeton** 

Araliaceae U, L B1, B2, G, R, M Old:

leaves of numerous individuals of Polyscias 

australiana tested negative for cyanide, and contained 

800 mgCNg 1 d. wt; Table 1). Overall, few tests were 

made of seeds or other reproductive tissues as part of the 

survey; however, where reproductive tissues were tested 

from species with cyanogenic leaves, they were typically 

cyanogenic and contained higher concentrations of cyanogenic 

glycosides, although the concentration varied with 

the maturity of the fruit/seed. For example, the immature 

seed and fruit (combined) from Prunus turneriana had 

>8mgCNg 1 d. wt compared with mature seed alone 

(

R. javanica occurs in a limited area within lowland rainforest 

north of the Daintree River. Cyanogenesis is reported 

for the first time in the genera Beilschmiedia, Cardwellia, 

Cleistanthus, Elaeocarpus, Embelia, Mischocarpus, 

Opisthiolepis, Parsonsia and Polyscias. The number of 

new reports at the generic level may in part reflect the 

high level of endemism among the Australian tropical rainforest 

flora. In addition, several species are from families in 

which cyanogenesis has been rarely reported, if at all. For 

example, cyanogenesis is rare in Elaeocarpaceae, 

Lauraceae, Apocynaceae, Myrsinaceae and Araliaceae 

families. Following are descriptions of each cyanogenic 

species (listed alphabetically by family) discussed in relation 

to previous reports of cyanogenesis within the relevant 

taxonomic groups. 

Ryparosa javanica (Blume) Kurz ex Koord. & 

Valeton (Achariaceae) 

Ryparosa javanica is currently the subject of taxonomic 

revision (Webber, 2005). The Queensland Ryparosa sp. 

currently known as R. javanica is endemic to lowland rainforest 

north of the Daintree River to Cape Tribulation, north 

east Queensland. This species was found to be cyanogenic 

early in this survey, and has subsequently been the focus 

of extensive population-level studies of cyanogenesis 

(Webber, 2005). Reports of cyanogenesis in this genus 

are common; for example, R. javanica (sensu stricto) and 

R. caesia are cyanogenic (Rosenthaler, 1919). Cyanogenesis 

has been reported among other genera in Achariaceae, 

e.g. Hydnocarpus, Calancoba, Ceratiosicyos, Gynocardia, 

Erythrospermum, Pangium and Kiggelaria (Rosenthaler, 

1919; Tjon Sie Fat, 1979a; Jensen and Nielsen, 1986). 

Many of these genera were previously in the Flacourtiaceae 

until recent revisions saw most cyanogenic genera assigned 

to the Achariaceae (Chase et al., 2002), including the tribe 

Pangieae of which Ryparosa is a member. Cyanogenesis 

was considered a useful taxonomic marker in Flacourtiaceae 

(Spencer and Seigler, 1985), this utility being apparent in 

the revision of the family (Chase et al., 2002). All individuals 

in sizeable populations of R. javanica were cyanogenic, 

and the cyanogenic glycoside in R. javanica has been 

identified as gynocardin (Webber, 2005), a cyclopentenoid 

cyanogenic glycoside typical of Achariaceae (Jaroszewski 

and Olafsdottir, 1987). Cyanogenic glycosides derived 

from valine/isoleucine have also been reported in 

species formerly in the Flacourtiaceae (Lechtenberg and 

Nahrstedt, 1999). 

Parsonsia latifolia (Benth.) S.T.Blake (Apocynaceae) 

This is the first report of cyanogenesis in the genus 

Parsonsia, a genus of woody or semi-woody climbers 

(130 species) distributed from Southeast Asia to Australia, 

New Caledonia and New Zealand. Members of the Apocynaceae 

family (220 genera, 2000 species) commonly have 

clear or milky latex, and frequently contain alkaloids 

(Mabberley, 1990; Collins et al., 1990). Parsonia latifolia 

(diameter up to 9 cm) has milky white latex, and is endemic 

to Australia (Forster and Williams, 1996). It is found in 

lowland and highland rainforest in noth east Queensland, 

parts of New South Wales and the Northern Territory 

(Hyland et al., 2003). Reports of cyanogenesis in Apocynaceae 

and the order Gentianales are few; Gibbs (1974), who 

reported negative results for Parsonsia eucalyptifolia and 

P. lanceolata, found only Alstonia scholaris (L.) R.Br. to be 

cyanogenic, and noted positive reports for four other 

species. Tests for cyanogenesis in the Apocynaceae are, 

however, apparently limited, with negative reports for 

only approx. 20 species (Gibbs, 1974; Adsersen et al., 

1988; Thomsen and Brimer, 1997). In this study, leaf 

samples of all ages from multiple A. scholaris trees and 

seedlings gave negative FA paper results; quantitative 

assaying confirmed the absence of cyanogenesis. No cyanogenic 

constituents in the family have been characterized. 

Polyscias australiana (F.Muell.) Philipson (Araliaceae) 


Polyscias, a genus of approx. 100 species distributed 

throughout Africa, Asia, Malesia, Australia and the Pacific 

Islands. In addition, cyanogenesis is very rare in the order 

Umbellales; Gibbs (1974) reported only negative results or 

some doubtful positive results in a few members of 

Araliaceae (Nothopanax sp. and Schefflera sp.) and the 

Umbelliferae. Subsequently, only Aralia spinosa L. 

(above-ground parts) has been reported as cyanogenic 

(Seigler, 1976b). Polyscias australiana is often considered 

a regrowth species in disturbed rainforest, and commonly 

occurs at rainforest margins. Concentrations of cyanogenic 

glycosides in this species were variable, and in mature 

leaves were commonly less than the threshold value used 

for classifying individuals as cyanogenic in this study 

(

considered common (Gibbs, 1974; Hegnauer, 1990; 

Mabberley, 1990). In the Elaeocarpaceae family, cyanogenesis 

has previously been reported in only two species: the 

leaves of Vallea stipularis ‘pyrifolia’ F.Ballard (Gibbs 

1974), and the leaves [Greshoff (1898) cited in Hegnauer 

(1973)] and the bark (Pammel, 1911; Rosenthaler, 1919) of 

Sloanea sigun (Blume) K.Schum (syn. Echinocarpus 

sigun), were found to be cyanogenic. Sambunigrin was 

isolated from the leaves of S. sigun [R. Hegnauer and 

L. H. Fikenscher, unpubl. data (1983) cited in Hegnauer 

(1990)], the only previous report of a cyanogenic constituent 

in Elaeocarpaceae and Malvales. Characterization of the 

principal foliar cyanogenic glycoside—an apparently 

unusual phenylalanine-derived glycoside with an organic 

acid residue—is ongoing (Miller, 2004). 

Cleistanthus myrianthus (Hassk.) Kurz (Euphorbiaceae) 

This appears to be the first report of cyanogenesis in 

the genus Cleistanthus, which consists of 100–140 species. 

Cleistanthus myrianthus is a subcanopy rainforest tree (to 

7 m), found in the lowland and foothills from the Daintree 

River to Rossville, north east Queensland (Cooper and 

Cooper, 1994). In Australia, it is classified as rare on the 

basis of this limited distribution, but it is also found in 

Southeast Asia and Malesia (Hyland et al., 2003). Cyanogenesis 

is especially common in Euphorbiaceae (300 

genera, 7500 species), and is found in many genera including 

Bridelia, Euphorbia and Phyllanthus, as well as the 

economically important species Hevea brasiliensis (rubber 

tree) and Manihot esculenta (cassava) (e.g. Rosenthaler, 

1919; Tjon Sie Fat, 1979a; Seigler et al., 1979; Adsersen 

et al., 1988). 

The chemotaxonomic utility of cyanogenesis, as well 

as other secondary metabolites (e.g. alkaloids and terpenes), 

has been demonstrated in Euphorbiaceae (Seigler, 1994). 

Cyanogenic glycosides are useful at the infra-familial 

level in the Euphorbiaceae (van Valen, 1978; Seigler, 

1994); the species in the subfamily Phyllanthoideae typically 

contain the tyrosine-derived cyanogenic glycosides 

dhurrin, taxiphyllin or triglochinin, while species in the 

subfamily Crotonoideae (sensu Pax and Hoffman, 1931; 

see van Valen, 1978), including Hevea and Manihot, produce 

cyanogenic glycosides derived from valine and isoleucine 

(e.g. linamarin and lotaustralin) (van Valen, 1978, 

Nahrstedt, 1987; Seigler, 1994). Given this pattern, one may 

predict Cleistanthus—assigned to the Phyllanthoideae—to 

contain cyanogens derived from tyrosine. 

Flagellaria indica L. (Flagellariaceae) 

Flagellaria indica (‘the supplejack’), a leaf tendril climber 

with cane-like stems, is known to be cyanogenic (Petrie, 

1912; Webb, 1948; Gibbs, 1974; Morley and Toelken, 

1983). Young shoots were suspected of poisoning stock 

in Australia (Everist, 1981), and it has also been reported 

to have medicinal properties (e.g. tumour inhibition; Collins 

et al., 1990). In other countries, it has been used as a hair 

wash, and as a contraceptive (see Hyland et al., 2003), 

but was not apparently used medicinally by aboriginal 

Australians (Morley and Toelken, 1983). Interestingly, 

monocotyledons are typically characterized by cyanogenic 

glycosides biosynthetically derived from tyrosine 

(Lechtenberg and Nahrstedt, 1999). Consistent with this, 

the tyrosine-derived cyanogenic glycoside triglochinin 

was isolated from the stem (and rhizome) of F. indica 

(L. H. Fikenscher unpubl. data, cited in Hegnauer, 1966), 

although meta-hydroxylated valine or isoleucine, and 

leucine-derived glycosides have also been found in 

monocotyledons (Nahrstedt, 1987; Lechtenberg and 

Nahrstedt, 1999). 

Clerodendrum grayi Munir (Lamiaceae) 

Clerodendrum grayi is a rare subcanopy tree endemic to 

the northern part of Queensland, Australia (Munir, 1989). 

Recent revisions of the division between Lamiaceae and 

Verbenaceae families (Cantino, 1992; Wagstaff et al., 

1997, 1998) transferred the genus Clerodendrum, and others 

historically in the Verbenaceae family, to the Lamiaceae 

family. Cyanogenesis in Lamiaceae, and also Verbenaceae, 

has rarely been reported. Even within the order Lamiales, 

cyanogenesis is considered rare (Gibbs, 1974). In the 

Lamiaceae, known for its culinary and medicinal herbs 

[e.g. Lavandula (lavender); Mentha (mint)], typical constituents 

are monoterpenoids, diterpenes or triterpenes, as 

well as flavonoids and iridoid glycosides (Gibbs, 1974; 

Hegnauer, 1989; Taskova et al., 1997). In a survey of the 

flora of the Galapagos Islands, Clerodendrum molle var. 

molle was found to be cyanogenic (Adsersen et al., 1988; 

see also Gibbs, 1974, Tjon sie Fat, 1979a). In addition, 

several species of Clerodendrum are known to be toxic 

(Hurst, 1942; Webb, 1948; CFSAN, 2003); however, the 

poison is not detailed. 

In this study, the extremely high foliar concentrations of 

cyanogenic compounds—up to 48mgCNg 1 d. wt in 

mature field-grown tree leaves—are among the highest 

reported for tree leaves (Table 1). Two cyanogenic glycosides 

were purified from the leaf tissue of C. grayi (Miller 

et al., 2006a). Prunasin and its primerveroside, the rare 

diglycoside lucumin (Eyjólfsson, 1971), were found in 

the ratio 1 : 158 (mol:mol) (Miller et al., 2006a), the first 

reported co-occurrence of these glycosides, and the first 

confirmed report of lucumin in vegetative tissue (see 

Thomsen and Brimer, 1997). Given the relatively rarity 

of reports of cyanogenic glycosides from the Lamiaceae, 

and even within the order Lamiales, it is difficult to draw 

any conclusions about the biogenetic origins of glycosides 

within these taxonomic groups. Refer to Miller et al. 

(2006a) for a detailed discussion of cyanogenesis in 

C. grayi and associated taxa. 

Beilschmiedia collina B.Hyland (Lauraceae) 

Beilschmiedia collina (‘the mountain blush walnut’) is a 

tree species endemic to Queensland rainforest (Cooper and 

Cooper, 1994). Cyanogenesis is very rare in Lauraceae 

(Gibbs, 1974; Hegnauer, 1989), having only been reported 

from Cinnamomum camphora and Litsea glutinosa (Gibbs, 

1974), with one other species (Nectranda megapotamica) 

reported to have cyanogenic glycosides but apparently 

lacks the catabolic enzymes, requiring further investigation

(Francisco and Pinotti, 2000). The Lauraceae is better 

known for producing a range of alkaloids (e.g. Gibbs, 

1974), and as a dominant family in the rainforest flora of 

Australia has been much studied for its toxic and potentially 

medicinal alkaloids (Webb, 1949, 1952; Collins et al., 1990). 

Of 39 species tested in the Lauraceae family in this survey, 

only B. collina was cyanogenic (Appendix). Given the 

apparent rarity of cyanogenesis in the family, and even 

the order Laurales, it is difficult to speculate as to the 

biosynthetic precursor of the cyanogenic constituent in 

B. collina. To the authors’ knowledge, only the tyrosinederived 

cyanogenic glycoside taxiphyllin has been reported 

in the Calycanthaceae family within the Laurales 

(Lechtenberg and Nahrstedt, 1999); tyrosine-derived 

glycosides are also found in Ranunculaceae in the order 

Ranunculales, which is in the same subclass Magnoliidae 

(sensu Cronquist, 1981) as Laurales. 

Embelia grayi S.T. Reynolds (Myrsinaceae) 


Embelia—a genus of approx. 130 species—and among 

the first in the family Myrsinaceae. Furthermore, Gibbs 

(1974) considered cyanogenesis to be unknown in the 

order Primulales. Subsequent to Gibbs (1974), only two 

reports of cyanogenesis in Myrsinaceae could be found— 

for Rapanea parviflora (Kaplan et al., 1983), and for 

R. umbellata, which needs confirmation as cyanogenesis 

was only detected after 24 h of tissue incubation 

(Francisco and Pinotti, 2000). Overall, there are even a 

few negative test records for the family. Embelia grayi is 

a vine with diameter up to 9 cm, endemic to upland and 

highland rainforest in north east Queensland. Embelia 

caulialata, and three Rapanea spp., also tested here, were 

not cyanogenic. The order Primulales (sensu Cronquist, 

1981) is in the subclass Dilleniidae which also includes 

the orders Malvales and Violales. Within the Violales, 

cyanogenesis is common in the Achariaceae (including 

Flacourtiaceae), Passifloraceae and Turneraceae families, 

which tend to contain cyanogenic glycosides of the cyclopentanoid 

series (Lechtenberg and Nahrstedt, 1999). 

Passiflora sp. (Kuranda BH12896) (Passifloraceae) 

Passiflora sp. (Kuranda BH12896) was the only species 

in the Passifloraceae tested in this study. It is a vine found 

in the lowland rainforests of north east Queensland; all 

Australian members of the Passifloraceae are climbers or 

sprawling shrubs (Morley and Toelken, 1983). Cyanogenesis 

is common within the family Passifloraceae (600 

species, two genera worldwide), and the genus Passiflora 

(e.g. Rosenthaler, 1919; Tjon Sie Fat, 1979a; Adsersen et al., 

1988; Olafsdottir et al., 1989). Several Australian Passiflora 

spp. were reported to be cyanogenic, including P. aurantia 

(Petrie, 1912; Smith and White, 1918; Webb, 1952; Gardner 

and Bennetts, 1956) and the endemic P. herbertiana Lindl. 

(Petrie, 1912), and implicated in poisoning stock (Smith 

and White, 1918). Within Passifloraceae, cyanogenesis 

has also been reported in Adenia spp. and Ophiocaulon 

spp. (Rosenthaler, 1919; Tjon Sie Fat, 1979a). The specific 

cyanogenic constituents may be taxonomically diagnostic at 

the infrafamilial level (e.g. occurrence of the rare glycoside 

passibiflorin; Adsersen et al., 1993). Cyanogenic Passifloraceae 

typically contain cyanogenic glycosides with 

cyclopentenoid ring structures (Seigler et al., 1982; 

Spencer and Seigler, 1985; Nahrstedt, 1987; Lechtenberg 

and Nahrstedt, 1999), although phenylalanine-derived glycosides 

(e.g. prunasin) have been isolated from Passiflora 

edulis (Spencer and Seigler, 1983; Chassagne et al., 1996; 

Seigler et al., 2002), and valine/isoleucine-derived glycosides 

(e.g. linamarin, lotaustralin) from several Passiflora 

spp. in the subgenus Plectostemma (Spencer et al., 1986). 

Cyanogenesis in the Proteaceae family 

Five of the 20 species tested in the Proteaceae family 

were found to be cyanogenic: C. sublimis F.Muell., 

O. heterophylla L.S.Sm., Helicia australasica F.Muell, 

H. blakei Foreman and H. nortoniana (F.M.Bailey) 

F.M.Bailey. The latter species was opportunistically tested 

as it was found only outside the plots. This is the first 

formal report of cyanogenesis in the monospecific genera 

Cardwellia and Opisthiolepis, while cyanogenesis has previously 

been reported in the genus Helicia (H. robusta; 

Gibbs, 1974). 

Cardwellia sublimis (‘the northern silky oak’) is the only 

species in the tribe Cardwelliinae (Hoot and Douglas, 1998). 

This canopy species (to 30 m), an important timber tree, is 

endemic to north east Queensland, being widely distributed 

throughout well-developed lowland to highland rainforest 

(Hyland et al., 2003). Cardwellia sublimis was common to 

all sites in this study. 

Opisthiolepis heterophylla (‘the blush silky oak’) is 

endemic and confined to north east Queensland, from the 

Kirrama Range to Cooktown. It grows to 30 m in lowland 

to highland rainforest, but is most common in upland and 

highland rainforest on the Atherton Tableland (Hyland et al., 

2003). The flowers of O. heterophylla have been found to 

be cyanogenic (E. E. Conn, University of California, Davis, 

CA, USA, pers. comm.). 

The genus Helicia (approx. 90 species) occurs throughout 

Asia and the Pacific, with nine species found naturally 

in Australia (Foreman, 1995). Helicia blakei (‘Blake’s silky 

oak’) is endemic to north east Queensland, occurring as an 

understorey tree in well-developed upland rainforest 

(Hyland et al., 2003). Helicia australasica is a shrub or 

tree (3–20 m) widespread throughout northern Australian 

through to Papua New Guinea. It occurs as an understorey 

tree in well-developed rainforest, monsoon forest and dry 

rainforest (Foreman, 1995; Hyland et al., 2003). The fruit is 

known to be eaten by aborigines (Foreman, 1995). Helicia 

nortoniana is also an understorey tree (to 20 m), endemic 

to north east Queensland, and found in well-developed 

lowland and highland rainforest (Cooper and Cooper, 

1994; Foreman, 1995; Hyland et al., 2003). These data 

support the preliminary results of tests on dried herbarium 

samples where only some samples of H. australasica and 

H. nortoniana tested positive for cyanide (E. E. Conn, 

University of California, Davis, CA, USA, pers. comm.). 

Tests of dried herbarium samples of H. blakei, however, 

gave only negative results (E. E. Conn, University of

California, Davis, CA, USA, pers. comm.). The inconsistent 

findings by Conn are probably the result of using dried 

herbarium material. 

Cyanogenesis is considered especially common in 

the Proteaceae (Swenson et al., 1989; Lechtenberg and 

Nahrstedt, 1999), known in a range of genera, but particularly 

in Grevillea and Hakea spp. In Australia, cyanogenic 

members of the Proteaceae have been implicated in 

stock poisoning (Gardner and Bennetts, 1956). Based on the 

reports of Gibbs (1974) and Tjon Sie Fat (1979a), and the 

study of Swenson et al. (1989) who found 44 of 155 proteaceous 

species tested to be cyanogenic, cyanogenesis 

is most widespread in the subfamily Grevilleoideae. For 

example, cyanogenesis has been reported in the genera 

Stenocarpus, Lomatia, Helicia, Xylomelum, Telopea, 

Macadamia, Hicksbeachia, Lambertia, Grevillea and 

Xylomelum (Petrie, 1912; Smith and White, 1918; Hurst, 

1942; Gardner and Bennetts, 1956; Gibbs, 1974; Swenson 

et al., 1989; Lamont, 1993; E. E. Conn, University of 

California, Davis, CA, USA, pers. comm.), all of which 

are in the Grevilloiedeae (Hoot and Douglas, 1998). Consistent 

with this pattern, the cyanogenic genera reported 

here are all in the subfamily Grevilleoideae; Cardwellia 

in the subtribe Cardwelliinae (tribe Knightieae), Opisthiolepis 

in the subtribe Buckinghamiinae (tribe Embothrieae), 

and Helicia in the subtribe Heliciinae (tribe Heliceae). In 

contrast, there are only a few reports of cyanogenesis within 

the Proteoideae subfamily; cyanogenesis was only reported 

in single species of Conospermum, Petrophile and Protea 

(Gibbs, 1974; Swenson et al., 1989; E. E. Conn, University 

of California, Davis, CA, USA, pers. comm.). 

The cyanogenic constituents, which have been identified 

in comparatively few species, appear to be biogenetically 

derived from tyrosine. Swenson et al. (1989) identified 

the cyanogenic glycosides in eight species; leaves and 

flowers of several Hakea, Leucadendron, Grevillea 

and Macadamia species were found to contain dhurrin 

and proteacin (see also Plouvier, 1964; Young and 

Hamilton, 1966). In this regard, the identity of the cyanogenic 

constituent in the monospecific genera in particular 

would be interesting. 

Prunus turneriana (F.M.Bailey) Kalkman (Rosaceae) 

Cyanogenesis is widespread within Rosaceae (Hegnauer, 

1990), and has been much studied in the subfamily 

Prunoideae in particular, as it contains many cyanogenic 

cultivated species [e.g. Prunus domestica L. (plum); 

Armeniaca vulgaris Lam. (apricot)]. Cyanogenic Rosaceae 

(in particular Prunus spp.) are also a common source of 

poisoning in domestic animals (e.g. Poulton, 1983; 

Schuster and James, 1988). Prunus turneriana is one of 

only two Prunus species native to Australia and is a late 

successional canopy tree species in the lowland and upland 

rainforests of far north Queensland, Australia. The fruits of 

this canopy species are known to be toxic, yet are also eaten 

by cassowaries, fruit pigeons, Herbert river ringtail possums 

and musky-rat kangaroos (Cooper and Cooper, 1994). 

The flesh of the fruit was used raw by the Ngadjonji 

people—the original inhabitants of the rainforests on 

the Atherton Tablelands, north Queensland—for treating 

toothache, while the toxic kernels were processed for a 

starchy food (Huxley, 2003). Despite its common name 

‘Almond bark’ and phytochemical surveys of Australian 

rainforest species (e.g. Webb, 1949), cyanogenesis in P. 

turneriana had not previously been reported. Cyanogenic 

glycosides were found to be distributed throughout all tissues 

and, consistent with other species in the Rosaceae, are 

biosynthetically derived from the amino acid phenylalanine 

(Møller and Seigler, 1999). Prunasin was identified as the 

major cyanogen in leaf, stem, root and seed tissues of 

P. turneriana, and amygdalin was restricted to the seed. 

What was unusual about P. turneriana was the presence 

of significant amounts of the (R)-prunasin epimer, (S)- 

sambunigrin, in leaf, stem and seed tissue, whereas root 

tissue contained only prunasin. Refer to Miller et al. 

(2004) for the detailed characterization of cyanogenesis 

in P. turneriana. 

Brombya platynema F.Muell. (Rutaceae) 


Brombya, which is a genus of 1–2 species endemic to 

Australia (Hyland et al., 2003). Brombya platynema is 

endemic to north east Queensland where it occurs as an 

understorey tree in well-developed forest (from sea level 

to 1100 m a.s.l.) (Hyland et al., 2003). The family (150 

genera, 1500 species) includes many strongly scented 

shrubs and trees, and rutaceous species are known for 

their terpenoids and alkaloids (Price, 1963; Gibbs, 1974; 

Everist, 1981). Cyanogenesis is rare in Rutaceae, and has 

only been reported in Boronia bipinnate Lindl. (leaves; 

Rosenthaler, 1919), Zieria spp. (Hurst, 1942; Gibbs, 1974; 

Fikenscher and Hegnauer, 1977), Zanthoxylum fagara 

(Adsersen et al., 1988) and Loureira cochinchinensis 

Meissa (Gibbs, 1974). Even within the order Rutales, 

cyanogenesis is rare, with only a few additional definitive 

reports of cyanogenesis in the Tremandraceae family 

(Gibbs, 1974). The phenylalanine-derived cyanogenic glycosides 

prunasin/sambunigrin and zierin have been isolated 

from leaves of two Zieria spp. (Finnemore and Cooper, 

1936; Fikenscher and Hegnauer, 1977). The rare metahydroxylated 

cyanogenic glycoside holocalin was recently 

identified as the principal cyanogen in leaves of 

B. platynema; traces of prunasin and amygdalin were 

also detected (Miller et al., 2006b). These data suggest 

the possibility that species in this family have cyanogenic 

glycosides biosynthetically derived from the amino acid 

phenylalanine. 

Mischocarpus grandissimus (F.Muell.) Radlk. and 

Mischocarpus exangulatus (F.Muell.) Radlk. (Sapindaceae) 

Two of the four species of Mischocarpus tested in this 

study were found to be cyanogenic—M. grandissimus and 

M. exangulatus—representing the first reports of cyanogenesis 

for the genus. Mischocarpus is a genus of 15 species 

found in Asia, Malesia and Australia; nine species occur 

naturally in Australia (Hyland et al., 2003). Both species 

are endemic to Queensland; M. grandissimus is restricted to 

north east Queensland, while M. exangulatus is also found 

on the Cape York Peninsula, Queensland. Mischocarpus

grandissimus occurs as an understorey tree in welldeveloped 

lowland and upland rainforest (sea level to 

750 m a.s.l.; Hyland et al., 2003). Similarly, M. exangulatus 

(‘the red bell mischocarp’) is an understorey tree to 15 m 

found in well-developed lowland and highland rainforest 

(sea level to 1100 m a.s.l.; Cooper and Cooper, 1994; 

Hyland et al., 2003). These were the only cyanogenic 

species identified among the 29 species in the Sapindaceae 

family tested in this study (Appendix). Cyanogenesis 

is known in the Sapindaceae family; however, the 

family is best known for cyanolipids in the seed oils of 

numerous species (e.g. Alectryon spp., Allophylus spp., 

Cardiospermum spp., Sapindus spp., Paullinia spp. and 

Ungnadia speciosa) (Mikolajczak et al., 1970; Seigler 

et al., 1971; Gowrikumar et al., 1976; Seigler and 

Kawahara, 1976). There are only a few reports of 

cyanogenesis in Australian indigenous members 

of Sapindaceae—for Dodonaea spp. (Hurst, 1942; Webb, 

1949) and Alectryon spp. (Smith and White 1918; 

Finnemore and Cooper, 1938)—and, with the exception 

of Heterodendrum oleifolium Desf. [syn. Alectryon oleifolius 

(Desf.) S.Reyn], the cyanogenic constituents in these 

species have not been characterized. In addition to cyanolipids 

in the seeds, leucine-derived cyanogenic glycosides 

have been characterized from vegetative parts of sapindaceous 

species (Seigler et al., 1974; Hübel and 

Nahrstedt, 1975, 1979; Nahrstedt, and Hübel, 1978). 

Further findings 

Alocasia brisbanensis (Araceae) was also found to be 

cyanogenic, consistent with reports of cyanogenesis in 

numerous congeneric species (Rosenthaler, 1919; Tjon 

Sie Fat, 1979a). However, as a herb, it was not included 

in the analysis. The tyrosine-derived cyanogenic glycoside 

triglochinin has been isolated from the closely related 

A. macrorrhiza (Nahrstedt, 1975). 

There were several findings which contradicted previous 

reports for species. In addition to negative results for 

A. scholaris noted above, Eupomatia laurina (Eupomatiaceae) 

was reported to be ‘doubtfully cyanogenic’ and 

Cananga odorata (Annonaceae) cyanogenic by Gibbs 

(1974), but neither species was found to be cyanogenic 

in this study (based on repeated tests of four and two individuals, 

respectively). Similarly, reports of cyanogenesis in 

fruit and leaves of the Australian endemic Davidson’s plum 

(Davidsonia pruriens) (Petrie, 1912; Rosenthaler, 1929) 

were not corroborated, both mature leaves and fruit testing 

negative in this study. Gibbs (1974) also only found negative 

results for leaves of D. pruriens. In addition, negative 

test results for cyanogenesis in this study corroborate previous 

negative reports for Neolitsea dealbata (syn. Litsea 

dealbata), Cayratia acris, Morinda jasminoides and Sarcopetalum 

harveyanum (Gibbs, 1974; Appendix). A screening 

of Australian Proteaceae family, conducted primarily using 

dry herbarium material, was carried out by E. E. Conn 

(University of California, Davis, CA, USA, pers. comm.) 

and included several species tested in this study. As noted 

above, Conn had some inconsistent, and possibly therefore 

inconclusive, results for a number of species, which were 

confirmed by fresh sampling in this study. One of eight 

samples of Musgravea heterophylla gave a positive reaction 

after 12 h in Conn’s survey, a species which was not found 

to be cyanogenic here. Further testing of M. heterophylla 

is warranted. As far as could be discerned, the majority of 

species tested in this study had not previously been tested, 

despite numerous screenings of Australian and Queensland 

plants, most notably by Webb (1948, 1949). 

General findings: frequency of cyanogenesis 

Of 401 species from 87 families tested in the survey, 

18 (45 %) species from 13 families were cyanogenic. 

The proportion of cyanogenic species at the sites ranged 

from 47to65 %, values similar to the frequency of cyanogenic 

species found in a substantial survey of woody species 

in Costa Rican rainforest (Thomsen and Brimer, 1997)—the 

only previous study to report the frequency of cyanogenesis 

standardized with respect to plant size (dbh >10 cm) and 

forest area (7 · 1 ha plots). Overall, Thomsen and Brimer 

(1997) found that 40 % (range 21–57 % for plots) of 

401 species from 68 families were cyanogenic, and that 

cyanogenic stems (dbh > 5 cm) accounted for 3 % of 

total basal area (range 16–51 %). Here, the overall proportion 

of total basal area in cyanogenic stems was 73 % and 

ranged from 12 to134 %. The highest proportions were in 

highland rainforest on basalt soil (134 %) and in lowland 

rainforest (116 %). Highland rainforest on rhyolite had the 

lowest proportion. 

Overall, at a community level, there are few studies with 

which to compare frequencies of cyanogenesis reported 

here for tropical rainforest in north east Queensland. In 

the first instance, there have been few studies in tropical 

systems, but also, the screening methodology varies 

depending on the research question being addressed, be it 

taxonomic (e.g. examining chemical differences in relation 

to proposed phylogenetic relationships) or ecological 

(e.g. the role of secondary compounds in plant–animal interactions). 

For example, while the survey of Thomsen and 

Brimer (1997) was standardized with respect to plant size 

and forest area, the few surveys in other tropical systems 

have focused on plant–animal interactions, and have not 

screened in a standardized fashion. Only 23 % (one species) 

was found to be cyanogenic in the screening of >90 % of the 

flora (n = 43 species) in a species-poor seasonal cloud forest 

in India (Mali and Borges, 2003). In contrast, in a survey 

examining the frequency of cyanogenesis in relation to 

environmental factors and insect density along a transect 

from the shoreline to an inland lagoon in a neotropical 

woodland (‘restinga’), Kaplan et al. (1983) found 25 species 

(23 %) of 108 species screened to be cyanogenic. They also 

reported variable test results for a further 49 species (for n = 

2–16 individuals), elevating the proportion of cyanogenic 

species to 68 %, a value which requires further examination 

before interpretation, as the sampling strategy (e.g. plant 

size, random sampling strategy, life form and transect area) 

was unclear, and some uncertainty with regard to picrate 

paper test results was expressed by the authors (Kaplan 

et al., 1983). 

Adsersen et al. (1988) compared frequencies of cyanogenesis 

within the endemic and non-endemic flora of the 

Galapagos Islands, two floras subject to different suites of

herbivores over a period of time. They screened fresh and 

herbarium specimens of a significant proportion (65 %) of 

the flora from the Archipelago, and reported 81% of 

endemic species, and 53 % of native species—those 

which also occur on the South American mainland—to 

be cyanogenic. Interestingly, they also reported a further 

22 % of native species and 30 % of endemic species to 

release cyanide in the presence of a crude mix of b- 

glycosidases (including 5 % b-glucuronidase from snails), 

suggesting that a large number of species contain cyanogenic 

glycosides but lack the catabolic b-glycoside enzyme. 

This contrasts with the findings of several other studies 

where the addition of non-specific b-glycosidases or 

pectinase during qualitative testing did not alter the frequency 

of positive results (e.g. Petrie, 1912; Thomsen 

and Brimer, 1997; Buhrmester et al., 2000; Lewis and 

Zona, 2000; but see Conn et al., 1985). Similarly, in this 

study, all samples were spontaneously cyanogenic without 

the addition of pectinase from Rhizopus spp., indicating 

that non-cyanogenic individuals probably lacked both the 

cyanogenic glycoside and b-glycosidase, or possibly that 

pectinase was not able to catalyse the cyanogenesis in these 

species. It is perhaps noteworthy that the greater frequency 

of cyanogenesis reported by Conn et al. (1985) in response 

to the addition of b-glycosidase (emulsin) was in a survey 

solely of the genus Acacia, indicating that such a response 

may vary among taxa. 

It is important to note that the frequencies reported in 

all of these surveys were apparently based on the testing of 

single specimens of the vast majority of species (Adsersen 

et al., 1988; Thomsen and Brimer, 1997; Mali and Borges, 

2003). Similarly, while the present study aimed to test at 

least three individuals of each species in duplicate (i.e. with 

and without added enzyme), only one individual of many 

species was encountered (Appendix). Furthermore most 

species here were also tested in both wet and dry seasons. 

A range of studies report variable positive and negative test 

results among sample sizes as small as n = 2 (e.g. Kaplan 

et al., 1983; Thomsen and Brimer, 1997). Given such polymorphism 

for cyanogenesis (see also Aikman et al., 1996), 

the reported frequencies in these surveys may underestimate 

overall the actual proportion of cyanogenic species in plant 

communities. The reported frequency of cyanogenesis may 

also vary with the plant part tested. In the Costa Rican study, 

Thomsen and Brimer (1997) reported a greater frequency of 

cyanogenesis among reproductive plant parts than leaves, 

as did Buhrmester et al. (2000) in populations of Sambucus 

canadensis (elderberry) in Illinois. Consistent with that 

trend, individuals of species with weakly cyanogenic leaves, 

including some which produced negative FA paper results, 

had higher concentrations of glycosides in flowers or fruits; 

however, overall, only a small number of reproductive 

tissues were tested, so limited comparison can be drawn 

(Table 1). 

The frequency of cyanogenesis varies between 

taxonomic groups and with life form. Cyanogenesis is considered 

especially common in some plant families (e.g. 

Rosaceae, Euphorbiaceae, Passifloraceae and Proteaceae; 

Lechtenberg and Nahrstedt, 1999), and rare or absent in 

others (e.g. Lauraceae and Araliaceae; Gibbs, 1974; 

Hegnauer, 1989), a trend which may in part reflect differential 

intensity of testing among taxonomic groups. In this 

study, the frequency of cyanogenesis among the dominant 

plant families varied. In the Proteaceae family, five of 

20 (25 %) species were cyanogenic, while two of 29 

(69 %) species in the Sapindaceae, and one of 39 (25%) 

species in the Lauraceae were cyanogenic (Table 1). In a 

screening of Australian Acacia, 69 % of 360 species were 

cyanogenic (Conn et al., 1985). Cyanogenesis appears to be 

rare among palms; only two species (12 %) of 155 species 

of palms (108 genera) were found to be cyanogenic 

(Lewis and Zona, 2000). No cyanogenic palm species 

were identified in this study. 

Concentrations of cyanogenic glycosides 

Several of the 18 cyanogenic species detected in this 

study contained concentrations of cyanogenic glycosides 

among the highest reported for leaves of woody species. 

Most notably, tree species E. sericopetalus, C. grayi and P. 

turneriana had foliar concentrations of cyanogenic glycosides 

up to 52, 49 and 48mgCNg –1 d. wt, respectively 

(Table 1). Similarly, Webber (1999) recorded concentrations 

up to 5 mg CN g –1 d. wt in the tree species R. javanica; 

individuals of that species occurring within the survey 

area of this study had a lower mean concentration of 

18mgCN g –1 d. wt (Table 1). These high concentrations 

are substantially greater than the majority of values reported 

for foliage from a range of tropical and temperate taxa. For 

example, concentrations up to 11 mg HCN g –1 d. wt were 

reported in the tropical shrub Turnera ulmifolia (Shore 

and Obrist, 1992; Schappert and Shore, 1999), while the 

highest concentrations in naturally occurring populations of 

Australian Eucalyptus spp. were 259 mg CN g –1 d. wt and 

316 mg CN g –1 d. wt for E. cladocalyx and E. yarraensis, 

respectively (Gleadow and Woodrow, 2000a; Goodger 

and Woodrow, 2002). Foliar concentrations of between 

166 and 378 mg HCN g 1 d. wt have been recorded in 

cultivated Prunus spp. (Santamour, 1998). To our knowledge, 

possibly the highest foliar cyanogenic glycoside 

concentration among naturally occurring woody species 

was reported in the tropical proteaceous species Panopsis 

costaricensis by Thomsen and Brimer (1997), who measured 

2150 mg HCN kg –1 f. wt (approximately equivalent to 

72 mg HCN g 1 d. wt using a conversion based on the mean 

foliar water content of several species in this study, which 

was 70 %). The age of the leaves analysed was not specified, 

however, and it should also be noted that this value was 

determined using picrate papers and reflectrometry, a less 

dependable method more sensitive to the presence of 

interfering substances (Brinker and Seigler, 1989). 

Assessing the ecological significance of the range 

of cyanogenic glycoside concentrations recorded here is 

difficult. While the high concentrations in several species 

(e.g. >2mgCNg 1 d. wt; Table 1) would almost certainly 

constitute toxic levels important in defence—plants with 

>600 mg HCN g 1 d. wt are considered potentially dangerous 

to livestock, for example (Haskins et al., 1987)—the 

ecological significance of lower concentrations (e.g. approx. 

8–50 mgCNg 1 d. wt) is harder to determine. This reflects

the fact that despite the well-documented effectiveness of 

cyanogenesis in defence against generalist herbivores 

(e.g. Jones, 1998; Gleadow and Woodrow, 2002), overall 

there is no known particular concentration at which 

cyanogenic compounds are effective in herbivore deterrence. 

This is in part because unfortunately many herbivory 

studies only report the presence or absence of cyanogenesis, 

and not the actual concentrations of cyanogenic glycosides. 

Moreover, the efficacy of cyanogenesis as a 

defence depends not only on the concentration of cyanogenic 

glycosides, but also on the physiology, morphology 

and behaviour of the consumer (Gleadow and Woodrow, 

2002). 

Intra-plant variation in cyanogenic glycoside content 

In tropical forests, it is estimated that up to 70 % of a leaf’s 

lifetime damage occurs while expanding (Coley and Barone, 

1996). This differential intensity of herbivory among old 

and young leaves, and the frequent observation that defence 

compounds tend to be concentrated in plant tissues of higher 

value to reproduction or growth (e.g. young leaves) is summarized 

in the Optimal Allocation Theory (OAT) of 

defence. This theory predicts that the most vulnerable 

and valuable plant parts—those susceptible to attack and 

most likely to contribute to growth and reproductive fitness 

such as reproductive structures and young leaves—will be 

more defended (McKey, 1974; Rhoades, 1979). 

Results here add to the already substantial body of work 

on mostly temperate cyanogenic species consistent with 

the predictions of the OAT (e.g. Martin et al., 1938; 

Dement and Mooney, 1974; Cooper-Driver et al., 1977; 

Shore and Obrist, 1992; Dahler et al., 1995; Thomsen 

and Brimer, 1997; Gleadow et al., 1998; Gleadow and 

Woodrow, 2000b). In all species where young leaves 

were sampled, they contained significantly higher concentrations 

of cyanogenic glycosides than old leaves. This 

trend was most apparent in species that had low cyanogen 

content in old leaves (e.g. C. sublimis, C. myrianthus, 

O. heterophylla and P. australiana) (Table 1). Moreover, 

in some cases, individuals of these species appeared only to 

invest in cyanogenic glycoside defence in young leaves; 

acyanogenic old leaves were seemingly reliant more on 

physical toughness. Thus, leaf age is an important consideration 

when assigning the cyanogenic phenotype. 

Again consistent with the OAT, reproductive tissues 

such as floral buds, flowers and fruits/seeds tended to 

have high total cyanogen content (Table 1). This pattern 

has been commonly reported among cyanogenic species 

(e.g. Spencer and Seigler, 1983; Selmar et al., 1991; 

Selmar, 1993b; Thomsen and Brimer, 1997; Webber, 

1999). One notable exception to this was the low to negligible 

concentrations of cyanogenic glycosides in mature 

seeds of C. sublimis; unlike the fleshy seeds of many rainforest 

species, C. sublimis seeds are dry and papery. The 

absence of cyanogenesis in mature seeds of proteaceous 

Grevillea spp. was reported by Lamont (1993) in species 

with cyanogenic foliage and flowers. The higher concentrations 

in floral tissues of C. sublimis is consistent with previous 

reports for proteaceous species which tend to have 

high concentrations of cyanogenic glycosides in flowers, 

while leaves may have low total cyanogen content or be 

acyanogenic (e.g. Smith and White, 1918; Tjon Sie Fat, 

1979a; Lamont, 1993). 

Intra-population variation in cyanogenic glycoside content 

All individuals of the majority of species were cyanogenic, 

albeit with low concentrations of cyanogenic 

glycosides in some instances. Negative results with FA 

papers for old leaves were only obtained for individuals 

of three of the 18 cyanogenic species. In the population 

of one of these species, B. platynema, 50 % of individuals 

were determined to be acyanogenic, with cyanogenic glycoside 

concentrations much less than the 8 mgCNg 1 d. wt 

threshold. The two other exceptions were C. myrianthus 

and P. australiana. Unlike B. platynema, cyanogenesis in 

individuals of these species varied qualitatively with leaf 

age and plant part. Thus, assigning the acyanogenic phenotype 

in these species was problematic. 

This developmental trend towards differences in 

expression of cyanogenic potential has been reported previously; 

cyanogenesis is known be affected by plant age, 

growth phase, as well as the plant part used (Jones, 1972; 

Gibbs, 1974; Seigler, 1991). Consequently, as noted earlier, 

studies have reported a greater frequency of cyanogenesis 

when testing reproductive tissues, young foliage and shoots 

compared with old leaves (e.g. Gibbs, 1974; Aikman et al., 

1996; Thomsen and Brimer, 1997; Buhrmester et al., 2000; 

Mali and Borges, 2003). These findings emphasize the 

importance of only comparing leaves of a similar age 

when classifying individuals according to the presence or 

absence of cyanogenesis. 

Aside from the three species mentioned above, no 

acyanogenic individuals were identified in populations of 

other cyanogenic species. While the small sample sizes for 

most species reduced the likelihood of encountering an 

acyanogenic individual, even within populations of the 

more abundant species such as B. collina (n = 46 all 

sites), C. sublimis (n = 31 at all sites) and R. javanica 

(n = 249; Webber, 1999), no acyanogenic individuals 

were detected. In the latter example, the quantitative screening 

of >800 individuals of R. javanica failed to detect an 

acyanogenic individual in several distinct populations 

(Webber, 2005). This number (800) was substantially 

greater than the number of individuals predicted by 

Gleadow and Woodrow (2000a) and Goodger et al. 

(2002) (n = 95–100) that would need to be sampled to 

capture an acyanogenic individual assuming a similar genetic 

system for cyanogenesis to Trifolium repens (Hughes 

et al., 1988) and an estimate of the rarity of a species (or 

polymorph) (McArdle, 1990). The floristic heterogeneity of 

the rainforest makes sampling large populations challenging; 

it is noteworthy, however, that others have detected 

polymorphism for cyanogenesis in tropical studies based 

on very small sample sizes (e.g. n = 2) (Kaplan et al., 

1983; Thomsen and Brimer, 1997). These acyanogenic 

morphs, determined only by indicator paper tests in these 

studies, were not verified by quantitative assay as for 

B. platynema here.

Conclusions 

In summary, the findings of this survey indicate that 

cyanogenesis is an important, yet little studied, chemical 

defence in tropical rainforests. The identification of specific 

cyanogens in but a few of the cyanogenic species first reported 

here has yielded novel findings. Given the large number 

of new reports for species belonging to plant families 

or orders in which cyanogenesis has been little reported, 

the ongoing characterization of cyanogenic constituents in 

these species will potentially be of both phytochemical and 

chemotaxonomic significance. In addition, preliminary data 

on intra-population variation in cyanogenesis here suggest 

that ontogenetic variation in cyanogenesis, and polymorphism 

for cyanogenesis merit further investigation in tropical 

rainforest species. 

ACKNOWLEDGEMENTS 

The authors thank Professor Eric Conn (University of 

California, Davis) for sharing unpublished data on the 

Proteaceae family, Nicole Middleton and volunteers at 

the University of Melbourne Herbarium for mounting 

and databasing specimens, Andrew Ford (CSIRO Tropical 

Forest Research Centre, Atherton, Queensland) for assistance 

with species name changes, and Wendy Cooper and 

Steve McKenna for additional taxonomic assistance during 

field work. We also thank Dr John Kanowski (Griffith 

University, Queensland) and Andrew Graham (TFRC, 

Atherton) for advice in the selection of field sites, and 

Marisa Collins, Bruce Webber, Ross Waller, Jennifer 

Fox, Gavin Smith and Joel Youl for assistance in the 

field. We thank the Australian Canopy Crane Research 

Facility for access to forest on the site. Field work was 

supported by an Individual Award from the Queen’s 

Trust for Young Australians to R.E.M., and was conducted 

under the Queensland Department of Environment and 

Heritage Scientific Purposes Permit number F1/000270/ 

99/SAA, and Department of Natural Resources Permits 

to Collect numbers 1542, 1716, 1409 and 1663. 

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APPENDIX 

Summary of all species tested for cyanogenesis within 

6 · 200 m 2 plots at five sites in upland/highland (U) or 

lowland (L) tropical rainforest, on sites contrasting in 

soil type: basalt sites at Lamins Hill (B1) and Longlands 

Gap (B2), and sites on granite at Mt Nomico (G), on rhyolite 

at Longlands Gap (R) and on metamorphic substrate near 

Cape Tribulation (M). Species are listed in alphabetical 

order. Life forms: tree (T), shrub (SH), herb (H), vine (V), 

treefern (TF), palm (P) and hemi-epiphyte (HE). The results 

(+ or –) for tests using Feigl–Anger papers and approx. 

1 g f. wt leaf tissue, with and without the addition of 

pectinase (+/– enz) for n individuals of each species are 

listed. All tests were carried out using most recently fully 

expanded leaves and in most instances also using young 

leaves. In some cases, leaf tips (tips), only a few fruit 

(ft) or flowers (flwr) were tested. Previous findings for 

species and in some cases genera are noted (e.g. Proteaceae 

species tested by E. E. Conn, University of California, 

Davis, CA, USA, pers. comm. based on herbarium specimens). 

Species included in phytochemical screenings of 

Queensland rainforest taxa (alkaloids, CN; Webb, 1948, 

Webb, 1949) are noted, most were not tested for CN 

(DNT). Canopy species or rare species for which no sample 

was obtained were included in floristic analysis, but were 

not tested for cyanogenesis (DNT). Lodgement numbers at 

Brisbane (BRI) and The University of Melbourne (MELU) 

herbaria are given.

A PPENDIX 

Lifeform Species Family HCN +/– enz (n tested) Upland/lowland Site (substrate) Previous reports Lodgement no. 

T Acacia celsa Tindale Mimosaceae / (3) U R Webb (1949) DNT 

T Acmena graveolens (F.M.Bailey) L.S.Sm. Myrtaceae / (3) L M MELU102318 

T Acmena resa B.Hyland Myrtaceae / (2) U B2, R 

T Acronychia acidula F.Muell. Rutaceae / (2) U G Webb (1949) DNT 

T Acronychia acronychioides (F.Muell.) T.G.Hartley Rutaceae / (3) U G 

T Acronychia crassipetala T.G.Hartley Rutaceae / (1) U B2 

SH Acronychia parviflora C.T.White Rutaceae / (5) U B1, R 

SH Actephila sp. (Wooroonooran NP 

P.I.Forster+ PIF17151) 

Euphorbiaceae / (5) U B1 

T Adenanthera pavonina L. Mimosaceae / (1) L M 

T Agathis atropurpurea B.Hyland Araucariaceae / (3) U R 

T Agathis microstachya J.F.Bailey & C.T.White Araucariaceae / (1) U G 

T Agathis robusta (C.Moore ex F.Muell.) F.M.Bailey Araucariaceae / (1) U B1 

SH Aglaia meridionalis C.M.Pannell Meliaceae / (13) U, L B1, M MELU102332 

T Aglaia sapindina (F.Muell.) Harms Meliaceae / (2) L M 

T Aglaia tomentosa Teijsm. & Binn. Meliaceae / (5) U, L B1, B2, G, M MELU102275 

T Alangium villosum subsp. polyosmoides 

Alangiaceae / (3) U B1 Webb (1949) DNT MELU102329 

(F.Muell.) Bloemb. 

T Alloxylon flammeum P.H.Weston & Crisp Proteaceae / (2) U B2 

T Alloxylon wickhamii (W.Hill ex F.Muell.) 

P.H.Weston & Crisp 

Proteaceae / (4) U G, B2 

H Alocasia brisbanensis (F.M.Bailey) Domin z Araceae +/+ U B1 Hurst (1942) 

T Alphitonia whitei Braid Rhamnaceae / (3) U G MELU102103 

H Alpinia arctiflora (F.Muell.) Benth. Zingiberaceae / (1) U B1 

H Alpinia modesta F.Muell. ex K.Schum. Zingiberaceae / (1) U B2 

T Alstonia scholaris (L.) R.Br. Apocynaceae / (10) L M Gibbs (1974) reported 

+CN; Webb (1949) DNT 

BRI 578809, 

MELU102127, 

102257 

SH Alyxia ilicifolia subsp. ilicifolia F.Muell. Apocynaceae / (3) U G 

V Alyxia spicata R.Br. Apocynaceae / (2) U B1, R Webb (1949) DNT 

T Antidesma erostre F.Muell. ex Benth. Euphorbiaceae / (6) U, L B1, G, R, M MELU102113 

SH Antirhea sp. (Mt Missery L.W.Jessup+ GJD 3136) Rubiaceae / (3) U B2 

T Antirhea tenuiflora F.Muell. ex Benth. Rubiaceae / (4) U, L B1, G, M 

T Apodytes brachystylis F.Muell. Icacinaceae / (6) U B1, G, R MELU102188 

T Archidendron ramiflorum (F.Muell.) Kosterm. Mimosaceae / (4) U, L B1, M 

T Archidendron vaillantii (F.Muell.) F.Muell. Mimosaceae / (3) U B2, G MELU102186 

T Archidendron whitei I.C.Nielsen Mimosaceae / (1) U B1 

SH Ardisia bifaria C.T.White & W.D.Francis Myrsinaceae / (2) U B1 

SH Ardisia brevipedata F.Muell. Myrsinaceae / (7); frt – U, L B1, B2, G, R, M MELU102148 

T Argyrodendron sp. (Boonjee BH 2139RFK) Sterculiaceae / (9) U B1 

T Arytera pauciflora S.T.Reynolds Sapindaceae / (4) U, L B1, M 

SH Atractocarpus hirtus (F.Muell.) Puttock Rubiaceae / (7) U, L B1, M Webb (1949) DNT 

SH Atractocarpus merikin (F.M.Bailey) Puttock Rubiaceae / (3) U B1, B2 

T Auranticarpa papyracea L.Cayzer, 

Crisp & I.Telford 

Pittosporaceae / (2) U B2, G 

V Austrobaileya scandens C.T.White Austrobaileyaceae / (6); flwr – U B1 

SH Austromatthaea elegans L.S.Sm. Monimiaceae / (3) U G, R 

T Austromuellera trinervia C.T.White Proteaceae / (4) U, L B1, M 

V Austrosteenisia stipularis (C.T.White) Jessup Fabaceae / (4) U B1, G

A PPENDIX Continued 


T Balanops australiana F.Muell. Balanopaceae / (3) U R, G Webb (1949) DNT MELU102216, 

102108 

T Beilschmiedia bancroftii (F.M. Bailey) C.T.White Lauraceae / (3) U, L B1, G, M Webb (1949) DNT MELU102219 

T Beilschmiedia castrisinensis B.Hyland Lauraceae / (2) L M 

T Beilschmiedia collina B.Hyland Lauraceae +/+ U B1, B2, G, R BRI 578804, 

MELU102299, 

102300 

T Beilschmiedia oligandra L.S.Sm. Lauraceae / (3) U G 

T Beilschmiedia recurva B.Hyland Lauraceae / (4) U B1, G 

T Beilschmiedia tooram (F.M.Bailey ) B.Hyland Lauraceae / (10) U B1, B2, G MELU102211 

T Bobea myrtoides (F.Muell.) Valeton Rubiaceae / (8) U G, R MELU102105, 102110 

SH Bowenia spectabilis Hook. ex Hook.f. Stangeriaceae / (6) U, L B1, M 

T Brachychiton acerifolius (A.Cunn. 

ex G.Don.) Macarthur 

Sterculiaceae / (3) U, L B2, M 

T Brackenridgea australiana F.Muell. Ochnaceae / (4) U, L G, R, M MELU102322 

SH/T Breynia cernua (Poir.) Muell.Arg Euphorbiaceae / (2) U B2, G Webb (1949) 1948 

B. oblongifolia Muell. 

Arg. +CN 

T Brombya platynema F.Muell. Rutaceae +/+ L M Webb (1949) DNT BRI 578818. 

MELU102283, 

102313 

T Bubbia semecarpoides (F.Muell.) B.L.Burtt Winteraceae / (5) U B1, G 

V (P) Calamus australis Mart. Arecaceae / (1); U B1 

V Calamus moti F.M.Bailey Arecaceae / (1) U B1 

T Caldcluvia australiensis (Schltr.) Hoogland Cunoniaceae / (1) U B1 

T Calophyllum costatum F.M.Bailey Clusiaceae / (3) U B2, R 

T Cananga odorata (Lam.) Hook.f. & Thomson Annonaceae / (2) L M 

T Canarium muelleri F.M.Bailey Burseraceae / (1) U G MELU102250 

T Canthium sp. Rubiaceae / (1) U G 

T Carallia brachiata (Lour.) Merr. Rhizophoraceae / (1) L M 

T Cardwellia sublimis F.Muell. Proteaceae +/+ U, L B1, B2, G, R, M E. E. Conn (pers. comm.) 

1of7+ve 

T Carnarvonia araliifolia var. montana B.Hyland Proteaceae / (3) U B2, G, R E. E. Conn 

(pers. comm.) –ve 

BRI 578806, 

MELU102280, 102281 

MELU102282 

V Carronia protensa (F.Muell.) Diels Menispermaceae / (5) U, L B1, B2, M MELU102213 

T Casearia costulata L.W.Jessup Salicaceae † / (4) U B1, B2, G, R Webb (1949) DNT 

T Casearia dallachii F.Muell. Salicaceae † / (1) L M 

T Casearia grayi L.W.Jessup Salicaceae † / (2) dry* U B2 

T Castanospermum australe A.Cunn. & 

C.Fraser ex Hook. 

Fabaceae / (3) L M Webb (1949) DNT 

T Castanospora alphandii (F.Muell.) F.Muell. Sapindaceae / (3) U B1, B2 MELU102130 

V Cayratia saponaria (Seem. & Benth.) Domin Vitaceae / (1) L M C. acris Gibbs (1974) –CN 

T Celtis paniculata (Endl.) Planch. Ulmaceae / (1) L M 

V Cephalaralia cephalobotrys (F.Muell.) Harms Araliaceae / (2) U B1, B2 

T Ceratopetalum succirubrum C.T.White Cunoniaceae / (4) U B2, G Webb (1949) DNT 

T Cerbera floribunda K.Schum. Apocynaceae / (1) U, L G, M Webb (1949) DNT 

T Chionanthus axillaris R.Br. Oleaceae / (4) U B1, G MELU102327 

T Cinnamomum laubatii F.Muell. Lauraceae / (3) U B1, G, R Webb (1949) DNT MELU102210 

V Cissus hypoglauca A.Gray Vitaceae / (2) U G 

V Cissus penninervis (F.Muell.) Planch. Vitaceae / (1) U, L R, M

V Cissus sterculiifolia (F.Muell. ex Benth.) Planch. Vitaceae / (4) U B1, B2, G MELU102325 

V Cissus vinosa Jackes Vitaceae / (3) U B1, B2, G MELU102195 

T Citronella smythii (F.Muell.) R.A.Howard Icacinaceae / (4) U, L B1, G, M MELU102143 

T Claoxylon tenerifolium (Baill.) F.Muell. Euphorbiaceae / (1) U B1 

T Cleistanthus myrianthus (Hassk.) Kurz Euphorbiaceae +/+ L M BRI 578811, 

MELU102122, 

102123, 102258 

T Clerodendrum grayi Munir Verbenaceae +/+ U B1, B2, G Other Clerodendrum 

spp. +CN (Gibbs, 1974; 

Tjon Sie Fat, 1979a; 

Adsersen et al., 1988) 

BRI 578815, 

MELU102115, 

102116, 102117 

T Cnesmocarpon dasyantha (Radlk.) Adema Sapindaceae / (2) U G 

V Connarus conchocarpus F.Muell. Connaraceae / (1) L M 

SH Cordyline petiolaris (Domin) Pedley Laxmanniaceae / (2) U B1 

SH Cordyline sp. Laxmanniaceae / (1) U G 

T Corynocarpus cribbianus (F.M.Bailey) L.S.Sm. Corynocarpaceae / (1) U B1 C. laevigata +CN 

(Webb, 1948) 

SH Crispiloba disperma (S.Moore) Steenis Alseuosmiaceae / (4) U G 

T Cryptocarya angulata C.T.White Lauraceae / (4) U B1, G, R MELU102102 

T Cryptocarya cocosoides B.Hyland Lauraceae / (3) U B2, G MELU102157, 102268 

T Cryptocarya corrugata 

C.T.White & W.D.Francis 

Lauraceae / (3) U B1, B2, G, R MELU102147 

T Cryptocarya densiflora Blume Lauraceae / (2) U G, R MELU102104, 102269 

T Cryptocarya grandis B.Hyland Lauraceae / (3); U, L B1, B2, G, M MELU102247 

T Cryptocarya hypospodia F.Muell. Lauraceae / (1) L M 

T Cryptocarya laevigata Blume Lauraceae / (2) U, L G, M 

T Cryptocarya leucophylla B.Hyland Lauraceae / (1) U B2, R MELU102248 

T Cryptocarya lividula B.Hyland Lauraceae / (2) U G, R MELU102095 

T Cryptocarya mackinnoniana F.Muell. Lauraceae / (5) U, L B1, G, M MELU102212 

T Cryptocarya melanocarpa B.Hyland Lauraceae / (5) U B1, B2 

T Cryptocarya murrayi F.Muell. Lauraceae / (2) U, L B1, M 

T Cryptocarya oblata F.M.Bailey Lauraceae / (2) U, L B1, M Webb (1949) DNT MELU102099 

T Cryptocarya pleurosperma C.T.White & 

W.D.Francis 

Lauraceae / (4) U B1, G Webb (1949) DNT MELU102270 

T Cryptocarya putida B.Hyland Lauraceae / (4) U G, R MELU102144 

T Cryptocarya saccharata B.Hyland Lauraceae / (3) U B2, R 

T Cryptocarya smaragdina B.Hyland Lauraceae / (3) U B2, R MELU102193 

T Cupaniopsis flagelliformis (F.M.Bailey) 

Sapindaceae / (4) U B1 MELU102129 

Radlk. var. flagelliformis 

T Cupaniopsis sp. Sapindaceae / (1) L M 

TF Cyathea rebeccae (F.Muell.) Domin Cyatheaceae / (2) U G, R 

T Cyclophyllum multiflorum 

S.T.Reynolds & R.J.F.Hend. 

Rubiaceae / (1) U B1 Canthium vaccinifolium 

F. Muell. +CN Webb 

(1949) 1948 

T Daphnandra repandula (F.Muell.) F.Muell. Monimiaceae / (2) U B1, B2, G 

T Darlingia darlingiana (F.Muell.) L.A.S.Johnson Proteaceae / (2) U G E. E. Conn (pers. comm.) –ve 

T Darlingia ferruginea J.F.Bailey Proteaceae / (3)– U B1, B2 E. E. Conn (pers. comm.) –ve 

T Davidsonia pruriens F.Muell. Davidsoniaceae / (2) U G Webb (1949) DNT; +CN 

(Rosenthaler, 1929; Hurst, 

1942); –CN Gibbs (1974) 

T Decaspermum humile (G.Don.) A.J.Scott Myrtaceae / (1) dry* L M 

T Delarbrea michieana (F.Muell.) F.Muell. Araliaceae / (4) U B1, G 

V Dendrotrophe varians (Blume) Miq. Santalaceae / (1) U R 

V Derris sp. (Daintree D.E. Boyland+469) Fabaceae / (1) dry* L M 

MELU102151, 102152



V Desmos goezeanus (F.Muell.) Jessup Annonaceae / (2) U G 

V Dichapetalum papuanum (Becc.) Boerl. Dichapetalaceae / (4) U B1 MELU102326 

T Dinosperma stipitatum (C.T.White & 

W.D.Francis) T.G. Hartley 

Rutaceae / (3) U B1 

T Diospyros cupulosa F.Muell. Ebenaceae / (1) L M 

T Diospyros hebecarpa A.Cunn. ex Benth. Ebenaceae / (1) dry* U B1 MELU102307 

T Diploglottis bernieana S.T.Reynolds Sapindaceae / (2) L M 

T Diploglottis bracteata Leenh. Sapindaceae / (1) U G 

T Doryphora aromatica (F.M.Bailey) L.S.Sm. Monimiaceae / (5) U, L B1, B2, M Webb (1949) DNT 

T Dysoxylum alliaceum Blume (Blume) Meliaceae / (1) L M 

T Dysoxylum arborescens (Blume) Miq. Meliaceae / (2) U, L B1, M 

T Dysoxylum klanderi F.Muell. Meliaceae / (3) U B1, B2, R 

T Dysoxylum oppositifolium F.Muell. Meliaceae / (3) U B2, G MELU102273 

T Elaeocarpus bancroftii F.Muell. & F.M.Bailey Elaeocarpaceae / (3) L M 

T Elaeocarpus eumundi F.M.Bailey Elaeocarpaceae / (1) U G, R MELU102125 

T Elaeocarpus foveolatus F.Muell. Elaeocarpaceae / (2) U B1, R 

T Elaeocarpus grahamii F.Muell. Elaeocarpaceae / (1) L M 

T Elaeocarpus largiflorens 

C.T.White subsp. largiflorens 

Elaeocarpaceae / (4) U B1, G 

T Elaeocarpus ruminatus F.Muell. Elaeocarpaceae / (3) U B2, G 

T Elaeocarpus sericopetalus F.Muell. Elaeocarpaceae +/+ U G, R BRI 578817, 

MELU102135, 

102304 

T Elaeocarpus sp. (Mt Bellenden Ker LJB 18336) Elaeocarpaceae / (3) U B2, G, R MELU102304 

V Embelia caulialata S.T.Reynolds Myrsinaceae / (1) dry* L M 

V Embelia grayi S.T.Reynolds Myrsinaceae +/+ U B1, B2 BRI 578803, 

MELU102267 

T Endiandra bessaphila B.Hyland Lauraceae / (2) U B1 

T Endiandra dielsiana Teschn. Lauraceae / (2) U G 

T Endiandra hypotephra F.Muell. Lauraceae DNT L M 

T Endiandra leptodendron B.Hyland Lauraceae / (7) U, L B1, M MELU102293 

T Endiandra microneura C.T.White Lauraceae / (3) L M 

T Endiandra monothyra B.Hyland subsp. monothyra Lauraceae / (3) U B1, B2 

T Endiandra montana C.T.White Lauraceae DNT U B2 

T Endiandra palmerstonii (F.M.Bailey) 

C.T.White & W.D.Francis 

Lauraceae / (2) U B2, G Webb (1949) DNT MELU102294 

T Endiandra sankeyana F.M.Bailey Lauraceae / (3) U B1, B2 MELU102155 

T Endiandra sideroxylon B.Hyland Lauraceae / (3) U B2 MELU102109 

T Endiandra wolfei B.Hyland Lauraceae / (3); sht – U B1, G MELU102295 

T Endospermum myrmecophilum L.S.Sm. Euphorbiaceae / (1) dry * L M 

V Entada phaseoloides (L.) Merr. Mimosaceae / (1) L M 

HE Epipremnum pinnatum (L.) Engl. Araceae / (2) L M 

V Erycibe coccinea (F.M.Bailey) Hoogl. Convolvulaceae / (3) L M 

T Erythroxylum ecarinatum Burck ex Hoch. Erythroxylaceae / (2) U B1 Webb (1949) DNT MELU102225 

SH Eupomatia barbata Jessup Eupomatiaceae / (2) dry* U, L B1, M 

SH/T Eupomatia laurina R.Br. Eupomatiaceae / (4) U B2, G Gibbs (1974) ‘doubtfully MELU102331 

cyanogenic’; Webb 

(1949) DNT 

V Eustrephus latifolius R.Br. ex Ker Gawl. Philesiaceae / (2) U, L B1, M 

T Fagraea cambagei Domin Gentianaceae / (4) L M MELU102317

T Fagraea fagraeacea (F.Muell.) Druce Gentianaceae / (3) U B1, B2, G, R MELU102323 

T Ficus congesta Roxb. Moraceae / (1) L M 

T Ficus crassipes F.M.Bailey Moraceae / (1) U B1 

T Ficus destruens F.Muell. ex C.T.White Moraceae / (1) U B2, G 

T Ficus leptoclada Benth. Moraceae DNT U B1, B2 MELU102107 

V Ficus pantoniana King Moraceae / (1) L M 

T Ficus pleurocarpa F.Muell. Moraceae / (1) U B1 

T Ficus triradiata Corner Moraceae / (1) L M 

V Flagellaria indica L. Flagellariaceae +/+ U, L B1, G, M +CN (1912) BRI 578816, 

MELU102259, 

102296 

T Flindersia bourjotiana F.Muell. Rutaceae / (4) U, L B2, G, R, M Webb (1949) DNT 

T Flindersia brayleyana F.Muell. Rutaceae / (1) U B2 Webb (1949) DNT 

T Flindersia laevicarpa C.T.White & W.D.Francis Rutaceae / (2) U G MELU102222 

T Flindersia pimenteliana F.Muell. Rutaceae / (2) U G, R Webb (1949) DNT MELU102132, 

102106 

T Fontainea picrosperma C.T.White Euphorbiaceae / (2) U B2 

T Franciscodendron laurifolium 

(F.Muell.) B.Hyland & Steenis 

Sterculiaceae / (10) U B1, G 

V Freycinetia excelsa F.Muell. Pandanaceae / (3) U B1, B2 

V Freycinetia scandens Gaudich. Pandanaceae / (1) U B1 

T Galbulimima baccata F.M.Bailey Himantandraceae / (3) U B1, B2, G, R Webb (1949) DNT; 

fruits –CN (Bailey, 1909 

in Webb, 1948) 

T Garcinia gibbsiae S.Moore Clusiaceae / (7) U B1 MELU102126 

T Garcinia warrenii F.Muell. Clusiaceae / (4) L M 

T Gardenia ovularis F.M.Bailey Rubiaceae / (5) U, L G, M Webb (1949) DNT MELU102319 

T Gevuina bleasdalei (F.Muell.) Sleumer Proteaceae / (3) U B1, G E. E. Conn (pers. comm.) –ve MELU102292 

T Gillbeea adenopetala F.Muell. Cunoniaceae / (3) U B1, G 

T Glochidion harveyanum Domin var. harveyanum Euphorbiaceae / (1) U G Webb (1949) DNT 

T Glochidion hylandii Airy Shaw Euphorbiaceae / (1) U B2, G Webb (1949) DNT 

T Gmelina fasciculiflora Benth. Lamiaceae / (2) U, L G, M Webb (1949) DNT MELU102261, 

102262,102263 

T Gomphandra australiana F.Muell. Icacinaceae / (2) L M MELU102333 

T Goniothalamus australis Jessup Annonaceae / (6) U B1 MELU102185, 

102290 

T Gossia dallachiana (F.Muell.) N.Snow & Guymer Myrtaceae / (5) U B1, B2, G MELU102220 

T Gossia myrsinocarpa (F.Muell.) 

N.Snow & Guymer 

Myrtaceae / (1) U B2 

T Gossia grayii N.Snow & Guymer Myrtaceae / (2) U G MELU102146 

T Grevillea baileyana McGill. Proteaceae / (1) L M Webb (1952); Hurst (1942) 

Grevillea spp. +CN; 

E. E. Conn (pers. comm.) –ve 

T Guioa acutifolia Radlk. Sapindaceae / (2) U, L B2, M 

T Guioa lasioneura Radlk. Sapindaceae / (2) U B1, G 

T Guioa montana C.T.White Sapindaceae / (2) U R MELU102150 

SH Gymnostachys anceps R.Br. Araceae / (2) U, L B2, M 

V Gynochtodes sp. (Lamb Range J.W.398) Rubiaceae / (2) U G, R 

T Halfordia kendack (Montrouz.) Guillaumin Rutaceae / (3) U B1, B2, G, R Webb (1949) DNT MELU102112 

SH Haplostichanthus sp. (Coopers Creek B. Gray 2433) Annonaceae / (7) L M MELU102291 

SH Haplostichanthus sp. (Topaz L.W.Jessup 520) Annonaceae / (7) U B1 MELU102328 

SH Harpullia frutescens F.M.Bailey Sapindaceae / (1) U B2 

SH/T Harpullia rhyticarpa C.T.White & W.D.Francis Sapindaceae / (10) U, L B1, G, M Webb (1949) DNT 

T Hedycarya loxocarya (Benth.) W.D.Francis Monimiaceae / (4) U G



T Helicia australasica F.Muell. Proteaceae +/+ L M E. E. Conn (pers. comm.) 

flwr +ve; H. robusta +CN 

Pammel, 1911 in Webb (1949) 

1948; Gibbs (1974) 

BRI 578805, 

MELU102284 

T Helicia blakei Foreman Proteaceae +/+ U B1 E. E. Conn (pers. comm.) –ve BRI 578807, 

MELU102285 

T Helicia lamingtoniana (F.M.Bailey) 

Proteaceae / (3) U B2 E. E. Conn (pers. comm.) –ve MELU102214 

C.T.White ex L.S.Sm. 

T Hernandia albiflora (C.T.White) Kubitzki Hernandiaceae / (4) L M Webb (1949) DNT MELU102314 

V Hibbertia scandens (Willd.) Dryand. Dilleniaceae / (1) U R 

V Hippocratea barbata F.Muell. Celastraceae / (4) U, L B1, M 

V Hoya sp. Asclepiadaceae / (1) L M 

T Hylandia dockrillii Airy Shaw Euphorbiaceae / (2) U B1 

V Hypserpa decumbens (Benth.) Diels Menispermaceae / (4) U B1, G MELU102316 

V Hypserpa laurina (F.Muell.) Diels Menispermaceae / (2) L M Webb (1949) DNT MELU102276 

V Hypserpa smilacifolia Diels Menispermaceae / (1) U B2 

T Hypsophila dielsiana Loes. Celastraceae / (3) U B1 MELU102311 

T Irvingbaileya australis (C.T.White) R.A.Howard Icacinaceae / (2) U B1, G MELU102324 

T Ixora biflora Fosberg Rubiaceae / (2) L M 

T Ixora sp. (North Mary LA B.P.Hyland 8618) Rubiaceae / (3) U B1, B2 

V Jasminum didymum G.Forst. Oleaceae / (2) U B1, B2, G 

V Jasminum kajewskii C.T.White Oleaceae / (2) U B2, G 

SH Lasianthus strigosus Wight Rubiaceae / (2) L M 

SH/T Leea indica (Burm.f.) Merr. Leeaceae / (1) L M 

SH/T Lepidozamia hopei (W.Hill) Regel Zamiaceae / (3) L M 

T Lethedon setosa (C.T.White) Kosterm. Thymelaeaceae / (2) U G 

P(T) Licuala ramsayi (F.Muell.) Domin Arecaceae / (3) L M 

P(SH) Linospadix minor (W.Hill) F.Muell. Arecaceae / (2) L M 

T Litsea bindoniana F.Muell. (F.Muell.) Lauraceae / (2) U G MELU102097 

T Litsea connorsii B.Hyland Lauraceae / (2) U G, R 

T Litsea leefeana (F.Muell.) Merr. Lauraceae / (7) U,L B1, B2, M MELU102145 

T Loganiaceae sp. Loganiaceae / (1) U G 

T Lomatia fraxinifolia F.Muell. ex Benth. Proteaceae / (2) U B2, G E. E. (Conn (pers. comm.) –ve; MELU102091 

L. silaifolia R.Br. ft, flwr +CN 

Webb (1949) 1948 

T Macaranga inamoena F.Muell. ex Benth. Euphorbiaceae / (7) U B1 Webb (1949) DNT MELU102156 

T Macaranga subdentata Benth Euphorbiaceae / (1) L M 

SH Mackinlaya confusa Hemsl. Araliaceae / (1) U R 

SH Mackinlaya macrosciadea (F.Muell.) F.Muell. Araliaceae / (4) U B1, G Webb (1949) DNT 

V Maesa dependens F.Muell. Myrsinaceae / (1) U R 

SH/T Melicope broadbentiana F.M.Bailey Rutaceae / (3) U B1, G MELU102131 

T Melicope jonesii T.G.Hartley Rutaceae / (1) U B2 

T Melicope vitiflora (F.Muell.) T.G.Hartley Rubiaceae / (1) U, L B1, G, M 

V Melodinus acutiflorus F.Muell. Apocynaceae / (1) L M Webb (1949) DNT 

V Melodinus australis (F.Muell.) Pierre Apocynaceae / (4) U B1, G, R 

V Melodinus baccellianus (F.Muell.) S.T.Blake Apocynaceae / (3) U B1, B2 MELU102230 

V Melodorum uhrii F.Muell. Annonaceae / (1) dry* L M 

T Mischarytera lautereriana (F.M.Bailey) H.Turner Sapindaceae / (2) U B2, G, R MELU102286

T Mischocarpus exangulatus (F.Muell.) Radlk. Sapindaceae +/+ U B1 BRI 578801, 

MELU102190, 

102288 

T Mischocarpus grandissimus (F.Muell.) Radlk. Sapindaceae +/+ U, L G, M BRI 578802, 

MELU102287 

T Mischocarpus lachnocarpus (F.Muell.) Radlk. Sapindaceae / (3) U B2, G 

T Mischocarpus macrocarpus S.T.Reynolds Sapindaceae / (3) U B1, B2 

T Mischocarpus pyriformis (F.Muell.) 

Radlk. subsp. pyriformis 

T Monimiaceae Gen.(AQ63687) sp. (Davies 

Creek L.J.Webb+ 6430) 

V Morinda Gen.(AQ124851) sp. 

(Boonjie L.J.Webb+ 6837A) 

Sapindaceae / (1) U B2 

Monimiaceae / (3) U B2, R 

Rubiaceae / (3) U B1 

V Morinda jasminoides A.Cunn. ex Hook. Rubiaceae / (1) U B2, G Webb (1949)–CN 

V Morinda sp. Rubiaceae / (1) U G 

V Morinda umbellata L. Rubiaceae / (1) U G 

T Musgravea heterophylla L.S.Sm. Proteaceae / (2) L M E. E. Conn (pers. comm.) 

1of8+ve 

T Musgravea stenostachya F.Muell. Proteaceae / (4) U G, R E. E. Conn (pers. comm.) –ve 

T Myristica globosa subsp. 

muelleri (Warb.) W.J.de Wilde 

Myristicaceae / (3) U B1 

T Myristica insipida R.Br. Myristicaceae / (3) L M MELU102312 

Apocynaceae / (2) U B2 

T Neisosperma poweri (F.M.Bailey) 

Fosberg & Sachet 

T Neolitsea dealbata (R.Br.) Merr. Lauraceae / (4) U B1, B2 Gibbs (1974) –CN 

V Neosepicaea jucunda (F.Muell.) Steenis Bignoniaceae / (2) L M 

T Niemeyera prunifera (F.Muell.) F.Muell. Sapotaceae / (4) U, L B1, G, M MELU102149 

P(T) Normanbya normanbyi (W.Hill) L.H.Bailey Arecaceae / (4) L M 

T Opisthiolepis heterophylla L.S.Sm. Proteaceae +/+ U B1, B2 E.E . Conn (pers. comm.) 

flwr and lf +ve 

P(T) Oraniopsis appendiculata 

(F.M.Bailey) J.Dransf., 

A.K.Irvine & N.W.Uhl 

Arecaceae / (1) U B1 

BRI 578808, 

MELU102298 

V Pachygone longifolia F.M.Bailey Menispermaceae / (2) L M 

T Palaquium galactoxylon (F.Muell.) H.J.Lam Sapotaceae / (1) dry* L M 

V Palmeria scandens F.Muell. Monimiaceae / (4) U G MELU102194 

SH/T Pandanus monticola F.Muell. Pandanaceae / (1) U G 

V Pandorea pandorana (Andrews) Steenis Bignoniaceae / (1) U B2 Webb (1949) DNT 

V Pararistolochia australopithecurus (F.Muell.) 

Michael J.Parsons 

Aristolochiaceae / (3) U B1, B2, G 

V Parsonsia latifolia (Benth.) S.T.Blake Apocynaceae +/+ U B1, G, R Webb (1949) DNT; 

other Parsonsia spp. –CN 

BRI 578800, 

MELU102128 

V Parsonsia sp. 1 Apocynaceae / (1) U G 

V Parsonsia langiana F.Muell Apocynaceae / (1) U R 

V Passiflora sp. (Kuranda BH12896) Passifloraceae +/+ L M BRI 578813, 

MELU102297 

T Perrottetia arborescens (F.Muell.) Loes. Celastraceae / (1) U G 

SH/T Pilidiostigma papuanum (Lauterb.) A.J.Scott Myrtaceae / (3) L M 

SH Pilidiostigma tetramerum L.S.Sm. Myrtaceae / (4) U B1 MELU102274 

T Pilidiostigma tropicum L.S.Sm. Myrtaceae / (2) U B1 

V Piper caninum Blume Piperaceae / (3) L M MELU102265 

V Piper novae-hollandiae Miq. Piperaceae / (3) U B1, G Webb (1949) DNT 

T Pitaviaster haplophyllus (F.Muell.) T.G.Hartley Rutaceae / (12) U, L B1, G, M MELU102100, 

102187



SH/T Pittosporum rubiginosum A.Cunn. Pittosporaceae / (3) U, L B1, B2, G, R, M P. undulatum 

(Bailey, 1909 in 

Webb, 1949) 1948 

T Pittosporum wingii F.Muell. Pittosporaceae / (2) U G 

Proteaceae / (2) U G 

T Placospermum coriaceum C.T.White & 

W.D.Francis 

T Podocarpus dispermus C.T.White Podocarpaceae / (2) U B1 

T Polyalthia michaelii C.T.White Annonaceae DNT U B1 

T Polyosma alangiacea F.Muell. Grossulariaceae / (3) U G, R 

T Polyosma hirsuta C.T.White Grossulariaceae / (1) U B1, B2 

T Polyosma rhytophloia C.T.White & W.D.Francis Grossulariaceae / (1) U B2, R Webb (1949) DNT 

T Polyosma rigidiuscula F.Muell. & 

F.M.Bailey ex F.M. Bailey 

Grossulariaceae / (3) U B1 

MELU102192 

T Polyscias australiana (F.Muell.) Philipson Araliaceae +/+ U, L B1, B2, G, R, M BRI 578812, 

MELU102301 

T Polyscias mollis (Benth.) Harms & C.T.White Araliaceae / (1) U B1 

T Polyscias murrayi (F.Muell.) Harms Araliaceae / (1) U B1 Webb (1949) DNT 

SH Polyscias purpurea C.T.White Araliaceae / (4) U G MELU102154 

HE Pothos longipes Schott Araceae / (4) U, L B1, M 

T Pouteria asterocarpon (P.Royen) Jessup Sapotaceae / (1) U B2 

T Pouteria brownlessiana (F.Muell.) Baehni Sapotaceae / (3) U, L B1, B2, G, M 

T Pouteria castanosperma (C.T.White) Baehni Sapotaceae / (4) U B1, G 

T Pouteria chartacea (F.Muell. ex Benth.) Baehni Sapotaceae / (3) 2 dry* L M 

T Pouteria myrsinodendron (F.Muell.) Jessup Sapotaceae / (7) U, L G, M MELU102221 

T Pouteria papyracea (P.Royen) Baehni Sapotaceae / (2) U B2 MELU102308, 

102309 

T Pouteria pearsoniorum Jessup Sapotaceae / (1) U R 

T Prunus turneriana (F.M.Bailey) Kalkman Rosaceae +/+ U, L B1, M BRI 578814, 

MELU102133, 

102134 

H Pseuderanthemum variabile (R.Br.) Radlk. Acanthaceae / (1) L M Webb (1949) DNT 

T Pseuduvaria froggattii (F.Muell.) Jessup Annonaceae / (3) L M MELU102310 

T Pseuduvaria mulgraveana Jessup 

var. glabrescens 

Annonaceae / (1) U G 

SH Psychotria dallachiana Benth. Sapindaceae / (1) U B2 

SH Psychotria loniceroides Sieber ex DC. Rubiaceae / (2) U G Webb (1949) DNT 

SH Psychotria nematopoda F.Muell. Rubiaceae / (2) L M 

SH Psychotria sp. Rubiaceae / (1) L M 

SH Psychotria sp. (Utchee Creek H. Flecker NQNC 5313) Rubiaceae / (3) U G 

SH Psychotria submontana Domin Rubiaceae / (2) U B1, G 

T Psydrax laxiflorens S.T.Reynolds & R.J.F.Hend. Rubiaceae / (2) U G 

T Pullea stutzeri (F.Muell.) Gibbs Cunoniaceae / (3) U G 

SH Randia sp. (Boonjie L.W.Jessup+ GJM264) Rubiaceae / (1) U B1 

SH Randia tuberculosa F.M.Bailey Rubiaceae / (7) U B1, G MELU102158 

T Rapanea achradifolia (F.Muell.) Mez Myrsinaceae / (3) U B2, G, R MELU102217 

SH/T Rapanea porosa (F.Muell.) Mez Myrsinaceae / (3) U, L R, M 

SH Rapanea sp. (Atherton K.J.White AQ91778) Myrsinaceae / (2) U R MELU102260 

HE Rhaphidophora petrieana A.Hay Araceae / (3) L M Webb (1949) DNT 

T Rhodamnia blairiana F.Muell. Myrtaceae / (2) U G 

T Rhodamnia spongiosa (F.M.Bailey) Domin Myrtaceae / (2) U R

T Pammel (1911) 

Rhodomyrtus pervagata Guymer Myrtaceae / (3) U B2, G, R 

T Rhysotoechia mortoniana (F.Muell.) Radlk. Sapindaceae / (1) U B2 

T Rhysotoechia robertsonii (F.Muell.) Radlk. Sapindaceae / (1) L M 

V Ripogonum album R.Br. Smilacaceae / (3) U B1, B2, G 

V Ripogonum elseyanum F.Muell. Smilacaceae / (3) U B1 

V Rourea brachyandra F.Muell. Connaraceae / (1) L M 

SH Rubus queenslandicus A.R.Bean Rosaceae / (1) U B2 

T Ryparosa javanica (Blume) Kurz ex 

Achariaceae † +/+ L M Koord. & Valeton D R. caesia +CN; 

Rosenthaler (1919) 

Ryparosa spp. +CN 

V Sarcopetalum harveyanum F.Muell. Menispermaceae / (1) B B1 Webb (1949) DNT; 

Hurst (1942) –CN 

T Sarcopteryx martyana (F.Muell.) Radlk. Sapindaceae DNT U G 

T Sarcopteryx reticulata S.T.Reynolds Sapindaceae / (2) L M 

T Sarcotoechia lanceolata (C.T.White) 

S.T.Reynolds 

Sapindaceae / (2) U G 

MELU102277 

T Sarcotoechia protracta Radlk. Sapindaceae / (3) U B1, G 

T Sarcotoechia sp. (Mt Carbine 

Sapindaceae / (2) U R MELU102303 

L.W.Jessup+ GJM995) 

V Scaevola enantophylla F.Muell. Goodeniaceae / (4) U B1, G Webb (1949) DNT 

T Schistocarpaea johnsonii F.Muell. Rhamnaceae / (5) U B1 

T Schizomeria whitei Mattf. Cunoniaceae / (1) U G 

T Scolopia braunii (Klotzsch) Sleumer Salicaceae † / (1) U B2 

T Siphonodon membranaceus F.M.Bailey Celastraceae / (3) U, L B1, M 

T Sloanea australis subsp. parviflora Coode Elaeocarpaceae / (4) U B1 

T Sloanea langii F.Muell. Elaeocarpaceae / (5) U, L B2, G, M 

T Sloanea macbrydei F.Muell. Elaeocarpaceae / (3) U B1, G 

V Smilax aculeatissima Conran Smilacaceae / (3) U B1 

V Smilax calophylla Wall. ex A.DC. Smilacaceae / (1) L M 

V Smilax glauca Walter Smilacaceae / (2) U B1 

V Smilax glyciphylla Sm. Smilacaceae / (2) U B2, R 

SH Solanum macoorai F.M.Bailey Solanaceae / (1) U B2 

SH Solanum sp. Solanaceae / (2) U B1 

SH Solanum viridifolium Dunal Solanaceae / (1) U G 

T Sphenostemon lobosporus (F.Muell.) L.S.Sm. Sphenostemonaceae / (3) U G, R MELU102111 

SH/T Steganthera australiana C.T.White Monimiaceae / (2) U B2 

T Steganthera macooraia (F.M.Bailey) P.K.Endress Monimiaceae / (4) U R MELU102231 

T Stenocarpus reticulatus C.T.White Proteaceae / (1) U R E. E. Conn (pers. comm.) –ve MELU102196 

T Stenocarpus sinuatus (Loudon) Endl. Proteaceae / (1); flwr – U B2 E. E. Conn (pers. comm.) –ve 

V Stephania japonica (Thunb.) Miers Menispermaceae DNT L M 

T Storckiella australiensis J.H.Ross & B.Hyland Caesalpiniaceae / (3) L M MELU102279 

T Streblus glaber var. australianus 

(C.T.White) Corner 

Moraceae / (3) U B2, R 

V Strychnos minor Dennst. Strychnaceae / (1) L M 

T Symplocos cochinchinensis subsp. 1 Symplocaceae / (2) U G 

T Symplocos cochinchinensis var. gittonsii Noot. Symplocaceae / (2) U B2, R 

T Symplocos cochinchinensis var. pilosiuscula Noot. Symplocaceae / (3) U B2, G, R MELU102218 

SH Symplocos hayesii C.T.White & W.D.Francis Symplocaceae / (3) U B1, B2, R MELU102320 

SH/T Symplocos paucistaminea F.Muell. & F.M.Bailey Symplocaceae / (3) U, L B1, G, M 

T Synima cordierorum (F.Muell.) Radlk. Sapindaceae / (1) U, L B2, G, M 

T Synima macrophylla S.T.Reynolds Sapindaceae / (4) U B1, B2, G MELU102289 

T Synoum muelleri C.DC. Meliaceae / (2) U B2 MELU102306 

T Syzygium canicortex B.Hyland Myrtaceae / (2) U B2, G MELU102272

T Syzygium cormiflorum (F.Muell.) B.Hyland Myrtaceae / (3) U, L B1, B2, G, M MELU102224 

T Syzygium endophloium B.Hyland Myrtaceae / (4) U B1, B2, G, R MELU102153 

T Syzygium gustavioides (F.M.Bailey) B.Hyland Myrtaceae / (1) U B1 

T Syzygium johnsonii (F.Muell.) B.Hyland Myrtaceae / (3) U B2, G 

T Syzygium kuranda (F.M.Bailey) B.Hyland Myrtaceae / (4) U, L B1, G, M 

T Syzygium luehmannii (F.Muell.) L.A.S.Johnson Myrtaceae / (1) U R 

T Syzygium monospermum Craven Myrtaceae / (1) L M 

T Syzygium papyraceum B.Hyland Myrtaceae / (3) U B1, G MELU102159 

T Syzygium trachyphloium (C.T.White) B.Hyland Myrtaceae / (1) U B1 

T Syzygium wesa B.Hyland Myrtaceae / (3) U B2, G MELU102096, 

102249 

T Syzygium wilsonii subsp. cryptophlebium 

Myrtaceae / (3) U B1, G MELU102223 

(F.Muell.) B.Hyland 

SH Tabernaemontana pandacaqui Lam. Apocynaceae / (1) L M 

SH Tapeinosperma sp. (Cedar Bay 

J.G.Tracey 14780) 

Myrsinaceae / (4) U G 

SH Tasmannia insipida R.Br. ex DC. Winteraceae / (3) dry* U R 

T Ternstroemia cherryi (F.M.Bailey) 

Theaceae / (2) L M 

Merr. ex J.F.Bailey & C.T.White 

V Tetracera nordtiana var. nordtiana F.Muell. Dilleniaceae / (2) U, L G, M 

T Tetrasynandra laxiflora (Benth.) J.R.Perkins Monimiaceae / (6) U, L B1, G, M MELU102229 

T Timonius singularis (F.Muell.) L.S.Sm. Rubiaceae / (1) U B1 

T Toechima erythrocarpum (F.Muell.) Radlk. Sapindaceae / (3) U, L B1, G, M MELU102321 

T Toechima monticola S.T.Reynolds Sapindaceae / (1) U G, R MELU102098 

T Triunia erythrocarpa Foreman Proteaceae / (3) U B1 E. E. Conn (pers. comm.) –ve 

V Trophis scandens (Lour.) Hook. & 

Arn. subsp. scandens 

Moraceae / (1) L M MELU102271 

V Uncaria lanosa var. appendiculata 

(Benth.) Ridsdale 

Rubiaceae / (1) L M 

T Uromyrtus metrosideros (F.M.Bailey) A.J.Scott Myrtaceae / (1) U R MELU102315 

V Ventilago ecorollata F.Muell. Rhamnaceae DNT L M 

T Wilkiea angustifolia (F.M.Bailey) J.R.Perkins Monimiaceae / (1) U B1 MELU102330 

SH Wilkiea sp. Monimiaceae / (2) U B1 

SH Wilkiea sp. (Barong L.W.Jessup 719) Monimiaceae / (14) U B1, B2, G 

SH Wilkiea sp. Gen. (AQ63687) sp. 

(Davies Creek L.J.Webb+ 6430) 

Monimiaceae / (3) U B1 MELU102302 

SH Wilkiea wardellii (F.Muell.) J.R.Perkins Monimiaceae / (3) U R 

T Xanthophyllum octandrum (F.Muell.) Domin Xanthophyllaceae / (3) U, L B1, B2, G, M MELU102101 

T Xylopia maccreae (F.Muell.) L.S.Sm. Annonaceae / (1) L M 

T Zanthoxylum veneficum F.M.Bailey Rutaceae / (2) U B2, G Webb (1949) DNT MELU102215 

† Species in the families Salicaceae and Achariaceae which, until recent revisions, were in the Flacourtiaceae (Chase et al., 2002) 

z Non-woody Alocasia brisbanensis was not included in the analysis of the frequency of cyanogenic species. 

D Ryparosa javanica is currently the subject of taxonomic review; this Queensland Ryparosa sp. is likely to be renamed (Webber, 2005). 

* Due to limited access to foliage, tests for cyanogenesis using fresh tissue were not conducted, instead, freeze-dried samples were assayed quantitatively for the presence of CN indicated as ‘dry’.

Frequency of Cyanogenesis in Tropical Rainforests of Far North ...

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