Natural products from plants targeting key enzymes for the future development of antidiabetic agents

R. Mata *, L. Flores-Bocanegra , B. Ovalle-Magallanes and M. Figueroa
Departamento de Farmacia, Facultad de Química, Universidad Nacional Autónoma de México, Mexico City, 04510, Mexico. E-mail: rachel@unam.mx

First published on 7th June 2023

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R. Mata

Rachel Mata was born in Caracas, Venezuela, in 1949. She received her B.S. degree in Pharmacy from the Universidad Central de Venezuela in 1971 and her M.S. (1976) and PhD (1979) degrees at Purdue University, IN, USA, under the direction of Dr Jerry L. McLaughlin. Following a postdoctoral stay at the Instituto de Química at Universidad Nacional Autónoma de México (UNAM), she became an Assistant Professor (1984–1987), Associate Professor (1987–1990), Visiting Professor at the University of Rhode Island, TRI, USA (1990–1991), and Professor (1991–2016) at Facultad de Química at UNAM, where she is currently Professor Emerita. Her work has contributed significantly to knowledge of Mexican traditional medicine by establishing the chemical composition and biological properties of many widely used herbal drugs.

L. Flores-Bocanegra

Laura Flores-Bocanegra received her PhD degree in Chemistry from the Universidad Nacional Autónoma de México (UNAM), and her research topic dealt mainly with the study of medicinal plants from Mexico. From 2018 to 2021, she worked as a Postdoctoral Fellow at the University of North Carolina at Greensboro, NC, USA, as a part of Professor Nicholas H. Oberlies's team. During this period, Dr Flores-Bocanegra worked in the chemical study of the medicinal plant Kratom and several filamentous fungi. She isolated diverse secondary metabolites with different activities such as opioid agonist, cytotoxic, and antibacterial. She became an Assistant Professor at the Facultad de Química at UNAM in 2023. The main research interests of Dr Flores-Bocanegra involve the isolation and characterization of bioactive compounds from medicinal plants as well as fungal species, their pharmacological evaluation, and the validation of analytical methods for the quality control of herbal drugs.

B. Ovalle-Magallanes

Berenice Ovalle-Magallanes received her PhD degree in Pharmacology at the Facultad de Química (FQ) of the Universidad Nacional Autónoma de México (UNAM) in 2016. After conducting research work at the Department of Pharmacology and Physiology of The Université de Montréal and the Department of Biology of the FQ, UNAM, she became interested in developing and applying in vitro cell-based models to elucidate mechanisms of action of natural products. Since 2019, she has worked as Associate Professor and project group leader under the Young Researchers Program at UNAM. Her research interests include molecular targets in liver and skeletal muscle (using murine and human cells) that regulate glycaemia and cholesterol synthesis and studying potential natural products–drug interactions from edible and medicinal plant species used in Traditional Mexican Cuisine and Medicine.

M. Figueroa

Mario Figueroa received his B.S. degree in Pharmaceutical and Biological Chemistry in 2004 and his M.S. (2006) and PhD (2009) degrees in Chemistry from the Universidad Nacional Autónoma de México (UNAM) under the mentorship of Professor Rachel Mata. Then, from 2009 to 2013, he worked as a Postdoctoral Fellow at Lehman College, NY, USA, and The University of North Carolina at Greensboro, NC, USA, in the laboratories of Professors Edward J. Kennelly and Nicholas H. Oberlies, respectively. During this period, Dr Figueroa discovered numerous new natural compounds with potential anticancer and anti-MRSA properties from plants and fungi. In 2013, Dr Figueroa returned to UNAM as Associate Professor at the Facultad de Química, and in 2021, he became a Full Professor. His research focuses on the chemistry and biology of fungi and bacteria from unexplored sources in Mexico.

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1. Introduction

Diabetes mellitus (DM) is a metabolic disorder characterized by chronic hyperglycemia resulting from insulin secretion and resistance defects. According to the International Diabetes Federation, DM is one of the major leading challenges of the 21^st century. In 2021, it was estimated that more than 537 million adults lived with diabetes, causing over 6.7 million deaths that year. The global prevalence of DM is expected to rise to 700 million by 2045. Type II DM (TIIDM) accounts for ∼90% of all cases, and obesity and lifestyle are the major risk factors for developing the disease.¹

TIIDM results from insulin resistance and pancreatic β-cells dysfunction, which can lead to micro and macro-vascular complications, often the major causes of mortality among patients.¹ Thus, the goal of the treatment of TIIDM is to control blood glucose as measured by glycated hemoglobin, preventing these complications. Usually, the therapy is patient-focused and depends on comorbidities and other factors, but most patients receive combined treatments. The American Diabetes Association and other organizations have designed a few combined-therapy schemes.² Drug treatments and insulin therapy are prescribed if the higher blood glucose levels cannot be regulated with diet, exercise, and standard drugs.

The most relevant drugs for TIIDM management and their risk/benefit profiles are summarized below:

(i) The biguanide metformin, the first-line oral blood glucose-lowering agent, decreases glucose production in the liver and improves insulin sensitivity, primarily by activating adenosine monophosphate-activated protein kinase (AMPK). Metformin is linked to the traditional herbal medicine Galega officinalis L. (Fabaceae), rich in galegine, another biguanide. It has been widely used to treat the symptoms of diabetes in Europe since the 18^th century. The ability of galegine to lower blood glucose was discovered in the 1920s, and after that, some galegine derivatives, including metformin, were prepared, and used to control diabetes during the 1930s. Later, metformin was revived in the search for antimalarial agents in the 1940s and, during clinical tests, proved helpful in lowering blood glucose. In 1994 was approved in the US as an antidiabetic agent.^3,4 Metformin possesses a good risk/benefit profile, as it induces minor side effects and has a low chance of provoking hypoglycemia or weight gain. For such characteristics, the American Diabetes Association and the European Association for the Study of Diabetes recommend metformin as initial antihyperglycemic therapy for patients with TIIDM.^3,4 Nonetheless, in one in three diabetic patients receiving metformin experiments drowsiness, vomiting, anorexia, diarrhea, nausea, and dyspepsia. These side effects can be overcome using extended-release preparations. Additional precautions should be taken when prescribing elderly patients or with severe renal insufficiency, as metformin can cause vitamin B12, folic acid deficiency, and lactic acidosis. The main problem to overcome when using metformin is its lost of efficacy when TIIDM progresses to a state of low insulin levels, limiting its use in patients with advanced pancreatic β-cells dysfunction.^3,4

(ii) Sulfonylureas promote insulin secretion by blocking K_ATP channels in pancreatic β-cells.³ Second-generation sulfonylureas glipizide, glimepiride, and glibenclamide are prescribed as second-line treatment options for the management of TIIDM. Although these are potent and effective antidiabetic drugs, concerns arise as they provoke hypoglycemia. Also, their combination with other medications, such as beta-adrenergic antagonists, aspirin, allopurinol, fibrates, and other antidiabetic agents like insulin, could increase the risk of hypoglycemic shock. These risks are diminished with a closure monitoring of glycemia and drug–drug interactions. However, typical side effects like weight gain, headaches, dizziness, and nausea provoke treatment abandonment and failure, thus leading to the prescription of another secretagogue medication, such as meglitinides, a class of non-sulfonylurea drugs, or others.³

(iii) Thiazolidinediones improve insulin sensitivity in different body tissues by acting as peroxisome proliferator-activated receptor (PPAR) agonists.³ The prescription of drugs in this class, i.e., troglitazone, rosiglitazone, and pioglitazone, is restricted and limited in most countries due to a poor risk/benefits profile. For instance, the significant side effects of these drugs include weight gain, edema, and hepatic steatosis.³ Although troglitazone was withdrawn from commercialization due to hepatotoxicity and other reports indicate rosiglitazone and pioglitazone induce cardiovascular toxicity and bladder cancer, respectively, recent analyses suggest these safety issues should be addressed again to assess risk/benefit when thiazolidinediones are prescribed.³

(iv) Sodium–glucose cotransporter type 2 (SGLT2) inhibitors, like the gliflozins, which were inspired by phlorizin, a dihydrochalcone glucoside found in many Rosaceae species of the genus Malus Mill, Pyrus L., and Fragaria L. Phlorizin is an inhibitor of SGLT1 and SGLT2 because it competes with D-glucose for binding to the carrier; this action reduces renal glucose transport, lowering the amount of glucose in the blood in an insulin-independent mechanism. As a result, glucose is excreted in the urine.^3,5 SGLT2 inhibitors canagliflozin, dapagliflozin, and empagliflozin are well tolerated and have demonstrated their capacity to modestly reduce body weight, blood pressure, and glucotoxicity by antioxidant mechanisms and to improve liver and pancreatic β-cells functioning. Common side effects such as genital mycosis and urinary tract infections, and less frequently, urosepsis and pyelonephritis appear. Therefore, SGLT2 inhibitors are regarded as an excellent alternative to use in combination with other antidiabetic agents, such as dipeptidyl peptidase-IV (DPP-IV) inhibitors.^3,5

(v) α-Glucosidase inhibitors hinder the digestion of complex carbohydrates to monosaccharides in the small intestine. The drugs used in therapeutic are natural products or derived from natural products. Thus, acarbose, a pseudo-tetrasaccharide, is a natural microbial product produced from cultures of Actinoplanes utahencis Couch (Micromonosporaceae), which possesses nitrogen between the first and second glucose molecule and has a structure like an oligosaccharide. It was introduced in 1990 for the treatment of TIIDM.⁶ Voglibose and miglitol are structurally unrelated to glucosidase substrates, possessing piperidine-3,4,5-triol and tetraol-cyclohexane moieties, respectively. Streptomyces hygroscopicus Jensen (Streptomycetaceae) produce valiolamine, the precursor of voglibose. Miglitol is produced from nojirimycin, first reported in 1966 as metabolite of S. roseochromogenes Sugai (Streptomycetaceae), and later from deoxynojirimycin, isolated from the roots of mulberry (Morus Alba L., Moraceae). Mulberry is a traditional Chinese herb with a long history of use.^6,7 Furthermore, mulberry leaves are widely used for treating diabetes. Miglitol has the potential as a therapeutic drug against obesity.^3,8 Acarbose, miglitol, and voglibose increased glucagon-like peptide-1 (GLP-1), therefore potentiating insulin secretion. Besides reducing postprandial hyperglycemia, acarbose also induces weight loss and lowers total postprandial triglycerides. Significant side effects of α-glucosidase inhibitors are related to gastrointestinal discomforts, such as nausea, abdominal pain, bloating, flatulence, and diarrhea.³

(vi) Incretin therapies (IT), comprise glucagon-like peptide-1 (GLP-1) receptor agonists and dipeptidyl peptidase-IV (DPP-IV) inhibitors. The GLP-1 receptor agonists (e.g., exenatide and liraglutide) stimulate insulin secretion and suppress glucagon synthesis. The first agonist of GLP-1 used in therapeutics was exenatide, a peptide isolated from the saliva of the Gila monster.³ This peptide is a potent antidiabetic agent that reduces body weight combined with metformin. Liraglutide also induces weight loss when given with metformin or a sulfonylurea and decreases glycosylated hemoglobin and systolic pressure. Both drugs induce mild gastrointestinal effects, and GLP-1 analogs, in general, are contraindicated in renal failure. Two significant setbacks in their use are that they must be given in the form of injection and that their cost makes them unavailable for most patients in developing countries.³

GLP-1 acts within 2–4 min and is rapidly inactivated by the DPP-IV enzyme. DPP-IV inhibitors delay the degradation of GLP-1 and the glucose-dependent insulinotropic peptide (GIP). Gliptins are the main class of DPP-IV inhibitors (sitagliptin, saxagliptin, linagliptin, and alogliptin). These drugs reduce glycosylated hemoglobin and have a low risk of hypoglycemia without affecting body weight. GLP-1 receptor agonists and DPP-IV inhibitors improve pancreatic β-cell physiology and reduce apoptosis-associated glucotoxicity. Furthermore, IT improves cardiovascular markers related to dyslipidemia, hypertension, and obesity, reasons for which IT accounts for 30% of antidiabetic drugs in clinical development. In particular, the use of DPP-IV inhibitors has grown significantly during the last decade. Despite its benefits, IT induces side effects such as infections in the respiratory tract, headaches, skin lesions, bladder infections, gastrointestinal problems, and nasopharyngitis, as well as raises concerns about the risk of developing pancreatic cancer. These undesired actions justify the search for new DPP-IV inhibitors.³

(vii) Finally, insulin, alone or in combination with other drugs, is prescribed if proper blood sugar levels are not reached with lifestyle changes and other medications.³ Insulin itself is a natural product. Currently, several insulin formulations possess different pharmacokinetic and pharmacodynamic profiles with high potency and low risks of hypoglycemia. Most prescribed insulin formulations must be given by injection (except for inhaled), which provokes lipohypertrophy due to repeated use of the same area for subcutaneous injection, thus leading to low insulin absorption and deficient glycemic control. Furthermore, insulin formulations induce weight gain, which is detrimental to TIIDM control, formulation excipients are potential allergens, and medication costs have increased exponentially in the last decade.³ These factors lead to treatment abandonment and the appearance of potentially deadly complications, such as diabetic ketoacidosis. Therefore, combination therapies using adequate antidiabetic agents of those mentioned above would contribute to improving insulin therapy.³

Understanding the intricate mechanisms implicated in TIIDM has prompted the search for novel therapeutic agents, including some crucial enzymes for glucose homeostasis. This review will focus on specific enzymes that play a critical role in the pathophysiology of diabetes. The enzymes selected were DPP-IV considering that IT accounts for 30% of antidiabetic drugs in clinical development, and some of its inhibitors (e.g., sitagliptin, saxagliptin, linagliptin, and alogliptin) are now leading drugs for treating diabetes, after insulin and metformin.³ Diacylglycerol acyltransferase (DGAT) is a critical enzyme in triacylglycerol synthesis (TAGS). The intensified activity of DGAT in patients with TIIDM can cause β-cell dysfunction. Therefore, selective DGAT inhibitors might be essential in managing TIIDM and other metabolic diseases. Fructose 1,6-bisphosphatase-1 (FBPase), an enzyme involved in gluconeogenesis, is relevant in glucose metabolism. In TIIDM, gluconeogenesis significantly contributes to an excess of glucose. Reducing this excess would mitigate high glucose concentrations in the blood and tissues. Glucokinase (GK) in the pancreatic β-cells serves as a glucose sensor, increasing insulin secretion as blood glucose rises. Fructokinase or hexokinase (KHK) is an enzyme that converts fructose into fructose 1-phosphate. The activation of the latter enzyme leads to the formation of uric acid, which causes inflammation in the cells of pancreatic islets and, thus, insulin resistance. The inhibition of this pathway could control hyperglycemia. No therapeutic agents affect DAGT, TAGS, FBPase, GK, or KHK, although some compounds targeting these enzymes are now in clinical studies.³ As pointed out in the above paragraphs, natural antidiabetic drugs from plants or other natural sources have inspired the development of some therapeutic agents in the past.⁹ Therefore, it is highly probable that other natural products targeting the selected enzymes but DPP-IV yield new drugs for treating diabetes.

Finally, it is important to point out that the enzyme-aimed treatments usually induce reversible inhibition, covalent changes of the objective enzymes, or bind non-covalently but so tightly that their inhibition is irreversible. In any case, the desired pharmacological action is achieved. One crucial advantage of targeting enzymes in drug discovery is that the required assays are usually simple, using biochemical experiments capable of analyzing enzyme activity. Enzyme-based drug discovery can also benefit from crystallography analysis of the enzymes and in silico assays.¹⁰ The assays must be completed with proper kinetics studies, demonstrating reagent and response stability.

2. Methods

A systematic and comprehensive search for literature was carried out in various scientific databases, including Google Scholar, PubMed, ScienceDirect, Scopus, and Springer Link, to obtain the desired information available in journals and books from 2000–2022. Searching keywords were “dipeptidyl peptidase IV inhibitors”, “diacylglycerol acyltransferase inhibitors”, “fructose 1,6-bisphosphatase inhibitors”, “glucokinase activators”, “hexokinase inhibitors” and “fructokinase inhibitors” over 14000, 680, 470, 680, and 80 hits, respectively, were found. After that, a comprehensive refinement using the keywords “Diabetes mellitus”, “medicinal plants”, “phytochemicals”, and “natural products”, alone and in combination, was performed. Synthetic and in silico studies and activity data from extracts or fractions were excluded. Chemical structures were drawn using ChemBioDraw Ultra 14.0 (Cambridge Software), and protein structures were rendered with PyMOL (Schrödinger).

3. Dipeptidyl peptidase IV inhibitors

3.1. Structure of DPP-IV

DPP-IV is a multifunctional type II glycoprotein of the prolyl oligopeptidase family. The enzyme cuts peptides with penultimate Pro or Ala residues from the N-terminus of at least 21 bioactive peptides.¹¹

The GIP and GLP-1 are the most relevant substrates of DPP-IV concerning TIIDM, but there are other biological ligands. DPP-IV is an integral membrane protein ubiquitously distributed and expressed in intestinal, kidney, vascular endothelium, immune system, liver, and pancreas cells.¹¹

Human DPP-IV is a homodimer with two subunits (PDB code: 4J3J; Fig. 1). Each subunit comprises 766 amino acids (110 kDa), with the β-propeller and the α/β-hydrolase domains surrounding a space containing the active site. Functionally, these two domains are extracellular (amino acids 29–766). The structure has two channels between the hydrolase and propeller domains next to the catalytic site, through which substrates and ligands could contact the active site. DPP-IV also possesses a small tail (intracellular, amino acids 1–6) and a transmembrane region (amino acids 7–28). The eight-bladed β-propeller domain (amino acids 55–497) consists of four-stranded antiparallel β-sheet motifs containing a short helix with a double glutamate residues motif (Glu205 and 206), inserted into the first sheet of the fourth propeller blade, and an S2-loop (Ser349-Glu361). The α/β-hydrolase domain contains a central β-sheet constituted by eight β-strands (six parallel and two antiparallels) surrounded by 12 α-helices, assembled by residues 506–766 at the C-terminus, and a short stretch of the N-terminal sequence (residues 39–51).^11,12 The C-terminal region contains a hydrophobic pocket (Tyr547, Ser630, Tyr631, Val656, Trp659, Asp663, Tyr666, Asn710, Val711 and Gly740). The active site, located in a large cavity at the interface of the two domains, contains a catalytic triad formed by Ser630, Asp708, and His740. The C-terminal loop of DPP-IV is essential for both dimer formation and optimal catalytic efficacy. Five disulfide bridges stabilize the protein: four are present in the β-propeller region between Cys444/447, 385/394, 454/472, and 328/339, and the last at the α/β-hydrolase domain between Cys649 and 762.¹¹

DPP-IV is produced as a monomer; nevertheless, it needs dimerization for its proteolytic activity. The body has two DPP-IV isoforms: the membrane-bound DPP-IV, composed of full-length amino acids, and soluble DPP-IV (residues 49–766), whose cytoplasmic and transmembrane regions are absent. The soluble form exerts its proteolytic action by clipping dipeptides from the penultimate position of its substrates.

Most inhibitors bind to the DPP-IV-GLP-1 interacting site as protein–protein interaction inhibitors. Residues from the hydrolase and propeller domains participate in binding some inhibitors, such as valine–pyrrolidine and a few gliptins. The Glu residues 205 and 206 form salt bridges to the free amino group of the substrates. Glu205 and Glu206 are situated in a small horizontal helix (residues 201–207), which narrows the active site, leaving room for only two amino acids, making DPP-IV a dipeptidyl peptidase. The spatial arrangement of serine 630 (active site) to glutamic acids 205 and 206 in DPP-IV makes these two amino acid residues the most critical feature for the alignment of the peptide before cleavage. According to the site of binding, the commercial DPP-IV inhibitors belong to three classes (Fig. 2).¹³

Class 1 (e.g., vildagliptin and saxagliptin) interacts with subsites S1 (a hydrophobic pocket containing Ser630, Val656, Trp659, Tyr662, Val711, and Asn710) and S2 (Arg125, Arg669, Glu205 and 206, Phe357, and Arg358). In the case of saxagliptin, a few studies, including NMR, suggest that its inhibition of DPP-IV most likely involves a reversible covalent enzyme–inhibitor complex formation.

Class 2 (e.g., alogliptin and linagliptin) attach to subsites S2, S1, S1′ (Phe357, Tyr547, Pro550, Ser630, and Tyr residues 631 and 666), and S2′ (Tyr547, Trp629, Ser630 and His740).

Class 3 (e.g., sitagliptin and teneligliptin) binds to extensive site S2 (Phe357, Arg358, Ser209, and Val207), S2, and S1 subsites. In general, the N-terminus of the substrate peptide or inhibitor is recognized by Glu205, Glu206, and Ser630 cleaves at the N-terminus penultimate position. Glu205 and Glu206 amino acids form salt bridges and hydrogen bonds with the ligands.

For class 2 and 3 inhibitors, interactions with Tyr547 and Trp629 and with Phe357 and Tyr666, respectively, are pivotal for their DPP-IV inhibitory activities. X-ray analyses have determined the structures of DPP-IV and DPP-IV-inhibitors complexes.¹⁴

3.2. Implication of DPP-IV in diabetes

The L-cells of the gut release the incretin hormone GLP-1 within minutes in response to food intake in two primary biological forms: GLP-1 amide and GLP-1. The latter is the most abundant under physiological conditions. Soluble DPP-IV rapidly degrades GLP-1 (half-life shorter than 2 min) into inactive GLP-1 amide. GLP-1 induced an increment of intracellular cAMP upon binding to its G-protein-coupled receptor on the pancreatic β-cells (Fig. 3). Then, after downstream signaling is accomplished, insulin secretion increases (incretin effect) and regulates postprandial glucose. In addition, GLP-1 inhibits glucagon secretion, regulates satiety and appetite, and increases α- and β-cell insulin sensitivity and gene expression. In diabetic patients, DPP-IV is widely expressed at the surface of epithelial and endothelial cells, causing the rapid degradation of GLP-1 (7–36) to GLP-1 (9–36) (Fig. 3).^15–19

On the other hand, GIP is synthesized and secreted from K-cells of intestinal epithelium located in the proximal duodenum. DPP-IV terminates its insulinotropic action, cleaving the N-terminal dipeptide of GIP (1–42) to inactive GIP (3–42). GIP in adipose tissue stimulates fatty acid synthesis, enhances insulin-stimulated incorporation of fatty acids into triglycerides, increases insulin receptor affinity, and increases insulin-stimulated glucose transport. Hence, GIP is considered more relevant for treating obesity and does not affect glucagon secretion (Fig. 3).^19,20

Concerning TIIDM, GLP-1 is more relevant because it accelerates glucose-induced insulin secretion. The increment of the half-lives of GLP-1 and GIP by DPP-IV inhibitors lowers fasting and postprandial hyperglycemia due to the increment of insulin secretion in a glucose-dependent manner and a decrease of excessive glucagon secretion observed in TIIDM. In addition, DPP-IV inhibitors are weight-neutral and well-tolerated.^18,19

3.3. Assay methods

In general, drug screening for DPP-IV inhibitors has followed a target-based strategy using enzymatic assays, which give a precise mode of action. Unfortunately, these assays cannot detect off-target or undesirable pharmacological effects. The assays to discover DPP-IV inhibitors use recombinant enzymes, usually human or porcine, combined with colorimetric or fluorometric detection methods.^21,22 The procedures are rapid, simple, sensitive, reliable, and usually performed with commercial kits. The colorimetric assay is based on the hydrolysis by DPP-IV of Gly-Pro-p-nitroanilide to yield a constant chromophore (4-nitroaniline) in the presence of the potential inhibitor.²¹ The production of 4-nitroaniline is quantified using UV-VIS spectroscopy in a microplate reader at λ_max 360 and 460 nm. The fluorometric assay uses a fluorogenic substrate [Gly-Pro-aminomethylcoumarin (AMC)] to measure DPP-IV activity. Cleavage of the peptide bond by DPP-IV releases the AMC group, resulting in fluorescence that can be measured using an excitation wavelength of 350–360 nm and an emission wavelength of 450–465 nm. In addition, other fluorescent probes with the function of aggregation-induced emission have been used to screen DPP-IV inhibitors.²³ The results for these bioassays are usually expressed as % of inhibition or IC₅₀ values.

Simple and rapid colorimetric and electrochemical methods using gold nanoparticles as the probe have been developed to discover DPP-IV inhibitors; however, they lack sensitivity.^24,25 Other methods for detecting DPP-IV inhibitors are liquid chromatography coupled to mass spectrometry and thin-layer chromatography-bioautography-mass spectrometry combined with a liquid chromatography-mass spectrometry-controlled purification system.^26–28

Over the past few years, in silico techniques (quantitative structure–activity relationship, QSAR), ligand-based and structure-based virtual screening, pharmacophore modeling, molecular docking, molecular dynamic, and ADMET predictions have been widely used in the search for compounds with DPP-IV inhibitory properties.^29,30

Finally, bioassays based on cell cultures or small animal studies, which are more relevant to humans, are currently being developed. In most assays, diprotin A (IPI), hemifumarate, or some gliptins are employed as positive controls.

3.4. Natural products with DPP-IV inhibitory properties

Natural DPP-IV inhibitors have been the topic of several review articles.^31–40 Some of these reviews describe mainly the testing of plant extracts or in silico studies.^{29,30,37,40,41} The evaluation of plant extracts is beneficial from the point of view of traditional medical practices, in which the efficacy of the treatments is due to the synergistic action of the mixture of compounds present in an extract (polypharmacy) or the combination of several herbs. An excellent review regarding the inhibitory action on DPP-IV activity of plant-based polyherbal formulations and nano phytomedicines was recently published.³⁷ Therefore, a comprehensive review focusing solely on natural enzymatic inhibitors would benefit the community. The natural DPP-IV inhibitors discovered during 2000–2022 are summarized and categorized in Tables 1–4 These include the compound type and name, inhibitory effect, positive control used in the enzymatic evaluation, natural sources, and the corresponding references. Unfortunately, the diversity of non-standardized assays employed, the lack of positive control in some studies, and the non-disclosure of inhibitors' concentration range used in evaluations preclude any critical structure–activity relationship analysis.

4. Diacylglycerol acyltransferase inhibitors

4.1. Structure of DGAT

Diacylglycerol acyltransferase (DGAT), an endoplasmic reticulum (ER)-membrane-bound enzyme, belongs to the membrane-bound O-acyltransferase (MBOAT) family. DGAT catalyzes the conversion from diacylglycerol (DAG) to triacylglycerol (TAG) in the glycerol phosphate pathway (Fig. 4). This enzyme has two isoforms, DGAT-1 and DGAT-2, present in white adipose tissue. DGAT-1 is mainly found in the small intestine, and DGAT-2 in the liver. The selective inhibition of DGAT-1 is actively investigated due to its potential for treating obesity and TIIDM. DGAT-2 may be a target for treating DGAT-2-related liver diseases, e.g., hepatic steatosis, hepatic injury, and fibrosis.¹¹⁷

DGAT isoforms are encoded by two different genes widely expressed in mammals. In humans, the DGAT-1 gene, found on chromosome 8, has 10.62 kb and 17 exons. This gene encodes a protein of ∼500 residues in most species with a calculated molecular mass of ∼55 kDa. The Dgat2 gene encodes a ∼42 kDa protein with a ‘hairpin-like’ structure. To date, the crystal structure of DGAT-2 has not been reported. The structure and catalytic mechanism of DGAT-1 remained unknown until 2020, when Sui et al.¹¹⁸ determined the structure of dimeric human DGAT-1 by cryo-electron microscopy with a resolution of approximately 3.0 Å (PDB code: 6VYI). DGAT-1 forms a homodimer through N-terminal segments and a hydrophobic interface. Its active sites are located within the membrane region. The protein has nine transmembrane helices, eight forming MBOAT fold structure. The crystal structure of DGAT-1 complexed with the oleoyl-CoA substrate (PDB code: 6VZ1; Fig. 5) shows that the CoA moiety binds DGAT-1 on the cytosolic side. The acyl group is in a hydrophobic channel, leaving the acyl-CoA thioester bond near a conserved catalytic histidine residue.¹¹⁹ The highly conserved catalytic residues exposed laterally to the membrane bilayer allow lipid access to the active site.

As mentioned above, we will focus on reviewing selective inhibitors of DGAT-1 as a potential target for treating obesity and type II diabetes. Several DGAT-1 inhibitors with promising therapeutic potential for obesity (Fig. 6) have gone into clinical trials, for example, LCQ-908 (Novartis), AZD-7687 (AstraZeneca), and PF-04620110 (Pfizer). Unfortunately, none has successfully concluded the clinical trials, some due to adverse skin and adipose tissue effects.¹¹⁷

4.2. Assay methods

DGAT-1 is ubiquitously expressed; however, its remarkably high expression in adipocytes and small intestine enterocytes indicates that this enzyme has a significant role in lipid metabolism. In adipocytes, DGAT-1 catalyzes the formation of TAG via the condensation of DAG and a range of fatty acids (FA). Accordingly, the synthesis of TAG should be quantitatively affected by the inhibition of this enzyme in adipocytes. Moreover, TAG synthesis should also decrease due to the inhibition of DGAT-1 in enterocytes. This, in turn, would affect TAG levels in the systemic circulation following an oral lipid load because of the following. FA and monoacylglycerol in the GI tract are absorbed by enterocytes and are then condensed into TAG. This is crucial for the assembly and secretion of chylomicrons that transport TAG into the bloodstream for further use by high-energy-requiring tissues, such as muscle.¹¹⁷

There are several animal models to assess DGAT inhibition by small molecules (adipose TAG synthesis in rats, oral lipid tolerance in rats, diet-induced obesity mice, etc.).¹²⁰ Yet, the rational discovery of potential inhibitors is often achieved by a combination of in vitro assays and in silico methods (docking, pharmacophore, 3D-QSAR models, machine learning methods, etc.).^117,121

HepG2 cells are used for in vitro assays because lipoprotein assembly heavily relies on high concentrations of exogenous fatty acids in these cells. DGAT inhibition is measured by the secretion of apolipoprotein B (apoB) in cells incubated with oleic acid (OA) and the test compound.¹¹⁷ Other methods use microsomes from rat liver. In both cases, whole-cell esterification of DAG is then quantified by adding [¹⁴C]palmitoyl-CoA and measuring the [¹⁴C]TAG formed by TLC or liquid scintillation counting.¹¹⁷

An in vitro assay for human DGAT-1 (hDGAT-1) inhibitors involves the expression of the enzyme using a baculovirus expression system in Sf9 insect cells.¹²² Once the enzyme is extracted, serial dilutions of the tested compounds and [¹⁴C]oleoyl coenzyme A are added to the reaction mixture in a 96-well plate. The lipids are then separated, and the hDGAT-1 activity is quantified by counting aliquots of the upper heptane layer by liquid scintillography.¹¹⁷ Finally, an in vitro and in vivo DGAT-selective assay to discern between the roles of DGAT-1 and DGAT-2 in TAG synthesis uses isotope-labeled [¹³C₁₈] oleic acid. Then LC-MS-MS analysis is used to measure the [¹³C₁₈] oleoyl incorporated into TAG and DAG.¹²³

4.3. Natural products with DGAT-1 inhibitory properties

Many natural products from plants and microorganisms, have been reported to inhibit DGAT activity (Tables 5 and 6). Like DPP-IV inhibitors, the largest group of DGAT inhibitors are flavonoids and other polyphenols (Table 5). For example, the flavonoids isoliquilitin (197), liquiritigenin (201), isoliquiritigenin (16), liquilitin (202), and glabrol (203) were isolated from the roots of Glycyrrhiza uralensis Fisch. (Fabaceae) and tested in an in vitro DGAT inhibitory assay. Glabrol (203) showed non-competitive inhibition against DGAT with an IC₅₀ value of 8 μM. The use of licorice roots for treating obesity and TIIDM patients could be related to the DGAT inhibitory activity of this flavonoid.¹²⁴

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The prenylflavonoids, kurarinone (204), kurarinol (205), kushenols H (206) and K (207), and the chalcone kuraridin (200), isolated from the roots of Sophora flavescens Aiton (Fabaceae), inhibited DGAT activity in a concentration-dependent manner with IC₅₀ values of 10.9, 8.6, 142, 250, and 9.8 μM. Interestingly, compounds without C3-OH (200, 204, and 205) showed more potent inhibition than those with C3-OH (206, and 207).¹²⁵ In addition, kurarinone (204) also inhibited the de novo synthesis of TAG in Raji cells. Compound 200 has been used as a positive control in many in vitro studies.

A bioassay-guided fractionation of an extract of the stem bark of Erythrina senegalensis led to the isolation of a series of prenylflavonoids (210–217). From these, 8-prenylleutone (210), auriculatin (211), erysenegalensein O (212), erysenegalensein D (213), derrone (215), and 6,8-diprenylgenistein (217) inhibited DGAT activity with IC₅₀ values ranging from 1.1 to 15.1 μg mL⁻¹.¹²⁶

The chalcone xanthohumol (198) was also described as an inhibitor of DGAT.¹²⁷ Compound 198 is a potent inhibitor of apoB secretion and enhances apoB degradation. It was also reported that xanthohumol (198) decreases the expression of DGAT-1 mRNA, has hypoglycemic effects, and decreases hepatic triglyceride in KK-Ay mice.¹²⁸

The ellagitannins rugosins B (221) and D (222) and eusupinin A (223) isolated from rose petals inhibited the DGAT activity by 96%, 82%, and 84%, respectively, at 10 μM.¹²⁹ Remarkably, the activities of other microsomal enzymes were not affected.

Plant alkaloids have also shown important DGAT inhibitory activity as demonstrated for the quinolone alkaloids 257–265 isolated from the fruit of Evodia rutecarpa. The evocarpine derivates 258–260 showed concentration-dependent inhibition with IC₅₀ values below 24 μM.¹³⁰ Other examples are the alkamides 261–265 isolated from the fruits of Piper longum and P. nigrum (Piperaceae). From these, piperrolein B (264) was the most potent, with an IC₅₀ value of 20.1 μM.

Among the plant terpenoids active against the activity of DGAT,^131–135 the dimeric diterpenoid, aphadilactone C (245), isolated from the leaves of Aphanamixis grandifolia (Meliaceae), is the most potent natural DGAT-1 inhibitor discovered to date (IC₅₀ = 0.46 μM, selectivity index > 217 against DGAT-2).¹³³A. grandifolia, is a tree that grows in Asia, and its leaves and roots are used in traditional Chinese medicine. In addition, a SAR analysis of diastereoisomers 242–245 revealed that the active binding site of the DGAT-1 enzyme is very selective to the stereochemistry of the substrate.¹³³ Therefore, compound 245 could lead to the development of potent and selective DGAT-1 inhibitors.

A series of polyacetylenes, diterpenes, tridepsides, fatty acid derivatives, and phenylpyropenes from algae¹³⁶ and fungi have shown DGAT inhibitory activity (Table 6). Particularly, from a series of amidepsines, roselipins, and phenylpyropenes, isolated from the fungal strains Humicola sp. FO-2942 and FO-5969 (soil isolates), Gliocladium roseum KF-1040 (marine isolate), and Penicillium griseofulvum F1959 (soil isolate),^137–139 amidepsine A (270, IC₅₀ = 10.2 μM) and phenylpyropene C (280, IC₅₀ = 11.04 μM) were the most potent inhibitors, with a potency comparable to kuraridin (200). Compound 280 showed an inhibitory effect in rat liver microsomes (IC₅₀ = 10.2 μM) and Raji cells (IC₅₀ = 15.5 μM) and is commercially available as a standard for the screening of DPP-IV inhibitors. While amidepsines B (271) and D (273) showed specific inhibition on TAG formation in Raji cells.¹³⁷ In the group of the phenylpyropenes, 280 showed a noncompetitive pattern of enzyme inhibition.¹³⁹

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5. Fructose 1,6-biphosphatase inhibitors

5.1. Structure of FBPase

Glycogenolysis and gluconeogenesis (GNG) are the main pathways for glucose formation in the human liver. The latter, where three-carbon precursors such as pyruvate, lactate, and glycerol are converted into glucose, is the main process for glucose output in a fasting state: over 50% of glucose production is formed via GNG during persistent fasting or starvation; this is even higher in TIIDM patients.¹⁴⁶ The GNG pathway involves the conversion of fructose 1,6-bisphosphate (FBP) into fructose 6-phosphate (F6P), which is then converted into glucose after catalysis by glucose 6-phosphate isomerase and glucose 6-phosphatase. The reaction from FBP to F6P is catalyzed by fructose 1,6-bisphosphatase (FBPase), a rate-limiting enzyme (Fig. 7). In mammals, two genes encode FBPase: FBP1, mainly found in the liver and kidney, and FBP2, found exclusively in muscle tissue.¹⁴⁷ FBP1 and FBP2 overall identity is ∼77%, and the regions involved in binding the substrate, regulatory molecules, and magnesium are almost identical. The physiological role of muscle FBPase remains unclear because muscle tissue has no glucose 6-phosphatase activity and is thus nongluconeogenic. Probably, muscle FBPase is needed for glycogenesis from lactate. The liver form of FBPase regulates blood glucose levels in the liver and kidney. Compared to the liver, the kidney is a minor source of de novo synthesized glucose except during extreme fasting, and, reportedly, in a diabetic state.¹⁴⁷ In this context, human liver FBPase has been considered a potential therapeutic target for the treatment of TIIDM. In patients and animal models of TIIDM, the expression levels and activities of FBPase were upregulated in the liver, indicating the importance of this enzyme for increasing blood glucose.

FBPase is a cytosolic homotetramer, where each monomer (36.7 kD) presents a substrate (FBP) binding site and an AMP allosteric regulatory site within 28 Å of the active site (Fig. 8).¹⁴⁸ The enzyme has two conformational states, active (R) and inactive (T). A conformational change from R to T or the stabilization of T result from the allosteric inhibition of AMP. Another allosteric binding site was recently found for the inhibitor OC252.¹⁴⁹ Also, the enzyme undergoes competitive substrate inhibition by fructose 2,6-bisphosphate.

The development of FBPase inhibitors as potential antidiabetic agents remains a challenging task. The difficulties arise from the complex hydrophobic region on the subunit interface of FBPase and the highly charged substrate-binding site. Potent and selective competitive and uncompetitive inhibitors are thus scarce.¹⁴⁷ Pharmaceutical companies and academic laboratories have investigated a wide array of noncompetitive (allosteric) inhibitors of the AMP binding site in FBPase (Fig. 9).¹⁵⁰ Some synthetic FBPase inhibitors effectively decrease hepatic glucose production and blood glucose levels in diabetic animal models. However, only AMP mimetic MB07803, a phosphonic acid-containing thiazole prodrug of MB07729, has reached phase II in clinical trials.

5.2. Assay methods

Since its original isolation from rabbit liver and kidney in 1943,¹⁵¹ the activity of FBPase has been assayed (a) spectrophotometrically by following the triphosphopyridine nucleotide (TNP) reduction or the reduction of NADP⁺ to NADPH in the presence of glucose 6-phosphate isomerase and glucose 6-phosphate dehydrogenase,^151–153 or by the release of inorganic phosphate (P_i) from FBPase using the Malachite Green assay¹⁵² or by the of Fiske and Subbarow¹⁵⁴ or Tausky and Shorr¹⁵⁵ methods; (b) radiometrically by measuring the release of ³²P_i from fructose l,6-[l-³²P]P₂;¹⁵⁶ and (c) by LC-ESI-MS analysis.¹⁵⁷

In addition, FBPase from pig kidneys and sheep liver have been purified and used for assays, and their complete amino acid sequences have been determined.¹⁵⁸ The homology of all mammal enzyme forms extends to those from plants, yeast, and Escherichia coli, indicating that FBPase has undergone minimal changes throughout evolution, even though regulation of the enzyme's activity differs in different cell types. In 1993, the isolation of the cDNA for the human liver enzyme, the construction of a plasmid for its expression in E. coli, and purification and characterization of the recombinant enzyme was reported, as well as the expression of mutant enzymes by site-directed mutagenesis of active-site residues.¹⁵⁸ The human enzyme has been used in the same protocols mentioned for the rabbit FBPase.^159–161 Recently, a new assay, the PicoProb™ FBPase inhibitor screening kit by BioVision, Inc., was developed to discover novel inhibitors. Briefly, human FBPase hydrolyzes fructose 1,6-bisphosphate to fructose 6-phosphate, which oxidizes and reacts with a fluorogenic probe that is detected at λ_Em/Ex = 535/587 nm.¹⁶²

5.3. Natural products with FBPase inhibitory properties

Several synthetic FBPase inhibitors bearing distinct scaffolds have been identified via either high-throughput screening of compound libraries or structure-guided design of AMP mimetics.¹⁶² These include: (1) uncompetitive inhibitors (anilinoquinazoline derivatives, pseudo-tetrapeptide, and quinoline derivatives); (2) non-competitive phosphorous based inhibitors (purine derivatives, benzimidazole derivatives, thiazole derivatives, and oxazole derivatives); and (3) other non-competitive inhibitors (indole derivatives, benzoxazole derivatives, various N, O, S containing heterocycles derivatives, sulfonyl derivatives, and phytochemicals). Among these, only the AMP mimetic MB07803 has advanced to phase II clinical trials. In contrast, natural products with FBPase inhibitory activity have been barely studied (Table 7). In the following lines, we will discuss the most active natural inhibitors discovered to date.

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Significant FBPase inhibition was reported for the flavonoids, baicalein (27), scutellarein (290), herbacetin (298), and gossypetin (293), with mean IC₅₀ = 29, 38.2, 8.7, and 24 μM, respectively. The positive control, AMP, exhibited an IC₅₀ value of 2.6 μM.¹⁶² Herbacetin (298), a natural flavonoid containing –OH groups at C-3 of ring C, at C-4′ of ring B, and at C-5, C-7, and C-8 of ring A, is found in flaxseed and other plants. This natural product also shows antihyperglycemic and antihyperlipidemic activities thus protecting against the effects of chronic high-fat diet consumption (associated with insulin resistance).¹⁶² Moreover, the potential of 298 for the management of TIIDM has been acknowledged because of its interaction with aldose reductase and the insulin receptor. Interestingly, ingesting flaxseed oil or flaxseed supplementation has proven to reduce glucose levels and glycosylated hemoglobin concentration and increase insulin sensitivity in humans.¹⁶²

Antitumor, anti-inflammatory, antioxidant, and antidiabetic properties have been reported for baicalein (27),¹⁶² a component of the Chinese plant Scutellaria baicalensis. Scutellarein (290), found in the perennial herb Scutellaria lateriflora, is the hydrolyzed, and highly soluble form of scutellarin. Scutellarin has been used in China since 1984 to treat acute cerebral infarction and paralysis induced by cerebrovascular diseases. Scutellarin is mainly absorbed in the form of 290, which is likely the actual bioactive component.¹⁶² Gossypetin (293), is usually found in the flowers and the calyces of Hibiscus sabdariffa. This plant has traditionally been used as a sedative and for treating nervous disorders. In addition to its anxiolytic and antidepressant activities, this flavonoid is associated with antimicrobial, antiatherosclerotic, and anti-inflammatory effects.¹⁶²

The flavonoid 8-prenylquercetin (296) showed potent α-glucosidase and FBPase dual inhibitory activity, with IC₅₀ values of 4.38 and 3.62 μM, respectively.¹⁶²

Finally, (+)-usnic acid (299), a naturally occurring dibenzofuran derivative found in several epiphytic lichen species, inhibits both human liver and pig kidney FBPases in vitro with IC₅₀ values of 0.37 mM and 0.93 mM, respectively.¹⁷⁰

6. Glucokinase activators

6.1. Structure of GK

Glucokinase (GK) is a 52 kDa monomeric enzyme comprising 465 residues arranged in two domains, a large and a small α/β domain, separated by a deep cleft that functions as the active site for phosphorylation. The small domain comprises of residues 66–202, and helix C (or C-terminus) contains residues 442–465. The large domain encloses the rest of the protein (Fig. 10).¹⁷¹ Crystallographic analysis of GK structure has revealed that this enzyme has three conformations: super-open, open, and closed.¹⁷² The super-open conformation is characterized by an opening angle of 100° between the large and small domains and a disordered loop formed by residues 151–180. In this state, the small domain residues are disorganized and unable to bind glucose. Thus, the super-open conformation predominates at low glucose concentration.^173,174 In the absence of either glucose or activator, GK adopts a “super-open”, inactive apo-conformation. The closed conformation, the most compact, has an opening angle of 40°. Glucose phosphorylation is then carried out in the presence of ATP. This process is facilitated by Lys169 residue, which acts as an acid catalyst and enhances the binding of GK with both ATP and glucose.¹⁷⁵ Once the phosphorylation is completed, GK shifts to the open conformation to release catalytic product, G6P (and ADP), and slowly equilibrates to the super-open conformation. The catalytic cycle re-starts if glucose binds to the open conformation (fast cycle) and slows if glucose concentrations decrease (slow cycle) as this state favors super-open conformation, thus explaining the sigmoidal saturation curve for glucose of GK.¹⁷³

6.2. GK and glucose homeostasis

After carbohydrate ingestion and hydrolysis in the gastrointestinal tract, dietary monosaccharides are transferred from the luminal intestinal surface across cellular membranes and then transported to other cells. Monosaccharides are phosphorylated by ATP-dependent phosphorylating enzymes known as kinases which are stored or used as substrates for cellular metabolic processes. Hexokinases (EC 2.7.1.1) phosphorylate hexoses such as 2-deoxyglucose, mannose, glucosamine, fructose, and glucose, usually at the hydroxyl at the 6′-position in a six-carbon sugar, using ATP as the phosphate donor (Fig. 11). Four hexokinase isoforms have been identified in mammals: hexokinase 1 (HK1, I or A), hexokinase 2 (HK2, II or B), hexokinase 3 (HK3, III or C), and hexokinase 4 or glucokinase (HK4, IV, D or GK).^174,176 GK is not inhibited by G6P, and its concentration is proportional to the extracellular glucose levels. Accordingly, GK activity can be regarded as a glucose sensor in several tissues, particularly those involved in glucose homeostasis, such as the liver and pancreas.^176–178 For instance, in hepatocytes, GK promotes glycogen synthesis, particularly during the postprandial state. On the other hand, in pancreatic β cells, GK regulates the entry of glucose into the glycolytic pathway and its subsequent metabolism, therefore controlling glucose-dependent insulin secretion.^179–186.

The hyperglycemic state characteristic of DM is due to a combination of impaired glucose utilization and enhanced liver gluconeogenesis, both regulated by GK. Thus, GK activators have been considered an alternative to treat TIIDM. These pharmacological agents are predicted to have a dual effect when controlling glucose metabolism by increasing insulin secretion in pancreatic β-cells or stimulating glycolysis and glycogen synthesis in the liver.¹⁷⁷ Development of synthetic GK activators have focused on identifying compounds that bind to allosteric sites in hepatic GK rather than in pancreatic β-cells GK, as in the latter case numerous early-developed GK activators induced severe hypoglycemia.^171,179,187

6.3. Assay methods

Most of the experimental evidence on GK activation by any compound derives from sub-chronic in vivo experiments using tissues from rodents with either chemically- or diet-induced diabetes.¹⁸⁸

Early GK activity determinations were carried out in cell cultures using D-(U-¹⁴C)-glucose and measured by scintillation counter. GK activity was proportional to radioactivity due to marked G6P.¹⁸⁹ Nowadays, GK activity is assessed by using a NADPH-coupled assay with G6P dehydrogenase. The end product of the reaction is 6-P-gluconate and NADPH, which possesses an intrinsic fluorescence intensity; GK activity is proportional to the fluorescence intensity of NADPH.¹⁹⁰ This procedure is useful for calculating the binding affinity constants (K_D) and kinetic characteristics of GK activators.¹⁹¹ Another method is the luminescence assay, which monitors the ATP consumption associated with GK catalytic activity and uses a detection system based on luciferin and luciferase to produce a luminescent signal that is proportional to ATP. Therefore, the GK activity is inversely proportional to the ATP concentrations in the GK reaction.¹⁹⁰ GK activity and expression is monitored by immunological and molecular biology techniques.^192,193 Finally, hepatic mRNA expression of GK can be determined using quantitative real-time PCR, using appropriate primers for GK amplification.^194,195

Since there are several methods to assess GK activation and GK protein expression, there are no consensus on units used to express the results, which makes it difficult to compare the efficacy of pure compound and extracts.

6.4. Plant products activators of GK

There are a few recent reports describing the effect of medicinal plant extracts on GK expression or activity using different assays in vitro or in vivo.^181,196 These reports do not specify the mechanism of GK activation. Furthermore, GK hepatic activation data is accompanied by information on increased insulin secretion and/or glycogen synthesis. Many natural products can modulate GK activity (Table 8) or expression (Table 9). As in previously discussed enzymes, flavonoids are largest group of GK activators for by catechin (6) administered orally (20 mg kg⁻¹ day⁻¹ per 6 weeks) to STZ-diabetic rats also provoked an augmentation in insulin levels (8.6 ± 1.9 μmol mL⁻¹) and liver glycogen (49.2 ± 5.3 mg per g tissue) compared to diabetic control (8.2 ± 1.4 μmol mL⁻¹ and 21.5 ± 3.8 mg g⁻¹, respectively).²⁰⁴ Likewise, flavanones hesperidin (20) and hesperetin (300) almost triplicated glycogen levels when given to STZ-diabetic rats (45.37 ± 3.14 mg per g tissue and 17.12 ± 1.31 mg per g tissue, respectively) compared to diabetic controls (26.19 ± 2.42 mg per g tissue and 10.75 ± 0.82 mg per g tissue, respectively) when administered either during 30 or 45 days at doses of 100 or 40 mg kg⁻¹ day⁻¹.^207,208 When amentoflavone (301) was given to obese and diabetic mice (40 mg kg⁻¹ day⁻¹ per 8 weeks), it increased insulin secretion (21.39 ± 3.62 μU mL⁻¹ respect to diabetic control, 16.12 ± 2.97 μU mL⁻¹).²⁰² In the same animal model, a daily dose of 100 mg kg⁻¹ per 30 days of polymethoxylated flavone tangeretin-1 (302) significantly increased insulin (15.98 ± 1.49 μU mL⁻¹) and liver glycogen levels (48.05 ± 4.14 mg per g tissue) compared to diabetic controls (6.26 ± 0.53 μU mL⁻¹ and 15.77 ± 1.45 mg per g tissue, respectively).²¹⁰ Similar effects were observed for the methoxylated isoflavone biochanin A (297) but at a lower dose (10 mg kg⁻¹ day⁻¹ per 45 days), with insulin and liver glycogen levels duplicated after treatment (13.34 ± 0.65 μU mL⁻¹versus 7.55 ± 0.39 μU mL⁻¹, and 45.56 ± 3.76 mg per g tissue versus 22.87 ± 2.18 mg per g tissue, respectively).²⁰³ Kaempferol (56, after 12 weeks of treatment, 50 mg kg⁻¹ day⁻¹) did not increase plasma insulin levels in diabetic mice but significantly augmented liver glycogen.²⁰⁹In silico studies predicted that flavonoids 301, 56, and 68 are allosteric GK activators.^235–237

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Among terpenoids, D-limonene (303) showed a significant hypoglycemic activity when given to STZ-diabetic rats (50–200 mg kg⁻¹ day⁻¹ per 45 days). GK activity was slightly increased compared to diabetic control.²¹² Geraniol (304) (200 mg kg⁻¹ day⁻¹ per 45 days), displayed moderated GK activation, since it increased the number of insulin-positive cells per islet in STZ-diabetic rats, thus significantly augmenting insulin levels.²¹³ Interestingly, trans-anethole (326), a phenylpropanoid, also ameliorates the β-cells architecture in a similar model (80 mg kg⁻¹ day⁻¹ per 45 days), which increases glycogen level (44.47 ± 4.22 mg per g tissue respect to 22.73 ± 0.02 mg per g of diabetic control).²¹⁴

The diterpenoid andrographolide (305) administered for 21 days (10 mg kg⁻¹) did not affect insulin levels in STZ-diabetic rats; instead, it significantly increased glycogen levels with respect to diabetic animals (54.2 ± 4.7 mg per g tissue versus 33.4 ± 4.1 mg per g tissue), therefore suggesting extrapancreatic effects of GK activation.²¹⁵ Rebaudioside A (306) showed potential GK activities, as its administration to STZ-diabetic rats (200 mg kg⁻¹ day⁻¹ per 45 days) protects pancreatic β-cells by decreasing oxidative stress, increasing insulin levels and glycogen content (45.01 ± 3.67 mg per g tissue versus 22.50 ± 1.82 mg per g tissue observed in diabetic control).²¹⁶

There are two contradictory reports on the activity of sesquilignans tatanans A–C (323–325). Ni and co-workers (2011) indicated that these compounds were potent GK allosteric activators in vitro,²¹⁷ while Xiao and co-workers (2013) refuted these findings.²³⁷ Thus, further investigations are required to clarify whether tatanans are GK activators, particularly under in vivo conditions.

Betulin (309),²²⁰ as well as 18β-glycyrrhetinic (307),²¹⁸ asiatic (308),²¹⁹ ursolic (332),²³⁴ and oleanolic (310)²²¹ acids augmented GK activation and increased glycogen levels in diabetic rodents. Compounds 307 and 308 induce a significant augmentation of insulin levels, whilst triterpenoid 310 activated GK in a similar manner as insulin (used as positive control). Docking studies predicted that these triterpenoids, and the related β-amyrin, could be GK allosteric activators, but showed different interaction pattern than flavonoids.²²⁰ The predominant observed interactions were hydrophobic.²³⁵ The tetranortriterpenoid meliacinolin (311), daily administered for 28 days at a dose of 4 mg kg⁻¹, increased both insulin and glycogen levels (17.86 ± 1.98 mg per g tissue and 3.34 ± 0.42 μU mL⁻¹, respectively) when compared with STZ-diabetic controls (9.25 ± 1.65 mg per g tissue and 1.42 ± 0.18 μU mL⁻¹, respectively) and in a similar fashion than glibenclamide, used as positive cotrol.²²²

GK activators modulates other metabolic pathways, as the amount of G6P is available for subsequent reactions. For example, in hepatocytes, G6P is dephosphorylated to glucose by glucose 6-phosphatase (G6Pase), a rate-limiting gluconeogenetic enzyme, which have an opposite effect than GK; thus GK and G6Pase have opposite actions and regulate each other activity.²³⁸ Kaempferol (56),²⁰⁹ berberine (191),¹⁹² amentoflavone (301),²⁰² rebaudioside A (306),²²² 18β-glycyrrhetinic acid (307),²³⁸ meliacinolin (311),²²² bellidifolin (316), methylswertianin (317),²²⁴ polydatin (331),²³³ and ursolic acid (332)²³⁴ activate GK and reduce both G6Pase activity and fasting glycemia.

In the liver, GK activators increase the activity of glycogen synthase (GS) and inhibit glycogen synthase kinase 3 (GSK-3β) promoting hepatic glycogen synthesis. There are reports of such activities for catechin (6),²⁰⁴ hesperidin (20),²⁰⁸ amentoflavone (301),²⁰² tangeretin-1 (302),²³⁹ and tyrosol (321).²¹¹ Finally, G6P can be used in the last step of glycolysis controlled by pyruvate kinase (PK), an enzyme whose activity is reduced in diabetic rodent models. Several GK activators, such as amentoflavone (301),²⁰² tangeretin-1 (302),²³⁹ asiatic acid (308),²¹⁹ and betanin (313),¹⁹⁹ activate PK.²⁴⁰

Since insulin controls hepatic GK expression, secondary metabolites able to active insulin pathways could increase GK levels. Thus, protein levels of different elements of the phosphatidylinositol-3-kinase (PI3K)/PKB canonical pathway can be monitored, including insulin receptor substrate-1 (IRS-1), PDK-1, Akt, and GSK-3β.²⁴¹ Curcumin (329) increased phosphorylation of insulin receptor (IR), IRS-1, PI3K and Akt.^193,222 Bellidifolin (316) and methylswertianin (317) increased the expression levels of IR-subunit α, IRS-1, and PI3K.²²⁴ Amentoflavone (301) also augmented the levels of phosphorylated Akt,²⁰² and berberine (191) increased PI3K expression.¹⁹²

Other studies on GK involved a series of in silico evaluations (molecular dynamics simulations and ADMET studies) of prenylated flavonoids, revealing their potential as GK activators.¹⁹⁷

7. Fructokinase inhibitors

7.1. Structure of KHK and metabolic actions

Fructokinase (ketohexokinase, KHK, E.C. 2.7.1.3) is a 39 kDa dimeric enzyme belonging to the ribokinase-like superfamily with a high substrate specificity (Fig. 12). KHK exists in two isoforms: fructokinase A (KHK-A) and fructokinase C (KHK-C). Each monomer of KHK isoforms has two distinct domains: a central α/β-fold and a four-stranded β-sheet. The α/β-fold comprises a nine-stranded β-sheet flanked on each side by five α-helices. There is one active site per subunit in a cleft between the α/β-domain, and the four-stranded β-sheet that forms the dimer interface. AMP-PNP (adenylyl-imidodiphosphate) and fructose bind to opposite sites at the active-site cleft, being fructose-binding closer to the four-stranded β-sheet.²⁴²

Although KHK-A is ubiquitous in most tissues, KHK-C is mainly expressed in the liver, kidney, and intestines and it is the main enzyme involved in fructose metabolism.^243,244 KHK catalyzes the phosphorylation of fructose to fructose-1-phosphate (F1P) using ATP and a potassium ion (K⁺) (Fig. 13). KHK-C has a higher affinity for fructose phosphorylation than KHK-A.^245,246 Unlike GK, KHK catalytic actions are not controlled by insulin but by fructose.²⁴⁷

Fructose enters rapidly into the liver using the glucose transporter-2 (GLUT-2) and is converted to F1P by KHK. F1P is a substrate of aldolase B, resulting in phosphotrioses used to synthesized glucose, lactate, and fatty acids. Increased fructose intake mainly due to the uncontrolled ingestion of sweetened beverages and highly processed foods, leads to an augmentation in triglyceride synthesis, as glycolysis intermediates are accumulated and then to glycerol-3-phosphate.²⁴⁷ Due to fructose phosphorylation, KHK triggers the depletion of ATP and facilitates uric acid liver and fat accumulation.²⁴⁸ Since KHK is not inhibited by its catalytic product nor regulated allosterically, there is a need to search for KHK inhibitors, as an alternative to diminishing fatty acids and triglyceride accumulation.²⁴⁹

7.2. Assay methods

The activity of KHK can be assessed in vitro using several assays. Lee and co-workers²⁴⁹ reported a 3-step cell-free enzymatic reactions which monitors (i) ADP formation related to fructose phosphorylation, (ii) pyruvate increase as a result of ADP and P-enol pyruvate reaction, and finally (iii) measures the stoichiometric decrease in NADH absorbance at 340 nm.²⁴⁹ Maryanoff and coworkers reported a method using HepG2 cells, which, after being exposed to test compounds, received fructose and were further processed to obtain cell lysates that were used to quantify F1P levels by means of liquid chromatography-mass spectrometry.^250,251 Another method to assess KHK activity is to determine fructose-induced ATP depletion. This assay is conducted in cell lysates of HepG2 cells overexpressing KHK-C. ATP level, measured using an ATP assay kit, and ATP depletion is proportional to KHK activity.²⁴⁹

7.3. KHK inhibitors from medicinal plants

In 2016, Le and coworkers published reported 406 plant extracts and 1200 purified phytochemicals activity against KHK-C. Promising result were obtained with Angelica archangelica (Apiaceae), Scutellaria baicalensis (Lamiaceae), Petroselinum crispum (Apiaceae), and Garcinia mangostana (Clusiaceae). Fourteen secondary metabolites derived from these extracts were also analyzed (Table 10).²⁴⁹

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The activity of the compounds ranged from 16.1 to 93.4% of inhibition. Among the prenylated chalcones 336 and 337, the methoxy isobavachalcone (336) was the most potent (IC₅₀ = 0.2 μM related to depletion of hepatic ATP).²⁴⁹ This metabolite also inhibits adipogenesis and contribute to triglycerides homeostasis, acting as a non-competitive inhibitor of acyl-coenzyme A: cholesterol acyltransferase (ACAT), and as an inhibitor of diacylglycerol acyltransferase (DGAT).²⁵² Four flavones (338–341) have been reported to be FK inhibitors, oroxylin A (338) is the most potent (42.1% inhibition, IC₅₀ = 8.2 μM). Other polyphenols such as cratoxyarborenone E (342) (61.7% inhibition, IC₅₀ = 1.0 μM), and a mixture of α-/γ-mangostin (318/343, 90.4% inhibition, IC₅₀ = 1.5 μM) showed prominent inhibitory activities against KHK.²⁴⁹ Prenylated coumarins such as osthenol (344), osthole (345) and the 2-methoxylated derivative (346) also displayed significant KHK inhibition, being 345 twice as potent as 344. Osthole (345) is a hypoglycemic molecule that acts as a dual PPARα/γ and AMPK activator, thus inducing the activities of acyl-CoA synthetases and acetyl CoA carboxylase, respectively, without lowering blood insulin or lipid levels.^253,254 Along these lines, prenylated xanthones α-mangostin (318) and γ-mangostin (343) evaluated as a mixture, inhibited almost all KHK activity. Investigations using STZ-diabetic animals suggest that 318 is a hypoglycemic agent that induces insulin secretion, increased GK activity, reduces fructose-1-6-biphosphatase and glucose-6-phosphatase activities,^226,255 and increases leptin and glucose transporter-4 (GLUT-4) expressions. γ-Mangostin has also shown to increase glucose uptake and diminishes saccharide digestion inhibiting intestinal α-amylase and α-glucosidase enzymes.²⁵⁶ A common structural feature among the most active compounds is the presence of a prenylated side chain (either an isoprenyl, a geranyl, or a 1,1-dimethylallyl moiety). Therefore, further studies on natural products should be conducted to assess binding interactions with this enzyme using computational or crystallographic tools.²⁵⁷

8. Concluding remarks and future directions

This review has summarized the natural compounds reported in the literature from 2000 to 2022 having in vitro or in vivo DPP-IV, DGAT, FBPase, GK, or KHK activities (Tables 1–10). No therapeutic agents affecting DGAT, FBPase, GK, or KHK have been developed yet.

The high number of reported flavonoids (Fig. 14) as inhibitors or activators of such enzymes and their non-selective actions is not surprising because it is well-documented that flavonoids are polypharmacological (polyvalent) agents; this is, they can interact with two or more molecular targets simultaneously, thus affecting multiple biological processes.²⁵⁸ Therefore modulating several mechanisms involved in glucose homeostasis, acting through different enzymes.²⁵⁸ Pharmacological polyvalence can arise from conserved binding sites in different proteins recognized by different compound scaffolds or from the compound's structural motifs (auxophoric moieties) prone to trigger multiple ligand-target interactions.²⁵⁹ In this context, flavonoids are listed as pan-assay interference compounds (PAINS) because most of them are phenolic. There are multiple reports where PAINS have shown to give false positives in assays, thus misleading investigations and generating several overstatements on observed activities. Therefore, a detailed structure–activity relationship analysis is necessary to identify such compounds' pharmacophoric and auxophoric moieties, thus leading to rational drug design. PAINS presence in drugs is common, as shown by the fact that almost 5% of FDA-approved drugs contain PAINS-recognized scaffolds, comprising both natural products and synthetic drugs. Concerns regarding PAINS in natural products must be overcome or confirmed using multiple validated and complementary experimental tools, such as in vitro, in silico, and in vivo testing.^259–261

For instance, we found significant differences in some compounds' inhibitory activity against DPP-IV during this revision. These discrepancies might be attributed to variations in the methodology, enzymes, and conditions to run the tests. Furthermore, different authors' docking analyses of many natural products evaluated in vitro, particularly flavonoids, yielded contradictory results (Table 1). This situation precluded any conclusion about the structure–activity relationship among the same type of compounds. Perhaps the next generation of DPP-IV inhibitors from natural sources could emerge from foods, as many articles emphasize the relevance of the consumption of dietary proteins from soybean, rice, amaranth, draft beer, broccoli, Brassica spp, macroalgae, yam species, and beans, which upon digestion, yield inhibitory DPP-IV peptides (Fig. 14).

Those compounds listed in Tables 5 and 6 targeting DGAT showed inhibitory activities from 0.46 to >250 μM. The lack of therapeutic drugs aiming DGAT made that the positive control used in most cases was kuridin (200), a natural chalcone from Sophora flavescens, which along with aphadilactone C (244), glabrol (203), kurarinone (204), kurarinol (205), amidepsine A (270), and phenylpyropenes C (280) are the most active inhibitory natural products against DGAT. Aphadilactone C (244) warrants further investigation as a DGAT-1 inhibitor due to its high selectivity, potency, and unusual scaffold.

Regarding the FBPase inhibitors, all compounds reviewed were flavonoids, but usnic acid (299), a well-known lichen metabolite. The most active were baicalein (27), scutellarein (300), herbacetin (308), and gossypetin (303), whose activities were comparable with the positive control, AMP (IC₅₀ of 2.6 μM). There is an urgent need for a more simple and robust in vitro analysis for detecting active compounds.

GK activators (Tables 8 and 9) could bind to the allosteric site or modulate protein expression. Flavonoids amentoflavone (301), kaempferol (56), and quercetin (68) are known allosteric activators of GK. In contrast, triterpenoids 307–318 could activate GK by an allosteric mechanism different from that of flavonoids. It is relevant to mention that secretagogue natural products that interfere with the insulin pathway could modify GK activity, as documented for compounds 6, 56, 29, 191, 297, 301, 302, 316, and 317. Finally, natural KHK inhibitors, such as 318, 336, 343, 344, and 354, showed in vivo hypoglycemic activity and regulated triglyceride synthesis.

Interestingly, multi-target activities displayed by natural products are an asset in developing drugs to treat complex and multi-factorial diseases such as TIIDM.²⁶² This review shows that natural products could improve glucose homeostasis by modulating related proteins and pathways. Therefore, a single chemical identity could effectively improve insulin resistance and hyperglycemia due to different etiologies. In the context of the worldwide spread of diabetic complications, an intensive search for new lead compounds for developing novel pharmacological therapeutics is extremely important, whereas clinical trials are still lacking. In this regard, it is important to remember that calculated IC₅₀ using in vitro models assesses ligand binding affinities to enzymes rather than ligand-target kinetics or ligand-target removal. Therefore, a straightforward correlation between preclinical (in vitro or in vivo) and clinical potency is not to be expected.²⁶³

For instance, in vivo level of receptor occupancy required to achieve maximal efficacy is related to the target's mechanism of action and its downstream signaling components rather than the quantity of drug needed for activation. Thus, compounds activating multiple elements in a signaling pathway have better chances to induce the desired pharmacological effect by acting additively or synergically, as proven for natural compounds. However, in vivo pharmacokinetics significantly impact the observed potency, as most of the natural compounds herein described possess low bioavailability. Thus, effective clinical concentrations will be hard to achieve without making structural modifications that could improve their absorption, reduce their protein-binding or modify their elimination, well-known optimizations conducted to obtain drugs from bioactive molecules.

Further studies are necessary to fully understand the mode of action of natural products targeting multiple enzymes, establish structure–activity relationships, therapeutical concentrations, and required scaffold modifications to improve pharmacokinetics or potency.

The continuous research of plants used in traditional and complementary medicines and other natural sources might lead to the discovery of new successful antidiabetic drugs, as before with metformin, acarbose, exenatide, and gliptins, all based on natural products, all belonging to different categories of natural products. As shown in Fig. 14, alkaloids and terpenoids could be good candidates for developing modulators of DPP-IV, DGAT, FBPase, GK, or KHK.

9. Conflicts of interest

There are no conflicts to declare.

10. Acknowledgements

This work was supported by grants of Dirección General de Asuntos de Personal Académico (DGAPA, IA205621 and IN203523), Consejo Nacional de Ciencia y Tecnología (CONACyT, CY011226), and L’Oréal-UNESCO-AMC-CONALMEX awarded to R.M.

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