Revalorizing Lignocellulose for the Production of ...

Send Orders for Reprints to [email protected] Current Medicinal Chemistry, 2018, 25, 1-9

1

REVIEW ARTICLE

Revalorizing Lignocellulose for the Production of Natural Pharmaceuticals and Other High Value Bioproducts Congqiang Zhang1,* and Heng-Phon Too1,2,* 1

Biotransformation Innovation Platform (BioTrans), Agency for Science, Technology and Research (A*STAR), Singapore; 2 Department of Biochemistry, National University of Singapore, Singapore

ARTICLE HISTORY Received: March 03, 2017 Revised: June 02, 2017 Accepted: August 25, 2017 DOI: 10.2174/0929867324666170912095755

Abstract: Lignocellulose is the most abundant renewable natural resource on earth and has been successfully used for the production of biofuels. A significant challenge is to develop cost-effective, environmentally friendly and efficient processes for the conversion of lignocellulose materials into suitable substrates for biotransformation. A number of approaches have been explored to convert lignocellulose into sugars, e.g. combining chemical pretreatment and enzymatic hydrolysis. In nature, there are organisms that can transform the complex lignocellulose efficiently, such as wood-degrading fungi (brown rot and white rot fungi), bacteria (e.g. Clostridium thermocellum), arthropods (e.g. termite) and certain animals (e.g. ruminant). Here, we highlight recent case studies of the natural degraders and the mechanisms involved, providing new utilities in biotechnology. The sugars produced from such biotransformations can be used in metabolic engineering and synthetic biology for the complete biosynthesis of natural medicine. The unique opportunities in using lignocellulose directly to produce natural drug molecules with either using mushroom and/or ‘industrial workhorse’ organisms (Escherichia coli and Saccharomyces cerevisiae) will be discussed.

Keywords: Lignocellulose, natural products, metabolic engineering, synthetic biology, secondary metabolites, drugs. 1. INTRODUCTION Lignocellulosic biomass or lignocellulose represents an enormous renewable resource (e.g. wheat stems, corn stover and wood shavings) with annual production rate of 150-170 x 109 tons [1]. Lignocellulosic biomass typically contains 50-80% complex carbohydrates consisting of C5 and C6 sugar units. Sugars exist in the form of two types of polymers - cellulose and hemicellulose. Structurally, cellulose exists in crystalline microfibrils that are attached to hemicellulose and is protected by surrounding lignin (Fig. 1) [2]. Cellulose chains are held tightly by numerous hydrogen bonds. Hemicellulose is linked to each other via covalent and *Address correspondence to these authors at the 8 Medical Drive, Blk MD7, Level 4, Department of Biochemistry, Yong Loo Lin School of Medicine, National University of Singapore, Singapore 117597; Tel: (65) 6516 3687; E-Mail: [email protected] and 61 Biopolis Drive, Proteos #04-13/14/15, Singapore 138673; E-mails: [email protected] or [email protected]

0929-8673/18 $58.00+.00

hydrogen bonds, to cellulose via hydrogen bonds and to lignin via ester linkages. Lignin is connected with hemicellulose but does not have a direct link with cellulose [3]. The structural complexity of lignocellulose in plants provides strength and rigidity but poses significant challenges to directly use it as carbon sources. To make full use of lignocellulose, pretreatments involving heat and corrosive chemicals (acids or bases) are required before enzymatic degradation of cellulose and hemicellulose. The present commercially available carbohydrate active enzymes (CAZymes) are inefficient and costly, accounting for significant cost contribution to the production of both biofuel [4] and other valuable compounds from lignocellulose. Thus, to effectively use lignocellulose, there is a demand to discover new sources of more efficient enzymes and to engineer more cost-effective enzymes and processes at industrial levels. Despite lignocellulose being recalcitrant to most enzymes, there are many species of naturally occurring

© 2018 Bentham Science Publishers

2 Current Medicinal Chemistry, 2018, Vol. 25, No. 00

Zhang and Too

Fig. (1). From lignocellulose to natural drug molecules. In lignocellulose, cellulose stores as crystalline microfibrils and is attached to hemicellulose. Cellulose and hemicellulose is further protected by surrounding lignin. In nature, lignocellulose can be degraded by diverse organisms, such as white-rot fungi, brown-rot fungi, termite and marine wood-boring isopod. Sugars obtained from lignocellulose hydrolysis can be biotransformed to produce valuable compounds, such as pharmaceuticals using microbial cells (such as Escherichia coli, Saccharomyces cerevisiae and fungi).

organisms which can efficiently use lignocellulose. Examples of such organisms include wood-degrading fungi (e.g. brown rot and white rot fungi) [5], wooddegrading bacteria (such as anaerobic bacterium Clostridium thermocellum) [6], some arthropods (e.g. termite and cockroaches [7]) and certain animals (such as ruminants and panda [8]). The biochemical pathways involved in wood decomposition in these organisms offer unprecedented opportunities in developing new generations of lignocellulose-degrading enzymes. From insights into their degradation mechanisms, it is possible to design and optimize better processes. In parallel, with the rapid developments in metabolic engineering and synthetic biology, it is now common to manipulate microbes to produce various natural products with complex structures beyond the small alcohol molecules (e.g. ethanol and butanol) from sugars. Such successful examples include artemisinin [9] and its precursor amorphadiene [10-12], taxol precursors [13, 14] and opioids (e.g. morphine and hydrocodone) [15-17]. For many years, sustainable production of secondgeneration biofuels using lignocellulose has been a main goal of research and development [18-20]. An attractive alternative is to harness lignocellulose for the biosynthesis and production of many other medicinal relevant compounds. This review intends to discuss the

recent studies in how lignocellulose can be used as a carbon source for the production of natural drugs in microbes using modern biotechnological tools. 2. BREAK DOWN LIGNOCELLUOSE As a consequence of the structural complexity of lignocellulose, effective breakdown of lignocellulose requires not only efficient cellulases and hemicellulases but also means to overcome the barriers imposed by the complex lignin structural matrix. Currently, in biofuel industry, pretreatment of the woody biomass is a prerequisite where heat and/or chemicals (acid, ammonia or ionic liquid) are used before enzymatic hydrolysis of cellulose and hemicellulose [21]. The pretreatments loosen the microstructure of cell walls and eliminate the strong interactions between various components in lignin to allow downstream enzymatic hydrolysis to occur efficiently [22]. In contrast, unmodified lignocellulose is naturally digested effectively by fungi and bacteria with synergistic participation of various enzymes (such as various carbohydrate active enzymes (CAZymes), lignin-degrading enzymes (FOLymes), and some cell wall loosening enzymes. In these processes, glucoside hydrolases (GHs) are the main enzymes that cleave glycosidic linkages present in cellulose and hemicellulose and GH-coding genes are abun-

Revalorizing Lignocellulose for the Production

dant and present in the vast majority of genomes corresponding to almost half of the enzymes classified in CAZy database [23]. GHs are assisted by polysaccharide esterases that remove methyl, acetyl and phenolic esters to function on hemicellulose and by other enzymes including lytic polysaccharide monooxygenases and polysaccharide lyases [24]. Lignin is modified by either secreted FOLymes (e.g. peroxidases and laccases) and/or oxidative radicals (hydroxyl radicals) [25]. These studies have inspired the industrial production and use of CAZymes in microbes and the use of natural biomass-degrading organisms as biocatalysts [21]. Thus, by understanding the mechanisms employed by these nature wood degraders, it is possible to develop enzyme cocktails to digest lignocellulose with limited or no pretreatments. 2.1. Wood-degrading Fungi Among the fungi and bacteria that are capable of degrading lignocellulosic biomass, basidiomycete white-rot and brown-rot fungi are the most efficient decomposers. White-rot fungi are capable of depolymerizing all three components (cellulose, hemicellulose and lignin) in woody cells. Despite derived from whiterot fungi, brown-rot fungi can only degrade polysaccharides and partially modify lignin. Recently, the comparative and functional genomics of white-rot and brown-rot fungi have elucidated the evolutionary relationships of the fungal families and have identified new types of lignocellulose degrading enzymes [24, 26], providing insights into the development of new processes to deal with lignocellulose materials. In particular, the genome of Schizophyllum commune, a white-rot fungus, has recently been sequenced. Genomic analysis indicates S. commune contains 16 FOLyme enzymes and 11 enzymes distantly related to FOLymes, along with one cellobiose dehydrogenase and two laccases. Unsurprisingly, S. commune possesses a more diverse assortment of FOLymes than brown-rot fungus Postia placenta and the fungi that do not have ligninolytic activities (such as Cryptococcus neoformans, Aspergillus nidulans, Neurospora crassa and Saccharomyces cerevisiae). It was found that the genome of S. commune contains 240 candidate GHs, 75 candidate glycosyl transferases, 16 candidate polysaccharide lyases and 30 candidate carbohydrate esterases [27]. Among all the basidiomycetes examined, S. commune has the most complete polysaccharide breakdown machinery with the largest number of polysaccharide lyases and glycoside hydrolases, especially rich in the glycosyl hydrolase families GH93 (hemicellulose degradation) and GH43 (hemicellulose and pectin degrada-

Current Medicinal Chemistry, 2018, Vol. 25, No. 00

3

tion), and the lyase families PL1, PL3 and PL4 (pectin degradation) [27]. Another white-rot fungus, Phanaerochaete chyrosporium secretes a range of enzymes such as lignin oxidases and peroxidases and its genome encodes more than 240 putative CAZymes (including 166 glycoside hydrolases, 14 carbohydrate esterases and 57 glycosyltransferases, comprising at least 69 distinct families) [28]. The ligninolytic system in white-rotters is attributed in part to extracellular oxidative enzymes, especially peroxidases, laccases, and other oxidases [29]. Intriguingly but still incompletely understood, brown-rot fungi are able to rapidly degrade cellulose in woody cells without the ligninolysis systems. It was hypothesized that the wood decomposition by brown-rot fungi (such as Serpula lacrymans) employs non-enzymatic disruption of lignocellulose with internal cleavage of cellulose chains catalyzed by highly localized hydroxyl free radical (·OH) [24, 30]. In the genome of Postia placenta, 242 CAZymes-encoding genes were identified but these do not possess the cellulose-binding domains and few glycoside hydrolases [29]. Elucidation of the degradative machinery in wood decay fungus potentially can facilitate the development of new generation of enzymes to break down lignocellulose. This is currently addressed by heterologously expressing and biochemically characterizing the functions of these enzymes. 2.2. Marine Wood Borer and Termite Guts Unlike other herbivores that rely on symbiotic gut microbes to digest lignocellulose, the marine woodboring isopod, Limnoria quadripunctata (or gribble), has an intestine devoid of microbes, indicating that this organism is able to natively produce the enzymes necessary for lignocellulose digestion. This hypothesis was confirmed by a study led by McQueen-Mason and his colleagues [29]. Analysis of expressed sequence tags (ESTs) indicated that there are 12 recognizable glycosyl hydrolase families belonging to CAZymes, of which the most abundant glycosly hydrolases are the GH7 family enzymes. GH7 enzymes, found only in fungi and protozoan mutualists in termites, are critical for cellulose depolymerization. The unique GH7 cellulase from L. quadripunctata (LqCel7B) has recently been produced in Aspergillus oryzae and Aspergillus niger and subsequently purified [31]. LqCel7B enzyme is a cellobiohydrolase and its activity remains or increases under conditions with high salt concentrations. In addition, it was hypothesized that hemocyanin, a highly abundant transcript in the hepatopancreas transcriptome, might be involved in the lignin degradation


as it can be readily transformed into phenoloxyidases via conformational changes induced by chemical treatment or limited proteolysis [29]. Unlike limnoriid wood borers, lignocellulose degradation in termite involves enzymatic activities of both the host and its mutualistic symbionts [32]. As a wooddegrading organisms, termites are far more capable and efficient than ruminants, they are able to digest up to 99% of cellulose and 87% of hemicellulose of wood [33]. The hydrolysis of cellulose is initiated by endoglucanases that are secreted by the salivary glands and facilitated by β-glucosidases. However, this process is far from complete possibly due to the absence of cellobiohydrolases and hemicellulolytic activities in the tubular midgut. The complete breakdown requires the help of more potent cellulases and hemicellulases (such as xylanases, arabinosidases, mannosidases and arabinofuranosidases) which are produced by bacterial symbionts [7]. Possibly, termites are not ideal organisms for the direct application in lignocellulose utilization. But termites are a reservoir of invaluable microbial symbionts and enzymes that have important biotechnological potentials. A better understanding of the wood-degrading processes in termites may help us to design a new generation of bioreactor with a consortia of symbionts [7]. 2.3. Anaerobic Cellulolytic Bacteria and Cellulosomes In addition to wood decay fungi and animals, there are groups of anaerobic bacteria, such as Clostridium thermocellum, that can also hydrolyze lignocellulose. These bacteria possess unique multi-enzyme complexes or cellulosome to degrade lignocellulose. Cellulosome is a very robust and high organized enzymatic system that can degrade lignocellulose very efficiently [34]. Cellulosome has two main types of building blocks, dockerin-containing enzymes and cohesioncontaining structural proteins or scaffoldins. Cohesins and dockerins are complementary modules that bind tightly to each other non-covalently. The thermophilic bacterium Clostridium clariflavum has as many as 160 enzymes in a single complex which includes numerous cellulases, CAZymes (notably, xylanases, pectinases, mannanases and xyloglucanases) [35]. Various cellulase and hemicellulase are well known for their synergistic catalytic capabilities. For example, exoglucanases, also known as cellodextrinases, would cleave more chain ends with the help of endoglucanases that concomitantly hydrolyze the oligosaccharides [36]. βglucosidases degrades the product of cellulase thus releasing the product inhibition on cellulase. In addition,

Zhang and Too

because of the physical proximity of the enzymatic components, substrate channeling effect in cellulosome makes it much more efficient than simple mixture of free enzyme solutions [37]. These studies in natural lignocellulose degrading species including fungi, animals and bacteria offer opportunities to develop next generation enzymatic systems for lignocellulose material biotransformation (Table 1). We can now concoct a combinatorial panel of enzyme mixtures tailored to digest different lignocellulosic biomass with very different structures. Even though still very challenging, engineering some natural degraders to convert lignocellulose into sugar or even the final products is promising as the availability of novel genetic tools have expanded greatly (such as zinc finger nucleases, CRISPR/Cas gene-editing tool) enabling precise manipulations of the genome [38]. Artificially designed cellulosome is yet another direction for lignocellulose utilization due to its flexibility and efficiency. Recent successful examples have lend further support that such a technology is indeed promising where the integration of a lignin-active enzyme into a designer cellulosome markedly enhanced degradation of plant-derived cellulose and xylan [39]. Traditionally, to produce metabolites at high yield, a single native or metabolically engineered strain of an organism is often used in monoculture fermentation conditions. An emerging expanded approach is to explore co-cultures where the metabolic burden can be distributed between various strains or different organisms and this allows specific modularization for optimization in ways not previously possible by traditional fermentation approaches [40-42]. 3. CONVERT SUGARS TO DRUGS Once sugars are obtained from lignocellulose, we can use them to produce complex natural products using microbes. Natural products produced by plants, bacteria, and fungi represent a rich source of bioactive compounds for drug discovery and development, and more than half of new drug chemicals approved by the Food and Drug Administration (FDA) in US are natural products or their derivatives and analogs [43]. Most bioactive compounds belong to three major classes, terpenoids (e.g. Taxol [13] and artemisinin [9]), alkaloids (e.g. morphine [17]) and flavonoids (e.g. naringenin [44, 45]). While the structural complexity renders chemical synthesis at high yield to be unattainable, microbial production of these compounds has emerged as an important renewable alternative approach facilitated by tools established in metabolic engineering and synthetic biology. Examples of some high-value



5

Table 1. A brief summary list for the enzymes found in lignocellulose degrading strains. GH – glycoside hydrolase, CAZymes - carbohydrate active enzymes, FOLymes - lignin-degrading enzymes. Wood-degrading Hosts/ Cellulosomes

Main CAZymes

Main FOLymes or oxidases

Reference

1

White-rot fungi e.g. chizophyllum commune

GH family 1, 2, 3, 5, 6, 7, 10, 11, 12, 26, 28, 43, 45, 51, 74, 61, 62, 88, 93, 105, 115

Laccases, glyoxal oxidase, Peroxidases, Cellobiose dehydrogenase, pyranose oxidase, glucose oxidase, benzoquinone reductase, alignocelluloseohol oxidases,

[9]

2

Brown-rot fungi e.g. Serpula lacrymans

GH family 1, 2, 3 , 5, 10, 12, 28, 31, 35, 47, 51, 61, 74, 92, 79

Multicopper oxidases, Glyoxaloxidases, Polyphenol Oxidase, Cellobiose dehydrogenase

[13]

3

Limnoriid wood borers, Limnoria quadripunctata

GH Family 5, 7, 9

Hemocynin

[14]

4

Termite and symbionts e.g. Nasutitermes spp.

GH family 1, 2, 3, 4, 5, 8, 9, 10, 11, 13, 16, 18, 20, 23, 25-28, 31, 35, 36, 38, 39, 42-45, 51-53, 57, 58, 65, 67, 74, 77, 88, 91, 92, 94, 95, 98, 103, 106, 109

No FOLymes were found

[7, 32]

5

Cellulolytic bacteria and Cellulosomes e.g. Clostridium clariflavum

The most highly expressed enzymes are GH family 48, followed by GH9 and GH5 for cellulose, and GH10, GH11 and GH26 for hemicellulose

No FOLymes were found

[35, 39]

No.

pharmaceuticals that have been produced by metabolic engineering and fermentations are discussed below, illustrating the prospects of revalorizing lignocellulose materials (Table 2). 3.1. Anti-malaria Artemisinin Artemisinin, produced by the plant Artemisia annua, is a sesquiterpenoid with potent antimalarial properties. Artemisinin-based combination therapies (ACTs) were recommended by World Health Organization to treat malaria caused by parasite Plasmodium falciparum. Tu Youyou [46] and her team initially identified its antimalaria activity and isolated pure artemisinin, were later awarded the Nobel prize in medicine in 2015. Commercially available artemisinin extracted from plant suffers from unstable supply, resulting in shortages and price fluctuation. Keasling and his colleagues have developed a Saccharomyces cerevisiae-based strain that produced artemisinic acid at a titer of 25 g/L [9]. Followed by a chemical conversion of artemisinic acid to artemisinin, they have developed an industrially viable process. In order to improve the yield and titer of artemisinic acid, the production of the precursor, amorphadiene, was first optimized with metabolic engineering tools in E. coli [10, 11, 47-49], yeast [12, 50, 51] and Bacillus [52]. In S. cerevisiae, native and modified versions of the genes (e.g. truncated HMG1) in the mevalonate (MVA) pathway were

overexpressed under the control of a strong galactoseregulated GAL1 promoter. In bacteria, the use of both native 1-deoxy-D-xylulose-5-phosphate pathway (DXP) pathway and heterologous MVA pathway have been explored for the production of amorphadiene. The highest amorphadiene production from the DXP pathway obtained was significantly lower than that of the mevalonate pathway despite DXP pathway has a higher theoretical yield [11]. 3.2. Anti-cancer Taxol Taxol (paclitaxel) represents one of the most clinically valuable natural products in the past decades. Paclitaxel was originally isolated from the bark of Pacific Yew, Taxus brevifolia. Later, Taxomyces andreanae, an endophytic fungus, isolated from the bark of Taxus brevifolia was shown to produce Taxol. Since then, there are about 200 endophytic fungi from various genera are known to produce Taxol [53]. However, many of these discoveries have yet to be successfully translated into industrial bioprocesses. Total chemical synthesis was certainly not an option as it is commercially unviable [54]. Therefore, it was imperative to develop alternative microbial sources of Taxol. In collaboration with Prof. Stephanopoulos, using a multivariant modular approach, we managed to produce about 1 g/L taxadiene, a precursor to Taxol, in Escherichia coli from glucose [13]. Further on, exploiting a ‘mutualistic


Zhang and Too

Table 2. A summary for selected natural drug molecules produced in microbial workhorses. No.

Drugs or Drug Precursors

Medical Application

Expressing Hosts

Starting Sources

Titer/ Yield

Reference

1

Artemisinic acid

Anti-malaria

Saccharomyces cerevisiae

Glucose

25 g/L

[9]

2

Taxadiene

Anticancer

Escherichia coli

Glucose

1 g/L

[13]

3

Oxygenated taxanes

Anticancer

Escherichia coli and Saccharomyces cerevisiae

Glucose

33 mg/L

[14]

4

Thebaine

Analgesic

Escherichia coli

glycerol

2.1 mg/L

[16]

5

Hydrocodone

Analgesic


glucose

0.3 µg/L

[17]

6

Opioids

Analgesic


Thebaine

131 mg/L

[17]

7

Noscapine

Anticancer


Norlaudanosoline

1.64 µM

[59]

microbial consortium’, Zhou and his colleagues produced about 33 mg/L oxygenated taxanes from glucose [14]. Here, the microbial co-culture was made up of Escherichia coli which produced taxadiene and Saccharomyces cerevisiae produced cytochrome P450 oxidases that converted taxadiene into oxygenated taxanes. The stability of such a consortium in bioreactors was achieved by designing a mutualistic relationship. All these promising studies demonstrated the production of Taxol sustainably by fermentation. Challenges remain in identifying downstream genes in the biosynthetic pathway and to enable the production at the commercial scales. 3.3. Analgesic and Anticancer Alkaloids Opioids are an important class of natural product molecules from opium poppy that include the analgesic morphine and the antitussive codeine. Currently, all natural opiates are derived from opium poppy [15]. However, like many agricultural production, poppy farming is susceptible to environmental factors such as pests, disease, and climate fluctuations that introduce instability and variability into the supply chain. A microbial-based process was recently developed by Christina and her colleagues to produce opioids from glucose [17]. To synthesize the final products, they engineered a yeast strain capable of producing (S)reticuline from glucose, which is a key intermediate for opiate alkaloids. A lot of efforts were directed at the biotransformation of (S)-reticuline to (R)-reticuline, an enzyme from Papaver bracteatum including 2-dehydroreticuline synthase (DRS) and 1, 2-dehydroreticuline reductase (DRR) enzyme (DRS-DRR). A chi-

meric engineering approach significantly increased salutaridine synthase activity and thus the yield from (R)reticuline to salutaridine. Two more enzymes were required to catalyze the conversion of salutaridine to thebaine. Thebaine was subsequently converted into hydrocodone and other opioids such as morphine. To date, semisynthetic production of artemisinin is the only natural drug that has entered the market, with the capability to meet up to one-third of global need [55]. With Taxol and opioids, the titers of oxygenated taxanes (30 mg/L) and hydrocodone (0.3 µg/L) obtained are still too low to achieve commercially viability currently (Table 2). The continual sequencing of genomes from different organisms (https://gold.jgi. doe.gov/) will enable the eventual identification of novel genes in biosynthetic pathways involved in the production of valuable natural drugs. Furthermore, the availability of such genetic sequences will provide an unprecedented opportunity to develop novel combinatorial modules of genes for engineering microbes toward these important natural drug molecules. 4. PERSPECTIVE AND CONCLUDING REMARKS With significant advances in the technical capabilities to degrade lignocellulose to sugars and to convert sugars into complex natural products, it is no longer a distant possibility to revalorizing lignocellulose for the production of high value natural products. Initial successful attempts at enhancing economic value of lignocellulose were in the production of biofuels. From such ventures, new approaches have been developed For example, consolidated bioprocessing (CBP) provides a


more efficient and more cost-effective method for production as it eliminates the requirement for exogenous enzymes and reduces the sugar inhibition on cellulases than bioprocesses involving separate hydrolysis and fermentation [18, 56]. Such an approach has yet to gain wide use in medicine production from lignocellulose. Currently, the main goal is to improve carbon yields and to elucidate the missing biosynthetic genes in the production of high value natural products. With recent advancements in the whole genome sequencing, metabolic engineering and synthetic biology, hitherto unknown biochemical pathways and genes are being discovered. This has enabled the recent high yield production of some of high-value substances and precursors where the consumption and cost of raw material may no longer be insignificant. And we believe it is more appealing and valuable to produce higher value compounds than only biofuels from lignocellulose if technically viable. Besides, the challenging scientific endeavor to valorize lignocellulosic waste, which accounts for a substantial proportion of the agroindustrial biomass, will also encourage the development and discovery of novel tools and offer new insights into the handling of lignocellulosic waste. Here, we foresee three approaches towards extending this application. First, metabolic engineering of mushrooms able to produce drug products like antitumor and immunostimulatory molecules directly from woody biomass [57]. Second, sophisticated engineering of industrial workhorse strains such as Escherichia coli and Saccharomyces cerevisiae by grafting the lignocellulose-degrading pathway together with biosynthetic pathways of drug molecules to produce heterologous drugs from lignocellulose. The example of complete biosynthesis of opioids in yeast sheds some light on this endeavor. Lastly, synthetic microbial consortium represents another promising direction [14, 58], for example, a fungus-bacterium mutualistic system can be designed where fungi provide bacteria with sugars by depolymerizing lignocellulose and the bacteria then convert the sugars into nutrients such as vitamins for fungi growth and simultaneously produce natural drugs. Unlike biofuels which require large volume of production to be competitive, natural drugs are high value-added products where scale of production is not as demanding. However, the intricate chemical nature of these natural drugs often require complex and multiple step biosynthesis. With the rapid progress in metabolic engineering and synthetic biology, designing and optimizing complex metabolic circuitry is no longer an unsurmountable objective. It is obvious then that lignocellulose material will serve as an important renewable


7

resource for the production of high value products like natural drugs or pharmaceuticals, beyond biofuels. CONSENT FOR PUBLICATION Not applicable. CONFLICT OF INTEREST The authors declare no conflict of interest, financial or otherwise. ACKNOWLEDGEMENTS Declared none. REFERENCES [1] [2] [3] [4]

[5] [6] [7] [8]

[9]

[10]

[11]

[12]

Pauly, M.; Keegstra, K., Cell-wall carbohydrates and their modification as a resource for biofuels. Plant J., 2008, 54, (4), 559-568. Sanderson, K., Lignocellulose: A chewy problem. Nature, 2011, 474, (7352), S12-14. Purchase, D. Fungal Applications in Sustainable Environmental Biotechnology. Springer, 2016. Zhang, C.; Qi, W.; Wang, F.; Li, Q.; Su, R.; He, Z., Ethanol from corn stover using SSF: an economic assessment. Energy Sources, Part B: Economics, Planning, and Policy, 2011, 6, (2), 136-144. Goodell, B.; Qian, Y.; Jellison, J.; American Chemical Society: Washington, DC, 2009; Vol. 982, pp 9-31. Blanchette, R.A.; Nilsson, T.; Daniel, G.; Abad, A.; American Chemical Society: Washington, DC, 2009; Vol. 225, pp 141-174. Brune, A., Symbiotic digestion of lignocellulose in termite guts. Nat. Rev. Microbiol., 2014, 12, (3), 168-180. Zhu, L.; Wu, Q.; Dai, J.; Zhang, S.; Wei, F., Evidence of cellulose metabolism by the giant panda gut microbiome. Proc. Natl. Acad. Sci. U. S. A., 2011, 108, (43), 1771417719. Paddon, C.J.; Westfall, P.J.; Pitera, D.J.; Benjamin, K.; Fisher, K.; McPhee, D.; Leavell, M.D.; Tai, A.; Main, A.; Eng, D.; Polichuk, D.R.; Teoh, K.H.; Reed, D.W.; Treynor, T.; Lenihan, J.; Fleck, M.; Bajad, S.; Dang, G.; Diola, D.; Dorin, G.; Ellens, K.W.; Fickes, S.; Galazzo, J.; Gaucher, S.P.; Geistlinger, T.; Henry, R.; Hepp, M.; Horning, T.; Iqbal, T.; Jiang, H.; Kizer, L.; Lieu, B.; Melis, D.; Moss, N.; Regentin, R.; Secrest, S.; Tsuruta, H.; Vazquez, R.; Westblade, L.F.; Xu, L.; Yu, M.; Zhang, Y.; Zhao, L.; Lievense, J.; Covello, P.S.; Keasling, J.D.; Reiling, K.K.; Renninger, N.S.; Newman, J.D., High-level semi-synthetic production of the potent antimalarial artemisinin. Nature, 2013. ZHANG, C.; Zou, R.; Chen, X.; Stephanopoulos, G.; Too, H.-P., Experimental design-aided systematic pathway optimization of glucose uptake and deoxyxylulose phosphate pathway for improved amorphadiene production. Appl. Microbiol. Biotechnol., 2015, 99, (9), 3825-3837. ZHANG, C.; Chen, X.; Stephanopoulos, G.; Too, H.-P., Efflux transporter engineering markedly improves amorphadiene production in Escherichia coli. Biotechnol. Bioeng., 2016, (113), 1755–1763. . Yuan, J.; Ching, C.B., Combinatorial engineering of mevalonate pathway for improved amorpha-4,11-diene production in budding yeast. Biotechnol. Bioeng., 2014, 111, (3), 608-617.

8 Current Medicinal Chemistry, 2018, Vol. 25, No. 00 [13]

[14]

[15] [16]

[17] [18]

[19]

[20]

[21] [22] [23]

[24]

[25]

[26]

Ajikumar, P.K.; Xiao, W.-H.; Tyo, K.E.; Wang, Y.; Simeon, F.; Leonard, E.; Mucha, O.; Phon, T.H.; Pfeifer, B.; Stephanopoulos, G., Isoprenoid Pathway Optimization for Taxol Precursor Overproduction in Escherichia coli. Science, 2010, 330, (6000), 70-74. Zhou, K.; Qiao, K.J.; Edgar, S.; Stephanopoulos, G., Distributing a metabolic pathway among a microbial consortium enhances production of natural products. Nat. Biotechnol., 2015, 33, (4), 377-U157. Thodey, K.; Galanie, S.; Smolke, C.D., A microbial biomanufacturing platform for natural and semisynthetic opioids. Nat. Chem. Biol., 2014, 10, (10), 837-844. Nakagawa, A.; Matsumura, E.; Koyanagi, T.; Katayama, T.; Kawano, N.; Yoshimatsu, K.; Yamamoto, K.; Kumagai, H.; Sato, F.; Minami, H., Total biosynthesis of opiates by stepwise fermentation using engineered Escherichia coli. Nat. Commun., 2016, 7, 10390. Galanie, S.; Thodey, K.; Trenchard, I.J.; Filsinger Interrante, M.; Smolke, C.D., Complete biosynthesis of opioids in yeast. Science, 2015, 349, (6252), 1095-1100. Salehi Jouzani, G.; Taherzadeh, M.J., Advances in consolidated bioprocessing systems for bioethanol and butanol production from biomass: a comprehensive review. Biofuel Research Journal, 2015, 2, (1), 152-195. Naik, S.N.; Goud, V.V.; Rout, P.K.; Dalai, A.K., Production of first and second generation biofuels: A comprehensive review. Renewable and Sustainable Energy Reviews, 2010, 14, (2), 578-597. Huang, R.; Su, R.; Qi, W.; He, Z., Bioconversion of lignocellulose into bioethanol: process intensification and mechanism research. Bioenergy Research, 2011, 4, (4), 225-245. Sanderson, K., Lignocellulose: A chewy problem. Nature, 2011, 474, (7352), S12-14. Kim, J.S.; Lee, Y.Y.; Kim, T.H., A review on alkaline pretreatment technology for bioconversion of lignocellulosic biomass. Bioresource technology, 2016, 199, 42-48. Cantarel, B.L.; Coutinho, P.M.; Rancurel, C.; Bernard, T.; Lombard, V.; Henrissat, B., The Carbohydrate-Active EnZymes database (CAZy): an expert resource for Glycogenomics. Nucleic Acids Res., 2009, 37, (Database), D233D238. Eastwood, D.C.; Floudas, D.; Binder, M.; Majcherczyk, A.; Schneider, P.; Aerts, A.; Asiegbu, F.O.; Baker, S.E.; Barry, K.; Bendiksby, M.; Blumentritt, M.; Coutinho, P.M.; Cullen, D.; de Vries, R.P.; Gathman, A.; Goodell, B.; Henrissat, B.; Ihrmark, K.; Kauserud, H.; Kohler, A.; LaButti, K.; Lapidus, A.; Lavin, J.L.; Lee, Y.H.; Lindquist, E.; Lilly, W.; Lucas, S.; Morin, E.; Murat, C.; Oguiza, J.A.; Park, J.; Pisabarro, A.G.; Riley, R.; Rosling, A.; Salamov, A.; Schmidt, O.; Schmutz, J.; Skrede, I.; Stenlid, J.; Wiebenga, A.; Xie, X.; Kues, U.; Hibbett, D.S.; Hoffmeister, D.; Hogberg, N.; Martin, F.; Grigoriev, I.V.; Watkinson, S.C., The plant cell wall-decomposing machinery underlies the functional diversity of forest fungi. Science, 2011, 333, (6043), 762-765. Kracher, D.; Scheiblbrandner, S.; Felice, A.K.; Breslmayr, E.; Preims, M.; Ludwicka, K.; Haltrich, D.; Eijsink, V.G.; Ludwig, R., Extracellular electron transfer systems fuel cellulose oxidative degradation. Science, 2016, 352, (6289), 1098-1101. Floudas, D.; Binder, M.; Riley, R.; Barry, K.; Blanchette, R.A.; Henrissat, B.; Martinez, A.T.; Otillar, R.; Spatafora, J.W.; Yadav, J.S.; Aerts, A.; Benoit, I.; Boyd, A.; Carlson, A.; Copeland, A.; Coutinho, P.M.; de Vries, R.P.; Ferreira, P.; Findley, K.; Foster, B.; Gaskell, J.; Glotzer, D.; Gorecki, P.; Heitman, J.; Hesse, C.; Hori, C.; Igarashi, K.; Jurgens, J.A.; Kallen, N.; Kersten, P.; Kohler, A.; Kues, U.; Kumar,

Zhang and Too

[27]

[28]

[29]

[30] [31]

[32]

T.K.; Kuo, A.; LaButti, K.; Larrondo, L.F.; Lindquist, E.; Ling, A.; Lombard, V.; Lucas, S.; Lundell, T.; Martin, R.; McLaughlin, D.J.; Morgenstern, I.; Morin, E.; Murat, C.; Nagy, L.G.; Nolan, M.; Ohm, R.A.; Patyshakuliyeva, A.; Rokas, A.; Ruiz-Duenas, F.J.; Sabat, G.; Salamov, A.; Samejima, M.; Schmutz, J.; Slot, J.C.; St John, F.; Stenlid, J.; Sun, H.; Sun, S.; Syed, K.; Tsang, A.; Wiebenga, A.; Young, D.; Pisabarro, A.; Eastwood, D.C.; Martin, F.; Cullen, D.; Grigoriev, I.V.; Hibbett, D.S., The Paleozoic origin of enzymatic lignin decomposition reconstructed from 31 fungal genomes. Science, 2012, 336, (6089), 1715-1719. Ohm, R.A.; de Jong, J.F.; Lugones, L.G.; Aerts, A.; Kothe, E.; Stajich, J.E.; de Vries, R.P.; Record, E.; Levasseur, A.; Baker, S.E.; Bartholomew, K.A.; Coutinho, P.M.; Erdmann, S.; Fowler, T.J.; Gathman, A.C.; Lombard, V.; Henrissat, B.; Knabe, N.; Kues, U.; Lilly, W.W.; Lindquist, E.; Lucas, S.; Magnuson, J.K.; Piumi, F.; Raudaskoski, M.; Salamov, A.; Schmutz, J.; Schwarze, F.W.; vanKuyk, P.A.; Horton, J.S.; Grigoriev, I.V.; Wosten, H.A., Genome sequence of the model mushroom Schizophyllum commune. Nat. Biotechnol., 2010, 28, (9), 957-963. Martinez, D.; Larrondo, L.F.; Putnam, N.; Gelpke, M.D.; Huang, K.; Chapman, J.; Helfenbein, K.G.; Ramaiya, P.; Detter, J.C.; Larimer, F.; Coutinho, P.M.; Henrissat, B.; Berka, R.; Cullen, D.; Rokhsar, D., Genome sequence of the lignocellulose degrading fungus Phanerochaete chrysosporium strain RP78. Nat. Biotechnol., 2004, 22, (6), 695700. Martinez, D.; Challacombe, J.; Morgenstern, I.; Hibbett, D.; Schmoll, M.; Kubicek, C.P.; Ferreira, P.; Ruiz-Duenas, F.J.; Martinez, A.T.; Kersten, P.; Hammel, K.E.; Vanden Wymelenberg, A.; Gaskell, J.; Lindquist, E.; Sabat, G.; Bondurant, S.S.; Larrondo, L.F.; Canessa, P.; Vicuna, R.; Yadav, J.; Doddapaneni, H.; Subramanian, V.; Pisabarro, A.G.; Lavin, J.L.; Oguiza, J.A.; Master, E.; Henrissat, B.; Coutinho, P.M.; Harris, P.; Magnuson, J.K.; Baker, S.E.; Bruno, K.; Kenealy, W.; Hoegger, P.J.; Kues, U.; Ramaiya, P.; Lucas, S.; Salamov, A.; Shapiro, H.; Tu, H.; Chee, C.L.; Misra, M.; Xie, G.; Teter, S.; Yaver, D.; James, T.; Mokrejs, M.; Pospisek, M.; Grigoriev, I.V.; Brettin, T.; Rokhsar, D.; Berka, R.; Cullen, D., Genome, transcriptome, and secretome analysis of wood decay fungus Postia placenta supports unique mechanisms of lignocellulose conversion. Proc. Natl. Acad. Sci. U. S. A., 2009, 106, (6), 1954-1959. Baldrian, P.; Valaskova, V., Degradation of cellulose by basidiomycetous fungi. FEMS Microbiol. Rev., 2008, 32, (3), 501-521. Kern, M.; McGeehan, J.E.; Streeter, S.D.; Martin, R.N.; Besser, K.; Elias, L.; Eborall, W.; Malyon, G.P.; Payne, C.M.; Himmel, M.E.; Schnorr, K.; Beckham, G.T.; Cragg, S.M.; Bruce, N.C.; McQueen-Mason, S.J., Structural characterization of a unique marine animal family 7 cellobiohydrolase suggests a mechanism of cellulase salt tolerance. Proc. Natl. Acad. Sci. U. S. A., 2013, 110, (25), 1018910194. Warnecke, F.; Luginbuhl, P.; Ivanova, N.; Ghassemian, M.; Richardson, T.H.; Stege, J.T.; Cayouette, M.; McHardy, A.C.; Djordjevic, G.; Aboushadi, N.; Sorek, R.; Tringe, S.G.; Podar, M.; Martin, H.G.; Kunin, V.; Dalevi, D.; Madejska, J.; Kirton, E.; Platt, D.; Szeto, E.; Salamov, A.; Barry, K.; Mikhailova, N.; Kyrpides, N.C.; Matson, E.G.; Ottesen, E.A.; Zhang, X.; Hernandez, M.; Murillo, C.; Acosta, L.G.; Rigoutsos, I.; Tamayo, G.; Green, B.D.; Chang, C.; Rubin, E.M.; Mathur, E.J.; Robertson, D.E.; Hugenholtz, P.; Leadbetter, J.R., Metagenomic and functional analysis of hindgut microbiota of a wood-feeding higher termite. Nature, 2007, 450, (7169), 560-565.

Revalorizing Lignocellulose for the Production [33] [34]

[35] [36]

[37]

[38] [39]

[40] [41] [42] [43]

[44]

[45] [46]


Watanabe, H.; Tokuda, G., Cellulolytic systems in insects. Annu. Rev. Entomol., 2010, 55, 609-632. Ding, S.-Y.; Liu, Y.-S.; Zeng, Y.; Himmel, M.E.; Baker, J.O.; Bayer, E.A., How does plant cell wall nanoscale architecture correlate with enzymatic digestibility? Science, 2012, 338, (6110), 1055-1060. Artzi, L.; Bayer, E.A.; Morais, S., Cellulosomes: bacterial nanomachines for dismantling plant polysaccharides. Nat. Rev. Microbiol., 2017, 15, (2), 83-95. Eriksson, T.; Karlsson, J.; Tjerneld, F., A model explaining declining rate in hydrolysis of lignocellulose substrates with cellobiohydrolase I (cel7A) and endoglucanase I (cel7B) of Trichoderma reesei. Appl. Biochem. Biotechnol., 2002, 101, (1), 41-60. You, C.; Zhang, Y.-H.P., Self-assembly of synthetic metabolons through synthetic protein scaffolds: one-step purification, co-immobilization, and substrate channeling. ACS. Synth. Biol., 2012, 2, (2), 102-110. Cox, D.B.; Platt, R.J.; Zhang, F., Therapeutic genome editing: prospects and challenges. Nat. Med., 2015, 21, (2), 121-131. Davidi, L.; Morais, S.; Artzi, L.; Knop, D.; Hadar, Y.; Arfi, Y.; Bayer, E.A., Toward combined delignification and saccharification of wheat straw by a laccase-containing designer cellulosome. Proc. Natl. Acad. Sci. U. S. A., 2016, 113, (39), 10854-10859. Zhang, H.; Wang, X., Modular co-culture engineering, a new approach for metabolic engineering. Metab. Eng., 2016, 37, 114-121. Gustavsson, M.; Lee, S.Y., Prospects of microbial cell factories developed through systems metabolic engineering. Microbial Biotechnology, 2016, 9, (5), 610-617. Newman, D.J.; Cragg, G.M., Natural Products as Sources of New Drugs over the Last 25 Years. J. Nat. Prod., 2007, 70, (3), 461-477. Eichenberger, M.; Lehka, B.J.; Folly, C.; Fischer, D.; Martens, S.; Simon, E.; Naesby, M., Metabolic engineering of Saccharomyces cerevisiae for de novo production of dihydrochalcones with known antioxidant, antidiabetic, and sweet tasting properties. Metab. Eng., 2017, 39, 80-89. Koopman, F.; Beekwilder, J.; Crimi, B.; van Houwelingen, A.; Hall, R.D.; Bosch, D.; van Maris, A.J.; Pronk, J.T.; Daran, J.M., De novo production of the flavonoid naringenin in engineered Saccharomyces cerevisiae. Microb. Cell. Fact., 2012, 11, 155. Tu, Y., The discovery of artemisinin (qinghaosu) and gifts from Chinese medicine. Nat. Med., 2011, 17, (10), 12171220. Ro, D.-K.; Paradise, E.M.; Ouellet, M.; Fisher, K.J.; Newman, K.L.; Ndungu, J.M.; Ho, K.A.; Eachus, R.A.; Ham,

[47]

[48]

[49]

[50] [51]

[52] [53] [54] [55]

[56] [57]

[58]

9

T.S.; Kirby, J.; Chang, M.C.Y.; Withers, S.T.; Shiba, Y.; Sarpong, R.; Keasling, J.D., Production of the antimalarial drug precursor artemisinic acid in engineered yeast. Nature, 2006, 440, (7086), 940-943. ZHANG, C.; Chen, X.; Zou, R.; Zhou, K.; Stephanopoulos, G.; Too, H.-P., Combining Genotype Improvement and Statistical Media Optimization for Isoprenoid Production in E. coli. PLoS ONE, 2013, 8, (10), e75164. Zou, R.; Zhou, K.; Stephanopoulos, G.; Too, H.P., Combinatorial engineering of 1-deoxy-D-xylulose 5-phosphate pathway using cross-lapping in vitro assembly (CLIVA) method. PLoS ONE, 2013, 8, (11), e79557. Westfall, P.J.; Pitera, D.J.; Lenihan, J.R.; Eng, D.; Woolard, F.X.; Regentin, R.; Horning, T.; Tsuruta, H.; Melis, D.J.; Owens, A.; Fickes, S.; Diola, D.; Benjamin, K.R.; Keasling, J.D.; Leavell, M.D.; McPhee, D.J.; Renninger, N.S.; Newman, J.D.; Paddon, C.J., Production of amorphadiene in yeast, and its conversion to dihydroartemisinic acid, precursor to the antimalarial agent artemisinin. Proc. Natl. Acad. Sci. U. S. A., 2012, 109, (3), E111-118. Yuan, J.; Ching, C.B., Dynamic control of ERG9 expression for improved amorpha-4,11-diene production in Saccharomyces cerevisiae. Microb. Cell. Fact., 2015, 14, 38. Zhou, K.; Zou, R.; ZHANG, C.; Stephanopoulos, G.; Too, H.-P., Optimization of amorphadiene synthesis in bacillus subtilis via transcriptional, translational, and media modulation. Biotechnol. Bioeng., 2013, 110, (9), 2556-2561. Hao, X.; Pan, J.; Zhu, X. In Natural Products; Springer, 2013, pp 2797-2812. Kusari, S.; Singh, S.; Jayabaskaran, C., Rethinking production of Taxol® (paclitaxel) using endophyte biotechnology. Trends Biotechnol., 2014, 32, (6), 304-311. Peplow, M., Synthetic biology's first malaria drug meets market resistance. Nature, 2016, 530, 389-390. Minty, J.J.; Singer, M.E.; Scholz, S.A.; Bae, C.H.; Ahn, J.H.; Foster, C.E.; Liao, J.C.; Lin, X.N., Design and characterization of synthetic fungal-bacterial consortia for direct production of isobutanol from cellulosic biomass. Proc. Natl. Acad. Sci. U. S. A., 2013, 110, (36), 14592-14597. Kothe, E., Mating-type genes for basidiomycete strain improvement in mushroom farming. Appl. Microbiol. Biotechnol., 2001, 56, (5-6), 602-612. Song, H.; Ding, M.Z.; Jia, X.Q.; Ma, Q.; Yuan, Y.J., Synthetic microbial consortia: from systematic analysis to construction and applications. Chem. Soc. Rev., 2014, 43, (20), 6954-6981. Li, Y.; Smolke, C.D., Engineering biosynthesis of the anticancer alkaloid noscapine in yeast. Nat. Commun., 2016, 7, 12137.

DISCLAIMER: The above article has been published in Epub (ahead of print) on the basis of the materials provided by the author. The Editorial Department reserves the right to make minor modifications for further improvement of the manuscript.

PMID: 28901274