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Appl Microbiol Biotechnol (2009) 83:799–808 DOI 10.1007/s00253-009-2039-z

MINI-REVIEW

Biosynthesis and biotechnological production of flavanones: current state and perspectives Zachary L. Fowler & Mattheos A. G. Koffas

Received: 7 April 2009 / Revised: 7 May 2009 / Accepted: 7 May 2009 / Published online: 28 May 2009 # Springer-Verlag 2009

Abstract Polyphenols produced in a wide variety of flowering and fruit-bearing plants have the potential to be valuable fine chemicals for the treatment of an assortment of human maladies. One of the major constituents within this chemical class are flavonoids, among which flavanones, as the precursor to all flavonoid structures, are the most prevalent. We review the current status of flavanone production technology using microorganisms, with focus on heterologous protein expression. Such processes appear as attractive production alternatives for commercial synthesis of these high-value chemicals as traditional chemical, and plant cell cultures have significant drawbacks. Other issues of importance, including fermentation configurations and economics, are also considered. Keywords Flavanones . Fermentation . Polyphenols . Cofactor engineering

Introduction Flavonoids are a highly diverse group of plant secondary metabolites, ubiquitously found throughout the plant kingdom with many flavonoid structures displaying a variety of biochemical activities, including estrogenic, antioxidant, antiviral, antibacterial, antiobesity, and anticancer. Typically, flavonoids are broken down into six major categories: isoflavones, flavanones, flavones, flavonols, catechins, and anthocyanins. Bond saturation, hydroxylation, and ring Z. L. Fowler : M. A. G. Koffas (*) Department of Chemical and Biological Engineering, University at Buffalo, Buffalo 14260 NY, USA e-mail: [email protected]

position are used to differentiate each class. The diverse pharmicodynamic abilities of flavonoids have drawn major attention to the molecules for personal health applications (Harborne and Williams 2000; Forkmann and Martens 2001) and also from pharmaceutical companies keen on their native nutraceutical properties or to use as starting formulations for market pharmaceuticals. While currently only being used as dietary supplements, flavonoids are being intensively investigated as treatment options to many chronic human pathological conditions, including cancer and diabetes (Zava and Duwe 1997; Potter et al. 1998; Caltagirone et al. 2000; Pouget et al. 2001; Greenwald 2004; Hou et al. 2004; Allister et al. 2005; McDougall et al. 2005; Popiolkiewicz et al. 2005). For example, using diabetic mice as model systems, the glucosylated flavanone hesperidin and the flavanone naringenin were found to be highly effective at improving lipid metabolism by altering hepatic enzyme activities while at the same time lowering blood sugar levels through the down-regulation of the hepatic GLUT2 and glucose-6phosphotase and simultaneously up-regulating the hepatic glucokinase and adipocyte GLUT4 (Ae Park et al. 2006). While this study and numerous others are promising, flavonoid availability in humans is a concerning problem due to diets, in many parts of the world, that are insufficient in the amount of fruits and vegetables ingested and therefore preventing sufficient flavonoid consumption (Birt et al. 2001). Thus, as a result of low consumption rates and to a lesser degree low flavonoid concentrations in planta, the commercialization of a large-scale production platform becomes more important for using flavonoids as nutraceutical supplements. Flavanones, the direct precursors of the vast majority of flavonoids, are synthesized from the amino acid phenylalanine or tyrosine (Fig. 1). The process begins with the

800 Fig. 1 Detailed biosynthetic steps for flavanones and the diversification of flavonoids. The three successive carbons added from malonyl-CoA by CHS are shown in green, red, and blue. The R-groups denote the hydroxylation patterns for the natural flavonoid compounds although unnatural substitutions can be performed at these positions. Abbreviations are in text or as follows: DFR dihydroflavanone reductase, LAR leucoanthocynanidin reductase, ANS anthocyanidin synthase, 3GT uridine, flavanone 3-glucoside transferase, FSI flanone synthase, CHR chalcone reductase, IFS isoflavanone synthase, FHT flavanone hydroxytransferase, FLS flavonol synthase

Appl Microbiol Biotechnol (2009) 83:799–808 OH OH

PAL/TAL HOOC

NH2

HOOC

Tyrosine

C4H

p-Coumaric Acid OH

PAL OH HOOC

HOOC

NH2

HOOC

4CL

Cinnamic Acid

Phenylalanine

R1

Mal-CoA

R1

Caffeic acid

CHS

R2

R2

CoAS

CoAS O

O

Mal-CoA

CHS

CoA CO2

H2C

Acid-CoA Complex

CoA CO2

O

COSCoA

Malonyl-CoA

R1 R1

COOH

Mal-CoA OH

HO

R2

R2

CHS

CoAS O

O

OH

O

Chalcones

O

2 CoA CO2

CHR IFS

CHI

R3 O+

HO

OH OH

R2

OH

Anthocyanidins

FHT DFR

HO

R1 O

OH O

R2

Flavanones

3GT

O

IFS

R2

LAR ANS

HO

R1

O

FHT FLS

FSI

Isoflavones R3

R3 O+

HO

OH O

OH

HO

R2

Glc

Anthocyanin 3-O-glucosides

enzyme phenylalanine/tyrosine ammonia lyase (PAL/TAL), converting these amino acid building blocks into phenylpropanoic acids. The flavanone-biosynthetic pathway also includes a cytochrome-P450 enzyme, cinnamate 4hydroxylase (C4H), which adds a 4′-hydroxyl group to the phenylalanine aromatic ring. The CoA esters are subsequently synthesized from phenylpropanoic acids by the action of phenylpropanoyl-CoA ligases, such as 4coumaryl:CoA ligase (4CL). The type III polyketide synthase chalcone synthase (CHS) then catalyzes the sequential condensation of three malonyl-CoA moieties with one CoA-ester molecule to form chalcones. This is the first committed step into flavonoid biosynthesis, as alternative type III polyketide synthases exist exhibiting high homology to CHS (>70%) and using these same precursors to form stilbenes (using three malonyl-CoA units), benzy-

R1

O

HO

R2

OH O

Flavones

OH

O OH

R2

OH O

Flavonols

lacetolactone (using only one malonyl-CoA unit), and other aromatic molecules. While the flavonoid biosynthetic enzymes were originally thought to be only plant-derived, reports have appeared demonstrating the presence of type III polyketide synthases in microorganisms (Ueda et al. 1995; Moore and Hopke 2001; Kaneko et al. 2003a, b). The final flavanone structure is formed only when chalcones are stereospecifically isomerized into (2S)-flavanones by chalcone isomerase (CHI), a reaction known to occur spontaneously in alkali environments. After generation of the flavanone, a myriad of enzymes can act to functionalize and/or alter conformations of the three-ring phenylpropanoid core resulting in the generation of over 8,000 different chemical structures. Functionalization can manifest as hydroxylations, reductions, alkylations, oxidations, and glucosylations alone or in combinations.

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This functionalization leads to many of the biologically active properties seen for flavonoids (Table 1). In this review, we will focus our attention on efforts to achieve efficient flavanone biosynthesis in microorganisms, the first and core flavonoid structure. As such, we will explore the current state of the biotechnological production of flavanones by examining both the current methodologies for attaining flavanones as well as the more recent bioengineering approaches. As such, we will place an emphasis on not only the introduction of the plant biosynthetic pathway into the host cell but also on any redesigning of its native metabolism.

Production in practice With interest rising due to their wide application potential toward a number of consumer products, developments in chemical and biotechnical methods for flavonoid synthesis have occurred. While many groups have demonstrated the feasibility of efficient production of a number of these plant-derived natural products through chemical synthesis (Furlong and Nudelman 1985; Tanaka et al. 2000; Kumazawa et al. 2001; Lim et al. 2001; Wan and Chan 2004; Tanaka et al. 2007), the use of harsh or toxic chemical solvents has limited this methodology to specialized small quantity production. More importantly, chemical synthesis of flavanones, like many other aromatic products, yields a mixture of two stereoisomers. This is a critical flaw since only the (2S)-flavanones have been shown to be biologically active (Andersen and Markham 2006). Since

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the use of flavonoids in general has been as supplements, plant extraction has been the dominant form of isolation for these diverse nutraceuticals providing a process that requires only minimal processing efforts. However, plant extraction also yields a mixture of various substituted flavonoids; thus generating an extract containing only one class, let alone a single substitution pattern, is challenging task that would require countless purification techniques adding to economical cost and environmental impacts. Alternatively, bioreactor-based systems for mass production of flavonoids from plant cell cultures have been described for a few species (Zhong et al. 1991; Kobayashi et al. 1993), but to date, economic feasibility has not been established, partly because of engineering challenges in large-scale cultivation. One major challenge is that plant cells tend to form aggregates that influence culture productivity (Hanagata et al. 1993) since cells within aggregates are not adequately exposed to the required lighting needed to induce flavonoid biosynthesis by plant tissue. For example, expression of PAL, a key enzyme in the biosynthetic pathway, is promoted in plants primarily by UV wavelengths, particularly those of the UV-B region (Wellmann 1975), while other enzymes, particularly those of the anthocyanin biosynthetic branch, appear to be regulated in part by UV and in part by the phytochromeactivating wavelengths of 700–800 nm (Meyer et al. 2002). In that respect, irradiance becomes a limiting factor to productivity not only when excluded from cells at the interior of an aggregate (Hall and Yeoman 1986) but also when in a dense cell culture, at reduced cell dosing, or

Table 1 Current and potential application of flavonoids or flavonoid based drugs Flavonoid

Current/potential applications (disease)

Reference

Flavanones and chalcones

LDL oxidation (arthrosclerosis) Cyclooxygenase inhibitory ability (cancer) Malaria chemotherapy (malaria) Inflammation response trigger (reduce inflammation) GLUT inhibitor (diabetes) Cyclooxygenase inhibitory ability (cancer) GLUT inhibitor (diabetes) Pancreatic lipase inhibitors (diabetes) Inhibitors of cell cycle control kinases (cancer) Alpha-glucosidase inhibitor (diabetes) GLUT inhibitor (diabetes) Improves cholesterol regulation (diabetes) Promote cardiovascular health (heart disease) Pancreatic lipase and glucosidase inhibitor (diabetes) Regulate adipocyte function (obesity) Improves glucose and lipid metabolism (diabetes) Modulate blood hormone levels (multiple sclerosis)

(Miranda et al. 2000; Kumar et al. 2003; Kinghorn et al. 2004; Kontogiorgis et al. 2008; Mojzis et al. 2008)

Flavone Flavonols

Isoflavonoids

Catechins and Anthocyanins

(Kinghorn et al. 2004; Kwon et al. 2007; Chemler et al. 2009) (Park and Levine 2000; Nakai et al. 2005; Hsu and Yen 2006; Kwon et al. 2007; Chemler et al. 2009) (Kim et al. 2000; Song et al. 2002; Lee 2006; Kwon et al. 2007; Chemler et al. 2009)

(Shi et al. 1998; Kim et al. 2000; Tokimitsu 2004; Wolfram et al. 2006; Sternberg et al. 2008; Tsuda 2008)

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when the vessel wall composition selectively restricts certain wavelengths (Smith and Spomer 1995). Expression of flavonoid biosynthetic genes in transgenic plants has also been investigated for production of these ubiquitous secondary metabolites. In a recent study, CHI from Saussurea medusa was transformed into Nicotiana tabacum, a non-leguminous species, and resulted in an up to 5-fold total flavonoid production increase when compared to wild-type plants (Li et al. 2006). A similar method was later used in separate studies to introduce isoflavone synthase (IFS), a cytochrome-P450 monooxgenase using flavanone precursors, into tobacco plants (Yu et al. 2000) and soybeans (Yu et al. 2003). Even though the transgenic plants successfully synthesized isoflavonoids, the amounts were low due to competing pathways leading to the biosynthesis of other flavonoid classes. This diversion of metabolic flux and issues inherent with plant cell suspension cultures (Hellwig et al. 2004) are some of the reasons why until now no such system has been established for the commercial, large-scale production of flavonoids despite a more than 10-year effort in this field. The emerging bioreactor-based systems for flavonoid production have and will continue to be focused on the heterologous biosynthesis of these natural products using well-characterized recombinant hosts, including the yeast Saccharomyces cerevisiae and the gram-negative bacterium Escherichia coli. Although other microorganisms may in time be identified as more suitable hosts, the tractability of these two hosts inclines in their favor. In actuality, much of the recent characterization and understanding of mechanisms and control within the flavonoid biosynthetic pathway has been a result of studies using recombinant protein expression. More importantly, the use of microbial factories poses several significant advantages over chemical synthesis or plant cell cultures, including rapid growth rates, ease of cultivation, and the availability of convenient genetic manipulation techniques. All these are factors that enable high-production levels to be maintained from continuous fermentative cultures. Moreover, this enzymebase production strategy enables increased product selectivity and reduced use in both toxic chemicals and energy.

Flavanone biosynthesis in microorganisms Engineering of the biosynthesis of flavanones has rapidly evolved in the last 6 years from the simple expression of the plant enzymes in a host to entail more advanced engineering strategies that improve factors such as precursor levels and expression stability. While yeast may be a more suitable expression platform, as we will discuss, a vast majority of the major biotechnological developments have been introduced into E. coli expression systems for

Appl Microbiol Biotechnol (2009) 83:799–808

different metabolic configurations (Fig. 2) and yielding a variety of production levels (Table 2). Nevertheless, it is important to consider that both systems have been developed cocurrently and have highlighted some important characteristics of phenylpropanoid biosynthesis. As more information about the specific flavonoid enzymes becomes available, achieving highly efficient production strategies will require the selection of optimal combinations in which to cluster the expressed proteins. More specifically, it has been noted already that there exists a great variability and flexibility in the types of activities phenylpropanoid biosynthetic genes have toward accepting different substrates (Jiang et al. 2005; Katsuyama et al. 2007). In fact, a recent study used the broad substrate specificity of Petroselinum crispum 4CL, Petunia X hybrida CHS, and Medicago sativa CHI to create a library of unnatural flavanones (Chemler et al. 2007). E. coli With a plethora of molecular biological techniques at hand and a history of a robust ability for recombinant expression, E. coli is traditionally the first choice as a production platform in the biotech industry. As such, it is not surprising that the first attempt to afford plant-specific flavanone biosynthesis was in E. coli. It was accomplished through the simultaneous expression of plant biosynthetic enzymes, namely PAL from Rhodotorula rubra, 4CL from Streptomyces coelicolor A3(2), and CHS from Glycyrrizha echinata using three different vector constructs with each construct varying the number and location of promoters and ribosome binding sites (RBS) for each gene. While an optimal pET expression vector construct consisting of both strong T7 promoters (PT7) and RBS sites before each gene produced the highest levels of production (Hwang et al. 2003; Kaneko et al. 2003a, b), the titers were still considerably low and thus would prohibit large-scale synthesis. Later, a similar strategy was employed to recombinantly express PAL, C4H, 4CL, and CHS from the model plant Arabidopsis thaliana (Watts et al. 2004). However, the C4H was inactive, therefore limiting production of naringenin to cultures supplemented with exogenous p-coumaric acid. In the end, a newly cloned and partially characterized TAL from Rhodobacter sphaeroides was used to generate naringenin from tyrosine-supplemented cultures resulting in production levels that reached 20.8 mg/L (Watts et al. 2004). Interestingly, the authors in both studies relied on alkali conditions for the conversion of the chalcone to the flavanone; thus, the inclusion of a chalcone isomerase could have significantly improved production titers. In fact, more recent studies have not only included the presence of a chalcone isomerase but also undergone a

Appl Microbiol Biotechnol (2009) 83:799–808 Fig. 2 Biosynthetic pathways used for engineering flavanone biosynthesis. Glucose, or another simple sugar, is metabolized into the major precursors for flavanone production. The multifunctional acetyl-CoA carboxylase requiring the action of biotin ligase is also shown. Engineered pathways discussed in the text are shown in orange while flavanone synthesis is in purple

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Extracellular

Cellular

CoaA ADP + Pi

Glucose (x)

PAL / TAL

Aromatic Amino Acids

Substrate (x)

Glucose

ATP + HCO3-

Esters

BirA

AccA AccD

Table 2 Engineered biosynthetic clusters for flavanone production in recombinant E. coli and yeast

Fatty Acids MatB

Expressed in E. coli Rhodotorula rubra PAL Streptomyces coelicolor A3(2) 4CL Glycyrrhiza echinata CHS Arabidopsis thaliana TAL, 4CL, CHS Rhodotorula rubra PAL Streptomyces coelicolor A3(2) 4CL Glycyrrhiza echinata CHS

a

Catalytic enzymes abbreviations: PAL, phenylalanine ammonia lyase; TAL, tyrosine ammonia lyase; 4CL, coumarate:coenzyme A ligase; CHS, chalcone synthase; CHI, chalcone isomerase; ACC; acetyl-coenzyme A carboxylase; BPL, biotin ligase; ACS, acetate synthase

b

Production conditions for pinocembrin (P), eriodictyol (E), and naringenin (N) vary between studies

Flavanones

Productionb (mg/L) P

+ Photorhabdus luminescens BPL + Escherichia coli ACS Petroselinum crispum 4CL Petunia X hybrida CHS Medicago sativa CHI Rhizobium trifolii MatB/MatC Expressed in S. cerevisiae Rhodosporidium toriloides PAL Arabidopsis thaliana 4CL Hypericum androsaemum CHS Petroselinum crispum 4CL Petunia X hybrida CHS, CHI Arabidopsis thaliana C4H

CHI

Malonate

Recombinant gene clustera

Pueraria lobata CHI Corynebacterium glutamicum ACC Petroselinum crispum 4CL Petunia X hybrida CHS Medicago sativa CHI Photorhabdus luminescens ACC

CHS

Chalcones

Acetate MatC

Biotin ATP

Malonyl -CoA

acs

Malonate (x)

4CL

PPi apoBCCP AMP apoBirA Propanoyl-CoA

holoBCCP

Acetyl -CoA

C4H

Coenzyme A

AccC holoBCCP*

Propanoic Acids

E

Reference N

(Hwang et al. 2003) 0.75 –

58

– –

0.45 20.8



60

(Watts et al. 2004) (Miyahisa et al. 2005)

(Leonard et al. 2007)

367 429

50 52

69 119 (Leonard et al. 2008)

480

50

155 (Jiang et al. 2005)

0.8



7 (Yan et al. 2005)

16.3

6.5

28.3

804

more extensive metabolic engineering effort to attain production-scale quantities of flavanones in E. coli. Specifically, various strategies were explored that amplify malonyl-CoA, the rate-limiting metabolite in E. coli, which serves as a flavonoid building block. The first approach was the most rational in design. Since malonyl-CoA is synthesized directly from acetyl-CoA by the enzyme acetyl-CoA carboxylase (ACC), its overexpression in E. coli was proposed to improve flavanone synthesis levels (Miyahisa et al. 2005). However, overexpression of E. coli’s native ACC has been reported to be detrimental to cell viability (Davis et al. 2000). To bypass this phenomenon, a plasmid harboring an artificial biosynthetic gene cluster that included PAL from a yeast, a cinnamate/coumarate:CoA ligase from S. coelicolor A3(2), licorice CHS, and Pueraria CHI along with a separate vector containing the two subunit genes of ACC from Corynebacterium glutamicum were expressed in the recombinant E. coli cells. The coexpression of these two vectors improved production titers to 60 mg/L of (2S)-naringenin (Miyahisa et al. 2005). In a related study, the four subunits, accABCD, of Photorhabdus luminescens ACC, were cloned with accA and accD individually controlled under separate PT7, while the transcription of the accBC operon was regulated under another PT7. Combining this strategy with the expression of P. crispum 4CL, P. hybrida CHS, and M. sativa CHI permitted flavanone synthesis to reach 196 mg/L of pinocembrin and 67 mg/L of naringenin. Since the functionality of the carboxylase domain of ACC requires biotinylation by the action of biotin ligase (BPL; encoded by birA), overexpression in concert with ACC overexpression was also investigated. Interestingly, BPL is known to recognize ACCs across species; therefore, the recombinant system was tested with the endogeneous E. coli BPL, the BPL from P. luminescens, and a chimeric BPL containing the N-terminus from E. coli and the C-terminus of P. luminescens. It was found that coexpression of ACC and BPL both from P. luminescens resulted in improved flavanone yields of up to 367 mg/L of pinocembrin (Leonard et al. 2007), although naringenin production only increased marginally to 69 mg/L. These results suggested that the protein interaction between ACC and BPL was important for optimum malonyl-CoA synthesis. The improved flavanone synthesis by coexpression of ACC and BPL also depended on biotin supplementation to the fermentation broth, a technique not viable for largescale synthesis especially considering the cost of biotin as an additive. As such, further metabolic engineering of the acetate pool was also explored to allow the conversion of inexpensive metabolites into high-value flavanones. In this case, the amplification of two acetate assimilation pathways in E. coli was explored, specifically the acetate kinase (ACK) and phosphotransacetylase (PTA) pathway and the

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acetyl-CoA synthetase (ACS) pathway (Leonard et al. 2007). While a strain harboring overexpressions for ACK and PTA, together with ACC only, resulted in only a moderate increase in flavanone synthesis, the coexpression of ACC with ACS resulted in flavanone production up to 383 mg/L of pinocembrin along with a decrease in acetate accumulation. Exogenous supplementation of acetate at a low level further improved flavanone synthesis to 429 mg/L of pinocembrin (Leonard et al. 2007). It is quite clear that the pool of intracellular malonyl-CoA is undersized for large-scale biosynthesis of flavanones from E. coli. Therefore, in order to increase malonyl-CoA levels, the culture supplementation using cheep exogenous sources of carbon was explored. For example, the identified malonate transport and assimilation pathway that forms malonyl-CoA by genes matB and matC from Rhizobium trifolii (An and Kim 1998) was proposed as a way to directly increase malonyl-CoA. Specifically, this malonate assimilation pathway was simultaneously overexpressed with the flavanone-biosynthetic genes 4CL, CHS, and CHI while supplementing with exogenous malonate. This resulted in production titers of 480 mg/L of pinocembrin and 155 mg/L of naringenin (Leonard et al. 2008). In an effort to retard the native degradation pathways of malonyl-CoA, the specific fatty acid inhibitor cerulenin was added to fermentations of E. coli strains harboring the flavanone-biosynthetic pathway. Doing so induced a dosedependent response of flavanone production with the maximum pinocembrin production of 710 mg/L accomplished when FabB/F was repressed using 0.2 mM of cerulenin (Leonard et al. 2008). Despite the near gram per liter productivity, the excessive cost of cerulenin makes this strategy ill-suited for an industrial process; therefore, an alternative strategy that focuses on re-routing native metabolic carbon flows to improve precursor and cofactor accessibility maybe more suitable for increasing production levels. Such an approach might utilize stoichiometric modeling (more commonly flux balance analysis) to identify unique and non-intuitive gene deletions within the host genotype, thereby significantly expanding the carbon flows from glucose into the metabolite pools needed for flavonoid biosynthesis. While flux balance analysis has been used to investigate the theoretical constraints of numerous metabolic systems, few examples of its practical application exist in the literature (Alper et al. 2005a, b; Fong et al. 2005). Recently, however, this approach to simultaneously balance carbon flows into the cofactor pools was employed to great success for flavanone biosynthesis (Fowler et al. 2009). It was shown that a stochastic model of E. coli metabolism, which also included the pathways for flavanone biosynthesis, could effectively identify gene deletions and overexpression targets required to increase availability

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of malonyl-CoA and other biosynthetic cofactors leading to improved production of flavanones (Fowler et al. 2009). To understand the connectivity of the metabolic network, an optimization routine termed CiED was utilized to evolve in silico strains and identify gene deletion targets leading to improved flavanone production. The authors demonstrated 3-fold increases in cellular malonyl-CoA levels in the mutant genotypes constructed from CiED predictions after which the maximum production of flavanones increased from 69 mg/L to in excess of 210 mg/L. By including overexpressions also predicted from the model, and the previously mentioned ACS, ACC, and BPL overexpressions, production levels rose to 270 mg/L of the flavanone naringenin (Fowler et al. 2009).

S. cerevisiae S. cerevisiae offers a number of distinct advantages over E. coli. Perhaps the one most important to commercial implementation is that the United States Food and Drug Administration classifies yeast as a GRAS organism, thus recognizing it as a safe production method for synthesis of consumed goods. On the other hand, as a eukaryote, yeast, has the ability to support functional expression of membrane-bound cytochrome P450 enzymes. P450 enzymes require the attachment to the eukaryotic cell’s endoplasmic reticulum (ER) membrane and a redox partner typically in the form of a P450 reductase that transports electrons from the NADPH donor to the heme-core of the P450 complex. More importantly, there are a number of flavonoid biosynthetic enzymes, including the C4H, which are of the P450 enzyme class. This feature, together with the expectation of better expression of the plant-derived enzymes due to the capability of posttranslational modifications and its phylogenetical similarity to plants, makes yeast an attractive host platform for flavonoid biosynthesis. The first time enzymes of the phenylpropanoid pathway, namely PAL and C4H, were positive expressed in S. cerevisiae was done by Ro and Douglas together with a cytochrome P450 reductase to aid the activity of the C4H monoxygenase (Ro and Douglas 2004). In this work, not only was the ability of yeast to express the flavonoid biosynthetic pathway demonstrated but the formation of metabolons or complexes between the two phenylpropanoid enzymes was also investigated. In their work, using radio-labeled substrates, the others showed that efficient carbon flux from phenylalanine to coumarate is not facilitated by the formation of metabolons as suspected, but rather by a biochemical coupling of the two enzymes (Ro and Douglas 2004). While production levels were fairly low—approximately 1.2 μmol of phenylpropanoids—the functional expression of the cytochrome P450s was permitted.

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A later study looked to enhance production levels using two engineered yeast strains along with a stronger GAL10 promoter system (Jiang et al. 2005). While only expressing a PAL from Rhodosporidium toriloides, 4CL from A. thaliana, and CHS from Hypericum androsaemum, the production levels reached nearly 7 mg/L of naringenin and 0.8 mg/L of pinocembrin. Interestingly, a deletion of PAD1 in one of the yeast strains caused a surprisingly different PAL (TAL) behavior that produced lower levels of flavanones, though with a different distribution. While the low 4CL activity toward trans-cinnamic acid may be attributed to the A. thaliana 4CLs Km value (6.3 mM), it was notably lower in the PAD1 mutant than in the parental strain, possibly due to the Pad1P decarboxylase (or another endogenous enzyme) out-competing 4CL for the transcinnamic acid (Jiang et al. 2005). In related study, the synthesis of the flavonoid naringenin from the phenylpropanoic acids was afforded in a recombinant S. cerevisiae strain expressing a P. crispum 4CL, P. hybrida CHS and CHI, and the P450 C4H from A. thaliana (Yan et al. 2005). Although this work used a similar GAL1 promoter system within a commercial yeast strain having a similar genetic background, production levels were increased 62% for naringenin and 22% for pinocembrin over the previous study. An extension of this work, also performed in yeast, involved the biosynthesis of the flavones that are derivatives of the flavanones (Leonard et al. 2005), which are known to have potent medicinal properties (Caltagirone et al. 2000).

Future challenges in flavonoid biosynthesis To achieve efficient production platforms of flavanones or flavonoids in general, improving the availability of starting materials used during the biosynthetic process is of chief concern. While many of the early studies resorted to the more practical approach of substrate feeding to improve the intracellular concentrations of the aromatic amino acids in an effort to increase production levels, the recent studies have bypassed the conversion of aromatics to the phenylpropanoic acids. This is because phenylpropanoid acids are readily available from abundant, renewable sources such as, lignin, one of the most abundant organic polymers in nature (Boerjan et al. 2003). The three typical monolignols used within the backbone of lignin are p-coumaryl, coniferyl, and sinapyl alcohols (Martone et al. 2009). A number of recent studies have investigated a variety of fungal enzymes linked with the degradation of plant matter to identify esterases and laccases that can be used to carry out the bioconversion of lignin (Bergbauer and Eggert 1994; Leonowicz et al. 2001; Benoit et al. 2006; Latha et al. 2007; Guglielmetti et al. 2008) to fermentable substrates

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and starting materials for flavonoid biosynthesis. As more degradation pathways are elucidated, other cellulostic materials may also have the potential to act as substrates for production of flavonoids. One of the biggest challenges however for engineering the flavonoid (and many other phytochemical’s) biosynthetic pathway in E. coli is the functional expression of ER-associated plant cytochromic P450 monoxygenases. Due to requirements of P450 systems, functional expression of these enzymes in E. coli is a daunting task since E. coli lacks ER. Efforts to engineer a soluble P450 monooxygenase have generally resulted in enzymes with very low solubility and formation of inclusion bodies resulting in extensive cell lysis, especially in cases with high expression levels. Such was the case for the generation of an active flavonoid 3′5′-hydroxylase (F3′5′H) derived from Catharanthus roseus for the biosynthesis of hydroxylated flavonols such as quercetin and myricetin (Leonard et al. 2006). Both the F3′5′H and a P450-reductase from C. roseus (the enzyme responsible for transferring electrons from NADPH to the P450) were modified by either removing terminal codons and/or changing existing codons in the N- and C-terminus regions then fused together using a linker sequence with no preference for secondary structure formation. When the engineered chimera was expressed in E. coli together with a grafted flavanone-biosynthetic pathway, small amounts of quercetin could be recovered from the culture media (Leonard et al. 2006). Recently, a series of artificial isoflavone synthases were designed that allowed robust production by E. coli (Leonard and Koffas 2007). In this work, by assembling the overall architecture to mimic that of a self-sufficient bacterial P450s and at the same time including tailor-made membrane recognition signals, one identified chimera had up to 20-fold higher in vivo production than that achieved by the native enzyme expressed in a eukaryotic host. Nevertheless, cell lysis was evident, indicating that there are still several other parameters, including the use of weaker promoters and lower copy number plasmids, that need to be tuned for biocatalysis optimization. Other challenges have still yet to be fully investigated, including the selection of the right cluster of flavonoid genes to allow a high kinetic turnover (and possibly a broad substrate specificity); codon optimization of recombinant plant proteins; and identification of suitable promoters for achieving optimal, as opposed to maximal, expression levels of the recombinant proteins. Additionally, methods that modify the flavanone-biosynthetic genes in order to improve the catalytic activity or relieve product inhibition, are viable directions for achieving production-scalable titers. These issues provide golden opportunities for the development of new methodologies and technologies in

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metabolic engineering that can be utilized in several other metabolic engineering projects related to natural product biosynthesis.

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