Appl Microbiol Biotechnol (2014) 98:5363–5374 DOI 10.1007/s00253-014-5726-3

MINI-REVIEW

Bacterial synthesis of D-amino acids Atanas D. Radkov & Luke A. Moe

Received: 10 February 2014 / Revised: 22 March 2014 / Accepted: 25 March 2014 / Published online: 22 April 2014 # Springer-Verlag Berlin Heidelberg 2014

Abstract Recent work has shed light on the abundance and diversity of D -amino acids in bacterial extracellular/ periplasmic molecules, bacterial cell culture, and bacteriarich environments. Within the extracellular/periplasmic space, D-amino acids are necessary components of peptidoglycan, and disruption of their synthesis leads to cell death. As such, enzymes responsible for D-amino acid synthesis are promising targets for antibacterial compounds. Further, bacteria are shown to incorporate a diverse collection of D-amino acids into their peptidoglycan, and differences in D-amino acid incorporation may occur in response to differences in growth conditions. Certain D-amino acids can accumulate to millimolar levels in cell culture, and their synthesis is proposed to foretell movement from exponential growth phase into stationary phase. While enzymes responsible for synthesis of Damino acids necessary for peptidoglycan (D-alanine and Dglutamate) have been characterized from a number of different bacteria, the D-amino acid synthesis enzymes characterized to date cannot account for the diversity of D-amino acids identified in bacteria or bacteria-rich environments. Free D-amino acids are synthesized by racemization or epimerization at the α-carbon of the corresponding L-amino acid by amino acid racemase or amino acid epimerase enzymes. Additionally, Damino acids can be synthesized by stereospecific amination of α-ketoacids. Below, we review the roles of D-amino acids in bacterial physiology and biotechnology, and we describe the known mechanisms by which they are synthesized by bacteria. Electronic supplementary material The online version of this article (doi:10.1007/s00253-014-5726-3) contains supplementary material, which is available to authorized users. A. D. Radkov : L. A. Moe (*) Department of Plant and Soil Sciences, College of Agriculture, Food, and Environment, University of Kentucky, 311 Plant Science Building, Lexington, KY 40546-0312, USA e-mail: [email protected]

Keywords D-amino acids . Peptidoglycan . Amino acid synthesis . Amino acid racemase . Amino acid epimerase . D-amino acid aminotransferase

Introduction With the exception of glycine, each of the 20 canonical proteinogenic amino acids is chiral about its α-carbon and therefore can exist in one of two stereoisomeric forms: the Lform and the D-form. Nature has effectively selected for Lamino acids (L-AAs) to serve as the building blocks of ribosomally produced polypeptides and as key metabolic intermediaries in biological systems. As such, L-AAs are typically found in much higher abundance than their mirror image counterparts. Nonetheless, recent work has begun to highlight both the abundance and potential roles of D-amino acids (DAAs) in nature. For example, D-AAs are abundant in soil, fermented foods, and beverages (Brückner and Hausch 1989; Kato et al. 2011; Vranova et al. 2011), and extracts of mammalian gut, rumen, and feces (Brückner and Schieber 2001; Schieber et al. 1999). The presence of D-AAs in environmental samples is typically attributed to the associated microbiota. This is because microbes are now known to be a rich source of D-AAs and because those environments that harbor relatively large quantities of D-AAs are typically microbe rich. The major sources of D-AAs in bacteria are generally extracellular or periplasmic polymeric biomolecules, including peptidoglycan (PG), teichoic acids (TAs), and poly-γglutamate (PGA). Among these polymers, PG is the major component of the bacterial cell wall and the most commonly cited source of D-AAs in bacteria. PG is comprised of long glycan chains (alternating N-acetylglucosamine and Nacetylmuramic acid moieties) linked by short peptide stems. PG is a plastic structure that provides a protective barrier for the cell, enabling cells to survive under variable

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physicochemical conditions (Barreteau et al. 2008; Vollmer et al. 2008). Stems from adjacent glycan strands can be linked either directly or by an interpeptide bridge, itself comprised of AAs. D-AAs can be incorporated into both the peptide stem and the interpeptide bridge (Fig. 1). The PG peptide stem most often contains a D-glu residue at the second position as well as D-ala at the fourth and fifth positions (Barreteau et al. 2008). Other D-AAs that are known to be incorporated into the peptide stem include D-ser (Arias et al. 1999), D-lys (Boniface et al. 2006), and the non-protein amino acid D-isoglutamine (which results from amidation of D-glu at the α-carbon carboxyl group) (Vollmer et al. 2008). In vitro incorporation of eight different D-AAs (gln, pro, phe, tyr, val, ile, leu, and met) at the fourth position of the peptide stem has also been observed (Cava et al. 2011). Nonetheless, the two D-ala residues at the fouth and fifth positions are nearly universal in the precursor peptide stem (the terminal D-ala is removed in mature PG). The primary mode of variation between PG structures is in the length and composition of the interpeptide bridge. D-AAs incorporated into this peptide can include D-ala, D-glu, D-ser, D-asp, D-asn, D-lys, and D-ornithine (Vollmer et al. 2008). A number of recent papers have highlighted the emerging roles of free D-AAs in alteration to PG structure and regulation of its synthesis (Cava et al. 2011; Hochbaum et al. 2011; Lam et al. 2009; Lupoli et al. 2011). Millimolar concentrations of free non-canonical D-AAs were detected in cell culture supernatant from a variety of bacteria (non-canonical indicating that they were not previously detected in PG (Lam et al. 2009)). The D-AAs were shown to impact PG functionality through their incorporation into the PG structure. Somewhat surprisingly, exogenous D-AAs provided in trans are incorporated into PG in a wide variety of bacteria—even those that do not synthesize the D-AA of note—leading to speculation about DAAs as paracrine regulators in microbial communities (Cava et al. 2011). A closer look at synthesis of non-canonical DAAs in Vibrio cholerae revealed high levels of D-met in the culture supernatant in stationary phase but not during exponential phase growth. The D-met was incorporated into the PG peptide stem, which had an impact on the functional properties of the PG (Cava et al. 2011). Periplasmic or extracellular transpeptidase enzymes are responsible for incorporation of DAAs into mature PG, and studies on the substrate specificity of

these enzymes suggest that they can incorporate a wide variety of D-AAs into PG (Cava et al. 2011; Lupoli et al. 2011). Because of the timing of D -met incorporation into V. cholerae PG, it was hypothesized that production of certain D-AAs foretells changes in PG structure as the cell moves into stationary phase (Lam et al. 2009). Subsequent work indicates that D-AA-induced changes in PG structure in V. cholerae are dependent on the stress-associated sigma factor RpoS, suggesting that these changes may be in response to multiple different environmental cues rather than just movement into stationary phase (Cava et al. 2011). In addition to their incorporation into PG, free D-AAs may directly modulate the activity of periplasmic transpeptidase enzymes (Lam et al. 2009), and certain D-AAs may serve as signaling compounds. D-AAs are proposed to disassemble biofilms in some cases (Hochbaum et al. 2011; KolodkinGal et al. 2010), but not in others (Leiman et al. 2013; Sanchez et al. 2013), and additional studies show that certain D-AAs inhibit spore germination (Hills 1949; Hu et al. 2007). As much of this work is relatively new, further experiments will likely uncover additional roles that D-AAs play in regulating bacterial physiology. Indeed, D-AA production appears to be dependent on the stress response-related sigma factor RpoS in V. cholerae (Cava et al. 2011), which links D-AA production with stress response-related genes as well as other notable physiological phenomena such as virulence (Fang et al. 1992). PGA is an extracellular polymeric material typically found among members of the Bacillales order that consists of γlinked glu residues (Candela and Fouet 2006). Depending on the species, the glu residues can be in the L-configuration, the D-configuration, or in both configurations (Candela and Fouet 2006). Among bacilli, PGA is comprised of D-glu residues in the noted pathogen Bacillus anthracis. PGA imparts the cell with a viscous layer that enables B. anthracis, and other PGAproducing pathogens, to evade the host immune system. Somewhat ironically, PGA also provides the slimy texture to the Japanese soybean delicacy natto, which is fermented with Bacillus subtilis, and is suggested to have numerous health benefits (Murooka and Yamshita 2008). PGA can be released from the exterior of the cell and can also serve as a carbon and nitrogen source to the producing organism under nutrient stress conditions (Schreier 1993).

Fig. 1 D-AAs in peptidoglycan (GlcNAc N-acetylglucosamine; MurNAc N-acetylmuramic acid, A2pm 2,6-diaminopimelic acid). a Direct cross-linkage of PG peptide stems. b Linkage of PG stems through an interpeptide bridge

a

b

Appl Microbiol Biotechnol (2014) 98:5363–5374 D-Ala is abundant in TAs, which are extracellular anionic molecules comprising polymeric glycerol phosphate moieties in Gram-positive bacteria. A D-ala moiety can be linked through an ester bond at the AA carboxylate position to the glycerol 2-hydroxyl group of the repeating glycerol phosphate monomer, yet the degree of D-alanyl substitution is highly variable among TAs according to organism and growth conditions (Neuhaus and Baddiley 2003). The D-alanylation of TAs is known to regulate autolytic activity, bind and assist in cation assimilation, and play a role in cellular defense (Reichmann et al. 2013; Volkman et al. 2001). While Dalanylation of TAs is not absolutely required, cells engineered to lack D-ala in their TAs are more susceptible to certain antimicrobials as well as human immune defenses (Collins et al. 2002; Kristian et al. 2005). D-AAs are commonly found as components of peptide antibiotics synthesized by bacterial non-ribosomal peptide synthetases (NRPSs) (Caboche et al. 2008). AAs are the most common monomer units in non-ribosomal peptides (NRPs) (representing 40 % of monomeric units), but these peptides also include lipids, polyketides, and carbohydrates (Caboche et al. 2010). A search of the NORINE non-ribosomal peptide database using individual D-AAs as the monomer query reveals that certain D-AAs are highly prevalent among the 1,164 curated peptides (accession date, 01/25/14) (Caboche et al. 2008). Table S1 shows that D-ala, D-ser, D-leu, and D-glu are the most commonly found D-AAs (each is present in more than 100 peptides), while the less abundant D-AAs include Dthr (found once), D-met, and D-his (not found at all). Considering the ratio of D-AA to L-AA incorporation for each amino acid, the D-AAs with the highest D/L ratio include D-glu, which represents 34 % of the total glu monomer units, and D-ornithine, which represents 43 % of the ornithine units. The database includes NRPs produced by prokaryotic and eukaryotic organisms. Incidentally, D-AA incorporation appears to be largely the domain of prokaryotic organisms, while L-AA incorporation is more evenly distributed among prokaryotes and eukaryotes (see, for example, Fig. 4 from Caboche et al. 2010).

Mechanisms for synthesis of D-AAs Bacteria synthesize D-AAs using one of two mechanisms: inversion of stereochemistry about the α-carbon of the corresponding L-AA, or stereospecific amination of the corresponding α-ketoacid. The former, catalyzed by AA racemase or epimerase enzymes, is a reversible reaction resulting in direct interconversion of the L - and D stereoisomers. The latter reaction, also reversible, relies on an amino-containing co-substrate and is catalyzed by D-AA aminotransferase enzymes.

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Both racemases and epimerases catalyze inversion of stereochemistry at the α-carbon of AAs, but this reaction is commonly (yet incorrectly) referred to simply as AA racemization. Epimerase enzymes, such as ile 2-epimerase (Mutaguchi et al. 2013), interconvert stereochemistry among AAs with more than one chiral center. Nonetheless, the same principles apply for synthesis of D-AAs, regardless of the number of chiral centers. This reaction relies on removal of the α-proton to form a planar, carbanion intermediate that is protonated on the opposite face to yield the opposite stereoisomer. By definition, an AA α-proton is activated by virtue of the adjacent carbonyl group that lowers the pKa of the corresponding proton. Nonetheless, the proton remains relatively difficult to remove by conventional base-catalyzed deprotonation [e.g., glu pKa is ∼21 (Tanner 2002)]. AA racemases/ epimerases can be separated into two classes; this is based on their use of a cofactor-dependent reaction or a cofactorindependent reaction. Cofactor-dependent AA racemase/epimerase enzymes use a pyridoxal-5′-phosphate (PLP) cofactor. The PLP cofactor enables the enzyme to lower the pKa of the α-proton by stabilizing the attendant carbanion using a substrate-bound external aldimine. In PLP-dependent AA racemases, the incoming AA substrate displaces the internal aldimine, forming an external aldimine. The α-proton is then removed by an active site base, resulting in a resonance-stabilized carbanion intermediate that is subsequently protonated on the opposite face (Fig. 2). The product is then displaced through reformation of the internal lys aldimine. The enzymes are proposed to use two active site bases for deprotonation/ reprotonation of the two faces of the planar intermediate, with one base being responsible for deprotonation and reprotonation of the α-proton of the L-AA and one base responsible for the α-proton of the D-AA (Toney 2011). The cofactor-independent racemase/epimerase enzymes also use a two-base mechanism, albeit without an external mechanism for stabilizing the carbanion intermediate (Glavas and Tanner 1999). Of those enzymes in which the identity of the catalytic bases has been studied, both bases are demonstrated to be cys residues (Buschiazzo et al. 2006; Liu et al. 2002; Tanner 2002). A mechanism whereby a cys thiolate deprotonates the Cα, followed by reprotonation from the alternate cys, is proposed for these enzymes (Tanner 2002). D-AAs that are incorporated into NRPs by NRPSs are typically epimerized during the peptide synthesis process (Stein et al. 2005). In certain cases [e.g., microcystin (Nishizawa et al. 2001)], an independent AA racemase is incorporated into the biosynthetic operon, and is responsible for synthesis of a D-AA that is subsequently loaded onto the synthetase enzyme. More generally, however, NRPS enzymes include an epimerization domain (E domain) in the NRPS that is necessary for conversion of the L-AA into the D-AA prior to

5366 Fig. 2 Biochemical mechanism for D-AA synthesis by a pyridoxal-5′-phosphatedependent racemase. a Resting state of the PLP moiety with an internal lys aldimine. b Addition of an L-AA substrate displaces the internal lys aldimine, forming an external L-AA aldimine. c Removal of the α-proton from the L-AA substrate results in a resonance stabilized carbanion intermediate. d Protonation of the planar carbanion intermediate on the opposite face yields an external D-AA aldimine. The DAA is displaced through reformation of the internal lys aldimine

downstream processing of the peptide. Using the wellcharacterized mechanism for gramicidin S synthesis as an example, the GrsA enzyme first adenylates L-phe in an ATPdependent process, followed by formation of a covalent phosphopantetheinylated L-phe thioester adduct. The L-phe moiety is epimerized by the E domain, and the D-phe thioester adduct is processed by the GrsB enzyme to produce gramicidin S. E domains are common among NRPS enzymes and are presumed to be the source of the vast majority of D-AAs present in bacterial non-ribosomal peptides (Stein et al. 2005). Nonetheless, certain NRPS enzymes directly load DAAs rather than relying on E domains (Tang et al. 2007). Other examples exist whereby a D-AA is formed from its LAA counterpart in a cofactor-independent manner. The enolase superfamily member N-acylamino acid racemase (NAAA R) of Amycolatopsis sp. TS-1-60 catalyzed racemization of Nacylated AAs (e.g., N-acetyl-met), but not of the free AA itself (Tokuyama and Hatano 1995). The N-acylated D-AA could then be converted to a free D-AA by the action of a N-acyl-DAA deacylase (Tokuyama et al. 1994). The physiological role of the NAAAR racemization reaction is not known, however, and the reaction itself was proposed to be an adventitious reaction catalyzed by what was actually a promiscuous osuccinylbenzoate synthase (Palmer et al. 1999). Members of the enolase superfamily are also shown to catalyze epimerization of dipeptides (Schmidt et al. 2001).

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a

b

d

c

Transamination of α-ketoacids is a common mechanism for synthesis and interconversion of L-AAs, and a number of bacterial L-AA aminotransferases have been characterized (Massey et al. 1976; Umbarger 1978). Using the same basic mechanism, D-AA aminotransferase enzymes catalyze a twosubstrate, two-product reaction that utilizes an amino donor substrate (typically D-ala) and an α-ketoacid substrate. The amino group is ultimately transferred to the keto group of the α-ketoacid in a stereospecific manner, resulting in formation of a new α-ketoacid (e.g., pyruvate from D-ala) and a D-AA product (e.g., D-glu from α-ketoglutarate) (Fig. 3). Similar to certain racemase/epimerase enzymes, the reaction is reversible and utilizes a PLP cofactor; however, the mechanisms differ significantly. PLP-dependent aminotransferases utilize a twostep reaction in which the amino donor transfers the amino group to PLP, generating pyridoxamine-5′-phosphate (PMP) from PLP in the first half reaction. The amino group of PMP is transferred to the amino acceptor substrate during the second half reaction, generating the D-AA product. The substrate specificity of the second half reaction determines the D-AA formed. In the majority of cases studied, the enzyme shows narrow substrate specificity for the amino acceptor [typically αketoglutarate (Walton and Chica 2013)]. However, in some cases, the enzyme is demonstrated to donate the amino group to a wide range of α-ketoacid acceptors (Kobayashi et al. 2013), potentially generating a wide range of D-AAs in the cell.

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5367 Table 1 Characterized enzymes involved in D-AA synthesis and known physiological role of the respective D-AAs DAA

Method of synthesis

Role of D-AA synthesized using this mechanism

Racemase

PG peptide stem and PG interpeptide bridge (Vollmer et al. 2008); fusaracidin (Li and Jensen 2008); catabolism of L-ala (He et al. 2011)

Ala

Lantibiotics (posttranslational modification) Two enzymes: (Cotter et al. 2005) dehydratase and dehydrogenase D-AA aminotransferase Not known if this is a physiologically relevant method for D-ala synthesis (Kuramitsu and Snoke 1962) NRPS E domain

Daptomycin (Baltz et al. 2005); enduracidin/ ramoplanin (Yin and Zabriskie 2006); leinamycin (Tang et al. 2007)

Racemase

Unknown (Kato et al. 2012; Radkov and Moe 2013)

Arg

Fig. 3 D-AA synthesis via transamination. Depending on the direction of the (reversible) reaction, α-ketoglutarate can accept the amino group from D-ala, or pyruvate can accept the amino group from D-glu

Individual D-AAs and enzymes responsible for their synthesis Table 1 shows the mechanisms for synthesis and roles, where known, of the 19 canonical D-AAs, as well as D-ornithine and cis-D-hydroxyproline (cis-D-hypro). D-ala

All known bacterial ala racemases are PLP-dependent. Because of the ubiquity and necessity of D-ala in PG, ala racemases are established drug targets and consequently have become a well-studied enzyme class. The presence of two loci encoding distinct ala racemases was first demonstrated in the model organism Salmonella typhimurium (Wasserman et al. 1983). Mutagenesis studies revealed that one ala racemase converts L-ala into D-ala as part of the L-ala catabolic route (D-ala is subsequently converted into pyruvate), while the second ala racemase provides D-ala for PG synthesis. Since the publication of this first report, several studies have demonstrated similar genomic organization and physiological function for ala racemases, including organisms such as Escherichia coli and Pseudomonas aeruginosa PAO1 (He et al. 2011; Lobocka et al. 1994). Consequently, it has been accepted that Gram-negative bacteria encode two ala racemases (Conti et al. 2011). However, studies on the substrate specificity of the two annotated ala racemases of Pseudomonas putida KT2440 indicated a divergence of function between the two enzymes (Radkov and Moe 2013). One enzyme (DadX) displayed narrow substrate specificity, clearly

Asn Unknown

PG interpeptide bridge (Bellais et al. 2006)

NRPS E domain

Daptomycin (Baltz et al. 2005); fusaracidin (Li and Jensen 2008)

Racemase

PG interpeptide bridge (resistance to vancomycin) (Bellais et al. 2006; Veiga et al. 2006)

NRPS E domain

Bacitracin (Konz et al. 1997)

No known mechanism

No known role

Racemase

PG peptide stem and PG interpeptide bridge (Vollmer et al. 2008); poly-D-γ-glutamate (Candela and Fouet 2006); microcystin (Nishizawa et al. 2001)

D-AA aminotransferase

PG peptide stem (Fotheringham et al. 1998; Pucci et al. 1995)

NRPS E domain

Daptomycin (Baltz et al. 2005)

NRPS E domain

Arthrofactin (Balibar et al. 2005); fusaracidin (Li and Jensen 2008)

No known mechanism

No known role

Predicted epimerase

PG modification (Lam et al. 2009)

Asp

Cys

Glu

Gln

His

Ile Epimerase

Unknown (Mutaguchi et al. 2013)

NRPS E domain

Fusaracidin (Li and Jensen 2008)

Leu Predicted racemase

PG modification (Lam et al. 2009)

NRPS E domain

Bacitracin (Konz et al. 1997)

Unknown

PG peptide stem and PG interpeptide bridge (Boniface et al. 2006; Guinand et al. 1969)

Lys

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Table 1 (continued) DAA

Method of synthesis

Role of D-AA synthesized using this mechanism

Racemase

Unknown (Kato et al. 2012; Kuan et al. 2011; Radkov and Moe 2013)

NRPS E domain

Daptomycin (Baltz et al. 2005)

Predicted racemase

PG modification (Lam et al. 2009)

Met Phe NRPS E domain

Gramicidin (Stein et al. 1995) Tyrocidine (Marahiel et al. 1985) Bacitracin (Konz et al. 1997)

Unknown

PG modification (Lam et al. 2009)

Pro/cis-D-Hypro Racemase/epimerase

Catabolism of the respective L-enantiomer (Adams 1959; Gryder and Adams 1969; Stadtman and Elliot 1957)

Epimerase

Unknown (Radkov and Moe 2013)

Racemase

PG peptide stem (resistance to vancomycin) (Arias et al. 1999, 2000)

NRPS E domain

Daptomycin (Baltz et al. 2005); enduracidin/ ramoplanin (Yin and Zabriskie 2006); arthrofactin (Balibar et al. 2005);

Unknown

Found in culture supernatant (Lam et al. 2009)

NRPS E domain

Enduracidin/ramoplanin (Yin and Zabriskie 2006); arthrofactin (Balibar et al. 2005)

NRPS E domain

Daptomycin (Baltz et al. 2005)

Unknown

PG modification (Lam et al. 2009)

NRPS E domain

Tyrocidine (Marahiel et al. 1985)

Ser

Thr

dal gene) whose disruption leads to severe growth defects (Pierce et al. 2008 and references therein). B. subtilis, however, encodes a second ala racemase (encoded by the yncD gene) (Pierce et al. 2008). The physiological role of this racemase remains unclear due to the fact that a B. subtilis dal deletion mutant cannot grow in the absence of exogenous D-ala. Phylogenetic analysis revealed that the presence of two genes encoding ala racemase activity might be more widely distributed among Gram-positive bacteria than previously thought (Pierce et al. 2008). Other Gram-positive bacteria demonstrated to encode two ala racemases include Lactobacillus reuteri and Bacillus licheniformis (Salifu et al. 2008; Thompson et al. 2002). While D-AA aminotransferases are more commonly known to synthesize D-glu, it is important to note that the reaction can also favor synthesis of D-ala (Fig. 3). This activity has been detected in cell free extracts from B. licheniformis (Kuramitsu and Snoke 1962) and has also been demonstrated more recently with purified D-AA aminotransferases isolated from Bacillus sphaericus and Lactobacillus salivarius, respectively (Kobayashi et al. 2013; Yonaha et al. 1975). Even though the physiological function of this activity of D-AA aminotransferases has not been explored, it is noteworthy that the purified enzymes from these two studies demonstrated equal in vitro activity with pyruvate and α-ketoglutarate as amino acceptors. D-glu

Trp Tyr

Val Predicted racemase

PG modification (Lam et al. 2009)

NRPS E domain

Actinomycin D (Keller and Schauwecker 2001); penicillin (Martín et al. 1994); fusaracidin (Li and Jensen 2008)

Ornithine NRPS E domain

Bacitracin (Konz et al. 1997)

Racemase

Enduracidin/ramoplanin (Yin and Zabriskie 2006) Unknown (Kato et al. 2012)

preferring either enantiomer of ala as a substrate. The other enzyme (Alr) exhibited broad substrate specificity with the highest Kcat/KM values with either enantiomer of lys. Regarding their activities using ala enantiomers as a substrate, the Kcat/KM values of DadX were ∼10-fold higher than those of Alr. For Gram-positive bacteria, several distantly related species are shown to have a single ala racemase (encoded by the

Bacteria can use two different types of enzymes for the synthesis of D-glu, a racemase, and/or a D-AA aminotransferase. All known glu racemases belong to the PLP-independent class. Some bacteria encode only one glu racemase, yet others may possess two such racemases, and still others may encode both a racemase and a D-AA aminotransferase (Barreteau et al. 2008). Comparative genomic analysis indicates that all bacteria that synthesize PG encode at least one glu racemase (Fisher 2008). Glu racemase activity was first detected in lactobacilli (Ayengar and Roberts 1952; Narrod and Wood 1952), and the enzyme responsible for this activity has been characterized in a number of Gram-positive and Gram-negative bacteria. The E. coli glu racemase gene (murI) has been used as a model for D-glu synthesis. This gene was revealed to be both necessary and sufficient for D-glu synthesis in E. coli (Doublet et al. 1993). Lactobacillus spp. and Pediococcus spp., where studied, resemble E. coli in that they also possess a single gene encoding glu racemase (Gallo and Knowles 1993; Nakajima et al. 1986). In contrast, B. subtilis encodes two glu racemases (Kimura et al. 2004). The two enzymes, racE and yrpC, both racemize glu yet appear to possess different physiological functions. Knockout mutants in either gene showed that only racE was

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necessary and sufficient for growth in rich medium, while either racE or yrpC could provide D-glu during minimal medium growth. The presence of either gene was sufficient for PGA production during early stationary phase. However, both genes were necessary for the degradation of D-glu released during the capsule decomposition that normally occurs in late stationary phase. A genomic screen of Staphylococcus haemolyticus in an E. coli D-glu auxotroph revealed two ORFs that rescued D-glu auxotrophy (Pucci et al. 1995). One of the ORFs encoded a protein that showed homology to Pediococcus pentosaceus glu racemase, while the second ORF showed homology to a Bacillus sp. D-AA aminotransferase. Crude enzyme extracts were assayed from the recombinant clones to confirm the nature of the enzymatic activity. The presence of a glu racemase and D-AA aminotransferase conferring D-glu synthesis was also noted in B. sphaericus 10208 (Fotheringham et al. 1998). D-enantiomers

of lys, arg, and ornithine

The synthesis of D-lys, D-arg, and D-ornithine occurs via a PLPdependent racemase often referred to as a BAR (broad-spectrum AA racemase). These enzymes show the highest activity towards arg and lys but can also act on other AAs such as ornithine, thr, met, ala, leu, ser, and gln (Matsui et al. 2009; Radkov and Moe 2013). Although arg/lys racemization has been detected in Oenococcus oeni (Kato et al. 2012) as well as Proteus spp. (P. mirabilis and P. vulgaris) (Kuan et al. 2011; Wu et al. 2012), the majority of the work on synthesis of D-lys, Darg, and D-ornithine has been done in pseudomonads. Periplasmic arg/lys racemase activity has been detected in P. putida and Pseudomonas taetrolens (Ichihara et al. 1960; Matsui et al. 2009). The gene responsible for P. putida KT2440 BAR activity was identified using a genomic screen (Radkov and Moe 2013). The recovered gene, alr, was annotated as an ala racemase but showed in vitro enzymatic activity consistent with a BAR. The physiological role of BARs in pseudomonads is not known, although they have been proposed to act in D-lys catabolism (Revelles et al. 2005, 2007). Independent pathways for the catabolism of D- and L-lys have been demonstrated in pseudomonads, however (Chang and Adams 1974; Muramatsu et al. 2005). Considering the periplasmic localization of BARs, and the recent discovery of D-lys in the PG peptide stem in Thermotoga maritima (Boniface et al. 2009), BARs may be involved in PG modification or synthesis of D-lys for signaling purposes, although this has not been experimentally determined. In addition to a putative biosynthetic role, this enzyme may also play a part in D-arg catabolism. The P. putida KT2440 alr gene is located between two genes, aruI and aruH, whose orthologs are involved in catabolism of D- and L-arg in P. aeruginosa PAO1 (Radkov and Moe 2013; Yang and Lu 2007). The location of the P. aeruginosa PAO1 alr ortholog is not conserved between the two organisms.

5369 D-asp D-asp

can be produced via cofactor-independent racemases and by D-AA aminotransferases. The first bacterial asp racemase was detected in Enterococcus faecalis (formerly Streptococcus faecalis) (Lamont et al. 1972), and it is now accepted that asp racemases are predominantly found in lactic acid bacteria due to the presence of D-asp in the interpeptide bridge of their PG layer (Conti et al. 2011). Further work has demonstrated the presence of asp racemases in Pyrococcus horikoshii, Bifidobacterium bifidum, and Enterococcus faecium (Bellais et al. 2006; Liu et al. 2002; Yamashita et al. 2004). A two-gene cluster was identified (encoding asp racemase and asp ligase) that facilitates the incorporation of D-asp into the PG of E. faecium (Bellais et al. 2006). Additionally, comparative genomic analysis identified homologs of the asp racemase in Lactobacillus spp., Lactococcus spp., and P. pentosaceus. Some of the first reports on D-AA aminotransferases came from Bacillus species, such as B. licheniformis (Kuramitsu and Snoke 1962), owing to a proposed role in synthesis of D-AAs for the antibiotic bacitracin. Using cellfree extracts, enzyme assays were performed with D-ala as the amino donor and a set of three ketoacids (α-ketoglutarate, pyruvate, and oxaloacetate) to establish that D-glu, D-asp, and a number of other D-AAs could be produced via transamination. Aside from lactic acid bacteria, however, little is known about the occurrence and physiological role of D-asp. D-ser

PLP-dependent ser racemases have been implicated in the resistance of enterococci towards the glycopeptide antibiotic vancomycin. Vancomycin binds to the terminal D-ala-D-ala moiety in the peptide stem, and replacement of the canonical dipeptide with D-ala-D-ser in the VanC phenotype limits binding of vancomycin to the peptide stem. The vanT gene from the VanC-encoding E. gallinarum BM4174 exhibits homology with known AA racemases and genetic and biochemical evidence supports its role as the ser racemase involved in the VanC phenotype (Arias et al. 1999). While VanT can also racemize ala, the enzyme exhibits a ∼10-fold greater efficiency for the racemization of D-ser (Arias et al. 2000). The VanT racemase is a membrane-bound enzyme and is the only known bacterial racemase with a transmembrane domain. The transmembrane domain may be involved in transport of L-ser from the external medium (Arias et al. 2003). D-Enantiomers

of branched-chain amino acids (D-BCAAs)

An ile epimerase was recently identified in the lactic acid bacterium Lactobacillus otakiensis JCM 15040 (Mutaguchi et al. 2013). The PLP-dependent enzyme epimerized both stereoisomers of ile (L-ile and D-allo-ile, 100 % of activity)

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but also reacted with D-/L-val (48 %) and D-/L-leu (30 %). Minimal activity was also confirmed with both enantiomers of phe and met. It was reported that L. otakiensis JCM 15040 releases high amounts of D-BCAAs into the growth medium, such as 200 μM D-leu, 150 μM D-allo-ile, and 50 μM D-val. Incidentally, V. cholerae has also been shown to secrete D-val, D-allo-ile, D-leu, and D-met (Lam et al. 2009). In the case of V. cholerae, synthesis was attributed to a periplasmic broadspectrum AA racemase (BsrV), as a bsrV knockout failed to produce the D-AAs. Subsequent biochemical characterization of the enzyme showed that it can racemize 12 AAs in vitro (ala, ser, met, leu, cys, gln, asn, his, lys, arg, val, and ile) and the highest overall efficiency was with L-lys (Kcat/KM of 52.9 mM−1 s−1; kinetic values with D-AAs were not determined) (Espaillat et al. 2014). Comparative genomic analysis revealed that BsrV orthologs can be found in multiple species. The bsrV gene exhibits homology with ala racemase genes, but shows little identity with ile epimerase. D-pro

and cis-D-hypro

Pro racemase and hypro epimerase are cofactor-independent enzymes that share a high degree of identity yet are very specific for their respective substrates. Extensive studies on the pro racemase from the eukaryotic pathogen Trypanosoma cruzi have provided a basis for identification of prokaryotic pro racemases (Buschiazzo et al. 2006; Chamond et al. 2003). However, studies on bacterial synthesis of D-pro are very limited. The only bacterial pro racemase activity to date was detected in Clostridium sticklandii (Stadtman and Elliot 1957), where D-pro is synthesized as an intermediate in the catabolism of L-pro. Despite the abundance of annotated pro racemase genes in bacterial genomes, it may be unlikely that these genes encode true pro racemases. Where studied, these genes are shown to encode hypro epimerase activity instead. This includes genes from B. anthracis, Pseudomonas spp., Brucella spp., and Burkholderia spp. (Goytia et al. 2007; Radkov and Moe 2013). Indeed, much more is known about bacterial hypro epimerases and their physiological role—a trend that may be explained by the fact that hypro is highly prevalent in collagen as well as plant cell walls (Bella et al. 1994; Cassab 1998). Similar to catabolism of pro described above, conversion to the D-epimer of hypro is a necessary step in L -hypro catabolism in Pseudomonas spp. (Adams 1959; Gryder and Adams 1969) and in Sinorhizobium meliloti (White et al. 2012). Hypro epimerases exhibit remarkably narrow substrate specificity when characterized in vitro, with effectively no activity on pro or either stereoisomer of the other canonical AAs (Goytia et al. 2007; Radkov and Moe 2013).

D-AAs

in biotechnology

Because of the importance of PG to most bacteria, enzymes involved in synthesis of its components and its superstructure are seen as key targets for antibacterials. Among the myriad reactions involved in this process, many successful drugs target the extracellular or periplasmic reactions necessary for cross-linking the glycan chains (e.g., β-lactams). Resistance to these drugs is abundant, and PG plasticity can limit efficacy. Drugs targeting the cytoplasmic reactions of PG synthesis, including D-AA synthesis, are gaining favor (Conti et al. 2011). Racemases responsible for synthesis of D-AAs required for PG have been targeted. Inhibitors of ala racemase include a collection of natural compounds (e.g., D-cycloserine) as well as synthetic ala analogs (e.g., β-haloalanines) that form irreversible adducts with PLP. While some successes are noted, inhibitors must be specific for the ala racemase enzyme as a general PLP-dependent enzyme inhibitor results in toxic human side effects. As the other racemase essential for PG, glu racemase is also considered a target, and some glu analogs exhibit promising results as antibacterials (Conti et al. 2011). D-AAs themselves exhibit potential as modulators of bacterial growth and persistence. Much has been made of their reported ability to disassemble mature biofilms (Hochbaum et al. 2011; Kolodkin-Gal et al. 2010). While there is some question about the roles of D-AAs in this process (Leiman et al. 2013; Sanchez et al. 2013), it is clear that their incorporation into PG is a method by which bacteria adjust to changes in environmental conditions (Lam et al. 2009). The prevalence of D-AAs in bioactive natural products (NPs) illustrates their importance as key functionalities in NPs as well as their potential as stereochemical building blocks in NP-like chemicals. Table S1 details the relative abundances of individual D-AAs in NRPs, and Table 1 indicates the known mechanisms by which they are synthesized for NRPs. D-AAs in NPs can provide chemical functionalities that confer unique properties on the molecule. For example, two bacteriocins that exhibit the same primary AA sequence and molecular weight but differ only in the stereochemistry of one AA display different bioactivities (Kawai et al. 2004). Because both D- and L-AAs exhibit similar chemistries (i.e., the incorporation of a D- versus an L-AA proceeds the same way), this provides a simple means for chemodiversification of chemical scaffolds. Further, D-AA-containing peptides are shown to confer a number of potent biological activities including antibacterial, antitumor, and antiangiogenic properties (Dawson et al. 1999; Dey et al. 2013; Fernandez-Lopez et al. 2001; Güell et al. 2011), they inhibit HIV entry into cells (Welch et al. 2007), reduce atherosclerosis (Navab 2002), and inhibit amyloid-β peptide fibrillogenesis, which is implicated in Alzheimer’s disease (Soto et al. 1996). D-AA containing peptides exhibit the same molecular weights as their L-

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counterparts as well as similar hydrophobicities. Further, most proteases do not recognize D-AAs, which renders these peptides more stable in biological systems (Güell et al. 2011; Sharma et al. 2010; Wei and Bobek 2005).

Conclusions and future directions The synthesis and catabolism of D-AAs, as well as the roles that they play in bacterial physiology and community structure, are all areas that warrant further study. Incorrectly annotated racemase and aminotransferase genes are currently an impediment to the study of D-AAs in bacteria. In particular, genes annotated as ala racemases and pro racemases are often discovered to encode activities that are not consistent with their designation (Goytia et al. 2007; Radkov and Moe 2013). Further biochemical characterization of these genes, across the prokaryotic domain, would enable a more accurate description of both the PLP-dependent and PLP-independent racemases based solely on gene sequence. The newly discovered ile epimerase and BARs are intriguing new additions to the racemase family. Unfortunately, these enzymes are typically misannotated, making it difficult to assess their abundance across genomes. Additional studies may also shed light on the differences between bacteria in the number of racemase genes encoded. There is some evidence suggesting different roles for paralogous racemase genes (e.g., catabolism versus synthesis of D-AAs). This research should be expanded to include biochemical and genetic studies of bacteria encoding multiple related racemases. An exploration of the role of D-AA aminotransferase enzymes in these processes should be considered as well. There is a dearth of information on the events that trigger DAA synthesis in bacteria. Certain D-AAs accumulate in large quantities in culture supernatant during stationary phase (Lam et al. 2009; Mutaguchi et al. 2013), yet much more remains to be discovered on the regulatory mechanisms that control their synthesis. Conversely, we know little about how the presence of D-AAs controls gene expression in bacteria. In at least one case, they are proposed to act as modulators of bacterial behavior based on their incorporation into PG (Cava et al. 2011; Lam et al. 2009). Much remains to be discovered on the scope and limits of D-AA incorporation into PG, as well as the genetics and biochemistry involved in sensing and responding to free D-AAs. Owing to the limited scope of this article, we have discussed only D-AA synthesis and have neglected D-AA catabolism. Levels of free D-AAs in the cell are impacted by the rates of their synthesis, incorporation into macromolecules, and by their degradation. D-AAs are degraded either by conversion to the corresponding L-AA or by direct oxidation to the α-ketoacid by a D-AA dehydrogenase or D-AA

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oxidase. Little is known about the intracellular levels of free DAAs and about the specific roles of these enzymes although DAA dehydrogenase activity appears to be more widespread than D-AA oxidase activity, which is considered to be a primarily eukaryotic enzyme (Pollegioni et al. 2008). In at least one case, D-AA dehydrogenase expression is induced by its D-AA substrates (Li et al. 2010). Advances in technology are making possible highresolution structural analyses of PG. This includes imaging studies of live cells (Siegrist et al. 2013), mass spectrometry of prepared PG layers (Boniface et al. 2009), as well as in vivo incorporation of fluorescently labeled D-AAs. Fluorescent DAAs were used to monitor incorporation of D-AAs into a diverse collection of bacteria, including E. coli, B. sublitis, Caulobacter crescentus, Streptomyces venezuelae, Agrobacterium tumefaciens, Staphylococcus aureus, Streptococcus pneumoniae, Listeria monocytogenes, and Chlamydia trachomatis (Kuru et al. 2012; Liechti et al. 2014). In concert with basic biochemical and genetic studies, as well as multiomic analyses, these techniques should provide an outstanding opportunity to study the roles of D-AAs in all aspects of bacterial physiology. Acknowledgments Research on D-amino acid synthesis in the PI’s lab is funded in part by a grant from the National Institute of Food and Agriculture of the US Department of Agriculture (grant 2011-6702030195).

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