Efficient Bioethanol Production by a Recombinant Flocculent Saccharomyces cerevisiae Strain with a Genome-Integrated NADP+-Dependent Xylitol Dehydrogenase Gene

The recombinant industrial Saccharomyces cerevisiae strain MA-R5 was engineered to express NADP⁺-dependent xylitol dehydrogenase using the flocculent yeast strain IR-2, which has high xylulose-fermenting ability, and both xylose consumption and ethanol production remarkably increased. Furthermore, the MA-R5 strain produced the highest ethanol yield (0.48 g/g) from nonsulfuric acid hydrolysate of wood chips.Successful fermentation of lignocellulosic biomass to ethanol is dependent on efficient utilization of d-xylose, which is the second most common fermentable sugar in the hydrolysate. Although the well-known fermentative yeast Saccharomyces cerevisiae is one of the most effective ethanol-producing organisms for hexose sugars, it is not able to ferment d-xylose. However, S. cerevisiae can metabolize an isomerization product of d-xylose, d-xylulose, which is phosphorylated to d-xylulose 5-phosphate, channeled through the pentose phosphate pathway and glycolysis.S. cerevisiae transformed with the XYL1 and XYL2 genes encoding xylose reductase (XR) and xylitol dehydrogenase (XDH) from Pichia stipitis (referred to as PsXR and PsXDH, respectively) acquires the ability to ferment d-xylose to ethanol (2, 5, 6, 9, 10, 12, 22). Furthermore, overexpression of the XKS1 gene encoding xylulokinase (XK) from S. cerevisiae (ScXK) has been shown to aid d-xylose utilization (4, 7, 11, 16, 23), with xylitol still a major by-product. Kuyper et al. (14) also demonstrated the successful fermentation of d-xylose to ethanol using recombinant S. cerevisiae strains expressing fungal xylose isomerase. However, these approaches are insufficient for industrial bioprocesses, mainly due to the low rate of d-xylose fermentation.Xylitol excretion has been ascribed mainly to the difference in coenzyme specificities between PsXR (with NADPH) and PsXDH (with NAD⁺), which creates an intracellular redox imbalance (1). Therefore, modifying the coenzyme specificity of XR and/or XDH by protein engineering is an attractive approach for achieving efficient fermentation of ethanol from d-xylose using recombinant S. cerevisiae. Watanabe et al. (24) previously succeeded in generating several PsXDH mutants (e.g., quadruple ARSdR mutant) with a complete reversal of coenzyme specificity toward NADP⁺ by multiple site-directed mutagenesis on amino acids from the coenzyme-binding domain. The ARSdR mutant (D207A/I208R/F209S/N211R) has more that 4,500-fold-higher catalytic efficiency (k_cat/K_m) with NADP⁺ than the wild-type PsXDH. In addition, we recently found that several laboratory recombinant S. cerevisiae strains, in which the ARSdR mutant, along with PsXR and ScXK, is expressed through a strong constitutive promoter, increased ethanol production from d-xylose at the expense of xylitol excretion (17, 18), probably by maintaining the intracellular redox balance. However, commercialization of lignocellulosic hydrolysate fermentation requires industrial strains that are more robust than laboratory strains (5, 19, 21).A potential host for developing d-xylose-fermenting strains requires an active and efficient pentose phosphate pathway linking the d-xylose-to-d-xylulose pathway to glycolysis. In the case of S. cerevisiae, strains have different d-xylulose fermentation abilities (3, 25), indicating inherent differences in the capacities of these strains to ferment pentose sugars. Furthermore, anaerobic d-xylulose fermentation was investigated to identify genetic backgrounds potentially beneficial to anaerobic d-xylose fermentation in strains not exhibiting product formation related to the redox imbalance generated by the first two steps of the eukaryotic d-xylose metabolism (3), although the physiological purpose of the different d-xylulose fermentation abilities of S. cerevisiae is not yet clear. Therefore, we selected an efficient industrial d-xylulose-fermenting S. cerevisiae strain as a host for constructing a recombinant strain through chromosomal integration of the NADP⁺-dependent XDH gene and the XR and endogenous XK genes. Using this recombinant strain, we characterized the enzyme activity and ability to ferment both d-xylose and lignocellulosic hydrolysate.     

Isolation and Characterization of a Novel Lysine Racemase from a Soil Metagenomic Library

A lysine racemase (lyr) gene was isolated from a soil metagenome by functional complementation for the first time by using Escherichia coli BCRC 51734 cells as the host and d-lysine as the selection agent. The lyr gene consisted of a 1,182-bp nucleotide sequence encoding a protein of 393 amino acids with a molecular mass of about 42.7 kDa. The enzyme exhibited higher specific activity toward lysine in the l-lysine-to-d-lysine direction than in the reverse reaction.Amino acids are the building blocks of proteins and play an important role in the regulation of the metabolism of living organisms. Among two enantiomers of naturally occurring amino acids, l-amino acids are predominant in living organisms, while d-amino acids are found in both free and bound states in various organisms like bacteria (36), yeasts (35), plants (47), insects (11), mammals (17), bivalves (39), and fish (28). The d-amino acids are mostly endogenous and produced by racemization from their counterparts by the action of a racemase. Thus, the amino acid racemases are involved in d-amino acid metabolism (29, 46). Since the discovery of alanine racemase in 1951 (42), several racemases toward amino acids, such as those for glutamate, threonine, serine, aspartate, methionine, proline, arginine, and phenylalanine, have been reported in bacteria, archaea, and eukaryotes, including mammals (1, 2, 15, 30, 31, 44). They are classified into two groups: pyridoxal 5′-phosphate (PLP)-dependent and PLP-independent enzymes (9, 36).Lysine racemase (Lyr, EC 5.1.1.5) was first reported in Proteus vulgaris ATCC 4669 (19) and proposed to be involved in the lysine degradation of bacterial cells (5, 19). Catabolism of lysine occurs via two parallel pathways. In one of the pathways, δ-aminovalerate is the key metabolite, whereas in the other l-lysine is racemized to d-lysine, and l-pipecolate and α-aminoadipate (AMA) are the key metabolites (5). d-Lysine catabolism proceeds through a series of cyclized intermediates which are necessary to regenerate an α-amino acid and comprise the following metabolites (AMA pathway): d-lysine→α-keto-ɛ-amino caproate→Δ¹-piperideine-2-carboxylate→pipecolate→Δ¹-piperideine-6-carboxylate→α-amino-δ-formylcaproate→α-AMA→α-ketoadipate (6, 7, 12, 27). The final product is converted to α-ketoglutarate via a series of coenzyme A derivatives and subsequently participates as an intermediate in the Krebs cycle. This pathway suggests that the biological function of d-lysine in the bacteria is that of a carbon or nitrogen source. Racemization of added l-lysine to d-lysine by whole cells of Proteus spp. and Escherichia spp. (19) and by the cell extract of Pseudomonas putida ATCC 15070 (5, 20) has been found. However, the enzyme has not been purified to homogeneity, and thus, its molecular and catalytic characteristics, including its gene structure, have not been elucidated. In this study, we explored a metagenomic library constructed from a garden soil to isolate a novel Lyr enzyme. After expression in Escherichia coli, the purified enzyme was characterized in terms of optimal pH and temperature, thermal stability, and racemization activity.     

Design of a Potent d-Peptide HIV-1 Entry Inhibitor with a Strong Barrier to Resistance

The HIV gp41 N-trimer pocket region is an ideal viral target because it is extracellular, highly conserved, and essential for viral entry. Here, we report on the design of a pocket-specific d-peptide, PIE12-trimer, that is extraordinarily elusive to resistance and characterize its inhibitory and structural properties. d-Peptides (peptides composed of d-amino acids) are promising therapeutic agents due to their insensitivity to protease degradation. PIE12-trimer was designed using structure-guided mirror-image phage display and linker optimization and is the first d-peptide HIV entry inhibitor with the breadth and potency required for clinical use. PIE12-trimer has an ultrahigh affinity for the gp41 pocket, providing it with a reserve of binding energy (resistance capacitor) that yields a dramatically improved resistance profile compared to those of other fusion inhibitors. These results demonstrate that the gp41 pocket is an ideal drug target and establish PIE12-trimer as a leading anti-HIV antiviral candidate.The HIV envelope protein (Env) mediates viral entry into cells (11). Env is cleaved into surface (gp120) and transmembrane (gp41) subunits that remain noncovalently associated to form trimeric spikes on the virion surface (16). gp120 recognizes target cells by interacting with cellular receptors, while gp41 mediates membrane fusion. Peptides derived from heptad repeats near the N and C termini of the gp41 ectodomain (N and C peptides) interact in solution to form a six-helix bundle, representing the postfusion structure (3, 55, 56). In this structure, N peptides form a central trimeric coiled coil (N trimer), creating grooves into which C peptides bind. This structure, in conjunction with the dominant-negative inhibitory properties of exogenous N and C peptides, suggests a mechanism for Env-mediated entry (10, 22, 58-60).During entry, gp41 forms an extended prehairpin intermediate that leaves the exposed N-trimer region vulnerable to inhibition for several minutes (18, 35). This intermediate ultimately collapses as the C-peptide regions bind to the N-trimer grooves to form a trimer of hairpins (six-helix bundle), juxtaposing viral and cellular membranes and inducing fusion. Enfuvirtide (Fuzeon), the only clinically approved HIV fusion inhibitor, is a C peptide that binds to part of the N-trimer groove and prevents six-helix bundle formation in a dominant-negative manner (61). Enfuvirtide is active in patients with multidrug resistance to other classes of inhibitors and is a life-prolonging option for these patients (30, 31). However, enfuvirtide use is restricted to salvage therapy due to several limitations, including (i) high dosing requirements (90 mg, twice-daily injections), (ii) high cost (∼$30,000/year/patient in the United States), and (iii) the rapid emergence of resistant strains (21, 47).A deep hydrophobic pocket at the base of the N-trimer groove is an especially attractive inhibitory target because of its high degree of conservation (3, 12, 48), poor tolerance to substitution (4, 34), and critical role in membrane fusion (2). Indeed, this region is conserved at both the amino acid level (for gp41 function in membrane fusion) and the nucleotide level (for the structured RNA region of the Rev-responsive element). Enfuvirtide binds to the N-trimer groove just N terminal to the pocket and is significantly more susceptible to resistance mutations than 2nd-generation C-peptide inhibitors, such as T-1249, that also bind to the pocket (8, 13, 29, 44, 46, 47, 58).Peptide design, molecular modeling, and small-molecule screening have produced a diverse set of compounds that interact with the gp41 pocket and inhibit HIV-1 entry with modest potency, but often with significant cytotoxicity (7, 14, 15, 17, 23, 24, 26, 34, 51, 54). The first direct evidence that pocket-specific binders are sufficient to inhibit HIV entry came with the discovery of protease-resistant d-peptides identified using mirror-image phage display (12). In this technique, a phage library is screened against a mirror-image version of the target protein (synthesized using d-amino acids) (50). By symmetry, mirror images (d-peptides) of the discovered sequences will bind to the natural l-peptide target. As the mirror images of naturally occurring l-peptides, d-peptides cannot be digested by natural proteases. Protease resistance provides d-peptides theoretical treatment advantages of extended survival in the body and possible oral bioavailability (41, 42, 49).These 1st-generation d-peptide entry inhibitors possess potency against a laboratory-adapted isolate (HXB2) at low to mid-μM concentrations (12). We previously reported an affinity-matured 2nd-generation d-peptide called PIE7, pocket-specific inhibitor of entry 7 (57). A trimeric version of PIE7 is the first high-affinity pocket-specific HIV-1 inhibitor and has potency against X4-tropic (HXB2) and R5-tropic (BaL) strains at sub-nM concentrations. However, significant further optimization is required to create a robust clinical candidate for two reasons. First, this d-peptide is much less potent (requiring high nM concentrations) against JRFL, a primary R5-tropic strain. Therefore, improved PIE potency is necessary to combat diverse primary strains. Second, by improving the affinity of our inhibitors for the pocket target, we hope to provide a reserve of binding energy that will delay the emergence of drug resistance, as described below.We and others have reported a potency plateau for some gp41-based fusion inhibitors that is likely imposed by the transient exposure of the prehairpin intermediate (9, 27, 53, 57). For very high-affinity inhibitors, association kinetics (rather than affinity) limits potency so that two inhibitors with significantly different affinities for the prehairpin intermediate can have similar antiviral potencies. We proposed that overengineering our d-peptides with substantial affinity beyond this potency plateau would provide a reserve of binding energy that would combat affinity-disrupting resistance mutations (57). Such a resistance capacitor should also prevent the stepwise accumulation of subtle resistance mutations in Env by eliminating the selective advantage that such mutants would otherwise confer.Here, we report on the design and characterization of a 3rd-generation pocket-specific d-peptide, PIE12-trimer, with ∼100,000-fold improved target binding compared to that of the best previous d-peptide, significantly broadened inhibitory potency, and an enhanced resistance capacitor that provides a strong barrier to viral resistance. We achieved this increased potency via structure-guided phage display and crosslinker optimization. PIE12-trimer has a dramatically improved resistance profile compared to the profiles of earlier d-peptides, as well as those of enfuvirtide and T-1249. These results validate the resistance capacitor hypothesis and establish PIE12-trimer as a leading anti-HIV therapeutic candidate.     

Molecular Basis of Vancomycin Dependence in VanA-Type Staphylococcus aureus VRSA-9

The vancomycin-resistant Staphylococcus aureus VRSA-9 clinical isolate was partially dependent on glycopeptide for growth. The responsible vanA operon had the same organization as that of Tn1546 and was located on a plasmid. The chromosomal d-Ala:d-Ala ligase (ddl) gene had two point mutations that led to Q260K and A283E substitutions, resulting in a 200-fold decrease in enzymatic activity compared to that of the wild-type strain VRSA-6. To gain insight into the mechanism of enzyme impairment, we determined the crystal structure of VRSA-9 Ddl and showed that the A283E mutation induces new ion pair/hydrogen bond interactions, leading to an asymmetric rearrangement of side chains in the dimer interface. The Q260K substitution is located in an exposed external loop and did not induce any significant conformational change. The VRSA-9 strain was susceptible to oxacillin due to synthesis of pentadepsipeptide precursors ending in d-alanyl-d-lactate which are not substrates for the β-lactam-resistant penicillin binding protein PBP2′. Comparison with the partially vancomycin-dependent VRSA-7, whose Ddl is 5-fold less efficient than that of VRSA-9, indicated that the levels of vancomycin dependence and susceptibility to β-lactams correlate with the degree of Ddl impairment. Ddl drug targeting could therefore be an effective strategy against vancomycin-resistant S. aureus.Methicillin-resistant Staphylococcus aureus (MRSA) bacteria that have acquired the vancomycin resistance vanA operon from glycopeptide-resistant enterococci are designated vancomycin-resistant S. aureus (VRSA) (29). Vancomycin acts by binding to the C-terminal acyl-d-Ala-d-Ala of the undecaprenol-diphosphate MurNAc-pentapeptide intermediate and inhibits transglycosylation and transpeptidation reactions in cell wall peptidoglycan polymerization and cross-linking (30). d-Ala-d-Ala is synthesized by the ATP-dependent d-Ala:d-Ala ligase (Ddl) (EC 6.3.2.4) before its incorporation in peptidoglycan precursors (26, 35). VanA-type vancomycin resistance results from the incorporation into peptidoglycan intermediates of a d-alanyl-d-lactate (d-Ala-d-Lac) depsipeptide, synthesized by a d-Ala:d-Lac ligase, which is responsible for diminished binding affinity of glycopeptides for their target. Kinetic analyses of Ddls have established two subsites in the active site for d-Ala binding (24, 27). The reaction mechanism culminates in the transfer of the γ-phosphoryl of ATP to the carboxyl group of d-Ala₁ to produce an acylphosphate and ADP. The acyl carbon atom of the acylphosphate then reacts with the amino group of d-Ala₂ to yield a tetrahedral intermediate. Finally, the intermediate releases phosphate to yield d-Ala-d-Ala.Mutants of Enterococcus faecium (8, 14), Enterococcus faecalis (34), and S. aureus (23) with an impaired Ddl are able to grow because they use the vancomycin resistance pathway for cell wall synthesis. Since resistance is inducible by the drug, these bacteria require the presence of vancomycin in the culture medium for growth. Ddls from vancomycin-dependent enterococci (14) have mutations affecting amino acids highly conserved in the d-Ala:d-Ala ligase superfamily (10). Molecular modeling based on the X-ray structure of Escherichia coli DdlB (11) revealed that all the mutated residues interact directly with one of the substrates of the enzymatic reaction or stabilize the position of critical residues in the active site. However, the degree of enzyme impairment was not evaluated biochemically. Recently, we reported the mechanism of vancomycin dependence in VanA-type S. aureus VRSA-7 and showed that the chromosomal Ddl had the single mutation N308K, which probably affects the binding of the transition-state intermediate, leading to a 1,000-fold decrease in activity relative to that of the wild-type enzyme (23). Glycopeptide-dependent mutants could therefore be considered useful tools to explore structure-activity relationships of the Ddl, which represents an attractive target for designing new drugs. Here we describe the partially vancomycin-dependent VanA-type S. aureus strain VRSA-9 and report the biochemical and structural characterization of its mutated Ddl.     

The Aminolysis Reaction of Streptomyces S9 Aminopeptidase Promotes the Synthesis of Diverse Prolyl Dipeptides

Prolyl dipeptide synthesis by S9 aminopeptidase from Streptomyces thermocyaneoviolaceus (S9AP-St) has been demonstrated. In the synthesis, S9AP-St preferentially used l-Pro-OBzl as the acyl donor, yielding synthesized dipeptides having an l-Pro-Xaa structure. In addition, S9AP-St showed broad specificity toward the acyl acceptor. Furthermore, S9AP-St produced cyclo (l-Pro-l-His) with a conversion ratio of substrate to cyclo (l-Pro-l-His) higher than 40%.Some proline-containing dipeptides and their cyclic analogs exhibit biological activity. For example, cyclo (l-arginyl-d-proline) [c(lR-dP)] is known to act as a specific inhibitor of family 18 chitinase (4, 10). A cyclic peptide, c(lP-lH), produced by the cleavage of thyrotropin-releasing hormone protects against oxidative stress, promotes cytoprotection (6, 7), and exhibits antihyperglycemic activity (11).Some serine peptidases exhibit peptide bond formation (i.e., aminolysis of esters, thioesters, and amides) in accordance with their hydrolytic activity (2, 14). The exchange of catalytic Ser for Cys to engineer the serine endopeptidase into “transpeptidase” for peptide bond formation has been well characterized (3, 5). Our recent approach confirmed the wide distribution of family S9 aminopeptidases that have catalytic Ser in actinomycetes (12). Of them, we obtained S9 aminopeptidase from Streptomyces thermocyaneoviolaceus NBRC14271 (S9AP-St). The enzyme was engineered into “transaminopeptidase” by exchange of catalytic Ser for Cys, and its aminolytic activity was evaluated (13). The engineered enzyme, designated as aminolysin-S, can synthesize hydrophobic dipeptides through an aminolysis reaction. However, aminolysin-S was unable to synthesize peptides containing proline. Although the report of aminolysin-S demonstrated that S9AP-St shows no aminolysis reaction toward limited substrates, details of its characteristics remain unknown. This study verified the peptide synthetic activity of S9AP-St, demonstrating that S9AP-St can synthesize widely varied prolyl dipeptides through an aminolysis reaction. The report also shows that S9AP-St is applicable to the synthesis of a biologically active peptide—c(lP-lH).     

Functional Expression of a Bacterial Xylose Isomerase in Saccharomyces cerevisiae

In industrial fermentation processes, the yeast Saccharomyces cerevisiae is commonly used for ethanol production. However, it lacks the ability to ferment pentose sugars like d-xylose and l-arabinose. Heterologous expression of a xylose isomerase (XI) would enable yeast cells to metabolize xylose. However, many attempts to express a prokaryotic XI with high activity in S. cerevisiae have failed so far. We have screened nucleic acid databases for sequences encoding putative XIs and finally were able to clone and successfully express a highly active new kind of XI from the anaerobic bacterium Clostridium phytofermentans in S. cerevisiae. Heterologous expression of this enzyme confers on the yeast cells the ability to metabolize d-xylose and to use it as the sole carbon and energy source. The new enzyme has low sequence similarities to the XIs from Piromyces sp. strain E2 and Thermus thermophilus, which were the only two XIs previously functionally expressed in S. cerevisiae. The activity and kinetic parameters of the new enzyme are comparable to those of the Piromyces XI. Importantly, the new enzyme is far less inhibited by xylitol, which accrues as a side product during xylose fermentation. Furthermore, expression of the gene could be improved by adapting its codon usage to that of the highly expressed glycolytic genes of S. cerevisiae. Expression of the bacterial XI in an industrially employed yeast strain enabled it to grow on xylose and to ferment xylose to ethanol. Thus, our findings provide an excellent starting point for further improvement of xylose fermentation in industrial yeast strains.It is widely acknowledged that fuels from regenerative resources are becoming increasingly important in times of a dwindling crude oil supply and the growing environmental concern of the public. Plant biomass, particularly when accruing as a waste product, is an attractive feedstock for bioethanol production. An important prerequisite for such an alternative strategy would be the complete conversion of all available sugars in biomass hydrolysates into ethanol. While the hexose sugars are easily fermentable, no suitable microorganism is available for fermenting pentose into ethanol. Calculations have resulted in an estimate that production of lignocellulosic ethanol would reduce the cost of producing ethanol by nearly 20% (3). Therefore, ethanol production from pentose sugars has received considerable attention (4, 9).Although some anaerobic fungi and bacteria are able to metabolize xylose, they are not suitable for industrial bioethanol production due to low and inefficient production rates and the mixed acid fermentation life-style (28), which generates too many by-products. The baker''s yeast Saccharomyces cerevisiae remains the organism of choice for industrial production of ethanol. However, while hexoses are converted rapidly to high yields of ethanol, wild-type S. cerevisiae strains are not able to ferment pentose sugars, such as d-xylose and l-arabinose, efficiently. Several different approaches in genetic engineering have been used to enable d-xylose fermentation in yeast.Successful xylose fermentation in recombinant S. cerevisiae strains was previously achieved by heterologous expression of the XYL1 and XYL2 genes (encoding xylose reductase [XR] and xylitol dehydrogenase [XDH], respectively) from Pichia stipitis (8, 12, 14, 15) or by expression of a xylA gene (encoding xylose isomerase [XI]) from Piromyces sp. strain E2 (17) or Thermus thermophilus (33). Both approaches resulted in strains growing on xylose and fermenting it into ethanol. Although expression of XR and XDH resulted in rapid fermentation of xylose, NADPH/NAD cofactor imbalance under anaerobic conditions led to considerable accumulation of xylitol (6, 14, 15, 30, 32). However, employing XI instead of XR/XDH avoids cofactor imbalance and xylitol accumulation, as d-xylose is converted directly into d-xylulose without a redox reaction being involved.Many attempts to express an active prokaryotic XI in S. cerevisiae have failed. None of the efforts to express XI from Escherichia coli (25), Bacillus subtilis (2), Lactobacillus pentosus (10), or Clostridium thermosulfurogenes (23) in S. cerevisiae resulted in active XI, arguing for the inability of yeast either to express xylA or to synthesize active enzyme (25). The first successful attempt was made with the xylA gene from the thermophilic bacterium Thermus thermophilus. XI from T. thermophilus could be expressed in S. cerevisiae in an active form, but the activity of this thermophilic enzyme, with a temperature optimum at 85°C, was very low at 30°C (33). In subsequent rounds of mutagenesis, the enzyme could be considerably improved but, however, still not enough for efficient xylose conversion in yeast (22).For the first time, Kuyper et al. (17) successfully expressed a xylA gene from the anaerobic fungus Piromyces sp. strain E2 in S. cerevisiae with high enzymatic activity. However, a drawback of this enzyme was its strong inhibition by xylitol. A laboratory haploid yeast strain which exhibited fast anaerobic growth on d-xylose and also high ethanol production rates was constructed (18, 20). Furthermore, mixed sugar utilization of d-glucose and d-xylose could recently be achieved by evolutionary engineering of recombinant yeast strains (19).In this paper, we report the cloning and successful expression of the first XI of prokaryotic origin with high activity in S. cerevisiae. As an advantage, the new enzyme is far less susceptible to inhibition by xylitol than is the enzyme from the Piromyces strain.     

Functional and Biochemical Analysis of the Chlamydia trachomatis Ligase MurE

Chlamydiae are unusual obligately intracellular bacteria that do not synthesize detectable peptidoglycan. However, they possess genes that appear to encode products with peptidoglycan biosynthetic activity. Bioinformatic analysis predicts that chlamydial MurE possesses UDP-MurNAc-l-Ala-d-Glu:meso-diaminopimelic acid (UDP-MurNAc-l-Ala-d-Glu:meso-A₂pm) ligase activity. Nevertheless, there are no experimental data to confirm this hypothesis. In this paper we demonstrate that the murE gene from Chlamydia trachomatis is capable of complementing a conditional Escherichia coli mutant impaired in UDP-MurNAc-l-Ala-d-Glu:meso-A₂pm ligase activity. Recombinant MurE from C. trachomatis (MurE_Ct) was overproduced in and purified from E. coli in order to investigate its kinetic parameters in vitro. By use of UDP-MurNAc-l-Ala-d-Glu as the nucleotide substrate, MurE_Ct demonstrated ATP-dependent meso-A₂pm ligase activity with pH and magnesium ion optima of 8.6 and 30 mM, respectively. Other amino acids (meso-lanthionine, the ll and dd isomers of A₂pm, d-lysine) were also recognized by MurE_Ct. However, the activities for these amino acid substrates were weaker than that for meso-A₂pm. The specificity of MurE_Ct for three possible C. trachomatis peptidoglycan nucleotide substrates was also determined in order to deduce which amino acid might be present at the first position of the UDP-MurNAc-pentapeptide. Relative k_cat/K_m ratios for UDP-MurNAc-l-Ala-d-Glu, UDP-MurNAc-l-Ser-d-Glu, and UDP-MurNAc-Gly-d-Glu were 100, 115, and 27, respectively. Our results are consistent with the synthesis in chlamydiae of a UDP-MurNAc-pentapeptide in which the third amino acid is meso-A₂pm. However, due to the lack of specificity of MurE_Ct for nucleotide substrates in vitro, it is not obvious which amino acid is present at the first position of the pentapeptide.Chlamydiae cause serious respiratory tract and genital infections in humans (9). They are obligately intracellular gram-negative bacteria, with a unique biphasic development cycle. Elementary bodies (EBs) are the infectious form of the organism and invade susceptible host cells. Once internalized, EBs differentiate into reticulate bodies (RBs), which have the capacity to divide (39, 40). The RBs are fragile and pleomorphic, whereas EBs are comparatively rigid and stable (19, 39). After repeated cycles of binary fission, the RBs differentiate into EBs, and the host cell lyses, releasing infectious EBs (1).In contrast to the vast majority of eubacteria, chlamydiae lack detectable amounts of peptidoglycan (PG), an essential polymer. PG is a giant macromolecule composed of alternating N-acetylglucosamine (GlcNAc) and N-acetylmuramic acid (MurNAc) residues cross-linked by short peptides. It determines the shape of bacteria, protects the cell from lysis due to internal osmotic pressure, and also plays a role in cell division. However, PG has not been detected in EBs (14, 20) or RBs (5).Although chlamydiae appear to lack PG, they contain penicillin-binding proteins and are sensitive to antibiotics that inhibit PG synthesis (5, 40). The Chlamydia trachomatis genome contains most of the genes coding for proteins involved in, or associated with, PG synthesis (54). Chlamydial MurA, MurC-Ddl, and the MurC domain of the latter fusion protein are active in vitro and complement Escherichia coli mutants deficient in the respective enzymes (23, 32, 33). Furthermore, proteomic analysis reveals that the murE gene product, which was assigned as UDP-MurNAc-l-Ala-d-Glu:meso-diaminopimelic acid ligase (UDP-MurNAc-l-Ala-d-Glu:meso-A₂pm ligase), is expressed in RBs (52).MurE ligases catalyze the addition of the third amino acid residue to the peptide chain of PG. This residue, generally a diamino acid, is usually meso-A₂pm for gram-negative bacteria and bacilli, and l-lysine for gram-positive bacteria, although other amino acids (for example, l-ornithine, meso-lanthionine, ll-A₂pm, l-diaminobutyric acid, or l-homoserine) occur in certain species (6, 50, 57). In many organisms, the third residue of the peptide chain participates in PG cross-linking; consequently, the MurE enzyme is highly specific for the relevant amino acid so as to avoid incorporation of incorrect amino acids into the macromolecule, which could result in deleterious morphological changes and cell lysis (35). Crystallization of MurE from E. coli (MurE_Ec) has permitted analysis of the structural basis for this high specificity (22). Sequence alignments of different MurE orthologues have also revealed the specific consensus sequences DNPR and D(D/N)P(N/A) located in the binding pockets for meso-A₂pm and l-Lys, respectively (11, 17). Chlamydia trachomatis MurE (MurE_Ct) possesses the DNPR motif, which suggests that it adds meso-A₂pm (17). However, there are no experimental data to confirm this prediction.In this paper we report for the first time the overproduction and purification of MurE_Ct, as well as a detailed investigation of its in vivo and in vitro biochemical properties. These studies contribute to our understanding of the nature and properties of the PG biosynthetic enzymes in chlamydiae and do indeed suggest that MurE_Ct has meso-A₂pm ligase activity.     

Substrate Specificity of a Mannose-6-Phosphate Isomerase from Bacillus subtilis and Its Application in the Production of l-Ribose

The uncharacterized gene previously proposed as a mannose-6-phosphate isomerase from Bacillus subtilis was cloned and expressed in Escherichia coli. The maximal activity of the recombinant enzyme was observed at pH 7.5 and 40°C in the presence of 0.5 mM Co²⁺. The isomerization activity was specific for aldose substrates possessing hydroxyl groups oriented in the same direction at the C-2 and C-3 positions, such as the d and l forms of ribose, lyxose, talose, mannose, and allose. The enzyme exhibited the highest activity for l-ribulose among all pentoses and hexoses. Thus, l-ribose, as a potential starting material for many l-nucleoside-based pharmaceutical compounds, was produced at 213 g/liter from 300-g/liter l-ribulose by mannose-6-phosphate isomerase at 40°C for 3 h, with a conversion yield of 71% and a volumetric productivity of 71 g liter⁻¹ h⁻¹.l-Ribose is a potential starting material for the synthesis of many l-nucleoside-based pharmaceutical compounds, and it is not abundant in nature (5, 19). l-Ribose has been produced mainly by chemical synthesis from l-arabinose, l-xylose, d-glucose, d-galactose, d-ribose, or d-mannono-1,4-lactone (2, 17, 23). Biological l-ribose manufacture has been investigated using ribitol or l-ribulose. Recently, l-ribose was produced from ribitol by a recombinant Escherichia coli containing an NAD-dependent mannitol-1-dehydrogenase (MDH) with a 55% conversion yield when 100 g/liter ribitol was used in a 72-h fermentation (18). However, the volumetric productivity of l-ribose in the fermentation is 28-fold lower than that of the chemical method synthesized from l-arabinose (8). l-Ribulose has been biochemically converted from l-ribose using an l-ribose isomerase from an Acinetobacter sp. (9), an l-arabinose isomerase mutant from Escherichia coli (4), a d-xylose isomerase mutant from Actinoplanes missouriensis (14), and a d-lyxose isomerase from Cohnella laeviribosi (3), indicating that l-ribose can be produced from l-ribulose by these enzymes. However, the enzymatic production of l-ribulose is slow, and the enzymatic production of l-ribose from l-ribulose has been not reported.Sugar phosphate isomerases, such as ribose-5-phosphate isomerase, glucose-6-phosphate isomerase, and galactose-6-phosphate isomerase, work as general aldose-ketose isomerases and are useful tools for producing rare sugars, because they convert the substrate sugar phosphates and the substrate sugars without phosphate to have a similar configuration (11, 12, 21, 22). l-Ribose isomerase from an Acinetobacter sp. (9) and d-lyxose isomerase from C. laeviribosi (3) had activity with l-ribose, d-lyxose, and d-mannose. Thus, we can apply mannose-6-phosphate (EC 5.3.1.8) isomerase to the production of l-ribose, because there are no sugar phosphate isomerases relating to l-ribose and d-lyxose. The production of the expensive sugar l-ribose (bulk price, $1,000/kg) from the rare sugar l-ribulose by mannose-6-phosphate isomerase may prove to be a valuable industrial process, because we have produced l-ribulose from the cheap sugar l-arabinose (bulk price, $50/kg) using the l-arabinose isomerase from Geobacillus thermodenitrificans (20) (Fig. ​(Fig.11).Open in a separate windowFIG. 1.Schematic representation for the production of l-ribulose from l-arabinose by G. thermodenitrificans l-arabinose isomerase and the production of l-ribose from l-ribulose by B. subtilis mannose-6-phosphate isomerase.In this study, the gene encoding mannose-6-phosphate isomerase from Bacillus subtilis was cloned and expressed in E. coli. The substrate specificity of the recombinant enzyme for various aldoses and ketoses was investigated, and l-ribulose exhibited the highest activity among all pentoses and hexoses. Therefore, mannose-6-phosphate isomerase was applied to the production of l-ribose from l-ribulose.     

Efficient Production of Optically Pure d-Lactic Acid from Raw Corn Starch by Using a Genetically Modified l-Lactate Dehydrogenase Gene-Deficient and α-Amylase-Secreting Lactobacillus plantarum Strain

In order to achieve direct and efficient fermentation of optically pure d-lactic acid from raw corn starch, we constructed l-lactate dehydrogenase gene (ldhL1)-deficient Lactobacillus plantarum and introduced a plasmid encoding Streptococcus bovis 148 α-amylase (AmyA). The resulting strain produced only d-lactic acid from glucose and successfully expressed amyA. With the aid of secreting AmyA, direct d-lactic acid fermentation from raw corn starch was accomplished. After 48 h of fermentation, 73.2 g/liter of lactic acid was produced with a high yield (0.85 g per g of consumed sugar) and an optical purity of 99.6%. Moreover, a strain replacing the ldhL1 gene with an amyA-secreting expression cassette was constructed. Using this strain, direct d-lactic acid fermentation from raw corn starch was accomplished in the absence of selective pressure by antibiotics. This is the first report of direct d-lactic acid fermentation from raw starch.Poly-lactic acid (PLA) is an important agro-based plastic that can be produced from inexpensive, renewable, and abundantly available biomass resources, including starchy materials. These resources have advantages over limited oil- and fossil-based sources, as they do not result in any net carbon dioxide release to the atmosphere (7). Recently, stereocomplex PLA, which is composed of both poly-l- and -d-lactic acid, has been attracting much attention due to its high thermostability. Stereocomplex-type polymers show a melting point (ca. 230°C) that is approximately 50°C higher than that of the respective single polymers (8). Therefore, d-lactic acid, in addition to l-lactic acid, which has been the focus of production to date, is of significant importance.Lactic acid bacteria (LAB) are promising microorganisms for the efficient production of lactic acid from various sugars, such as glucose, sucrose, and lactose. However, when starchy materials are used as a carbon source, they must be saccharified by physicochemical and enzymatic treatment because most LAB cannot utilize starchy materials directly (13). This makes the whole process less economically viable. Therefore, many researchers have examined the direct production of lactic acid from starchy materials by using wild amylolytic LAB (ALAB) (6, 24, 25) or genetically modified amylase-producing LAB (15, 16). Although d-lactic acid has been produced by fermentation from pretreated substrates such as rice starch (5) and by simultaneous saccharification and fermentation from cellulose (23), there have been no reports on the direct production of d-lactic acid from starchy materials. This is due to a lack of d-lactic acid-producing ALAB and difficulties in gene manipulation of d-lactic acid-producing LAB, such as Lactobacillus delbrueckii (22).We focused on Lactobacillus plantarum, which is an industrially important strain due to its environmental flexibility and its ability to assimilate a wide range of carbohydrates (9). In recent years, several gene manipulation methods for Lactobacillus plantarum have been established (18, 19). Moreover, the complete genome sequence has been decoded for L. plantarum NCIMB 8826 (9). Based on whole-genome analysis, L. plantarum possesses two types of lactate dehydrogenase (LDH), l-LDH and d-LDH, which convert pyruvate into l- and d-lactic acid, respectively. Ferain et al. (4) reported that chromosomal deletion in the ldhL1 gene of L. plantarum NCIMB 8826 provoked an absence of l-LDH activity and produced d-lactic acid from glucose.In the present study, to produce d-lactic acid directly from starch, we constructed an l-LDH-deficient, α-amylase-secreting L. plantarum strain. The engineered strain expressed α-amylase from Streptococcus bovis 148 (AmyA) (20) and efficiently degraded raw starch with the aid of a C-terminal starch-binding domain (11). Using this strain, we achieved the direct and efficient fermentation of optically pure d-lactic acid from raw corn starch.     

Zif,the Zoocin A Immunity Factor,Is a FemABX-Like Immunity Protein with a Novel Mode of Action

Producer cell immunity to the streptococcolytic enzyme zoocin A, which is a d-alanyl-l-alanine endopeptidase, is due to Zif, the zoocin A immunity factor. Zif has high degrees of similarity to MurM and MurN (members of the FemABX family of proteins), which are responsible for the addition of amino acids to cross bridges during peptidoglycan synthesis in streptococci. In this study, purified peptidoglycans from strains with and without zif were compared to determine how Zif modifies the peptidoglycan layer to cause resistance to zoocin A. The peptidoglycan from each strain was hydrolyzed using the streptococcolytic phage lysin B30, and the resulting muropeptides were separated by reverse-phase high-pressure liquid chromatography, labeled with 4-sulfophenyl isothiocyanate, and analyzed by tandem mass spectrometry in the negative-ion mode. It was determined that Zif alters the peptidoglycan by increasing the proportion of cross bridges containing three l-alanines instead of two. This modification decreased binding of the recombinant target recognition domain of zoocin A to peptidoglycan. Zif-modified peptidoglycan also was less susceptible to hydrolysis by the recombinant catalytic domain of zoocin A. Thus, Zif is a novel FemABX-like immunity factor because it provides resistance to a bacteriolytic endopeptidase by lengthening the peptidoglycan cross bridge rather than by causing an amino acid substitution.During streptococcal peptidoglycan synthesis, monomer subunits are generated inside the cell, with nonribosomal peptidyl transferases responsible for the addition of amino acids onto the epsilon amino group of lysine in the subunits. These nonribosomal peptidyl transferases are part of the FemABX family of proteins, some of which have been implicated in penicillin resistance (5, 26). In Streptococcus pneumoniae peptidoglycan synthesis, MurM attaches either an l-alanine or an l-serine to the epsilon amino group of lysine, and MurN then adds an l-alanine (11, 26).Zoocin A is a d-alanyl-l-alanine endopeptidase produced by Streptococcus equi subsp. zooepidemicus 4881 that hydrolyzes peptidoglycan cross bridges of susceptible streptococci (12). Zoocin A has two functional domains (18). The N-terminal catalytic domain (CAT) has high degrees of similarity to several other bacteriolytic endopeptidases, including the staphylolytic enzyme lysostaphin. The C-terminal target recognition domain (TRD), which facilitates binding of the enzyme to peptidoglycan (1), has very little similarity to any characterized conserved domain.Producer cell immunity to zoocin A is due to zif (zoocin A immunity factor), which is adjacent to zooA on the chromosome and is transcribed divergently (4). Zif has high degrees of similarity to MurM and MurN and also to the lysostaphin resistance protein and other FemABX-like immunity proteins (23). Previously characterized FemABX-like immunity proteins provide resistance to peptidoglycan cross-bridge hydrolases by inserting an amino acid different from those specified by the normal FemABX-like proteins (6, 9, 15, 25), whereas Zif does not (4). It has been shown previously that Zif-specified resistance to zoocin A is an intrinsic characteristic of the peptidoglycan layer (12). Therefore, Zif must modify the peptidoglycan layer in a novel way that provides resistance to zoocin A. In the present study, Zif was shown to insert an additional l-alanine into the peptidoglycan cross bridges, which inhibited both binding of the zoocin A TRD and the ability of the zoocin A CAT to hydrolyze the cross bridge.     

A Role for the Class A Penicillin-Binding Protein PonA2 in the Survival of Mycobacterium smegmatis under Conditions of Nonreplication

Class A penicillin-binding proteins (PBPs) are large, bifunctional proteins that are responsible for glycan chain assembly and peptide cross-linking of bacterial peptidoglycan. Bacteria in the genus Mycobacterium have been reported to have only two class A PBPs, PonA1 and PonA2, that are encoded in their genomes. We report here that the genomes of Mycobacterium smegmatis and other soil mycobacteria contain an additional gene encoding a third class A penicillin-binding protein, PonA3, which is a paralog of PonA2. Both the PonA2 and PonA3 proteins contain a penicillin-binding protein and serine/threonine protein kinase-associated (PASTA) domain that we propose may be involved in sensing the cell cycle and a C-terminal proline-rich region (PRR) that may have a role in protein-protein or protein-carbohydrate interactions. We show here that an M. smegmatis ΔponA2 mutant has an unusual antibiotic susceptibility profile, exhibits a spherical morphology and an altered cell surface in stationary phase, and is defective for stationary-phase survival and recovery from anaerobic culture. In contrast, a ΔponA3 mutant has no discernible phenotype under laboratory conditions. We demonstrate that PonA2 and PonA3 can bind penicillin and that PonA3 can partially substitute for PonA2 when ponA3 is expressed from a constitutive promoter on a multicopy plasmid. Our studies suggest that PonA2 is involved in adaptation to periods of nonreplication in response to starvation or anaerobiosis and that PonA3 may have a similar role. However, the regulation of PonA3 is likely different, suggesting that its importance could be related to stresses encountered in the environmental niches occupied by M. smegmatis and other soil-dwelling mycobacteria.The cell envelope of mycobacteria is a complex carbohydrate- and lipid-rich entity and is a major factor contributing to the success of these organisms as saprophytic and pathogenic bacteria (7, 8, 29, 35). The innermost layer of the cell envelope is a peptidoglycan (PG) composed of N-acylmuramic acid and N-acetylglucosamine with l-alanyl (or glycyl in the case of Mycobacterium leprae)-d-isoglutaminyl-meso-diaminopimelyl-d-alanyl-d-alanine pentapeptides attached to the muramic acid residues (13, 16, 54). While some of the muramyl residues are N acetylated, as they are in most other bacteria, a majority of the muramyl residues are N glycolylated (2, 37, 48, 49), a modification that confers lysozyme resistance (53) and also influences the innate immune response to mycobacterial cell walls (10). The pentapeptide chains of the mycobacterial PG can be modified by amidation, glycylation, or methylation, but the functional significance of these modifications is unknown (28, 31, 32, 38, 54).Approximately 80% of the pentapeptides in mycobacterial PG are cross-linked, and a majority of the links are between the carboxyl group of a penultimate d-Ala residue in a pentapeptide precursor and the amino group of the side chain d center of a meso-diaminopimelic acid (DAP) residue from an adjacent peptide (referred to as a 4-3 cross-link), while approximately one-third of the links are between the carboxyl group of the l center of a DAP residue of one peptide and the amino group of the side chain d center of the DAP residue in an adjacent peptide (referred to as a 3-3 cross-link) (17, 65). The 4-3 linkage is considered the “standard” linkage and is catalyzed by classical, penicillin-sensitive dd-transpeptidases, while the novel 3-3 linkage is thought to be catalyzed by the concerted action of dd-carboxypeptidases and novel ld-transpeptidases (31, 34, 39-41). The reasons why bacteria produce both 4-3 and 3-3 linkages are unknown. Some workers have suggested that the 3-3 linkages might reinforce the wall during times of stress and under nonreplicating conditions or stabilize complex cell envelopes (17, 50, 51, 55). In this regard, the high percentage of 3-3 linkages found in the PG of mycobacteria and their predominance in stationary-phase M. tuberculosis cells (31) suggest that these linkages may have an important role in maintaining cell envelope integrity during periods of growth and under nonreplicating conditions.The enzymes involved in peptidoglycan assembly, the penicillin-binding proteins (PBPs), have a triad sequence motif that forms the transpeptidation active site ([SxxK]——[S/YxN/C]——[K/H][T/S]G), which is the target of the β-lactam class of antibiotics (for a review, see reference 18). The PBPs have been grouped into several classes based on this motif, surrounding sequences, and other structural features (18). Of interest here are the class A PBPs, which are high-molecular-weight (HMW) proteins with both a transglycosylase domain (also called a non-penicillin-binding module [n-PB]) and a transpeptidase domain (also called a penicillin-binding module [PB]) (18). These proteins are tethered to the cytoplasmic membrane by a transmembrane helix with the catalytic domains facing the outside of the cell. Mycobacteria have been reported to have only two genes that encode class A PBPs, ponA1 and ponA2, which are annotated Rv0051 and Rv3682 in the sequence genome of M. tuberculosis H37Rv (9, 17). Previous studies that analyzed collections of transposon mutants to obtain clones with various phenotypes identified strains with insertions in these two genes. The phenotypes of these mutants have clearly shown that these PBPs play a complex role in mycobacterial physiology. One group of workers found a ponA1 mutant of M. smegmatis in a search for mutants with an altered dye-binding phenotype (an indicator of changes in the cell envelope) and showed that this slowly growing mutant was hypersusceptible to β-lactam antibiotics and had altered permeability (6). A ponA2 mutant of M. smegmatis was discovered in a screen for mutants defective for survival during long-term culture (25), while other workers isolated an M. tuberculosis ponA2 mutant in a screen for mutants sensitive to low pH (61). The same group of workers also showed that the M. tuberculosis mutant was more sensitive to heat, H₂O₂, and NO and was attenuated for persistence in the mouse model of inhalation tuberculosis (62). We previously identified an M. tuberculosis ponA2 mutant in a screen for mutants hypersusceptible to β-lactam antibiotics (14). All of these studies identified transposon mutants in searches for mutants with specific phenotypes, but there have been no direct genetic studies that have specifically examined the function of these PBPs in peptidoglycan metabolism.In this study we demonstrated that M. smegmatis has three class A PBPs. We show here that a newly recognized protein, which we designated PonA3, is a paralog of the PonA2 protein and is found only in certain environmental species of mycobacteria. We analyzed the phenotypes of M. smegmatis mutants with in-frame deletions of ponA2 and ponA3 singly and in combination to increase our understanding of the role that these PBPs play in mycobacterial peptidoglycan biology.     

The d-2-Hydroxyacid Dehydrogenase Incorrectly Annotated PanE Is the Sole Reduction System for Branched-Chain 2-Keto Acids in Lactococcus lactis

Hydroxyacid dehydrogenases of lactic acid bacteria, which catalyze the stereospecific reduction of branched-chain 2-keto acids to 2-hydroxyacids, are of interest in a variety of fields, including cheese flavor formation via amino acid catabolism. In this study, we used both targeted and random mutagenesis to identify the genes responsible for the reduction of 2-keto acids derived from amino acids in Lactococcus lactis. The gene panE, whose inactivation suppressed hydroxyisocaproate dehydrogenase activity, was cloned and overexpressed in Escherichia coli, and the recombinant His-tagged fusion protein was purified and characterized. The gene annotated panE was the sole gene responsible for the reduction of the 2-keto acids derived from leucine, isoleucine, and valine, while ldh, encoding l-lactate dehydrogenase, was responsible for the reduction of the 2-keto acids derived from phenylalanine and methionine. The kinetic parameters of the His-tagged PanE showed the highest catalytic efficiencies with 2-ketoisocaproate, 2-ketomethylvalerate, 2-ketoisovalerate, and benzoylformate (V_max/K_m ratios of 6,640, 4,180, 3,300, and 2,050 U/mg/mM, respectively), with NADH as the exclusive coenzyme. For the reverse reaction, the enzyme accepted d-2-hydroxyacids but not l-2-hydroxyacids. Although PanE showed the highest degrees of identity to putative NADP-dependent 2-ketopantoate reductases (KPRs), it did not exhibit KPR activity. Sequence homology analysis revealed that, together with the d-mandelate dehydrogenase of Enterococcus faecium and probably other putative KPRs, PanE belongs to a new family of d-2-hydroxyacid dehydrogenases which is unrelated to the well-described d-2-hydroxyisocaproate dehydrogenase family. Its probable physiological role is to regenerate the NAD⁺ necessary to catabolize branched-chain amino acids, leading to the production of ATP and aroma compounds.Hydroxyacid dehydrogenases catalyze the stereospecific and reversible reduction of 2-keto acids to 2-hydroxyacids. These NAD(H)-dependent oxidoreductases are of interest in a variety of fields. Firstly, they are valuable catalysts for the production of the stereospecific isomers of 2-hydroxyacids that are used in the production of semisynthetic antibiotics or pharmaceuticals (32). Secondly, in lactic acid bacteria, hydroxyacid dehydrogenases are believed to be negatively involved in flavor production, since they compete with other enzymes generating flavor compounds from 2-keto acids derived from amino acids (66), while 2-hydroxyacids are not aroma compounds or precursors of flavor compounds. Indeed, overexpression of the d-2-hydroxyisocaproate dehydrogenase (d-2-HicDH) of Lactobacillus casei has been shown previously to decrease the production of aroma compounds and to delay flavor formation in low-fat cheddar cheese (12). Additionally, hydroxyacid dehydrogenases are involved in the biopreservation properties of lactic acid bacteria, because certain 2-hydroxyacids exhibit antifungal and antilisterial activities (15, 37, 64). Several hydroxyacid dehydrogenases in lactic acid bacteria have been characterized previously (21, 27). Lactate dehydrogenases (LDH), responsible for the specific reduction of pyruvate to lactic acid, have been studied extensively (2, 7, 13, 16, 21, 40, 49, 54). HicDHs and mandelate dehydrogenases (manDHs), active toward a broad range of 2-keto acids, including straight-chain aliphatic 2-keto acids, branched-chain 2-keto acids, and 2-keto acids with aromatic side chains, in several lactic acid bacteria have also been characterized previously (5, 6, 27, 28, 29, 38, 39, 50). Although manDHs and HicDHs of lactic acid bacteria prefer 2-ketoisocaproate (KIC) among 2-keto acid substrates, they differ in their activities toward C-3-branched substrates. In particular, manDHs exhibit high levels of activity toward 2-ketoisovalerate (KIV) and benzoylformate, unlike HicDHs. LDHs, HicDHs, and manDHs are divided into two groups, the l and d groups, depending on the stereoisomer produced. d-LDHs and d-HicDHs are members of the same family of 2-hydroxyacid dehydrogenases, which is distinct from the l-LDH family (13, 54). In general, each lactic acid bacterium contains several hydroxyacid dehydrogenases.In Lactococcus lactis, which is widely employed as a starter in cheese production, the main LDH is an l-LDH activated by fructose 1,6-bisphosphate (FBP) (23). In addition to the ldh gene that encodes the l-LDH (40), L. lactis contains other genes showing significant levels of similarity to the hydroxyacid dehydrogenases of other lactic acid bacteria (9), but their functions remain unknown, except for that of the recently identified ldhB gene. ldhB is a silent gene that can be activated in ldh-deficient strains via an IS981 element-specific insertion to produce a functional LDH (10). However L. lactis produces 2-hydroxyacids from the 2-keto acids derived from amino acids not only in vitro, using resting cells, but also in cheese (65, 66). In L. lactis, the catabolism of the aromatic and branched-chain amino acids that are precursors of aroma compounds is initiated by aminotransferases, producing 2-keto acids. These 2-keto acids can be further catabolized into carboxylic acids via a 2-keto acid dehydrogenase, a transacetylase, and a kinase or reduced to 2-hydroxyacids by a hydroxyacid dehydrogenase (66) (Fig. ​(Fig.1).1). In some strains, branched-chain 2-keto acids can also be decarboxylated to aldehydes that are potent aroma compounds. The aim of the present study was to identify and characterize the enzyme(s) involved in the production of 2-hydroxyacids from the 2-keto acids derived from phenylalanine and branched-chain amino acids and to evaluate its impact on amino acid catabolism.Open in a separate windowFIG. 1.Leucine catabolism pathway in L. lactis TIL46. AT, aminotransferase; HADH, hydroxyacid dehydrogenase; TA, transacetylase; KDH, keto acid dehydrogenase; HIC, 2-hydroxyisocaproate.     

Escherichia coli Strains Engineered for Homofermentative Production of d-Lactic Acid from Glycerol

Given its availability and low price, glycerol has become an ideal feedstock for the production of fuels and chemicals. We recently reported the pathways mediating the metabolism of glycerol in Escherichia coli under anaerobic and microaerobic conditions. In this work, we engineer E. coli for the efficient conversion of glycerol to d-lactic acid (d-lactate), a negligible product of glycerol metabolism in wild-type strains. A homofermentative route for d-lactate production was engineered by overexpressing pathways involved in the conversion of glycerol to this product and blocking those leading to the synthesis of competing by-products. The former included the overexpression of the enzymes involved in the conversion of glycerol to glycolytic intermediates (GlpK-GlpD and GldA-DHAK pathways) and the synthesis of d-lactate from pyruvate (d-lactate dehydrogenase). On the other hand, the synthesis of succinate, acetate, and ethanol was minimized through two strategies: (i) inactivation of pyruvate-formate lyase (ΔpflB) and fumarate reductase (ΔfrdA) (strain LA01) and (ii) inactivation of fumarate reductase (ΔfrdA), phosphate acetyltransferase (Δpta), and alcohol/acetaldehyde dehydrogenase (ΔadhE) (strain LA02). A mutation that blocked the aerobic d-lactate dehydrogenase (Δdld) also was introduced in both LA01 and LA02 to prevent the utilization of d-lactate. The most efficient strain (LA02Δdld, with GlpK-GlpD overexpressed) produced 32 g/liter of d-lactate from 40 g/liter of glycerol at a yield of 85% of the theoretical maximum and with a chiral purity higher than 99.9%. This strain exhibited maximum volumetric and specific productivities for d-lactate production of 1.5 g/liter/h and 1.25 g/g cell mass/h, respectively. The engineered homolactic route generates 1 to 2 mol of ATP per mol of d-lactate and is redox balanced, thus representing a viable metabolic pathway.Lactic acid (lactate) and its derivatives have many applications in the food, pharmaceutical, and polymer industries (13, 30). An example is polylactic acid, a renewable, biodegradable, and environmentally friendly polymer produced from d- and l-lactate (19). In this context, biological processes have the advantage of being able to produce chirally pure lactate from inexpensive media containing only the carbon source and mineral salts (43). While lactic acid bacteria traditionally have been used in the production of d-lactate from carbohydrate-rich feedstocks, several laboratories recently have reported alternative biocatalysts (13, 30), many of which are engineered Escherichia coli strains that produce d- or l-lactate (4, 8, 50, 51, 52).Unlike the aforementioned reports, which have dealt with the use of carbohydrates, our work focuses on the use of glycerol as a carbon source for the production of d-lactate. Glycerol has become an inexpensive and abundant substrate due to its generation in large amounts as a by-product of biodiesel and bioethanol production (18, 32, 47). The conversion of glycerol to higher-value products has been proposed as a path to economic viability for the biofuels industry (47). One such product is lactate, whose production could be readily integrated into existing biodiesel and bioethanol facilities, thus establishing true biorefineries.Although many microorganisms are able to metabolize glycerol (25), the use of industrial microbes such as E. coli could greatly accelerate the development of platforms to produce fuels and chemicals from this carbon source. We recently reported on the ability of E. coli to metabolize glycerol under either anaerobic or microaerobic conditions and identified the environmental and metabolic determinants of these processes (9, 11, 28). In one of the studies, the pathways involved in the microaerobic utilization of glycerol were elucidated, and they are shown in Fig. ​Fig.11 (9). A common characteristic of glycerol metabolism under either anaerobic or microaerobic conditions is the generation of ethanol as the primary product and the negligible production of lactate (6, 9, 11, 28). In the work reported here, the knowledge base created by the aforementioned studies was used to engineer E. coli for the efficient conversion of glycerol to d-lactate in minimal medium. The engineered strains hold great promise as potential biocatalysts for the conversion of low-value glycerol streams to a higher-value product like d-lactate.Open in a separate windowFIG. 1.Pathways involved in the microaerobic utilization of glycerol in E. coli (9). Genetic modifications supporting the metabolic engineering strategies employed in this work are illustrated by thicker lines (overexpression of gldA-dhaKLM, glpK-glpD, and ldhA) or cross bars (disruption of pflB, pta, adhE, frdA, and dld). Broken lines illustrate multiple steps. Relevant reactions are represented by the names of the gene(s) coding for the enzymes: aceEF-lpdA, pyruvate dehydrogenase complex; adhE, acetaldehyde/alcohol dehydrogenase; ackA, acetate kinase; dhaKLM, dihydroxyacetone kinase; dld, respiratory d-lactate dehydrogenase; fdhF, formate dehydrogenase, part of the formate hydrogenlyase complex; frdABCD, fumarate reductase; gldA, glycerol dehydrogenase; glpD, aerobic glycerol-3-phosphate dehydrogenase; glpK, glycerol kinase; hycB-I, hydrogenase 3, part of the formate hydrogenlyase complex; ldhA, fermentative d-lactate dehydrogenase; pflB, pyruvate formate-lyase; pta, phosphate acetyltransferase; pykF, pyruvate kinase. Abbreviations: DHA, dihydroxyacetone; DHAP, DHA phosphate; G-3-P, glycerol-3-phosphate; PEP, phosphoenolpyruvate; PYR, pyruvate; P/O, amount of ATP produced in the oxidative phosphorylation per pair of electrons transferred through the electron transport system; QH₂, reduced quinones.