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Construction of an in vitro bypassed pyruvate decarboxylation pathway using thermostable enzyme modules and its application to N-acetylglutamate production
© Krutsakorn et al.; licensee BioMed Central Ltd. 2013
Received: 10 July 2013
Accepted: 17 September 2013
Published: 7 October 2013
Metabolic engineering has emerged as a practical alternative to conventional chemical conversion particularly in biocommodity production processes. However, this approach is often hampered by as yet unidentified inherent mechanisms of natural metabolism. One of the possible solutions for the elimination of the negative effects of natural regulatory mechanisms on artificially engineered metabolic pathway is to construct an in vitro pathway using a limited number of enzymes. Employment of thermostable enzymes as biocatalytic modules for pathway construction enables the one-step preparation of catalytic units with excellent selectivity and operational stability. Acetyl-CoA is a central precursor involved in the biosynthesis of various metabolites. In this study, an in vitro pathway to convert pyruvate to acetyl-CoA was constructed and applied to N-acetylglutamate production.
A bypassed pyruvate decarboxylation pathway, through which pyruvate can be converted to acetyl-CoA, was constructed by using a coupled enzyme system consisting of pyruvate decarboxylase from Acetobacter pasteurianus and the CoA-acylating aldehyde dehydrogenase from Thermus thermophilus. To demonstrate the applicability of the bypassed pathway for chemical production, a cofactor-balanced and CoA-recycling synthetic pathway for N-acetylglutamate production was designed by coupling the bypassed pathway with the glutamate dehydrogenase from T. thermophilus and N-acetylglutamate synthase from Thermotoga maritima. N-Acetylglutamate could be produced from an equimolar mixture of pyruvate and α-ketoglutarate with a molar yield of 55% through the synthetic pathway consisting of a mixture of four recombinant E. coli strains having either one of the thermostable enzymes. The overall recycling number of CoA was calculated to be 27.
Assembly of thermostable enzymes enables the flexible design and construction of an in vitro metabolic pathway specialized for chemical manufacture. We herein report the in vitro construction of a bypassed pathway capable of an almost stoichiometric conversion of pyruvate to acetyl-CoA. This pathway is potentially applicable not only to N-acetylglutamate production but also to the production of a wide range of acetyl-CoA-derived metabolites.
The integration of diverse biocatalytic modules to expand the versatility of fermentation-based industries has been widely employed for the production of biofuel, pharmaceuticals, and other useful chemicals . However, these “metabolic engineering” approaches often suffer from flux imbalances because the naturally occurring translational and transcriptional regulation mechanisms fail to appropriately function in artificially engineered cells [2, 3]. One possible approach to overcome this limitation is to construct an in vitro metabolic pathway in which only a limited number of enzymes involved in the pathway-of-interest are used as catalytic modules. In vitro systems offer a number of potential advantages over the conventional fermentation-based production processes such as predictable production yield, operational flexibility, and scalability of the reaction [4, 5]. However, the enzyme isolation generally requires laborious, costly, and time-consuming procedures, and thus little attention has been paid to the practical application of in vitro metabolic engineering approaches. Recently, we reported a simple approach to construct an in vitro artificial metabolic pathway using thermostable biocatalytic modules [6, 7]. The basic procedure consists of the following four steps: 1) selection of suitable thermostable enzymes; 2) expression of the enzymes in mesophilic hosts (e.g. Escherichia coli); 3) preheating the cell suspension at high temperature; and 4) assembly of the catalytic modules at an experimentally optimized ratio to achieve stoichiometric production. The denaturation of indigenous enzymes at high temperatures results in a one-step preparation of highly selective and thermostable biocatalytic modules. The membrane structure of E. coli cells is partially or entirely disrupted at high temperatures, and consequently, better accessibility between the enzymes and substrates can be achieved .
Acetyl-CoA is a key precursor for the biosynthesis of a wide variety of industrially useful metabolites. In the mevalonate pathway, acetyl-CoA is carboxylated and conjugated to form various isoprenoid and terpenoid compounds that can be used as flavors, fragrances, and anticancer drugs . Condensation of two acetyl-CoA units leads to the formation of acetoacetyl-CoA, which serves as a C4 backbone for polyhydroxyalkanoates and butanol biosynthesis [10–12]. Acetyl-CoA production tends to be a rate-limiting step of artificially engineered pathways to produce target compounds because acetyl-CoA serves a central intermediate of a wide range of naturally occurring metabolic pathways and thus is subjected to depletion when it is routed into co-existing pathways .
In natural metabolism, acetyl-CoA is mainly generated from pyruvate via three routes. Most aerobic microorganisms use the large protein complex of pyruvate dehydrogenase (PDH) to produce acetyl-CoA . Alternatively, pyruvate-ferredoxin oxidoreductase (PFOR) plays an important role in the oxidative decarboxylation of pyruvate in some anaerobic bacteria and hyperthermophilic achaea [14–16]. Pyruvate formate-lyase (PFL) is another key enzyme involved in the carbohydrate metabolism of anaerobic microorganisms [17, 18]. However, none are readily available in in vitro production systems. PDH is one of the largest protein complexes and consists of multiple copies of three or four subunits . For example, the PDH of E. coli consists of a central cubic core composed of 24 molecules of dihydrolipoamide acetyltransferase (E2), onto which up to 24 copies of pyruvate dehydrogenase (E1) and 12 copies of dihydrolipoamide dehydrogenase (E3) are assembled . To our knowledge, there have been no reports on the functional expression of the complete form of PDH in heterologous hosts probably owing to their highly complex nature. PFORs require specific redox partner proteins, namely ferredoxin and ferredoxin reductase, to exert their catalytic abilities. In in vitro systems, the enzymes and the partner proteins freely diffuse in the reaction mixture, resulting in less frequent interactions between them. In addition, the poor oxygen tolerance of PFOR results in operational limitations for its in vitro use. Similarly, PFL is highly sensitive to oxygen stress .
To overcome these limitations, in this study, we constructed an in vitro bypassed pathway capable of converting pyruvate to acetyl-CoA using an enzyme couple of thermostable pyruvate decarboxylase and CoA-acylating aldehyde dehydrogenase. Owing to the absence of a co-existing pathway, an almost stoichiometric production of acetyl-CoA could be achieved through the in vitro pathway. Furthermore, the performance of this pathway was assessed by integration to an artificial pathway for the production of N-acetylglutamate (NAG), which can be used as an ingredient in the cosmetic industry as an anti-odor compound and skin moisturizer [21, 22].
Construction of the bypassed pyruvate decarboxylation pathway
Design of the NAG production pathway
The bypassed pyruvate decarboxylation pathway was integrated into a newly designed synthetic pathway for NAG production to evaluate its applicability to chemical production (Figure 1). In the designed pathway, NAG is produced through transacetylation from acetyl-CoA to glutamate catalyzed by N-acetylglutamate synthase (NAGS). CoA molecules released by the acetylation of glutamate can be recycled for the synthesis of acetyl-CoA in the bypassed pyruvate decarboxylation pathway. The thermostable NAGS, which is involved in the arginine biosynthesis pathway of Thermotoga maritima, is well characterized , therefore, the NAGS from T. maritima was selected as the module to construct the pathway and designated Tm NAGS.
Unlike natural metabolic pathways, which are equipped with a complete enzyme apparatus for the de novo synthesis of cofactors, indigenous regeneration of redox cofactors is necessary to facilitate the economic feasibility of in vitro production processes [4, 6]. To achieve balanced NAD+/NADH consumption and regeneration, the glutamate dehydrogenase of T. thermophilus (Tt GDH) was integrated into the NAG production pathway. The enzyme can use α-ketoglutarate, an inexpensive material, as a cosubstrate for the re-oxidation of NADH and to produce glutamate, which can be directly used as the substrate for Tm NAGS. Thermal stability analysis of Tm NAGS and Tt GDH demonstrated that they can be used at 50°C for at least 4 h without a significant loss of their activities (Figure 3c and d).
pH profile of enzyme activity
Specific activity (U mg-1total protein)a
Total protein concentration (mg ml-1)b
The flux through the bypassed pyruvate decarboxylation pathway could be determined by incubating an enzyme mixture of Ap PDC and Tt ADDH with 0.2 mM each of pyruvate, CoA, and NAD+, and monitoring the concomitant production rate of NADH at 340 nm. When equal units (0.04 U ml-1) of the enzymes were used, the initial NADH production rate was lower than the expected value of 0.04 μmol min-1 ml-1. In the constructed pathway, the actual concentration of acetaldehyde, which is generated by Ap PDC and then used as the substrate for Tt ADDH, was kept at a lower level than those used in the standard assay, and thus Tt ADDH failed to catalyze the reaction at the expected rate. We increased the Tt ADDH concentration in the reaction mixture in a stepwise manner, and found that the expected NADH production rate of 0.04 μmol min-1 ml-1 could be achieved by using 0.09 U ml-1 of Tt ADDH.
Acetyl-CoA is an essential intermediate for the production of a variety of metabolites; therefore intensive studies have been conducted to increase the intracellular level of acetyl-CoA in artificially engineered microorganisms. Atsumi et al. reported that the increases in the intracellular pool size of acetyl-CoA, which can be achieved by deleting the gene sets of ldh A, adh E, and frd BC of E. coli, leads to a two-fold increase in 1-butanol production by the engineered cells . Shiba et al. have engineered a pyruvate dehydrogenase bypass pathway in Saccharomyces cerevisiae to increase intracellular acetyl-CoA and to improve the production level of amorphadiene, a sesquiterpene precursor for the biosynthesis of the anti-malarial drug artemisinin . Furthermore, Chen et al. have developed an engineered yeast platform for improved acetyl-CoA production and demonstrated its application for α-santalene production .
Previously, we reported a simple approach to construct an in vitro artificial metabolic pathway using thermostable biocatalytic modules [6, 7]. This approach is, in principle, applicable to any thermostable enzyme as long as it can be functionally expressed in a mesophilic host, and thus is potentially applicable to the biocatalytic manufacture of a wide range of useful compounds. By using this approach, we have constructed a non-ATP-forming chimeric glycolytic pathway and applied it to the direct conversion of glucose to lactate  and malate . The construction of an in vitro pyruvate decarboxylation pathway to form acetyl-CoA was one of the key challenges remaining to expand the applicability of this system.
Another possible approach to mitigate the effect of product inhibition is the employment of a continuous product recovery system. Recently, we reported that the pretreatment of recombinant E. coli cells having a thermophilic fumarase with a low concentration glutaraldehyde (~0.15 vol%) can consolidate the membrane structure of the cells and prevent the heat-induced leakage of the enzyme . The glutaraldehyde-treated cells can be used at 70°C for more than 10 h in a continuous reactor equipped with a cell-separation membrane filter. In the conversional fermentation-based processes, the culture broth contains a number of impurities including residual sugars, nutrients, and other organic materials, apart from cell mass. By contrast, in in vitro production systems, the reaction can be performed in a simple buffer solution, and thus the product separation step can be considerably simplified. Development and integration of an on-line product separation system would be a key challenge to eliminate product inhibition and to achieve the long-term operation of the in vitro production system.
We designed and constructed an in vitro pyruvate decarboxylation pathway capable of converting pyruvate to acetyl-CoA using an enzyme couple of Ap PDC and Tt ADDH. To demonstrate the applicability of the bypassed pathway to chemical production, the pathway was coupled with Tt GDH and Tm NAGS to construct a cofactor-balanced and CoA-recycling pathway for NAG production. The highly traceable nature of the in vitro pathway enabled us to readily identify the rate-limiting enzyme. Consequently, 5.4 mM of NAG could be produced through the pathway with a molar yield of 55% and a CoA-recycling number of 27. The bypassed pathway constructed in this study would be widely applicable to the production of other acetyl-CoA-derived chemicals including n-butanol, polyhydroxyalkanoates, and isoprenoids.
Materials and methods
Bacterial strain and plasmid
The expression plasmids encoding Tt ADDH (Genebank accession number, YP_145486.1) and Tt GDH (YP_144842.1) were obtained from the RIKEN T. thermophilus HB8 expression plasmid set . The gene coding for Ap PDC (YP_005499956.1) was amplified by PCR from the chromosomal DNA of A. pasteurianus using the following primers: 5′-AACATATG ACCTATACTGTTGGCAT-3′ (the Nde I restriction site is shown in italics) and 5′-TTCTCGAG TCAGGCCAGAGTCGTCTTGC-3′ (the Xho I restriction site is shown in italics). The amplified DNA was digested with Nde I and Xho I and inserted into the corresponding restriction sites of pET21a (Novagen, Madison, WI, USA). Similarly, the gene encoding Tm NAGS (NP_229580.1) was amplified from the chromosomal DNA of T. maritima using a primer pair of 5′-AACATATG TTCACTCCCAGGGGTTT-3′ (the Nde I restriction site is shown italics) and 5′-TTGAATTC TCATGTTCTGTACCTCCCGT-3′ (the EcoR I restriction site is shown italics), and then introduced into pET21a. The expression vectors were separately introduced in E. coli Rosetta2 (DE3) (Novagen). Recombinant cells were cultivated aerobically at 37°C in Luria-Bertani (LB) media supplemented with 50 μg ml-1 ampicillin and 30 μg ml-1 chloramphenicol. Isopropyl-β-D-1-thiogalactopyranoside was added to the culture at a final concentration of 0.2 mM in the late-log phase. The cells were harvested by centrifugation and washed once with 50 mM 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid (HEPES)-NaOH buffer (pH 7.0).
Recombinant cells were suspended in an appropriate volume of 50 mM HEPES-NaOH (pH 7.0) and disrupted using an UD-201 ultrasonicator (Kubota, Osaka, Japan) at 80 W for 2 min. Cell debris were removed by centrifugation at 12,000 × g at 4°C for 10 min. Protein concentration in the crude lysates was measured using a Bio-Rad assay system (Bio-Rad, Hercules, CA, USA) with bovine serum albumin as the standard. The resulting crude lysates were incubated at 70°C for 30 min to inactivate the indigenous enzymes. An incubation temperature of 60°C was employed for the heat treatment of the lysate with Ap PDC. The heat-treated lysates were centrifuged again to remove the denatured protein and then the supernatants were used as enzyme solutions.
All assays were performed at 50°C in 50 mM HEPES-NaOH (pH 7.0) supplemented with 100 mM KCl and 10 mM MgCl2 unless otherwise stated. Activities of Ap PDC, Tt ADDH, and Tt GDH were spectrophotometrically determined by monitoring the consumption or formation rate of NADH at 340 nm. Tt ADDH was assayed in the buffer containing 0.2 mM NAD+, 0.2 mM CoA, and an appropriate amount of the enzyme. After the preincubation at 50°C for 2 min, the reaction was started by the addition of acetaldehyde at a final concentration of 0.2 mM, and the reduction of NAD+ was monitored at 340 nm. The activity of Tt GDH was assessed in a similar manner by monitoring the oxidation of NADH upon the reduction of 0.2 mM α-ketoglutarate. Ap PDC activity was assessed by coupling with Tt ADDH. After the preincubation of the enzymes at 50°C in the buffer containing 0.2 mM NAD+, 0.2 mM CoA, and an excess amount of Tt ADDH, the reaction was initiated by the addition of 0.2 mM pyruvate and monitored at 340 nm.
Tm NAGS activity was assessed by determining the NAG production rate by high-performance liquid chromatography (HPLC). The appropriate amount of enzyme solution was incubated at 50°C in the buffer containing 0.2 mM each of L-glutamate and acetyl-CoA. The reactions were quenched by the addition of an equal volume of ice-cold methanol. HPLC analysis was performed as described below. One unit (U) of the enzyme was defined as the amount capable of product formation at a rate of 1 μmol min-1.
The basal reaction mixture was composed of 100 mM KCl, 10 mM MgCl2, 0.2 mM CoA, 0.2 mM NAD+, 0.2 mM NADH, and 50 mM HEPES-NaOH (pH 7.0). Recombinant cells were suspended in 50 mM HEPES-NaOH (pH 7.0) and heated at 70°C for 30 min, although an incubation temperature of 60°C was employed for the heat-treatment of E. coli having Ap PDC. The heat-treated suspensions of E. coli having Ap PDC, Tt ADDH, Tt GDH, and Tm NAGS were added into the reaction mixture at final concentrations of 16, 7, 50, and 10 mg (wet weight cells) ml-1, respectively. The mixture (8 ml) was placed in a 10-ml rubber-capped cylindrical vessel and stirred in a water bath kept at 50°C. The reaction was initiated by adding the indicated concentrations of substrates and the intermediates. The feeding of a substrate solution consisting of 100 mM each of α-ketoglutarate, pyruvate, and NH4Cl was started using a LC-20 AD solvent delivery unit (Shimadzu, Kyoto, Japan). The feeding rate was kept at 3.2 μl min-1 (= 0.04 μmol ml-1 min-1). Simultaneously, 0.2 mM pyruvate, 0.2 mM acetyl-CoA, 0.2 mM α-ketoglutarate, 0.2 mM L-glutamate, and 100 mM NH4Cl were added to the reaction mixture to maintain the constant pool sizes. The reaction was quenched at timed intervals by the addition of an equal volume of pre-cooled methanol and analyzed to determine the product and intermediates.
Acetate, α-ketoglutarate, pyruvate, and NAG were determined by HPLC on two ion exclusion columns connected in tandem as described previously . CoA and acetyl-CoA were quantified by HPLC using a Cosmosil 5C18-ARII column (4.6 × 250 mm, Nacalai Tesque, Kyoto, Japan). The column was eluted with 0.1% (v/v) orthophosphate solution for the initial 2 min and then with a linear gradient of methanol (0–30 vol%) for 10 min. The column was maintained at 40°C and the flow rate of 1.0 ml min-1 was employed for the elution. The eluent was analyzed with a SPD-20AV UV-VIS detector (Shimadzu) at 254 nm. Aldehydes were fluorescence-labeled with 1,3-cyclohexanedione as described elsewhere  and determined using HPLC on the same column. The column was eluted at 40°C using 30% (v/v) acetonitrile at a flow rate of 0.5 ml min-1. The eluent was analyzed with an RF10A fluorometric detector (Shimadzu) at excitation and emission wavelengths of 366 and 440 nm, respectively. NAD(H) and glutamate were quantified using commercial kits as described in the instructions provided by the manufacturers (Biovision, Mountain View, CA, USA and Cosmo Bio, Tokyo, Japan, respectively).
This work was supported in part by the Japan Science and Technology Agency (JST), PRESTO program. This work was also partly supported by the Japan Society for the Promotion of Science (JSPS), Japanese-German Graduate Externship Program.
- Alper H, Stephanopoulos G: Engineering for biofuels: exploiting innate microbial capacity or importing biosynthetic potential?. Nat Rev Microbiol. 2009, 7: 715-723. 10.1038/nrmicro2186View ArticleGoogle Scholar
- Atsumi S, Cann AF, Connor MR, Shen CR, Smith KM, Brynildsen MP, Chou KJY, Hanai T, Liao JC: Metabolic engineering of Escherichia coli for 1-butanol production. Metab Eng. 2008, 10: 305-311. 10.1016/j.ymben.2007.08.003View ArticleGoogle Scholar
- Kwok R: Five hard truths for synthetic biology. Nature. 2010, 463: 288-290. 10.1038/463288aView ArticleGoogle Scholar
- Zhang Y-HP: Production of biocommodities and bioelectricity by cell-free synthetic enzymatic pathway biotransformation: challenges and opportunities. Biotechnol Bioeng. 2010, 105: 663-677.Google Scholar
- Hodgman CE, Jewett MC: Cell-free synthetic biology: thinking outside the cell. Metab Eng. 2012, 14: 261-269. 10.1016/j.ymben.2011.09.002View ArticleGoogle Scholar
- Ye X, Honda K, Sakai T, Okano K, Omasa T, Hirota R, Kuroda A, Ohtake H: Synthetic metabolic engineering-a novel, simple technology for designing a chimeric metabolic pathway. Microb Cell Fact. 2012, 11: 120- 10.1186/1475-2859-11-120View ArticleGoogle Scholar
- Ye X, Honda K, Morimoto Y, Okano K, Ohtake H: Direct conversion of glucose to malate by synthetic metabolic engineering. J Biotechnol. 2013, 164: 34-40. 10.1016/j.jbiotec.2012.11.011View ArticleGoogle Scholar
- Ninh PH, Honda K, Yokohigashi Y, Okano K, Omasa T, Ohtake H: Development of a continuous bioconversion system using a thermophilic whole-cell biocatalyst. Appl Environ Microbiol. 2013, 79: 1996-2001. 10.1128/AEM.03752-12View ArticleGoogle Scholar
- Dewick PM: The biosynthesis of C5–C25 terpenoid compounds. Nat Prod Rep. 2002, 19: 181-222. 10.1039/b002685iView ArticleGoogle Scholar
- Steinbüchel A, Füchtenbusch B: Bacterial and other biological systems for polyester production. Trends Biotechnol. 1998, 16: 419-427. 10.1016/S0167-7799(98)01194-9View ArticleGoogle Scholar
- Bond-Watts BB, Bellerose RJ, Chang MCY: Enzyme mechanism as a kinetic control element for designing synthetic biofuel pathways. Nat Chem Biol. 2011, 7: 222-227. 10.1038/nchembio.537View ArticleGoogle Scholar
- Chen Y, Daviet L, Schalk M, Siewers V, Nielsen J: Establishing a platform cell factory through engineering of yeast acetyl-CoA metabolism. Metab Eng. 2013, 15: 48-54.View ArticleGoogle Scholar
- Patel MS, Roche TE: Molecular biology and biochemistry of pyruvate dehydrogenase complexes. FASEB J. 1990, 4: 3224-33.Google Scholar
- Blamey JM, Adams MWW: Purification and characterization of ferredoxin oxidoreductase from the hyperthermophilic archaeon Pyrococcus furiosus. Biochim Biophys Acta. 1993, 1161: 19-27. 10.1016/0167-4838(93)90190-3View ArticleGoogle Scholar
- Furdui C, Ragsdale SW: The role of pyruvate ferredoxin oxidoreductase in pyruvate synthesis during autotrophic growth by the Wood-Ljungdahl pathway. J Biol Chem. 2000, 275: 28494-28499.View ArticleGoogle Scholar
- Furdui C, Ragsdale SW: The roles of coenzyme A in the pyruvate:ferredoxin oxidoreductase reaction mechanism: rate enhancement of electron transfer from a radical intermediate to an iron − sulfur cluster. Biochemistry. 2002, 41: 9921-9937. 10.1021/bi0257641View ArticleGoogle Scholar
- Knappe J, Blaschkowski HP, Grobner P, Schmitt T: Pyruvate formate-lyase of Escherichia coli: the acetyl-enzyme intermediate. Eur J Biochem. 1974, 50: 253-263. 10.1111/j.1432-1033.1974.tb03894.xView ArticleGoogle Scholar
- Külzer R, Pils T, Kappl R, Hüttermann J, Knappe J: Reconstitution and characterization of the polynuclear iron-sulfur cluster in pyruvate formate-lyase-activating enzyme. J Biol Chem. 1998, 273: 4897-4903. 10.1074/jbc.273.9.4897View ArticleGoogle Scholar
- Yang H, Hainfeld JF, Wall JS, Frey PA: Quaternary structure of pyruvate dehydrogenase complex from Escherichia coli. J Biol Chem. 1985, 260: 16049-16051.Google Scholar
- Knappe J, Neugebauer FA, Blaschkowski HP, Ganzler M: Post-translational activation introduces a free radical into pyruvate formate-lyase. Proc Natl Acad Sci U S A. 1984, 81: 1332-1335. 10.1073/pnas.81.5.1332View ArticleGoogle Scholar
- Friedman HH, Halik LK, Schwarz MH: Skin moisturizer. 1976, US: Patent 3932622 AGoogle Scholar
- Van Scott-Eugene J, Yu-Ruey J: Anti-odor compositions and therapeutic use. World Patent. 2006, 119283: A2-Google Scholar
- Dawes EA, Ribbons DW, Large PJ: The route of ethanol formation in Zymomonas mobilis. Biochem J. 1966, 98: 795-803.View ArticleGoogle Scholar
- Raj KC, Ingram LO, Maupin-Furlow JA: Pyruvate decarboxylase: a key enzyme for the oxidative metabolism of lactic acid by Acetobacter pasteurianus. Arch Microbiol. 2001, 176: 443-451. 10.1007/s002030100348View ArticleGoogle Scholar
- Gocke D, Graf T, Brosi H, Frindi-Wosch I, Walter L: Comparative characterization of thiamine diphosphate-dependent decarboxylases. J Mol Catal B Enzym. 2009, 61: 30-35. 10.1016/j.molcatb.2009.03.019.View ArticleGoogle Scholar
- Yokoyama S, Matsuo Y, Hirota H, Kigawa T, Shirouzu M, Kuroda Y, Kurumizaka H, Kawaguchi S, Ito Y, Shibata T, Kainosho M, Nishimura Y, Inoue Y, Kuramitsu S: Structural genomics projects in Japan. Nat Struct Biol. 2000, 7: 943-945.View ArticleGoogle Scholar
- Van de-Casteele M, Demarez M, Legrain C, Glansdorff N, Piérard A: Pathways of arginine biosynthesis in extreme thermophilic archaeo- and eubacteria. J Gen Microbiol. 1990, 136: 1177-1183. 10.1099/00221287-136-7-1177.View ArticleGoogle Scholar
- Shiba Y, Paradise EM, Kirby J, Ro D-K, Keasling JD: Engineering of the pyruvate dehydrogenase bypass in Saccharomyces cerevisiae for high-level production of isoprenoids. Metab Eng. 2007, 9: 160-168. 10.1016/j.ymben.2006.10.005View ArticleGoogle Scholar
- Stahovec WL, Mopper K: Trace analysis of aldehydes by pre-column fluorigenic labeling with 1, 3-cyclohexanedione and reversed-phase high-performance liquid chromatography. J Chromatogr A. 1984, 298: 399-406.View ArticleGoogle Scholar
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