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Artificial cofactor regeneration with an iron(III)porphyrin as NADH-oxidase mimic in the enzymatic oxidation of l -glutamate to α-ketoglutarate
Journal of Molecular Catalysis B: Enzymatic 103 (2014) 10–15
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Artificial cofactor regeneration with an iron(III)porphyrin as NADH-oxidase mimic in the enzymatic oxidation of l-glutamate to ␣-ketoglutarate Wilko Greschner a , Carsten Lanzerath b , Tina Reß a , Katharina Tenbrink a,c , Sonja Borchert c , Andreas Mix a , Werner Hummel b,∗ , Harald Gröger a,c,∗∗ a
Faculty of Chemistry, Bielefeld University, Universitätsstraße 25, 33615 Bielefeld, Germany Institute of Molecular Enzyme Technology, Research Centre Jülich, Wilhelm-Johnen-Straße, 52426 Jülich, Germany c Department of Chemistry and Pharmacy, University of Erlangen-Nürnberg, Henkestr. 42, 91054 Erlangen, Germany b
a r t i c l e
i n f o
Article history: Available online 28 December 2013 Keywords: Artificial cofactor recycling Amino acid dehydrogenase Biomimetic catalysis ␣-Ketoglutarate Oxidation
a b s t r a c t In this contribution the use of an artificial in situ-cofactor regeneration with [Fe(III)TSPP]Cl for enzymatic amino acid oxidation, exemplified for the l-glutamate dehydrogenase-catalyzed synthesis of ␣-ketoglutarate from sodium l-glutamate, is reported. In comparison of two l-glutamate dehydrogenases, the one isolated from Clostridium difficile turned out to be the preferred enzyme. At a substrate concentration of 15 mM of l-glutamate in situ-cofactor regeneration using [Fe(III)TSPP]Cl as an “artificial NADH-oxidase” proceeded smoothly, leading to up to >99% overall conversion and 88% conversion related to the formation of ␣-ketoglutarate after 24 h. At an increased concentration of 50 mM of l-glutamate, a somewhat decreased conversion of 43% was observed (which, however, corresponds to a nearly doubled volumetric productivity of 3.95 g/(L d) compared to the experiments at 15 mM). Thus, the iron complex [Fe(III)TSPP]Cl turned out to be capable to be used for cofactor regeneration of the cofactor NAD+ for enzymatic amino acid oxidation. © 2014 Elsevier B.V. All rights reserved.
1. Introduction In the last decades an impressive number of enzymatic redox processes has been established on industrial scale for the synthesis of a broad variety of commercial products [1,2]. In particular this is true for reductive processes, and, e.g., today production routes of chiral alcohols and in part l-amino acids required in the pharmaceutical industry are typically based on dehydrogenase-catalyzed reactions such as reduction and reductive amination, respectively. A key prerequisite for technical feasibility in all of such processes is efficiency of the in situ-cofactor regeneration. Due to the high price of the cofactor its use in low (catalytic) amount is required. Whereas for reductive processes a range of in situ-cofactor regenerations are available with several of them being scalable as well [1–4], in the oxidative mode only a few strategies have been studied more in detail for synthetic purpose. The most prominent approach is based on the use of an NADH-oxidase [3–9].
∗ Corresponding author. Tel.: +49 2461 61 3790. ∗∗ Corresponding author at: Faculty of Chemistry, Bielefeld University, Universitätsstraße 25, 33615 Bielefeld, Germany. Tel.: +49 521 106 2057. E-mail addresses: [email protected] (W. Hummel), [email protected] (H. Gröger). 1381-1177/$ – see front matter © 2014 Elsevier B.V. All rights reserved. http://dx.doi.org/10.1016/j.molcatb.2013.12.015
Attractive features are the use of molecular oxygen for oxidation of NADH (which is formed in the dehydrogenase-catalyzed oxidation process from NAD+ ) and the formation of water as by-product at least in case of some NADH-oxidases. On the other hand, the number of available NADH-oxidases is still limited and some of them form hydrogen peroxide as a by-product, which is disadvantageous since H2 O2 typically causes deactivation of enzymes. Alternatively, artificial in situ-cofactor regeneration has been studied. An early breakthrough was achieved by the Steckhan group by means of a rhodium complex as a biomimetic NADH-oxidase [10]. In this case hydrogen peroxide is again formed as a by-product, thus requiring a catalase as an additional enzyme component in the reaction system for cleavage of hydrogen peroxide to molecular oxygen and water. Recently we demonstrated as a proof of concept the ability of an iron(III)porphyrin complex (3; [Fe(III)TSPP]Cl) for in situ-regeneration of the oxidized cofactors NAD+ and NADP+ through reduction of molecular oxygen and the coupling of this type of cofactor regeneration with alcohol dehydrogenase- and glucose dehydrogenase-catalyzed oxidation reactions [10]. Thus, this iron(III)porphyrin-complex represents an “artificial NAD(P)Hoxidase” for in situ-cofactor regeneration in enzymatic oxidation processes. Furthermore we did not observe hydrogen peroxide as a side product, which indicates that water is formed as a byproduct. Since this initial study [10] with the iron porphyrin 3 has
W. Greschner et al. / Journal of Molecular Catalysis B: Enzymatic 103 (2014) 10–15
Scheme 1. Concept for the biocatalytic synthesis of ␣-ketoglutarate with “artificial” in situ-cofactor regeneration using an ironporphyrin as an NADH-oxidase substitute.
focused on oxidation of carbohydrates and an alcohol, we became interested to explore if this iron porphyrin also is compatible with other dehydrogenase-catalyzed oxidations. In particular the use of [Fe(III)TSPP]Cl for artificial in situ-cofactor regeneration in glutamate dehydrogenase (GluDH)-catalyzed oxidation of l-glutamate toward ␣-ketoglutarate found our interest due to the industrial relevance of ␣-ketoglutarate (2). In the following we report our results on the use of the artificial in situ-cofactor regeneration with [Fe(III)TSPP]Cl 3 in amino acid oxidation, exemplified for the lglutamate dehydrogenase-catalyzed synthesis of ␣-ketoglutarate (2) from l-glutamate (l-1) according to Scheme 1. 2. Experimental
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2.2.2. Molecular cloning of the l-glutamate dehydrogenases from C. difficile and F. nucleatum Genomic DNA from C. difficile (DSM 12056) or F. nucleatum subsp. nucleatum (DSM 15643) (purchased by the German Collection of Microorganisms and Cell Cultures (DSMZ)) was used as a template for the amplification of the gludh genes. For cloning steps following primers were used: GluDH from C. difficile forward: 5 -TGCCGGTCTCGCATGTCAGGAAAAGATGTAAATGTCTTCGAGATGGCGCAATC-3 , GluDH from C. difficile reverse: 5 -CATGGAGCTCTTAGTACCATCCTCTTAATTTCATAGCTTCAGCAACTTTCTTAATTGAA TGCATG-3 , GluDH from F. nucleatum forward: 5 -TGCCGGTCTCGCATGAGTAAAGAAACTTTAAACCCATTAGAAAGCGGACAAAAACAA GTTAAAAAAG-3 GluDH from F. nucleatum reverse: 5 -CATGGAGCTCTTAGTACCATCCTCTAATTTTCATAGCTTCAGCTATTCTCTTAATAGAT TTCATATAAGTAGCTTGTCTA-3 Forward primers contain a recognition site for the restriction enzyme BsaI, reverse primer for the restriction enzyme SacI. Primers were products of MWG Biotech. PCR was performed with a DNA Thermal Cycler from Eppendorf in a total volume of 0.05 mL containing 0.2 mM dNTPs, 30 pmol of each primer, 5 ng template DNA, 0.010 mL of 5x reaction buffer (HV, Fermentas) and 2.5YU Phusion polymerase (Fermentas). After an initial denaturation for 3 min at 98 ◦ C PCR was run for 30 cycles with the following program: denaturation by incubation at 98 ◦ C for 20Ys; annealing at 64 ◦ C for 20Ys; elongation at 72 ◦ C for 1 min, finished by an final elongation for 5 min. After amplification PCR fragments were digested with BsaI/SacI and purified by preparative agarose gel and gel extraction kit (AnalytikJena). 2.2.3. Construction of l-glutamate dehydrogenases GluDH expression vectors The GluDH genes from C. difficile and F. nucleatum, respectively, were cloned into the first multiple cloning site of the expression vector pETDuet-1 (Novagen) under the control of the T7-promoter yielding pDGluDHCD and pDGluDHFN using T4 DNA ligase (Fermentas) according to the manufacturer’s instructions. Resulting vectors were transformed into competent E. coli DH5 cells and transformants were selected on LB agar plates supplemented with 100 g ampicillin per mL. Correct amplification was confirmed by restriction analysis and DNA sequencing. Sequencing was carried out by MWG Germany. For protein production, vectors pDGluDHCD and pDGluDHFN were transformed in the E.coli host strain BL21(DE3).
2.1. Materials The catalyst [Fe(III)TSPP]Cl (3; 5,10,15,20-tetrakis-(4sulfonatophenyl)-porphyrin-iron(III)chloride) is commercially available and was purchased from TriPorTech GmbH (product number: tpt00142617). The sodium l-glutamate monohydrate (1) was purchased from Tokyo Chemical Industry, the ␣-ketoglutarate sodium salt (2) was available from Applichem GmbH and both were used as received. 2.2. Preparation of the NAD+ -dependent l-glutamate dehydrogenases from Clostridium difficile and Fusobacterium nucleatum and determination of their enzyme activities (according to Table 1 and Scheme 1) 2.2.1. Microorganisms and culture conditions E. coli BL21(DE3) and E. coli DH5 were grown in a 1 L-scale in LB medium containing 100 g/mL of ampicillin, 1.0% of tryptone, 1.0% of NaCl, and 0.5% of yeast extract as well as for non liquid medium 1.5% of agar. The pH was measured and adjusted to 7.5.
2.2.4. Production and purification of the recombinant l-glutamate dehydrogenases from C. difficile and F. nucleatum The strains E. coli BL21(DE3)/pDGluDHCD and E. coli BL21(DE3)/pDGluDHFN were cultivated in 10 mL of LB medium containing 100 g/mL ampicillin overnight at 37 ◦ C. These cultures were used to inoculate 1YL of LB medium for expression containing 100 g/mL ampicillin at a final optical density 600 nm of 0.05. Culturing was done in shaking flasks (120 rpm) at 30 ◦ C and the production of recombinant GluDH under the T7 -promoter was induced by the addition of IPTG (isopropyl thio--d-galactoside) in a final concentration of 0.2 mM when the optical density at 595 nm reached 0.5, followed by further growing for enzyme production for 16 h. The following steps were done in the same way for the GluDHs from C. difficile and F. nucleatum. Cells were harvested by centrifugation at 5000 × g for 30 min at 4 ◦ C (centrifuge by Hettich), suspended for lysis (25% cell suspension) in 50 mM phosphate buffer (pH 7.5) and disrupted by three sonification cycles of 1 min (cooling periods for 3 min on ice between) (Sonopuls HD 60 ultrasonic oscillator, power: 80%, cycles: 50; Bandelin, Germany). After sonification, separation of cell debris from the cell lysate
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was done by centrifugation at 14,000 × g for 30 min at 4 ◦ C (centrifuge by Hettich). After obtaining the clear supernatant a heat precipitation was performed by incubating the enzyme solution at a temperature of 59 ◦ C (according to Anderson et al. [11] and Gharbia and Haroun [12] where the high thermal stability of both GluDHs are described). The residence time was adjusted to 60 min. Separation of denaturated proteins was done by centrifugation at 14,000 rpm for 30 min at 4 ◦ C and the supernatant was taken as enzyme solution for the subsequent experiments. 2.2.5. Determination of the enzyme activity of l-glutamate dehydrogenase Enzyme activity was determined for the oxidative deamination of l-glutamate. The standard reaction mixture contained 50 mM lglutamate, 2.5 mM of NAD+ in 100 mM Tris–HCl buffer (pH 9.0), and enzyme in a final volume of 1 mL. For product inhibition studies, 50 mM ␣-ketoglutarate was added. The substrate was replaced by water in a blank experiment. Enzyme solution (0.01 mL) was added and the change in absorbance at 340 nm was monitored for 1 min at 30 ◦ C. One unit of the GluDH is defined as the amount of enzyme that catalyzes the formation of 1 mol of NADH per min in the oxidative deamination of l-glutamate under the conditions mentioned above with a molar absorption coefficient of 6.22 mM−1 cm−1 for NADH. The specific activity is expressed as units per mg of protein. 2.2.6. NADH oxidase from Lactobacillus brevis Preparation of NADH oxidase from L. brevis as a recombinant enzyme in E. coli BL21(DE3) and the determination of its activity using 0.1 mM NADH at pH 7.0 were carried out as described by Geueke et al. [5]. 2.2.7. Determination of the amount of protein in enzyme crude extracts The determinations of the amount of protein in the crude and partially purified extracts were done by means of the method of Bradford [13] using bovine serum albumin as a standard. The expression levels were monitored by means of a polyacrylamide gel electrophoresis using commercial Bis/tris-gels (4–12%) and the NuPAGE-System from Invitrogen. Prestained SeeBlue Protein Standard from Invitrogen was used as molecular weight standard. 2.3. Determination of time-dependent stability of ˛–ketoglutarate in buffered aqueous solution To a solution of sodium l-glutamate monohydrate (l-1; 22.5 mg; 0.12 mmol) and sodium ␣–ketoglutarate (2; 5.0 mg; 0.03 mmol) in phosphate buffer (3.0 mL; pH 8; 100 mM) or in carbonate buffer (3.0 mL; pH 9.6; 100 mM) the GluDH from C. difficile or F. nucleatum (15YU referring to the activity for l-glutamate oxidation) was added. The mixture was stirred with 550 rpm at room temperature for 48 h. Samples of 0.3 mL were periodically taken and the protein was removed through ultrafiltration-centrifugation (10 kDa MWCO; 12.000 rpm). To each sample 0.05 mL of a freshly prepared 50 mM pivalic acid solution and 0.250 mL deuterium oxide were added and analyzed by 1 H-NMR spectroscopy using pivalic acid (trimethylacetic acid) as an internal standard in combination with a presaturation technique for suppression of the water signal. 2.4. Typical procedure for the synthesis of ˛-ketoglutarate (2) (according to Tables 2 and 3) To a solution, consisting of sodium l-glutamate monohydrate (l-1; 14.0 mg; 75 mol respectively 46.8 mg; 250 mol) and [Fe(III)TSPP]Cl (3; 5.1 mg; 5 mol) or NADH-oxidase from L. brevis (22.5YU referring to the activity for NADH) in phosphate (or carbonate/bicarbonate) buffer (5.0 mL; 100 mM), the GluDH from
Table 1 Enzyme activities of the prepared recombinant l-GluDHs from F. nucleatum and C. difficile
. Entry
Original strain of recombinant l-GluDH
Enzyme activity of crude extracta [U/mg]
Enzyme activity after heat treatment [U/mg]b
1 2
Fusobacterium nucleatum Clostridium difficile
6 8
16 (32)c 20 (146)c
a For preparation of the crude extracts and conditions of spectrophotometric activity determination, see Section 2. b Heat treatment for 1 h at 60 ◦ C; for details, see Section 2. c In parentheses literature values for the specific activities of the purified proteins are given (in U/mg).
C. difficile or from F. nucleatum (15YU referring to the activity for l-glutamate oxidation) was added. With the addition of NAD+ (6.6 mg; 10 mol) the reaction was started and stirred with 550 rpm at room temperature. Workup and determination of the conversion were done as described in Section 2.3. 3. Results and discussion First, we searched for an l-glutamate dehydrogenase (l-GluDH) synthetically suitable for oxidative deamination of sodium lglutamate (l-GluDH, l-1) since our own preliminary experiments and literature data [8,14] revealed strong product inhibition of commercially available l-glutamate dehydrogenases for this purpose. For example, GluDH from C. symbiosum shows a severe product inhibition (leading to a reduction of activity of 70%) caused by only 3 mM of ␣-ketoglutarate (2) when converting 5 mM of l-1 [8]. The resulting productivities with 1.07 g/(L d) at the highest [8] limit preparative applicability of this enzymatic approach based on glutamate dehydrogenase and NADH-oxidase. Thus, the established technical process is based not on biocatalysis but on oxidation of l-1 in air in the presence of a copper complex [8]. Since in our initial literature search for alternative l-GluDHs we recognized that data with respect to product inhibition of l-GluDHs are very rare, we focused on l-GluDHs with a catabolic function in microbial metabolisms, since the natural function of such enzymes is the conversion (degradation) of l-Glu (l-1) to ␣-ketoglutarate (2). Among about 30 catabolic l-GluDHs in literature we priorized the enzymes of F. nucleatum and C. difficile. These GluDHs are reported to have a high specific activity of about 32YU/mg (GluDH from F. nucleatum) [12] and 146YU/mg (GluDH from C. difficile) [11] in comparison to GluDHs from other sources like such as Peptostreptococcus asaccharolyticus (7.8YU/mg) [15], Bacteroides fragilis (7.3YU/mg) [16], Pyrobacterium islandicum (3.5YU/mg) [17], and Janthinobacterium lividum (3.1YU/mg) [18]. Recombinant expression and production of sufficient amounts of both biocatalyst were conducted applying an E. coli expression system as mentioned above. Both enzymes, namely the GluDHs from C. difficile and F. nucleatum, were successfully expressed in E. coli as a soluble and fully active protein. The specific enzyme activities in the crude extracts of 6 and 8YU/mg of protein for the GluDHs from F. nucleatum and C. difficile, respectively, could be increased nearly threefold up to 16 and 20YU/mg after heat precipitation for the recombinant l-GluDHs as summarized in Table 1. From 4 g of cells (wet weight) we obtained 1.580YU of the recombinant l-GluDH from F. nucleatum and 1.390YU of the recombinant l-GluDH from C. difficile (corresponding to yields of 76% and 88%, respectively referred to activities of the crude extracts). The
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Table 2 Biotransformation of sodium glutamate (l-1, 15 mM) into 2 with a l-GluDH and [Fe(III)TSPP]Cl (3, 1 mM) or NADH-oxidase from Lactobacillus brevis at 20 ◦ C and pH 8
. Entry
l-GluDH [U/mL]
NOX[U/mL]
Conv.a 2 h [%]
Conv.a 4 h [%]
Conv.a 8 h [%]
Conv.a 24 h [%]
1 2 3
1.5 5.0 1.5
– – 4.5
51(50) 49(51) 36(37)
65(64) 77(60) 60(60)
76(78) 91(89) 77(76)
86(66) >99(88) 82(77)
a The conversion is defined as the amount of consumed substrate 1 related to the original amount of substrate 1 (in parentheses the amount of generated product 2 related to the original amount of substrate 1 (in %) is given).
biochemical data in terms of enzyme activities of the two recombinant l-GluDHs are listed in Table 1. In addition, our expectation that such recombinant catabolic GluDHs show product inhibition to a less extend than above mentioned commercially l-GluDHs has been fulfilled by the two prioritized enzymes. Product inhibition caused by 50 mM of 2 was in the range of 30% for the l-GluDH from C. difficile and around 55% for the l-GluDH from F. nucleatum at pH 9.0, thus leading to residual enzyme activities of 70% and around 45%, respectively (Fig. 1). The lowered activities of both enzymes are caused by the addition of ␣-ketoglutarate, not by an instability of the enzymes themselves because stability studies showed that no deactivation at all could be observed after a period of 24 h. In comparison to the published data obtained by l-GluDH from C. symbiosus the results clearly indicate a significantly lowered product inhibition level, even at higher concentrations in the range of 50 mM. With the two recombinant l-GluDHs in hand, we next focused on the desired preparative enzymatic synthesis of ␣-ketoglutarate (2). Initial work addressed the set up of a robust and reliable analytical method for determination of the conversion. Due to the known potential of ␣-keto acids to decompose, determination of conversion (related to the formation of 2) through the absolute amount of ␣-ketoglutarate (2) via comparison with an internal standard appeared to us more attractive compared to calculation of conversion based on a relative comparison of substrate/product ratio. As an analytical method we chose proton NMR spectroscopy using pivalic acid (trimethylacetic acid) as an internal standard in combination with a presaturation technique (to improve the
Fig. 1. Product inhibition of the l-GluDHs from C. difficile (GluDH CD) and F. nucleatum (GluDH FN) by ␣-ketoglutarate (2) (assay: 50 mM l-1; pH 9.0; 50 mM 2; 2.5 mM NAD+ ).
quality of the baseline of the spectra due to suppression of the large signal resulting from the solvent water) [19]. Next we studied the stability of ␣-ketoglutarate (2) under reaction conditions starting from an ␣-ketoglutarate (2, 10 mM)/l-glutamate mixture (l-1, 40 mM) which simulates a 20% conversion of an enzymecatalyzed oxidation of 50 mM L-glutamate. After optimization of sampling and work-up for analytical determination (showing a deviation in the range of ca. 3%), we found high recovery rates of 97% or more for ␣-ketoglutarate (2) within a time range of 48 h (Fig. 2). Notably, recovery rates were high not only for the ␣-ketoglutarate/l-glutamate mixture but also in the presence of a crude extract of the recombinant l-GluDH from C. difficile. However, when using a crude extract of the l-GluDH from F. nucleatum we observed a significantly higher loss of 2 probably caused by an ␣-ketoglutarate decomposing enzyme present in the partially purified sample of the L-GluDH from F. nucleatum. In general stability of the samples appeared to be high at pH 8.0 as well as at 9.6. Selected examples of this stability study are given in Fig. 2. Next we conducted biotransformations with in situ-cofactor regeneration based on the use of the iron(III)porphyrin [Fe(III)TSPP]Cl (3) as an artificial NADH-oxidase according to Scheme 1. A first proof of concept for the compatibility of glutamate dehydrogenases with the iron(III)porphyrin complex 3 has been demonstrated when conducting the process at pH 8.0 and with a substrate concentration of 15 mM (Table 2). High conversions
Fig. 2. Time-dependent stability of ␣-ketoglutarate (2) determined out of a mixture of ␣-ketoglutarate (2, 10 mM) and l-glutamate (1, 40 mM) (aqueous solution) at pH 8.0 or 9.6. Additionally, crude extracts of GluDH from C. difficile (C.d.) or from F. nucleatum (F.n.) were added to the ␣-ketoglutarate/glutamate mixture.
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Table 3 Biotransformation of l-glutamate (l-1, 50 mM) into ␣-ketoglutarate (2) with l-GluDH and Fe(III)TSPP (3, 1 mM) at pH 8 (according to Scheme 1). Entry
l-GluDH
l-GluDH [U/mL]
Temp. [◦ C]
Conv.a 1 h [%]
Conv.a 4 h [%]
Conv.a 24 h [%]
1 2 3
C. difficile F. nucleatum F. nucleatum
5 5 5
20 20 12
22(22) 5(5) 8(8)
33(33) 9(8) 11(11)
47(43)b 16(5) 28(27)c
a The conversion is defined as the amount of consumed substrate 1 related to the original amount of substrate 1 (in parentheses the amount of generated product 2 related to the original amount of substrate 1 (in %) is given). b Maximum at 49% after 48 h. c Maximum at 33% after 48 h.
were obtained in particular in the presence of the l-GluDH from C. difficile (entries 1 and 2). For example, after optimization and a reaction time of 8 h a conversion of 91% (based on consumed substrate l-1) as well as a product-related conversion of 89% (based on the amount of formed 2, entry 2) were observed, whereas using a reduced amount of the enzyme resulted in a somewhat decreased conversion of 76% (entry 1). The total turnover numbers for the cofactor NAD+ calculated from entries 1 and 2 (Table 2) are within the range of 6.5–7.4. Furthermore, we carried out experiments with NADH-oxidase (NOX) from Lactobacillus brevis in order to compare the “artificial NADH oxidase” (iron(III)porphyrin [Fe(III)TSPP]Cl (3)) with the enzyme NADH oxidase (Table 2; entry 3). Unfortunately, the activity of the enzyme NADH oxidase is relatively low at higher pH values, therefore an excess of activity was added to obtain complete regeneration of NAD+ . Table 2 shows that, in terms of efficiency, the conversion rates of the biomimetic [Fe(III)TSPP]Cl (3) are in the same order of magnitude as the enzyme NADH oxidase. Subsequently, we increased the substrate concentration of l-1 to 50 mM and operated at a pH-value of 8 which turned out to be the preferred pH in terms of product stability. When using the lGluDH from C. difficile we gained after optimization a conversion of 47% at room temperature (Table 3, entry 1). In the case of l-GluDH from F. nucleatum we faced the problem of an immense loss of 2 and low conversions (entry 2). In order to suppress chemical degradation of ␣-ketoglutarate (2), experiments were done at a somewhat decreased temperature of 12 ◦ C. When using the l-GluDH from F. nucleatum as a biocatalyst at this reaction temperature, a product-related conversion of ␣-ketoglutarate (2) (often defined as so-called “reaction yield”) of 27% was found after 24 h (Table 3, entry 3). Further experiments under different conditions like varying the pH value, more intensive cooling during the reaction or stepwise addition of the substrate l-1 did not show higher conversions than those presented in Table 3. Moreover the cooling was the only way to prevent the chemical degradation of ␣-ketoglutarate on prolonged reaction times, especially when the GluDH from F. nucleatum was used.
volumetric productivity of 3.95 g/(L d) compared to the experiments at 15 mM). In addition, this productivity is almost four times higher as the one reported previously in literature when using an NADH-oxidase [8]. Thus, the iron complex [Fe(III)TSPP]Cl turned out to be capable to be used for cofactor regeneration of the cofactor NAD+ for enzymatic amino acid oxidation. The regeneration of NAD+ is a prerequisite if NAD+ -dependent dehydrogenases are designated for oxidation reactions, for example, converting hydroxy acids, amino acids or alcohols into the corresponding keto compounds. Only a few enzymatic methods are described in the literature to achieve this aim, but all of them present also disadvantages: the use of glutamate dehydrogenase [20,21], lactate dehydrogenase [22], or horse liver alcohol dehydrogenase [23] always leads to the formation of by-products, thus complicating the isolation of the desired product. The reactions catalyzed by these enzymes are reversible therefore it is difficult to reach a complete conversion regarding the primary desired oxidizing reaction. Peroxide forming NADH oxidases require the addition of catalase. Furthermore, all enzymatic systems depend typically on one coenzyme species, either NADH or NADPH, which hampers the development of a generalizable reaction platform. Complementing enzymes as catalysts to regenerate the oxidized coenzyme, the iron complex [Fe(III)TSPP]Cl can be used successfully for this purpose. In addition, it can be stored over a long period of time and it accepts NADH as well as NADPH. Furthermore, peroxide is not formed by the reaction catalyzed by this complex. Currently, further extension of this type of “artificial cofactor regeneration system” by means of the [Fe(III)TSPP]Cl (3) is in progress as well as studies with respect to the clarification of the reaction mechanism. Acknowledgements The authors thank the German Federal Environmental Foundation (Deutsche Bundesstiftung Umwelt, DBU) for generous support within the DBU-network “ChemBioTec” of the funding priority “Biotechnology” (Project AZ 13234-32). We further thank. Dr. Philipp Böhm for helpful discussions and technical support.
4. Conclusion
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In conclusion, we reported the use of an artificial in situ-cofactor regeneration with [Fe(III)TSPP]Cl (3) for enzymatic amino acid oxidation, exemplified for the l-glutamate dehydrogenase-catalyzed synthesis of ␣-ketoglutarate (2) from l-glutamate (l-1). In comparison of the two investigated dehydrogenases, the l-glutamate dehydrogenase from C. difficile turned out to be the preferred enzyme. At a substrate concentration of 15 mM of l-glutamate (l-1) in situ-cofactor regeneration using [Fe(III)TSPP]Cl (3) as an “artificial NADH-oxidase” proceeded smoothly, leading to up to >99% overall conversion and 88% conversion related to the formation of desired product 2 after 24 h. At an increased concentration of 50 mM of substrate l-1, a somewhat decreased conversion of 43% was observed (which, however, corresponds to a nearly doubled
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R. Kawakami, H. Sakuraba, T. Ohshima, J. Bacteriol. 189 (2007) 5626–5633. P.J. Hore, Method. Enzymol. 176 (1989) 64–77. N. Sakamoto, A.M. Kotre, M.A. Savageau, J Bacteriol. 124 (1975) 775–783. F.M. Veronese, J.F. Nyc, Y. Degani, D.M. Brown, E.L. Smith, J. Biol. Chem. 249 (1974) 7922–7928. [22] E.I. Garvie, Microbiol. Rev. 44 (1980) 106–139. [23] G.L. Lemiere, J.A. Lepoivre, F.C. Alderweireldt, Tetrahedron Lett. 26 (1985) 4527–4528.
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Journal of Molecular Catalysis B: Enzymatic journal homepage: www.elsevier.com/locate/molcatb
Artificial cofactor regeneration with an iron(III)porphyrin as NADH-oxidase mimic in the enzymatic oxidation of l-glutamate to ␣-ketoglutarate Wilko Greschner a , Carsten Lanzerath b , Tina Reß a , Katharina Tenbrink a,c , Sonja Borchert c , Andreas Mix a , Werner Hummel b,∗ , Harald Gröger a,c,∗∗ a
Faculty of Chemistry, Bielefeld University, Universitätsstraße 25, 33615 Bielefeld, Germany Institute of Molecular Enzyme Technology, Research Centre Jülich, Wilhelm-Johnen-Straße, 52426 Jülich, Germany c Department of Chemistry and Pharmacy, University of Erlangen-Nürnberg, Henkestr. 42, 91054 Erlangen, Germany b
a r t i c l e
i n f o
Article history: Available online 28 December 2013 Keywords: Artificial cofactor recycling Amino acid dehydrogenase Biomimetic catalysis ␣-Ketoglutarate Oxidation
a b s t r a c t In this contribution the use of an artificial in situ-cofactor regeneration with [Fe(III)TSPP]Cl for enzymatic amino acid oxidation, exemplified for the l-glutamate dehydrogenase-catalyzed synthesis of ␣-ketoglutarate from sodium l-glutamate, is reported. In comparison of two l-glutamate dehydrogenases, the one isolated from Clostridium difficile turned out to be the preferred enzyme. At a substrate concentration of 15 mM of l-glutamate in situ-cofactor regeneration using [Fe(III)TSPP]Cl as an “artificial NADH-oxidase” proceeded smoothly, leading to up to >99% overall conversion and 88% conversion related to the formation of ␣-ketoglutarate after 24 h. At an increased concentration of 50 mM of l-glutamate, a somewhat decreased conversion of 43% was observed (which, however, corresponds to a nearly doubled volumetric productivity of 3.95 g/(L d) compared to the experiments at 15 mM). Thus, the iron complex [Fe(III)TSPP]Cl turned out to be capable to be used for cofactor regeneration of the cofactor NAD+ for enzymatic amino acid oxidation. © 2014 Elsevier B.V. All rights reserved.
1. Introduction In the last decades an impressive number of enzymatic redox processes has been established on industrial scale for the synthesis of a broad variety of commercial products [1,2]. In particular this is true for reductive processes, and, e.g., today production routes of chiral alcohols and in part l-amino acids required in the pharmaceutical industry are typically based on dehydrogenase-catalyzed reactions such as reduction and reductive amination, respectively. A key prerequisite for technical feasibility in all of such processes is efficiency of the in situ-cofactor regeneration. Due to the high price of the cofactor its use in low (catalytic) amount is required. Whereas for reductive processes a range of in situ-cofactor regenerations are available with several of them being scalable as well [1–4], in the oxidative mode only a few strategies have been studied more in detail for synthetic purpose. The most prominent approach is based on the use of an NADH-oxidase [3–9].
∗ Corresponding author. Tel.: +49 2461 61 3790. ∗∗ Corresponding author at: Faculty of Chemistry, Bielefeld University, Universitätsstraße 25, 33615 Bielefeld, Germany. Tel.: +49 521 106 2057. E-mail addresses: [email protected] (W. Hummel), [email protected] (H. Gröger). 1381-1177/$ – see front matter © 2014 Elsevier B.V. All rights reserved. http://dx.doi.org/10.1016/j.molcatb.2013.12.015
Attractive features are the use of molecular oxygen for oxidation of NADH (which is formed in the dehydrogenase-catalyzed oxidation process from NAD+ ) and the formation of water as by-product at least in case of some NADH-oxidases. On the other hand, the number of available NADH-oxidases is still limited and some of them form hydrogen peroxide as a by-product, which is disadvantageous since H2 O2 typically causes deactivation of enzymes. Alternatively, artificial in situ-cofactor regeneration has been studied. An early breakthrough was achieved by the Steckhan group by means of a rhodium complex as a biomimetic NADH-oxidase [10]. In this case hydrogen peroxide is again formed as a by-product, thus requiring a catalase as an additional enzyme component in the reaction system for cleavage of hydrogen peroxide to molecular oxygen and water. Recently we demonstrated as a proof of concept the ability of an iron(III)porphyrin complex (3; [Fe(III)TSPP]Cl) for in situ-regeneration of the oxidized cofactors NAD+ and NADP+ through reduction of molecular oxygen and the coupling of this type of cofactor regeneration with alcohol dehydrogenase- and glucose dehydrogenase-catalyzed oxidation reactions [10]. Thus, this iron(III)porphyrin-complex represents an “artificial NAD(P)Hoxidase” for in situ-cofactor regeneration in enzymatic oxidation processes. Furthermore we did not observe hydrogen peroxide as a side product, which indicates that water is formed as a byproduct. Since this initial study [10] with the iron porphyrin 3 has
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Scheme 1. Concept for the biocatalytic synthesis of ␣-ketoglutarate with “artificial” in situ-cofactor regeneration using an ironporphyrin as an NADH-oxidase substitute.
focused on oxidation of carbohydrates and an alcohol, we became interested to explore if this iron porphyrin also is compatible with other dehydrogenase-catalyzed oxidations. In particular the use of [Fe(III)TSPP]Cl for artificial in situ-cofactor regeneration in glutamate dehydrogenase (GluDH)-catalyzed oxidation of l-glutamate toward ␣-ketoglutarate found our interest due to the industrial relevance of ␣-ketoglutarate (2). In the following we report our results on the use of the artificial in situ-cofactor regeneration with [Fe(III)TSPP]Cl 3 in amino acid oxidation, exemplified for the lglutamate dehydrogenase-catalyzed synthesis of ␣-ketoglutarate (2) from l-glutamate (l-1) according to Scheme 1. 2. Experimental
11
2.2.2. Molecular cloning of the l-glutamate dehydrogenases from C. difficile and F. nucleatum Genomic DNA from C. difficile (DSM 12056) or F. nucleatum subsp. nucleatum (DSM 15643) (purchased by the German Collection of Microorganisms and Cell Cultures (DSMZ)) was used as a template for the amplification of the gludh genes. For cloning steps following primers were used: GluDH from C. difficile forward: 5 -TGCCGGTCTCGCATGTCAGGAAAAGATGTAAATGTCTTCGAGATGGCGCAATC-3 , GluDH from C. difficile reverse: 5 -CATGGAGCTCTTAGTACCATCCTCTTAATTTCATAGCTTCAGCAACTTTCTTAATTGAA TGCATG-3 , GluDH from F. nucleatum forward: 5 -TGCCGGTCTCGCATGAGTAAAGAAACTTTAAACCCATTAGAAAGCGGACAAAAACAA GTTAAAAAAG-3 GluDH from F. nucleatum reverse: 5 -CATGGAGCTCTTAGTACCATCCTCTAATTTTCATAGCTTCAGCTATTCTCTTAATAGAT TTCATATAAGTAGCTTGTCTA-3 Forward primers contain a recognition site for the restriction enzyme BsaI, reverse primer for the restriction enzyme SacI. Primers were products of MWG Biotech. PCR was performed with a DNA Thermal Cycler from Eppendorf in a total volume of 0.05 mL containing 0.2 mM dNTPs, 30 pmol of each primer, 5 ng template DNA, 0.010 mL of 5x reaction buffer (HV, Fermentas) and 2.5YU Phusion polymerase (Fermentas). After an initial denaturation for 3 min at 98 ◦ C PCR was run for 30 cycles with the following program: denaturation by incubation at 98 ◦ C for 20Ys; annealing at 64 ◦ C for 20Ys; elongation at 72 ◦ C for 1 min, finished by an final elongation for 5 min. After amplification PCR fragments were digested with BsaI/SacI and purified by preparative agarose gel and gel extraction kit (AnalytikJena). 2.2.3. Construction of l-glutamate dehydrogenases GluDH expression vectors The GluDH genes from C. difficile and F. nucleatum, respectively, were cloned into the first multiple cloning site of the expression vector pETDuet-1 (Novagen) under the control of the T7-promoter yielding pDGluDHCD and pDGluDHFN using T4 DNA ligase (Fermentas) according to the manufacturer’s instructions. Resulting vectors were transformed into competent E. coli DH5 cells and transformants were selected on LB agar plates supplemented with 100 g ampicillin per mL. Correct amplification was confirmed by restriction analysis and DNA sequencing. Sequencing was carried out by MWG Germany. For protein production, vectors pDGluDHCD and pDGluDHFN were transformed in the E.coli host strain BL21(DE3).
2.1. Materials The catalyst [Fe(III)TSPP]Cl (3; 5,10,15,20-tetrakis-(4sulfonatophenyl)-porphyrin-iron(III)chloride) is commercially available and was purchased from TriPorTech GmbH (product number: tpt00142617). The sodium l-glutamate monohydrate (1) was purchased from Tokyo Chemical Industry, the ␣-ketoglutarate sodium salt (2) was available from Applichem GmbH and both were used as received. 2.2. Preparation of the NAD+ -dependent l-glutamate dehydrogenases from Clostridium difficile and Fusobacterium nucleatum and determination of their enzyme activities (according to Table 1 and Scheme 1) 2.2.1. Microorganisms and culture conditions E. coli BL21(DE3) and E. coli DH5 were grown in a 1 L-scale in LB medium containing 100 g/mL of ampicillin, 1.0% of tryptone, 1.0% of NaCl, and 0.5% of yeast extract as well as for non liquid medium 1.5% of agar. The pH was measured and adjusted to 7.5.
2.2.4. Production and purification of the recombinant l-glutamate dehydrogenases from C. difficile and F. nucleatum The strains E. coli BL21(DE3)/pDGluDHCD and E. coli BL21(DE3)/pDGluDHFN were cultivated in 10 mL of LB medium containing 100 g/mL ampicillin overnight at 37 ◦ C. These cultures were used to inoculate 1YL of LB medium for expression containing 100 g/mL ampicillin at a final optical density 600 nm of 0.05. Culturing was done in shaking flasks (120 rpm) at 30 ◦ C and the production of recombinant GluDH under the T7 -promoter was induced by the addition of IPTG (isopropyl thio--d-galactoside) in a final concentration of 0.2 mM when the optical density at 595 nm reached 0.5, followed by further growing for enzyme production for 16 h. The following steps were done in the same way for the GluDHs from C. difficile and F. nucleatum. Cells were harvested by centrifugation at 5000 × g for 30 min at 4 ◦ C (centrifuge by Hettich), suspended for lysis (25% cell suspension) in 50 mM phosphate buffer (pH 7.5) and disrupted by three sonification cycles of 1 min (cooling periods for 3 min on ice between) (Sonopuls HD 60 ultrasonic oscillator, power: 80%, cycles: 50; Bandelin, Germany). After sonification, separation of cell debris from the cell lysate
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was done by centrifugation at 14,000 × g for 30 min at 4 ◦ C (centrifuge by Hettich). After obtaining the clear supernatant a heat precipitation was performed by incubating the enzyme solution at a temperature of 59 ◦ C (according to Anderson et al. [11] and Gharbia and Haroun [12] where the high thermal stability of both GluDHs are described). The residence time was adjusted to 60 min. Separation of denaturated proteins was done by centrifugation at 14,000 rpm for 30 min at 4 ◦ C and the supernatant was taken as enzyme solution for the subsequent experiments. 2.2.5. Determination of the enzyme activity of l-glutamate dehydrogenase Enzyme activity was determined for the oxidative deamination of l-glutamate. The standard reaction mixture contained 50 mM lglutamate, 2.5 mM of NAD+ in 100 mM Tris–HCl buffer (pH 9.0), and enzyme in a final volume of 1 mL. For product inhibition studies, 50 mM ␣-ketoglutarate was added. The substrate was replaced by water in a blank experiment. Enzyme solution (0.01 mL) was added and the change in absorbance at 340 nm was monitored for 1 min at 30 ◦ C. One unit of the GluDH is defined as the amount of enzyme that catalyzes the formation of 1 mol of NADH per min in the oxidative deamination of l-glutamate under the conditions mentioned above with a molar absorption coefficient of 6.22 mM−1 cm−1 for NADH. The specific activity is expressed as units per mg of protein. 2.2.6. NADH oxidase from Lactobacillus brevis Preparation of NADH oxidase from L. brevis as a recombinant enzyme in E. coli BL21(DE3) and the determination of its activity using 0.1 mM NADH at pH 7.0 were carried out as described by Geueke et al. [5]. 2.2.7. Determination of the amount of protein in enzyme crude extracts The determinations of the amount of protein in the crude and partially purified extracts were done by means of the method of Bradford [13] using bovine serum albumin as a standard. The expression levels were monitored by means of a polyacrylamide gel electrophoresis using commercial Bis/tris-gels (4–12%) and the NuPAGE-System from Invitrogen. Prestained SeeBlue Protein Standard from Invitrogen was used as molecular weight standard. 2.3. Determination of time-dependent stability of ˛–ketoglutarate in buffered aqueous solution To a solution of sodium l-glutamate monohydrate (l-1; 22.5 mg; 0.12 mmol) and sodium ␣–ketoglutarate (2; 5.0 mg; 0.03 mmol) in phosphate buffer (3.0 mL; pH 8; 100 mM) or in carbonate buffer (3.0 mL; pH 9.6; 100 mM) the GluDH from C. difficile or F. nucleatum (15YU referring to the activity for l-glutamate oxidation) was added. The mixture was stirred with 550 rpm at room temperature for 48 h. Samples of 0.3 mL were periodically taken and the protein was removed through ultrafiltration-centrifugation (10 kDa MWCO; 12.000 rpm). To each sample 0.05 mL of a freshly prepared 50 mM pivalic acid solution and 0.250 mL deuterium oxide were added and analyzed by 1 H-NMR spectroscopy using pivalic acid (trimethylacetic acid) as an internal standard in combination with a presaturation technique for suppression of the water signal. 2.4. Typical procedure for the synthesis of ˛-ketoglutarate (2) (according to Tables 2 and 3) To a solution, consisting of sodium l-glutamate monohydrate (l-1; 14.0 mg; 75 mol respectively 46.8 mg; 250 mol) and [Fe(III)TSPP]Cl (3; 5.1 mg; 5 mol) or NADH-oxidase from L. brevis (22.5YU referring to the activity for NADH) in phosphate (or carbonate/bicarbonate) buffer (5.0 mL; 100 mM), the GluDH from
Table 1 Enzyme activities of the prepared recombinant l-GluDHs from F. nucleatum and C. difficile
. Entry
Original strain of recombinant l-GluDH
Enzyme activity of crude extracta [U/mg]
Enzyme activity after heat treatment [U/mg]b
1 2
Fusobacterium nucleatum Clostridium difficile
6 8
16 (32)c 20 (146)c
a For preparation of the crude extracts and conditions of spectrophotometric activity determination, see Section 2. b Heat treatment for 1 h at 60 ◦ C; for details, see Section 2. c In parentheses literature values for the specific activities of the purified proteins are given (in U/mg).
C. difficile or from F. nucleatum (15YU referring to the activity for l-glutamate oxidation) was added. With the addition of NAD+ (6.6 mg; 10 mol) the reaction was started and stirred with 550 rpm at room temperature. Workup and determination of the conversion were done as described in Section 2.3. 3. Results and discussion First, we searched for an l-glutamate dehydrogenase (l-GluDH) synthetically suitable for oxidative deamination of sodium lglutamate (l-GluDH, l-1) since our own preliminary experiments and literature data [8,14] revealed strong product inhibition of commercially available l-glutamate dehydrogenases for this purpose. For example, GluDH from C. symbiosum shows a severe product inhibition (leading to a reduction of activity of 70%) caused by only 3 mM of ␣-ketoglutarate (2) when converting 5 mM of l-1 [8]. The resulting productivities with 1.07 g/(L d) at the highest [8] limit preparative applicability of this enzymatic approach based on glutamate dehydrogenase and NADH-oxidase. Thus, the established technical process is based not on biocatalysis but on oxidation of l-1 in air in the presence of a copper complex [8]. Since in our initial literature search for alternative l-GluDHs we recognized that data with respect to product inhibition of l-GluDHs are very rare, we focused on l-GluDHs with a catabolic function in microbial metabolisms, since the natural function of such enzymes is the conversion (degradation) of l-Glu (l-1) to ␣-ketoglutarate (2). Among about 30 catabolic l-GluDHs in literature we priorized the enzymes of F. nucleatum and C. difficile. These GluDHs are reported to have a high specific activity of about 32YU/mg (GluDH from F. nucleatum) [12] and 146YU/mg (GluDH from C. difficile) [11] in comparison to GluDHs from other sources like such as Peptostreptococcus asaccharolyticus (7.8YU/mg) [15], Bacteroides fragilis (7.3YU/mg) [16], Pyrobacterium islandicum (3.5YU/mg) [17], and Janthinobacterium lividum (3.1YU/mg) [18]. Recombinant expression and production of sufficient amounts of both biocatalyst were conducted applying an E. coli expression system as mentioned above. Both enzymes, namely the GluDHs from C. difficile and F. nucleatum, were successfully expressed in E. coli as a soluble and fully active protein. The specific enzyme activities in the crude extracts of 6 and 8YU/mg of protein for the GluDHs from F. nucleatum and C. difficile, respectively, could be increased nearly threefold up to 16 and 20YU/mg after heat precipitation for the recombinant l-GluDHs as summarized in Table 1. From 4 g of cells (wet weight) we obtained 1.580YU of the recombinant l-GluDH from F. nucleatum and 1.390YU of the recombinant l-GluDH from C. difficile (corresponding to yields of 76% and 88%, respectively referred to activities of the crude extracts). The
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Table 2 Biotransformation of sodium glutamate (l-1, 15 mM) into 2 with a l-GluDH and [Fe(III)TSPP]Cl (3, 1 mM) or NADH-oxidase from Lactobacillus brevis at 20 ◦ C and pH 8
. Entry
l-GluDH [U/mL]
NOX[U/mL]
Conv.a 2 h [%]
Conv.a 4 h [%]
Conv.a 8 h [%]
Conv.a 24 h [%]
1 2 3
1.5 5.0 1.5
– – 4.5
51(50) 49(51) 36(37)
65(64) 77(60) 60(60)
76(78) 91(89) 77(76)
86(66) >99(88) 82(77)
a The conversion is defined as the amount of consumed substrate 1 related to the original amount of substrate 1 (in parentheses the amount of generated product 2 related to the original amount of substrate 1 (in %) is given).
biochemical data in terms of enzyme activities of the two recombinant l-GluDHs are listed in Table 1. In addition, our expectation that such recombinant catabolic GluDHs show product inhibition to a less extend than above mentioned commercially l-GluDHs has been fulfilled by the two prioritized enzymes. Product inhibition caused by 50 mM of 2 was in the range of 30% for the l-GluDH from C. difficile and around 55% for the l-GluDH from F. nucleatum at pH 9.0, thus leading to residual enzyme activities of 70% and around 45%, respectively (Fig. 1). The lowered activities of both enzymes are caused by the addition of ␣-ketoglutarate, not by an instability of the enzymes themselves because stability studies showed that no deactivation at all could be observed after a period of 24 h. In comparison to the published data obtained by l-GluDH from C. symbiosus the results clearly indicate a significantly lowered product inhibition level, even at higher concentrations in the range of 50 mM. With the two recombinant l-GluDHs in hand, we next focused on the desired preparative enzymatic synthesis of ␣-ketoglutarate (2). Initial work addressed the set up of a robust and reliable analytical method for determination of the conversion. Due to the known potential of ␣-keto acids to decompose, determination of conversion (related to the formation of 2) through the absolute amount of ␣-ketoglutarate (2) via comparison with an internal standard appeared to us more attractive compared to calculation of conversion based on a relative comparison of substrate/product ratio. As an analytical method we chose proton NMR spectroscopy using pivalic acid (trimethylacetic acid) as an internal standard in combination with a presaturation technique (to improve the
Fig. 1. Product inhibition of the l-GluDHs from C. difficile (GluDH CD) and F. nucleatum (GluDH FN) by ␣-ketoglutarate (2) (assay: 50 mM l-1; pH 9.0; 50 mM 2; 2.5 mM NAD+ ).
quality of the baseline of the spectra due to suppression of the large signal resulting from the solvent water) [19]. Next we studied the stability of ␣-ketoglutarate (2) under reaction conditions starting from an ␣-ketoglutarate (2, 10 mM)/l-glutamate mixture (l-1, 40 mM) which simulates a 20% conversion of an enzymecatalyzed oxidation of 50 mM L-glutamate. After optimization of sampling and work-up for analytical determination (showing a deviation in the range of ca. 3%), we found high recovery rates of 97% or more for ␣-ketoglutarate (2) within a time range of 48 h (Fig. 2). Notably, recovery rates were high not only for the ␣-ketoglutarate/l-glutamate mixture but also in the presence of a crude extract of the recombinant l-GluDH from C. difficile. However, when using a crude extract of the l-GluDH from F. nucleatum we observed a significantly higher loss of 2 probably caused by an ␣-ketoglutarate decomposing enzyme present in the partially purified sample of the L-GluDH from F. nucleatum. In general stability of the samples appeared to be high at pH 8.0 as well as at 9.6. Selected examples of this stability study are given in Fig. 2. Next we conducted biotransformations with in situ-cofactor regeneration based on the use of the iron(III)porphyrin [Fe(III)TSPP]Cl (3) as an artificial NADH-oxidase according to Scheme 1. A first proof of concept for the compatibility of glutamate dehydrogenases with the iron(III)porphyrin complex 3 has been demonstrated when conducting the process at pH 8.0 and with a substrate concentration of 15 mM (Table 2). High conversions
Fig. 2. Time-dependent stability of ␣-ketoglutarate (2) determined out of a mixture of ␣-ketoglutarate (2, 10 mM) and l-glutamate (1, 40 mM) (aqueous solution) at pH 8.0 or 9.6. Additionally, crude extracts of GluDH from C. difficile (C.d.) or from F. nucleatum (F.n.) were added to the ␣-ketoglutarate/glutamate mixture.
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Table 3 Biotransformation of l-glutamate (l-1, 50 mM) into ␣-ketoglutarate (2) with l-GluDH and Fe(III)TSPP (3, 1 mM) at pH 8 (according to Scheme 1). Entry
l-GluDH
l-GluDH [U/mL]
Temp. [◦ C]
Conv.a 1 h [%]
Conv.a 4 h [%]
Conv.a 24 h [%]
1 2 3
C. difficile F. nucleatum F. nucleatum
5 5 5
20 20 12
22(22) 5(5) 8(8)
33(33) 9(8) 11(11)
47(43)b 16(5) 28(27)c
a The conversion is defined as the amount of consumed substrate 1 related to the original amount of substrate 1 (in parentheses the amount of generated product 2 related to the original amount of substrate 1 (in %) is given). b Maximum at 49% after 48 h. c Maximum at 33% after 48 h.
were obtained in particular in the presence of the l-GluDH from C. difficile (entries 1 and 2). For example, after optimization and a reaction time of 8 h a conversion of 91% (based on consumed substrate l-1) as well as a product-related conversion of 89% (based on the amount of formed 2, entry 2) were observed, whereas using a reduced amount of the enzyme resulted in a somewhat decreased conversion of 76% (entry 1). The total turnover numbers for the cofactor NAD+ calculated from entries 1 and 2 (Table 2) are within the range of 6.5–7.4. Furthermore, we carried out experiments with NADH-oxidase (NOX) from Lactobacillus brevis in order to compare the “artificial NADH oxidase” (iron(III)porphyrin [Fe(III)TSPP]Cl (3)) with the enzyme NADH oxidase (Table 2; entry 3). Unfortunately, the activity of the enzyme NADH oxidase is relatively low at higher pH values, therefore an excess of activity was added to obtain complete regeneration of NAD+ . Table 2 shows that, in terms of efficiency, the conversion rates of the biomimetic [Fe(III)TSPP]Cl (3) are in the same order of magnitude as the enzyme NADH oxidase. Subsequently, we increased the substrate concentration of l-1 to 50 mM and operated at a pH-value of 8 which turned out to be the preferred pH in terms of product stability. When using the lGluDH from C. difficile we gained after optimization a conversion of 47% at room temperature (Table 3, entry 1). In the case of l-GluDH from F. nucleatum we faced the problem of an immense loss of 2 and low conversions (entry 2). In order to suppress chemical degradation of ␣-ketoglutarate (2), experiments were done at a somewhat decreased temperature of 12 ◦ C. When using the l-GluDH from F. nucleatum as a biocatalyst at this reaction temperature, a product-related conversion of ␣-ketoglutarate (2) (often defined as so-called “reaction yield”) of 27% was found after 24 h (Table 3, entry 3). Further experiments under different conditions like varying the pH value, more intensive cooling during the reaction or stepwise addition of the substrate l-1 did not show higher conversions than those presented in Table 3. Moreover the cooling was the only way to prevent the chemical degradation of ␣-ketoglutarate on prolonged reaction times, especially when the GluDH from F. nucleatum was used.
volumetric productivity of 3.95 g/(L d) compared to the experiments at 15 mM). In addition, this productivity is almost four times higher as the one reported previously in literature when using an NADH-oxidase [8]. Thus, the iron complex [Fe(III)TSPP]Cl turned out to be capable to be used for cofactor regeneration of the cofactor NAD+ for enzymatic amino acid oxidation. The regeneration of NAD+ is a prerequisite if NAD+ -dependent dehydrogenases are designated for oxidation reactions, for example, converting hydroxy acids, amino acids or alcohols into the corresponding keto compounds. Only a few enzymatic methods are described in the literature to achieve this aim, but all of them present also disadvantages: the use of glutamate dehydrogenase [20,21], lactate dehydrogenase [22], or horse liver alcohol dehydrogenase [23] always leads to the formation of by-products, thus complicating the isolation of the desired product. The reactions catalyzed by these enzymes are reversible therefore it is difficult to reach a complete conversion regarding the primary desired oxidizing reaction. Peroxide forming NADH oxidases require the addition of catalase. Furthermore, all enzymatic systems depend typically on one coenzyme species, either NADH or NADPH, which hampers the development of a generalizable reaction platform. Complementing enzymes as catalysts to regenerate the oxidized coenzyme, the iron complex [Fe(III)TSPP]Cl can be used successfully for this purpose. In addition, it can be stored over a long period of time and it accepts NADH as well as NADPH. Furthermore, peroxide is not formed by the reaction catalyzed by this complex. Currently, further extension of this type of “artificial cofactor regeneration system” by means of the [Fe(III)TSPP]Cl (3) is in progress as well as studies with respect to the clarification of the reaction mechanism. Acknowledgements The authors thank the German Federal Environmental Foundation (Deutsche Bundesstiftung Umwelt, DBU) for generous support within the DBU-network “ChemBioTec” of the funding priority “Biotechnology” (Project AZ 13234-32). We further thank. Dr. Philipp Böhm for helpful discussions and technical support.
4. Conclusion
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In conclusion, we reported the use of an artificial in situ-cofactor regeneration with [Fe(III)TSPP]Cl (3) for enzymatic amino acid oxidation, exemplified for the l-glutamate dehydrogenase-catalyzed synthesis of ␣-ketoglutarate (2) from l-glutamate (l-1). In comparison of the two investigated dehydrogenases, the l-glutamate dehydrogenase from C. difficile turned out to be the preferred enzyme. At a substrate concentration of 15 mM of l-glutamate (l-1) in situ-cofactor regeneration using [Fe(III)TSPP]Cl (3) as an “artificial NADH-oxidase” proceeded smoothly, leading to up to >99% overall conversion and 88% conversion related to the formation of desired product 2 after 24 h. At an increased concentration of 50 mM of substrate l-1, a somewhat decreased conversion of 43% was observed (which, however, corresponds to a nearly doubled
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