Engineering NAD+ availability for Escherichia coli whole-cell biocatalysis: a case study for dihydroxyacetone production
© Zhou et al.; licensee BioMed Central Ltd. 2013
Received: 10 July 2013
Accepted: 5 November 2013
Published: 9 November 2013
Whole-cell redox biocatalysis has been intensively explored for the production of valuable compounds because excellent selectivity is routinely achieved. Although the cellular cofactor level, redox state and the corresponding enzymatic activity are expected to have major effects on the performance of the biocatalysts, our ability remains limited to predict the outcome upon variation of those factors as well as the relationship among them.
In order to investigate the effects of cofactor availability on whole-cell redox biocatalysis, we devised recombinant Escherichia coli strains for the production of dihydroxyacetone (DHA) catalyzed by the NAD+-dependent glycerol dehydrogenase (GldA). In this model system, a water-forming NAD+ oxidase (NOX) and a NAD+ transporter (NTT4) were also co-expressed for cofactor regeneration and extracellular NAD+ uptake, respectively. We found that cellular cofactor level, NAD+/NADH ratio and NOX activity were not only strain-dependent, but also growth condition-dependent, leading to significant differences in specific DHA titer among different whole-cell biocatalysts. The host E. coli DH5α had the highest DHA specific titer of 0.81 g/gDCW with the highest NAD+/NADH ratio of 6.7 and NOX activity of 3900 U. The biocatalyst had a higher activity when induced with IPTG at 37°C for 8 h compared with those at 30°C for 8 h and 18 h. When cells were transformed with the ntt4 gene, feeding NAD+ during the cell culture stage increased cellular NAD(H) level by 1.44 fold and DHA specific titer by 1.58 fold to 2.13 g/gDCW. Supplementing NAD+ during the biotransformation stage was also beneficial to cellular NAD(H) level and DHA production, and the highest DHA productivity reached 0.76 g/gDCW/h. Cellular NAD(H) level, NAD+/NADH ratio, and NOX and GldA activity dropped over time during the biotransformation process.
High NAD+/NADH ratio driving by NOX was very important for DHA production. Once cofactor was efficiently cycled, high cellular NAD(H) level was also beneficial for whole-cell redox biocatalysis. Our results indicated that NAD+ transporter could be applied to manipulate redox cofactor level for biocatalysis. Moreover, we suggested that genetically designed redox transformation should be carefully profiled for further optimizing whole-cell biocatalysis.
Cofactor-dependent redox biocatalysis has been shown as a powerful strategy for the production of valuable chemicals that are otherwise difficult to be synthesized [1, 2]. Whole cells are preferred for industrial application because cofactors can be regenerated more efficiently . To drive the redox chemistry to a specified direction, it is essential to manipulate intracellular redox state as well as cofactor levels . Thus, various strategies have been applied to control the cofactor regeneration system or balance the enzyme activities of redox reactions. For example, H2O-forming NADH oxidase (NOX) has been applied for cofactor regeneration by engineered whole-cell biocatalyst for chiral compound production . The intracellular cofactor concentration was also important to attain high efficiency especially in the case that the redox enzymes had high apparent Km values to the cofactor . Previous reports showed that exogenous supplied cofactors could improve the reaction rates under whole-cell catalysis conditions, although cells were permeated and external cofactor concentrations were applied at concentrations of over 0.5 mM [6–9]. We recently found that the nucleotide transporter NTT4 encoded by the ntt4 gene from the chlamydial endosymbiont Protochlamydia amoebophila UWE25  could enable Escherichia coli cells to uptake NAD(H) from the culture broth . The NAD+ auxotrophic E. coli YJE003 cells expressing NTT4 cultivated in the media containing 40 μM NAD+ could realize the intracellular NAD(H) pool of 5.1 mM, which was 5.8-fold more than that of the wild-type cells . We reasoned that such a unique NAD(H) supplementation system could be further explored to drive cellular redox chemistry.
Construction of plasmids for DHA production
As no discernible DHA was observed when GldA alone was overexpressed in E. coli whole cells (date not shown), we introduced NOX as well as the NAD+ transporter NTT4 to increase NAD+ availability. Thus, two plasmids were constructed for DHA production from glycerol (Figure 1). For the plasmid pTrc99A-gldA-nox, the gldA gene from E. coli and the nox gene from Enterococcus faecalis were cloned into the vector pTrc99A with ribosome binding sites (RBS), under the control of the Trc promoter and the lacI repressor (Figure 1B). An rrnbT transcription terminator was located downstream of nox. For the plasmid pTrc99A-gldA-nox + ntt4, the NTT4 expression cassette in which the ntt4 gene from P. amoebophila UWE25 was regulated by the promoter gntT105 P was constructed, and inserted into the pTrc99A-gldA-nox backbone downstream of the rrnBT terminator using the RF cloning strategy . Both plasmids ensured the expression of GldA for the oxidation of glycerol to DHA and NOX for NAD+ regeneration. In the case of pTrc99A-gldA-nox + ntt4, NTT4 was expressed to enable NAD+ uptake.
Strain dependence of DHA production
Effects of cell growth on DHA production
Promoter selection for NAD+ transporter expression
Intracellular NAD(H) levels of E. coli DH5α cells a harboring NTT4 expression and corresponding empty plasmids
Cellular NAD(H) level (mM)b
1.72 ± 0.03
1.85 ± 0.01
1.96 ± 0.00
2.13 ± 0.01
1.86 ± 0.05
4.47 ± 0.14
Enhancing NAD+ supply for DHA production by using a NAD+ transporter NTT4
DHA production under shake-flask conditions
Cellular redox state, roughly indicated by the NAD+/NADH ratio, is important for whole-cell redox biocatalysis. In E. coli, NAD+/NADH ratios ranging from 3 to 10 were previously documented . It was expected that cellular NAD+/NADH ratio as well as NAD(H) level should have major effects on whole-cell biocatalysis. And co-expressing the nox gene encoding a H2O-forming NOX with GldA is expected to be helpful for reversing GldA activity toward DHA biosynthesis by efficient regenerating NAD+. Indeed, NOX has been widely used to enhance NAD+-dependent biosynthesis in both resting cells  and growing cells . In this study, we found NOX activity was not only strain-dependent (Figure 2B), but also growth condition-dependent (Figure 3B), though cells were transformed with the same plasmid pTrc99A-gldA-nox. This may be attributed to the adaption ability difference against NOX expression among these strains because high NOX activity retarded cell growth and affected the metabolism . Indeed, we also found that cell growth was negatively correlated with NOX activity (date not shown). Thus, while NOX expression provided driving force to maintain a higher NAD+ level and cellular NAD+/NADH ratio, it should be carefully tuned to avoid toxic effects on cellular physiology. It was found that DHA specific titer was positively correlated with the cellular NAD+/NADH ratio but not the cellular NAD(H) level (Figure 2). Similarly, introducing robust cofactor regeneration system increased the whole-cell biocatalysts efficiency [19, 27]. However, enhancing the cellular NADP(H) pool alone without efficient cofactor regeneration failed to improve the redox biocatalyst efficiency . Together, NAD+/NADH ratio driven by a cofactor regeneration system is more important than cellular cofactor level for an efficient oxidative bioreaction.
Once cofactor was efficiently regenerated, cellular cofactor level could also affect the biocatalyst efficiency . External cofactor supplementation has been applied to increase intracellular NAD+ levels and NAD+/NADH ratios [6–8]. In those cases, cells were treated with permeating agents and external cofactor concentrations were high. We recently showed that NTT4 expression in E. coli led to uptaking NAD(H) from the culture broth . Thus, ntt4 was used to improve E. coli whole-cell biocatalytic efficiency by enhancing the cellular NAD(H) level. Actually, ntt4 expression in YJE006 increased cellular NAD(H) level by 1.44 fold, indicating that NTT4 was functional in terms of uptaking external NAD+. As NTT4 could transport NADH efficiently  and increase the cellular reduction state with external NADH supplementation , it could also be used for increase the cellular NADH level for reduction biocatalyst such as asymmetric reduction of o-chloroacetophenone .
Although NTT4 expression strain YJE006 had a higher DHA production, it appeared to lose enzyme activities and cellular cofactor much more rapidly than YJE005. One possible reason was that NTT4 expression led to increased membrane permeability and other toxic effects. Therefore, it seemed challenging to maintain good NAD(H) durability for NTT4 expressions cells that were capable of uptaking external NAD+. As it requires alive cells to mediate continuous NAD+ transportation, NTT4 may have better potential profiting for NAD+-dependent multi-step biosynthesis in living cells such as 1-butanol production from glucose . Although the NAD+ feeding strategy is cost prohibitive for bulk chemicals production, it may be useful for the production of high-value chemicals and pharmaceuticals. Alternatively, reconstruction of efficient heterologous NAD+ biosynthesis pathway  may increase cellular NAD(H) level for enhanced NAD(H)-dependent biosynthesis.
The highest specific DHA productivity reached 0.76 g/gDCW/h under shake-flask conditions for the first 2 h (Figure 5A), which was lower compared with other studies of more than 2 g/gDCW/h using G. oxydans as the hosts in small scale bioreactor , but much higher than that recombinant Saccharomyces cerevisiae of 0.03 g/gDCW/h from sugar . The high DHA productivity of G. oxydans is attributed to the membrane-bound GDH, which can directly oxidize glycerol to DHA without material transfer across cell membrane . However, our attempt, in reversing an endogenous cytosolic GldA catalysis for DHA production in recombinant E. coli by engineering NAD+ availability, provided some insights on optimizing whole-cell redox biocatalyst for other valuable chemicals and pharmaceuticals production.
As DHA concentration dropped over time after 2 h, it was possible that other metabolic steps consumed DHA. In this regard, disruption of the DHA and glycerol catabolic pathway related genes such as dhaK and glpK be useful (Figure 1A). Interestingly, external NAD+ supplementation during the whole-cell catalysis ensured a higher NOX activity and continuous accumulation of DHA for up to 10 h. These results indicated that NOX activity was beneficial to maintain oxidative ability of the NAD+-dependent whole-cell biocatalysis. It is worth mentioning that the initial specific biocatalyst activity (141 U/gDCW) was half to the initial GldA activity of 249 U/gDCW. This might be attributed to the substrate diffusion, product consumption, etc.
In summary, using oxidation of glycerol to DHA by recombinant E. coli whole-cells as a model system, we enhanced oxidation state and cellular cofactor level for increasing the catalytic efficiency by expressing NADH oxidase and NAD(H) transporter, respectively. As the overall biocatalytic performance is dependent upon the cellular cofactor level, redox state and the corresponding enzymatic activity, genetically designed redox transformation should be systematically profiled to identify optimal whole-cell biocatalysis.
Material and methods
Bacterial strains and plasmids
Strains and plasmids used in this study
Strains or plasmids
Genotype or characteristic
Resources or references
E. coli Strains
F-, φ80d/lacZ∆M15, ∆(lacZYA-argF)U169, deoR, recA1, endA1, hsdR17(rk-, mk+), phoA, supE44, λ-, thi-1, gyrA96, relA1
F-,mcr A, ∆(mrr-hsd RMS-mcr BC), φ80lac Z∆M15, ∆lac X74, rec A1, end A1, ara D139, ∆ (ara, leu)7697, gal U, gal K, λ-, rps L, nup G/pMON14272/pMON7124
F-, glnV44(AS), λ - , rfbC1, gyrA96(NalR), recA1, endA1, thi-1, hsdR17
F-, λ - , rph-1
rrnB3, ∆lacZ4787, hsdR514, ∆(araBAD)567, ∆(rhaBAD)568 rph-1
F–, dcm, ompT, hsdS(rB–, mB–), gal, λ(DE3)
DH5α/pTrc99A-gldA-nox + ntt4
lacZ, pBR322 ori, bla, cloning vector
lacI, pBR322 ori, bla, expression vector
ntt4 inserted within Nde I and Bam H I sites, kan
ntt4 inserted within Sac I and Bam H I sites, kan
ntt4 inserted within Sac I and Bam H I sites, kan
gldA and nox transcription under Trc promoter
pTrc99A-gldA-nox + ntt4
gldA and nox transcription under Trc promoter, ntt4 under gntT105p promoter.
Primers used in this study a
Promoter gapAP1 amplification
ACCGAATTC GATCTCATATGTTCCA CCAGCTATTTGTTAG
GGAATTC TATCTCCT TATTCATTTGCGCTGGGTAACGTCAATTT
gntT105P + ntt4 amplification
Terminator BBa_B0015 amplification
The gldA (NCBI GeneID: 6058353) gene was amplified from E. coli DH5α genomic DNA using primer pair gldA-F0/gldA-R0 and cloned into pMD18T to give pMD18T-gldA. The nox (NCBI GeneID: 1200486) gene was amplified from pMD18T-nox, which was constructed by cloning the nox gene from Enterococcus faecalis (CGMCC 1.130) into the pMD18T. The gldA-nox co-expression cassette was constructed with the modified one-step overlap extension (SOE) PCR strategy described previously . Briefly, gldA and nox were amplified with primer pairs gldA-F1/gldA-R1 and nox-F1/nox-R1, and purified gldA and nox fragments (molar ratio 1:1, about 200 ng each) were mixed. To the mixture were added 3 μL of dNTP (2.5 mM each), 5 μL 5 × PrimerStar buffer, 1.25 U PrimeSTAR HS DNA polymerase, and H2O to a total volume of 25 μL. PCR amplification was performed according to the thermocycle conditions of 95°C for 5 min, 15 cycles of 98°C for 10 s, 68°C for 3 min, and 68°C for 10 min. Next, 2 μL of unpurified PCR products was used as the template using the primer pair gldA-F1/nox-R1 for normal PCR amplification in a total volume of 100 μL. The purified gldA-nox cassette was cloned into the pTrc99A using the restriction-free (RF) cloning strategy  to give plasmid pTrc99A-gldA-nox.
The ntt4 (NCBI GeneID: 2780098) gene containing the 3'-end 6 × His-tag encoding sequence was cloned from the vector pET15k-ntt4. The constitutive glyceraldehyde-3-phosphate dehydrogenase promoter P1 gapA P1  and the internal operator of gluconate transporter promoter 105 mutant gntT105 P  was cloned from E. coli DH5α genomic DNA using primer pairs gapAP1-F/gapAP1-R and gntT105P-F1/gntT105P-R1, respectively. Then, Eco R V-Eco R I digested promoters were cloned into the Eco R V-Eco R I site of pTrc99A to substitute the Trc promoter, resulting in the constitutive expression vectors pBCTA and pBCTB. The function of these two vectors was checked by constitutive expression of red fluorescent protein (date not shown). Lastly, ntt4 was cloned into the Sac I-Bam H I site locating downstream of the constitutive promoter of pBCTA and pBCTB after cloning with the primer pair ntt4-F1/ntt4-R1, and the bla was replaced by the kan using a RF cloning strategy, resulting in plasmids pBCTC-ntt4 and pBCTD-ntt4, respectively.
The ntt4 constitutive expression cassette was also constructed and cloned into pTrc99A-gldA-nox. The gntT105P-ntt4 was amplified from the pBCTD-ntt4 using the primer pair gntT105P-F2/ntt4-R2 and terminator B0015 cloned using the primer pair T-F1/T-R1 from the international genetically engineered machine competition (IGEM, http://partsregistry.org/Part:BBa_B0015). These two DNA fragments were fused with a modified SOE PCR approach (33), and cloned into pTrc99A-gldA-nox locating downstream of the rrnBT terminator, resulting in plasmid pTrc99A-gldA-nox + ntt4.
Whole-cell biocatalyst preparation
Recombinant E. coli cells harboring appropriate plasmid were cultivated in LB medium supplemented with appropriate antibiotics at 37°C, 200 rpm, to the early exponential phase (OD600 = 0.2–0.4). Cultures were induced by adding IPTG to a final concentration of 0.1 mM (and 0.2 mM NAD+ if needed), and cultivated for additional 8 h or 18 h at appropriate temperature (37°C or 30°C), 200 rpm. Cells were harvested by centrifugation (2,000 g, 5 min) and washed twice with 0.1 M potassium phosphate buffer (pH 9.0).
As GldA had a higher activity toward glycerol dehydrogenation  and our previous study  showed DHA production reached the highest at pH 9.0. The IPTG induced E. coli cells were resuspended in 5 mL of 0.1 M potassium phosphate buffer (pH 9.0) for 10 h in 5 mL of potassium phosphate buffer containing 20 g/L glycerol in 50-mL test tubes; or in 20 mL of the buffer containing 2–5 g/L glycerol in 500-mL shake flasks. NAD+ was added into the reaction to a final concentration of 0.2 mM when necessary. All reactions were performed at 37°C, 200 rpm. All the data represent the averages standard deviations from at least three independent samples.
DHA was assayed according to a known method with minor modifications . Briefly, biotransformation mixtures were centrifuged at 10,000 g for 2 min. Exactly 20 μL of supernatants were mixed with 180 μL diphenylamine reagent containing 1% (w/v) diphenylamine and 10% (v/v) sulfuric acid in acetic acid, and heated at 105°C for 20 min. Then, the absorbance at 620 nm were recorded after cooling to room temperature, and DHA concentrations were quantified according to a standard curve obtained under identical conditions.
Enzyme activity assay
E. coli dry cell weight (DCW) was weight by converting OD600 value with a coefficient of 0.275 gDCW/(L × OD600), which was determined by freezer drying the E. coli cells according to our recently report . As the DHA production was performed at 0.1 M potassium phosphate buffer (pH 9.0) due to GldA activity has higher activity as mentioned above, all enzymatic assays were performed at consistent pH of 9.0. About 2 × 109E. coli cells were harvested, washed twice with 0.1 M potassium phosphate buffer (pH 9.0), and stored as cell pellets at -80°C. For enzyme assays, cell pellets were resuspended in 0.2 mL of lysis buffer (10 mM Tris-Cl, 1.0 mM MgCl2, 1 mg/mL lysozyme and 0.1 mg/mL DNase, pH 8.0) and incubated at 37°C for 30 min. GldA activity was estimated by recording the absorbance increase at 340 nm and assays were performed at 25°C, in 100 μL of 0.1 M potassium carbonate buffer (pH 9.0) containing 5 mM NAD+, 100 mM glycerol and 5 μL of crude cell lysates, which was with minor modifications from a previous report . NOX activity was measured by recording the absorbance decrease at 340 nm and assays were performed at 25°C, in 100 μL of 0.1 M potassium phosphate buffer (pH 9.0) containing 0.4 mM NADH and 1 μL (if the cell NOX activity was more than 1000 U/gDCW) or 5 μL of (if the cell NOX activity was less than 1000 U/gDCW) crude cell lysates.
Cell pellets (containing about 2 × 109 cells) were washed twice with 0.1 M potassium phosphate buffer, and then treated at 55°C for 10 min in 150 μl of 0.2 M NaOH (for NADH extraction) or 150 μl of 0.2 M HCl (for NAD+ extraction). The extracts were neutralized by adding 150 μl of 0.1 M HCl (for NADH extraction) or 150 μl of 0.1 M NaOH (for NAD+ extraction). The cellular debris was removed by centrifuging at 12,000 g for 5 min. Supernatants were transferred to new tubes and stored at -80°C until assay. NAD(H) was quantified using a sensitive enzymatic cycling assay as reported previously .
We are indebted to Prof. Qin Ye (East China University of Science and Technology, China) for kindly providing pTrc99A. This work was supported by the National Basic Research and Development Program of China (No. 2012CB721103) and the State Key Laboratory of Catalysis, Dalian Institute of Chemical Physics, CAS (R201306).
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