Determining the Extremes of the Cellular NAD(H) Level by Using an Escherichia coli NAD+-Auxotrophic Mutant
ABSTRACT
NAD (NAD+) and its reduced form (NADH) are omnipresent cofactors in biological systems. However, it is difficult to determine the extremes of the cellular NAD(H) level in live cells because the NAD+ level is tightly controlled by a biosynthesis regulation mechanism. Here, we developed a strategy to determine the extreme NAD(H) levels in Escherichia coli cells that were genetically engineered to be NAD+ auxotrophic. First, we expressed the ntt4 gene encoding the NAD(H) transporter in the E. coli mutant YJE001, which had a deletion of the nadC gene responsible for NAD+ de novo biosynthesis, and we showed NTT4 conferred on the mutant strain better growth in the presence of exogenous NAD+. We then constructed the NAD+-auxotrophic mutant YJE003 by disrupting the essential gene nadE, which is responsible for the last step of NAD+ biosynthesis in cells harboring the ntt4 gene. The minimal NAD+ level was determined in M9 medium in proliferating YJE003 cells that were preloaded with NAD+, while the maximal NAD(H) level was determined by exposing the cells to high concentrations of exogenous NAD(H). Compared with supplementation of NADH, cells grew faster and had a higher intracellular NAD(H) level when NAD+ was fed. The intracellular NAD(H) level increased with the increase of exogenous NAD+ concentration, until it reached a plateau. Thus, a minimal NAD(H) level of 0.039 mM and a maximum of 8.49 mM were determined, which were 0.044× and 9.6× those of wild-type cells, respectively. Finally, the potential application of this strategy in biotechnology is briefly discussed.
INTRODUCTION
NAD (NAD+; its reduced state is NADH) plays an essential role in cellular metabolism due to its function as a cofactor in over 300 redox reactions (10). Thus, the NAD+ level and the NADH/NAD+ ratio determine the metabolic fluxes of many pathways as well as the transcriptional level of many genes in vivo (13, 14). For example, a high NADH/NAD+ ratio triggers overflow metabolism (33), and a low NADH/NAD+ ratio enhances the glycolytic flux (38). Manipulating the NADH/NAD+ ratio is considered one of the most effective strategies for improving the productivity of some products (19, 23). There are two metabolic pathways for NAD+ biosynthesis in Escherichia coli, the de novo pathway and the salvage pathway (3, 15). The de novo pathway starts with the oxidation of aspartate to iminosuccinic acid, which is catalyzed by NadB and in turn reacts with dihydroxyacetone phosphate to give quinolinic acid (QA), followed by phosphoribosylation and decarboxylation, resulting in nicotinic acid mononucleotide (NAMN). The salvage pathway converts several precursors (nicotinic acid [NA], nicotinamide [NM], and nicotinamide mononucleotide [NMN]) to NAMN, which is further converted to NAD+ through adenylation and amidation catalyzed by NadD and NadE, respectively (Fig. 1A). Like most of the precursors, such as NA, NMN can be taken up from the medium (25); however, extracellular NAD+ is degraded to NMN and adenosine in the periplasm before passing through the inner membrane, because NAD+ per se is impermeable to the bacterial cytoplasmic membrane (Fig. 1B) (21). The first NAD+ transporter, NTT4, was discovered in the prokaryote chlamydial endosymbiont Protochlamydia amoebophila UWE25, which could transport intact NAD(H) in counterexchange with ADP (11). Recently, other NAD+ carriers located at chloroplasts or mitochondria have been discovered in the eukaryotes Saccharomyces cerevisiae and Arabidopsis thaliana (20, 28). These transporters import NAD+, in counterexchange with nucleotides (ADP, AMP, or GMP), for cells or organelles that lack the NAD+ biosynthesis capacity to support some biological process (Fig. 1B).
Fig. 1.
The level of intracellular NAD(H) is tightly controlled by a variety of mechanisms, including its biosynthesis and salvage through NadR regulation (10). When the internal NAD(H) level is low, NadR interacts with the integral membrane protein PnuC, which is responsible for the transport of NMN across the cytoplasmic membrane, promoting a net NMN transportation into the cell. When the NAD(H) level increases, NadR binds to the operator regions of the target genes (nadA, nadB, and pncB) and represses their transcription to decrease the biosynthesis. However, several studies have succeeded in enhancing the NAD(H) pool by manipulating the biosynthetic pathway as well as the regulation system. With the overexpression of pncB under the control of the native operator and promoter sequences, the intracellular NAD(H) increased 5-fold when cells were cultured in the presence of 0.1 mM exogenous nicotinic acid (34). The pncB was also mutated in the operator region to protect against the regulatory effect of NadR and overexpressed under the lac promoter, resulting in a 2.2-fold increase of the NAD(H) level (22). Cooverexpression of the pncB and nadE gene led to a more dramatic 7-fold increase of the NAD(H) level (12). The NAD(H) level was also reduced with the nadD mutation, which encoded the NAMN adenylyltransferase (25).
Although the NAD(H) pool can be adjusted by manipulating the biosynthetic pathway, it remains difficult to investigate the effects of extremes of the NAD(H) level on cell growth due to the corresponding self-regulation mechanism. In this study, we manipulated the NAD(H) level by feeding NAD(H) to an NAD+-auxotrophic E. coli mutant that was constructed by disrupting the essential gene nadE, which is responsible for the last step of NAD+ biosynthesis in cells with an NAD+ transporter NTT4 expression background. This led to successful determination of the extremes of the intracellular NAD(H) level. Finally, we discuss the potential application of our results in the fields of biotechnology related to cofactor engineering.
MATERIALS AND METHODS
Bacterial strains and plasmids.
Cloning and plasmid propagation were performed with E. coli DH5α. E. coli BW25113(pKD3, pKD4, pKD46) was kindly provided by B. L. Wanner (Purdue University). E. coli JW0105 was obtained from the KEIO knockout collection and has the E. coli BW25113 background (2). The strains and plasmids we used are listed in Table 1.
Table 1.
| Strain or plasmid | Genotype or characteristic | Resource or reference |
|---|---|---|
| E. coli strains | ||
| DH5α | F− φ80dlacZΔM15 Δ(lacZYA-argF)U169 deoR recA1 endA1 hsdR17(rk− mk+) phoA supE44 λ− thi-1 gyrA96 relA1 | TaKaRa |
| BW25113 | rrnB3 ΔlacZ4787 hsdR514 Δ(araBAD)567 Δ(rhaBAD)568 rph-1 | 8 |
| JW0105 | BW25113/nadC::kan | 2 |
| YJE001 | BW25113/nadC::kan/pET15b-NTT4 | This study |
| YJE002 | BW25113/pKD46/pET15K-NTT4 | This study |
| YJE003 | BW25113/pET15K-NTT4/nadE::cat | This study |
| Plasmids | ||
| pMD18-T | lacZ pBR322 ori bla; cloning vector | TaKaRa |
| pET15b | T7 promoter, lacI pBR322 ori bla | Novagen |
| pET15b-NTT4 | ntt4 inserted at NdeI-BamHI site, NTT4 expression, bla | This study |
| pET15K-NTT4 | ntt4 inserted at NdeI-BamHI site, NTT4 expression, kan | This study |
| pKD3 | oriR6Kγ bla cat rgnB(Ter) | 8 |
| pKD4 | oriR6Kγ bla kan rgnB(Ter) | 8 |
| pKD46 | araBp-gam-bet-exo bla repA101(Ts) oriR101 | 8 |
Growth media and conditions.
E. coli was routinely cultivated with agitation at 37°C or 30°C (for gene disruption) and 200 rpm in either LB broth (10 g tryptone [Oxoid], 5 g yeast extract [Oxoid], 10 g NaCl per liter water), SOB broth (20 g tryptone [Oxoid], 5 g yeast extract [Oxoid], 0.5 g NaCl, 2.5 mM KCl, 1 mM MgCl2, 1 mM MgSO4 per liter water), or M9 minimal medium (6 g Na2HPO4, 3 g KH2PO4, 0.5 g NaCl, 1 g NH4Cl, 1 mM MgSO4, 0.1 mM CaCl2 per liter water) containing 2% glucose, and 1,000× trace metal mix (27 g FeCl3·6H2O, 2 g ZnCl2·4H2O, 2 g CaCl2·2H2O, 2 g Na2MoO4·2H2O, 1.9 g CuSO4·5H2O, 0.5 g H3BO3 per liter water). The strains were inoculated from overnight cultures at 37°C, washed 2 times with M9 medium, and transferred into 50 ml of M9 medium supplemented with NAD(H) to an optical density at 600 nm (OD600) of 0.02. Samples were regularly taken for measurement of the OD600 and NAD(H).
Reagents.
Pfu DNA polymerase was supplied by Tiangen Biotech (Beijing, China). All primers used in this study (Table 2) were purchased from Invitrogen (Shanghai, China), and DNA sequencing was performed by TaKaRa (Dalian, China). A DNA gel purification kit and plasmid extraction kit were purchased from Beyotime (Haimen, China). DpnI was purchased from New England BioLabs (Beijing, China), and PrimeSTAR HS DNA polymerase and all other reagents were purchased from TaKaRa (Dalian, China). All chemicals were purchased from Sigma (Shanghai, China).
Table 2.
| Primer | Sequence (5′-3′)a | Function |
|---|---|---|
| NTT4-F | GAGCATATGAGTAAAACAAACCAG | ntt4 cloning |
| NTT4-R | TACGGATCCTTAGTGATGATGATGATGATGTTTTTTTATAAAAG | |
| Kan-F | TTAAACATGCCAGTGATGCAAAGGTAGTGCAAGAGCTATGACATATACCTGCCGTTCAC | kan cloning |
| Kan-R | TCAGGCGGTCAGTGTATCATCACTCATACTCTGCCCGACACCATGGTCATAGCTGTTTC | |
| nadE-F | GTTAATACCGCCGCTGCACTCAG | nadE cloning |
| nadE-R | TCACCATGAAACAGATACACCCCTG | |
| cat(E)-F | TGTCTTTTCTGTCTGGAGGGGTTCAATGATGGGAATTAGCCATGGTCC | cat amplification |
| cat(E)-R | CAAATTATTACTTTTTCCAGAAATCATCGTGTAGGCTGGAGCTGCTTC |
a
The restriction sites are underlined.
DNA manipulation.
The ntt4 gene (11) was custom synthesized by TaKaRa (Dalian, China) and then cloned into the NdeI-BamHI site of pET15b with the primer pair NTT4-F/NTT4-R, resulting in the NTT4 expression vector pET15b-NTT4. pET15K-NTT4 was constructed by substituting the bla gene with kan amplified with Kan-F/Kan-R, using the restriction-free (RF) cloning strategy (32). The RF cloning experiment was done according to the procedure described below. Briefly, the reaction mixture consisted of plasmid (200 ng), megaprimer Kan (600 ng), 5× PrimeSTAR buffer (10 μl), 2.5 mM deoxynucleoside triphosphates (6 μl), PrimeSTAR HS DNA polymerase (1.25 U) in a total volume of 50 μl. The thermocycling conditions were 95°C for 5 min, 35 cycles of 95°C for 1 min, 55°C for 45 s, and 68°C for 6 min, and a final 68°C for 20 min. The reaction mix was treated with DpnI at 37°C for 4 h to digest the methylated parental plasmid and then transformed into E. coli DH5α competent cells.
nadE with about 200-bp flanking regions were amplified from E. coli BW25113 genomic DNA by using the primer pair nadE-F/nadE-R and then cloned into pMD18-T. Disruption cassettes for nadE were constructed by replacing nadE with cat amplified from pKD3 with the primer pair cat(E)-F/cat(E)-R, resulting in pMD18T-nadE::CAT.
Deletion of nadE was performed as described previously (8) with a minor modification. The nadE disruption cassette nadE::CAT was PCR amplified from pMD18T-nadE::CAT with the prime pair nadE-F/nadE-R and then transformed into E. coli YJE002 with electroporation. The mutants were selected on SOB plates supplemented with kanamycin sulfate (50 μg/ml), chloramphenicol (30 μg/ml) and NAD+ (100 μM).
Cofactor measurements.
Cofactor recycling assay was used to determine intracellular concentrations of NAD(H) and NADP(H) with minor modifications (7, 22). For the assay, about 5 × 109 cells were collected by centrifugation at 8,000 × g for 2 min and the cell pellets were washed twice with ice-cold phosphate-buffered saline (PBS) buffer. Then, 300 μl of 0.2 M NaOH [for NAD(P)H extraction] or HCl [for NAD(P)+ extraction] was added to the cell pellets, which were heated for 10 min at 55°C. The extracts were neutralized by adding 300 μl of 0.1 M HCl [for NAD(P)H extraction] or NaOH [for NAD(P)+ extraction]. The cellular debris was removed by centrifuging at 12,000 × g for 5 min. Supernatants were transferred to new tubes and stored at −20°C for no more than 24 h.
The cycling assay was performed by using a reagent mixture consisting of equal volumes of 1.0 M Bicine buffer (pH 8.0), absolute ethanol [for the NAD(H) assay] or 30 mM glucose-6-phosphate [for the NADP(H) assay], 40 mM EDTA (pH 8.0), 4.2 mM 3-(4,5-dimethyl-2-thiazolyl)-2,5-diphenyl-2H-tetrazolium bromide, twice the volume of 16.6 mM phenazine ethosulfate, and 3 volumes of water, previously incubated at 25°C. The following volumes were added to 1-ml cuvettes: 50 μl neutralized extract, 0.9 ml reagent mixture. For the NAD(H) assay, the reaction was started by adding 50 μl of yeast ADH II (500 U/ml in Bicine buffer). Fifty microliters of yeast glucose-6-phosphate dehydrogenase (10 U/ml) was added to start the NADP(H) assay. The absorbance at 570 nm was recorded for 2 min at 25°C. The volume of a cell is 10−12 ml, and the cell concentrations were calculated based on the assumption that an OD600 of 1.0 corresponds to 109 cells/ml (5); thus, the cell density was correlated to the cell volume, with 1 ml/liter equivalent to an OD600 of 1.
Determination of the minimal NAD(H) level.
The colonies of NAD+-auxotrophic mutant YJE003 were picked into LB medium containing 100 μM NAD+ and cultivated for 12 h with agitation at 37°C, 200 rpm. Then, the cells were inoculated into 100 ml of M9 medium supplemented with 100 μM NAD+ to an OD600 of 0.02 and cultivated to the early exponential phase (OD600, ≈0.8). The cells were collected and washed twice with M9 medium and then inoculated into 50 ml of M9 medium, with the initial OD600 being 0.1, 0.2, or 0.3, respectively, and cultivated until stationary phase, in which the cofactor level was the minimal level for E. coli growth. The cells were collected and washed with ice-cold PBS buffer and used for intracellular NAD(H) measurements.
Determination of the maximal NAD(H) level.
The NAD+-auxotrophic mutant YJE003 was picked into LB medium containing 100 μM NAD+ and cultivated for 12 h with agitation at 37°C, 200 rpm. The cells were collected and washed twice with M9 medium, inoculated into 50 ml of M9 medium supplemented with different concentrations of NAD+, with an initial OD600 of 0.02 and cultivated with agitation at 37°C, 200 rpm to stationary phase. The cells were used for intracellular NAD(H) measurements after washing twice with ice-cold PBS buffer.
RESULTS
Verification of NTT4 function in the ΔnadC mutant.
NAD(H) per se is impermeable to the bacterial cytoplasmic membrane (1, 21), and usually only those intermediates, such as AMP, NMN, and NA, can be directly assimilated. Recently, the first NAD+ transporter, NTT4, was discovered in chlamydial endosymbiont P. amoebophila UWE25 (11). The properties of this transporter were studied by cloning the ntt4 gene into E. coli and analyzing the transportation activity with 32P-labeled nucleotides and cells in the resting state. However, it remains unknown whether this transportation activity of NTT4 is sufficient to support cell growth on exogenous NAD+. We constructed E. coli YJE001, which had the ntt4 gene expressed in E. coli JW0105, a strain that lacks the de novo NAD+ biosynthesis pathway due to nadC disruption (Fig. 2A). It was found that YJE001 grew about 20% faster in the exponential phase in the presence of NAD+ in the medium (Fig. 2B), while the wild-type BW25113 cells were insensitive to exogenous NAD+ (data not shown), suggesting that the NTT4 protein was functional as an NAD+ transporter in E. coli. We surmised that the activity of NTT4 might be able to rescue cells in which NAD+ biosynthesis was completely blocked. To test this conjecture, we decided to disrupt the essential gene nadE, which is involved in the last step of NAD+ biosynthesis.
Fig. 2.
Construction of the NAD+-auxotrophic mutant by nadE disruption.
In order to manipulate the intracellular NAD+ level, we attempted to remove the NAD+ regulation effect by disrupting the essential gene nadE, which is responsible for the last step of NAD+ biosynthesis. However, this was impractical if cells did not have access to exogenous NAD+ (2). As the ΔnadC mutant carrying the ntt4 gene showed an advantageous growth profile in the presence of exogenous NAD+, we thought it might be possible to construct an NAD+-auxotrophic mutant if an NTT4 expression background were generated (Fig. 3A). Thus, we expressed the ntt4 gene in E. coli BW25113 and obtained disruption of the nadE gene to obtain the NAD+-auxotrophic mutant YJE003. The mutant strain grew well on LB plates supplemented with either NAD+ or NADH but failed to grow on plates with NAD+ intermediates, such as NA or NMN (Fig. 3B).
Fig. 3.
Comparison of the cofactor levels between exogenous NAD+- and NADH-fed cells.
A previous study showed that NTT4 could transport both NAD+ and NADH and that the NADH uptake rate was a little lower in resting cells (11), but in growing cells the NTT4 transportation activity differences between NAD+ and NADH remained undetermined. Therefore, we examined the effects of the concentration of exogenous NAD+ or NADH on the intracellular cofactor levels as well as the cell growth profile. It turned out that cells grew faster when fed NAD+, instead of NADH. In cells fed NAD+, total intracellular NAD(H) levels were 4.82 mM at 14 h and 7.55 mM at 26 h, which were much higher than levels in those fed the same amount of NADH (Table 3). These results suggested that NTT4 prefers NAD+ over NADH, which was consistent with the results observed using resting cells (11). It was found that the NAD+/NADH ratio was higher in the NAD+-fed cells than the NADH-fed cells, indicating that NAD+-fed cells had a higher oxidative state.
Table 3.
| Feed | Duration (h) of feeding | Intracellular levela (mM) | NAD+/NADH ratio | ||
|---|---|---|---|---|---|
| NAD+ | NADH | NAD(H) | |||
| NAD+ | 14 | 4.26 ± 0.55 | 0.56 ± 0.09 | 4.82 ± 0.62 | 7.66 |
| 26 | 6.23 ± 0.13 | 1.32 ± 0.10 | 7.55 ± 0.17 | 4.72 | |
| NADH | 14 | 2.21 ± 0.21 | 0.41 ± 0.12 | 2.62 ± 0.32 | 5.33 |
| 26 | 2.46 ± 0.06 | 0.68 ± 0.04 | 3.14 ± 0.09 | 3.63 | |
a
Values are means ± standard deviations.
Determination of the minimal cofactor level for E. coli cell proliferation.
Many studies have successfully perturbed the intracellular cofactor level, especially for increasing the NAD+ level (34) and manipulation of the NAD+/NADH ratio (22), but it was difficult to determine the minimal cofactor level for cell proliferation. This is because the NAD+ level is regulated by NAD+ biosynthesis pathways, which are activated to supply NAD+ when the NAD+ level is low. With the NAD+-auxotrophic mutant, the minimal cofactor level for cell growth could be determined based on the NAD+ distribution strategy (Fig. 4A), which was inspired by the plasmid distribution model (31). Briefly, the auxotrophic mutant YJE003 was preloaded with a certain amount of NAD+ and then transferred into minimal medium without NAD+. The cells divided and the cofactor was distributed into daughter cells until the cellular cofactor level reached the minimum that limited cell division. It was interesting that no matter whether the initial OD600 was 0.1, 0.2, or 0.3, the YJE003 cells reached stationary phase after cultivation for about 4 h, and the cell density increased about 4-fold. For example, when the initial OD600 was 0.3, the maximal OD600 was about 1.7 (Fig. 4B), which was much lower than the OD600 of 2.7 obtained when YJE003 cells were cultivated in the presence of 100 μM NAD+ (Fig. 5A). These observations suggested that the cofactor level was indeed limiting cell division.
Fig. 4.
Fig. 5.
As the cell density of YJE003 increased 4-fold in M9 medium without an NAD+ supply, we estimated the total cofactor level should be reduced to less than 20% that of the parent cells. The results of the cofactor measurements were consistent with our expectations. The total NAD(H) and NADP(H) level was 0.12 mM, which was about 20% that in the parent cells (Fig. 4C) and only 10% that of the wild-type BW25113 cells. The NAD(H) level was only 0.04 mM, which was about 4.4% that of the wild-type cells (Fig. 4C and Table 4). The cellular NADP(H) level was 0.08 mM, which was about 50% and 20% that of the parent cells and the wild-type cells, respectively (Fig. 4C). It should be pointed out that the NAD(H) levels of the wild-type cells were consistent with a previous report (9), indicating that our data were accurate.
Table 4.
| Cofactor | NAD(H) level (mM)a | ||
|---|---|---|---|
| Maximal | Minimal | Wild type | |
| NAD+ | 7.50 ± 0.10 | 0.019 ± 0.002 | 0.64 ± 0.04 |
| NADH | 0.99 ± 0.15 | 0.020 ± 0.004 | 0.24 ± 0.01 |
| Total | 8.49 ± 0.24 | 0.039 ± 0.002 | 0.88 ± 0.02 |
a
Data are means ± standard deviations.
Determination of the maximal cofactor level in live E. coli cells.
With the NAD+-auxotrophic mutant, the minimal cofactor was determined with the cell division strategy. However, does the NAD(H) level change in response to the concentration of exogenous NAD+? Is there a maximal NAD(H) level that live E. coli cells can uphold? In order to answer these questions, we cultivated YJE003 cells in the presence of NAD+ at different concentrations (10 μM to 10 mM). The lag phase of the cells decreased with increases in the exogenous NAD+ concentration, but the growth rate of the exponential phase remained the same (5.1 to 5.3 h−1). This suggested that the higher NAD+ transportation rate resulting from a higher exogenous NAD+ concentration was beneficial for the early stage of cell growth but had no advantages for exponential growth (Fig. 5A). The intracellular NAD(H) levels increased with the increase of initial exogenous NAD+ concentration and reached maxima of 7.50 mM NAD+ and 1.51 mM NADH with an initial feeding NAD+ concentration of 1 mM and 60 μM, respectively. The total NAD(H) level reached an extremely high value of 8.49 mM when the initial feeding NAD+ concentration was 1 mM (Fig. 5B), which was 9.6-fold that of the wild-type cells.
DISCUSSION
In a previous study, NTT4 from chlamydial P. amoebophila UWE25 was expressed in E. coli and characterized as an NAD+ transporter by incubating the resting recombinant E. coli cell with 32P-labeled nucleotides (11), but the means to determine the effect of transport of NAD+ in growing cells remained unavailable. In the present study, we first confirmed the function of NTT4 in E. coli by disrupting the nadC gene, which is responsible for the de novo biosynthesis of NAD+ (Fig. 2). We further showed that the NAD+ transportation activity of NTT4 could support cell growth fully, depending on the uptake of extracellular NAD+ (Fig. 3). It is known that disruption of the essential genes nadD and nadE is prohibitive (2, 25), but we were successful as we used an E. coli strain that carried the ntt4 gene (Fig. 3B). Therefore, this strategy could be explored as an alternative for identification of an NAD+ transporter without utilization of radioactive nucleotides. It could also be used to identify and characterize other NAD+ biosynthetic pathways, such as the recently discovered one in Francisella tularensis (24). ATP is an essential energy cofactor that determines the energy metabolism and metabolic flux (17). Several ATP transporters have been identified and heterologously expressed in E. coli (27, 29, 30). The extremes of cellular ATP levels may be determined by using a similar strategy. The impacts of the extreme cofactor levels on the metabolism, transcriptome, and interactions between cofactor and enzyme (35) will be investigated in the future, and these studies should give new insights into microbial cell biology.
The level of NAD is tightly controlled by a variety of mechanisms, including its biosynthesis and salvage. Studies have successfully manipulated the NAD+ level by overexpressing pncB or changing the activity of NAMN adenylyltransferase by constructing an nadD mutant library (25, 26). However, the extremes of the NAD(H) level, i.e., the minimal and maximal level, remain undetermined for live E. coli cells. We constructed an NAD+-auxotrophic mutant in which NAD+ biosynthesis was completely blocked, and we designed an NAD+ distribution strategy to determine the minimal NAD+ level for cell growth (Fig. 4). With the mutant strain YJE003, we showed that the minimal NAD(H) level was 4.4% that of wild-type cells. It has been demonstrated that about 20% total cellular NAD(H) in wild-type cells is freely accessible and the majority (80%) is bound to enzymes (35). The fact that the minimal NAD(H) level in the mutant strain was far lower than the freely accessible NAD(H) level in wild-type cells suggests that the binding affinity between NAD(H) and enzymes should be much higher than observed previously. Moreover, the minimal cellular NADP(H) level was 2-fold that of the minimal NAD(H) level, while the NADP(H) level was lower than the NAD(H) level in wild-type cells (Fig. 4C), indicating that E. coli cells need more NADP(H) for biosynthesis of biomass precursors under a cofactor starvation state, as NADP(H) is the main electron carrier in anabolism (4).
In several studies, the cellular cofactor concentration has been increased through manipulating the NAD+ biosynthetic pathway (12, 16) or feeding permeabilized cells (18, 36, 37). The NAD(H) level was increased up to 6-fold previously (12). Yet, what is the maximal NAD(H) level that live cells can tolerate? To address this fundamental question, we increased the intracellular NAD(H) level by exposing the NAD+-auxotrophic mutant YJE003 cells to higher exogenous NAD+ concentrations in the M9 medium. We observed a 9.6-fold increase in the NAD(H) level compared to that of the wild-type BW25113 cells in the presence of 1 mM exogenous NAD+. The intracellular NAD(H) level did not increase further with exogenous NAD+ concentrations up to 10 mM, indicating that the maximal NAD(H) level in E. coli are around 8.49 mM (Fig. 5B). In wild-type cells, NAD+ biosynthesis is repressed and the salvage for NAD+ degradation is activated when the intracellular NAD(H) level is high. In contrast, NAD+ transportation in the mutant strain YJE003 was likely independent of the biosynthesis regulation mechanism, leading to accumulation of intracellular NAD(H) to a very high level.
Cellular cofactor concentration is important in whole-cell redox biocatalysis (4, 6), which determines whether the biocatalyst operates effectively under a given condition. If the cellular NAD(H) concentration is lower than the Km, the biocatalyst is not working under the optimal condition, based on the assumption that the apparent kinetics of NAD(H) follows the classic rate equation V = Vmax [NAD(H)]/(Km + [NAD(H)]). Therefore, determination of the apparent kinetics of NAD(H) is necessary for optimizing and designing biocatalysts. It has been difficult to acquire such apparent kinetics data using wild-type cells, because the self-regulation mechanism limits the fluctuation of the cellular cofactor level. However, the apparent kinetics of a given biocatalyst can be determined with the NAD+-auxotrophic mutant described here, due to the accessibility of a wide range of the cellular NAD(H), which should provide valuable information for evaluating the effects of application of biocatalysts.
In summary, we constructed an NTT4-expressed, NAD+-auxotrophic E. coli mutant by disrupting the essential gene responsible for NAD+ biosynthesis and determined the extremes of NAD(H) levels in growing cells. This NAD(H) manipulation strategy should be helpful for increasing our understanding of the biological significance of these redox cofactors and for guiding the design of cofactor-dependent biotransformations.
ACKNOWLEDGMENTS
We are indebted to Ilka Haferkamp (Technische Universität Kaiserslautern, Germany) for her kindly discussion on NTT4 function. We thank the National BioResource Project (NIG, Japan) for providing ΔnadC mutant JW0106 from the KEIO Collection and Barry L. Wanner (Purdue University) for providing E. coli BW25113(pKD3, pKD4, pKD46).
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Information & Contributors
Information
Published In
Applied and Environmental Microbiology
Volume 77 • Number 17 • 1 September 2011
Pages: 6133 - 6140
PubMed: 21742902
Copyright
© 2011 American Society for Microbiology.
History
Received: 21 March 2011
Accepted: 28 June 2011
Published online: 24 August 2011
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Citation
2011.
Determining the Extremes of the Cellular NAD(H) Level by Using an Escherichia coli NAD+-Auxotrophic Mutant . Appl Environ Microbiol
77:.
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