ABSTRACT
Shewanella oneidensis MR-1 is a facultative anaerobe that respires using a variety of electron acceptors. Although this organism is incapable of fermentative growth in the absence of electron acceptors, its genome encodes LdhA (a putative fermentative NADH-dependent d-lactate dehydrogenase [d-LDH]) and Dld (a respiratory quinone-dependent d-LDH). However, the physiological roles of LdhA in MR-1 are unclear. Here, we examined the activity, transcriptional regulation, and traits of deletion mutants to gain insight into the roles of LdhA in the anaerobic growth of MR-1. Analyses of d-LDH activity in MR-1 and the ldhA deletion mutant confirmed that LdhA functions as an NADH-dependent d-LDH that catalyzes the reduction of pyruvate to d-lactate. In vivo and in vitro assays revealed that ldhA expression was positively regulated by the cyclic-AMP receptor protein, a global transcription factor that regulates anaerobic respiratory pathways in MR-1, suggesting that LdhA functions in coordination with anaerobic respiration. Notably, we found that a deletion mutant of all four NADH dehydrogenases (NDHs) in MR-1 (ΔNDH mutant) retained the ability to grow on N-acetylglucosamine under fumarate-respiring conditions, while an additional deletion of ldhA or dld deprived the ΔNDH mutant of this growth ability. These results indicate that LdhA-Dld serves as a bypass of NDH in electron transfer from NADH to quinones. Our findings suggest that the LdhA-Dld system manages intracellular redox balance by utilizing d-lactate as a temporal electron sink under electron acceptor-limited conditions.
IMPORTANCE NADH-dependent LDHs are conserved among diverse organisms and contribute to NAD+ regeneration in lactic acid fermentation. However, this type of LDH is also present in nonfermentative bacteria, including members of the genus Shewanella, while their physiological roles in these bacteria remain unknown. Here, we show that LdhA (an NADH-dependent d-LDH) works in concert with Dld (a quinone-dependent d-LDH) to transfer electrons from NADH to quinones during sugar catabolism in S. oneidensis MR-1. Our results indicate that d-lactate acts as an intracellular electron mediator to transfer electrons from NADH to membrane quinones. In addition, d-lactate serves as a temporal electron sink when respiratory electron acceptors are not available. Our study suggests novel physiological roles for d-LDHs in providing nonfermentative bacteria with catabolic flexibility under electron acceptor-limited conditions.
INTRODUCTION
Members of the genus Shewanella, which belongs to the class Gammaproteobacteria, are widely distributed in marine, freshwater, and sedimentary environments (1, 2). Shewanella spp. are known to be able to respire using various terminal electron acceptors, including oxygen, fumarate, nitrite, nitrate, dimethyl sulfoxide, trimethylamine N-oxide (TMAO), iron and manganese oxides, and electrodes in microbial fuel cells (1, 2). Shewanella oneidensis MR-1 is the most extensively studied strain in the genus Shewanella in terms of its annotated genome sequence (3), ease of genetic manipulation (4), and respiratory versatility (5, 6). The strain MR-1 was isolated from anaerobic sediment in a shallow eutrophic lake (5). The versatile respiratory capacity of this strain is thus thought to have evolved in eutrophic environments where no abundant electron acceptors are present. Studies have also suggested that this strain possesses well-developed environment-sensing systems for regulating catabolic and respiratory pathways in response to environmental changes, including those associated with shifts in external redox potentials and/or available electron acceptors (7, 8).
Previous studies have reported that MR-1 uses a cyclic-AMP (cAMP) receptor protein (CRP) to regulate the expression of genes involved in anaerobic respiration, including those for the reduction of fumarate (fccA), nitrite (nrf), nitrate (nap), and metal oxides (mtrC, mtrA, mtrB, and omcA) (9–12). CRP is therefore regarded as a master regulator of diverse anaerobic respiratory pathways in MR-1 (7). Moreover, a recent study found that CRP is also required for the transcriptional activation of dld, a gene encoding a respiratory d-lactate dehydrogenase (d-LDH) that catalyzes the oxidation of d-lactate to pyruvate with the reduction of membrane quinones (13). This finding suggests that CRP is used to coordinately regulate d-lactate oxidation and anaerobic respiration.
In addition to dld, MR-1 possesses the ldhA gene, which encodes a putative NADH-dependent d-LDH that catalyzes the fermentative production of d-lactate from pyruvate (3, 14). A previous study found that an engineered S. oneidensis strain produced d-lactate from glucose under electron acceptor (fumarate)-limited conditions, and that LdhA was the primary catalyst for this reaction (15). Orthologs of the ldhA gene are present in other members of Shewanella (https://www.ncbi.nlm.nih.gov/genome/13542), suggesting that the ability to produce d-lactate is a common feature of this genus. However, given that this strain is incapable of fermentative growth in the absence of external electron acceptors (1, 16) and that it utilizes Dld for the oxidation of d-lactate in the presence of an electron acceptor (14, 15), the physiological roles of LdhA in MR-1 remain unclear.
Previous studies have shown that an imbalance in NADH/NAD+ affects a variety of cellular processes, including catabolic pathways (17), stress resistance (18), and conditional viability (19). In the present study, we hypothesized that the strain MR-1 uses d-LDHs to control the intracellular NADH/NAD+ ratio during anaerobic respiration-associated sugar catabolism, in which d-lactate serves as an intracellular electron carrier and/or temporal electron sink. To test this hypothesis, we analyzed the activity of LdhA, transcriptional regulation of the ldhA gene, and the growth traits of ldhA deletion mutants.
RESULTS
NADH oxidation activity of LdhA.The ldhA gene (SO_0968) exhibits homology to fermentative d-LDH genes of fermentative bacteria (14) and is thought to be involved in d-lactate production under electron acceptor-limited conditions (15). However, there is no experimental evidence of whether or not LdhA catalyzes d-lactate production coupled to NADH oxidation. To determine the enzymatic activity of LdhA, we measured the NADH-dependent LDH activity of cell extracts obtained from wild-type (WT) MR-1 and the ldhA deletion mutant (ΔldhA) (Fig. 1). Our results showed that the LDH activity of the ΔldhA mutant was markedly lower than that of the WT, indicating that LdhA functions as an NADH-dependent LDH.
NADH-dependent LDH activity of WT, ΔldhA mutant, and Δcrp mutant strains. Cells were anaerobically grown in NAG minimal medium (NMM) containing 20 mM NAG and 5 mM TMAO and harvested at stationary-growth phase. The LDH activity of cells was normalized based on their total protein content. Error bars represent standard deviations calculated from the results from at least three independent experiments.
Involvement of CRP in ldhA expression.A previous study identified a transcription start site (TSS) of the ldhA gene located 29 bp upstream of the ATG start codon (TSSldhA; Fig. 2A) (20). Given that a putative CRP-binding motif is located upstream of TSSldhA (Fig. 2A) (21), we hypothesized that CRP is involved in the expression of ldhA. To investigate the involvement of CRP in the expression of ldhA, we compared the expression levels of ldhA in the WT and a crp-deletion mutant (Δcrp) using quantitative reverse transcription-PCR (qRT-PCR). When cells were anaerobically grown under TMAO-reducing conditions, the expression level of ldhA in the Δcrp mutant decreased to approximately 40% of that in the WT (Fig. 2B). A comparison of the NADH-dependent LDH activity of WT and Δcrp mutant cells (Fig. 1) showed that the Δcrp mutant exhibited a lower level of LDH activity than did WT. These results demonstrate that CRP is involved in the transcriptional activation of ldhA.
Regulation of ldhA expression by CRP. (A) Nucleotide sequence upstream of ldhA. The TSSldhA is indicated in bold. Putative –10 and –35 promoter sequences are underlined. The putative CRP-binding motif is indicated by the boxed region. (B) qRT-PCR analyses of ldhA in WT and Δcrp mutant. Cells were anaerobically grown in NMM containing 20 mM NAG and 5 mM TMAO and harvested at logarithmic-growth phase. Results are expressed as relative levels of mRNA expression in WT cells. Error bars represent standard deviations calculated from the results from at least three independent experiments. (C) DNA fragments used as probes for EMSA. Positions of the 5′ and 3′ ends of the fragments relative to TSSldhA (+1) are shown. The mutated sequences in PBldhAm are shown in bold. (D) Binding of CRP to each probe. DNA-binding reactions were performed in the presence (+) or absence (–) of CRP and cAMP.
To confirm whether CRP directly regulates the expression of ldhA, we performed an electrophoretic mobility shift assay (EMSA) using purified CRP. When a Cy3-labeled DNA probe containing the upstream region of TSSldhA (PBldhA; Fig. 2C) was mixed with CRP, a shifted band was detected in the presence of cAMP (Fig. 2D). To examine the binding of CRP to the putative CRP-binding motif contained in this probe, we also performed an EMSA using a mutated probe in which the core motif sequence was modified (PBldhAm; Fig. 2C). Our results revealed that CRP did not interact with this mutated probe even in the presence of cAMP (Fig. 2D). We therefore concluded that CRP binds to this motif and directly regulates the transcription of ldhA. Given that CRP is also involved in the transcriptional regulation of dld (13), our results indicate that CRP coordinately regulates the expression of these two d-LDH genes and thereby plays a central role in the regulation of d-lactate metabolism in MR-1.
d-Lactate production during sugar catabolism.Although WT MR-1 is unable to grow on glucose, it has the ability to utilize N-acetylglucosamine (NAG) as a growth substrate (22). Given that MR-1 catabolizes NAG using the Entner-Doudoroff (ED) pathway, which employs NADH as a redox cofactor (7), we speculated that this strain may use LdhA to produce d-lactate during NAG catabolism. To test this hypothesis, the WT and ΔldhA mutant were anaerobically grown on NAG under TMAO-respiring conditions, and their abilities to produce d-lactate were compared (Fig. 3). We found that the ΔldhA mutant showed a lower ability to produce d-lactate than did the WT, demonstrating that MR-1 utilizes LdhA to produce d-lactate during NAG catabolism. We also found that the Δcrp mutant produced less d-lactate than did the WT, supporting our finding that CRP regulates d-lactate production in MR-1 (Fig. 3).
d-Lactate production from NAG in WT, ΔldhA mutant, and Δcrp mutant strains. Cells were anaerobically grown in NMM containing 20 mM NAG and 5 mM TMAO and harvested at the stationary phase. Error bars represent standard deviations calculated from the results from at least three independent experiments.
Roles of d-LDHs in respiratory NAG utilization.NADH-dependent LDH is known to be involved in NADH oxidation (NAD+ regeneration) during lactic acid fermentation. However, MR-1 did not grow on NAG in the absence of external electron acceptors (see Fig. S1 in the supplemental material), indicating that this strain cannot grow during lactic acid fermentation. We were therefore interested in investigating the physiological role of LdhA in MR-1. Given that MR-1 possesses a respiratory d-LDH (Dld), we speculated that this strain utilizes LdhA and Dld for NADH oxidation and quinone reduction as a bypass of respiratory NADH dehydrogenases (NDHs). To test this hypothesis, we used gene knockout experiments to examine the contribution of these d-LDHs to respiratory NAG utilization. We first compared the growth of the WT and ΔldhA mutant on NAG under electron acceptor (fumarate)-limited (EAL) and electron acceptor-rich (EAR) conditions (Fig. 4). We found that the ΔldhA mutant exhibited growth trends similar to those of the WT (Fig. 4), indicating that LdhA is not essential for growth under these conditions. However, this result is reasonable given that MR-1 possesses four NDH complexes, Nuo, Nqr1, Nqr2, and Ndh (3). We therefore examined the growth of a mutant in which the genes for all four NDH complexes were disrupted (ΔNDH mutant) (8) on NAG under EAL and EAR conditions. The results revealed that the ΔNDH mutant retained the ability to grow on NAG, although its growth was slower than that of the WT (Fig. 4). Given that catabolism of NAG requires NAD+ (23), our results indicate that MR-1 possesses an additional NADH-oxidizing system(s) on top of the four NDHs. To investigate whether LdhA and Dld function as this alternative NADH-oxidizing system, we generated a deletion mutant of ldhA and dld in a ΔNDH mutant background (ΔNDH ΔldhA and ΔNDH Δdld mutants, respectively) and examined the growth on NAG under EAL and EAR conditions (Fig. 4). These mutants completely lost the ability to grow on NAG (Fig. 4), demonstrating that LdhA and Dld are both required for NAG catabolism in a ΔNDH mutant background. These d-LDHs likely have a significant role in growth under EAL conditions, given that the growth of the ΔNDH mutant was more severely impaired under the EAR conditions (Fig. 4B) than the EAL conditions (Fig. 4A).
Growth of S. oneidensis derivatives on NAG under EAL (A) and EAR (B) conditions. Cells were grown in NMM containing 20 mM NAG and 10 mM fumarate (for EAL [A]) or 10 mM NAG and 100 mM fumarate (for EAR [B]). Error bars represent standard deviations calculated from the results from at least three independent experiments.
The results support the idea that MR-1 utilizes d-lactate as an electron carrier in NAG catabolism, particularly when electron acceptors are limited. To further examine this hypothesis, we measured NAG consumption and d-lactate production in the cultures of the WT and the ΔNDH, ΔNDH Δdld, and ΔNDH ΔldhA mutants grown under EAL and EAR conditions (Fig. 5). Under the EAL conditions, the ΔNDH and ΔNDH Δdld mutants consumed NAG and produced significant amounts of d-lactate, the WT produced a moderate amount of d-lactate, and the ΔNDH ΔldhA mutant showed no NAG consumption or d-lactate production (Fig. 5A and C). This result suggests that some pyruvate produced during NAG catabolism in WT MR-1 is converted to d-lactate by LdhA under EAL conditions. It is noteworthy that the ΔNDH and ΔNDH Δdld mutants produced markedly increased amounts of d-lactate compared to the WT (Fig. 5C). The increased d-lactate production by the ΔNDH mutant is likely related to an increased accumulation of intracellular NADH in this mutant, as shown previously (8), suggesting the importance of the LdhA-Dld system in electron transfer in this mutant. The accumulation of d-lactate in the ΔNDH Δdld mutant is reasonable, given that this mutant is unable to oxidize d-lactate produced by LdhA. This result, together with the observation that the ΔNDH Δdld mutant did not grow under these culture conditions (Fig. 4A), indicates that MR-1 is unable to conserve sufficient energy from lactic acid fermentation only, and that the growth of this strain on NAG requires respiratory electron transfer via NDHs and/or LdhA-Dld. Under the EAR condition, however, d-lactate concentrations in the cultures of WT, ΔNDH mutant, and ΔNDH Δdld mutant were markedly decreased compared to those under the EAL conditions (Fig. 5D). This result is consistent with a previous observation that ldhA expression is reduced under EAR conditions (21). Under both culture conditions, acetate was detected as the major metabolite from NAG (Fig. S2), indicating that the respiratory growth of MR-1 on NAG under anaerobic conditions depends on the partial oxidation of this carbohydrate. Taken together, our results indicate that d-lactate serves as an electron carrier and/or a temporal electron sink when MR-1 catabolizes NAG under EAL conditions.
NAG consumption (A and B) and d-lactate production (C and D) by S. oneidensis derivatives grown under EAL (A and C) and EAR (B and D) conditions. Cells were grown in NMM containing 20 mM NAG and 10 mM fumarate (for EAL [A and C]) or 10 mM NAG and 100 mM fumarate (for EAR [B and D]). Error bars represent standard deviations calculated from the results from at least three independent experiments.
To investigate the contribution of LdhA to NAG catabolism in the WT background, we examined the anaerobic growth of the ΔldhA mutant in minimal medium containing on 20 mM NAG and 5 mM TMAO. When cells were grown under this severe EAL condition, the growth of the ΔldhA mutant was slower than that of the WT (Fig. 6). This growth retardation was not observed when the ΔldhA mutant was complemented with a plasmid expressing the ldhA gene (Fig. S3). These results, together with the observation that the ΔldhA mutant produced a smaller amount of d-lactate than WT under this growth condition (Fig. 3), indicate that d-lactate production by LdhA facilitates the growth of MR-1 on NAG when electron acceptors are severely limited.
Growth of WT and ΔldhA mutant in NMM containing 20 mM NAG and 5 mM TMAO. Error bars represent standard deviations calculated from the results from at least three independent experiments.
DISCUSSION
The present work shows that MR-1 uses LdhA and Dld as a bypass of NDH for transferring electrons from NADH to quinones during NAG catabolism under EAL conditions (termed the d-LDH-dependent NDH bypass; Fig. 7). It is likely that d-lactate serves as a temporal electron sink under EAL conditions. We also demonstrated that the d-LDH-dependent bypass is regulated by CRP, indicating that this bypass is regulated in coordination with anaerobic respiratory pathways in MR-1. Intracellular NADH/NAD+ ratios have been shown to affect a variety of cellular processes in bacteria (17–19). We therefore propose that MR-1 utilizes the d-LDH-dependent NDH bypass as a method for controlling the NADH/NAD+ ratio and enhancing its catabolic flexibility under eutrophic EAL conditions.
Proposed NADH oxidation and quinone reduction pathway consisting of LdhA and Dld. MQ, oxidized form of menaquinone; MQH2, reduced form of menaquinone; G3P, glyceraldehyde 3-phosphate. A blue dashed box indicates d-LDH-dependent NDH bypass consisting of LdhA and Dld. Red dotted arrows indicate the expression of the lldP (putative lactate permease), dld, and ldhA genes that are regulated by CRP.
Members of the genus Shewanella are known to preferentially utilize low-molecular-weight organic acids, such as lactate, as sources of carbon and energy (24). Many researchers therefore regard d-lactate as the major growth substrate for these bacteria. The present study, however, shows that MR-1 produces d-lactate as an intermediate metabolite during NAG catabolism (Fig. 3 and 5). NAG is the main sugar component of chitin and is abundant in the natural environment (25). Given that most known members of Shewanella, including MR-1, have the ability to utilize NAG (26), it is likely that they utilize this carbohydrate as a major growth substrate in their natural habitat. Although anaerobic lactate oxidation in Shewanella spp. does not require NADH-dependent enzymes, including NDHs or NADH-dependent LDHs (27), NAG catabolism is associated with the redox cycling of NADH (7, 23) (Fig. 7). Considering that members of Shewanella have acquired versatile respiratory pathways to survive in eutrophic EAL environments, it is reasonable to speculate that they may also harbor multiple NADH oxidation pathways for responding to changes in environmental conditions. Given that the present study shows that MR-1 possesses five systems for respiratory NADH oxidation, i.e., four NDHs (Nuo, Nqr1, Nqr2, and Ndh) and a d-LDH-dependent system, it would be interesting to examine the differences in the physiological roles of these NADH oxidation systems.
Our findings indicate that the regulation of lactate metabolism in S. oneidensis MR-1 is substantially different from those in bacteria capable of fermentative growth on sugars, such as Escherichia coli, Bacillus subtilis, and Lactococcus lactis (28–30). For example, E. coli uses multiple regulatory systems, including Mlc and CsrAB, to regulate ldhA expression (28). Mlc is a global regulator that negatively regulates the transcription of genes involved in sugar utilization, including those for the phosphoenolpyruvate-dependent sugar phosphotransferase system (31). The CsrAB system, consisting of the RNA-binding protein CsrA and the noncoding RNA molecule CsrB, functions as a posttranscriptional regulatory system that balances glycolysis and sugar storage as glycogen (32). These findings indicate that LdhA is regulated in coordination with the glycolytic pathway in E. coli. However, E. coli uses FnrS, an anaerobically induced small RNA that also regulates the expression of genes involved in aerobic respiration, to regulate the expression of the dld gene (33). The differences in the regulation of LdhA (d-lactate-producing LDH) and Dld (d-lactate-oxidizing LDH) likely reflect the fact that E. coli uses these d-LDHs in different physiological processes (fermentation and respiration, respectively). In contrast, the present study demonstrates that MR-1 uses CRP to coordinately regulate the expression of ldhA and dld, suggesting that Shewanella spp. use these d-LDHs in concert for NADH oxidation (NAD+ regeneration) during respiratory carbohydrate catabolism. The difference in the physiological roles of LdhA in E. coli and MR-1 may reflect the difference in the abilities of these bacteria for fermentative growth on sugars. We propose two possible explanations for the inability of MR-1 to grow under fermentative conditions: (i) low ATP yields result from glycolysis through the ED pathway (34), and (ii) the excessive reduction of membrane quinones results from the simultaneous expression of ldhA and dld in the absence of external electron acceptors. The latter hypothesis is supported by the observation that the deletion of formate dehydrogenases allows MR-1 to grow on pyruvate under fermentative conditions, probably by suppressing the irreversible reduction of membrane quinones that causes the cessation of cell growth (35).
Based on the results presented here, we propose that Shewanella spp. have evolved the ability to utilize d-LDHs for respiratory electron transfer in eutrophic EAL environments. Fermentative NADH-dependent LDHs exemplified by LdhA are also found in other nonfermentative bacteria, including members of the genera Pseudomonas and Alteromonas (36, 37), and we hypothesize that the d-LDH-dependent NDH bypass may similarly be present in these bacteria. Further investigations are needed to address the ecological significance of the d-LDH-dependent NDH bypass as a common physiological mechanism for bacteria to cooperatively survive in anaerobic environments.
MATERIALS AND METHODS
Bacterial strains, plasmids, and growth conditions.The bacterial strains and plasmids used in this study are listed in Table 1. S. oneidensis strains were cultivated at 30°C in lysogeny broth (LB) medium or minimal medium (15) containing NAG as the carbon and energy source (NAG minimal medium [NMM]). The Δcrp mutant strain was cultured aerobically in LB or anaerobically in NMM supplemented with 20 mM NAG and 5 mM TMAO as the electron acceptor because this strain cannot utilize other electron acceptors (9). The ΔNDH mutant strain was cultivated under anaerobic fumarate-reducing conditions because this strain cannot grow under aerobic conditions (8). For anaerobic cultivation under EAL conditions, NMM supplemented with 20 mM NAG and 10 mM fumarate or 5 mM TMAO in a screw-top test tube or vial was inoculated with S. oneidensis strains. For anaerobic cultivation under EAR conditions, NMM supplemented with 10 mM NAG and 100 mM fumarate was used. Test tubes or vials containing the anaerobic cultures were sealed with butyl rubber septa and purged with high-purity nitrogen gas (99.99%). The optical density at 600 nm (OD600) of the cultures was measured using a UH5300 spectrometer (Hitachi, Tokyo, Japan) or a mini-photo 518R photometer (Taitec, Tokyo, Japan). Escherichia coli strains were cultivated in LB or 2× yeast extract-tryptone (YT) medium at 37°C. The E. coli mating strain (WM6026) required 100 μg/ml 2,6-diaminopimelic acid for growth. When necessary, 50 μg/ml kanamycin (Km) was added to the culture medium. Agar plates contained 1.6% Bacto agar (Difco, Franklin Lakes, NJ).
Bacterial strains and plasmids used in this study
To construct plasmid pBBRldhA, the ldhA gene was amplified using primers ldhA-F-KpnI and ldhA-R-EcoRI (Table S1). The PCR product was digested using KpnI and EcoRI and cloned between the corresponding sites of pBBR1MCS-2 (38). The resulting plasmid, pBBRldhA, was introduced into ΔldhA mutant cells by filter mating with E. coli WM6026.
LDH assay.S. oneidensis cells were anaerobically grown in 50 ml of MM containing 20 mM NAG and 5 mM TMAO and harvested at stationary-growth phase. The cells were washed with 50 mM KH2PO4 buffer (pH 7.4) and resuspended in 1 ml of phosphate-buffered saline (PBS; 137 mM NaCl, 2.7 mM KCl, 10 mM Na2HPO4·12H2O, 1.8 mM KH2PO4 [pH 7.4]). The cell suspensions were ultrasonicated using a Misonix S4000 sonicator (Farmingdale, NY) and centrifuged at 1,000 × g for 5 min to remove cell debris. The protein concentration of the extracted solution was determined using a Micro bicinchoninic acid (BCA) protein assay kit (Thermo Fisher Scientific, Waltham, MA).
The NADH-dependent LDH activity of cell-free extracts was spectrophotometrically measured using the SH-1200Lab absorbance microplate reader (Corona Electric, Ibaraki, Japan), according to a method described previously (39–41). The assay measured the decrease in absorbance at 340 nm as NADH was oxidized to NAD+ by pyruvate, as catalyzed by LDH. The reaction mixture (250 μl) contained 30 mM sodium pyruvate, 2 mM 3-(N-morpholino)propanesulfonic acid (MOPS) buffer (pH 7.0), 192 μM NADH, and 10 μl of a cell extract. One unit of enzyme activity was defined as the amount of enzyme required to oxidize 1 nmol NADH per min (39–41). All measurements were performed in triplicate at the minimum.
Mutant construction.In-frame gene deletion mutants of S. oneidensis MR-1 were generated using a two-step homologous recombination method with the suicide plasmid pSMV-10, as described previously (42, 43). The pSMV-10 derivatives pSMV-ldhA and pSMV-dld-II (15) were used for the disruption of ldhA and dld, respectively. Each plasmid was introduced into the WT or ΔNDH mutant by filter mating with E. coli WM6026. Transconjugants (single-crossover clones) were selected on LB plates containing 50 μg/ml Km and were further cultivated for 20 h in LB medium without antibiotics. The cultures were then spread onto LB plates containing 10% (wt/vol) sucrose to isolate Km-sensitive double-crossover mutants. Disruption of the target gene in the obtained strains was confirmed by PCR.
RNA extraction.S. oneidensis cells were anaerobically grown in NMM containing 20 mM NAG and 5 mM TMAO and were harvested at the logarithmic-growth phase (OD600, 0.05 to 0.07). Total RNA was extracted from cells using TRIzol reagent (Thermo Fisher Scientific) and subsequently purified using an RNeasy minikit and a RNase-free DNase set (Qiagen, Valencia, CA), according to the manufacturer’s instructions. The quality of the purified RNA was evaluated using an Agilent 2100 Bioanalyzer with RNA 6000 Pico reagents and RNA Pico chips (Agilent Technologies, Santa Clara, CA), according to the manufacturer’s instructions.
qRT-PCR.qRT-PCR was performed using a LightCycler 1.5 instrument (Roche, Indianapolis, IN), according to a method described previously (44). Briefly, the PCR mixture (20 μl) contained 15 ng RNA, 1.3 μl of 50 mM Mn(OAc)2 solution, 7.5 μl of LightCycler RNA master SYBR green I (Roche), and 0.15 μM primers listed in Table S1. To generate standard curves, DNA fragments of the ldhA and 16S rRNA genes were PCR amplified from the total DNA of MR-1. The expression level of ldhA was normalized based on the expression level of the 16S rRNA gene. All measurements were performed in triplicate at the minimum.
Metabolite analysis.S. oneidensis strains were anaerobically cultivated in NMM, and the cells were removed by filtration through a membrane filter unit (0.20-μm pore size, DISMIC-25HP; Advantec, Tokyo, Japan). The amount of d-lactate in the supernatants was measured using an F-kit (J. K. International, Tokyo, Japan), according to the manufacturer’s instructions. The amounts of NAG and acetate in the supernatants were measured using high-performance liquid chromatography (Agilent 1100 series), as described elsewhere (45).
EMSA.N-terminal histidine-tagged CRP (N-ht-CRP) was purified and used for EMSA according to a method described previously (12, 13). Cy3-labeled DNA probes PBldhA and PBldhAm were generated by annealing the complementary single-strand oligonucleotides listed in Table S1. DNA-binding reactions were performed in 20 μl of a previously described reaction mixture (12) containing 2 nM Cy3-labeled DNA probe, 2 μg N-ht-CRP, and 50 nM cAMP. The mixtures were incubated on ice for 30 min and then loaded onto a nondenaturing 12.5% polyacrylamide gel. Electrophoresis was conducted at 150 V in 0.5× Tris-borate-EDTA buffer. Fluorescent gel images were obtained using a Typhoon FLA 9000 imager (GE Healthcare, Chicago, IL).
ACKNOWLEDGMENTS
We thank Nanako Amano for technical assistance.
This work was supported by a grant-in-aid for JSPS Fellows (to T.K., grant 16J08653), Grant-in-Aid for Young Scientists (B) (to A.K., grant 26850056), and a Grant-in-Aid for Scientific Research (C) (to A.K., grant 18K05399).
T.K. carried out the majority of the experimental work and drafted the manuscript. Y.S. constructed the plasmid pBBRldhA and carried out the complementation experiments of the ΔldhA mutant. A.K. conceived of the study, participated in its design and coordination, and drafted the manuscript. K.W. supervised the study and performed manuscript editing. All authors read and approved the final manuscript.
We declare no conflicts of interest.
FOOTNOTES
- Received 5 November 2018.
- Accepted 18 November 2018.
- Accepted manuscript posted online 30 November 2018.
Supplemental material for this article may be found at https://doi.org/10.1128/AEM.02668-18.
- Copyright © 2019 American Society for Microbiology.
