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. 2022 Nov 24;16(3):560–568. doi: 10.1111/1751-7915.14175

NADH dehydrogenases drive inward electron transfer in Shewanella oneidensis MR‐1

Nicholas M Tefft 1, Kathryne Ford 1,2, Michaela A TerAvest 1,✉
PMCID: PMC9948175  PMID: 36420671

Abstract

Shewanella oneidensis MR‐1 is a promising chassis organism for microbial electrosynthesis because it has a well‐defined biochemical pathway (the Mtr pathway) that can connect extracellular electrodes to respiratory electron carriers inside the cell. We previously found that the Mtr pathway can be used to transfer electrons from a cathode to intracellular electron carriers and drive reduction reactions. In this work, we hypothesized that native NADH dehydrogenases form an essential link between the Mtr pathway and NADH in the cytoplasm. To test this hypothesis, we compared the ability of various mutant strains to accept electrons from a cathode and transfer them to an NADH‐dependent reaction in the cytoplasm, reduction of acetoin to 2,3‐butanediol. We found that deletion of genes encoding NADH dehydrogenases from the genome blocked electron transfer from a cathode to NADH in the cytoplasm, preventing the conversion of acetoin to 2,3‐butanediol. However, electron transfer to fumarate was not blocked by the gene deletions, indicating that NADH dehydrogenase deletion specifically impacted NADH generation and did not cause a general defect in extracellular electron transfer. Proton motive force (PMF) is linked to the function of the NADH dehydrogenases. We added a protonophore to collapse PMF and observed that it blocked inward electron transfer to acetoin but not fumarate. Together these results indicate a link between the Mtr pathway and intracellular NADH. Future work to optimize microbial electrosynthesis in S. oneidensis MR‐1 should focus on optimizing flux through NADH dehydrogenases.

This study elucidates the role of NADH dehydrogenases in electron uptake from a cathode to drive intracellular acetoin reduction in Shewanella oneidensis.

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INTRODUCTION

As sustainable methods of electricity generation have expanded, so has interest in using electrical energy to power the electrochemical reduction of CO2 into useful products, a process called electrosynthesis. However, electrosynthesis is still in early stages of development and there are currently no catalysts with the efficiency and longevity to make electrosynthesis economically viable (Pappijn et al., 2020). Microbes are an attractive option for catalysing electrosynthesis because they are self‐renewing and can be genetically modified to produce molecules with unparalleled specificity. To be a good catalyst for electrosynthesis, a microbe must be able to accept electrons efficiently from an extracellular electrode and use them to generate intracellular reducing equivalents that can be used to drive metabolic pathways (Rabaey & Rozendal, 2010). Some attempts have been made to isolate an organism from nature with the capability for electrosynthesis, but none of the isolates have proved easy to culture or genetically modify (Eddie et al., 2017; Rowe et al., 2015). Some natural candidates are iron‐oxidizing bacteria and acetogens (which fix CO2 using inorganic electron donors), and some success in electrosynthesis has been demonstrated with both groups of organisms (Bose et al., 2014; Nevin et al., 2011).

Another avenue to microbial electrosynthesis is to focus on organisms that are known as metal reducers and support much higher extracellular electron transfer rates, although in the outward rather than inward direction. Metal‐reducing bacteria use solid metal oxides as terminal electron acceptors for respiration in nature and can use electrodes as an electron acceptor in the laboratory (Bond & Lovley, 2003; Hernandez & Newman, 2001; Myers & Nealson, 1988). Model metal reducers, such as Shewanella oneidensis and Geobacter sulfurreducens have well‐understood biochemical pathways that connect intracellular electron carriers with extracellular electrodes (Coursolle & Gralnick, 2010; Levar et al., 2014; Marritt et al., 2012; Richardson et al., 2012; Shi et al., 2012, 2014). In S. oneidensis, the reversibility of this pathway has been demonstrated by multiple research groups (Ross et al., 2011; Rowe et al., 2018, 2021). Recently, Rowe et al. (Rowe et al., 2021) have proposed that there may also be a dedicated pathway in S. oneidensis MR‐1 for electron uptake in natural settings. In all previous studies, electron transfer proceeded from the cathode through the electron transport chain to a native terminal electron acceptor, such as oxygen or fumarate (Figure 1).

FIGURE 1.

FIGURE 1

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Shewanella oneidensis MR‐1 inward electron transfer pathways. Black arrows indicate electron transfer steps. CCCP, Ccyanide m‐chlorophenyl hydrazone; LED, Ggreen light light‐emitting diode; MQ, Mmenaquinone; MQH2, Mmenaquinol; PEC, Pperiplasmic electron carriers (e.g., FccA, CctA); PMF, Pproton motive force; PR, Pproteorhodopsin; SMF, sodium motive force.

To use S. oneidensis for microbial electrosynthesis, electrons must be transferred from the cathode to intracellular electron carriers, such as NADH. We previously developed a system in S. oneidensis MR‐1 to enable electrode‐driven reduction of exogenous acetoin to 2,3‐butanediol in the cytoplasm, a process that is NADH‐dependent. We engineered a strain of S. oneidensis carrying butanediol dehydrogenase (Bdh) and proteorhodopsin (PR) on a plasmid with constitutive expression (Figure 1). We observed that the modified S. oneidensis catalysed acetoin reduction to 2,3‐butanediol and that the reduction was dependent on both an active cathode and PR (Tefft & TerAvest, 2019). These results indicated that electrons from the cathode can be used to generate intracellular NADH.

Here, we hypothesize that native NADH dehydrogenases form the link between quinones (which are reduced directly by CymA) and intracellular NADH and that a source of proton motive force (PMF) is necessary to drive NADH dehydrogenases in reverse. The redox potential for the menaquinone/menaquinol pair (−80 mVSHE) is more positive than the redox potential for the NADH/NAD+ pair (−320 mVSHE), making the electron transfer thermodynamically unfavourable (Flamholz et al., 2012). Although ubiquinone could theoretically also be used as an electron carrier for inward electron transfer, it would be even more unfavourable to transfer electrons to NAD+ because of the more positive redox potential of ubiquinone (+80 mVSHE). Further, menaquinone is the redox partner of CymA, suggesting that it is likely the main quinone involved in extracellular electron transfer (McMillan et al., 2012). In the ‘forward’ reaction during respiration, electrogenic NADH dehydrogenases catalyse the favourable electron transfer from NADH to quinones and conserve energy by coupling the electron transfer with proton or sodium movement against their gradients, thus forming a proton motive force (PMF) or sodium motive force (SMF). These complexes are also capable of reverse electron transfer in some contexts, coupling the reverse (unfavourable) electron transfer with the favourable movement of protons or sodium ions with the gradient, thus dissipating PMF or SMF (Spero et al., 2015). The S. oneidensis MR‐1 genome encodes four NADH dehydrogenases, Nuo (SO_1009 to SO_1021), Nqr1 (SO_1103 to SO_1108), Nqr2 (SO_0902 to SO_0907) and Ndh (SO_3517). Nuo couples electron transfer with proton transfer, while Nqr1 and Nqr2 couple electron transfer with sodium ion transfer, and Ndh does not transfer any ions across the membrane.

To test whether NADH dehydrogenases participate in inward electron transfer to acetoin, we measured the ability of strains lacking NADH dehydrogenases to carry out this process. We previously studied the NADH dehydrogenases of S. oneidensis under a range of growth conditions and found that only Nuo and Nqr1 have a significant impact on growth (Duhl et al., 2018; Duhl & Ter Avest, 2019; Madsen & TerAvest, 2019). Therefore, we focused only on those two complexes in this work. Further, we evaluated the role of PMF in this system by testing the effect of the protonophore cyanide m‐chlorophenyl hydrazone (CCCP) on inward electron transfer.

EXPERIMENTAL PROCEDURES

Strains, plasmids and growth conditions

Strains and plasmids used in this study are listed in Table 1 and grown as previously described (Duhl et al., 2018) with the following modifications. M5T medium was additionally supplemented with 0.1% tryptone. M5 medium with the following modifications was used in the working electrode chamber during bioelectrochemical experiments: 100 mM HEPES, no carbon source, no casamino acids. Working chambers were further supplemented with 0.2 μM riboflavin prior to inoculation. For aerobic 2,3‐butanediol experiments, 50 ml of LB supplemented with 50 μg/ml kanamycin and 10 mM acetoin were inoculated with 100 μl of bacterial culture standardized to OD = 1 then grown at 30°C shaking in a 250 ml flask. Samples were taken at 18, 22 and 46 h.

TABLE 1.

Strains and plasmids used in this study.

Strain or plasmid Description Source
E. coli
WM3064 Conjugation strain for S. oneidensis Saltikov and Newman (2003)
S. oneidensis Myers and Nealson (1988)
∆hyaB∆hydA Hydrogenase double knockout mutant Marshall et al. (2008)
∆nuoN∆nqrF1 NADH dehydrogenase double KO strain Duhl et al. (2018)
∆nqrF1 Na+‐dependent NADH dehydrogenase KO strain Duhl et al. (2018)
∆nuoN H+‐dependent NADH dehydrogenase KO strain Duhl et al. (2018)
∆nuoN∆nqrF1 ∆hyaB∆hydA ∆nqrF1 ∆hyaB∆hydA ∆nuoN ∆hyaB∆hydA

NADH dehydrogenase double KO strain with Hydrogenase knockout

Na+‐dependent NADH dehydrogenase KO strain with Hydrogenase knockout

H+‐dependent NADH dehydrogenase KO strain with Hydrogenase knockout

This study

This study

This study
Plasmids
pSMV3.0 R6K origin vector
pSMV3.0∆hyaB In‐frame deletion plasmid for hyaB This study
pSMV3.0∆hydA In‐frame deletion plasmid for hydA This study
pBBR1‐MCS2 Kan resistance, broad host vector Yazynin et al. (1996)
pBdh‐PR pBBR1MCS2 bearing butanediol dehydrogenase and proteorhodopsin Tefft and TerAvest (2019)
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Deletion of hydrogenase genes and transformation of BDH‐PR plasmid

Hydrogenase knockout mutants were prepared in existing NADH dehydrogenase backgrounds, (Duhl et al., 2018) using pSMV3.0 (Saltikov & Newman, 2003). Either hyaB or hydA was deleted from target strains using a standard protocol for a standard protocol for S. oneidensis (Saltikov & Newman, 2003). Deletion strains were initially screened via PCR using gene‐specific forward and reverse primers flanking the deletion site and then sequenced (Sanger sequencing, RTSF Genomics Core, Michigan State University) to verify deletion. Bdh‐PR plasmids were conjugated into S. oneidensis strains using E. coli WM3064 using a standard protocol for S. oneidensis MR‐1 (Coursolle & Gralnick, 2010).

Bioelectrochemical system construction and operation

Bioelectrochemical measurements were performed in custom two‐chambered bioreactors as previously described (Tefft & TerAvest, 2019). Riboflavin was added to working chamber prior to inoculation to a final concentration of 0.2 μM using a 0.1 mM stock solution. Green LED lights were attached to reactors as previously described (Tefft & TerAvest, 2019). Reactors were connected to a potentiostat (VMP, BioLogic USA) and the working electrode poised at +0.2 VAg/AgCl. Current was measured every 1 s for the duration of the experiment. Current measurements were collected for at least 16 h prior to inoculation. Reactors were inoculated with cultures grown in 50 ml of either M5 or M5T medium supplemented with 20 mM d, l‐lactate. Cultures were grown in 250‐ml flasks at 30°C for 17 h shaking at 275 rpm. After 17 h, 50 μl 20 mM all‐trans‐retinal (vitamin A aldehyde; Sigma‐Aldrich, R2500), the essential proteorhodopsin cofactor, (Fuhrman et al., 2008) was added to the flasks to a final concentration of 20 μM and all flasks were returned to incubator, shaking, for 1 h. Adjustments were made to account for the growth defect in ∆nuoN∆nqrF1, preculture growth time was extended to 20 h for this strain (Duhl et al., 2018).

Absorbance at 600 nm was determined for each culture using a biophotometer (Eppendorf, D30) before the volume was transferred to a 50 ml conical tube (VWR, 89039–664) and centrifuged for 5 min at 8000 rpm (Thermo Scientific ST8R; Rotor: 75005709). For M5 cultures 100 ml of culture was pelleted. For M5T cultures, 30 ml of culture was pelleted, except for ∆nuoN∆nqrF1∆hyaB∆hydA where 100 ml was pelleted to account for the growth defect in this strain. Cells were washed in 50 ml of M5 (100 mM HEPES, no carbon source, no tryptone) the supernatant removed and then resuspended in M5 (100 mM HEPES, no carbon source, no tryptone) to OD600 = 3.6.

The working chamber of each bioelectrochemical system was inoculated with 9 ml of standardized cell suspension using an 18 g needle (Beckton Dickson, 305196) and 10‐ml syringe (Beckton Dickson, 302995). Inoculated reactors were exposed to ambient oxygen for 6 h before N2 gas, 99.9% (Airgas) was bubbled into the reactors through a 0.2 μm filter. The rate of flow of nitrogen gas was observed using a bubbler attached to the gas outlet line from the reactors. The rate of gas flow was maintained so there was positive pressure against the water in the bubbler and bubble rate was kept constant between reactors.

Reactors remained at +0.2 VAg/AgCl for 40 h before the potential was changed to −0.5 VAg/AgCl. After 2 h, sterile, anoxic acetoin solution was added to a final concentration of 1 mM. A 2‐ml sample was taken approximately every 24 h using a 21 g needle (Beckton Dickson, 305167) and 3‐ml syringe (Beckton Dickson, 309657). One millilitre was used for determination of OD600. One millilitre was transferred to a micro‐centrifuge tube (VWR, 20170–038) and frozen at −20°C until preparation for HPLC analysis. Experiments without a set potential on the working electrode were set up as above excepting that immediately after acetoin injection, the potentiostat was disconnected from the working electrodes. Anoxic sodium fumarate was injected into each reactor to a final concentration of 20 mM at 72 h post acetoin injection.

For CCCP experiments, reactors were prepared as above but 24 h after acetoin injection CCCP was added to a final concentration of 125 μM from a stock dissolved in 100% ethanol via syringe and needle. For control conditions, an equal volume of 100% ethanol was added without CCCP.

HPLC analysis

HPLC analysis was performed as previously described with the following changes (Tefft & TerAvest, 2019). A 0.5 ml/min flow rate was used for better separation of 2,3 butanediol and acetoin at measured concentrations. 2,3‐butanediol standards were prepared at 0.01, 0.025, 0.05, 0.1, 0.5, 1 and 5 mM.

Electrode protein quantification

Immediately upon cessation of each experiment, working electrodes were removed from the working chamber and placed in a 50 ml conical tube. The electrodes were frozen at −20°C until protein extraction. Electrodes were prepared by suspending harvested electrodes in 20 ml 0.2 M NaOH. Samples were heated at 95°C in a drying oven (VWR, 89511–404) for 30 min, inverting every 10 min to mix. After heating, 25 μl of prepared samples were transferred to a clear 96 well plate (VWR, 10062–900). Standards and reagents were prepared using a Pierce BCA protein assay kit (Thermo Scientific) as per protocol. A working reagent volume of 200 μl was added to each well and incubated at 37°C for 30 min. Absorbance at 562 nm was measured using a synergy H1M plate reader (BioTek) and exported using Gen5 analysis software (BioTek). Protein concentration was determination using a standard bovine serum albumin (BSA) curve.

Data analysis

Analysis of HPLC, protein concentration, OD and current data was performed using Rstudio using the following packages: ggplot2, reshape2, dplyr and TTR (Ulrich, 2017; Wickham, 2007, 2009; Wickham & Francois, 2015).

RESULTS

To determine the role of NADH dehydrogenases in inward electron transfer in S. oneidensis MR‐1, we compared a background strain without hydrogenases (∆hyaB∆hydA) to strains lacking the hydrogenases and one or more NADH dehydrogenase. The hydrogenase deletion strain was used as the background to rule out H2‐mediated electron transfer, as in our previous work (Tefft & TerAvest, 2019). We focused on Nuo and Nqr1 because previous work from our group demonstrated the importance of these two complexes under a range of conditions, including aerobic growth, soluble iron reduction and electrode reduction (Duhl et al., 2018; Madsen & TerAvest, 2019). We have not observed significant phenotypes for Nqr2 or Ndh mutants under any tested condition, therefore, they were not included in this study.

As in our previous work, PR and Bdh were constitutively expressed from a plasmid in all strains to enable light‐driven PMF generation and the use of acetoin reduction as an NADH sink (Figure 1). We then pre‐grew all strains under aerobic conditions and inoculated into two‐chambered electrochemical cells at an optical density (OD600) of 0.2. Because flavins are essential for effective extracellular electron transfer in S. oneidensis MR‐1, 0.2 μM riboflavin was added to all bioreactors before inoculation (Kotloski & Gralnick, 2013; Marsili et al., 2008). To facilitate cell attachment to the electrode and oxidation of any stored or carryover organic carbon, the electrochemical cells were passively aerated, and the electrode was poised at an anodic potential of +0.2 VAg/AgCl. After 6 h, N2 sparging was started and all strains generated anodic current during this phase of the experiment, with ∆nuoN∆nqrF1∆hyaB∆hydA generating a higher peak anodic current than the other strains (Figure 2A). Anodic current generation increases after N2 sparging because the cells switch from using O2 to the electrode as the terminal electron acceptor. After an additional 40 h, the electrode was switched to a cathodic potential of −0.5 VAg/AgCl (Figure 2B). After the potential switch, a spike in cathodic current was observed, and current was allowed to stabilize for 2 h before acetoin injection (time = 0). Under these conditions, the cells do not grow and OD600 decreases over time due to electrode attachment and possibly cell death (Figure S1). However, the removal of organic carbon and alternative electron acceptors is essential to interpretation of the experiments. The total protein on the electrodes was evaluated at the end of the experiment and was not significantly different among any of the strains (Figure S2).

FIGURE 2.

FIGURE 2

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Current generation by S. oneidensis strains under operation of electrochemical cells. (A) Electrode poised at anodic potential. (B) Electrode poised at cathodic potential. Injection of acetoin to a final concentration of 1 mM occurred at time 0. (C) Black arrows indicate when sodium fumarate was injected to a concentration of 20 mM (second arrow is for ∆nuoN∆nqrF1∆hyaB∆hydA only). The disparity in time of fumarate injection in ∆nuoN∆nqrF1∆hyaB∆hydA conditions was due to the fumarate being injected slightly later during the experiment. ∆hyaB∆hydA is shown in black, ∆nuoN∆hyaB∆hydA is shown in green, ∆nqrF1∆hyaB∆hydA is shown in orange, ∆nuoN∆nqrF1∆hyaB∆hydA is shown in magenta. Conditions where no cells were added (sterile) are shown in grey. Lines represent the average of three replicates with standard error shown in transparent ribbons. Sterile and ∆nqrF1∆hyaB∆hydA conditions are an average of two replicates.

As in our previous study, we observed that ∆hyaB∆hydA with PR and Bdh generated cathodic current over the course of 3 days and accumulated 2,3‐butanediol, while sterile controls produced background cathodic current of less than −5 μA and no 2,3‐butanediol (Figure 2). In general, cathodic current was unchanged in the ∆nuoN∆hyaB∆hydA strain but decreased in the ∆nqrF1∆hyaB∆hydA and double knockout strains for the first ~1.5 days after acetoin injection (Figure 2B). Deletion of nuoN did not affect 2,3‐butanediol production, while deletion of nqrF1 caused a 25% decrease compared with the parent strain at day three (two‐tailed t‐test, p < 0.05) (Figure 3). Combined deletion of nuoN and nqrF1 completely blocked electrode‐driven 2,3‐butanediol production. 2,3‐butanediol accumulation was below detectable levels for all strains when the working electrode was not connected to the potentiostat (data not shown).

FIGURE 3.

FIGURE 3

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2,3‐butanediol accumulation. ∆hyaB∆hydA is shown in black, ∆nuoN∆hyaB∆hydA is shown in green, ∆nqrF1∆hyaB∆hydA is shown in orange. Reactors with an unpoised electrode and ∆nuoN∆nqrF1∆hyaB∆hydA had 2,3‐butanediol values that fell below the limit of detection shown in the dotted black line. Lines represent the average of three replicates with standard error shown in transparent ribbons.

Because the ∆nuoN∆nqrF1∆hyaB∆hydA strain generally grows more poorly than ∆hyaB∆hydA, we made several comparisons to determine whether the lack of 2,3‐butanediol production was due to a general defect in the strain. First, we compared anodic current generated by all strains in the first phase of the experiment, before switching the electrode to a cathodic potential. We observed that ∆nuoN∆nqrF1∆hyaB∆hydA produced a peak anodic current over 2‐fold higher than the other strains and the total charge transfer in the anodic phase of the experiment was 2.8 times higher than the ∆hyaB∆hydA strain (Figure 2A). We also measured total protein in each BES to rule out possible disparity in cell attachment to the electrode. We found no significant differences between strains in measured protein from electrode or media samples (Figure S2).

We also tested the ability of all strains to transfer electrons from a cathode to fumarate. This activity requires MtrCAB, CymA and FccA, but is not expected to require NADH dehydrogenases (Ross et al., 2011; Rowe et al., 2018, 2021). After 3 days of cathodic operation with acetoin, we injected fumarate to a final concentration of 20 mM and observed an immediate increase in cathodic current (Figure 2C). Strains lacking single NADH dehydrogenases exhibited moderate decreases in fumarate‐dependent cathodic current compared with the ∆hyaB∆hydA strain; however, the difference in charge transfer was not significant. The ∆nuoN∆nqrF1∆hyaB∆hydA strain generated over 3‐fold less fumarate‐dependent cathodic current. However, all strains exhibited fumarate‐dependent current >5‐fold higher than acetoin‐dependent current.

We evaluated the ability of the ∆nuoN∆nqrF1∆hyaB∆hydA strain to reduce acetoin to 2,3‐butanediol under conditions where it grows well, that is, in oxic lysogeny broth (LB). Cells were grown in flasks to ensure oxygen availability and sampled periodically to measure growth and 2,3‐butanediol. We observed that both the control and NADH dehydrogenase double mutant strain produced ~1 mM 2,3‐butanediol from 10 mM acetoin under this condition (Figure 4A). In this condition, the NADH dehydrogenase mutant strain produced slightly higher levels of 2,3‐butanediol, possibly due to a build‐up of NADH in the cytoplasm, as previously observed (Duhl & Ter Avest, 2019). The 2,3‐butanediol concentration decreased during stationary phase for both strains, possibly due to back conversion to acetoin after major electron donors in the medium were consumed.

FIGURE 4.

FIGURE 4

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(A) Cell density and (B) 2,3‐butanediol accumulation of S. oneidensis strains grown in LB with shaking in 250‐ml flasks. ∆hyaB∆hydA is shown in black, ∆nuoN∆nqrF1∆hyaB∆hydA is shown in magenta. Each point represents the average of three replicates for ∆nuoN∆nqrF1∆hyaB∆hydA and two replicates for ∆hyaB∆hydA with standard error or range, respectively, shown in error bars.

We hypothesized that a source of PMF would be essential for inward electron transfer to acetoin if the Nuo and Nqr1 were functioning in reverse to generate NADH. The experimental conditions used for inward electron transfer do not provide a known route for PMF generation by any native process, that is, there is no source of ATP that could be hydrolysed by the FOF1 ATP synthase and no electron acceptor energetically ‘downhill’ of menaquinone (such as fumarate or oxygen) that could be used for forward reactions of the electron transport chain. We previously observed that the presence of functional PR and green light enhanced inward electron transfer, supporting the hypothesis that PMF is important for electron transfer from a cathode to acetoin (Tefft & TerAvest, 2019).

Here, we tested whether PMF is required for inward electron transfer by adding the protonophore uncoupler carbonyl cyanide m‐chlorophenyl hydrazone (CCCP) in experiments with the ∆hyaB∆hydA strain (all NADH dehydrogenases present). In these experiments, CCCP dissolved in ethanol was injected into the bioelectrochemical systems to a final concentration of 125 μM approximately 24 h after acetoin injection (Figure 5). CCCP has poor solubility in water which necessitated the use of ethanol as a carrier to achieve target concentrations. Upon CCCP injection, cathodic current decreased and 2,3‐butanediol accumulation slowed (Figure 5B). In systems with ethanol injection alone as a negative control, 2,3‐butanediol concentration continued to increase for another 2 days, at a faster rate than in systems without ethanol or CCCP injection (Figure 2). To test whether the effect of CCCP was caused specifically by the loss of PMF or simply by cell death and lysis caused by CCCP, we injected fumarate into the electrochemical cells on day 3 as described above. We observed that fumarate‐dependent cathodic current increased similarly in both conditions (Figure 5C).

FIGURE 5.

FIGURE 5

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(A) Cathodic current in electrochemical cells with modified S. oneidensis. (B) 2,3‐butanediol accumulation in the same experiment. (C) Cathodic current after addition of 20 mM fumarate. Black arrows indicate when CCCP was injected to a concentration of 125 μM. Dotted lines indicate reactors with CCCP injection and solid lines represent reactors with ethanol only injection. Each point represents the average of three replicates with standard error shown in error bars or transparent ribbons.

DISCUSSION

Our previous work provided proof of the concept that the extracellular electron transfer pathway of S. oneidensis can be driven in reverse to power NADH‐dependent reactions in the cytoplasm. Here, we showed that deletion of the two key NADH dehydrogenases, Nuo and Nqr1, prevented accumulation of 2,3‐butanediol to detectable levels, indicating that these complexes are critical to inward electron transfer from the electrode. We observed that deletion of either NADH dehydrogenase alone had little to no effect, consistent with our previous observations that Nuo and Nqr1 have overlapping functions but one or the other is required for aerobic growth or current generation at an anode (Duhl et al., 2018; Madsen & TerAvest, 2019).

Although the ∆nuoN∆nqrF1 NADH dehydrogenase deletion strain has a growth defect compared with the parent strain, the lack of acetoin reduction cannot be attributed to this general defect. The ∆nuoN∆nqrF1 NADH dehydrogenase mutant was capable of acetoin reduction in oxic‐rich medium (Figure 5) and had cell mass on the electrode consistent with other strains (Figure S2). Further, the double NADH dehydrogenase deletion strain produced a higher peak anodic current than any other strain (Figure 2A). We speculate that this was due to a greater amount of stored organic carbon accumulated in the cells during aerobic growth in flasks. The ∆nuoN∆nqrF1 NADH dehydrogenase mutant has a reduced respiration rate which reduces its ability to oxidize residual organic carbon during the aerobic phases of the experiment (Duhl & Ter Avest, 2019). It is not likely that cell death was responsible for the lack of acetoin reduction as the strain was capable of elevated electron transfer to fumarate (Figure 2C), albeit at a reduced rate compared to the hydrogenase mutant. These results point to a role of the NADH dehydrogenases specifically in electron transfer from a cathode to acetoin.

While the NADH dehydrogenases had a major impact on 2,3‐butanediol production, anodic current and fumarate‐dependent current, the changes in cathodic current were smaller. This is due in part to abiotic processes occurring at the cathode, which account for ~5 μA. We also observed biotic current that was not associated with 2,3‐butanediol production in ∆nuoN∆nqrF1. At this time, it remains unclear what other biotic process is occurring, although one possibility is reduction of trace oxygen in the system.

We also found that the electron transfer rate to acetoin is orders of magnitude lower than the rate for outward electron transfer to an anode or inward electron transfer to a native terminal electron acceptor, such as fumarate or oxygen (Ringeisen et al., 2006; Ross et al., 2011; Rowe et al., 2018, 2021). This observation provides evidence that the Mtr pathway is not the rate‐limiting step for electrode‐driven acetoin reduction because electron transfer to fumarate occurred at a much faster rate than electron transfer to acetoin although both are dependent on the Mtr pathway (Figure 1). Thermodynamic analysis suggests that electron transfer from menaquinol to NAD+ could be the rate‐limiting step, considering that this reaction is highly unfavourable with ΔrG'm = +46.2 kJ/mol (Flamholz et al., 2012). We hypothesize that electron transfer from menaquinol to NAD+ can only occur at a significant rate by coupling it to the thermodynamically favourable movement of protons from the periplasm to the cytoplasm via electrogenic NADH dehydrogenases. Favourable proton movement down the gradient is coupled with many otherwise unfavourable but necessary cellular processes, including ATP synthesis, flagellar rotation and transport of solutes against their gradients. There are also confirmed examples of SMF dissipation driving unfavourable redox reactions, such as electron transfer from NADH to ferredoxin catalysed by the Rnf complex (Westphal et al., 2018). Our results with NADH deletion strains support the hypothesis that PMF is needed to drive reverse electron transfer; when the two key electrogenic NADH dehydrogenases were deleted, electrode‐driven reduction was dramatically inhibited, although Ndh (the uncoupling NADH dehydrogenase) was still present.

The hypothesis that PMF drives inward electron transport is further supported by results with CCCP injection, which showed that when the uncoupler is added, electrode‐driven acetoin reduction is greatly reduced. CCCP increases membrane permeability to protons and inhibits cells from maintaining PMF, therefore, it blocks processes that require PMF, such as ATP synthesis. Fumarate injection after CCCP injection indicated that the uncoupler did not inhibit electron transfer to an energetically downhill electron acceptor, suggesting that the cessation of 2,3‐butanediol production was not due to a general effect of CCCP, such as toxicity. Surprisingly, an increase in 2,3‐butanediol production (relative to experiments shown in Figure 2) was observed with ethanol alone and was not accompanied by an increase in current (Figure 5). We speculate that S. oneidensis oxidized the ethanol to gain NADH that was used to reduce acetoin. We also observed that cathodic current recovered somewhat after ~1 day in the bioreactors with CCCP, although no further 2,3‐butanediol was accumulated. Previous reports indicate that CCCP can be degraded by light (Pistorius et al., 1975). It is not clear why cathodic current would recover but not 2,3‐butanediol production. We speculate that regulatory changes caused by the CCCP may have activated an alternate electron transfer pathway, although the identity of the terminal electron acceptor remains unknown. In future work, we will investigate alternate methods for dissipating PMF that do not involve introduction of CCCP or an organic carbon source.

AUTHOR CONTRIBUTIONS

Nicholas M Tefft: Formal analysis (equal); investigation (lead); methodology (equal); visualization (lead); writing – original draft (lead); writing – review and editing (equal). Kathryne C Ford: Formal analysis (supporting); investigation (supporting); methodology (supporting); visualization (supporting); writing – review and editing (supporting). Michaela TerAvest: Formal analysis (equal); funding acquisition (lead); investigation (supporting); methodology (equal); project administration (lead); visualization (supporting); writing – original draft (supporting); writing – review and editing (equal).

FUNDING INFORMATION

This material is based upon work supported by the National Science Foundation Graduate Research Fellowship under Grant No. (DGE‐1848739) to Kathryne C. Ford, and a National Science Foundation CAREER Award (1750785) and 2018 Beckman Young Investigator Award to Dr. Michaela TerAvest.

CONFLICT OF INTEREST

The authors declare no conflict of interest.

Supporting information

Figure S1.

Click here for additional data file. (103.9KB, docx)

ACKNOWLEDGMENTS

This material is based upon work supported by the National Science Foundation Graduate Research Fellowship under Grant No. (DGE‐1848739) to Kathryne C. Ford, and a National Science Foundation CAREER Award (1750785) and 2018 Beckman Young Investigator Award to Dr. Michaela TerAvest.

Tefft, N.M. , Ford, K. & TerAvest, M.A. (2023) NADH dehydrogenases drive inward electron transfer in Shewanella oneidensis MR‐1. Microbial Biotechnology, 16, 560–568. Available from: 10.1111/1751-7915.14175

DATA AVAILABILITY STATEMENT

All data are available upon request.

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Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Supplementary Materials

Figure S1.

Click here for additional data file. (103.9KB, docx)

Data Availability Statement

All data are available upon request.

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