Electrodriven H2 Production in Escherichia coli: Rational Design and Mechanistic Studies of the Electron Uptake Process

doi:10.21203/rs.3.rs-5108109/v1

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Electrodriven H2 Production in Escherichia coli: Rational Design and Mechanistic Studies of the Electron Uptake Process

https://doi.org/10.21203/rs.3.rs-5108109/v1

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Electromicrobial production systems, which use electrons from renewable energy sources to drive microbial metabolism towards desired products, are considered a promising strategy for future energy conversion and sustainable synthesis technologies. However, electron transport to microbes remains a critical yet poorly understood process. This hinders the rational design of these systems for viable energy efficiencies. Here, we report the construction of an efficient redox power-transport unit that converts electrons into internally generated H₂, which can either be harvested directly or used as reducing power within the cell to drive Escherichia coli metabolism towards more complex products. Efficient conversion of electrons supplied by the electronic circuit to H₂ within the cell is achieved by engineering E. coli with functional [Fe-Fe] hydrogenase from the green algae Chlamydomonas reinhardtii (CrHydA1) and selecting a suitable electron transfer mediator. The system's design is guided by a developed kinetic model, which provides insights into the mechanism and kinetics of the electron uptake process.

Physical sciences/Chemistry/Electrochemistry/Electrocatalysis

Physical sciences/Chemistry/Catalysis/Biocatalysis

Biological sciences/Biotechnology/Industrial microbiology

Physical sciences/Chemistry/Green chemistry/Biofuels

Meeting the continuously increasing global energy demand while mitigating global warming requires alternative sustainable pathways for fuel and chemical production. Electromicrobial production has garnered considerable attention as a promising strategy for future energy conversion technologies. The basic principle behind electromicrobial production involves using reducing equivalents generated extracellularly to drive microbial metabolism facilitating the conversion of substrates like CO₂, H₂O, and/or N₂ into a desired product. This field has branched into two primary research directions: microbial electrosynthesis ¹ and biohybrid ^2–4 systems. Systems where microbial production is driven by electrons directly supplied by electrodes are usually referred to as microbial electrosynthesis (MES), ^5,6 while those where microbe metabolism is driven photoelectrochemically are termed biohybrids.^7–9 Despite this distinction, the two approaches have significant overlaps, both with regard to general system design and fundamental chemical processes.

For both MES and biohybrid systems, the transfer of reducing power from the abiotic to the biotic component poses a significant challenge.¹⁰ Depending on the approach used to deliver redox equivalents, both MES and biohybrid systems can broadly be further divided into two types.^6,11 In the first type there is a direct electron transfer (DET) from the abiotic (photo)electrode component to the cells.^12,13 In the second type, electron shuttles, such as H₂ produced (photo)electrochemically ^4,14,15 or other small molecule redox mediators are used.^16–18 This type is referred to as mediated electron transfer (MET) systems. For both DET- and MET-type systems, the most employed strategy so far was to use natively “electroactive” microorganisms possessing the natural ability to uptake electrons.^1,2,19–21 However, due to limited genetic tools currently available for commonly used autotrophs, ²² production of targeted products is limited.⁶ Thus, about 75% of all studies report production of acetate, as a very high percentage of metabolic flux in typically utilized acetogens is intrinsically directed toward this compound.²³

Recently, there has been interest in electromicrobial production with non-electroactive organisms, particularly Escherichia coli (E. coli), for which a wide variety of genetic tools are available, thereby enabling the production of almost any desirable product. ²⁴ Chen and co-workers introduced the Phenazine-1-carboxylic acid (PCA) pathway of P. aeruginosa into E. coli. ¹⁶ Engineered E. coli using self-excreted PCA as an electron transfer mediator allowed electrons on the cathode to be transferred to the inner membrane and utilized for improved succinate production. Similarly, Wu et. al improved production of succinate in engineered electroactive strains of E. coli by introducing proteins from the electron transport pathway of Shewanella oneidensis MR-1 into E. coli. ²⁵ However, in this case, Neutral red was introduced as an electron mediator to transfer redox equivalents from an electrode to the biological carriers inside the cell. Ye et al. compared the effectiveness of various strategies of electron transfer inside E. coli cells either by ″building bridges″ through the introduction of membrane electron transfer proteins or ″digging holes″ through the insertion of membrane porins. ²⁶ The authors compared strains engineered with electron transfer pathways from either S. oneidensis MR-1 or S. lithotrophicus ES-1 alone and in combination with the OprF porin from P. aeruginosa, and/or three overexpressed endogenous OmpA, OmpC, and OmpF porins. The strain containing the S. lithotrophicus ES-1 electron transfer pathway and overexpressed OmpF showed the highest yield of succinate production, but again with Neutral red as the mediator.

In addition to studying the effect of supplying electrochemical reducing power on the natural anaerobic metabolism of E. coli, the possibility of using this organism as a platform for the synthesis of a wider range of products has been investigated. Mayr et. al introduced an alcohol dehydrogenase from Lactobacillus brevis (LbADH) to E. coli previously engineered with electron-transfer pathway c-type-cytochromes from S. oneidensis. ^27,28 Using methyl viologen as a mediator, electrons from an electrode were transferred to the LbADH and utilized for the synthesis of (R)-1-phenylethanol from acetophenone. We and others have engineered E. coli cells with [Fe-Fe] hydrogenases, yielding H₂-producing biohybrid systems when combined with both intra-^29,30 and extracellular^17,31,32 light harvesters. In the latter case, electron transfer from the photosensitizer to the enzyme was enabled by redox mediators such as methyl viologen or diquat derivates. Despite the use of redox mediators, an efficiency of about 85% has been reported with regard to converting electrons from the sacrificial electron donor to H₂.³¹

While there is a vast body of literature on anodic electron transfer from microorganisms to the electrode, the mechanism of the reverse process—cathodic electron transfer from the electrode to the microorganisms—remains poorly understood. ^23,33 To the best of our knowledge only one study has attempted to model this process,³⁴ while another report offered a general conceptual approach towards such modeling.³³ The reported model as well as the offered modeling approach considered a configuration where microbial cells are attached to a cathode in the MES reactor.

Here, we construct an electromicrobial production system based on suspended E. coli cells engineered with functional [Fe-Fe] hydrogenase from the green algae Chlamydomonas reinhardtii (CrHydA1) produced by whole-cell artificial maturation.^35,36 We demonstrate that, with a suitable choice of mediator, redox equivalents can be transferred from the electrode to the [Fe-Fe] hydrogenases within the cells, resulting in H₂ production. Using cyclic voltammetry, we investigate the kinetics of electron transfer within the system and propose a kinetic model that allows us to simulate the experimental response. By applying the developed model to experimental data collected under varying conditions, we find that electron transfer from the mediator to the hydrogenase inside the cell is the rate-limiting step of the overall electron supply process. This electron transfer rate depends on the driving force provided by the difference in potentials between the mediator and the hydrogenase active site. This finding challenges the common assumption that, at the cell level, mediator mass transport across the abiotic/biotic interface limits the energetic efficiency of the system.

The approach presented here opens up the possibility of systematically investigating the contribution of each system component down to the (intra)cellular level to the rate of electron consumption, thereby enabling the informed use of synthetic biology tools. Moreover, the developed system can be used as an efficient redox power-transport unit, converting electrons to internally generated H₂, for a modular design of microbial production in E. coli engineered towards the desired product. This electron transport unit can bypass constraints related to energy efficiency and safety linked with the limited solubility of externally produced H₂. ^6,37

Electrochemical characterization of mediators. From a thermodynamic perspective, the formal reduction potential of the mediator must be lower than that of the reaction catalyzed by the [Fe-Fe] hydrogenase, which involves the interconversion of H⁺/H₂ (E⁰′= -0.636 V vs. Ag/AgCl at pH 6.8), to facilitate efficient transfer of the reducing power towards the desired product. We selected four organic mediators, previously demonstrated by some of us to transfer electrons to [Fe-Fe] hydrogenases inside E. coli cells in a biohybrid system,³¹ as possible candidates for the construction of the system (Fig. 1A). These mediators were then examined through cyclic voltammetry (CV). As depicted in Fig. 1B, each mediator, starting from its (+ 2) oxidation state, undergoes two quasi-reversible reduction events (Table 1). Thus, each mediator has two redox couples, M²⁺/M¹⁺ and M¹⁺/M⁰, with reduction potentials that are thermodynamically favorable for driving H⁺/H₂ interconversion. The more reduced couple of each mediator provides a large driving force for the reaction at the cost of a higher overpotential. Considering only the first reduction step, the reductive power of the mediators follows the order: DQ > DQ-CO₂H > DQ-OH > MV.

Table 1

Formal reduction potentials of the investigated mediators.
Mediator	E⁰′_M2+/M1+ (V vs Ag/AgCl)	E⁰′_M1+/M0 (V vs Ag/AgCl)
DQ	-0.873	-1.065
DQ-CO₂H	-0.845	-1.110
DQ-OH	-0.808	-1.035
MV	-0.660	-1.010

Selection of suitable mediator. The capacity of the mediators to drive H₂ production by E. coli BL21 DE3 cells containing [Fe-Fe] hydrogenase (E. coli (CrHydA1)) was evaluated by CV analysis. The E. coli (CrHydA1) cells heterologously expressed the CrHydA1 hydrogenase, and the enzyme was activated through artificial maturation to ensure a high and well-defined intracellular concentration.^36,38 Fig. 2A shows CVs of 1 pM E. coli (CrHydA1) cells suspended in a deaerated 150 mM phosphate-buffered saline (PBS) at pH 6.8 (orange trace) in comparison to that of the same electrolyte without the cells (black trace). The CVs overlap, indicating (i) the absence of direct electronic interaction between the electrode and the cells; and (ii) the experimental conditions did not compromise cell viability during the measurement period, as cell lysis would result in the release of hydrogenase into the electrolyte leading to an increase in the cathodic current near E⁰′(H⁺/H₂) arising from DET to the enzyme (vide infra).

Initially, MV was assessed as the electron transfer mediator as the formal potential of its first reduction (E⁰′ = -0.660 V vs. Ag/AgCl) lies closest to the E⁰′(H⁺/H₂) among the selected mediators and therefore a successful mediation by MV²⁺/MV¹⁺ will lead to the lowest overpotential requirement. The voltammogram recorded immediately after the addition of cells to a 2 mM solution of MV in the electrolyte showed a gradual increase in the anodic peak current at -0.620V V vs. Ag/AgCl, assigned to MV¹⁺ → MV²⁺ transformation with no corresponding changes in the cathodic wave (Fig. S1A). Over time (7–30 min), this change in the voltammograms became more pronounced. This observation contradicts the anticipated response for MET from the electrode to the cytoplasmic hydrogenase. Instead, it suggests that in the presence of cells, MV²⁺ is converted into MV¹⁺ by some cellular component. We attribute this to the reduction of the mediator by ferredoxin. Ferredoxin is abundant in E. coli and has a formal reduction potential in the range of -0.636 V to -0.734 V vs. Ag/AgCl. ^39,40 Thus, ferredoxin having E^0′ < -0.636 V can reduce MV²⁺ (Fig. S1B), leading to a rise in MV¹⁺ concentration near the electrode surface, reflected in the gradual increase in the anodic current over time. No changes in the cathodic peak current corresponding to the second reduction of MV (E^0′ = -1.010V vs. Ag/AgCl) were observed either, suggesting that MV¹⁺/MV⁰ also cannot mediate electron transfer from the electrode to the hydrogenase inside the cells (Fig. S1A) under tested conditions. A sharp anodic peak at -0.973 V vs. Ag/AgCl was observed, which was previously assigned to the one-electron oxidation of the surface-adsorbed MV0 species. ⁴¹ Thus, MV0 cannot transfer electrons to the cells as it adsorbs to the electrode surface.

Among the diquat-based molecules investigated in this study, DQ has the lowest formal reduction potential in the first reduction event DQ²⁺ → DQ¹⁺. It was therefore evaluated as the next mediator. Upon the addition of 2.2 pM of E. coli (CrHydA1) cells to a 200 µM solution of DQ in a deaerated 150 mM PBS at pH 6.8, an increase in the cathodic currents with a concurrent decrease in anodic peak currents compared to CVs of DQ in the absence of cells were observed for both DQ redox couples (black vs orange trace in Fig. 2B and Fig. S2A). With increasing E. coli (CrHydA1) cell concentration, the cathodic current augmented further and the anodic peak almost disappeared (purple trace in Fig. 2B and Fig. S2A). To quantify the current increase, we compared values of the cathodic peak current at -0.925 V vs. Ag/AgCl, corresponding to the DQ²⁺ → DQ¹⁺ transformation, upon addition of cells to the value recorded without the cells (Fig. 2C). At the highest added concentration of E. coli (CrHydA1) of 8.8 pM, a current enhancement of 39 ± 5% was observed. This current enhancement did not depend on the concentration of the mediator (Fig. S3). It is worth mentioning here that the extent of the peak current increase started to plateau beyond 4.4 pM. This may arise from the precipitation of the cells, which was observed at cell concentrations > 4.4 pM. The current enhancement for the second reduction peak, assessed at -1.126 V vs. Ag/AgCl and assigned to the DQ¹⁺ → DQ⁰ transformation, was 40 ± 8%, i.e. practically identical to that of the first reduction peak (Fig. S2B). Despite the larger driving force for the more reduced redox couple of DQ for electron transfer to the hydrogenase inside the cells, no additional current enhancement was observed. The observed changes in the CV are consistent with the expected response of a MET system.

To confirm that the changes in current are due to the catalytic activity of hydrogenase and not arising from some other process(es), we substituted the natural cofactor [Fe₂(adt)(CO)₄(CN)₂]²⁻ ([2Fe]^adt, adt = azadithiolate) with [Fe₂(pdt)(CO)₄(CN)₂]²⁻ ([2Fe]^pdt, pdt = propanedithiolate) during the enzyme maturation stage, (yielding E. coli (CrHydA1-PDT)). ⁴² In [2Fe]^pdt, the amine bridgehead of the adt ligand is replaced with a methylene group that prevents [2Fe] subsite protonation and impedes catalytic turnover. ^35,43 Upon performing electrochemical measurements with E. coli (CrHydA1-PDT), both cathodic and anodic currents of DQ were found to slightly decrease with increasing cell concentration (Fig. 2D), most probably due to blocking of the active electrode surface by cells. This observation underscores that the active form of the cytoplasmic CrHydA1 hydrogenase is necessary to manifest the changes illustrated in Fig. 2B and confirm that the observed changes in the CVs upon E. coli (CrHydA1-ADT) addition are indeed appearing from MET from the electrode to the cytoplasmic hydrogenase.

Following successful mediation by DQ, the reductive capability of DQ-CO₂H was assessed. Upon the addition of E. coli (CrHydA1), cathodic currents emerged from − 0.636 V vs. Ag/AgCl, i.e. from E⁰′(H⁺/H₂) (Fig. S4A). The cathodic current also increased with increasing concentration of E. coli (CrHydA1). However, this was approximately 98 mV higher compared to the onset of the reductive current corresponding to DQ-CO₂H²⁺ → DQ-CO₂H^+ 1 reduction in DQ-CO₂H (-0.734 V vs Ag/AgCl, black trace). Such a response is unexpected in a MET system, where changes are typically constrained to the CV features of the mediator. ⁴⁴ Instead, we attribute this behaviour to direct electron transfer from the electrode to the hydrogenase, resulting in currents starting from the formal potential of the H⁺/H₂ couple. ⁴⁵ Therefore, the observed current response suggests that DQ-CO₂H is adversely impacting the cellular integrity leading to the release of the cytoplasmic hydrogenase into the electrolyte. To test this, we first measured the CV of DQ-CO₂H in the presence of 4.4 pM E. coli (CrHydA1) (Fig. S4B, red trace) and thereafter exposed the electrolyte to air for 1 minute. Any hydrogenase present in the electrolyte will become irreversibly inhibited due to its extreme sensitivity to oxygen. ^46,47 The electrolyte was subsequently deoxygenated before being transferred to the glovebox and subjected to CV measurements again. The previously observed current increase completely disappeared (Fig. S4B, pink vs red trace), confirming that the changes indeed stemmed from the hydrogenases released into the electrolyte (contrary to the behavior observed in the presence of DQ, as discussed below). Consequently, DQ-CO₂H was excluded from further consideration in this study.

Finally, DQ-OH was evaluated as the mediator. As for DQ, an increase in the cathodic current with a concurrent decrease in the anodic current was again observed in CVs with the addition of E. coli (CrHydA1) to the 200 µM DQ-OH solution for both DQ-OH redox couples (Fig. S5A). Similar to DQ, the changes in the currents were confined to the CV features of DQ-OH, suggesting that, unlike DQ-CO₂H, DQ-OH does not adversely impact cellular integrity. Fig. S5 shows the current increase at both the first (-0.855 V vs. Ag/AgCl) and second (-1.105 V vs. Ag/AgCl) reduction peak potentials upon the addition of cells in comparison to the value recorded without the cells. The maximum increase of 27% was observed at the first peak potential upon the addition of 8.8 pM of cells. As in the case of DQ, the current enhancement for the first and second reduction peaks of DQ-OH was similar. Overall, the current enhancements observed with DQ were consistently higher compared to DQ-OH. Therefore, further stability studies of the system were performed with DQ as the mediator.

Cell integrity analysis. As DQ-CO₂H prompted the release of hydrogenase into the electrolyte, we opted to verify cellular integrity following electrochemical studies with DQ to ensure that the observed responses originated from cytoplasmic hydrogenase. After recording CVs of DQ in the presence of 8.8 pM E. coli (CrHydA1), we centrifuged the electrolyte to precipitate the cells. The resulting supernatant is devoid of E. coli (CrHydA1) but will still contain hydrogenase released from the cells in the event of cell lysis. The supernatant was subjected to CV (Fig. S6). The CV traces of the supernatant overlapped with that of DQ in isolation (light purple vs. black trace in Fig. S6) suggesting that (i) all current changes were arising from the interaction between DQ and cytoplasmic hydrogenase present in E. coli (CrHydA1), and (ii) no hydrogenase is present in the electrolyte. Further, SDS-PAGE analysis on both the supernatant and the centrifuged cells was performed to check the presence of hydrogenase. While the centrifuged cells showed a strong band close to 50 kDa corresponding to hydrogenase, no corresponding band was present in the supernatant (Fig. S7). Indeed, the SDS-PAGE analysis showed no indication of proteins in the electrolyte. This further confirms that in measurements using DQ as a mediator, the cells remain intact under our experimental conditions and the observed current enhancements are entirely due to MET between the electrode and cytoplasmic hydrogenases.

To assess the impact of DQ on the cellular integrity and bioelectrochemical performance of E. coli (CrHydA1) over an extended duration, we incubated 8.8 pM E. coli (CrHydA1) cells with 200 µM DQ in a sealed glass vial at 4 ˚C under strictly anaerobic conditions for 24 hours. We then compared the CVs of the solution before and after 24-hour storage (Fig. S8A purple vs light purple traces). After 24 hours, the system retained ~ 95% of its initial activity with no significant changes in the CV features (Fig. S8B). The latter observation further confirms that the cells remained intact as the release of cytoplasmic hydrogenase into the electrolyte is expected to change CV features (Fig. S4A), as observed with DQ-CO₂H. The minimal change in activity, coupled with the preservation of cellular integrity, suggests that DQ can serve as an effective and safe mediator for electromicrobial production with suspended E. coli cells engineered with functional [Fe-Fe] hydrogenases.

Oxygen tolerance of the electromicrobial production system. It has been previously observed that E. coli provides protection to cytoplasmic hydrogenase from the external environment, particularly oxygen. ^29,48 CrHydA1 is an extremely oxygen-sensitive enzyme and in our measurements using DQ-CO₂H as the mediator, we observed that exposing the electrolyte to air for one minute leads to the complete disappearance of the current enhancements originating from hydrogenase present in the electrolyte (Fig. S4B). To check the oxygen sensitivity of the electromicrobial production system based on DQ-mediated E. coli (CrHydA1), we exposed the electrolyte to air for 10 or 20 minutes. We then compared CVs of 200 µM DQ in the presence of 8.8 pM E. coli (CrHydA1) before and after oxygen exposure (Fig. S9, purple vs light purple traces). As shown in Fig. S9B, the activity drops to 74% and 70% of the initial enhancement after 10 and 20 minutes, respectively. Such retention of activity after 20 minutes of air exposure suggests that the cytoplasm of E. coli is able to partially protect CrHydA1 from oxygen, most likely due to the high respiration rate of the bacteria resulting a low intracellular concentration.

Modeling of electrochemical data. To unravel the kinetics of MET to cytoplasmic hydrogenase and pinpoint the rate-determining step of the electromicrobial synthesis platform at the level of cells, we constructed a kinetic model allowing us to simulate the experimental cyclic voltammograms. We considered two different approaches to model microbial cells: (I) by modeling the activity of individual cells ^49–52; or (II) by using a continuum approach, where the behavior of single cells is neglected, and the average concentration of components is used instead (Scheme 1). ³³ The difference between these two approaches lies in what is considered the catalyst and how the concentration and activity of the catalyst are calculated. In the first approach, E. coli cells are the catalyst. This approach is based on the adaptation of early reported modeling frameworks for modeling CVs of biofilm anodes.^49–52 In the second approach hydrogenase is the catalyst and it is assumed that it is homogenously distributed in the reaction volume. Both approaches are based on molecular electrochemistry methods commonly used to model enzymatic kinetics.^53,54 Further, they incorporate steps reported for the modeling of biofilm-modified anodes but adapted to model the cathodic process.⁴⁹ For both approaches the same steps with slight modifications were used (Scheme 1).

Step 1 represents the reversible reduction of the mediator at the electrode surface characterized by the formal potential and standard heterogeneous electron transfer rate constant (k₀), both of which can be obtained by simulating CVs of the mediator without the addition of cells (Fig. 1, Table S1). Mediator oxidation and reduction rates at the electrode surface depend on the potential applied relative to the formal potential of the mediator as described by the Butler-Volmer equation. ⁵⁵ Initially, only the DQ response was modeled and only one redox couple of the mediator, DQ²⁺/DQ¹⁺, was considered. The model was subsequently further extended to include the second redox couple, M^+ 1/M⁰ and DQ-OH.

Step 2 describes the reduction of the biocatalyst (either the E. coli cells or hydrogenase) by electron transfer from the mediator. The biocatalyst in its native state was designated as BioCat(Ox), while its states following successive reductions were labeled as BioCat (Red) and BioCat (Red2). The electron transfer from the mediator to hydrogenase could occur directly or via an intermediate intracellular species, which undergoes initial reduction before transferring the electron to hydrogenase. Thus, in the context of [Fe-Fe] hydrogenase, BioCat(Ox), BioCat (Red), and BioCat (Red2) can be interpreted as the well-known H_ox, H_redH⁺ and H_Hyd states, all of which have been shown to accumulate in vivo.³⁸ However, the model does not a priori make any assumption on the exact nature of the three redox states. For simplicity, the rates associated with the electron transfer steps (k_ET), regardless of their actual complexity, were denoted as k_ET1 and k_ET2, respectively, where k_ET1 and k_ET2 were assumed to represent the rate-limiting step of the entire electron transfer process from the mediator to the biocatalyst for both approaches. In the modeling, the magnitudes of k_ET1 and k_ET2 were assumed to be equal (k_ET1= k_ET2= k_ET)

Step 3 is the catalytic turnover of the biocatalyst in which protons are converted to hydrogen. In models of biofilm-modified anodes, this step is usually assumed to follow Michaelis-Menten kinetics. In our modeling for both approaches it was assumed that there is excess of the substrate and this step was modeled as a single catalytic step with the rate constant k_cat. This assumption should be valid as the pKa of the proton-donating Cys169 in the active site of CrHydA1 is 12,^56,57 so it should always be protonated at the pH 6.8 used in the experiments. While the internal pH of the cell might differ from the pH of the buffer used, and it was reported to be approximately 0.5 pH units higher in E. coli cytoplasm, ^58,59 it is still sufficiently low to maintain effective protonation of the Cys169.

Mass transport of the mediator, cells, and hydrogen occurs only by diffusion in the model and is described by Fick’s law of diffusion, ⁵⁵ parametrized by the corresponding values of the diffusion coefficients (D) taken from the literature (Table S1). Further, experimental conditions are selected in such a way that mass transport of the mediator does not limit the catalytic response as the current enhancement does not depend on the concentration of the mediator (Fig. S3) and this condition is reproduced in the model. To simplify the model, the reactions of Steps 2 and 3 were considered as unidirectional occurring only in the direction of H₂ generation. Considering the reduction potentials, this assumption should hold for all diquat derivates studied here. In short, current enhancement in the presence of E. coli cells results from the reduction of the excess of the oxidized mediator at the electrode surface (Step 1). This excess is generated by oxidation of the reduced mediator by the biocatalyst via electron transfer between the reduced mediator and the biocatalyst (Step 2), which in turn is sustained by the catalytic activity of the biocatalyst (Step 3).

Approach I. To model the activity of individual E. coli cells we first estimated the k_cat of individual cells using the earlier calculated concentration of hydrogenase inside the cell (43 000 molecules or 70 µM) and the estimated catalytic activity of a single hydrogenase enzyme, 400 s^− 1.³² Assuming a linear correlation between the total enzyme amount and rate, a calculated value of k_cat of 1.72×10⁷ s^− 1 is obtained. Using the calculated k_cat, 8.8 pM concentration of the cells, and values of other parameters specified in Table S1, simulated CV traces reproduced the experimentally observed current response (Fig. 3A vs Fig. 2B, purple trace) for k_ET ~ 400–500 k_cat. Higher values of k_ET relative to the k_cat resulted in a larger enhancement of the cathodic current. To simulate the gradual addition of E. coli (CrHydA1) during the measurements (Fig. 2B), we varied the concentration of the biocatalyst from 2.2 to 8.8 pM for k_ET = 500 k_cat (Fig. 3B). A gradual increase in the cathodic current with a subsequent decrease in the anodic current was observed in the simulated CVs (Fig. 3B), reproducing the experimental data.

We further performed simulations to mimic control experiments (Fig. 2D). Hydrogenases artificially matured with [2Fe]^pdt can undergo a single electron reduction ⁶⁰ and this was used in the simulations. In this case, both k_ET2 and k_cat were set to 0, while k_ET1 was set to 6.88×10⁹ s^− 1 (i.e. 500 × 1.72×10⁷ s^− 1). The simulated catalytic CV does not show any current enhancements compared to the mediator CV (black vs. purple curves in Fig. S10A), in agreement with the experimental data.

Approach II. The average concentration of hydrogenase in solution at the added concentration of 8.8 pM cells, when neglecting the existence of individual E. coli cells but considering that each cell on average contains 43000 CrHydA1, ³² was calculated to be ~ 0.4 µM. Using the k_cat of 400 s^{− 1 32} and values of other parameters shown in Table S1, simulated catalytic CV also reproduced the experimentally observed current enhancement (Fig. 3C vs Fig. 2B, purple trace) when k_ET ~ 300–400 k_cat. Similar to the results obtained with Approach I, increasing the value of k_ET relative to the k_cat value led to an increase in the catalytic current. Figure 3D shows the dependence of the simulated current responses on the concentration of the hydrogenase corresponding to experimental changes in E. coli (CrHydA1) cells concertation (Fig. 2B). As with Approach I the simulation reproduced the experimental data. The control experiment using E. coli (CrHydA1-PDT) (Fig. 2D) was also successfully reproduced using Approach II (Fig. S10B).

Thus, both Approach I and II allowed to reproduce the experimental data. In both cases, the current enhancement increased with increasing the value of k_ET relative to k_cat (up to very large values of k_ET, at which point k_cat became limiting, as shown in Fig. S11), demonstrating that this is the rate-limiting step of the process. While values of k_ET that reproduced the experimental data were somewhat different for Approach I and II (k_ET ~ 400–500 k_cat vs. k_ET ~ 300–400 k_cat, respectively), when used to calculate the initial rate of the electron transfer reaction (Step 2), they gave the same value of ~ 10^− 5 M/s (Note S1).

We further investigated the effect of the driving force on the rate-limiting step of the process. For this, simulations of experimental current enhancements for both redox couples of DQ and DQ-OH were performed (Note S2, Fig. S12). Since Approach I and II gave the same results, only Approach I was used for these simulations. Figure 4C shows simulated CVs reproducing experimental data (Fig. S2A) and current enhancements for two cathodic peaks (Fig. 4A) obtained with DQ. In order to reproduce experimentally observed shapes of catalytic CVs for both redox couple of DQ (equivalent current increase at both cathodic peaks) for all concentrations of the cells, the electron transfer rate from the fully reduced DQ⁰ to the E. coli (CrHydA1) should be at least an order of magnitude lower than the electron transfer rate from DQ¹⁺ (Fig. S13A). This effectively means that mediation occurs primarily by the DQ²⁺/DQ¹⁺ redox couple. The contribution of the DQ¹⁺/DQ⁰ redox couple to the mediation process would result in a large current enhancement for the second redox peak (Fig S13A, light purple traces), which was not observed experimentally. Figure 4D depicts simulated CVs reproducing experimental data (Fig. S5A) and current enhancements for two cathodic peaks of DQ-OH (Fig. 4C). Cathodic current enhancements in the case of DQ-OH were significantly lower than for DQ for both cathodic peaks and all concentrations of the E. coli (CrHydA1) cells. To reproduce these experimental data k_ET~200 k_cat is required. Similar to DQ, mediation occurs primarily by the DQ-OH²⁺/DQ-OH¹⁺ redox couple (Fig. S13B). The driving force for electron transfer by the M²⁺/M¹⁺ redox couple for DQ-OH is 0.065V lower compared to DQ (Table 1), while the rate of the electron transfer to the E. coli (CrHydA1) is ~ 2.5 times lower for DQ-OH than for DQ.

We have tested four redox compounds as possible mediators for electron transfer from a glassy carbon electrode to the [Fe-Fe] hydrogenase located in the cytoplasm of E. coli cells (Fig. 1). MV, one of the most common mediators used with CrHydA1, did not support cathodic electron transfer but instead resulted in anodic electron flow (Fig. S1). It should be noted that MV has been shown to be an efficient mediator in a photo-biocatalytic system using the same E. coli (CrHydA1) as a whole-cell catalyst.^29,31 This apparent discrepancy can be easily understood by the differences in experimental setup. During CV, the potential applied to the electrode controls the relative concentration of the oxidized and reduced species close to the electrode surface. However, in the bulk solution mostly MV²⁺ is present. In contrast, a large excess of the reduced MV species forms in the photochemical system over time, and this huge excess leads to a significant lowering of the MV solution reduction potential (Fig. S1B).

Under our experimental conditions, all DQ derivatives supported cathodic electron transfer to the cells. However, DQ-CO₂H affected cell integrity leading to the release of hydrogenase to the solution, as evident from the appearance of an electrochemical signal resembling DET to the enzyme (Fig. S4). Between DQ and DQ-OH the highest catalytic activity (39 ± 5% of current enhancement at the peak current) was recorded using DQ as the mediator (Fig. 4A). This finding is in line with DQ featuring the lowest reduction potentials of the investigated mediators (Table 1), and therefore the largest driving force for the electron transfer to hydrogenase. However, when comparing cathodic current increases associated with the two reduction events: M²⁺ →M¹⁺ and M¹⁺ →M⁰ for DQ (Fig. 4A) and DQ-OH (Fig. 4B) at different concentrations of E. coli (CrHydA1) they were found to be nearly identical for each mediator at the two reduction events.

To understand the kinetics of the electron transfer in the developed electromicrobial production system we developed a kinetic model for modeling cyclic voltammetry data, consisting of 3 steps as depicted in Scheme 1. Two different approaches to modeling E. coli cells were tested. In Approach I, the catalytic activity of individual E. coli cells was modeled, while in Approach II, the existence of individual cells was neglected and the average concentration of CrHydA1 was used. Both approaches accurately reproduced experimental data (Fig. 3 vs. Figure 2), leading to the same conclusion that electron transfer from the mediator to the biocatalyst is the rate-limiting step of the overall process. Approach I and Approach II gave different values of k_ET for this step. This is not surprising considering that in Approach I, the cell can be regarded the biocatalyst with an overall k_cat reflecting the total amount of hydrogenase in the cell and a reduction potential defined by the initial cellular electron acceptor. Thus, in Approach I the electron transfer is considered to occur from the mediator to the cell, while in Approach II, the electron transfer is modeled as a reaction between the mediator and hydrogenase. While values of the rate constants are different for the two approaches, the rates of the electron-transfer step (Step 2) calculated using values of these rate constants are strikingly similar for the two approaches (Note S1).

The simulations of DQ and DQ-OH experimental data (Fig. 4), showed that the rate of the electron transfer step depends on the driving forcefor electron transfer from the one-electron reduced form of the mediators, M¹⁺, to the cells. The decrease in the driving force by 0.065 V for DQ-OH²⁺/DQ-OH¹⁺ compared to DQ²⁺/DQ¹⁺ resulted in a corresponding decrease of the electron transfer rate from the M¹⁺ to the E. coli cells by a factor of 2.5. Conversely, no increase in the current enhancement was observed for electron transfer from M⁰ compared to M¹⁺, despite the driving force increasing by ~ 0.2 V for both mediators,, (Fig. 4A and B). In line with observation, the simulations of CV traces recorded in the presence of different concentrations of E. coli (CrHydA1) (Fig. 4C and D), we found that neither DQ⁰ nor DQ-OH⁰ transfer electrons to the cells at experimentally detectable rates. This could be attributed to the limited solubility of fully reduced diquats and their adsorption on the electrode, as previously reported,⁶¹ and observed experimentally in our results (Note S2). Another possibility is unfavorable transport of uncharged species through the E. coli cell membrane.⁶²

Assuming that both DQ¹⁺ and DQ-OH¹⁺ transfer electrons to the active site of CrHydA1, the dependence of the electron transfer rate on the reduction potential of the redox mediator can be calculated using Marcus theory (Note S3). The experimentally observed dependence of the electron transfer rate on the driving force is fully consistent with predictions from Marcus theory. This suggests that the mediator transfers electrons directly to the active site of the hydrogenase inside the cells, making this step the rate-limiting one for the entire process. We note that our experimental data and simulations do not completely rule out the possibility that electrons are first transferred from the mediator to other species inside the cells, which has the redox potential and reorganization energy strikingly similar to those of the active site of CrHydA1, but this scenario is highly unlikely. The importance of the electron transfer step on overall catalytic performance was further supported by the correlation between total H₂ production during bulk electrolysis and the potential of electrolysis, using DQ as the mediator (Fig. S14).

We have designed a mediated electromicrobial production system based on E. coli cells containing CrHydA1 [Fe-Fe] hydrogenase. Using diquat as a mediator we have shown that electrons can be efficiently transferred from the electrode to the cytoplasmic hydrogenase resulting in production of H₂ within the cells. This system can be used as an electron uptake and transport unit, converting electrons into intracellular H₂, which can be readily harvested directly or further serve as a reducing power within the cell to drive E. coli metabolism toward a desired more complex product by employing a suitable uptake hydrogenase. Using cyclic voltammetry and kinetic modeling, we studied the kinetics of electron transfer within the developed system and we propose that the electron is transferred directly from the mediator to the hydrogenase. The rate of this step controls the overall rate of H₂ production, which in turn depends on the driving force provided by the difference between the potential of the mediator couple and the active site of the hydrogenase. The developed systematic approach for investigating the kinetics of electron transfer in electromicrobial production allows us to decipher the contribution of each component of the system to overall performance and can enable more informed use of synthetic biology tools for the production of desired products using microbial electrosynthesis.

Materials and Instruments. All chemicals were purchased from Sigma-Aldrich or VWR and used as received unless otherwise stated. DQ, DQ-OH and DQ-CO₂H were synthetized and characterized as described previously. ^31,63All anaerobic work was performed in an MBRAUN glovebox ([O₂] < 10 ppm). The expression vector encoding the hydA1 gene was kindly provided by Prof. Marc Fontecave (College de France, Paris/CEA, Grenoble). All electrochemical measurements were performed using an Eco/Chemie PGSTAT10 and the GPES software (Metrohm/Autolab).

Preparation of E. coli (CrHydA1)

Synthesis of (Et4N)2[Fe2(pdt)(CO)4(CN)2] ([2Fe]pdt]) and - (Et4N)2[Fe2(adt)(CO)4(CN)2] ([2Fe]adt]). Synthesis of (Et₄N)₂[Fe₂(pdt)(CO)₄(CN)₂] ([2Fe]^pdt]) and - (Et₄N)₂[Fe₂(adt)(CO)₄(CN)₂] ([2Fe]^adt]). The complexes, [2Fe]^{pdt 64–66} and [2Fe]^{adt 67} were synthesized following literature protocols with minor modifications, and verified by FTIR spectroscopy. The complexes were dissolved in anaerobic potassium phosphate buffer (100 mM, pH 6.8) at 1 µg/µL concentration and used directly.

Overexpression of the apo-HydA1 hydrogenase. E. coli BL21 (DE3) cells containing the HydA1 plasmid were grown in 400 mL supplemented M9 medium (22 mM Na₂HPO₄, 22 mM KH₂PO₄, 85 mM NaCl, 18 mM NH₄Cl, 0.2 mM MgSO₄, 0.1 mM CaCl₂, 0.4% (v/v) glucose) under aerobic conditions at 37°C (shaking speed at 170 rpm) until O.D.600 = 0.5 in the presence of antibiotics (100 mg/L ampicillin). The protein overproduction was induced with 1mM Isopropyl-ß-D-thiogalactopyranoside and persisted at 20°C for 15 h with continuous aeration. The final O.D.600 of the cultures was 0.8 ± 0.1. Thereafter, the culture was kept at 4°C for 4 h. This cooling step is critically important to ensure that the cells do not lyse during electrochemical measurements.

In vivo formation of [2Fe]pdt - and [2Fe]adt -HydA1. The preparation of the semi-synthetic hydrogenase was performed following a literature protocol with minor modifications. ⁴²Following overexpression of the apo-HydA1 hydrogenase and subsequent cooling down of the culture, the cells were collected by centrifugation at 5000 rpm. The centrifuge was pre-cooled at 4°C to ensure that the cells did not lyse during the centrifugation process. The cells were gently washed in M9 followed by centrifugation in a pre-cooled centrifuge (4°C). Finally, the cells were resuspended in supplemented M9 solution (4 ml final volume), deaerated, and transferred into a glovebox. Formation of [2Fe]^pdt - or [2Fe]^adt–HydA1 was achieved by treating the cell cultures with [2Fe]^pdt or [2Fe]^adt (final concentration 70–80 µM) at 37°C under strict anaerobic conditions.

Washing of E. coli (CrHydA1) cells. Following the complex insertion, cell suspensions were washed three times in Tris-HCl buffer supplemented with 150 mM NaCl (pH 8). The washed cells were then suspended in 4 ml phosphate-buffered saline (PBS, 100 mM NaH₂PO₄/Na₂HPO₄ with 150 mM NaCl and, pH 6.8) and kept at 4°C overnight (approx. 12h) before performing electrochemical measurements.

Electrochemical Measurements

Cyclic Voltammetry. All CV measurements were carried out in a three-electrode setup, consisting of a glassy carbon (diameter 3 mm) working electrode, a graphite counter electrode, and an Ag/AgCl (1 M KCl) reference electrode. The glassy carbon electrode was polished with 0.5 µm alumina before measurements inside the glovebox. All electrochemical measurements were performed in a glass electrochemical cell inside a nitrogen-purged glove box (O2 < 5 ppm) unless otherwise specified. Each reported data set is a representative of at least two, and typically three or more, biological replicates. At least three CV traces were recorded for each sample, with the second cycle reported. First, 4 ml of oxygen-free 150 mM PBS (pH 6.8) was introduced in the electrochemical cell, and CV was performed at a scan rate of 5mV/s from − 0.4 V to -1 V vs. Ag/AgCl (1 M KCl). Thereafter, 1 ml of 1 mM mediator solution was added to the electrolyte leading to a total electrolyte volume of 5 ml and the final mediator concentration of 200 µM. CV was again performed using the same protocol to record the electrochemical response of the mediators. To this electrolyte solution, the requisite amount of E. coli (CrHydA1) was gently added to obtain a cellular concentration of 2.2 pM. Before this cellular addition, the glassy carbon electrode and Ag/AgCl (1 M KCl) electrodes were taken out of the electrolyte. The electrolyte solution with the added E. coli (CrHydA1) cells was gently pumped using a 1 ml micropipette several times and left undisturbed for 10 minutes. Thereafter, the working and reference electrodes were reintroduced, the electrolyte was pumped again, and CVs were recorded after a rest period of 30 seconds. The second cycle of the CVs was considered for data analysis. After the measurement, the working and reference electrodes were again taken out. The glassy carbon electrode was polished with 0.5 µm alumina. This step is critical as some of the E. coli (CrHydA1) cells stick to the electrode surface and if not cleaned thoroughly, they block the electrode surface in subsequent measurements leading to loss/distortion of the electrochemical behavior of the mediators. The reference electrode is also cleaned with water and stored in 1 M KCl. Following the above-mentioned protocol, appropriate amounts of E. coli (CrHydA1) were added to the electrolyte to achieve final concentrations of 4.4 pM and 8.8 pM, respectively and the measurements were repeated in the same manner. Beyond 8.8 pM, significant precipitation of the E. coli (CrHydA1) cells was found to occur during the electrochemical measurement timescale.

For cell integrity analysis the electrolyte containing cells was centrifuged at 5000 rpm in a pre-cooled (4 °C) centrifuge inside the glovebox for 5 minutes. Thereafter, the supernatant was carefully pipetted out to a clean electrochemical cell and subjected to CV measurements. Both the supernatant and the precipitate were further subjected to SDS-PAGE analysis. The protein contents in the supernatant and the cells were analyzed by 10% SDS-PAGE mini gels in a SE250 Mighty Small II unit (Hoefer) system. The proteins were stained with Page Blue protein staining solution (Thermo Fisher Scientific) according to the suppliers’ instructions. To check the oxygen sensitivity of the system, we performed CV of DQ in the presence of 8.8 pM E. coli (CrHydA1) following the above protocol and thereafter exposed the electrolyte to air by taking it out of the glovebox for 10 or 20 minutes (Fig. S7). The electrolyte was thereafter purged with nitrogen (the needle was kept above the electrolyte surface) for 30 minutes and brought back inside the glovebox to perform CVs. We would like to point out here that though the electrolyte was directly exposed to air for 10 and 20 minutes, the actual exposure time is longer as the deoxygenation took time.

Bulk electrolysis. A gas-tight electrochemical cell with a three-electrode setup, consisting of a glassy carbon rod working electrode, a platinum-wire counter electrode, and an Ag/AgCl (1 M KCl) reference electrode, was used for bulk electrolysis measurements. A 5 mM solution of DQ was prepared with the electrolyte (150 mM PBS, pH 6.8) and degassed by bubbling with N2 for 30 minutes. An O₂-free solution of E. coli (CrHydA1) cells was added to the mediator solution via cannula to achieve a final cell concentration of 20 pM and a total volume of 5 mL in the electrochemical cell. The setup was sealed with a rubber septum to enable gas chromatography sampling.

Chronoamperometry was performed at a potential of -0.9 V or -1.1 V vs. Ag/AgCl (1 M KCl) for 30 minutes under gentle stirring of the solution. The exposed surface area of the glassy carbon rod working electrode was 1.45 cm². Gas chromatography measurements were performed with a PerkinElmer (LLC, MA, USA) instrument equipped with a thermal conductivity detector (TCD) and a stainless steel column with a 60/80 molecular sieve mesh. The operational oven temperature was set to 80°C, while the temperatures of the injection port and TCD were set to 100°C. The GC instrument utilized Ar as the carrier gas (flow rate of 35 mL·min⁻¹) and N₂ as the reference gas. The measured amounts of H₂ were extrapolated to the total headspace in the electrochemical cell (14.2 mL).

Modeling

Simulations of CVs were performed using DigiElch-Professional. A 1-D semi-infinite diffusion model was used for a 0.0707 cm² area planar electrode, with one data point per mV and an x-grid expansion factor of 0.5. The temperature was set to 298.2 K. Parameters used for simulations are presented in Tables S1 and S2.

Acknowledgments

ESA Programme Discovery & Preparation (Contract No. 4000137773/22/NL/GLC/my), the European Innovation Council (Pathfinder project ECOMO, Grant No. 101115403 to N.P.) and Göran Gustafsson Foundation (grant number 2409) are gratefully acknowledged for funding.

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Electrodriven H2 Production in Escherichia coli: Rational Design and Mechanistic Studies of the Electron Uptake Process

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