Energy transfer between the nicotinamide nucleotide transhydrogenase and ATP synthase of Escherichia coli

Membrane bound nicotinamide nucleotide transhydrogenase (TH) catalyses the hydride transfer from NADH to NADP+. Under physiological conditions, this reaction is endergonic and must be energized by the pmf, coupled to transmembrane proton transport. Recent structures of transhydrogenase holoenzymes suggest new mechanistic details, how the long-distance coupling between hydride transfer in the peripheral nucleotide binding sites and the membrane-localized proton transfer occurs that now must be tested experimentally. Here, we provide protocols for the efficient expression and purification of the Escherichia coli transhydrogenase and its reconstitution into liposomes, alone or together with the Escherichia coli F1F0 ATP synthase. We show that E. coli transhydrogenase is a reversible enzyme that can also work as a NADPH-driven proton pump. In liposomes containing both enzymes, NADPH driven H+-transport by TH is sufficient to instantly fuel ATP synthesis, which adds TH to the pool of pmf generating enzymes. If the same liposomes are energized with ATP, NADPH production by TH is stimulated > sixfold both by a pH gradient or a membrane potential. The presented protocols and results reinforce the tight coupling between hydride transfer in the peripheral nucleotide binding sites and transmembrane proton transport and provide powerful tools to investigate their coupling mechanism.

NADH and NADPH and their oxidized counterparts NAD + and NADP + are important cofactors in countless cellular reactions. While NADH is formed during catabolism by many different enzymes and consumed mainly during oxidative phosphorylation, NADPH is majorly regenerated by only a handful of cellular systems found in the cytosol or the mitochondria 1 , their individual contribution depending on the respective tissue 2 , but also the growth conditions 3 . However, it should be noted that there are indications that also other NADPH generating reactions seem to play a role in maintaining the NADPH/NADP + redox balance 3 . A unique system is the proton pumping transhydrogenase (TH) in the mitochondrial inner membrane or the bacterial cytoplasmic membrane which couples the reduction of NADP + to the oxidation of NADH via direct hydride transfer according to the following equation: with "out" meaning the periplasmic space and "in" meaning the cytosol. Although membrane bound TH is not essential in E. coli, it appears to be the major source of NADPH in contrast to most other investigated microbes that rely majorly on the oxidative pentose phosphate pathway and the isocitrate dehydrogenase for NADPH generation 4 . Under physiological conditions, the TH forward reaction is endergonic and requires energization by the pmf 5 . Therefore, TH employs transmembrane proton translocation to energize hydride transfer between NADH and NADP + .
The reaction is completely reversible and can thus also occur in the reverse direction, converting TH to a redox-driven proton pump without redox cofactor 6 . In mitochondria and bacteria, the enzyme exists as a functional dimer with each protomer containing three structural domains (Fig. 1). The hydrophilic domains dI and dIII protrude into the cytoplasmic space and bind NAD(H) (Fig. 1, yellow) and NADP(H) (Fig. 1, red) (1) NADH + NADP + + H + out ↔ NAD + + NADPH + H + www.nature.com/scientificreports/ NADP controls protonation of the critical histidine (βH91 in E. coli) in the channel by influencing its pKa and that conformational changes within the channel are directly linked to dIII attachment or detachment from dII 24 . Both models are based on structures of solubilized enzymes and require experimental verification, especially the proposed interaction between the face-down dIII domain and the presumable proton loading site histidine in the membrane. It is further unclear in which step the pmf, consisting of a pH gradient and the membrane potential Δψ, affects the molecular mechanism. The investigation of the pmf dependent coupling mechanism requires the presence of a tight membrane and the protein inserted in this membrane. Critical amino acid residues are investigated using mutant variants that requires a robust experimental system allowing convenient genetic manipulation and production of the enzyme. In the following, we describe the construction, over-expression and purification of E. coli transhydrogenase, as well as a basic functional characterization based on functional assays in solution.
In addition, we provide protocols for the straight-forward reconstitution of TH alone and in presence of E. coli ATP synthase. We show for the first time that in liposomes containing both enzymes, TH generates a sufficiently large pmf to energize ATP synthesis by the ATP synthase. While some of these experiments have been described before [17][18][19][20] , we are convinced that our modern straight-forward experimental setup is timely to help researchers elucidating the enigmatic molecular mechanism of energy coupled transhydrogenase.

Reconstitution of transhydrogenase into liposomes. Soybean lecithin (90% PC content, Alfa Aesar)
was resuspended in 50 mM HEPES, pH 7.5 100 mM KCl at 5 mg/ml, followed by seven cycles of flash freezing in liquid nitrogen and thawing at 29.4 °C and extruded 21 × through a 200 nm membrane (Whatman® Nuclepore Track-Etched Membrane). Liposomes were destabilized by the addition of octyl β-D-glucopyranoside (OG) to a final concentration of 0.6% followed by addition of 28 μl of purified E. coli transhydrogenase (108 µM) to 1 ml of liposomes and incubation at RT for 15 min on a rotating wheel. The detergent was removed by addition of BioBeads SM 2 (BioRad) to a final concentration of 400 mg/ml and incubation at 4 °C on a rotating wheel for 4 h. Before measurements, the proteoliposomes were mixed 1:1 with the same buffer and kept on ice.
Orientation of reconstituted TH in proteoliposomes was determined using alamethicin which forms a membrane pore allowing NADH to pass through and thereby also reach nucleotide binding sites located on the inside of the proteoliposome. Reverse and cyclic activity of proteoliposomes containing transhydrogenase were determined as described above. Briefly, reverse activity was measured in 1.5 ml 50 mM HEPES, pH 7.5, 100 mM KCl, containing 50 μl proteoliposomes, 1 μM CCCP, 0.2 μM valinomycin, 100 μM NADPH and in presence or absence of 6.7 μg/ml alamethicin. The reaction was started with the addition of 200 μM APAD + and progress of the reaction was monitored at 375 nm. Cyclic activity was measured in 1.5 ml 50 mM HEPES, pH 7.5, 100 mM KCl, containing 50 μl proteoliposomes, 1 μM CCCP, 0.2 μM valinomycin, 200 μM NADH, 10 μM NADP + and in presence or absence of 6.7 μg/ml alamethicin. The reaction was started with the addition of 200 μM APAD + and progress of the reaction was monitored at 375 nm.
NADPH driven proton pumping. Proton pumping during reverse activity of transhydrogenase was measured following ACMA (9-Amino-6-Chloro-2-Methoxyacridine) fluorescence at λ ex/em 419/483 nm in a PTI Quantamaster. For each measurement, 100 μl of liposomes were resuspended in 1.5 ml buffer containing 50 mM HEPES pH 7.5, 100 mM KCl, 180 mU/ml lactate dehydrogenase, 2 μM ACMA, 0.2 μM valinomycin. Depending on the starting conditions, 200 μM NADPH or 500 μM NAD + were also present or added to start the reaction after recording a baseline. The proton gradient was dissipated by addition of NH 4 Cl to a final concentration of 33 mM.
Co-reconstitution of transhydrogenase and ATP synthase. ATP synthase was purified as described 42 . Liposomes were prepared as above and destabilized by addition of 0.6% OG, followed by addition of 28 μl of E. coli transhydrogenase (108 µM) and 28 µl of E. coli ATP synthase (44 μM) per ml liposome. Proteoliposomes were incubated at RT for 15 min on a rotating wheel. Subsequently, BioBeads (SM 2 , BioRad) were added to a final concentration of 400 mg/ml and the sample was incubated at 4 °C on a rotating wheel for 4 h. Before measuring the proteoliposomes were mixed 1:1 with reconstitution buffer (50 HEPES, pH 7.5, 100 mM KCl).
Transhydrogenase forward reaction in the co-reconstituted system. ACMA dequenching assay. ACMA dequenching was measured using proteoliposomes containing E. coli transhydrogenase and ATP synthase. Measurements were done in 1.5 ml 50 mM HEPES, pH 7.5, 100 mM KCl, 100 μl liposomes, 0.2 µM valinomycin, 4 mU/ml glutathione reductase, 1 mM oxidized glutathione, 2 µM ACMA, 500 µM NADP + , and 2 mM MgCl 2 . After recording the baseline, an acidic inside proton gradient was established by addition of 250 µM ATP. After a stable plateau was reached, transhydrogenase was switched on in the forward direction by addition of NADH to a final concentration of 500 µM. The proton gradient was dissipated by addition of 33 mM NH 4 Cl.
Transhydrogenase driven ATP synthesis. ATP synthesis using proteoliposomes containing E. coli transhydrogenase and ATP synthase was measured using a Glomax Luminometer (Promega). For each measurement, 100 μl proteoliposomes were suspended in 400 µl 50 mM HEPES, pH 7.5, 100 mM KCl, 5 mM MgCl 2 , 40 mM KH 2 PO 4 , 0.4 mg/ml CLS II luciferase kit reagent (Roche), 250 µM ADP, and 1.5 mM NAD + . After recording basal ATP levels, the reaction was started by addition of 750 µM NADPH. www.nature.com/scientificreports/ pmf induced switch from reverse to forward transhydrogenase activity. Using liposomes containing ATP synthase and transhydrogenase, the measurements were performed in 1.5 ml 50 mM HEPES, pH 7.5, 100 mM KCl containing 100 μl liposomes along with 4 mM MgCl 2 , and 150 µM NADPH. First, the transhydrogenase reaction was started in reverse direction by addition of 150 µM APAD + . After a plateau was reached, indicating chemical equilibrium, 250 µM ATP was added. The reaction was followed by measuring absorbance at 375 nm to monitor reduction of APAD + on a Cary UV/Vis spectrophotometer.

Results
Cloning and purification. The pntAB operon for E. coli transhydrogenase was subcloned into a pET16b vector, placing the genes under the control of the IPTG-inducible T7 promoter, yielding pET16b-EcTH. The plasmid was transformed into a transhydrogenase knockout E. coli strain (THO), and the protein was overexpressed following IPTG induction. The expression of E. coli transhydrogenase exhibited no significant toxicity, and the yield of purified protein was 5-6 mg/L. Protein expression was verified in native membranes ( Fig. 2A), followed by purification using NTA-affinity chromatography. After a single purification step, the protein was essentially pure as judged by SDS-PAGE and was used for functional experiments without further purification (Fig. 2B). The detergent solubilized protein showed reverse, forward and cyclic activity. In contrast to forward and reverse activity, cyclic activity is not accompanied by proton transport, but the hydride is transferred from NADH over NADP + back to NAD + . Specific activity during the reverse reaction of the purified detergent solubilized enzyme was 5.4 ± 1 μmol/mg/min (n = 7).
Activity measurements and K M determination using detergent solubilized enzyme. In a first set of experiments, we determined the apparent K M of NADPH and NAD + during the reverse reaction under Michaelis-Menten conditions according to the following equation: To this end, NAD + was held constant at 0.5 mM using lactate dehydrogenase and pyruvate, and the K M of NADPH was determined measuring catalytic activity at different NADPH concentrations. For the K M of NAD + , different NAD + concentrations were added in the presence of 100 μM NADPH which was held constant using glucose-6-phosphate dehydrogenase and D-glucose-6-phosphate as a regenerating system (Fig. S1).
Next, we determined the apparent K M values of substrates NADH and NADP + during the forward reaction according to the following equation: www.nature.com/scientificreports/ To determine the K M of NADH, NADP + was held constant at 0.5 mM using glutathione reductase and oxidized glutathione while various concentrations of NADH were added. Determination of K M for NADP + , on the other hand, was obtained using alcohol dehydrogenase to hold NADH concentration constant at 500 μM while adding various concentrations of NADP + (Fig. S1). In all measurements, the activity was followed over time measuring absorption at 340 nm where both NADH and NADPH show an absorption maximum. The K M values determined from these experiments are summarized in Table 1.
Reconstitution of purified TH into preformed liposomes. As described above for the solubilized enzyme, the hydride transfer reaction catalysed by TH is fully reversible and the direction of the reaction is dictated by the relative concentrations of the present substrates. The same is true for the membrane embedded enzyme, but in addition to hydride transfer, proton translocation across the membrane takes place in either direction, depending on the directionality of the reaction. To investigate the coupling of hydride transfer to proton translocation, the enzyme must be embedded into tight vesicles. We prepared soybean lecithin liposomes and incorporated TH via detergent-mediated reconstitution. Trying several reconstitution approaches 43 , we were most successful using 0.6% octyl glucoside to destabilize preformed liposomes and Bio-Beads to remove the detergent after a short incubation of the enzyme with the destabilized liposomes. Other detergents (DDM or cholate for instance) or methods for detergent removal (gel filtration or cyclodextrin) yielded proteoliposomes with lower proton pumping activities. Success of reconstitution was tested monitoring NADPH-driven proton pumping using an ACMA quench assay (Fig. 3A). Reconstitution efficiency of transhydrogenase was estimated using SDS-PAGE, visualizing the steps of the reconstitution process (soluble enzyme, low spin supernatant, high spin supernatant and pellet). Following this approach, an incorporation efficiency of 65 ± 10% is estimated. Orientation of the reconstituted TH was estimated comparing activities of outwards oriented enzymes and total enzyme. To this end, reverse TH activity was measured in the absence and in presence of the membrane poreforming peptide alamethicin that allows the membrane impermeable nucleotides to reach the binding sites of inwards-oriented TH. The same experiment was also performed following cyclic activity (Fig. S2). These measurements revealed a preferential orientation of TH with the nucleotide binding sites situated on the outside of the liposomes (75 ± 5% using reverse activity, 85 ± 5% using cyclic activity).

NADPH-driven pmf generation and ATP synthesis. We start with the description of experiments
where liposome-reconstituted TH was run in the reverse direction, i.e. where transhydrogenase acts as a NADPH driven proton pump. Proton pumping was followed using the fluorescent dye 9-amino-6-chloro-2-methoxyacridine (ACMA) which is quenched upon the build-up of an acidic inside pH gradient (Fig. 3A). The reverse reaction is either started by adding NADPH or NAD + (while the other is present from the beginning), leading to a decrease in ACMA fluorescence. In Fig. 3A, the observed ACMA quench was slightly slower and progressed to a lesser maximal quench when the reaction was started with NADPH instead of NAD + , however this was not observed in all measurements. If mutant variant H91A was reconstituted, no ACMA quench was observed, supporting the essential role of the H91 for proton pumping (Fig. 3A).
In many bacteria and mitochondria, the membrane potential is the dominant component of the pmf (~ − 120 to − 160 mV) while the pH gradient is typically < 0.5 pH units. This makes the membrane potential generated by respiratory complexes the primary driving force for the endergonic forward reaction under cellular conditions, where the NADPH/NADP + is much larger than the NADH/NAD + ratio. We have therefore mimicked this scenario by co-reconstituting TH with ATP synthase from E. coli. ATP synthase has the advantage that it can readily be energized with the non-redox active substrate ATP and shows a high unidirectionality of incorporation into liposomes (F 1 head located to the outside) 42 . We have confirmed the successful co-reconstitution of the two enzymes by a series of experiments. First, we utilized TH as a redox-driven proton pump to establish a pmf (inside positive and acidic, as described above) and followed if the established pmf is sufficient to support ATP synthesis by ATP synthase. As seen in Fig. 3B, upon addition of NADPH, continuous ATP synthesis is observed using a luciferin/luciferase-based ATP detection system. No ATP synthesis was observed, if H91A variant was reconstituted alongside ATP synthase.

ATP hydrolysis driven pmf generation and NADPH generation by TH.
As stated in the introduction, under cellular conditions, the redox potential of the NADPH/NADP + couple is more negative than the NADH/NAD + couple, and the pmf is required to push the reaction towards NADPH generation (forward reac- www.nature.com/scientificreports/ tion). If the pmf is thereby not constantly supplied, the reaction will halt once it has reached its thermodynamic equilibrium. ATP hydrolysis driven proton pumping could represent such as source of pmf generation and was mimicked in the following experiment. NADPH and APAD + (an NAD + analogue and substrate of TH) were added to proteoliposomes containing both enzymes and the formation of APADH was followed spectrophotometrically at 375 nm. After a few minutes, the increase in absorption stopped indicating that the reaction reached its chemical equilibrium, i.e. rates of the forward and reverse reactions are identical due to the relative substrate and product concentrations and the established pmf. At this point, addition of ATP energized inward proton pumping by ATP synthase, increasing the inside positive membrane potential. Consequently, the reaction moved away from equilibrium, catalysing the forward reaction of TH and thus NADPH formation and APADH consumption which can be observed as a decrease in absorption at 375 nm (Fig. 3C).

NADH-driven Δψ generation by TH.
Next, we designed experiments to monitor the forward activity of TH in proteoliposomes under different conditions. Using suitable substrate concentrations that shift the equilibrium of the reaction (high NADH and NADP + , low NADPH and NAD + ), the transhydrogenase can be run in the forward direction. Since only proteoliposomes with TH enzymes having their nucleotide binding domains on the outside will be activated, addition of NADH and NADP + will result in proton ejection from the proteoliposomes. Forward activity is therefore expected to establish a positive outside membrane potential, which can be monitored using the fluorescent potential-sensitive dye tetramethylrhodamine ethyl ester (TMRE). As depicted in Fig. 4A, the generation of a membrane potential is observed as a decrease in TMRE fluorescence. No signal shift was observed with H91A. www.nature.com/scientificreports/ TH forward activity driven by an ATP-driven proton pump. To mimic a more natural situation, a proton pump such as the ATP synthase can be used as a constant pmf generator that energizes forward activity of the transhydrogenase enzyme. In the following, two different experimental setups using proteoliposomes containing both TH and ATP synthase from E. coli have been used to follow forward TH activity. In the first, proton pumping by either enzyme was detected by ACMA fluorescence. First, ATP synthase was run as a proton pump, establishing an inside acidic pH gradient observed as a quench in ACMA fluorescence (See Fig. 4B). Addition of valinomycin in the presence of potassium ensured that no opposing membrane potential was established during proton pumping. Addition of the substrates NADH and NADP + drives TH in the forward direction which is accompanied by extrusion of protons from the lumen that is in turn observed as an increase in ACMA fluorescence (dequenching) (Fig. 4B). As both systems are continuously working, a new equilibrium is established.
In the second setup, we have reproduced earlier findings 19,22,44 that demonstrate the direct effect of a pmf on the rate of NADPH formation during the forward reaction. The experiments were started by addition of NADH and NADP + either in the absence and presence of valinomycin, as well as in presence of valinomycin and CCCP. In all cases, a continuous production of NADPH was observed (Fig. 4C). Initial NADPH production was fastest in proteoliposomes with supplied valinomycin and CCCP (no opposing pmf was generated), whereas liposomes having no ionophores or valinomycin alone showed both lower but very similar NADPH production rates (opposing pmf generated in form of a ΔpH which is negative and alkaline inside). Addition of ATP triggered  www.nature.com/scientificreports/ proton influx into the liposomes by the ATP synthase. This established a supportive pmf (acidic and positive inside) in form of a ∆Ψ and ∆pH in the proteoliposomes without ionophores, whereas in the proteoliposomes supplied with valinomycin the pmf was established in form of a ∆pH alone. In the proteoliposomes supplied with valinomycin and CCCP, addition of ATP is not expected to have a significant effect since the formation of a ∆Ψ and/or a ∆pH is prevented. Addition of ATP increased the rate of NADPH synthesis 7 ± 1.8-fold in the sample without any ionophore and 6.4 ± 0.8-fold in the sample with added valinomycin (Fig. 4C). In presence of both valinomycin and CCCP the stimulation of the forward reaction was essentially lost (1.6 ± 0.3-fold) (Fig. 4C).

Discussion
Here, we describe the efficient overexpression and Ni-NTA affinity-based purification of the E. coli transhydrogenase. The enzyme is obtained in good yields and shows a specific activity of around 6 µmol/mg/min. We have determined the K M values for its natural substrates under similar conditions in the reverse and forward direction, which to our knowledge, has not been described so far. The values are in the same range as previously published values for the bovine enzyme 18 47 . In general, the found values deviate from each other considerably, but they typically agree on the observation that the K M is lower for the reduced species. Direct comparison of such values is difficult, as the experimental conditions among reports vary significantly, e.g. purified enzyme or membrane vesicles, use of nucleotide analogues or regenerative systems which are required to allow the use of the natural substrates 47 . In summary, the high yield and specific activity as well as the sensible range of K M values obtained with our preparation makes it a suitable expression and purification system for functional (and structural) investigations. Most importantly, it is critical to better understand the functional coupling between the hydride transfer in the soluble dI/dIII interface and the proton transfer through the membrane embedded dII domain. These experiments require the presence of a lipid environment, and our protocols for efficient reconstitution of TH (and ATP synthase) facilitate this critical step. Our measurements showed that a majority (75-85%) of the enzymes is oriented with its large soluble nucleotide binding domains towards the outside. Given the method used for reconstitution, destabilization of preformed liposomes using a defined detergent concentration, this preferred orientation is not surprising 43 . Using TMRE and ACMA fluorescence we show that transhydrogenase establishes a pmf by translocating protons across the membrane, both when run in the forward and reverse direction. The establishment of a positive inside proton gradient in proteoliposomes during the reverse reaction has been shown before for the bovine enzyme 17,19,21,48 as well as for the E. coli enzyme 21,22,31,32,46,47,49,50 . We now show that the pmf generated during the reverse reaction is sufficient to sustain ATP synthesis by ATP synthase co-reconstituted in the same liposomes. In our experiments, we have used rather high numbers of enzymes per liposomes, yielding only liposomes containing several copies of both types of proteins. The use of non-membrane permeable substrates ensures that only one type of proteoliposomes is functionally relevant. In those, the relative orientation of TH and ATP synthase (both nucleotide binding sites on the same side of the membrane) also reflects the physiological situation. An interesting observation is the almost immediate start of ATP synthesis after NADPH addition. This is noteworthy as compared to typical respiratory enzymes such as complex IV (300 to 1000 H + /s), the turnover rate of TH is rather slow (90 ± 17 H + /s). The lack of an initial delay indicates that the requirement of ATP synthesis (pmf > 120 mV and an inside pH < 6.5) is met very quickly (much quicker than the corresponding ACMA signal), supporting the idea of a local distribution of the pumped protons along the membrane surface 51,52 . Co-reconstitution of TH and ATP synthase from bovine mitochondria was described already earlier by Rydström et al. 18 . According to them, despite repeated attempts they were not able to observe ATP synthesis driven by NADPH driven proton pumping and potential issues are discussed 8 . Our data thus fill this important gap, and demonstrates that TH could potentially act as a pmf generator sufficient to energize ATP synthesis in the absence of other suitable reducing equivalents. Using TMRE fluorescence, we also show the formation of a positive outside membrane potential during the forward reaction by reconstituted transhydrogenase using the appropriate substrate concentrations, reinforcing the view that hydride transfer is coupled to proton transport also under the most favourable conditions e.g. nucleotide concentrations.
Our experimental system also allows to observe the stimulating effect of a membrane potential by ATP synthase on transhydrogenase forward activity that has been demonstrated before for the reconstituted bovine enzymes. We find a stimulating rate of approximately 7 in absence of valinomycin which is lower than that found for the reconstituted bovine enzymes by Eytan et al. 19 but similar to the one found for the reconstituted bovine enzymes by Jolla et al. 21 . Such experiments are important for the investigation of mutant variants that affect the coupling between the two spatially separated reactions. Thanks to the new enzyme structures, more refined mechanistic models have been postulated that now must be biochemically validated. Yet another question, e.g. which step of the proton transport (loading and/or unloading of the proton loading site) is affected by which component of the pmf has not yet been discussed or approached at all. Here, we experimentally show that the direction of the reaction can be switched from reverse to forward by applying a pmf, once the reaction has reached equilibrium. This finding supports the idea that under physiologic conditions, when the relevant nucleotide concentration are rather constant, transhydrogenase direction and activity is mainly controlled by the presence and extent of the pmf. This experimental setup should be useful to assess mutant variants that have an altered proton pathway. www.nature.com/scientificreports/ Finally, we present an experiment in which TH forward activity was assessed using ACMA fluorescence. This setup has the unique advantage that both reaction directions can be observed using the natural substrates and followed via transmembrane proton transport, while otherwise nucleotide analogues such as APAD were necessary. In co-reconstituted liposomes, ATP synthase first pumps protons into the lumen which is seen as a large quench in ACMA fluorescence. Addition of NADH and NADP + enables TH to catalyse the forward reaction which is coupled to proton translocation to the outside, seen as a dequench in ACMA fluorescence. It should be noted that in this experiment (Fig. 4B), the rate and the level of pmf established by the ATP synthase are identical with either co-reconstituted wt or the non-pumping βH91A mutant. The latter lacks the key histidine and has an interrupted proton translocation pathway that prohibits transmembrane proton transfer 24,26 . Only upon addition of substrates for transhydrogenase forward activity, a change in the signal is observed for the wt enzyme, implying that the transhydrogenase proton translocation channel is gated (by the nucleotides binding sites) to prevent short circuiting of protons and thus loss of pmf. In other words, there are no substantial leaks in the proton channel of wt transhydrogenase when it is not activated e.g. in absence of all the required substrates.

Concluding remarks
In summary, we used established as well as new techniques to monitor transhydrogenase activity, including proton translocation as well as dependence on a membrane potential. The described protocols are straightforward allowing fast handling of this rather delicate enzyme and should provide a good basis for biochemical, biophysical and structural experiments. The different experimental setups provide a broad toolset to investigate different functional aspects of potentially interesting transhydrogenase mutant variants. Liposome experiments are crucial for the investigation of the capability of transhydrogenase to establish and maintain a membrane potential as well as the sensibility towards the presence of a membrane potential when certain key residues are altered. An interesting experiment in this perspective would be to investigate the sensitivity towards the presence of a membrane potential of transhydrogenase mutant variants which have been shown to have impaired activity and/or proton pumping capability, such as mutations of residue αY226 in E. coli which is part of the NAD(H) binding pocket 53 or the critical histidine (βH91) which lies in the proton translocation channel 26,27,54 . Another intriguing setup would be to coreconstitute the membrane portion of transhydrogenase alone and together with ATP synthase. This would allow to investigate whether the channel is open or closed when the soluble domains are absent. Despite the large set of functional data and the recent protein structures, intimate details of this unique molecular mechanism are still to be discovered.