Proton transfer reactions in the red light-activatable channelrhodopsin variant ReaChR and their relevance for its function

Channelrhodopsins (ChRs) are light-gated ion channels widely used for activating selected cells in large cellular networks. ChR variants with a red-shifted absorption maximum, such as the modified Volvox carteri ChR1 red-activatable channelrhodopsin (“ReaChR,” λmax = 527 nm), are of particular interest because longer wavelengths allow optical excitation of cells in deeper layers of organic tissue. In all ChRs investigated so far, proton transfer reactions and hydrogen bond changes are crucial for the formation of the ion-conducting pore and the selectivity for protons versus cations, such as Na+, K+, and Ca2+ (1). By using a combination of electrophysiological measurements and UV-visible and FTIR spectroscopy, we characterized the proton transfer events in the photocycle of ReaChR and describe their relevance for its function. 1) The central gate residue Glu130 (Glu90 in Chlamydomonas reinhardtii (Cr) ChR2) (i) undergoes a hydrogen bond change in D → K transition and (ii) deprotonates in K → M transition. Its negative charge in the open state is decisive for proton selectivity. 2) The counter-ion Asp293 (Asp253 in CrChR2) receives the retinal Schiff base proton during M-state formation. Starting from M, a photocycle branching occurs involving (i) a direct M → D transition and (ii) formation of late photointermediates N and O. 3) The DC pair residue Asp196 (Asp156 in CrChR2) deprotonates in N → O transition. Interestingly, the D196N mutation increases 15-syn-retinal at the expense of 15-anti, which is the predominant isomer in the wild type, and abolishes the peak current in electrophysiological measurements. This suggests that the peak current is formed by 15-anti species, whereas 15-syn species contribute only to the stationary current.

photocycle model is well based on experimental data, the actual determinants for the equilibrium between syn and anti cycle remain elusive.
The light-induced photocycles of ChRs involve a number of changes in hydrogen bonding and proton transfers that affect a variety of channel properties, such as ion selectivity, conductivity, and photocycle kinetics. The photocycle of the best studied ChR, ChR2 from Chlamydomonas reinhardtii (CrChR2), involves proton dynamics of the central gate residue Glu 90 (Glu 130 in ReaChR; Fig. 1), the counter-ion residue Asp 253 (Asp 293 in ReaChR), and the DC pair residue Asp 156 (Asp 196 in ReaChR). Glu 90 plays a major role for both formation of the conducting pore and ion selectivity (20,21). Asp 253 , in the following referred to as Ci2 (counter-ion 2), receives the RSBH ϩ proton that is released prior to channel opening (22). Alternatively, it was proposed that both Ci1 (Glu 123 ) and Ci2 (Asp 253 ) serve as proton acceptors (21). Furthermore, it was suggested that the DC pair residue Asp 156 reprotonates the RSB in the P390 3 P520 transition (M 3 N in ReaChR) (22) as mutations of this residue highly decelerate the photocycle kinetics (23). However, it is unclear whether the proton dynamics of CrChR2 also apply to red-shifted ChRs, such as ReaChR, as the bathochromically shifted absorption maximum points to molecular alterations near the retinal-binding pocket and the active site as compared with CrChR2; e.g. threonine 159, in the direct environment of Asp 156 , is exchanged for a cysteine in ReaChR (Cys 199 ).
The present study, based on a combination of FTIR difference spectroscopy, time-resolved UV-visible spectroscopy, electrophysiological measurements, and site-directed mutagenesis, showed that Glu 130 , Ci2, and the DC pair residue Asp 196 (Fig. 1) also experience proton dynamics in ReaChR. Our data show that the Glu 90 -helix 2-tilt-model for CrChR2 (21) applies to ReaChR as well. Additionally, we now present a more detailed mechanism for ReaChR and show that deprotonation of Glu 130 occurs in two distinct mechanistic steps. It undergoes a change in hydrogen bonding in the D 3 K transition and deprotonates before formation of the M-state. Consistent with CrChR2, the M-state (P390 in CrChR2) is formed by a proton transfer from the RSBH ϩ to Ci2, but after M formation a photocycle branching occurs that was not observed in CrChR2 WT. The main branch involves M 3 N 3 O transitions, and water is likely to serve as the proton donor for the reprotonation of the RSB during M 3 N transition because no amino acid residue could be identified as proton donor. In parallel, a direct M 3 D transition, involving reprotonation of the RSB by Ci2, takes place.
In either case, Asp 196 does not serve as the proton donor for reprotonation of the RSB in ReaChR as its homologue Asp 156 in CrChR2 does (22). Instead, Asp 196 deprotonates in the N 3 O transition. Neutralization of Asp 156 as in the D196N mutant increases DЈ at the expense of D under conditions of continuous illumination and abolishes the peak current in electrophysiological measurements. This interesting correlation provides experimental evidence for the concept that the peak current is mainly formed by a conducting state with 13-cis,15-anti-retinal conformation, whereas the stationary current contains contri-butions of both 13-cis,15-anti-and 13-trans,15-syn-retinal isomers.

Electrophysiological characterization of key residues
To characterize ion conductance, we expressed ReaChR WT and selected mutants in HEK cells. We mutated the two RSB counter-ions Ci1 (Glu 163 ) and Ci2 (Asp 293 ), the central gate residue Glu 130 , and the DC pair residue Asp 196 . A structural model of the protein region with these residues is given in Fig. 1. The amplitudes of both transient and stationary photocurrent (I t and I s , respectively), of E163T resemble WT-like currents, whereas I s of E130Q and D293N at standard conditions decreased to 41 and 8%, respectively (Fig. 2, A and B). In E163T, the decay of the photocurrent after activation was accelerated ( Fig. 2C; 34 Ϯ 2 ms versus 132 Ϯ 7 ms of WT) in line with findings for homologous mutations in CrChR2 (WT, off ϭ 9.8 ms; E123T, off ϭ 5.2 ms (24)) and the CrChR1/2 chimera (C1C2-E162A) (25). Off-kinetics were moderately accelerated in E130Q (89 Ϯ 7 ms) similar to E90A in CrChR2 (26).
Most striking, however, was the lack of the transient current I t during illumination in the DC pair mutant D196N, whereas the off-kinetics (202 Ϯ 23 ms) of wild type and mutant are in the same order of magnitude. This is in contrast to the homologous mutations in CrChR2 where both the off-kinetics (WT, off ϭ 10 ms; D156A, off Ͼ 1.5 ϫ 10 5 ms (23)) and the photocurrents are significantly different (23,27).
As expected from previous studies on CrChR2 (20,28), the central gate mutant E130Q conducts less protons in favor of Na ϩ , most clearly seen from the enlarged current and shifted reversal potential upon replacement of Na ϩ by N-methyl-Dglucamine (NMG) at extracellular pH 9 (Fig. 2, D-F). This suggests that Glu 130 , like in other ChRs, is part of the central selectivity filter. Based on this functional characterization, we asked whether there are internal alterations of the proton conducting network within the conducting pore and how these changes might be linked to changes of selectivity and to photocurrent inactivation and channel closure after light switched off.

Steady-state and time-resolved UV-visible spectroscopy of selected mutants
Next, light adaptation and photocycle intermediates of selected mutants were characterized by UV-visible spectros-  (50)) is based on the 3D structure of C1C2 (Protein Data Bank code 3ug9). Protonation states were estimated based on a pK a calculation (64).

Proton transfer reactions in ReaChR
copy and compared with the wild type (11). The IDA, i.e. the state before any light exposure of the protein, is slightly redshifted for the counter-ion mutants, E163T ( max ϭ 530 nm) and D293N ( max ϭ 528 nm), with respect to the wild type ( max ϭ 527 nm; Fig. 3A). However, the bathochromic shift is minor compared with CrChR2, C1C2, and ChR2 from Platymonas subcordiformis (PsChR2), which show 10 -25-nm shifts upon counter-ion neutralization (29,30). In contrast, the IDA of E130Q ( max ϭ 513 nm) and D196N ( max ϭ 523 nm) are blue-shifted, which was not reported for analogous mutations in CrChR2 (23,31). Upon extended illumination (60 s, 530 nm) and subsequent recovery in the dark (10 min), the DA app is formed. Corresponding UV-visible spectra of all samples except E163T are shifted toward shorter wavelengths. A likely explanation for these hypsochromic shifts from IDA to DA app is a relative increase of 15-syn-retinal (DЈ) compared with 15-anti-retinal (D) in DA app assuming a blue-shifted absorption of the underlying 13-cis,15-syn chromophore as observed in bacteriorhodopsin (32)(33)(34)(35). This effect is most pronounced in D196N (shift from 523 to 506 nm).
To further evaluate this interpretation, FTIR difference spectra of the wild type (gray filled curves), E130Q, E163T, D293N, and D196N were recorded (Fig. 3B). In the retinal fingerprint region, two negative bands at 1234 and 1199 cm Ϫ1 in the WT spectrum reflect depletion of the 13-trans,15-anti dark state (D) (18,36), and the positive band at 1176 cm Ϫ1 is assigned to 13-trans-retinal (37)(38)(39). This band pattern is only slightly altered by mutations of Glu 130 , Ci1, and Ci2. However, in the D196N spectrum, an additional negative band at 1183 cm Ϫ1 is present, indicating a 13-cis,15-syn-retinal dark state (DЈ) (39 -42). Additionally, a more pronounced positive band at 1173 cm Ϫ1 points to the 13-trans photoproduct of a photoconverted DЈ state. This finding implies that 1) DA app is composed of a mixture of 13-trans,15-anti-retinal (D) and 13-cis,15-syn-retinal (DЈ) isomers and 2) the contribution of the syn conformation (DЈ) to DA app is relatively increased in the D196N mutant. D and DЈ, which together form DA app , undergo light-induced photocycles comprising the intermediates K, L, M, N, and O as well as KЈ, LЈ, MЈ, NЈ, and OЈ, respectively (Fig. 3C, left) (11). Due to similar absorption maxima, which are not more  D196N. A, representative photocurrent traces recorded upon 500-ms illumination. B, average size of peak (left) and stationary (right) photocurrents; significance was tested with the Mann-Whitney U test. *, 0.05 Ͼ p Ͼ 0.01; **, 0.01 Ͼ p Ͼ 0.001; ns, not significant. C, time constants of apparent closing kinetics after light switched off; wild type, E130Q, and D196N were fitted monoexponentially; E163T was fitted biexponentially; and due to decreased currents there is no value for D293N. D, current-voltage relation of wild type (black) and E130Q (gray) under different extracellular ionic conditions (intracellular NaCl, pH 7.2) with corresponding I-E plot (normalized to standard conditions); holding potential (E hold ) ϭ 0 mV in red. E, calculated reversal potentials (E rev ) for wild type (filled bars) and E130Q (empty bars) (liquid junction potentialcorrected; n ϭ 5-8). F, reversal potential shifts (⌬E rev ) with respect to standard conditions (NaCl, pH 7.2) after substitution of extracellular Na ϩ for nonpermeable NMG (NMG, pH 7.2; black), reduction of extracellular protons (NaCl, pH 9.0; dark gray), and reduction of both extracellular NaCl and protons (NMG, pH 9.0; light gray) (n ϭ 5-8; liquid junction potential-corrected). Error bars represent standard errors (S. E.).

Proton transfer reactions in ReaChR
than 10 nm shifted (39), it is not a simple task to separately investigate both photocycles. Additionally, the contribution of the syn photocycle is small as compared with the anti photocycle because 1) ReaChR WT populates only around 20% of DЈ in DA app (11) and 2) a lower quantum efficiency of the photoactivation of the 13-cis,15-syn DЈ is assumed (43). Therefore, the observed photocycle dynamics and half-life times are pri-marily assigned to the anti photocycle branch. The influence of key residues on the photocycle was studied by flash photolysis experiments on the mutants E130Q, E163T, D196N, and D293N (Fig. 3, D-G). Purified proteins were excited with green laser flashes (10 ns, 530 nm, 5 mJ), and UV-visible absorption changes were recorded. Half-life times (t1 ⁄ 2 ) were derived from global analysis (table in Fig. 3C; wild type values are taken from  Ref. 11 (11), whereas it is superimposed by the strong-absorbing dark state at pH 7.4 ( max ϭ 527 nm). This general reaction sequence of the wild type is also found in the mutants at pH 7.4 but with significant variations. L is not observed in E130Q (Fig. 3D) and D196N (Fig.  3F), but all mutants show the N-state. The latter finding could again be explained by the hypsochromic shift of DA app compared with the N-state for E130Q and D196N and by the reduced full width at half-maximum of the chromophore peaks of E163T and D293N. Neutralization of Ci1 or Ci2, either E163T (

Monitoring proton transfer processes and changes in hydrogen bonding by FTIR difference spectroscopy
The formation of the photocycle intermediates is accompanied by changes of hydrogen bonding and/or proton transfer processes involving acidic and alkaline amino acid side chains. To address the question in which stage of the photocycle these events occur and to elucidate the role of specific amino acids, FTIR difference spectra of WT and selected mutants were recorded at cryogenic temperatures to stabilize photocycle intermediates. The steady-state spectrum obtained at 80 K mainly represents the transition from the dark state to the early K intermediate (11). Fig. 4A shows the FTIR difference spectra of the D 3 K transition in H 2 O (gray filled curves) and in D 2 O buffer (black) of ReaChR WT and the mutants E130Q, E163T, D293N, and D196N.

Early proton dynamics in Glu 130
Bands in the spectral region between 1800 and 1700 cm Ϫ1 are predominantly caused by carbonyl CϭO stretching modes of protonated acidic amino acid side chains, and therefore corresponding difference bands reflect changes in hydrogen bonding strength and/or proton transfer processes. In the WT spectrum (Fig. 4A), a prominent difference band at 1729(ϩ)/1721(Ϫ) cm Ϫ1 and a positive band at 1713 cm Ϫ1 are observed. In D 2 O, the negative band at 1721 cm Ϫ1 is downshifted to 1713 cm Ϫ1 , whereas its positive counterpart at 1729 cm Ϫ1 is shifted to 1721 cm Ϫ1 . However, this band maximum is superimposed by the negative band at 1721 cm Ϫ1 due to incomplete deuteration so that two apparent maxima at 1725 and 1719 cm Ϫ1 are seen. The band intensities and positions of the 1729(ϩ)/ 1721(Ϫ) cm Ϫ1 difference band in the wild type are only slightly affected by mutations of the counter-ions, E163T and D293N, or by the DC pair mutation D196N. As this difference band is completely absent in E130Q, it is assigned to a vibration of Glu 130 . The characteristic pattern with one positive and one negative band suggests that Glu 130 undergoes a hydrogen bond change in the D 3 K transition rather than a deprotonation, which would be indicated by only one single negative band.
The impact of the D196N mutation on this difference band might be explained by its influence on the ratio of 15-anti and 15-syn retinal (see Fig. 3B). The slight effect of E163T and D293N on the Glu 130 difference band is explained by an altered interaction of these residues with Glu 130 . In the E163T spectrum, a negative band at 1712 cm Ϫ1 instead of the positive WT band at 1713 cm Ϫ1 is observed. The origin of the positive wild type band is unclear; however, protonation of Ci1 is unlikely because in that case the positive band would be absent in the spectrum of the E163T mutant rather than being replaced by a negative band.

Proton transfer reactions in ReaChR Proton transfer reactions in later photocycle intermediates
In the FTIR difference spectrum of the wild type at 293 K representing the O-state with minor contributions of M, N, and L (Fig. 4B, gray filled curve), we observe a band pattern at 1753(Ϫ)/1738(ϩ)/1723(Ϫ) cm Ϫ1 . The spectra of the mutants (Fig. 4B) support the assignment of the 1723 cm Ϫ1 band to Glu 130 because it is absent only in E130Q. In D 2 O, this negative band is downshifted by 11 cm Ϫ1 to 1712 cm Ϫ1 . The residual band at 1723 cm Ϫ1 indicates only partial sample deuteration. The existence of only one band assigned to Glu 130 instead of a difference band as observed in the spectrum at 80 K (Fig. 4A) now shows deprotonation of Glu 130 . Deprotonation of the homologous residue Glu 90 was observed by various authors for the photocycle of CrChR2, although it is still under debate in which intermediate this deprotonation occurs (20 -22).
The positive band at 1738 cm Ϫ1 is affected by all mutants analyzed. Its intensity is significantly reduced in both the E163T and D293N mutants and thus reflects protonation of Ci2 during M-state formation. The influence of both mutations on the 1738 cm Ϫ1 band can be explained by an accelerated proton release from the unmutated counter-ion residue due to a lowered pK a value if the other respective residue is mutated. In E130Q, instead of one positive band at 1738 cm Ϫ1 , two positive bands arise at 1739 and 1731 cm Ϫ1 . This observation is explained by an overlap of the positive counter-ion band with a negative band that arises in E130Q only, possibly due to deprotonation of a so far unidentified residue.
In the spectrum of the D196N mutant, the band at 1738 cm Ϫ1 is upshifted to 1748 cm Ϫ1 . The high upshift of this band can be explained by the influence of this mutation on the ratio of the 15-anti and 15-syn photocycle branches (see Fig. 3B), leading to a weaker chromophore-counter-ion interaction in line with the accumulation of the red-shifted N-intermediate in the steady state (Fig. 3F).
At 293 K, a negative band arises at 1753 cm Ϫ1 in the WT spectrum that is not observed in the spectrum of any mutant. An assignment of this band is nevertheless possible based on the spectra at 260 K. In these spectra, this vibration of the WT is slightly altered in E130Q, E163T, and D293N, but D196N is the only mutant in which it is not observed (Fig. 4B, inset column). Thus, we assign this band to a vibration of Asp 196 , reflecting deprotonation of this residue. Interestingly, this band is not D 2 O-sensitive, indicating that Asp 196 is buried within the receptor and not accessible to bulk water, similar to observations made for C1C2 (36). The absence of this band in any mutant at 293 K indicates that Asp 196 is already reprotonated in the steady state of the mutants at this temperature, which is in agreement with faster photocycle progression of the mutants compared with WT (Fig. 3C).

M-state-related proton transfer
Although proton transfer reactions involving Glu 130 , the counter-ion complex, and Asp 196 were shown to occur in the photocycle of ReaChR, it is still unclear in which photocycle intermediate(s) these transfers actually occur. Accordingly, we investigated proton transfers related to M-state formation and decay. The slow-cycling DC pair mutant C168S (C128S in CrChR2) is suitable to investigate proton transfer reactions in the M-state because it highly accumulates M upon illumination (60 s, 530 nm) in the steady state at 293 K (Fig. 5A, inset). In the FTIR difference spectra of C168S in H 2 O (orange) and D 2 O (magenta) reflecting D 3 M transition (Fig. 5A), the positive band at 1738 cm Ϫ1 , which was assigned to the protonation of Ci2 (Fig. 4B), is split into two positive bands arising at 1742 and 1729 cm Ϫ1 , similar to the E130Q mutant (Fig. 4B). This observation can again be explained by an overlap of a single positive counter-ion band with an additional negative band arising in the M-state. The negative band at 1721 cm Ϫ1 assigned to deprotonation of Glu 130 is downshifted to 1711 cm Ϫ1 in D 2 O, similar to WT. This indicates that Glu 130 deprotonates at the latest during M-formation. An early deprotonation of the homologous residue Glu 90 was observed during the photocycle of CrChR2 (21). Asp 196 is not deprotonated in M because the spectra lack the negative band at 1753 cm Ϫ1 .
Ion conductance of channelrhodopsins can be rapidly switched on and off with alternating illumination of suitable wavelengths as shown in electrical studies (44,45). We applied a similar illumination protocol with alternating green (520 nm) and UV (390 nm) light to the wild type. To estimate the independent spectral components contributing to the signal change, we evaluated the results using a combination of singular value decomposition (SVD) and a rotation procedure (46,47) as performed for CrChR2-C128T (15) (Fig. 5B and see "Experimental procedures" for details). The first component represents formation of the late intermediates, mainly O, which can be derived from similarity of this spectral component to the steady-state spectrum at 293 K (Fig. 4B, gray filled curves). This component displays a negative band at 1753 cm Ϫ1 , indicating deprotonation of Asp 196 . It arises with green illumination, remains throughout the whole illumination period, and slowly decays after the end of the illumination. A further component strictly follows the illumination protocol. Green illumination induces a steady state including M, which can be depopulated by UV light, so that formation and decay of M, which contains contributions of both M 1 and M 2 (see Fig. 3, H-J), can be directly observed. This component involves a difference band at 1766(ϩ)/1756(Ϫ) cm Ϫ1 , which demonstrates a change of hydrogen bonding of still protonated Asp 196 coincident with formation and decay of M, whereas the negative band in the first component confirms later deprotonation of Asp 196 , i.e. after formation of the N-state. Thus, it can be excluded that Asp 196 serves as proton donor for the RSB during M decay in contrast to the homologue Asp 156 in CrChR2 (22). Another likely candidate for reprotonation of the RSB is Ci2, which receives the RSBH ϩ proton during M formation. Indeed, the component following the illumination protocol comprises a band pattern with two maxima at 1744 and 1731 cm Ϫ1 that was already assigned to Ci2 protonation (see Fig. 5A). This shows that Ci2 receives a proton during M formation and releases it during light-induced decay. In contrast, the band at 1737 cm Ϫ1 in the first component shows that Ci2 remains partially protonated in the O state. This finding shows that a photocycle branching takes place after M formation, involving a light-induced shortcut from M to D, as observed earlier in ReaChR (11).

Proton transfer reactions in ReaChR
To elucidate the role of the observed proton transfers during the thermal M 3 D and O 3 D transitions, respectively, we investigated the decay process starting from the steady state at 293 K (Fig. 5C). As revealed by SVD and rotation procedure, two components, which were fitted by a global analysis procedure using a biexponential function (31), contributed to the decay process. The first decay component almost mirrors the component following alternating green and UV illumination (Fig. 5B) and thus reflects the direct thermal M 3 D transition involving Ci2 deprotonation. The second component, which represents the O 3 D transition as can be inferred from its slow kinetics ( Ϸ 71 s; see Fig. 3C), comprises a band at 1738 cm Ϫ1 that implies late deprotonation of Ci2. These findings support the above mentioned concept that the photocycle branches after the M-state, involving 1) a direct M 3 D transition where the RSB is presumably reprotonated by Ci2 and 2) a late O 3 D transition. In the latter case, both Asp 196 and Ci2 are excluded as proton donors to the RSB.

Discussion
In this study, the electrical properties and the photocycle dynamics of ReaChR WT were compared with properties of selected mutants by a combined spectroscopic and electrophysiological approach. In the following, the key proton trans-fers are characterized with respect to their functional relevance (Fig. 6).

Glu 130 is a determinant for ion selectivity
Our observation that replacement of Glu 130 by Gln reduces H ϩ conductance in favor of Na ϩ (Fig. 2D) raised questions about the dynamics of this residue during the photocycle, which were then addressed by spectroscopic measurements. We show that Glu 130 undergoes a hydrogen bond change in the D 3 K transition and deprotonates in the K 3 M transition (Figs. 4B and 5A). This finding agrees well with the Glu 90 -helix 2-tilt (EHT) model, that we postulated for the homologue Glu 90 of CrChR2 based on quantum mechanics/molecular mechanics calculations (21). The low-temperature measurements on ReaChR allowed us now for the first time to observe hydrogen bond change and deprotonation as separate events and thus provide experimental evidence for validity of the EHT model to ChRs other than CrChR2.
The EHT model suggests that in CrChR2 one of the two hydrogen bonds between Glu 90 and Asn 258 is disrupted after retinal isomerization. Subsequently, Glu 90 moves outward and deprotonates, eventually leading to the preformation of the conducting pore. Based on the presented experiments and because of the far reaching sequence homologies of the Glu 130

Proton transfer reactions in ReaChR
environment in ReaChR as compared with the environment of Glu 90 in CrChR2, we propose a similar scenario for ReaChR. Accordingly, the observed upshift of the Glu 130 band by 8 cm Ϫ1 in the K spectrum (Fig. 4A) reflects weakening of hydrogen bonding of Glu 130 and is in agreement with a reduction of the number of hydrogen bonds between Glu 130 and Asn 298 from two (weak hydrogen bonds) to one (48). Subsequently, Glu 130 deprotonates in the K 3 M transition, moves outward, and presumably forms a hydrogen bond to Lys 133 (Lys 93 in CrChR2) as it does in CrChR2 (21). Now we focus on how the change of the protonation state of Glu 130 is linked to ion selectivity. We showed that, upon light activation of ReaChR, Glu 130 deprotonates, and therefore a negative charge in the central gate is created that persists in the conducting M 2 -and N-states (11). The E130Q mutation has two consequences. First, in the dark state, no hydrogen bond to Asn 298 is formed, and Gln 130 is already oriented outward as proposed for the chloride-conducting ChRs "ChloC" and derivatives (27,49). This would explain the about 10-fold accelerated M formation in the E130Q mutant (Fig. 3D). Outward orientation of Gln 130 and thus a preactive conformation might be stabilized by a hydrogen bond of its ␥-NH 2 group to Ci1 as estimated from a structural model (50) and molecular dynamics calculations on CrChR2-E90Q (20). The second consequence of the ReaChR-E130Q mutation is neutralization of the negative charge normally present in the conducting state(s). It was shown previously that mutation of the homologous Glu 90 in CrChR2 alters Na ϩ selectivity in a positive or negative sense depending on the charge of the introduced residue (20, 28). Exchange for aspartate hardly affects Na ϩ selectivity as the charge remains unchanged. Neutralization of Glu 90 by glutamine or alanine increases Na ϩ selectivity, whereas lysine or arginine eliminates the Na ϩ conductance in favor of Cl Ϫ conductance (27). Accordingly, increased Na ϩ selectivity in the E130Q mutant of ReaChR as in E90Q appears to be predominantly achieved by altered electrostatics within the ion pore rather than changes of pore geometry near the restriction site.

Asp 196 is a determinant of dark adaptation
The two-photocycle model, comprising two closed (D and DЈ) and two conducting states, was proposed to explain the photocurrent kinetics and amplitudes of the transient and photostationary photocurrents (1,(12)(13)(14)51). This model was supported by spectroscopic data (52) and chromophore isomer analysis and extended with the finding that during one photocycle the retinal remains in 15-anti conformation and during the other the retinal remains in 15-syn conformation (11,15,16). Similarly, in bacteriorhodopsin, two different dark states exist, BR 548 with 13-cis,15-syn-retinal and BR 568 with 13-trans,15-anti-retinal (32,53). Based on the observed differences between IDA and DA app (Fig. 3A), we assume that this applies to ReaChR as well. The photocycle of ReaChR is anticipated to start from IDA with 100% 13-trans,15-anti-retinal (D) by both trans/cis and, with lower efficiency, C13ϭC14 and C15ϭN double isomerization of the retinal (16). Incomplete thermal back-reaction leads to an altered syn/anti ratio in DA app as compared with IDA (see Fig. 3A). This effect is most prominent in D196N (see Fig. 3, A and B). The observation that the D196N mutation abolishes the transient current I t ( Fig. 2A) can thus be explained by the assumption that the 15-anti branch, which is reduced in D196N, causes the peak current, whereas both the 15-anti and 15-syn branches contribute to the stationary current. Accordingly, we provide further evidence that the two open states observed in electrophysiological experiments represent the conducting states of the respective photocycle branches as described earlier (15).
For the question how Asp 196 affects the retinal syn/anti ratio, especially because it is located at more than 9-Å distance from the RSB (Fig. 6), the protonation state of Asp 196 is decisive: Asp 196 deprotonates during N 3 O transition, rendering a negative charge close to the chromophore. The electrostatic interaction of deprotonated Asp 196 with the retinal polyene chain facilitates the back-reaction from 15-syn-to 15-anti-retinal during dark adaptation. Due to the D196N mutation, this negative charge at the chromophore is neutralized, and the syn 3 anti reaction is impaired. Moreover, the DC pair residues or other amino acid pairs (DT in bacteriorhodopsin) at this position in other microbial rhodopsins form a hydrogen bond between helix 3 and helix 4 that is essential for the protein stability and the retinal-binding pocket in particular. Destruction of this bond in many cases destabilizes the retinal-binding pocket and the protein stability in general (54), resulting in a slowdown of both M formation and decay, and might also alter the syn/anti ratio accordingly.

Proton transfer reactions in ReaChR
It remains unclear whether the influence of Asp 196 on the syn/anti equilibrium is applicable to CrChR2 as well: although mutations of Asp 156 abolish the peak current of CrChR2 wild type similarly to D196N in ReaChR (23,27), the retinal fingerprint region in FTIR spectra does not imply a significant alteration of the retinal isomer composition (22). The environment of Asp 196 is different from its homologues in CrChR2 or C1C2, which is reflected by the higher frequency of the CϭO stretch vibration (1753 cm Ϫ1 ) as compared with the respective vibrations at 1738 -1737 cm Ϫ1 (22,55). This could, among other things, be due to the adjacent Cys 199 , which is exchanged for a threonine in these blue-absorbing ChRs.

Proton transfers in the RSBH ؉ counter-ion complex coincide with photocycle branching
The counter-ion complex of the RSBH ϩ is formed by Ci1 (Glu 163 ), Ci2 (Asp 293 ), and presumably water molecules (56 -58). Ci2 receives the RSB proton during M-state formation (Fig.  3G), which is in agreement with the findings for Ci2 in CrChR2 (22) but different to CaChR1 where Ci1 was proposed as proton acceptor (59). Although in both CrChR2 and ReaChR the counter-ion complex interacts with a lysine (Lys 133 in ReaChR) that is exchanged for a phenylalanine in CaChR1, the frequency of the Ci2 stretch vibration (1738 cm Ϫ1 ) is more similar to that of CaChR1 (1753-1740 cm Ϫ1 ) (59) as compared with CrChR2 (1695 cm Ϫ1 ) (22).
Reprotonation of the RSB coincides to M-state decay. As shown by UV/visible and FTIR spectroscopy, the M-state decays thermally by two distinct parallel paths: a major fraction undergoes an M 3 N 3 O transition (main path), eventually leading back to D, and a minor fraction directly reacts back to the dark state in a shortcut. A similar bifurcation of the late photocycle was stated for CrChR2 with the P520 intermediate (homologue to N) as the branching point (39) or for its slowcycling mutants (P390 3 D470) (23). Similarly to the proposed P520 3 D470 shortcut in CrChR2, the actual molecular determinant of this branching process as well as its functional relevance remains elusive in ReaChR. Nevertheless, the branching process is correlated with the protonation state of Ci2: in the side path, Ci2 deprotonates during dark-state recovery and might thus serve as proton donor for reprotonation of the RSB. In the main path, Ci2 deprotonates in the O 3 D transition (Fig. 5C), which excludes Ci2 as proton donor for reprotonation of the RSB. The DC pair residue Asp 196 is excluded as well as proton donor for the RSB (see Figs. 3F and 5B) because RSB reprotonation occurs prior to deprotonation of Asp 196 . This is in contrast to its homologue Asp 156 that was proposed to be the proton donor in CrChR2 (22). As no further carboxylic amino acids could be identified as proton donors to the RSB, the most likely proton source is bulk water, which is abundant during the conducting state.

Conclusion
The proton dynamics observed in the early stage of the ReaChR photocycle, i.e. until formation of the M intermediate, are similar to the blue-absorbing CrChR2, involving (i) hydrogen bond change and subsequent deprotonation of Glu 130 and (ii) proton transfer from the RSBH ϩ to Ci2 during M formation.
Neutralization of Glu 130 enhances Na ϩ conductivity by altered electrostatics of the central gate.
After formation of the M-state, important mechanistic differences between ReaChR and CrChR2 become evident: (i) the photocycle branches with M-state decay, (ii) Asp 196 is excluded as proton donor for RSB reprotonation, and (iii) deprotonation of Asp 196 facilitates thermal syn 3 anti isomerization of the chromophore during dark-state recovery. These differences are explained by distinct structural differences between ReaChR and blue-absorbing ChRs at the active site and Asp 196 environment, respectively. Asp 196 represents an interesting target for elucidating the actual functional relevance of 15-syn-retinal species in ChRs that is poorly understood so far.

Electrical measurements
HEK293 cells were seeded (0.75 ϫ 10 5 cells/ml) in Petri dishes on poly-D-lysine-coated glass coverslips and supplemented with 1 M all-trans-retinal. One day later, cells were transiently transfected with DNA encoding the respective construct using FuGENE HD (Promega, Madison, WI). Recordings took place 48 h after transfection. Signals were amplified and digitized using an AxoPatch 200B and a DigiData 1440 (Molecular Devices, Sunnyvale, CA). Light for activation was provided by a Polychrome V (TILL Photonics, Planegg, Germany) coupled to the optical path of an inverted IX-70 microscope (Olympus, Shinjuku, Japan) and controlled with a programmable shutter (Vincent Associates, Rochester, NY). Standard buffer conditions were as follows: internal 140 mM NaCl, 2 mM MgCl 2 , 2 mM CaCl 2 , 1 mM KCl, 1 mM CsCl, 10 mM HEPES, 10 mM EGTA, pH 7.2, and external 140 mM NaCl, 2 mM MgCl 2 , 2 mM CaCl 2 , 1 mM KCl, 1 mM CsCl, 10 mM HEPES. If not indicated otherwise, data were recorded under that conditions in the whole-cell configuration at a holding potential of Ϫ60 mV. Raw traces were baseline-corrected and filtered with a lowpass Gaussian filter (1000-Hz cutoff). Data are displayed as mean Ϯ S.E.

Proton transfer reactions in ReaChR
were calculated from the respective stationary currents via linear interpolation between the two nearest data points and corrected for the respective liquid junction potential (23°C) except for NMG, pH 9.0, where the values were obtained through linear extrapolation from the currents recorded at Ϫ60 and Ϫ40 mV.

Expression and purification of ReaChR
Expression in HEK293T and purification via immunoaffinity chromatography (1D4) were performed as described before (11).

UV-visible spectroscopy
Absorption spectra were recorded by a Cary 50 Bio spectrophotometer (Varian Inc., Palo Alto, CA) at 22°C. IDA spectra represent spectral properties of protein purified under safe light (Ͼ600 nm), but DA app spectra were measured after preillumination (530 nm, 60 s, 1 ϫ 10 20 photons m Ϫ2 s Ϫ1 ; Luxeon lightemitting diode, Phillips, Amsterdam, Netherlands) and subsequent recovery phase in the dark (10 min). The difference spectrum of ReaChR-C168S was achieved upon prolonged illumination (60 s, 530 nm). Flash photolysis experiments and data processing were performed as described elsewhere (49). In short, a tunable optical parametric oscillator (Rainbow, Mag-icPrism TM , Opotek Inc., Carlsbad, CA) was pumped by the third harmonic (355 nm) of a neodymium-doped yttrium aluminium garnet laser (Rainbow, BrilliantB, Quantel, Les Ulis, France). Excitation wavelength was adjusted manually by a micrometer drive and calibrated with an Andor iStar intensified charge-coupled device camera (Andor Technology Ltd., Belfast, Ireland). Absorption changes were probed by a 150watt xenon short-arc XBO lamp (Osram, München, Germany) and detected by the intensified charge-coupled device camera. Samples with an optical density of A 280 ϭ 1.0 were excited with green flashes (10 ns, 530 nm, 5 mJ/flash). Global analysis is based on a sequential model with four or five spectral components and was performed with Glotaran (62,63).

FTIR measurements
FTIR samples in Dulbecco's PBS, pH 7.4, and 0.03% (w/v) n-dodecyl ␤-D-maltopyranoside were prepared on a BaF 2 window by repeated drying under a nitrogen stream and subsequent rehydration. After preparation, the sample was sealed with a second BaF 2 window. Until use, the samples were stored at Ϫ40°C for cryostatic samples and at 4°C for room temperature samples. For deuteration experiments, pD was adjusted to 7.8. Samples were deuterated by repeated buffer exchange using Centricon centrifugal filter units (GE Healthcare) and subsequent equilibration for 3 days minimum.
Samples were illuminated with light-emitting diodes (maximum emission wavelengths of ϳ530 nm and ϳ390 nm for alternating illumination experiments). For cryostatic measurements, the cryostat DN (Oxford Instruments, Abingdon, UK) was used. Samples were equilibrated at the respective temperature for at least 45 min. After measurement, the sample was heated up again to a minimum 20°C to allow relaxation. FTIR measurements were performed using an ifs66v/s FTIR spectrometer (Bruker Optics, Karlsruhe, Germany) with an LN 2 -cooled mercury cadmium telluride detector (Kolmar Technologies, Newburyport, MA). A 1850 cm Ϫ1 optical cutoff filter was used. Spectra were recorded with a 200-kHz sampling rate and a spectral resolution of 2 cm Ϫ1 . For each data set, Ͼ2500 spectra of the samples were collected and averaged in the dark and after illumination. At cryotemperatures, this procedure was performed for every sample at least twice (n Ն 2), and measurements at 293 K were conducted at least 18 times (n Ն 18) to exclude instabilities of pH and temperature that might affect the reproducibility of the data set.
The difference spectra were corrected for baseline drifts using a spline algorithm and the baseline correction mode implemented in the OPUS 6.5 software package (Bruker Optics). The "steady state" was defined by the steady state of the kinetics of the strongest absorption band in the amide I region (ϳ1660 cm Ϫ1 ). Results obtained from alternating illumination were evaluated by a combination of SVD and a rotation procedure to allow for the estimation of independent spectral components (15). Briefly, the first set of independent spectral components (b-spectra; S i ) and corresponding kinetics components (V i ) were estimated by SVD. These data were then subjected to a rotation procedure based on the autocorrelation function of V i to increase the signal content in a smaller number of vectors as described previously (47). Recovery kinetics were obtained by application of a method combining SVD with a rotation procedure and a global fitting approach using a multiexponential function, the latter supplying the time constants of the decay of the steady state (46).