Unifying photocycle model for light adaptation and temporal evolution of cation conductance in channelrhodopsin-2

Jens Kuhne; Johannes Vierock; Stefan Alexander Tennigkeit; Max-Aylmer Dreier; Jonas Wietek; Dennis Petersen; Konstantin Gavriljuk; Samir F. El-Mashtoly; Peter Hegemann; Klaus Gerwert

doi:10.1073/pnas.1818707116

New Research In

Edited by F. Ulrich Hartl, Max Planck Institute of Biochemistry, Martinsried, Germany, and approved March 27, 2019 (received for review November 5, 2018)

Significance

Understanding the mechanisms of photoactivated biological processes facilitates the development of new molecular tools, engineered for specific optogenetic applications, allowing the control of neuronal activity with light. Here, we use a variety of experimental and theoretical techniques to examine the precise nature of the light-activated ion channel in one of the most important molecular species used in optogenetics, channelrhodopsin-2. Existing models for the photochemical and photophysical pathway after light absorption by the molecule fail to explain many aspects of its observed behavior, including the inactivation of the photocurrent under continuous illumination. We resolve this by proposing a branched photocycle explaining electrical and photochemical channel properties and establishing the structure of intermediates during channel turnover.

Abstract

Although channelrhodopsin (ChR) is a widely applied light-activated ion channel, important properties such as light adaptation, photocurrent inactivation, and alteration of the ion selectivity during continuous illumination are not well understood from a molecular perspective. Herein, we address these open questions using single-turnover electrophysiology, time-resolved step-scan FTIR, and Raman spectroscopy of fully dark-adapted ChR2. This yields a unifying parallel photocycle model integrating now all so far controversial discussed data. In dark-adapted ChR2, the protonated retinal Schiff base chromophore (RSBH⁺) adopts an all-trans,C=N-anti conformation only. Upon light activation, a branching reaction into either a 13-cis,C=N-anti or a 13-cis,C=N-syn retinal conformation occurs. The anti-cycle features sequential H⁺ and Na⁺ conductance in a late M-like state and an N-like open-channel state. In contrast, the 13-cis,C=N-syn isomer represents a second closed-channel state identical to the long-lived P₄₈₀ state, which has been previously assigned to a late intermediate in a single-photocycle model. Light excitation of P₄₈₀ induces a parallel syn-photocycle with an open-channel state of small conductance and high proton selectivity. E90 becomes deprotonated in P₄₈₀ and stays deprotonated in the C=N-syn cycle. Deprotonation of E90 and successive pore hydration are crucial for late proton conductance following light adaptation. Parallel anti- and syn-photocycles now explain inactivation and ion selectivity changes of ChR2 during continuous illumination, fostering the future rational design of optogenetic tools.

In neuroscience, light-activated proteins are utilized to modify the membrane potential and intracellular signal transduction processes of selected cells precisely and noninvasively with light (1, 2). The first and most widely used optogenetic tool is channelrhodopsin-2 (ChR2), a light-gated ion channel from the green alga Chlamydomonas reinhardtii (3, 4).

ChRs are structurally similar to the well-studied prototype of microbial rhodopsins, bacteriorhodopsin (BR) (5, 6). In both proteins, similar arranged clusters of protein-bound water molecules along pathways are crucial for proton conductance (7, 8). However, only a very few tiny alterations are required to switch the proton pump BR into an ion channel. In ChR2, light absorption of the retinal triggers a photocycle involving spectroscopically distinguishable intermediates as outlined in Fig. 1A. After blue-light excitation (λ = 470 nm) of the dark-state D₄₇₀, retinal isomerizes from all-trans to 13-cis, resulting in the red-shifted P₅₀₀ intermediate that corresponds to K in BR. Proton transfer from the protonated retinal Schiff base chromophore (RSBH⁺) to the counter-ion complex leads to P₃₉₀ in ChR2 (M state in BR), possibly also split into an early and late P₃₉₀ state (such as M₁ and M₂ in BR). P₃₉₀ is succeeded by P₅₂₀ (N in BR) after reprotonation of the RSB. Considering the time constants of channel opening observed in electrophysiological experiments, it has been suggested that both states, the late P₃₉₀ (M₂) and P₅₂₀ (N), contribute to ion conductance of the open channel (9 ⇓–11). Finally, a long-lived nonconducting state P₄₈₀ appears after channel closing. P₄₈₀ is considered to be the last photocycle intermediate, and D₄₇₀ is recovered from this species with a time constant of ∼40 s. The unbranched photocycle is reasonably well suited to describe a single-turnover transition starting from the dark-adapted protein, but fails to explain photocurrent changes during extended light application. During continuous illumination, photocurrents of ChR2 inactivate within milliseconds from a transient peak to a stationary level, and the initial peak current is recovered only after many seconds in darkness (3). Furthermore, the photocurrent decays biexponentially with two distinct time constants that differ by an order of magnitude and an amplitude ratio that depend on the preillumination time, excitation wavelength, and membrane voltage. Thus, the above-mentioned single-cycle model was extended to a parallel two-cycle model comprising two closed (C₁ and C₂) and two open (O₁ and O₂) states that are populated differently in the dark and during repetitive or continuous illumination (12, 13) (Fig. 1B). Photocurrent inactivation and differences in conductance were explained by a higher quantum efficiency for the transition from C₁ to O₁ compared with that from C₂ to O₂, consistent with recent theoretical calculations (14).

Fig. 1.

Proposed photocycle models of ChR2. (A) Single nonbranched photocycle model starting from the IDA has been most widely used to explain time-resolved UV/VIS and FTIR experiments (9, 10, 15 ⇓⇓–18). (B) To explain both moderate photocurrent inactivation and slow dark-state recovery, however, a branched model with two open states is required (12, 13).

Time-resolved FTIR spectroscopy was originally established as a powerful approach for the determination of the molecular reaction mechanism of BR (19). Accordingly, the dark-adapted ChR2 photocycle was recorded between 50 ns and 140 s after exposure to a light pulse by step-scan and rapid-scan FTIR. These measurements revealed an ultrafast all-trans to 13-cis isomerization and subsequent deprotonation of the RSBH⁺ in parallel with protonation of the counter-ion residues E123 and D253 (18). Deprotonation of D156 coincides with P₃₉₀ depletion, which was previously considered as indicative of RSB reprotonation (17, 18). FTIR studies paired with HPLC analysis of the slow-cycling step-function variant C128T provided spectroscopic evidence for two distinct closed states with different retinal isomers (20). NMR-spectroscopic data of the ChR2 (WT) and WT-like variant H134R showed that although different closed states exist, the fully dark-adapted state [called the initial dark-adapted state (IDA)] of ChR2 is composed of 100% all-trans,C=N-anti retinal (21, 22). Raman experiments on ChR2-H134R revealed that illumination of the IDA at 80 K produced an apparent dark state (DA_app) containing a second retinal isomer (22). Following double isomerization around the C₁₃ = C₁₄ and the C=N double bonds, 13-cis,C=N-syn retinal is formed, and this was proposed as the transformation step for forming the second “metastable” dark state (22). Both retinal isomers in the DA_app were proposed to initiate distinct photocycles, with both involving homologous P₅₀₀-, P₃₉₀-, P₅₂₀-, and P₄₈₀-like intermediates.

The central gate residue E90 is one of the key determinants of proton selectivity in ChR2 (16, 18, 23) and related cation-conducting ChRs (24). During the photocycle, E90, which is located in the central gate in the middle of the putative pore, is deprotonated and remains deprotonated until P₄₈₀ decays (16 ⇓–18). From experiments with high laser pulse repetition frequencies preventing complete dark adaptation, a late deprotonation of E90 exclusively in P₄₈₀ was proposed for ChR2 (17). In contrast, E90 deprotonation within submicroseconds after light excitation was observed in single-turnover experiments on fully dark-adapted ChR2 (18). Thus, there seemed to be a controversy between fully dark-adapted and non–dark-adapted FTIR experiments on the timing of E90 deprotonation in a single photocycle model.

Here, we present a unifying functional study of dark- and light-adapted ChR2 by integrating single-turnover electrical recordings and FTIR measurements on ChR2, Raman spectroscopy with ¹³C-labeled retinal, and molecular dynamics (MD) simulations. The controversies observed between single-turnover experiments and recordings under continuous illumination are resolved by developing an extended model, including two parallel photocycles with C=N-anti and C=N-syn retinal conformations. The light-adapted 13-cis,C=N-syn state is the P₄₈₀ intermediate, which was formerly assigned to the last intermediate of the anti-cycle in a linear photocycle model. Within the anti-cycle, ion conductance evolves in two subsequent steps, resulting in two different conducting states of distinct ion selectivity (O_1-early and O_1-late). Interestingly, E90 stays protonated in the anti-cycle. In contrast, the syn-cycle initiated by photoexcitation of P₄₈₀, which represents the second C₂ in Fig. 1B_, comprises a third slowly decaying O₂ of high proton selectivity but low overall ion conductance. Conductance of O₂ depends on deprotonation of E90 and is completely abolished in the ChR2 E90Q mutant. Our results resolve the former discrepancies. In the anti-photocycle, E90 stays protonated and channel opening of O_1-early and O_1-late is observed, whereas in the syn-cycle, including P₄₈₀, E90 is deprotonated and favors proton conductance of O₂.

Results

Single-Turnover Patch-Clamp Recordings Identify Three Conducting ChR2 States.

To examine functional changes during light adaptation of ChR2, we recorded single-turnover photocurrents in HEK293 cells following 7-ns laser excitation before and after light adaptation. We addressed changes in ion selectivity by reducing either the extracellular sodium (110 mM → 1 mM) or extracellular proton concentration (pH_e 7.2 → pH_e 9.0) (Fig. 2 A and B).

Fig. 2.

Voltage-clamp recordings in HEK293 cells of photocurrents from dark- and light-adapted ChR2 WT. (A) Experimental scheme of the whole-cell patch-clamp experiment in different extracellular buffers and under different illumination conditions. (B, Left) Representative photocurrents of ChR2 with intracellular 110 mM NaCl and pH_i 7.2 and extracellular 110 mM Na⁺ and pH_e 7.2 (Top), 1 mM Na⁺ and pH_e 7.2 (Middle), and 110 mM Na⁺ and pH_e 9.0 (Bottom) at different holding potentials as indicated. Photocurrents were excited before and after light adaptation with a 470-nm, 7-ns laser pulse. For light adaptation, cells were continuously illuminated for 500 ms with 470-nm light. (B, Right) Normalized, log-binned, and averaged photocurrents of the dark-adapted (DA) or light-adapted (LA) protein (mean ± SEM, n = 5–8). (C) Time evolution of estimated proton and sodium fluxes in the DA protein at −60 mV either directly measured in extracellular 1 mM Na⁺ and pH_e 7.2 (H⁺ current) or calculated by subtraction of proton fluxes from combined inward flux of sodium and protons measured in symmetrical conditions (Na⁺ current) (I [110 mM Na⁺ (pH 7.2)] − I [1 mM Na⁺ (pH 7.2)]; mean ± SEM; n = 7). (D) Relative photocurrent changes upon light adaptation at different extracellular voltages and pH_e [I (LA) − I (DA)]/I (DA); mean ± SEM; n = 5–8]. (E) Current-voltage dependency of normalized photocurrents at 0.1 ms (Left), 5 ms (Center), and 100 ms (Right) after excitation in different extracellular buffer compositions before (DA) and after (LA) light adaptation (mean ± SD; n = 5–8).

Under symmetrical sodium and proton concentrations, the dark-adapted ChR2 pore opens biexponentially with two almost voltage-independent time constants (150 μs and 2.5 ms). The photocurrents decline, with a dominant voltage-dependent time constant of 10–22 ms and a second, minor, slow time constant of 70–220 ms (Fig. 2 A and B, Top), in general agreement with previous reports (11). Decreasing extracellular Na⁺ not only reduces inward-directed photocurrent amplitudes but also affects the temporal evolution of inward currents (Fig. 2B, Middle). Whereas inward currents in low extracellular Na⁺ are predominantly carried by protons (H⁺ flux), inward currents under symmetrical conditions are mediated by both H⁺ and Na⁺ ions. Subtraction of photocurrents at high and low Na⁺ at pH 7.2 allows an approximation of the pure inward Na⁺ flux (Fig. 2C). Strikingly, the proton flux peaks as early as 300 μs after excitation, significantly earlier than Na⁺ flux (2.5 ms). This observation is indicative of two open states with distinct ion selectivity following single excitation of dark-adapted ChR2.

During continuous illumination, photocurrents peak within milliseconds (dependent on the light intensity) and subsequently decline to a stationary level. Inactivation is more pronounced at positive voltages, contributing to the increased inward rectification of stationary photocurrents compared with the initial peak current (3). After light adaptation, laser pulse-induced photocurrents are significantly reduced in amplitude. The photocurrent still rises and decays biexponentially, however, reaching a maximal amplitude at the same postflash time point as photocurrents in the dark-adapted protein (Fig. 2B). The relative photocurrent change at different time points after excitation shows a homogeneous photocurrent reduction of 60–80% between 0.2 ms and 10 ms (Fig. 2D). However, notably, relative photocurrent changes differ at early time points (<200 μs) and during the slow photocurrent decline (after 50–100 ms), indicating different open-state conformations in the early and late stages of the photocycle of light- and dark-adapted ChR2. In particular, at 100 ms after the laser excitation, the photocurrents even increase in amplitude for light-adapted ChR2 (especially at pH 9.0), indicative of an additional slowly decaying open state. In summary, there are at least three different conductive states: O_1-early and O_1-late in the dark-adapted photocycle and O₂ after light adaptation.

The proton versus sodium selectivity of all three open states is analyzed at three different time points after the actinic laser pulses (0.1 ms, 5 ms, and 100 ms; Fig. 2E). Although the photocurrents at 0.1 ms and 100 ms are barely distinguishable in high and low extracellular Na⁺ in either the dark- or light-adapted protein, reduction of the extracellular proton concentration causes a strong shift in reversal potential and an increase in outward-directed photocurrent amplitudes that is even more pronounced in the light-adapted channel than in the dark-adapted channel. In contrast, 5 ms after excitation of photocurrents, both ionic changes, a reduction in extracellular sodium or proton concentration, shift the reversal potential and decrease the inward photocurrent amplitude. We conclude that after channel opening, the short-lived highly proton-selective O_1-early is followed by the more Na⁺-selective but still highly proton-permeable O_1-late. After multiphoton excitation and light adaptation, the contribution of O_1-early and O_1-late decreases in favor of the third highly proton-selective O₂, which, although small in amplitude, significantly contributes to stationary photocurrents at alkaline pH due to its long lifetime.

Single-Turnover Time-Resolved FTIR Measurements Reveal a Splitting of the Photocycle After Light Activation of Fully Dark-Adapted ChR2.

The single-turnover electrophysiology data recall that dark adaptation and light adaptation need to be compared thoroughly for the correct interpretation of time-resolved measurements. However, most time-resolved spectroscopy studies are performed with barely dark-adapted samples at rather high repetition rates to avoid long measurement times. To elucidate the underlying molecular mechanism of the observed channel-gating transitions and different ion conductance, we performed single-turnover time-resolved FTIR measurements of the fully dark-adapted ChR2 WT-like H134R variant with a time resolution of 50 ns over nine orders of magnitude (Fig. 3A), similar to our data from 2015 (18). The ChR2 WT-like H134R mutant shows higher protein expression in Pichia pastoris compared with the WT protein and has been used for the examination of light adaptation before (22). Electrical properties and photocycle kinetics are comparable, although slightly slower than those of the WT protein (25), and the same IR bands are observed in WT and in H134R. However, some crucial IR marker bands are more pronounced in H134R, which simplifies the presentation of the dataset. Dark adaptation of D₄₇₀ was achieved by long dark periods of 140 s between pulsed excitation (temperature = 15 °C), which increased the advanced step-scan measurement time to about 4 wk (18), whereas light-adapted samples take a few hours only (17). The appearance of the marker band at 1188 cm⁻¹ (not time-resolved) indicates the all-trans to 13-cis,C=N-anti isomerization because it represents the C₁₄-C₁₅ stretching vibration of 13-cis retinal as originally assigned in BR by site-specific isotopic labeling (26). The decay of the 1,188 cm⁻¹ marker band within a microsecond (green time trace in Fig. 3A) indicates the formation of the M-like P₃₉₀ intermediate with a deprotonated RSB (18). As in other microbial rhodopsins, the subsequent rise and decay of the N-like P₅₂₀ intermediate with a reprotonated Schiff base can be monitored by its reappearance and the decay reflects formation of the all-trans isomer on the time scale of a few milliseconds (19). Comparing the time course of this marker band (1,188 cm⁻¹) with the single-turnover electrical measurements, we can now assign the described conducting states O_1-early and O_1-late to the late part of P₃₉₀ (M₂) and P₅₂₀ (N), respectively, which is in line with earlier reports on the WT protein (11, 18). Due to these similarities and the abundance of spectroscopic data on BR, we decided to name the ChR2 intermediates as follows: P₅₀₀^K, P_390a^M1, P_390b^M2, and P₅₂₀^N. Global fitting of the whole dataset (solid lines, Fig. 3A) describes the data adequately. The apparent rate constants of H134R are similar to those of earlier reports for the dark-adapted ChR2 WT (18).

Fig. 3.

FTIR measurements on H134R and WT. (A) Kinetic transients of the marker bands in WT-like H134R variant recorded by step-scan FTIR. The P₄₈₀ C=N-syn (red) and E90 (black) marker bands, 1,154 cm⁻¹ and 1,718 cm⁻¹, respectively, are observed not time-resolved at the very beginning of the reaction. Their decay occurs with the decay of P₄₈₀ (t_1/2 = 40 s). In parallel, the D₄₇₀ → P₅₀₀^K → P₃₉₀^M → P₅₂₀^N reaction is monitored by the marker band for protonated 13-cis,C=N-anti retinal at 1,188 cm⁻¹ (green). The continuous lines are the result of a global fit analysis using five rate constants that sufficiently describe the dataset. (B) Comparison of unlabeled (black), ¹³C₁₄-¹³C₁₅ labeled (red), and unlabeled but deuterated (green) WT samples in the P₄₈₀-D₄₇₀ difference spectrum. The marker band at 1,188 cm⁻¹ from A is not seen at this late photocycle intermediate. The red arrows indicate the isotope-induced downshifts of the C₁₄-C₁₅ stretching vibration at 1,186 cm⁻¹ in D₄₇₀ and 1,154 cm⁻¹ in P₄₈₀. The green arrows denote the large upshift of the C₁₄-C₁₅ stretching vibration at 1,154 cm⁻¹ induced by deuteration to 1,180 cm⁻¹, indicating the syn-conformation. Also, the C₁₀-C₁₁ stretching vibration at 1,176 cm⁻¹ is upshifted. The large upshift of the P₄₈₀ bands indicates a C=N-syn conformation of the retinal in P₄₈₀. In contrast the C₁₄-C₁₅ stretching vibration at 1,186 cm⁻¹ in D₄₇₀ is only slightly upshifted in D₂O to 1,191 cm⁻¹, indicating a trans-conformation. More details are provided in SI Appendix, Supplementary Notes 1 and 3–5, Figs. S1–S8, and Table S1.

Light-Induced Splitting in 13-cis,C=N-anti and 13-cis,C=N-syn RSBH⁺ Conformations.

Interestingly, an additional retinal band at an unusual low wavenumber, 1,154 cm⁻¹, appears parallel to the 1,188-cm⁻¹ 13-cis,C=N-anti marker band (not time-resolved). Because the low-wavenumber band is more pronounced in H134R than in WT (SI Appendix, Fig. S1), we discuss the data here for the reader on the mutant, although the results are also valid for the WT. The 1,154-cm⁻¹ band persists from nanoseconds to seconds after a single pulse of excitation light (Fig. 3A). The band is assigned here to the C₁₄-C₁₅ stretching vibration of retinal because of the characteristic 14-cm⁻¹ downshift upon retinal ¹³C₁₄-¹³C₁₅ carbon-specific labeling (Fig. 3B, upper part). The band assignment is confirmed by additional Raman experiments (SI Appendix, Fig. S6). The 22-cm⁻¹ upshift of the C₁₄-C₁₅ band in D₂O indicates a 13-cis,C=N-syn conformation (Fig. 3B, lower part). In 13-cis,C=N-syn retinal, the C₁₄-C₁₅ stretching vibration is strongly coupled to the N-H bending vibration, which is decoupled in D₂O (N-D) and results in a deuteration-induced large upshift in the syn-conformation, but not in the anti-conformation (27, 28). Therefore, the band at 1,154 cm⁻¹ represents a 13-cis,C=N-syn marker band. The band assignments are confirmed by extended Raman experiments shown in SI Appendix in more detail for P₄₈₀ (SI Appendix, Supplementary Notes 1 and 3–5, Figs. S1–S6, and Table S1).

The negative difference bands in Fig. 3B reflect vibrations of dark-adapted ChR2 WT (D₄₇₀). The negative band at 1,186 cm⁻¹ is also assigned to the C₁₄-C₁₅ stretching vibration of D₄₇₀ because of the characteristic isotope downshift. From the additional analysis of the D₄₇₀ Raman spectrum (SI Appendix, Supplementary Notes 3 and 4), we conclude that the retinal of dark-adapted ChR2 is in a 100% all-trans,C=N-anti conformation, in agreement with NMR data (21, 22). A detailed band assignment of the P₄₈₀ and D₄₇₀ vibrational spectra and retinal conformations is provided in SI Appendix, Supplementary Notes 1 and 3–5, Figs. S1–S7, and Table S1.

The parallel but temporally unresolved appearance of the bands at 1,188 cm⁻¹ and 1,154 cm⁻¹ in single-turnover experiments in Fig. 3A indicates that light absorption induces parallel isomerization of all-trans,C=N-anti retinal in D₄₇₀ into either a 13-cis,C=N-anti or a 13-cis,C=N-syn conformation. The splitting ratio into parallel syn- and anti-pathways can be estimated as 1:1 under our measurement conditions (SI Appendix, Fig. S3).

Considering that the 13-cis,C=N-syn isomerization occurs in parallel to the 13-cis,C=N-anti isomerization, we conclude that the 13-cis,C=N-syn retinal conformation observed in P₄₈₀ is therefore not the last intermediate of the 13-cis,C=N-anti photocycle, as proposed in the single-cycle model. P₄₈₀ reflects a long-lived 13-cis,C=N-syn state, which appears in parallel to 13-cis,C=N-anti state instantaneously.

The conclusion that the P₄₈₀ is not the last intermediate of the 13-cis, C=N-anti single photocycle but appears in parallel to the light-adapted state is furthermore strongly supported by detailed Raman experiments, as described in SI Appendix. The Raman results are in agreement with former Raman studies on light adaption (22). Upon complete light adaptation of ChR2 due to long illumination periods, a D_app state evolves. It is composed of a 40:60 mixture of the all-trans,C=N-anti species in D₄₇₀ and the 13-cis,C=N-syn species in P₄₈₀ (SI Appendix, Fig. S3). The bands observed in the Raman spectra of D₄₇₀ and P₄₈₀ correlate with retinal bands seen in the IR difference spectra in Fig. 3B and are in agreement with the published Raman spectra of the all-trans,C=N-anti and 13-cis,C=N-syn bands of the D_app state (22) (SI Appendix, Supplementary Note 3). The Raman data confirm the ultrafast C=N-syn formation in P₄₈₀ as seen in Fig. 3A at 1,154 cm⁻¹.

E90 Deprotonates upon 13-cis,C=N-syn Formation.

The E90-deprotonation marker band (1,718 cm⁻¹) (18) and the C=N-syn marker band (1,154 cm⁻¹) (Fig. 3A) appear instantaneously, and are not time-resolved. Both marker bands persist alongside the dark-adapted anti-cycle intermediates (P₅₀₀^K, P₃₉₀^M, and P₅₂₀^N). Both decay with a slow t_1/2 of ∼40 s. We therefore conclude that E90 remains deprotonated during the entire C=N-syn pathway. However, E90 does not deprotonate in the anti-cycle. This resolves the former discrepancies between Lórenz-Fonfría et al. (17) and Kuhne et al. (18). In agreement with the former findings, E90 deprotonates in P₄₈₀, but this intermediate and E90 deprotonation appear not late in the last intermediate, as proposed in a linear photocycle (17), but much faster, with the appearance of P₄₈₀ during light adaptation in a parallel photocycle (18). In addition, E90 deprotonation appears to be closely connected to helix hydration, as indicated by the helix hydration marker bands (1,662 cm⁻¹ in D₄₇₀ and 1,650 cm⁻¹ in P₄₈₀) that were assigned in an earlier study, but only for P₄₈₀ (11). Interestingly, the same P₄₈₀ hydration marker bands that are present in the WT were no longer observed in the mutants E90Q, E123T, and K93S that prevent E90 deprotonation (SI Appendix, Supplementary Note 2 and Fig. S1). Therefore, E90 deprotonation seems to be induced by the all-trans,C=N-anti → 13-cis,C=N-syn isomerization and modulates the water influx in P₄₈₀.

Excitation of P₄₈₀ Induces a Parallel Photocycle.

After light adaptation, both D₄₇₀ and P₄₈₀ serve as parent states for parallel photocycles. At flash frequencies of 0.2 Hz that do not allow sufficient dark-state recovery, a biexponential decay with a fast process (t_1/2 = 30 ms) and a slow process (t_1/2 = 250 ms) was observed. The corresponding amplitude spectra are shown in Fig. 4C (also SI Appendix, Supplementary Note 6 and Fig. S8). The 30-ms amplitude spectrum exhibits positive ultraviolet/visible (UV/VIS) bands at 380 nm and 520 nm (Fig. 4B), indicative of a mixture of P₃₉₀^M and P₅₂₀^N intermediates in the C=N-anti cycle. In contrast, if the sample is sufficiently dark-adapted (0.005-Hz flash repetition rate), the 250-ms component is no longer observed. We therefore conclude that the slow time constant reflects the conducting state of the syn-cycle (Fig. 4A). Because the t_1/2 = 30 ms process reflects an apparent, but not an intrinsic, rate constant, the decay of both intermediates, which follows different intrinsic rate constants, is described by the integrated apparent rate.

Fig. 4.

Kinetic behavior of ChR2-WT at different laser pulse repetition rates. (A) Time-evolution of the amide-I band at 1,544 cm⁻¹ at low (0.005 Hz, black) and high (0.2 Hz, red) pulse repetition frequency. Upon higher pulse repetition frequency, the decay switches from mono- to biexponential. (B) Comparison of O₁ (green) and O₂ (black) decay-associated UV/VIS amplitude spectra. The two positive bands at 380 nm and 520 nm in O₁ (green) indicate a mixture of P₃₉₀^M2 (O_1-early) and P₅₂₀^N (O_1-late). No evidence for RSBH⁺ deprotonation is visible in the slow component (O₂). (C) Same processes as monitored by FTIR.

The amplitude spectrum of the 250-ms apparent rate constant indicates decaying intermediates of the syn-cycle. As there is only a positive band at 520 nm, it was designated the P*₅₂₀^N intermediate to distinguish it from the P₅₂₀^N of the anti-cycle. The P*₅₂₀^N kinetics shown in Fig. 4 fit well to the slow photocurrent component O₂ shown in Fig. 2, and are assigned to it in the following. In contrast, no P₃₉₀^M-like intermediate is observed in the syn-cycle (Fig. 4B).

FTIR Amplitude Spectra Indicate Structural Differences of O₁ and O₂.

The O₁ and O₂ FTIR amplitude spectra of the 30-ms and 250-ms apparent rates are shown in Fig. 4C. Both decay-associated amplitude spectra exhibit negative D₄₇₀ marker bands (Fig. 4C and SI Appendix, Figs. S8 and S10), indicating a direct transition from the anti- and syn-photocycles into the all-trans,C=N-anti configuration of D₄₇₀.

The t_1/2 = 30-ms FTIR decay-associated amplitude spectrum of the C=N-anti cycle exhibits carbonyl bands at 1,760 cm⁻¹ (+)/1,736 cm⁻¹ (−) and 1,728 cm⁻¹ and 1,695 cm⁻¹, which were assigned to protonation of the counter-ion D253 (18, 29) (1,728 cm⁻¹) and deprotonation of D156 (1,736 cm⁻¹). Furthermore, the helix hydration marker bands at 1,662 cm⁻¹ (−)/1,650 cm⁻¹ are present, which are now assigned to both O_1-early and O_1-late. In the t_1/2 = 250-ms decay-associated amplitude spectrum, all carbonyl bands are strongly reduced, including the bands of the Schiff base proton acceptor D253 (at 1,728 cm⁻¹), as well as D156 (1,736 cm⁻¹), which has been proposed to be the RSB reprotonation donor (17). Because the Schiff base deprotonation is not observed, the corresponding counter-ion D253 protonation is not seen either. Also, reprotonation of D156 (1,736-cm⁻¹ band) is no longer observed. The negative P₄₈₀ band (1,154 cm⁻¹) in the t_1/2 = 250-ms decay-associated amplitude spectrum indicates an additional O₂ → P₄₈₀ backreaction within the syn-photocycle (SI Appendix, Supplementary Note 6 and Figs. S8 and S10).

Isomerization of Retinal Leads to a Rearrangement of the Central Gate.

To visualize the structural changes within the protein, we performed MD simulations based on the recently published crystal structure of ChR2 WT [Protein Data Bank (PDB) ID code 6EID] (30). The structure of ChR2 is highly similar to the structure of the ChR1-like chimera C1C2; however, in contrast to the latter, in the ChR2 structural model, E90 is already “downward”-oriented in the dark-adapted state and at the same position as observed for the C1C2 chimera structural model after isomerization (18, 31). Within our simulations of the ChR2 WT structure presented here, the retinal isomerization was changed from the dark-adapted all-trans,C=N-anti conformation (Fig. 5C, Left) to either a 13-cis,C=N-anti single isomerization (SI Appendix, Supplementary Note 7 and Fig. S13 A and B) or a 13-cis,C=N-syn double isomerization (Fig. 5C, Right and SI Appendix, Fig. S13C). The observed changes in hydrogen bond interaction and water distribution are shown in Fig. 5 B and C and SI Appendix, Figs. S12–S14. It is noteworthy that the single isomerization induces an upward orientation of the RSB proton, whereas the position of the RSB proton is only slightly changed in the double isomerization (16, 22) (Fig. 5A). Starting from the WT structure, E90 keeps its initial downward orientation in the dark-adapted state (Fig. 5 B and C). Very recently, a more advanced method to perform such isomerization simulations was introduced by Ardevol and Hummer (31). They simulated a homology model of ChR2 based on the C1C2 chimera crystal structure (PDB ID code 3UG9) (32) and obtained a downward flip of the initially upward-orientated E90. We have already observed the same downward movement in our model based on the same crystal structure using a classical approach (18). This proves that even our classical approach correctly predicts alterations of the hydrogen bond pattern of E90 due to retinal isomerization. It seems that E90 is trapped in a local minimum in both models but finds the correct position for ChR2 [as observed in the PDB ID code 6eid crystal structure (30)] after disturbance by isomerization.

Fig. 5.

Retinal conformations and formation of P₄₈₀. (A) Modeled representation of the calculated retinal configurations. The Schiff base orientations in the D₄₇₀ structure all-trans,C=N-anti (gray) and in the modeled 13-cis,C=N-anti (green) and 13-cis,C=N-syn (blue) retinal structures are shown. (B) Overview of E90 hydrogen bond pattern for five independent simulations, with two monomers forming one dimer based on the ChR2 WT crystal structure [PDB ID code 6EID (30)]. Bars indicate the frequency of the respective hydrogen bond (percentage) during the 100-ns simulation. (C) Representative structure of the simulations is depicted. (Left) D₄₇₀ dark state. (Right) Structure after all-trans,C=N-anti → 13-cis,C=N-syn double isomerization and E90 deprotonation (P₄₈₀). After deprotonation of E90, the central gate opens and water invades.

Following 13-cis,C=N-syn double isomerization, helices 2 and 7 stay connected via E90 and D253 as long as E90 remains protonated (SI Appendix, Fig. S12). Deprotonation of E90 leads to an alternative contact between E90 and K93 (Fig. 5 B and C and SI Appendix, Fig. S12) that opens the central gate and results in an influx of water molecules into the pore (Fig. 5C). This water influx is in agreement with the channel opening due to E90 deprotonation proposed formerly in the E90-Helix2-tilt (EHT) model in the 13-cis,C=N-anti conformation (18). We now attribute E90 deprotonation and pore hydration to the light-adapted closed-state P₄₈₀. In this light-adapted state, the inner gate still remains closed and ion permeation is hindered, in agreement with the electrophysiology results (SI Appendix, Fig. S14). As 13-cis,C=N-syn isomerization accumulates during light adaptation, we could attribute E90 deprotonation and pore hydration to the light-adapted closed-state P₄₈₀.

Deprotonation of E90 Is Essential for Proton Conductance of O₂ Following Light Adaptation.

As we have shown that the deprotonation of E90 is responsible for pore hydration in the light-adapted dark-state P₄₈₀, an important role of E90 deprotonation for channel conductance in the syn-cycle appeared likely from our model. Consequently, we mutated E90 to glutamine and analyzed photocurrent changes before and after light adaptation (Fig. 6A and SI Appendix, Fig. S11). In general agreement with previous results of steady-state measurements (16, 33), the E90Q mutation reduces proton conductance of O_1-late of the anti-cycle. Accordingly, upon reduction of extracellular sodium, photocurrent amplitudes are more decreased compared with the WT (Fig. 6B), and the reversal potential shifts are larger 2 ms after excitation (Fig. 6C and SI Appendix, Fig. S11C). In addition to the effect on the anti-cycle, the E90Q mutation completely abolishes the late photocurrent increase upon light adaptation, which was observed in the WT channel (SI Appendix, Fig. S11D). Instead, slow photocurrents are reduced in the E90Q mutant following continuous illumination (Fig. 6 D and E). This indicates that E90 facilitates proton conductance of O₂ by deprotonation, rendering it completely impermeable in the E90Q mutant. The results on the E90Q mutation validate our photocycle model with a parallel syn-cycle that involves E90 deprotonation and populates during light adaption.

Fig. 6.

Proton and sodium conductance of the dark- and light-adapted ChR2 mutant E90Q. (A) Representative photocurrents of ChR2-E90Q with intracellular 110 mM NaCl and pH_i 7.2 and extracellular 110 mM Na⁺ and pH_e 7.2 (Top), 1 mM Na⁺ and pH_e 7.2 (Middle), 110 mM Na⁺ and pH_e 9.0 (Bottom) at different holding voltages as indicated. Photocurrents were excited before and after light adaptation by 7-ns laser pulses of 470-nm wavelength light. For light adaptation, cells were illuminated for 500 ms with continuous 470-nm light. (B) Time evolution of estimated proton and sodium fluxes in the dark-adapted protein at −60 mV either directly measured in extracellular 1 mM Na⁺ and pH_e 7.2 (“H⁺ current”) or calculated by subtraction of proton fluxes from combined inward flux of sodium and protons measured in symmetrical conditions (“Na⁺ current”) (I [110 mM Na⁺ (pH 7.2)] − I [1 mM Na⁺ (pH 7.2)]; mean ± SE; WT: n = 7, E90Q: n = 6). (C) Reversal potential shift (ΔE_rev) 2 ms after laser light excitation of the dark-adapted protein upon reduction of extracellular sodium (110 mM NaCl → 1 mM NaCl) or proton (pH_e 7.2 → pH_e 9.0) concentration (mean ± SD; E90Q: n = 5–6, WT: n = 6–7; corrected for liquid junction potentials). (D) Equally scaled representative photocurrents of ChR2 WT and E90Q at +30 mV and extracellular 110 mM Na⁺ and pH_e 9.0. (E) Normalized, log-binned, and averaged photocurrents of the dark-adapted (DA) or light-adapted (LA) WT and E90Q at +30 mV and extracellular 110 mM Na⁺ and pH_e 9.0 (mean ± SEM; WT: n = 6, E90Q: n = 5).

Discussion

Microbial rhodopsins are excellent optogenetic tools (1 ⇓⇓–4). Understanding of their detailed mechanisms is catalyzed by earlier extended studies on BR providing detailed insight on how tiny light-induced protein alterations induce a proton transfer by an interplay of catalytic key residues and clusters of protein-bound water molecules along the proton transfer pathway (5 ⇓–7). This detailed understanding paved the way for studies of several other microbial rhodopsins, like halorhodopsins, sensory rhodopsins, and especially ChRs. In the present work, we combined single laser pulses and continuous or repetitive illumination in an advanced biophysical approach to analyze the fully dark- and light-adapted ChR2 in single-turnover electrophysiological recordings, time-resolved FTIR, and resonance Raman spectroscopic measurements, complemented by MD simulations. We verified early branching into two parallel photocycles with distinct retinal isomerization and alternative configurations of the central gate, and elaborated a unifying photocycle model shown in Fig. 7 that addresses light adaptation and temporal changes in cation conductance on a functional and molecular level.

Fig. 7.

Unifying photocycle model. D₄₇₀ and P₄₈₀ represent the closed states C₁ and C₂ of Fig. 1B. D₄₇₀ is the state with all-trans,C=N-anti retinal, which is populated to almost 100% in fully dark-adapted ChR2. Upon light activation, two pathways are observed. In path 1, all-trans,C=N-anti → 13-cis,C=N-anti isomerization initiates the anti-photocycle, with the K-like P₅₀₀^K that converts into M-like P₃₉₀^M and N-like P₅₂₀^N assigned to the open states (O_1-early) and (O_1-late). In path 2, all-trans,C=N-anti → 13-cis,C=N-syn isomerization leads to P₄₈₀ formation with deprotonated E90. The nonconducting state P₄₈₀ slowly relaxes back to D₄₇₀. At high flash frequencies or under continuous illumination, P₄₈₀ (C₂) accumulates. The light-adapted DA_app represents a mixture of D₄₇₀ and P₄₈₀ (C₁ + C₂). The photoproduct of the photoreactive P₄₈₀ (C₂) is the N-like P*₅₂₀^N, which is regarded as the open state O₂. Due to its relatively long decay time P*₅₂₀^N accumulates under continuous illumination with bright light as commonly used in electrophysiology. During such conditions, both P₄₈₀ and P*₅₂₀^N contribute to the inactivation of ChR2 as they accumulate at the expense of the highly conductive O₁ and its parent state D₄₇₀. E90 remains protonated during the anti-cycle and deprotonated during the syn-cycle.

Following longer dark periods, the IDA comprises only D₄₇₀ containing 100% all-trans,C=N-anti retinal, which is in agreement with previous reports (21, 22). The inner gate and the central gate are closed, and interhelical hydrogen bonding of D253 with the RSBH⁺ and the protonated E90 (31) prevents the invasion of water molecules from the extracellular bulk phase. After illumination of D₄₇₀ (C₁), early branching of the photocycle due to an alternative retinal single or double isomerization occurs.

In the classical reaction path starting from D₄₇₀ (designated the anti-cycle) all-trans,C=N-anti retinal isomerizes to 13-cis,C=N-anti, leading to P₅₀₀^K, deprotonation of the RSBH⁺ in P₃₉₀^M and reprotonation of the RSB in P₅₂₀^N, and direct monoexponential recovery of D₄₇₀. Channel opening occurs at the UV/VIS silent transition from P_390a^M1 to P_390b^M2 in two subsequent steps (11) that we can now attribute to different ion selectivities pinpointing different pore conformations. Whereas during P_390b^M2, the short-lived O_1-early conducts almost exclusively protons, photocurrents of O_1-late, which evolve upon reprotonation of the RSB and formation of P₅₂₀^N, are also carried by cations. Curiously, we observe a small positive charge displacement during P₅₂₀^N at 0 mV and under symmetrical ionic conditions (Fig. 2) that could result from an outward-directed proton displacement following reprotonation of the RSB. This might reflect the earlier proposed residual proton transfer in ChR2 (34) that was later associated with protonation changes of D156 (17). We note that the observed small charge transfer is barely visible after light adaptation or any applied membrane potential. It occurs more than one order of magnitude later than the peak displacement current in the proton pump BR (35) and differs from fast charge transfer observed in other ChRs (36).

In the second reaction path, illumination of D₄₇₀ (C₁) results in all-trans,C=N-anti → 13-cis,C=N-syn isomerization and direct formation of P₄₈₀ (C₂), which is also photoreactive. Alternative photoreactions have been considered before to explain the biexponential decay of the conductive-state P₅₂₀ (17, 37, 38), and were assumed to involve an early all-trans,C=N-anti → 13-cis,C=N-syn isomerization based on NMR and low-temperature Raman measurements (22). Using isotopically labeled retinal and vibrational spectroscopy, we proved that the early all-trans,C=N-anti → 13-cis,C=N-syn double isomerization (also at ambient temperatures) causes early formation of P₄₈₀ (C₂). Consequently, the slowly decaying P₄₈₀ does not represent a late photocycle intermediate of the anti-cycle, as proposed in several previous publications (9, 11, 17, 37), but is the result of a reaction branching that occurs directly after photoexcitation of D₄₇₀ (C₁), possibly already during the excited-state lifetime. As shown by MD simulations, P₄₈₀ features a preopening of the central gate, allowing water influx, but remains nonconductive because the inner gate is still closed as previously proposed for the E90R chloride-conducting mutant (23).

In a third reaction path, photoactivation of P₄₈₀ (C₂) initiates the syn-cycle. Here, we identified P*₅₂₀^N as the conductive-state O₂. Under continuous illumination, P*₅₂₀^N accumulates due to its slow decay rate and significantly contributes to the stationary photocurrent, especially at high pH. Consequently, the parallel formation of P₅₂₀^N and P*₅₂₀^N accumulation accounts for the biexponential channel-closing kinetics and the evolution of proton conductance during continuous illumination (3, 39). Comparing P*₅₂₀^N accumulation in our FTIR measurements with the small photocurrent amplitude of O₂ in our electrophysiological recordings indicates a significantly reduced conductance of O₂ (P*₅₂₀^N) compared with O_1-early (P_390b^M2) and O_1-late (P₅₂₀^N), as previously predicted (12, 13). This now explains the ChR2 photocurrent inactivation during continuous illumination. It also explains the remarkably small shifts of the action spectra of the dark-adapted (D₄₇₀) and light-adapted (P₄₈₀) protein (40). The reduced sodium selectivity of P*₅₂₀^N (O₂) compared with P₅₂₀^N (O_1-late) indicates substantial differences in the open-pore structures of both conducting states that are further supported by distinct FTIR spectra for P*₅₂₀^N and P₅₂₀^N. In summary, the slower decay of the syn-cycle with reduced conductance leads to the accumulation of P₄₈₀ and now explains photocurrent inactivation.

Multiple dark states and different retinal isomers have been observed in other microbial rhodopsins, such as BR (41) and anabaena sensory rhodopsin (ASR) (42), before. Whereas we herein confirm photoactive all-trans,C=N-anti → 13-cis,C=N-syn as an alternative photoisomerization and an important step for light adaptation in ChRs, such a photoreaction has not been described for BR. What has been described for BR is a thermal equilibration between all-trans,C=N-anti (λ_max = 568 nm) and 13-cis,C=N-syn (λ_max = 548 nm) toward a ratio of roughly 6:4. Photoactivation of BR₅₄₈ initiates the syn-cycle that branches either early or late to the all-trans,C=N-anti state, accumulating all molecules in the BR₅₆₈ isoform (43, 44). The situation is different in ASR. Here, the fully dark-adapted state again contains an all-trans,C=N-anti chromophore. Photoactivation is thought to first cause a classic 13-trans to 13-cis isomerization and then a thermal obligatory C=N isomerization late during the photocycle, ending up with a 13-cis,C=N-syn light-adapted second dark state (45). Illumination of this second dark state is thought to cause 13-cis to trans-isomerization and, at the end of the syn-cycle, a thermal C=N syn- to anti-isomerization. In summary, in microbial rhodopsins, photochemical as well as thermal single isomerization around the C₁₃ = C₁₄ bond and photochemical and thermal double isomerization around C₁₃ = C₁₄ and C₁₅ = N are possible, but the efficiency of both reactions in light and in darkness, as well as the preferences of the directions, vary substantially within the diverse family.

Finally, we show that E90 is crucial for proton conductance in both photocycles and constitutes one key determinant for ion selectivity changes during continuous illumination with an intriguing double function depending on its protonation state before and after light adaptation. In the anti-cycle, E90 might be directly involved in proton transport as a proton shuttle or in the organization of water molecules, both of which bridge the distinct water-filled cavities seen in the dark-state crystal structure of ChR2 (31). In this scenario, E90 would favor proton selectivity, forming either a direct or indirect shortcut for protons that cannot be taken by larger cations at a similar efficiency. By similar means, the outer pore glutamates E139 and E143 in the highly proton-selective ChR Chrimson (46) or D112 in the voltage-gated proton channel Hv1 (47) were also shown to contribute to proton conductance and selectivity. Although the selectivity filter of ChR2, localized in the central gate, might be more permissive for larger cations than that of Chrimson, localized in the outer gate (48), in both cases, substitution of essential glutamates (E90 in ChR2 and E139 in Chrimson) by equally titratable histidines preserved proton selectivity, whereas substitution with the nontitratable glutamine or alanine impaired proton conductance (16, 46). In the anti-cycle, channel opening occurs with E90 staying protonated for the entire gating process. Accordingly, recent 4-μs molecular mechanics (MM) simulations on an ChR2 homology model based on the C1C2 chimera structure [PDB ID code 3ug9 (32)] showed impressively that minor hydrogen bond changes of E90 due to protonation of the counter-ion in the central gate region were sufficient to promote water invasion in the same time range (31). This water invasion weakens the electrostatic interactions of helix 2 and leads to channel opening. In our P₃₉₀ simulations for the ChR2 WT structure (30) (SI Appendix, Fig. S12), we observed a similar hydrogen bond rearrangement of E90 toward E123 that allowed water influx at the longer microsecond simulation times (30). Once deprotonated in the syn-cycle, E90 forms a salt bridge with the adjacent K93 and completely opens the central gate (as found for C₂), promoting helix hydration from the extracellular site, as shown in our MD simulations (Fig. 5). Although the proton acceptor of E90 has not been identified yet, the close proximity to water molecules, as indicated by our MD simulations, might allow fast proton diffusion into the bulk phase. Water molecules are expected to serve as proton shuttles as they can become transiently protonated, as previously shown for BR (7). It is essential to note that pore hydration due to hydrogen bond changes of E90 in P₃₉₀ differs from the pore hydration due to E90 deprotonation and salt bridge formation with K93 that we proposed earlier in our E90-Helix2-tilt model (18). Without considering parallel photocycles, we initially attributed early pore hydration due to E90 deprotonation to a pregating step in a linear photocycle. At the same time, early E90 deprotonation has been challenged by measurements of partly light-adapted ChR2, arguing that E90 is only deprotonated during the lifetime of P₄₈₀, which was assumed to be a late intermediate of a linear photocycle or a late branching reaction (11, 17, 49). Comparing FTIR measurements of dark- and light-adapted ChR2, we were able to resolve the controversy regarding E90 deprotonation. P₄₈₀ (C₂) is formed in an ultrafast branching reaction that leads to the very fast deprotonation of E90. The splitting of the photocycle into the anti- and syn-branches forms the basis for the light adaptation of ChR2. In the E90Q mutant, all-trans,C=N-anti → 13-cis,C=N-syn isomerization still occurs but can no longer trigger deprotonation of residue E90. Consequently, in E90Q, pore hydration in the syn-cycle is reduced and conformational changes during formation of P*₅₂₀^N are no longer sufficient to support passive proton flux in the syn-cycle. As the syn-cycle is still populated in the E90Q mutant but nonconductive, photocurrents still inactivate during continuous illumination at a degree determined by the relative rate constants. As an essential revision of our previous E90-Helix2-tilt model, we therefore reassign early pore hydration to the light-dependent transition to the syn-cycle. Deprotonation of E90 and subsequent pore hydration prepare proton conductance of P*₅₂₀^N in the syn-cycle. However, even if E90 is not deprotonated in the anti-cycle, it seems still crucial for ion selectivity.

In a combined study of single-turnover electrophysiology and FTIR and Raman spectroscopy with isotopic retinal labeling, site-directed mutagenesis, and MD simulations, we developed a unifying two-photocycle model that simplifies and embraces previous kinetic models and completely resolves the channel gating, light adaptation, and temporal changes in ion selectivity. Identifying the corresponding molecular transitions, we may facilitate future protein engineering of ChR variants with reduced or improved photocurrent inactivation for optogenetic applications, requiring either a stable response to continuous illumination or a transient response to light switching. Early photocycle branching by alternative retinal isomerization and the corresponding large conformational protein changes that do not directly lead to channel opening will need careful consideration for the interpretation of molecular gating transitions observed in time-resolved spectroscopy, crystallography, and optogenetic experiments.

It turns out that only tiny light-induced alterations are crucial for the specific protein function. The elucidation of the molecular reaction mechanisms of proteins therefore deserves vibrational spectroscopic techniques like time-resolved FTIR with nanoscale spatiotemporal resolution.

Materials and Methods

Yeast Culture.

P. pastoris strain SMD1163 cells (kindly gifted by C. Bamann, Max Planck Institute of Biophysics, Frankfurt) containing the pPIC9KChR2His10 construct were precultured in buffered glycerol complex (BMGY) medium (50). Expression of ChR2 was induced in buffered methanol complex (BMMY) medium containing 2.5 μM all-trans retinal (either ¹²C, ¹³C₁₄,¹³C₁₅-labeled or ¹³C₁₀,¹³C₁₁-labeled) and 0.00004% biotin at an initial OD₆₀₀ of 1 and at 30 °C and 120 rpm. Cells were harvested at an OD₆₀₀ of 20 by centrifugation.

Membrane Preparation and Protein Purification.

Cells were disrupted using a BeadBeater (Biospec Products), and membranes were isolated by ultracentrifugation. Homogenized membranes were solubilized with 1% decylmaltoside overnight. ChR2 purification was done by nickel-nitrilotriacetic acid affinity chromatography and subsequent gel filtration using a HiLoad 16/600 Superdex 200-pg column (General Electric).

Reconstitution of ChR2 into DPPC or EggPC.

The purified ChR2 was reconstituted into DPPC (Avanti Polar Lipids) or EggPC (Avanti Polar Lipids). The lipids were solubilized with 2% cholate in 20 mM Hepes (pH 7.5), 100 mM NaCl, and 1 mM MgCl₂ by incubation at 50 °C for 10 min. Solubilized lipids and purified ChR2 were mixed at a 2:1 ratio [lipid/protein (wt/wt)] and incubated for 20 min. Detergent was removed overnight either by adsorption on Bio-Beads SM 2 (BioRad) or by dialysis.

The resulting suspension containing proteoliposomes and buffer was ultracentrifuged at 200,000 × g for 2 h, and the pellet was then transferred and squeezed between two CaF₂ slides to obtain an optical path length between 5 μm and 10 μm. This sample was then placed in a vacuum-tight cuvette.

Preparation of HEK Cells.

Electrophysiological recordings were performed on stably expressing the ChR2-mVenus fusion construct HEK cell line (34), as previously described in detail (51). Briefly, HEK cells were cultured at 5% CO₂ and 37 °C in DMEM supplemented with 10% FBS, 100 μg/mL penicillin/streptomycin (Biochrom), 200 μg/mL zeocin, and 50 μg/mL blasticidin (Thermo Fisher Scientific). Cells were seeded onto polylysine-coated glass coverslips at a concentration of 1 × 10⁵ cells per milliliter and supplemented with a final concentration of 1 μM all-trans retinal (Sigma–Aldrich). Induction of ChR2-mVenus expression was induced by addition of 0.1 µM tetracycline (Thermo Fisher Scientific).

Patch-Clamp Experiments in HEK293 Cells.

Patch pipettes were pulled using a P1000 micropipette puller (Sutter Instruments) and fire-polished. Pipette resistance was 1.5–2.5 MΩ. A 140 mM NaCl agar bridge served as a reference (bath) electrode. In whole-cell recordings, membrane resistance was typically >1 GΩ, while access resistance was below 10 MΩ. Pipette capacity, series resistance, and cell capacity compensation were applied. All experiments were carried out at 23 °C. Signals were amplified (AxoPatch200B), digitized (DigiData1400), and acquired using Clampex 10.4 software (all from Molecular Devices). Holding potentials were varied in 15-mV steps between −60 and +30 mV. Extracellular buffer exchange was performed manually by adding at least 5 mL of the respective buffer to the recording chamber (500-μL chamber volume), while a Ringer Bath Handler MPCU (Lorenz Messgerätebau) maintained a constant bath level. Standard bath solutions contained 110 mM NaCl, 1 mM KCl, 1 mM CsCl, 2 mM CaCl₂, 2 mM MgCl₂, and 10 mM Hepes at extracellular pH (pH_e) 7.2 (with glucose added up to 310 mOsm). Standard pipette solutions contained 110 mM NaCl, 1 mM KCl, 1 mM CsCl, 2 mM CaCl₂, 2 mM MgCl₂, 10 mM EGTA, and 10 mM Hepes at pH_i 7.2 or 10 mM Tris at intracellular pH (pH_i) 9.0 (glucose was added up to 290 mOsm). For ion selectivity measurements, either NaCl was replaced by 110 mM N-methyl-d-glucamine or extracellular pH was adjusted to pH_e 9.0 by buffering with 10 mM Tris instead of Hepes.

Continuous light was generated using a Polychrome V light source (TILL Photonics) set to 470 ± 7 nm. Light exposure was controlled with a programmable shutter system (VS25 and VCM-D1; Vincent Associates). The Polychrome V light intensity was 3.4 mW/mm² in the sample plane, measured with a calibrated optometer (P9710; Gigahertz Optik). Light intensities were calculated for the illuminated field of the W Plan-Apochromat 40×/1.0 differential interference contrast objective (0.066 mm²; Carl Zeiss). For delivery of 470-nm⋅ns⁻¹ laser pulses, an Opolette polette HENd:YAG laser/OPO system (OPOTEK) was coupled/decoupled into a M37L02 multimode fiber patch cable with a modified KT110/M free space-to-fiber coupler using AC127 019 A ML achromatic doublets (Thorlabs). Single pulses were selected using a LS6ZM2 shutter (Vincent Associates). Laser intensity was set to 5% using the built-in motorized variable attenuator, resulting in a pulse energy of 100 ± 20 μJ/mm². Pulse energies were measured with a calibrated S470C thermal power sensor and a PM100D power and energy meter (Thorlabs) after passing through all of the optics. Actinic light was coupled into an Axiovert 100 microscope (Carl Zeiss) and delivered to the sample using a 90/10 beamsplitter (Chroma). To toggle between activation with the laser and the Polychrome V light source, a BB1 E02 broadband dielectric mirror mounted on an MFF101/M motorized filter flip mount (Thorlabs) was used. Data were filtered at 100 kHz and sampled at 250 kHz. Due to minimal timing uncertainties, each acquired sweep was time-shifted after measurements to align it with the rising edges of the Q-switch signals of the activating laser pulses. Photocurrents were binned to 50 logarithmically spaced data points per temporal decade with custom-written MATLAB script (MathWorks).

FTIR Experiments.

To gain insight into the changes upon illumination, we performed time-resolved FTIR difference spectroscopy at 15 °C. For the continuous light experiments, the sample was illuminated with a blue LED (λ_max = 465 nm) for 5 s. Spectra were recorded before switching on the light (reference), during illumination [accumulation of the stationary photoproduct (P_Stat) for 5 s] and after switching off the light (decay of P_Stat for 500 s) using the conventional rapid scan mode of the spectrometer. Difference spectra were calculated using the Beer–Lambert law, which results in positive photo product bands and negative educt bands in the difference spectra. For the single-turnover measurements, the sample was illuminated with a short laser pulse of an excimer laser-driven dye laser (Coumarin 102 dye; λ_max = 475 nm, pulse width ∼ 50 ns). Conventional rapid scan experiments (time resolution ∼ 10 ms, spectral resolution = 4 cm⁻¹) were performed with a sufficient relaxation time (t_relax = 200 s) and flash frequency (f_flash = 0.005 Hz) between the flashes to allow the D₄₇₀ to significantly repopulate ([D₄₇₀] became ∼96%).

For a comparison with the photocycle under the “shortcut condition,” the flash frequency was increased (t_relax = 5 s, f_flash = 0.2 Hz). Using this approach, equilibrium between D₄₇₀ and P₄₈₀ emerges and the ChR2 molecules start the photocycle from both states. The datasets were then analyzed by a global-fitting routine as presented previously (16, 18, 52, 53) to isolate the decay-associated amplitude spectra of the transitions involved in D₄₇₀ recovery. To get access to the earlier intermediates of the dark-adapted ([D₄₇₀] ∼ 91%) photocycle of H134R (Fig. 3A), step-scan measurements were performed with a light pulse repetition rate of 0.007 Hz (t_relax = 140-s, detector rise time = 50 ns, resolution = 8 cm⁻¹, wavenumber range: 0–1974 cm⁻¹) as already published for the WT (18). One measurement was completed after 22 h, and ∼15 measurements were averaged to give the final result. H134R used for the step scan was expressed in COS (abbreviation for CV-1 in Origin with SV40 genes) cells and prepared as described in our earlier publication (18).

Raman Experiments.

The Raman experiments were performed with samples that were prepared exactly the same as those for the rapid scan FTIR experiments, but with a higher optical path length (20–50 μm). The room temperature was ∼18 °C. We used the Raman microscope XPloRA One (HORIBA Scientific) to scan the sample. To prevent sample degradation due to long illumination, we performed measurements at a 785-nm excitation wavelength to ensure the lowest possible photoexcitation of the sample and a sufficient enhancement of the Raman signal due to the preresonant Raman effect. Laser power at the sample position was 28 mW. A 50× objective (Olympus LCPLN-IR) was used, resulting in a confocal volume of ∼1 μm³ in the sample plane.

To excite the D₄₇₀ state of the sample and create a photoproduct with a high P₄₈₀ fraction, the sample was illuminated with an external blue-light source (100-W halogen lamp filtered with a 470-nm filter coupled to the observation beam path of the microscope). Sample illumination was controlled by a shutter between the lamp and sample.

To ensure that the measured photoproduct spectra are free of contamination by their preceding intermediates (P₅₂₀^N and P₃₉₀^M) under continuous illumination, a controlled illumination/relaxation experiment was performed:

Laser on + illumination on (formation of P_Stat)
Wait 0.5 s
Acquisition of spectrum (t_integration = 2 s); P_Stat is measured
Laser on + illumination off (slow relaxation of P_Stat: t_1/2 ∼ 40 s)
Wait 0.5 s
Acquisition of spectrum (t_integration = 2 s); high fraction of P₄₈₀ free of illumination artifacts is measured (P′_Stat)
Go to next position on the sample

This procedure was repeated for a 15 × 15 spot matrix (pixel spacing = 5 μm) of the sample, and the acquired spectra were averaged for each illumination condition.

Next, the illumination was stopped and a relaxation phase of at least 5 min was commenced to allow full relaxation of the generated P₄₈₀. The dark-state D₄₇₀ was measured for the same spot matrix (t_integration = 2 s).

To obtain the pure lamp artifact, the same area was measured without the laser with only illumination of the sample. This spectrum was later subtracted from the spectra measured under continuous illumination.

The complete protocol was performed for three spectral regions (center wavenumbers: 900 cm⁻¹, 1,250 cm⁻¹, and 1,550 cm⁻¹), which were then combined to obtain the complete spectrum ranging from 650 cm⁻¹ to 1,700 cm⁻¹ with a wavenumber spacing of ∼0.2 cm⁻¹ and a nominal resolution of 0.6 cm⁻¹.

MD Simulations.

The MD simulations were performed according to our previous reports (16, 18), except for the force field and the GROMACS version used. We used the Optimized Potentials for Liquid Simulations/All-Atom (OPLS/AA) force field and GROMACS version 2016.3. A series of 5 × 100-ns independent and unrestrained MD simulations was performed for each protonation state of E90 with the respective chromophore configuration. The MD simulations were performed consecutively using the resulting structures of dark-adapted ChR2 (all-trans,C=N-anti with protonated E90) given by MD simulations as a starting point for the isomerization (discussed below). Each MD simulation was initiated using a different temperature seed number to generate the random distribution of starting velocities.

Water Dynamic and Run-Average Structure.

The water dynamic and run-average structures were calculated according to our previous report (18).

Retinal Isomerization.

Retinal all-trans,C=N-anti to 13-cis,C=N-anti isomerization was performed as described earlier (18), achieved via the following scheme. The torsion angles of the C₁₃ = C₁₄ and C=N double bonds were tilted counterclockwise in 20° steps, starting in the range 0–180°. For each tilting step, the retinal + K257 (without backbone) atoms were maintained as a freeze group and the rest of the simulation system was allowed to relax in a 10-ns unrestrained MD simulation as described above.

The resulting 13-cis retinal structures served as starting structures for the MD simulations of the different intermediates (P₅₀₀^K, P₄₈₀, and P₄₈₀-E90p) with protonated and deprotonated E90. For the starting structures for the P_390a^M1 intermediate, we used the final structures of the P₅₀₀^K simulations after Schiff base deprotonation and D253 protonation.

Acknowledgments

We thank Harald Chrongiewski and Gabi Smuda for technical assistance. We also thank Mathias Lübben and Till Rudack for helpful discussions. We thank Maila Reh, Altina Klein, and Tharsana Tharmalingam for technical assistance. We also thank Christiane Grimm, Joel Kaufmann, and Franz Bartl for fruitful discussions. This work was supported by the Deutsche Forschungsgemeinschaft (DFG) Priority Programme SPP 1926 and by DFG Grant SFB1078 (B2) and the Cluster of Excellence Unifying Concepts in Catalysis Berlin International Graduate School of natural Science and Engineering (BIG-NSE) (to J.V.) and E4 (to P.H.). P.H. is a Senior Research Professor of the Hertie Foundation. The spectroscopic and molecular dynamic part of this work was first presented at the 17th International Conference on Retinal Proteins in September 2016 in Potsdam, Germany. The electrophysological work was presented for the first time at the 18th International Conference on Retinal Proteins in September 2018 in Ontario, Canada. This work was prepublished on the preprint server bioRxiv in December 2018.

Footnotes

↵¹J.K. and J.V. contributed equally to this work.
↵²To whom correspondence may be addressed. Email: hegemann{at}rz.hu-berlin.de or klaus.gerwert{at}bph.rub.de.

Author contributions: J.K., J.V., P.H., and K. Gerwert designed research; J.K., J.V., and S.A.T. performed research; M.-A.D., J.W., D.P., K. Gavriljuk, and S.F.E.-M. contributed new reagents/analytic tools; J.K., J.V., S.A.T., and S.F.E.-M. analyzed data; and J.K., J.V., S.A.T., M.-A.D., J.W., P.H., and K. Gerwert wrote the paper.
The authors declare no conflict of interest.
This article is a PNAS Direct Submission.
This article contains supporting information online at www.pnas.org/lookup/suppl/doi:10.1073/pnas.1818707116/-/DCSupplemental.

This open access article is distributed under Creative Commons Attribution-NonCommercial-NoDerivatives License 4.0 (CC BY-NC-ND).

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References

↵
1. Zhang F, et al.
(2011) The microbial opsin family of optogenetic tools. Cell 147:1446–1457.
OpenUrl CrossRef PubMed
↵
1. Scheib U, et al.
(2015) The rhodopsin-guanylyl cyclase of the aquatic fungus Blastocladiella emersonii enables fast optical control of cGMP signaling. Sci Signal 8:rs8.
OpenUrl Abstract/FREE Full Text
↵
1. Nagel G, et al.
(2003) Channelrhodopsin-2, a directly light-gated cation-selective membrane channel. Proc Natl Acad Sci USA 100:13940–13945.
OpenUrl Abstract/FREE Full Text
↵
1. Boyden ES,
2. Zhang F,
3. Bamberg E,
4. Nagel G,
5. Deisseroth K
(2005) Millisecond-timescale, genetically targeted optical control of neural activity. Nat Neurosci 8:1263–1268.
OpenUrl CrossRef PubMed
↵
1. Gerwert K,
2. Hess B,
3. Soppa J,
4. Oesterhelt D
(1989) Role of aspartate-96 in proton translocation by bacteriorhodopsin. Proc Natl Acad Sci USA 86:4943–4947.
OpenUrl Abstract/FREE Full Text
↵
1. Lanyi JK
(1992) Proton transfer and energy coupling in the bacteriorhodopsin photocycle. J Bioenerg Biomembr 24:169–179.
OpenUrl CrossRef PubMed
↵
1. Garczarek F,
2. Gerwert K
(2006) Functional waters in intraprotein proton transfer monitored by FTIR difference spectroscopy. Nature 439:109–112.
OpenUrl CrossRef PubMed
↵
1. Gerwert K
(2017) Channelrhodopsin reveals its dark secrets. Science 358:1000–1001.
OpenUrl Abstract/FREE Full Text
↵
1. Ritter E,
2. Stehfest K,
3. Berndt A,
4. Hegemann P,
5. Bartl FJ
(2008) Monitoring light-induced structural changes of Channelrhodopsin-2 by UV-visible and Fourier transform infrared spectroscopy. J Biol Chem 283:35033–35041.
OpenUrl Abstract/FREE Full Text
↵
1. Bamann C,
2. Kirsch T,
3. Nagel G,
4. Bamberg E
(2008) Spectral characteristics of the photocycle of channelrhodopsin-2 and its implication for channel function. J Mol Biol 375:686–694.
OpenUrl CrossRef PubMed
↵
1. Lórenz-Fonfría VA, et al.
(2015) Temporal evolution of helix hydration in a light-gated ion channel correlates with ion conductance. Proc Natl Acad Sci USA 112:E5796–E5804.
OpenUrl Abstract/FREE Full Text
↵
1. Hegemann P,
2. Ehlenbeck S,
3. Gradmann D
(2005) Multiple photocycles of channelrhodopsin. Biophys J 89:3911–3918.
OpenUrl CrossRef PubMed
↵
1. Nikolic K, et al.
(2009) Photocycles of channelrhodopsin-2. Photochem Photobiol 85:400–411.
OpenUrl CrossRef PubMed
↵
1. Valentini A, et al.
(2017) Optomechanical control of quantum yield in trans-cis ultrafast photoisomerization of a retinal chromophore model. Angew Chem Int Ed Engl 56:3842–3846.
OpenUrl
↵
1. Verhoefen M-K, et al.
(2010) The photocycle of channelrhodopsin-2: Ultrafast reaction dynamics and subsequent reaction steps. ChemPhysChem 11:3113–3122.
OpenUrl CrossRef PubMed
↵
1. Eisenhauer K, et al.
(2012) In channelrhodopsin-2 Glu-90 is crucial for ion selectivity and is deprotonated during the photocycle. J Biol Chem 287:6904–6911.
OpenUrl Abstract/FREE Full Text
↵
1. Lórenz-Fonfría VA, et al.
(2013) Transient protonation changes in channelrhodopsin-2 and their relevance to channel gating. Proc Natl Acad Sci USA 110:E1273–E1281.
OpenUrl Abstract/FREE Full Text
↵
1. Kuhne J, et al.
(2015) Early formation of the ion-conducting pore in channelrhodopsin-2. Angew Chem Int Ed Engl 54:4953–4957.
OpenUrl CrossRef PubMed
↵
1. Gerwert K,
2. Freier E,
3. Wolf S
(2014) The role of protein-bound water molecules in microbial rhodopsins. Biochim Biophys Acta 1837:606–613.
OpenUrl CrossRef PubMed
↵
1. Ritter E,
2. Piwowarski P,
3. Hegemann P,
4. Bartl FJ
(2013) Light-dark adaptation of channelrhodopsin C128T mutant. J Biol Chem 288:10451–10458.
OpenUrl Abstract/FREE Full Text
↵
1. Becker-Baldus J, et al.
(2015) Enlightening the photoactive site of channelrhodopsin-2 by DNP-enhanced solid-state NMR spectroscopy. Proc Natl Acad Sci USA 112:9896–9901.
OpenUrl Abstract/FREE Full Text
↵
1. Bruun S, et al.
(2015) Light-dark adaptation of channelrhodopsin involves photoconversion between the all-trans and 13-cis retinal isomers. Biochemistry 54:5389–5400.
OpenUrl PubMed
↵
1. Wietek J, et al.
(2014) Conversion of channelrhodopsin into a light-gated chloride channel. Science 344:409–412.
OpenUrl Abstract/FREE Full Text
↵
1. Kaufmann JCD, et al.
(2017) Proton transfer reactions in the red light-activatable channelrhodopsin variant ReaChR and their relevance for its function. J Biol Chem 292:14205–14216.
OpenUrl Abstract/FREE Full Text
↵
1. Berndt A, et al.
(2011) High-efficiency channelrhodopsins for fast neuronal stimulation at low light levels. Proc Natl Acad Sci USA 108:7595–7600.
OpenUrl Abstract/FREE Full Text
↵
1. Gerwert K,
2. Siebert F
(1986) Evidence for light-induced 13-cis, 14-s-cis isomerization in bacteriorhodopsin obtained by FTIR difference spectroscopy using isotopically labelled retinals. EMBO J 5:805–811.
OpenUrl
↵
1. Smith SO, et al.
(1984) Determination of retinal Schiff base configuration in bacteriorhodopsin. Proc Natl Acad Sci USA 81:2055–2059.
OpenUrl Abstract/FREE Full Text
↵
1. Babitzki G,
2. Mathias G,
3. Tavan P
(2009) The infrared spectra of the retinal chromophore in bacteriorhodopsin calculated by a DFT/MM approach. J Phys Chem B 113:10496–10508.
OpenUrl PubMed
↵
1. Lórenz-Fonfría VA,
2. Muders V,
3. Schlesinger R,
4. Heberle J
(2014) Changes in the hydrogen-bonding strength of internal water molecules and cysteine residues in the conductive state of channelrhodopsin-1. J Chem Phys 141:22D507.
OpenUrl CrossRef PubMed
↵
1. Volkov O, et al.
(2017) Structural insights into ion conduction by channelrhodopsin 2. Science 358:eaan8862.
OpenUrl Abstract/FREE Full Text
↵
1. Ardevol A,
2. Hummer G
(2018) Retinal isomerization and water-pore formation in channelrhodopsin-2. Proc Natl Acad Sci USA 115:3557–3562.
OpenUrl Abstract/FREE Full Text
↵
1. Kato HE, et al.
(2012) Crystal structure of the channelrhodopsin light-gated cation channel. Nature 482:369–374.
OpenUrl CrossRef PubMed
↵
1. Ruffert K, et al.
(2011) Glutamate residue 90 in the predicted transmembrane domain 2 is crucial for cation flux through channelrhodopsin 2. Biochem Biophys Res Commun 410:737–743.
OpenUrl CrossRef PubMed
↵
1. Feldbauer K, et al.
(2009) Channelrhodopsin-2 is a leaky proton pump. Proc Natl Acad Sci USA 106:12317–12322.
OpenUrl Abstract/FREE Full Text
↵
1. Keszthelyi L,
2. Ormos P
(2001) Electric signals associated with the photocycle of bacteriorhodopsin. FEBS Lett 109:189–193.
OpenUrl
↵
1. Sineshchekov OA,
2. Govorunova EG,
3. Wang J,
4. Li H,
5. Spudich JL
(2013) Intramolecular proton transfer in channelrhodopsins. Biophys J 104:807–817.
OpenUrl CrossRef PubMed
↵
1. Bamann C,
2. Gueta R,
3. Kleinlogel S,
4. Nagel G,
5. Bamberg E
(2010) Structural guidance of the photocycle of channelrhodopsin-2 by an interhelical hydrogen bond. Biochemistry 49:267–278.
OpenUrl CrossRef PubMed
↵
1. Stehfest K,
2. Hegemann P
(2010) Evolution of the channelrhodopsin photocycle model. ChemPhysChem 11:1120–1126.
OpenUrl CrossRef PubMed
↵
1. Berndt A,
2. Prigge M,
3. Gradmann D,
4. Hegemann P
(2010) Two open states with progressive proton selectivities in the branched channelrhodopsin-2 photocycle. Biophys J 98:753–761.
OpenUrl CrossRef PubMed
↵
1. Lin JY,
2. Lin MZ,
3. Steinbach P,
4. Tsien RY
(2009) Characterization of engineered channelrhodopsin variants with improved properties and kinetics. Biophys J 96:1803–1814.
OpenUrl CrossRef PubMed
↵
1. Sperling W,
2. Carl P,
3. Rafferty Ch,
4. Dencher NA
(1977) Photochemistry and dark equilibrium of retinal isomers and bacteriorhodopsin isomers. Biophys Struct Mech 3:79–94.
OpenUrl CrossRef PubMed
↵
1. Sineshchekov OA,
2. Spudich EN,
3. Trivedi VD,
4. Spudich JL
(2006) Role of the cytoplasmic domain in Anabaena sensory rhodopsin photocycling: Vectoriality of Schiff base deprotonation. Biophys J 91:4519–4527.
OpenUrl CrossRef PubMed
↵
1. Bryl K,
2. Taui M,
3. Yoshizawa M,
4. Kobayashi T
(1992) Light adaptation of dark-adapted bacteriorhodopsin studied by nanosecond time-resolved absorption spectroscopy. Photochem Photobiol 56:1013–1018.
OpenUrl
↵
1. Hofrichter J,
2. Henry ER,
3. Lozier RH
(1989) Photocycles of bacteriorhodopsin in light- and dark-adapted purple membrane studied by time-resolved absorption spectroscopy. Biophys J 56:693–706.
OpenUrl CrossRef PubMed
↵
1. Kawanabe A,
2. Furutani Y,
3. Jung K-H,
4. Kandori H
(2007) Photochromism of Anabaena sensory rhodopsin. J Am Chem Soc 129:8644–8649.
OpenUrl CrossRef PubMed
↵
1. Vierock J,
2. Grimm C,
3. Nitzan N,
4. Hegemann P
(2017) Molecular determinants of proton selectivity and gating in the red-light activated channelrhodopsin Chrimson. Sci Rep 7:9928.
OpenUrl
↵
1. Dudev T, et al.
(2015) Selectivity mechanism of the voltage-gated proton channel, H_V1. Sci Rep 5:10320.
OpenUrl CrossRef PubMed
↵
1. Oda K, et al.
(2018) Crystal structure of the red light-activated channelrhodopsin Chrimson. Nat Commun 9:3949.
OpenUrl
↵
1. Saita M, et al.
(2018) Photoexcitation of the P₄⁴⁸⁰ state induces a secondary photocycle that potentially desensitizes channelrhodopsin-2. J Am Chem Soc 140:9899–9903.
OpenUrl
↵
1. Radu I, et al.
(2009) Conformational changes of channelrhodopsin-2. J Am Chem Soc 131:7313–7319.
OpenUrl CrossRef PubMed
↵
1. Grimm C,
2. Vierock J,
3. Hegemann P,
4. Wietek J
(2017) Whole-cell patch-clamp recordings for electrophysiological determination of ion selectivity in channelrhodopsins. J Vis Exp, e55497.
↵
1. Müller K-H,
2. Plesser T
(1991) Variance reduction by simultaneous multi-exponential analysis of data sets from different experiments. Eur Biophys J 19:231–240.
OpenUrl
↵
1. Hessling B,
2. Souvignier G,
3. Gerwert K
(1993) A model-independent approach to assigning bacteriorhodopsin’s intramolecular reactions to photocycle intermediates. Biophys J 65:1929–1941.
OpenUrl CrossRef PubMed

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Article Classifications

Biological Sciences
Biophysics and Computational Biology

[1] ↵
Zhang F, et al.
(2011) The microbial opsin family of optogenetic tools. Cell 147:1446–1457.
OpenUrl CrossRef PubMed

[2] Zhang F, et al.

[3] ↵
Scheib U, et al.
(2015) The rhodopsin-guanylyl cyclase of the aquatic fungus Blastocladiella emersonii enables fast optical control of cGMP signaling. Sci Signal 8:rs8.
OpenUrl Abstract/FREE Full Text

[4] Scheib U, et al.

[5] ↵
Nagel G, et al.
(2003) Channelrhodopsin-2, a directly light-gated cation-selective membrane channel. Proc Natl Acad Sci USA 100:13940–13945.
OpenUrl Abstract/FREE Full Text

[6] Nagel G, et al.

[7] ↵
Boyden ES,
Zhang F,
Bamberg E,
Nagel G,
Deisseroth K
(2005) Millisecond-timescale, genetically targeted optical control of neural activity. Nat Neurosci 8:1263–1268.
OpenUrl CrossRef PubMed

[8] Boyden ES,

[9] Zhang F,

[10] Bamberg E,

[11] Nagel G,

[12] Deisseroth K

[13] ↵
Gerwert K,
Hess B,
Soppa J,
Oesterhelt D
(1989) Role of aspartate-96 in proton translocation by bacteriorhodopsin. Proc Natl Acad Sci USA 86:4943–4947.
OpenUrl Abstract/FREE Full Text

[14] Gerwert K,

[15] Hess B,

[16] Soppa J,

[17] Oesterhelt D

[18] ↵
Lanyi JK
(1992) Proton transfer and energy coupling in the bacteriorhodopsin photocycle. J Bioenerg Biomembr 24:169–179.
OpenUrl CrossRef PubMed

[19] Lanyi JK

[20] ↵
Garczarek F,
Gerwert K
(2006) Functional waters in intraprotein proton transfer monitored by FTIR difference spectroscopy. Nature 439:109–112.
OpenUrl CrossRef PubMed

[21] Garczarek F,

[22] Gerwert K

[23] ↵
Gerwert K
(2017) Channelrhodopsin reveals its dark secrets. Science 358:1000–1001.
OpenUrl Abstract/FREE Full Text

[24] Gerwert K

[25] ↵
Ritter E,
Stehfest K,
Berndt A,
Hegemann P,
Bartl FJ
(2008) Monitoring light-induced structural changes of Channelrhodopsin-2 by UV-visible and Fourier transform infrared spectroscopy. J Biol Chem 283:35033–35041.
OpenUrl Abstract/FREE Full Text

[26] Ritter E,

[27] Stehfest K,

[28] Berndt A,

[29] Hegemann P,

[30] Bartl FJ

[31] ↵
Bamann C,
Kirsch T,
Nagel G,
Bamberg E
(2008) Spectral characteristics of the photocycle of channelrhodopsin-2 and its implication for channel function. J Mol Biol 375:686–694.
OpenUrl CrossRef PubMed

[32] Bamann C,

[33] Kirsch T,

[34] Nagel G,

[35] Bamberg E

[36] ↵
Lórenz-Fonfría VA, et al.
(2015) Temporal evolution of helix hydration in a light-gated ion channel correlates with ion conductance. Proc Natl Acad Sci USA 112:E5796–E5804.
OpenUrl Abstract/FREE Full Text

[37] Lórenz-Fonfría VA, et al.

[38] ↵
Hegemann P,
Ehlenbeck S,
Gradmann D
(2005) Multiple photocycles of channelrhodopsin. Biophys J 89:3911–3918.
OpenUrl CrossRef PubMed

[39] Hegemann P,

[40] Ehlenbeck S,

[41] Gradmann D

[42] ↵
Nikolic K, et al.
(2009) Photocycles of channelrhodopsin-2. Photochem Photobiol 85:400–411.
OpenUrl CrossRef PubMed

[43] Nikolic K, et al.

[44] ↵
Valentini A, et al.
(2017) Optomechanical control of quantum yield in trans-cis ultrafast photoisomerization of a retinal chromophore model. Angew Chem Int Ed Engl 56:3842–3846.
OpenUrl

[45] Valentini A, et al.

[46] ↵
Verhoefen M-K, et al.
(2010) The photocycle of channelrhodopsin-2: Ultrafast reaction dynamics and subsequent reaction steps. ChemPhysChem 11:3113–3122.
OpenUrl CrossRef PubMed

[47] Verhoefen M-K, et al.

[48] ↵
Eisenhauer K, et al.
(2012) In channelrhodopsin-2 Glu-90 is crucial for ion selectivity and is deprotonated during the photocycle. J Biol Chem 287:6904–6911.
OpenUrl Abstract/FREE Full Text

[49] Eisenhauer K, et al.

[50] ↵
Lórenz-Fonfría VA, et al.
(2013) Transient protonation changes in channelrhodopsin-2 and their relevance to channel gating. Proc Natl Acad Sci USA 110:E1273–E1281.
OpenUrl Abstract/FREE Full Text

[51] Lórenz-Fonfría VA, et al.

[52] ↵
Kuhne J, et al.
(2015) Early formation of the ion-conducting pore in channelrhodopsin-2. Angew Chem Int Ed Engl 54:4953–4957.
OpenUrl CrossRef PubMed

[53] Kuhne J, et al.

[54] ↵
Gerwert K,
Freier E,
Wolf S
(2014) The role of protein-bound water molecules in microbial rhodopsins. Biochim Biophys Acta 1837:606–613.
OpenUrl CrossRef PubMed

[55] Gerwert K,

[56] Freier E,

[57] Wolf S

[58] ↵
Ritter E,
Piwowarski P,
Hegemann P,
Bartl FJ
(2013) Light-dark adaptation of channelrhodopsin C128T mutant. J Biol Chem 288:10451–10458.
OpenUrl Abstract/FREE Full Text

[59] Ritter E,

[60] Piwowarski P,

[61] Hegemann P,

[62] Bartl FJ

[63] ↵
Becker-Baldus J, et al.
(2015) Enlightening the photoactive site of channelrhodopsin-2 by DNP-enhanced solid-state NMR spectroscopy. Proc Natl Acad Sci USA 112:9896–9901.
OpenUrl Abstract/FREE Full Text

[64] Becker-Baldus J, et al.

[65] ↵
Bruun S, et al.
(2015) Light-dark adaptation of channelrhodopsin involves photoconversion between the all-trans and 13-cis retinal isomers. Biochemistry 54:5389–5400.
OpenUrl PubMed

[66] Bruun S, et al.

[67] ↵
Wietek J, et al.
(2014) Conversion of channelrhodopsin into a light-gated chloride channel. Science 344:409–412.
OpenUrl Abstract/FREE Full Text

[68] Wietek J, et al.

[69] ↵
Kaufmann JCD, et al.
(2017) Proton transfer reactions in the red light-activatable channelrhodopsin variant ReaChR and their relevance for its function. J Biol Chem 292:14205–14216.
OpenUrl Abstract/FREE Full Text

[70] Kaufmann JCD, et al.

[71] ↵
Berndt A, et al.
(2011) High-efficiency channelrhodopsins for fast neuronal stimulation at low light levels. Proc Natl Acad Sci USA 108:7595–7600.
OpenUrl Abstract/FREE Full Text

[72] Berndt A, et al.

[73] ↵
Gerwert K,
Siebert F
(1986) Evidence for light-induced 13-cis, 14-s-cis isomerization in bacteriorhodopsin obtained by FTIR difference spectroscopy using isotopically labelled retinals. EMBO J 5:805–811.
OpenUrl

[74] Gerwert K,

[75] Siebert F

[76] ↵
Smith SO, et al.
(1984) Determination of retinal Schiff base configuration in bacteriorhodopsin. Proc Natl Acad Sci USA 81:2055–2059.
OpenUrl Abstract/FREE Full Text

[77] Smith SO, et al.

[78] ↵
Babitzki G,
Mathias G,
Tavan P
(2009) The infrared spectra of the retinal chromophore in bacteriorhodopsin calculated by a DFT/MM approach. J Phys Chem B 113:10496–10508.
OpenUrl PubMed

[79] Babitzki G,

[80] Mathias G,

[81] Tavan P

[82] ↵
Lórenz-Fonfría VA,
Muders V,
Schlesinger R,
Heberle J
(2014) Changes in the hydrogen-bonding strength of internal water molecules and cysteine residues in the conductive state of channelrhodopsin-1. J Chem Phys 141:22D507.
OpenUrl CrossRef PubMed

[83] Lórenz-Fonfría VA,

[84] Muders V,

[85] Schlesinger R,

[86] Heberle J

[87] ↵
Volkov O, et al.
(2017) Structural insights into ion conduction by channelrhodopsin 2. Science 358:eaan8862.
OpenUrl Abstract/FREE Full Text

[88] Volkov O, et al.

[89] ↵
Ardevol A,
Hummer G
(2018) Retinal isomerization and water-pore formation in channelrhodopsin-2. Proc Natl Acad Sci USA 115:3557–3562.
OpenUrl Abstract/FREE Full Text

[90] Ardevol A,

[91] Hummer G

[92] ↵
Kato HE, et al.
(2012) Crystal structure of the channelrhodopsin light-gated cation channel. Nature 482:369–374.
OpenUrl CrossRef PubMed

[93] Kato HE, et al.

[94] ↵
Ruffert K, et al.
(2011) Glutamate residue 90 in the predicted transmembrane domain 2 is crucial for cation flux through channelrhodopsin 2. Biochem Biophys Res Commun 410:737–743.
OpenUrl CrossRef PubMed

[95] Ruffert K, et al.

[96] ↵
Feldbauer K, et al.
(2009) Channelrhodopsin-2 is a leaky proton pump. Proc Natl Acad Sci USA 106:12317–12322.
OpenUrl Abstract/FREE Full Text

[97] Feldbauer K, et al.

[98] ↵
Keszthelyi L,
Ormos P
(2001) Electric signals associated with the photocycle of bacteriorhodopsin. FEBS Lett 109:189–193.
OpenUrl

[99] Keszthelyi L,

[100] Ormos P

[101] ↵
Sineshchekov OA,
Govorunova EG,
Wang J,
Li H,
Spudich JL
(2013) Intramolecular proton transfer in channelrhodopsins. Biophys J 104:807–817.
OpenUrl CrossRef PubMed

[102] Sineshchekov OA,

[103] Govorunova EG,

[104] Wang J,

[105] Li H,

[106] Spudich JL

[107] ↵
Bamann C,
Gueta R,
Kleinlogel S,
Nagel G,
Bamberg E
(2010) Structural guidance of the photocycle of channelrhodopsin-2 by an interhelical hydrogen bond. Biochemistry 49:267–278.
OpenUrl CrossRef PubMed

[108] Bamann C,

[109] Gueta R,

[110] Kleinlogel S,

[111] Nagel G,

[112] Bamberg E

[113] ↵
Stehfest K,
Hegemann P
(2010) Evolution of the channelrhodopsin photocycle model. ChemPhysChem 11:1120–1126.
OpenUrl CrossRef PubMed

[114] Stehfest K,

[115] Hegemann P

[116] ↵
Berndt A,
Prigge M,
Gradmann D,
Hegemann P
(2010) Two open states with progressive proton selectivities in the branched channelrhodopsin-2 photocycle. Biophys J 98:753–761.
OpenUrl CrossRef PubMed

[117] Berndt A,

[118] Prigge M,

[119] Gradmann D,

[120] Hegemann P

[121] ↵
Lin JY,
Lin MZ,
Steinbach P,
Tsien RY
(2009) Characterization of engineered channelrhodopsin variants with improved properties and kinetics. Biophys J 96:1803–1814.
OpenUrl CrossRef PubMed

[122] Lin JY,

[123] Lin MZ,

[124] Steinbach P,

[125] Tsien RY

[126] ↵
Sperling W,
Carl P,
Rafferty Ch,
Dencher NA
(1977) Photochemistry and dark equilibrium of retinal isomers and bacteriorhodopsin isomers. Biophys Struct Mech 3:79–94.
OpenUrl CrossRef PubMed

[127] Sperling W,

[128] Carl P,

[129] Rafferty Ch,

[130] Dencher NA

[131] ↵
Sineshchekov OA,
Spudich EN,
Trivedi VD,
Spudich JL
(2006) Role of the cytoplasmic domain in Anabaena sensory rhodopsin photocycling: Vectoriality of Schiff base deprotonation. Biophys J 91:4519–4527.
OpenUrl CrossRef PubMed

[132] Sineshchekov OA,

[133] Spudich EN,

[134] Trivedi VD,

[135] Spudich JL

[136] ↵
Bryl K,
Taui M,
Yoshizawa M,
Kobayashi T
(1992) Light adaptation of dark-adapted bacteriorhodopsin studied by nanosecond time-resolved absorption spectroscopy. Photochem Photobiol 56:1013–1018.
OpenUrl

[137] Bryl K,

[138] Taui M,

[139] Yoshizawa M,

[140] Kobayashi T

[141] ↵
Hofrichter J,
Henry ER,
Lozier RH
(1989) Photocycles of bacteriorhodopsin in light- and dark-adapted purple membrane studied by time-resolved absorption spectroscopy. Biophys J 56:693–706.
OpenUrl CrossRef PubMed

[142] Hofrichter J,

[143] Henry ER,

[144] Lozier RH

[145] ↵
Kawanabe A,
Furutani Y,
Jung K-H,
Kandori H
(2007) Photochromism of Anabaena sensory rhodopsin. J Am Chem Soc 129:8644–8649.
OpenUrl CrossRef PubMed

[146] Kawanabe A,

[147] Furutani Y,

[148] Jung K-H,

[149] Kandori H

[150] ↵
Vierock J,
Grimm C,
Nitzan N,
Hegemann P
(2017) Molecular determinants of proton selectivity and gating in the red-light activated channelrhodopsin Chrimson. Sci Rep 7:9928.
OpenUrl

[151] Vierock J,

[152] Grimm C,

[153] Nitzan N,

[154] Hegemann P

[155] ↵
Dudev T, et al.
(2015) Selectivity mechanism of the voltage-gated proton channel, H_V1. Sci Rep 5:10320.
OpenUrl CrossRef PubMed

[156] Dudev T, et al.

[157] ↵
Oda K, et al.
(2018) Crystal structure of the red light-activated channelrhodopsin Chrimson. Nat Commun 9:3949.
OpenUrl

[158] Oda K, et al.

[159] ↵
Saita M, et al.
(2018) Photoexcitation of the P₄⁴⁸⁰ state induces a secondary photocycle that potentially desensitizes channelrhodopsin-2. J Am Chem Soc 140:9899–9903.
OpenUrl

[160] Saita M, et al.

[161] ↵
Radu I, et al.
(2009) Conformational changes of channelrhodopsin-2. J Am Chem Soc 131:7313–7319.
OpenUrl CrossRef PubMed

[162] Radu I, et al.

[163] ↵
Grimm C,
Vierock J,
Hegemann P,
Wietek J
(2017) Whole-cell patch-clamp recordings for electrophysiological determination of ion selectivity in channelrhodopsins. J Vis Exp, e55497.

[164] Grimm C,

[165] Vierock J,

[166] Hegemann P,

[167] Wietek J

[168] ↵
Müller K-H,
Plesser T
(1991) Variance reduction by simultaneous multi-exponential analysis of data sets from different experiments. Eur Biophys J 19:231–240.
OpenUrl

[169] Müller K-H,

[170] Plesser T

[171] ↵
Hessling B,
Souvignier G,
Gerwert K
(1993) A model-independent approach to assigning bacteriorhodopsin’s intramolecular reactions to photocycle intermediates. Biophys J 65:1929–1941.
OpenUrl CrossRef PubMed

[172] Hessling B,

[173] Souvignier G,

[174] Gerwert K

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Unifying photocycle model for light adaptation and temporal evolution of cation conductance in channelrhodopsin-2

Significance

Abstract

Results

Single-Turnover Patch-Clamp Recordings Identify Three Conducting ChR2 States.

Single-Turnover Time-Resolved FTIR Measurements Reveal a Splitting of the Photocycle After Light Activation of Fully Dark-Adapted ChR2.

Light-Induced Splitting in 13-cis,C=N-anti and 13-cis,C=N-syn RSBH⁺ Conformations.

E90 Deprotonates upon 13-cis,C=N-syn Formation.

Excitation of P₄₈₀ Induces a Parallel Photocycle.

FTIR Amplitude Spectra Indicate Structural Differences of O₁ and O₂.

Isomerization of Retinal Leads to a Rearrangement of the Central Gate.

Deprotonation of E90 Is Essential for Proton Conductance of O₂ Following Light Adaptation.

Discussion

Materials and Methods

Yeast Culture.

Membrane Preparation and Protein Purification.

Reconstitution of ChR2 into DPPC or EggPC.

Preparation of HEK Cells.

Patch-Clamp Experiments in HEK293 Cells.

FTIR Experiments.

Raman Experiments.

MD Simulations.

Water Dynamic and Run-Average Structure.

Retinal Isomerization.

Acknowledgments

Footnotes

References

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Unifying photocycle model for light adaptation and temporal evolution of cation conductance in channelrhodopsin-2

Significance

Abstract

Results

Single-Turnover Patch-Clamp Recordings Identify Three Conducting ChR2 States.

Single-Turnover Time-Resolved FTIR Measurements Reveal a Splitting of the Photocycle After Light Activation of Fully Dark-Adapted ChR2.

Light-Induced Splitting in 13-cis,C=N-anti and 13-cis,C=N-syn RSBH+ Conformations.

E90 Deprotonates upon 13-cis,C=N-syn Formation.

Excitation of P480 Induces a Parallel Photocycle.

FTIR Amplitude Spectra Indicate Structural Differences of O1 and O2.

Isomerization of Retinal Leads to a Rearrangement of the Central Gate.

Deprotonation of E90 Is Essential for Proton Conductance of O2 Following Light Adaptation.

Discussion

Materials and Methods

Yeast Culture.

Membrane Preparation and Protein Purification.

Reconstitution of ChR2 into DPPC or EggPC.

Preparation of HEK Cells.

Patch-Clamp Experiments in HEK293 Cells.

FTIR Experiments.

Raman Experiments.

MD Simulations.

Water Dynamic and Run-Average Structure.

Retinal Isomerization.

Acknowledgments

Footnotes

References

Citation Manager Formats

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Article Classifications

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Information

Light-Induced Splitting in 13-cis,C=N-anti and 13-cis,C=N-syn RSBH⁺ Conformations.

Excitation of P₄₈₀ Induces a Parallel Photocycle.

FTIR Amplitude Spectra Indicate Structural Differences of O₁ and O₂.

Deprotonation of E90 Is Essential for Proton Conductance of O₂ Following Light Adaptation.