1. Structural Biology and Molecular Biophysics
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Time-resolved serial femtosecond crystallography reveals early structural changes in channelrhodopsin

  1. Kazumasa Oda
  2. Takashi Nomura
  3. Takanori Nakane
  4. Keitaro Yamashita
  5. Keiichi Inoue
  6. Shota Ito
  7. Johannes Vierock
  8. Kunio Hirata
  9. Andrés D Maturana
  10. Kota Katayama
  11. Tatsuya Ikuta
  12. Itsuki Ishigami
  13. Tamaki Izume
  14. Rie Umeda
  15. Ryuun Eguma
  16. Satomi Oishi
  17. Go Kasuya
  18. Takafumi Kato
  19. Tsukasa Kusakizako
  20. Wataru Shihoya
  21. Hiroto Shimada
  22. Tomoyuki Takatsuji
  23. Mizuki Takemoto
  24. Reiya Taniguchi
  25. Atsuhiro Tomita
  26. Ryoki Nakamura
  27. Masahiro Fukuda
  28. Hirotake Miyauchi
  29. Yongchan Lee
  30. Eriko Nango
  31. Rie Tanaka
  32. Tomoyuki Tanaka
  33. Michihiro Sugahara
  34. Tetsunari Kimura
  35. Tatsuro Shimamura
  36. Takaaki Fujiwara
  37. Yasuaki Yamanaka
  38. Shigeki Owada
  39. Yasumasa Joti
  40. Kensuke Tono
  41. Ryuichiro Ishitani
  42. Shigehiko Hayashi
  43. Hideki Kandori
  44. Peter Hegemann
  45. So Iwata
  46. Minoru Kubo  Is a corresponding author
  47. Tomohiro Nishizawa  Is a corresponding author
  48. Osamu Nureki  Is a corresponding author
  1. Department of Biological Sciences, Graduate School of Science, The University of Tokyo, Japan
  2. Graduate School of Life Science, University of Hyogo, Japan
  3. Graduate School of Engineering, Nagoya Institute of Technology, Japan
  4. Institute of Biology, Experimental Biophysics, Humboldt-Universität zu Berlin, Germany
  5. RIKEN SPring-8 Center, Japan
  6. Precursory Research for Embryonic Science and Technology (PRESTO), Japan Science and Technology Agency, Japan
  7. Department of Bioengineering Sciences, Graduate School of Bioagricultural Sciences, Nagoya University, Japan
  8. Department of Cell Biology, Graduate School of Medicine, Kyoto University, Japan
  9. Department of Chemistry, Graduate School of Science, Kobe University, Japan
  10. Japan Synchrotron Radiation Research Institute, Japan
  11. Department of Chemistry, Graduate School of Science, Kyoto University, Japan
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Cite this article as: eLife 2021;10:e62389 doi: 10.7554/eLife.62389

Abstract

Channelrhodopsins (ChRs) are microbial light-gated ion channels utilized in optogenetics to control neural activity with light . Light absorption causes retinal chromophore isomerization and subsequent protein conformational changes visualized as optically distinguished intermediates, coupled with channel opening and closing. However, the detailed molecular events underlying channel gating remain unknown. We performed time-resolved serial femtosecond crystallographic analyses of ChR by using an X-ray free electron laser, which revealed conformational changes following photoactivation. The isomerized retinal adopts a twisted conformation and shifts toward the putative internal proton donor residues, consequently inducing an outward shift of TM3, as well as a local deformation in TM7. These early conformational changes in the pore-forming helices should be the triggers that lead to opening of the ion conducting pore.

Introduction

Channelrhodopsins (ChRs) are light-gated cation channels first identified as sensory photoreceptors in green algae (Nagel et al., 2003). ChRs consist of seven transmembrane helices and a retinal chromophore covalently attached to a conserved lysine residue via a Schiff base linkage. ChRs allow fast ion flux across cell membranes upon illumination and thus are widely utilized as optogenetic tools to precisely control neural activity (Schneider et al., 2015; Ernst et al., 2014; Zhang et al., 2011). As in the well-studied bacterial proton pump bacteriorhodopsin (BR), light absorption triggers all-trans to 13-cis isomerization of the retinal, and induces the transitions of optically distinguishable intermediates that lead to channel opening and closing (photocycle process) (Kuhne et al., 2019; Bamann et al., 2010; Bamann et al., 2008; Figure 1). Spectroscopic studies have provided detailed characterizations of the respective intermediates in the photocycle of ChRs. In the best studied variant, Chlamydomonas reinhardtii ChR2 (CrChR2) (Bamann et al., 2008; Stehfest and Hegemann, 2010; Lórenz-Fonfría et al., 2013), retinal isomerization leads to an intermediate with a red-shiftedabsorption peak, P1520 (K or L in BR), and sequential deprotonation and reprotonation of the Schiff base generate P2390 and P3520 (M and N in BR), respectively (Figure 1). Photocycle transitions are determined in most ChRs by an aspartate–cysteine pair in the retinal binding pocket (DC-pair) (Berndt et al., 2009; Yizhar et al., 2011; Figure 1—figure supplement 1), but the mechanism by which ion pore formation is associated with these residues is still under debate. Recent studies have suggested that the ion conducting pore is formed during the late P2390 intermediate Kuhne et al., 2019, involving the rearrangement of the pore-forming TM helices (Müller et al., 2011), and a two-dimensional crystal study indeed revealed low resolution projection maps of the light-induced helix movement in ChR (Müller et al., 2011; Müller et al., 2015). A detailed mechanism for how retinal isomerization causes conformational changes and how these changes induce ion pore opening has remained elusive.

Figure 1 with 1 supplement see all
Photocycle model of channelrhodopsin.

Schematic model of the C1C2 photocycle. Superscript of each reaction intermediate (P1(520), P2(390), P3(520), P4(480)) indicates the wavelength of maximum absorption. Open/close states of the channel pore and the proposed model of the retinal and protonation of Schiff base are indicated. The time range in which the P1–P3 intermediates were observed in the spectroscopic studies in solution is shown.

The recent development of the X-ray free electron laser (XFEL) technology has enabled high-resolution visualization of the time-resolved molecular conformational changes in crystals, thus profoundly advancing our understanding of biological reactions (Tosha et al., 2017; Nango et al., 2016; Suga et al., 2017; Shimada et al., 2017). In this study, we performed the time-resolved serial femtosecond crystallography (TR-SFX) analysis of ChR, which revealed critical conformational changes occurring early in the photocycle that probably induce the cation permeation in the later stages.

Results

Time-resolved spectroscopy of C1C2

We used a chimeric construct between Chlamydomonas reinhardtii ChR1 (CrChR1) and ChR2 (CrChR2) for the TR-SFX experiments (hereafter referred to as C1C2) (Kato et al., 2012; Figure 1—figure supplement 1), which has normal ChR1 photocycle properties (Hontani et al., 2017; Inaguma et al., 2015; Figure 2a,b). C1C2 yielded high-density microcrystals that diffracted XFEL to about 2.3 Å at SACLA (Figure 3a,b). The dark state structure of C1C2 determined at SACLA is almost the same as the previous structure solved by the synchrotron analysis, including the chromophore retinal, the interhelical interactions of the inner and central gates, and the extracellular water access channel in the dark state (Figure 3c,d, and Figure 3—figure supplement 1; Hontani et al., 2017). The C1C2 photocycle was reported in detail recently (Hontani et al., 2017), revealing different accumulations of the P1 and P2 photo-intermediates, as compared to CrChR2. Preceding the TR-SFX experiment, we investigated the photocycle and ion pore formation processes of C1C2. In the flash-photolysis experiments, C1C2 showed similar photo-intermediates as observed in the previous study (Hontani et al., 2017), in detergent micelles or reconstituted lipid membranes (Figure 2a,b and Figure 2—figure supplement 1). The results indicate a rapid rise of 520 nm absorption after photo-excitation, occurring before 10 μs, which corresponds to the formation of the P1 intermediate with an isomerized retinal (Hontani et al., 2017; Nogly et al., 2018), and the subsequent increases at 390 nm and 520 nm show formation of the P2 and P3 intermediates with a deprotonated and reprotonated retinal Schiff base, respectively (Figure 2a). The absorption spectra of P1 and P3 resemble each other and cannot be distinguished in C1C2, but the delayed rise of the 530 nm absorption after 360 μs shows the formation of the P3 intermediate at the later time point. To correlate the different photocycle intermediates to formation of the ion conducting pore, we next measured the C1C2 photocurrents evoked by 7-ns laser flash (Figure 2c and Figure 2—figure supplement 2). The C1C2 photocurrents are initiated by a fast, inward directed charge transfer that likely coincides with the retinal deprotonation (P2 formation), which has not been fully time-resolved in the whole cell patch clamp recordings. Afterwards, passive photocurrents rise with a time constant of 70–140 µs (rather faster at acidic pH) and change direction depending on the membrane voltage. The photocurrent remains after reprotonation of the retinal Schiff base and declines in a biexponential manner, with two voltage dependent time constants of 8–40 ms and 60–130 ms and a kinetic amplitude ratio that depends on the pH, correlating neither precisley with the P2nor the P3 intermediate. The absorption spectrum of ChR depends mostly on the microenvironment of the chromophore retinal, especially on the Schiff base environment (Ritter et al., 2008), and does not need to reflect changes in the ion pore structure . As recently shown for CrChR2 (Kuhne et al., 2019), also C1C2 channel opening takes place only after deprotonation of the retinal Schiff base, during further confirmational changes of the P2 intermediate.

Figure 2 with 3 supplements see all
Flash photolysis and flash photo activation measurement of C1C2.

(a) Transient absorption spectra of C1C2 reconstituted in POPE/POPG (protein/lipid molar ratio = 1/50), 150 mM NaCl, 50 mM Tris-HCl (pH 8.0), 5% glycerol, and 0.01% cholesteryl hemisuccinate (CHS). (b) Time trace of absorption changes of C1C2 reconstituted in POPE/POPG, at 445 (green), 375 (blue), and 520 (red) nm probe wavelengths, which roughly estimate C1C2 (ground state), P2, and P3 formation, respectively. (c) Normalized, log-binned and averaged photocurrents of the C1C2 protein (mean ± SEM, n = 3 - 5). The black arrow indicates the inward directed current caused by retinal Schiff base deprotonation. (d) Transient difference absorption spectra recorded from the multiple C1C2 crystals. Each time-resolved spectrum was averaged over 21 crystal data sets. The time evolution is indicated from red to purple.

Figure 3 with 3 supplements see all
Microcrystals for serial femtosecond crystallography (SFX) experiment and C1C2 SFX structure.

(a) Lipidic cubic phase (LCP) crystals of C1C2 optimized for the time-resolved SFX (TR-SFX) experiments. The orange scale bar on the lower right indicates 50 μm, with 5 μm sub-scaling lines. The size of the crystals ranged from 2 to 5 μm. (b) A diffraction image from a C1C2 crystal, obtained with a single SACLA-XFEL pulse. (c) The structure of dark state C1C2 determined by serial femtosecond crystallography. A water accessible cavity is illustrated, with the putative ion pathway indicated by an arrow. The five glutamic acid residues lining the ion pore (E1–5) and the two counterion residues (Ci1 and Ci2) are indicated by sticks, and the three constriction sites, the inner, central, and outer gates, are enclosed in boxes. (d) Comparisons of the constriction sites of the TR-SFX structure (left panels) and the synchrotron structure (right panels; PDB code 3UG9) of C1C2, for the inner (upper panels), central (middle panels), and outer (lower panels) gates. The constituent residues are shown as sticks, and the TM helix number is indicated on each helix.

To determine the time points for the TR-SFX study, we next investigated the photocycle of C1C2 in the lipidic cubic phase (LCP) crystals. The flash illumination induced a rapid elevation at 530 nm, confirming retinal isomerization (P1 formation) (Figure 2d and Figure 2—figure supplement 3a, c). The elevation of the 390 nm absorption, which is most prominent in 2.1–21 ms, showed the P2 formation, but the subsequent increase at the 530 nm absorption was not observed in the crystal (Figure 2d and Figure 2—figure supplement 1 and Figure 2—figure supplement 3a,c), suggesting that the transition from P2 to P3 is hindered in the crystal, although the C1C2 crystal has only minimal interactions at the extracellular membrane region (Figure 3—figure supplement 2). Nonetheless, the accumulation of the P2 intermediate is more prominent in the crystals, as the spectral increase at 390 nm is relatively higher, as compared to the similar increase detected in micelles and reconstituted membranes (Figure 2a,d and Figure 2—figure supplement 1, Figure 2—figure supplement 3b,d). In addition, the flash photo-activation experiment using HEK cells showed that the photocurrent rose in between 100–1000 μs, regardless of pH, which corresponds to a second late P2 intermediate (Figure 2c and Figure 2—figure supplement 2), in a good agreement with the previous studies in ChR2 (Kuhne et al., 2015; Kuhne et al., 2019). Therefore, we selected the time points corresponding to the P1 to P2 transition in the crystals for the TR-SFX study, to understand the detailed mechanism of the light-dependent channel opening in ChR.

Structural change in ChR

The LCP matrix containing C1C2 microcrystals was introduced to the XFEL chamber in a continuous flow. Since the crystallized sample was swollen and almost in the liquid sponge phase, we added paraffin and solid LCP matrix to enable stable and constant sample flow to the FEL beam (detailed in Materials and methods). The diffraction images were collected at 30 Hz, with the 470 nm pump-pulse laser at 15 Hz and the delayed time points at 1 μs, 50 μs, 250 μs, 1 ms, and 4 ms (Figure 3—figure supplement 3a,c and Supplementry file 1). The absence of light contamination was checked by the second dark data set, obtained with a 10 Hz pump laser pulse (Figure 3—figure supplement 3c,d). The difference-Fourier maps were calculated at 2.5 Å for each delayed time point, with the dark state SFX structure as the reference (Figure 4), and they clearly revealed the structural changes in the protein core region of C1C2 (Figure 4—figure supplement 1; Figure 4—figure supplement 2 and Video 1). Due to the low quantum efficiency of the retinal isomerization, which is estimated as only about 30% in C1C2 (Hontani et al., 2017), we calculated the extrapolated structure factor amplitudes and the models were refined against the extrapolated maps (detailed in Materials and methods and Figure 4—figure supplement 3). We also calculated the Fclight–Fcdark maps using the models, which shows a similar appearance with the difference-Fourier map and thus confirms the conformational change (Figure 4—figure supplement 4).

Figure 4 with 5 supplements see all
Difference Fourier electron density map and structural changes around TM7 and TM3.

Views of the |Fobs|light− |Fobs|dark difference Fourier electron density maps and the structural changes around TM7 (a–e) and TM3 (f–j) for 1 μs (a and f), 50 μs (b and g), 250 μs (c and h), 1 ms (d and i), and 4 ms (e and j). Green and purple meshes indicate positive and negative difference electron densities, respectively (contoured at ±3.1 to 3.3σ, see also Figure 4—figure supplement 1). The difference Fourier maps were calculated by using the phases from the coordinates of the dark-adapted C1C2 structure (gray). Paired negative and positive difference electron densities indicate the downward shift of TM7 and the outward shift of TM3. The structural model (blue) was refined against the extrapolated data of each time point and the model was superimposed upon the initial-state C1C2 model (gray). Movements of the residues and the TM helices are indicated by arrows. Probably due to the variations in the crystal quality (Table 1), the difference density in the 250 μs time-delay seems rather weak.

Video 1
Overview of the |Fobs|light− |Fobs|dark difference.

Fourier electron density maps for five time points and structural changes. Views of the |Fobs|light− |Fobs|dark difference Fourier electron density maps and the structural changes. Green and purple meshes indicate positive and negative difference electron densities, respectively (contoured at ±3.1 to 3.3σ, see also Figure 4—figure supplement 1). The difference Fourier maps were calculated by using the phases from the coordinates of the dark-adapted C1C2 structure (gray). Paired negative and positive difference electron densities indicate the downward shift of TM7 and the outward shift of TM3. The structural model (blue) was refined against the extrapolated data of each time point and the model was superimposed upon the initial-state C1C2 model (gray). Movements of the residues and the TM helices are indicated by arrows. Probably due to the variations in the crystal quality (Supplementary file 1), the difference density in the 250 μs time-delay seems rather weak.

Since the retinal isomerization reaction is generally completed within a few picoseconds (Hontani et al., 2017; Neumann-Verhoefen et al., 2013; Bühl et al., 2018), the observed changes correspond to the events following the retinal isomerization. The isomerization of the retinal induces conformational changes of the retinal-linked residue, Lys296, and TM7 (Figure 4, left panels). The positive densities along Lys296 suggest its downward shift (toward the extracellular side), which is detected from 1 μs and becomes most prominent at 4 ms (Figure 4, left panels). In accordance with this shift, positive and negative difference densities are observed around the proximal residues in TM7, such as Trp299, indicating an overall downward shift of the middle portion of TM7. The similar downward shift of TM7 (Helix G) was the most distinct feature in the L-like intermediate of BR, observed in both the cold-trapped synchrotron structure (Neutze et al., 2002) and the recent TR-SFX study (Nango et al., 2016). In the TR-SFX experiment of BR, the primary retinal isomerization reaction induced local distortion around the C13–C14 double bond of the retinal, which was subsequently propagated as conformational changes to other protein regions, also involving a downward shift of TM7. Therefore, the retinal-attached residue in C1C2 probably undergoes similar conformational changes, although the primary retinal isomerization was not visualized, due to the limited resolution of the current TR-SFX experiment. Notably, at 50 μs we observed positive and negative difference densities at the putative internal proton acceptor residue, Asp292 (Figure 4, left panels), indicating a slight downward shift of this residue, which might be associated with the proton accepting reaction.

Retinal kink-induced outward shift of TM3

The most prominent change occurs in TM3 (Figure 4, right panels). A strong negative density appears at Cys167, accompanied by a complementary positive density at the distal site from the retinal, suggesting an outward shift of Cys167. In addition, a strong positive density appears near Pro168 on the outside of the TM3 segment, which indicates an outward shift of the cytoplasmic segment of TM3, originating at the helix kink at the Pro168 residue. Along with these changes, negative and positive densities appear at the C13 methyl group and on the lateral side of the polyene chain of the retinal, respectively. This kink and lateral shift of retinal allows lateral movement of TM6 as well as TM3 (Figure 4, right panels). The current difference Fourier map is in excellent agreement with the recent MD simulation combined with QM/MM calculations (Cheng et al., 2018), in which the retinal adopts a kinked conformation toward TM3 upon the Schiff base deprotonation, probably due to the destabilized resonance structure of the polyene chain (Figure 5), although the QM/MM simulation focuses only on the molecular events after the Schiff base deprotonation and thus does not exactly match the experimental time scale. Together, the sequential changes in the difference-Fourier maps suggest that the retinal kink occurs, followed by the consequent outward shift of TM3. Consistently, the positive/negative peaks at the retinal appear slightly earlier, whereas the change in TM3 becomes slower and reaches the maximum at the later time points of 1 ms and 4 ms (Figure 5).

Conformational change of retinal and comparison with QM/MM model.

(a) Intracellular view of the |Fobs|light− |Fobs|dark difference Fourier electron density map and the structural changes around the retinal. Green and purple meshes indicate positive and negative difference electron densities, respectively. The structural model (blue) was refined against the extrapolated data of 1 ms and the model was superimposed upon the initial-state C1C2 model (gray). Movements of residues or TM helices are indicated by arrows. (b) The retinal pocket of dark-adapted C1C2 (gray), the model at the 1 ms delayed-time point (blue), and the model at 400 ns of the QM/MM simulation (Cheng et al., 2018) (red) are superimposed together.

The current TR-SFX experiment revealed two critical conformational changes occurring early in the photocycle process: the outward shift of TM3 and the downward shift of TM7, both of which constitute gate constriction sites along the ion permeation pathway in the dark state (Figure 3). TM7 contributes essentially to the ‘central gate’, in which Asn297 forms hydrogen bond interactions with Ser102 (TM1) and Glu129 (TM2). Therefore, the TM7 shift, as well as the outward shift of Asp292 upon proton acceptance, should lead to rearrangements of the central gate residues. The intracellular end of TM3 shapes the ‘inner-gate’, in which His173 forms ionic interactions with Arg307 in TM7 and Glu121 and Glu122 in TM2. The recent MD simulation showed a similar retinal kink and subsequent water influx from the intracellular side (Figure 5; Cheng et al., 2018). Therefore, the conformational change in the middle of TM3 might propagate to the intracellular segment and induce the opening of the inner gate.

Cys167 and Asp195 have been identified as important residues for channel gating and are thus termed DC-gate or DC-pair, which is conserved among most ChRs. FTIR and structural studies have suggested that Asp195 functions as the internal proton donor for the deprotonated Schiff base, via an internal hydrated water molecule and Cys167 (Nack et al., 2010; Bamann et al., 2010; Lórenz-Fonfría et al., 2013). A previous FTIR study showed an increase of the S-H stretching frequency of the Cys167 side chain upon illumination (Ito et al., 2014). The DC-pair residues are critical for the fast channel kinetics of ChRs, as mutations of any of them drastically slowed the open/close kinetics of the channel, resulting in step function opsin (SFO) mutants, with long-lasting photocurrents that can be switched on and off by different wavelengths of light (Berndt et al., 2009; Yizhar et al., 2011). Previous 2D crystallographic studies of the CrChR2 C128T mutant (corresponding to Cys167 in C1C2 [Figure 1—figure supplement 1]) suggested rearrangements of TM2, 6, and 7 in the open channel conformation, that was here induced by the stationary light (Müller et al., 2011; Müller et al., 2015). These conformational changes are partly consistent with the current TR-SFX analysis. However, the TM3 shift is only observed in the current study, indicating the critical role of the Cys residue in the TM3 shift. The movement of TM7, on the other hand, is similarly observed in both experiments, although to different degrees. We therefore propose that the conformational changes in TM7 and TM3 are the two important triggers that cooperatively induce channel opening by eliciting sequential conformational changes in the surrounding helices, such as TM2 and TM6 (Figure 6), as observed in the ‘open’ conformation of the SFO mutant of CrChRs (Müller et al., 2015). This model is also in good agreement with the results of the vibrational spectroscopic study showing a two-phasic helix hydration occurring during the early and late stages of the P2 intermediate, in which only the latter leads to the fully-open channel pore (Lórenz-Fonfría et al., 2015). Furthermore, site-directed labeling with infrared sensitive, unnatural amino acids subsequently showed more specificly, that espacially hydration changes in the intracellular gate are correlated with channel opening (Krause et al., 2019). In summary, our TR-SFX experiment has revealed two critical conformational changes in TM7 and TM3 that occur during the early part of the photocycle, and these triggers would induce water and proton influx and subsequent pore dilation, although only slight structural perturbations in the gate constrictions were observed in the current study, as they were probably affected by the crystal packing interactions (Figure 4—figure supplement 5).

Schematic model of the C1C2 channel opening.

The inner gate formed by Glu121, Glu122, His173, and Arg307 prevents water influx in the initial dark-adapted state. After the retinal isomerization reaction, the retinal twists and shifts toward Cys167 and induces an outward shift of TM3, originating at the kink introduced by Pro168. Isomerization of the retinal also induces a downward shift of the middle portion of TM7, especially at the retinal-attached residue, Lys296 (transition between the P1 and P2 intermediates). These movements are likely to induce water influx, by destabilizing the intracellular constriction, and further induce rearrangement of the surrounding helices (Open model).

Discussion

Channel pore gating in ChR

The current results highlight the diversity and similarity of rhodopsin proteins. TM7 is covalently attached to the retinal and thus undergoes a similar shift in other rhodopsin proteins, such as the proton pump BR (Nango et al., 2016), and the Na+ pump KR2 (Skopintsev et al., 2020), albeit with some variations. Therefore, this structural movement is probably conserved in rhodopsin proteins. Most notably, the retinal kink and TM3 shift were also observed in KR2, even though it is evolutionally and functionally distant from ChRs, while such a conformational change was not observed in BR. In BR, the retinal is transiently kinked toward the opposite direction, but only during the very early time after photoexcitation (around 10 ps to 16 ns), and causes totally different changes in the protein part. Therefore, the retinal kink and subsequent conformational changes are likely to be affected by the residues constituting the retinal pocket. Indeed, the outward shift of TM3 was not observed in the Cys167 mutant of C1C2, as elucidated by the light-activated projection maps on the 2D crystallographic study (Müller et al., 2015). A recent MD simulation demonstrated the hydrogen bonding interactions between the deprotonated Schiff base and the hydroxyl group of Thr166, located beside the Cys167 residue (Cheng et al., 2018), which are likely to affect the retinal conformation. The same threonine is highly conserved among microbial rhodopsins, and in CrChR2 it was also proposed to couple the protonation changes in the counterion region to the hydrogen bonding changes in the DC-pair and the conformational changes of TM3 (Watanabe et al., 2012). Together, the hydrogen-bonding network beside the retinal, involving the DC-pair, may affect the retinal conformation after isomerization, in good agreement with the experimental evidence that the residues lining the retinal pocket affect channel kinetics (Oda et al., 2018). These results would pave the way for the development of novel optogenetic tools with different kinetics.

Materials and methods

Laser flash photolysis experiment

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The time-evolution of the transient absorption changes of photo-excited C1C2 was measured as previously described (Inoue et al., 2013; Inoue et al., 2016). The purified protein sample was solubilized in buffer containing 150 mM NaCl, 50 mM Tris-HCl (pH 8.0), 0.05% n-dodecyl-β-D-maltoside (DDM), 5% glycerol, and 0.01% cholesteryl hemisuccinate (CHS). For the measurement of C1C2 in liposomes, the purified C1C2 protein was mixed with a lipid mixture of POPE and POPG (molar ratio = 3: 1) at a protein-to-lipid molar ratio of 1: 50 (the final concentrations of C1C2 and lipids were adjusted to 5 μM and 250 μM, respectively), and the DDM was removed by BioBeads (SM-2, Bio-Rad). The sample solution was placed in a quartz cuvette with a 1 cm optical path length and illuminated with a 480 nm beam generated by an OPO system, pumped by the third harmonics generation of a nanosecond Nd3+: YAG laser (LT-2214, LOTIS TII, Minsk, Republic of Belarus). The intensities of the transmitted probe light from a Xe arc lamp (L8004, Hamamatsu Photonics, Japan) were measured before and after laser excitation with an ICCD linear array detector (C8808-01, Hamamatsu, Japan) at 24°C, and transient absorption spectra were obtained by calculating the ratios between them. Ninety spectra were averaged to improve the signal-to-noise (S/N) ratio. To obtain the detailed time evolution of the transient absorption change after photoexcitation, the change in the intensity of the monochromated output of a Xe arc lamp (L9289-01, Hamamatsu Photonics, Japan), passed through the sample at 24°C, was observed by a photomultiplier tube (R10699, Hamamatsu Photonics, Japan) at 375, 445, and 520 nm. The signal from the photomultiplier tube was averaged to improve the S/N ratio and stored in a digital-storage-oscilloscope (DPO7104, Tektronix, Japan). The obtained kinetics data were fitted with a multi-exponential function.

Dark adaptation kinetics of C1C2 was measured for the 0.05% DDM-solubilized sample at 24°C, as previously described (Inoue et al., 2016). The dark-adapted sample was illuminated for 1 min by using the output from a 1-kW tungsten–halogen projector lamp (Master HILUX-HR, Rikagaku, Japan), through an interference filter at 460 nm. After turning off the light, the spectra or the absorption at a specific wavelength were measured by a UV-visible spectrometer (V-730, JASCO).

Flash photo-activation current measurement

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HEK293 cells were cultured in Dulbecco’s Modified Medium (Biochrom), supplemented with 10% (v/v) fetal bovine serum (FBS Superior; Biochrom), 1 μM all-trans retinal, and 100 µg/ml penicillin/streptomycin (Biochrom, Berlin, Germany). Cells were seeded on poly-lysine coated glass coverslips at a concentration of 1 × 105 cells/ml, and transiently transfected using the FuGENE HD Transfection Reagent (Promega) 28–48 hr before measurement. C1C2 was fused to an mCherry-fluorophor and expressed under the control of the CMV-promotor.

Patch pipettes were prepared from borosilicate glass capillaries (G150F-3; Warner Instruments) using a P-1000 micropipette puller (Sutter Instruments), and fire polished with a final pipette resistance of 1.5–3.0 MΩ. Fluorescent cells were identified using an Axiovert 100 inverted microscope (Carl Zeiss). Single ns laser pulses were generated by an Opolette Nd:YAG laser/OPO system (OPOTEK), selected by an LS6ZM2 shutter system (Vincent Associates), and delivered to the cells through the optics of the microscope and a 90/10 beamsplitter (Chroma). The laser intensity was set to 5% using the built-in motorized variable attenuator, resulting in a pulse energy of 100 ± 20 µJ/mm2. Photocurrents were recorded with an AxoPatch 200B amplifier (Molecular Devices) at a maximal bandwidth of 100 kHz and digitized using a DigiData 1440A digitizer (Molecular Devices) at a sampling rate of 250 kHz. Membrane resistance was typically >1 GΩ, while access resistance was below 10 MΩ. Pipette capacity, series resistance, and cell capacity compensations were applied. A 140 mM NaCl agar bridge was employed for the reference bath electrode. Intracellular pipette and extracellular bath solutions contained 110 mM NaCl, 1 mM KCl, 1 mM CsCl, 2 mM CaCl2, and 2 mM MgCl2, and were buffered to pH 6.0, 7.2, or 8.0 by 10 mM MES, HEPES, or TRIS (with glucose added to 290 mOsm for intracellular and 310 mOsm for extracellular solutions). Measurements were controlled by the pCLAMP software (Molecular Devices). Post measurement photocurrent recordings were first baseline corrected and time shifted to align with the rising edges of the Q-switch signals of the laser pulse, using the Clampfit 10.4 software (Molecular Devices). They were then binned to 50 logarithmically spaced data points per temporal decade, normalized to peak photocurrents at −60 mV, and averaged for up to 10 individual repetitions for each cell and voltage condition with a custom-written Matlab script (MathWorks). Kinetic analysis was performed in Origin 9.1 (OriginLab). Passive photocurrents were calculated by subtracting the photocurrents at 0 mV, which were recorded in the absence of an electrochemical gradient and represent the inner protein charge transfer reactions. For sufficient statistical significance, each measurement was repeated multiple times on different biological replicates, in at least three independent experiments. The exact number of biological replicates for each measurement is provided in the figure legend.

Time-resolved visible absorption spectroscopy

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The photocycle reaction was induced by a 6-ns, 470 nm pump pulse from an optical parametric oscillator (OPO) (NT230, EKSPLA), and the spectral changes during the reaction were measured by using a fiber-coupled spectrometer (Flame-S, Ocean Optics), with a microsecond white-light probe pulse from a Xe-flash lamp (L11316-11-11, Hamamatsu Photonics). A portion of the probe light (5%) was used to correct the pulse-to-pulse fluctuation of the probe light intensity. The pump pulse and the probe light were focused on the sample point, with beam diameters of 300 μm and 40 μm, respectively. The pump energy was reduced to 15 μJ (0.21 nJ/μm [Schneider et al., 2015]), to avoid sample damage by the repetitive pump illuminations during the data accumulation. The pump pulse, the probe light source, and the spectrometer were synchronized by pulse generators (DG645, Stanford Research Systems), and the pump-probe delay time was adjusted with a timing jitter of ±20 ns. The repetition rates of the pump and the probe were 0.5 Hz and 1 Hz, respectively. In the solution measurement, the sample was packed in a spinning quartz cell with an optical path length of 1 mm and spun at 50 rpm during the measurement. The sample concentration was ~10 mg/ml, dissolved in 50 mM Tris-HCl, pH 8.0, containing 150 mM NaCl and 5% glycerol. In the microcrystal measurement, we used the C1C2 crystals obtained under similar conditions as in the TR-SFX (below). Specifically, a pillar of 30 μl protein-laden LCP, approximately 2 cm in length, was extruded from the syringe and soaked in 900 μl of the crystallization buffer, containing 100 mM HEPES, pH 7.0, 280 mM KH2PO4, and 30% PEG500DM, and incubated for 1 week in the dark at 20°C, which yielded crystals in a liquid sponge phase floating on the buffer solution. The sponge phase layers were transferred by pipetting and packed in two quartz windows with a 50 μm-thick spacer. Only a few absorption signals from the non-crystallized area were observed, and we used the transmitted light intensity from the non-crystallized area as the reference to calculate the absorption spectra of C1C2. The sample temperature was kept at 20°C for both the solution and microcrystal measurements. To decrease the noise level of the time-resolved spectra, singular value decomposition (SVD) (Shrager and Hendler, 1998) was used with Igor Pro (Wave Matrix). In the SVD, matrix A comprising the intensity changes at each wavelength and time (column and row, respectively) was decomposed according to A = USVT, where S represents the diagonal matrix with the singular values in decreasing order. This allows an evaluation of the number of significant orthonormal basis vectors for the wavelength (columns of U) and the time (columns of V). By reconstructing the matrix A’ from only the most significant components, random noise was eliminated from the raw spectra. The time-resolved spectra for the microcrystal and solution samples were well reconstructed by two and three components, respectively. Based on the SVD, global fitting was then performed for the time-resolved reconstructed spectra with Igor Pro (Wave Matrix). In the global fitting, the spectra were fitted with a sum of exponentials with apparent time constants τi as global parameters, where i is the number of components used for the spectral reconstruction in the SVD analysis.

Crystallization of C1C2

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The vector construction and protein production and purification were performed according to the previous report (Kato et al., 2012). C1C2 was finally purified by size-exclusion chromatography, in 150 mM NaCl, 50 mM Tris-HCl, pH 8.0, 5% glycerol, 0.05% DDM, and 0.01% CHS, and concentrated to ~15 mg/ml for crystallization. C1C2 was mixed with monoolein (Sigma) in a 2:3 protein to lipid ratio (w/w), using a 100 μl volume Gastight syringe (Hamilton). A pillar of 30 μl protein-laden LCP, approximately 2 cm in length, was extruded from the syringe and soaked in 900 μl of the crystallization buffer, containing 100 mM MES, pH 6.9, 100 mM Na formate, and 30% PEG500DM, and incubated for 2–3 weeks in the dark at 20°C. Crystals were observed with a Hirox KH8700 digital microscope.

Sample preparation for LCP-SFX

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100 μl of LCP pillars with grown C1C2 microcrystals were collected and loaded into a Hamilton syringe and mixed with an equivalent volume of buffer-containing LCP (2:3 ratio of the buffer containing 150 mM NaCl and 50 mM Tris-HCl, pH 8.0, to monoolein). A 10 μl aliquot of liquid paraffin (Wako) was further added to the LCP material and homogenously mixed to enable smooth flow in the SFX experiment. All of these procedures were performed in the dark under red LED lights (over 600 nm wavelength).

Diffraction experiment at SACLA-FEL

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Single-shot XFEL data collection was performed, using femtosecond X-ray pulses from the SACLA at BL3. The pulse parameters of SACLA were as follows: pulse duration, <10 fs; X-ray energy, 7 keV; energy bandwidth, 33.5013 eV (FWHM); pulse flux, 2.0 × 1011 photon/pulse photons per pulse; beam size 1.5 μm (H) × 1.5 μm (W); repetition rate, 30 Hz. The C1C2 crystals were mixed with protein-less LCP and paraffin oil, loaded into an injector with a 75 μm inner diameter nozzle, and set in a diffraction chamber filled with helium gas, in a setup called Diverse Application Platform for Hard X-ray Diffraction in SACLA (DAPHNIS). The flow rate was set to 2.5 μl/min (9.4 mm/s). The diffraction patterns were recorded on a multiport CCD detector with an eight sensor module (Kameshima et al., 2014). The excitation laser pulses were provided at 15 Hz, and the XFEL pulses had a repetition rate of 30 Hz. Each ‘pump-on’ image was followed by ‘pump-off’ images, which were recorded separately. The pump-on images were used to analyze the structural differences, whereas the diffraction data for the dark-state were collected in a separate run to avoid light contamination.

Pump laser setup

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To advance the C1C2 samples to the excited state, a 6 ns visible light pump pulse with a wavelength of 470 nm was provided to the samples at Δt = 1 μs, 50 μs, 250 μs, 1 ms, and 4 ms, from an OPO (NT230, EKSPLA), which was Q-switched at 15 Hz using a pulse generator (DG645, Stanford Research Systems). The pulse generator was also used to synchronize the pump pulse with the XFEL pulse and control the delay time (Δt) with a timing jitter of <1 ns. Samples were thus irradiated from both sides of the LCP microjet, a geometry that was chosen to optimize the crystal excitation efficiency. The pump focal size was set to 40 μm (FWHM) and the pump energy was 20 μJ (10 μJ from each direction) at the sample point. The pump beam center was aligned 6 μm upstream from the XFEL spot. The pump light was scattered, as the LCP microjet is semi-transparent. To avoid the interference from the pump scattering, the sample flow rate (9.4 mm/s) was set to be the same as that in the TR-SFX analysis of BR, in which each pump-illuminated crystal for the pump-on XFEL images was separated by 630 µm. A portion of the pump beam (5%) was isolated with a beam sampler, and its photodiode-detected signal was used to monitor the laser status and to tag the diffraction image with ‘light-on’. Since the repetition rate of the pump laser was half of that of the XFEL, the ‘light-on’ data and the ‘dark’ data were collected alternately.

Data processing and structural analysis

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Data collection was monitored by a real-time data processing pipeline (Nakane et al., 2016), based on Cheetah (Barty et al., 2014). Diffraction images with more than 20 spots were considered as hits and processed in CrystFEL, version 0.6.3 (White et al., 2012), with the parameters of hitfinderAlgorithm = 6, hitfinderADC = 40, hitfinderMinSNR = 5. Diffraction spots for indexing were detected by the built-in spot finding algorithms ‘zaef’ (Zaefferer, 2000) or ‘peakfinder8’ (Barty et al., 2014) in CrystFEL, and indexing was performed with DirAx (Duisenberg, 1992). Intensities were merged by Monte Carlo integration with the process_hkl command in the CrystFEL suite, with the per-image resolution cutoff (--push-res 1.2). Data collection statistics are summarized in Supplementary file 1. As the cell parameters do not vary among the different delayed-time points, the difference Fourier electron density maps (|Fobs|light–|Fobs|dark)·exp[iΦcalc] were calculated by FFT (Ten Eyck, 1973) in the CCP4 suite (Winn et al., 2011), with phases calculated from the refined dark state model.

Structural refinement

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The structure was solved by the molecular replacement method implemented in Phenix Phaser-MR, using the model of C1C2 (PDB: 3UG9). The structure was manually modified to fit into the electron density maps, using the program Coot (Emsley et al., 2010), and then refined with the programs Phenix (Adams et al., 2010) and Refmac5 in the CCP4 suite (Vagin et al., 2004). The model structures of the respective delayed time points were generated by real-space refinement against the difference-Fourier maps by COOT, with the restraints of protein geometry and retinal planarity between C5and C13. Extrapolated data were calculated by a linear approximation, as follows: FExtra = (Folight − Fodark)/R + Fodark, where R represents the activation ratio of molecules and FExtra represents the extrapolated structure factor amplitude. Folight was scaled to Fodark by using the scaleit program in the CCP4 suite (Howell and Smith, 1992) before calculating FExtra. The activation ratio R (the ratio of molecules in the excited state after light illumination) was determined by calculating extrapolated maps with phases of the dark state at different activation ratios in steps of 0.01, until a feature of the dark state at Cys167 disappeared. Next, 4 ms model was manually adjusted to fit the extrapolated map and refined by Refmac5 against FExtra and its estimated error. Then, each time point model was refined by Refmac5 with external structure restraints using 4 ms structure by the Geman–McClure function (Murshudov et al., 2011) the refinement statistics are. Figures were prepared with Cuemol (http://www.cuemol.org).

Data availability

Coordinates and structure factors for the TR-SFX structures have been deposited in the PDB with the accession codes 7C86 (dark), 7E6Y (1 μs), 7E6Z (50 μs), 7E70 (250 μs), 7E71 (1 ms), and 7E6X(4 ms). Raw diffraction images have been deposited in the Coherent X-ray Imaging Data Bank (accession ID 150, https://doi.org/10.11577/1768368).

The following data sets were generated

References

    1. Emsley P
    2. Lohkamp B
    3. Scott WG
    4. Cowtan K
    (2010) Features and development of coot
    Acta Crystallographica. Section D, Biological Crystallography 66:486–501.
    https://doi.org/10.1107/S0907444910007493

Decision letter

  1. Andrew C Kruse
    Reviewing Editor; Harvard Medical School, United States
  2. Kenton J Swartz
    Senior Editor; National Institute of Neurological Disorders and Stroke, National Institutes of Health, United States
  3. Doeke Hekstra
    Reviewer

In the interests of transparency, eLife publishes the most substantive revision requests and the accompanying author responses.

Acceptance summary:

This paper describes the use of time-resolved serial femtosecond crystallography to investigate light-induced changes in the conformation of channelrhodopsin. The authors identified initial conformational changes that occur upon illumination, including a shift in the position of retinal as well as additional changes in the conformation of transmembrane helices 3 and 7. Using these results, the authors propose a model for how initial conformational changes may culminate in channel opening. This work advances our understanding of the molecular mechanisms of channelrhodopsin activation and of light-induced conformational change more generally.

Decision letter after peer review:

Thank you for submitting your article "Time-resolved serial femtosecond crystallography reveals early structural changes in channelrhodopsin" for consideration by eLife. Your article has been reviewed by three peer reviewers, and the evaluation has been overseen by a Reviewing Editor and Kenton Swartz as the Senior Editor. The following individual involved in review of your submission has agreed to reveal their identity: Doeke Hekstra (Reviewer #1).

The reviewers have discussed the reviews with one another and the Reviewing Editor has drafted this decision to help you prepare a revised submission.

As the editors have judged that your manuscript is of interest but revised data analysis is required before it is published, we would like to draw your attention to changes in our revision policy that we have made in response to COVID-19 (https://elifesciences.org/articles/57162). First, because many researchers have temporarily lost access to the labs, we will give authors as much time as they need to submit revised manuscripts. We are also offering, if you choose, to post the manuscript to bioRxiv (if it is not already there) along with this decision letter and a formal designation that the manuscript is "in revision at eLife". Please let us know if you would like to pursue this option. (If your work is more suitable for medRxiv, you will need to post the preprint yourself, as the mechanisms for us to do so are still in development.)

Summary:

This manuscript by K. Oda et al. describes the use of time-resolved serial femtosecond crystallography to investigate light-induced changes in the conformation of channelrhodopsin. The manuscript identifies initial conformational changes that occur upon illumination, including a shift in the position of retinal as well as additional changes in the conformation of transmembrane helices 3 and 7. The authors propose a model for how these initial changes may culminate in channel opening.

Essential revisions:

Several technical points were raised during the review process, but the most significant concern centers on the approach to model refinement. The description of the refinement process in the Materials and methods section is very brief, and is missing essential details such as the choice of restraints and the exact details of which difference maps were used. More generally, the use of real-space refinement directly against difference maps is not sufficiently described, and is potentially confounded by a variety of issues. For example, motion of an amino acid side chain into a region that would otherwise be occupied by an ordered water molecule could lead to little change in Fo-Fo density, or changes that are not readily interpretable. Reciprocal space refinement against extrapolated structure factors or reciprocal space refinement of partial occupancy models for each conformational state would be more appropriate (see below). At least one of these approaches should be included in a revised manuscript, along with appropriate statistical benchmarks such as real or reciprocal space correlation coefficients or R-factors. The reviewers also wish to emphasize that it is more appropriate to refine against structure factors corresponding to a conventional electron density (2Fo-Fc type) map than to a difference map. A second significant concern is the mismatch between the timescales of the QM/MM simulations and the experiments, which is not adequately explained. There are a variety of smaller technical points raised, which are discussed in detail below.

1) The authors mentioned low quantum efficiency of the retinal isomerization (~30% in C1C2), and then they built models onto the observed peaks in the difference maps. This can be problematic without validation, and the difference map signals can be misleading. In order to justify the correctness of the refined structures, the difference maps calculated between structures of light and dark states should be presented. Furthermore, more details should be included in structure refinement section, such as restraints, weighting factors, and validation metrics. Alternatively, the extrapolation approach has been adapted by several recent TR-SFX studies, such as the bR or KR2 studies (Science 365, 61-65, 2019); Nature 583, 314-318, 2020). Authors may want to try these approaches. In principle, it may also be possible to refine models of different states with partial occupancy based on the known photoconversion efficiency, which would also provide useful refinement statistics to assess model quality.

2) A vexing complication of the experiments is that the spectroscopic kinetics in crystal form differ markedly from those in solution. Clearly, the crystallographic data show that retinal isomerization is induced in the crystal and leads to conformational change at nearby positions. Spectroscopically, only two states are clearly distinguishable within the time scales of interest (P1/3 versus P2), reflecting the protonation state of the Schiff base. The authors speculate that they see a longer-lived P2 state in the crystal, apparently concurrent with a P1 or P3 state. This is reasonable, but it is not clear whether the authors really see a "P1 to P2 transition" (inclusion of shorter pump-probe delays might have been more convincing). The spectroscopic data likely miss detail of the structural transitions following photoexcitation and do not shed much light on how conformational dynamics may differ between crystal and solution.

3) The comparison between refined structures and the QM/MM simulation model is not convincing. QM/MM calculation results are based on short simulations. The timescales do not match with the ones studied in this TR-SFX. It is hard to make a fair comparison.

https://doi.org/10.7554/eLife.62389.sa1

Author response

Essential revisions:

Several technical points were raised during the review process, but the most significant concern centers on the approach to model refinement. The description of the refinement process in the Materials and methods section is very brief, and is missing essential details such as the choice of restraints and the exact details of which difference maps were used. More generally, the use of real-space refinement directly against difference maps is not sufficiently described, and is potentially confounded by a variety of issues. For example, motion of an amino acid side chain into a region that would otherwise be occupied by an ordered water molecule could lead to little change in Fo-Fo density, or changes that are not readily interpretable. Reciprocal space refinement against extrapolated structure factors or reciprocal space refinement of partial occupancy models for each conformational state would be more appropriate (see below). At least one of these approaches should be included in a revised manuscript, along with appropriate statistical benchmarks such as real or reciprocal space correlation coefficients or R-factors. The reviewers also wish to emphasize that it is more appropriate to refine against structure factors corresponding to a conventional electron density (2Fo-Fc type) map than to a difference map. A second significant concern is the mismatch between the timescales of the QM/MM simulations and the experiments, which is not adequately explained. There are a variety of smaller technical points raised, which are discussed in detail below.

According to the comments, we performed the reciprocal space refinement against the extrapolated structure factors, which is included below as a response to the reviewers.

1) The authors mentioned low quantum efficiency of the retinal isomerization (~30% in C1C2), and then they built models onto the observed peaks in the difference maps. This can be problematic without validation, and the difference map signals can be misleading. In order to justify the correctness of the refined structures, the difference maps calculated between structures of light and dark states should be presented. Furthermore, more details should be included in structure refinement section, such as restraints, weighting factors, and validation metrics. Alternatively, the extrapolation approach has been adapted by several recent TR-SFX studies, such as the bR or KR2 studies (Science 365, 61-65, 2019); Nature 583, 314-318, 2020). Authors may want to try these approaches. In principle, it may also be possible to refine models of different states with partial occupancy based on the known photoconversion efficiency, which would also provide useful refinement statistics to assess model quality.

We appreciate the reviewer’s critical comments on our study. We calculated the extrapolation maps according to the recent TR-SFX papers (Figure 4—figure supplement 3). First, we roughly estimated the ratio of the excited state by calculating extrapolated maps with different activation ratios until a feature of the dark state at Cys167 disappeared, and then modeled the excited states and performed the reciprocal refinement against the extrapolated maps. The obtained models confirmed the structural changes in TM3 and TM7, one of the major issues in the current manuscript (Figure 4—figure supplements 4, 5). However, this approach could not provide detailed information about the retinal conformation, because it showed only a weak density around the C13-C14 double bond of the retinal, probably due to its intrinsic flexibility (Figure 4—figure supplements 4, 5). Despite the lack of strong evidence, we speculate that the retinal adopts a kink toward TM3, based on the difference Fourier map that showed strong positive density beside the retinal, as well as the results of the QM/MM simulation. It should also be noted that a similar retinal kink and TM3 shift were visualized in recent work on another rhodopsin protein, KR2 (Skopintsev et al., 2020). Therefore, we supposed that the series of conformational changes is somehow conserved in the rhodopsin family proteins, although it was not observed in the representative member, bacteriorhodopsin. We would like to include both the normal difference Fourier maps (Figure 4) and the extrapolated maps (Figure 4—figure supplement 3), because high resolution diffraction data are required to calculate reliable extrapolation maps, and our maps thus contain a lot of noise.

2) A vexing complication of the experiments is that the spectroscopic kinetics in crystal form differ markedly from those in solution. Clearly, the crystallographic data show that retinal isomerization is induced in the crystal and leads to conformational change at nearby positions. Spectroscopically, only two states are clearly distinguishable within the time scales of interest (P1/3 versus P2), reflecting the protonation state of the Schiff base. The authors speculate that they see a longer-lived P2 state in the crystal, apparently concurrent with a P1 or P3 state. This is reasonable, but it is not clear whether the authors really see a "P1 to P2 transition" (inclusion of shorter pump-probe delays might have been more convincing). The spectroscopic data likely miss detail of the structural transitions following photoexcitation and do not shed much light on how conformational dynamics may differ between crystal and solution.

Conformational transitions are generally slower in crystals than in solution, and our spectroscopic study showed the same trend in C1C2. Figure 2—figure supplement 3 shows clear evidence that the P1 formation is completed within 10 μs, and the subsequent P2 formation is most prominent in 10 msec in crystals, which is slower than the corresponding steps in solution. Also, the lack of the P3 formation is consistent with the idea that the later reaction is more likely to be affected by crystal packing interactions. This information is sufficient to support the proposal that the dominant changes along the TR-SFX (occurring between 10 μs – 2.1 ms) are derived from the conformational transition from P1 to P2, although slight contamination by other intermediates (such as P3) might exist. It would be informative if we could understand the details of the early changes (including the precise population of photo-intermediates) after photoexcitation, but this is technically difficult because the spectrum changes in the tiny crystals of C1C2 are very weak. In addition, the spectroscopic studies only provide information about the micro-environmental changes around the retinal Schiff base and do not directly correlate with the global conformational changes of the protein. The previous bR TR-SFX study also suggested the lack of later conformational changes in the crystals, and suggested that the local changes in the Schiff base and the global structural changes are not necessarily coincident, especially in the later reactions. Therefore, the conclusion in the current manuscript is minimally dependent on the spectroscopic study. The observed changes in TM3 may occur during the P1 to P2 transition, and probably more quickly in solution.

3) The comparison between refined structures and the QM/MM simulation model is not convincing. QM/MM calculation results are based on short simulations. The timescales do not match with the ones studied in this TR-SFX. It is hard to make a fair comparison.

The molecular dynamics simulations in the QM/MM free energy geometry optimization by Cheng et al., 2018, showed the precursor state of the channel opening (early P2 state) on the timescale of tens to hundreds of nanoseconds. Although this time scale is much longer than that of the conventional QM/MM simulations (typically tens of picoseconds; e.g., Ardevol and Hummer, PNAS 115, 3557 (2018)), it is still shorter than the experimental time scale of the pore formation, 2 ~10 ms (Lorenz-Fonfria et al., 2013). However, please note that the timescales of the QM/MM simulations do not directly represent the experimental timescale of the early P2 formation. Spectroscopic measurements showed that the time constant of the early P2 state formation coincides with that of the proton translocation from the protonated Schiff base to its counterion, glutamate (Lorenz-Fonfria et al., 2013), exhibiting a deuterium kinetic isotope effect (Resler et al., Biophys. J. 109, 287 (2015)). These studies indicate that the early P2 formation is kinetically governed by the proton translocation. In contrast, the QM/MM calculations simulated the conformational changes of the protein after the proton translocation event. The proton translocation has already been well characterized experimentally. Therefore, it was omitted in the simulation to enable the modeling of the conformational changes triggered by the proton translocation, at the unprecedentedly high level of theory achieved by the combination of highly accurate ab initio quantum chemistry. The key chemical feature revealed by the QM/MM simulations was the torsional motion from a locally twisted conformation in the Schiff base, due to the increased electronic double bond character upon the proton translocation. This can be properly described by the ab initio QM treatment of the QM/MM simulations and is in excellent agreement with the current TR-SFX observation. The molecular events described in the simulations are distinctly different from those in the experiments. Therefore, although the simulation and current TR-SFX both indicated the retinal kink upon the proton transfer, the timescales of these results do not exactly match. These points are clarified in the revised manuscript (although the QM/MM simulation focuses only on the molecular events after the Schiff base deprotonation and thus does not exactly match the experimental time scale.).

https://doi.org/10.7554/eLife.62389.sa2

Article and author information

Author details

  1. Kazumasa Oda

    Department of Biological Sciences, Graduate School of Science, The University of Tokyo, Tokyo, Japan
    Contribution
    Conceptualization, Data curation, Investigation, Visualization, Methodology, Writing - original draft, Project administration, Writing - review and editing
    Contributed equally with
    Takashi Nomura, Takanori Nakane and Keitaro Yamashita
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0003-0996-0393
  2. Takashi Nomura

    Graduate School of Life Science, University of Hyogo, Hyogo, Japan
    Contribution
    Conceptualization, Data curation, Investigation, Methodology, Writing - original draft, Writing - review and editing
    Contributed equally with
    Kazumasa Oda, Takanori Nakane and Keitaro Yamashita
    Competing interests
    No competing interests declared
  3. Takanori Nakane

    Department of Biological Sciences, Graduate School of Science, The University of Tokyo, Tokyo, Japan
    Present address
    Structural Studies Division, MRC Laboratory of Molecular Biology, Cambridge Biomedical Campus, Cambridge, United Kingdom
    Contribution
    Data curation, Software, Supervision
    Contributed equally with
    Kazumasa Oda, Takashi Nomura and Keitaro Yamashita
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0003-2697-2767
  4. Keitaro Yamashita

    Department of Biological Sciences, Graduate School of Science, The University of Tokyo, Tokyo, Japan
    Present address
    Structural Studies Division, MRC Laboratory of Molecular Biology, Cambridge Biomedical Campus, Cambridge, United Kingdom
    Contribution
    Software and model refinement
    Contributed equally with
    Kazumasa Oda, Takashi Nomura and Takanori Nakane
    Competing interests
    No competing interests declared
  5. Keiichi Inoue

    Graduate School of Engineering, Nagoya Institute of Technology, Nagoya, Japan
    Present address
    Institute for Solid State Physics, University of Tokyo, Kashiwa, Japan
    Contribution
    Data curation, Validation, Investigation, Writing - original draft, Writing - review and editing
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-6898-4347
  6. Shota Ito

    Graduate School of Engineering, Nagoya Institute of Technology, Nagoya, Japan
    Contribution
    Investigation
    Competing interests
    No competing interests declared
  7. Johannes Vierock

    Institute of Biology, Experimental Biophysics, Humboldt-Universität zu Berlin, Berlin, Germany
    Contribution
    Data curation, Formal analysis, Visualization, Writing - original draft, Writing - review and editing
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0001-7368-5539
  8. Kunio Hirata

    1. RIKEN SPring-8 Center, Hyogo, Japan
    2. Precursory Research for Embryonic Science and Technology (PRESTO), Japan Science and Technology Agency, Kawaguchi, Japan
    Contribution
    Investigation
    Competing interests
    No competing interests declared
  9. Andrés D Maturana

    Department of Bioengineering Sciences, Graduate School of Bioagricultural Sciences, Nagoya University, Nagoya, Japan
    Contribution
    Data curation, Investigation
    Competing interests
    No competing interests declared
  10. Kota Katayama

    Graduate School of Engineering, Nagoya Institute of Technology, Nagoya, Japan
    Contribution
    Investigation
    Competing interests
    No competing interests declared
  11. Tatsuya Ikuta

    Department of Biological Sciences, Graduate School of Science, The University of Tokyo, Tokyo, Japan
    Contribution
    Investigation
    Competing interests
    No competing interests declared
  12. Itsuki Ishigami

    Department of Biological Sciences, Graduate School of Science, The University of Tokyo, Tokyo, Japan
    Contribution
    Investigation
    Competing interests
    No competing interests declared
  13. Tamaki Izume

    Department of Biological Sciences, Graduate School of Science, The University of Tokyo, Tokyo, Japan
    Contribution
    Investigation
    Competing interests
    No competing interests declared
  14. Rie Umeda

    Department of Biological Sciences, Graduate School of Science, The University of Tokyo, Tokyo, Japan
    Contribution
    Investigation
    Competing interests
    No competing interests declared
  15. Ryuun Eguma

    Department of Biological Sciences, Graduate School of Science, The University of Tokyo, Tokyo, Japan
    Contribution
    Investigation
    Competing interests
    No competing interests declared
  16. Satomi Oishi

    Department of Biological Sciences, Graduate School of Science, The University of Tokyo, Tokyo, Japan
    Contribution
    Investigation
    Competing interests
    No competing interests declared
  17. Go Kasuya

    Department of Biological Sciences, Graduate School of Science, The University of Tokyo, Tokyo, Japan
    Present address
    Division of Integrative Physiology, Department of Physiology, Jichi Medical University, Shimotsuke, Japan
    Contribution
    Investigation
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0003-1756-5764
  18. Takafumi Kato

    Department of Biological Sciences, Graduate School of Science, The University of Tokyo, Tokyo, Japan
    Contribution
    Investigation
    Competing interests
    No competing interests declared
  19. Tsukasa Kusakizako

    Department of Biological Sciences, Graduate School of Science, The University of Tokyo, Tokyo, Japan
    Contribution
    Investigation
    Competing interests
    No competing interests declared
  20. Wataru Shihoya

    Department of Biological Sciences, Graduate School of Science, The University of Tokyo, Tokyo, Japan
    Contribution
    Investigation
    Competing interests
    No competing interests declared
  21. Hiroto Shimada

    Department of Biological Sciences, Graduate School of Science, The University of Tokyo, Tokyo, Japan
    Contribution
    Investigation
    Competing interests
    No competing interests declared
  22. Tomoyuki Takatsuji

    Department of Biological Sciences, Graduate School of Science, The University of Tokyo, Tokyo, Japan
    Contribution
    Investigation
    Competing interests
    No competing interests declared
  23. Mizuki Takemoto

    Department of Biological Sciences, Graduate School of Science, The University of Tokyo, Tokyo, Japan
    Present address
    Preferred Networks, Inc, Tokyo, Japan
    Contribution
    Investigation
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-6339-6431
  24. Reiya Taniguchi

    Department of Biological Sciences, Graduate School of Science, The University of Tokyo, Tokyo, Japan
    Contribution
    Investigation
    Competing interests
    No competing interests declared
  25. Atsuhiro Tomita

    Department of Biological Sciences, Graduate School of Science, The University of Tokyo, Tokyo, Japan
    Contribution
    Investigation
    Competing interests
    No competing interests declared
  26. Ryoki Nakamura

    Department of Biological Sciences, Graduate School of Science, The University of Tokyo, Tokyo, Japan
    Contribution
    Investigation
    Competing interests
    No competing interests declared
  27. Masahiro Fukuda

    Department of Biological Sciences, Graduate School of Science, The University of Tokyo, Tokyo, Japan
    Contribution
    Investigation
    Competing interests
    No competing interests declared
  28. Hirotake Miyauchi

    Department of Biological Sciences, Graduate School of Science, The University of Tokyo, Tokyo, Japan
    Contribution
    Investigation
    Competing interests
    No competing interests declared
  29. Yongchan Lee

    Department of Biological Sciences, Graduate School of Science, The University of Tokyo, Tokyo, Japan
    Present address
    Department of Structural Biology, Max Planck Institute of Biophysics, Frankfurt, Germany
    Contribution
    Investigation
    Competing interests
    No competing interests declared
  30. Eriko Nango

    1. RIKEN SPring-8 Center, Hyogo, Japan
    2. Department of Cell Biology, Graduate School of Medicine, Kyoto University, Kyoto, Japan
    Present address
    Institute of Multidisciplinary Research for Advanced Materials, Tohoku University, Sendai, Japan
    Contribution
    Conceptualization, Supervision, Investigation
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0001-9851-7355
  31. Rie Tanaka

    1. RIKEN SPring-8 Center, Hyogo, Japan
    2. Department of Cell Biology, Graduate School of Medicine, Kyoto University, Kyoto, Japan
    Contribution
    Supervision, Project administration
    Competing interests
    No competing interests declared
  32. Tomoyuki Tanaka

    1. RIKEN SPring-8 Center, Hyogo, Japan
    2. Department of Cell Biology, Graduate School of Medicine, Kyoto University, Kyoto, Japan
    Contribution
    Investigation
    Competing interests
    No competing interests declared
  33. Michihiro Sugahara

    RIKEN SPring-8 Center, Hyogo, Japan
    Contribution
    Investigation, Project administration
    Competing interests
    No competing interests declared
  34. Tetsunari Kimura

    Department of Chemistry, Graduate School of Science, Kobe University, Kobe, Japan
    Contribution
    Investigation
    Competing interests
    No competing interests declared
  35. Tatsuro Shimamura

    Department of Cell Biology, Graduate School of Medicine, Kyoto University, Kyoto, Japan
    Contribution
    Investigation
    Competing interests
    No competing interests declared
  36. Takaaki Fujiwara

    Department of Cell Biology, Graduate School of Medicine, Kyoto University, Kyoto, Japan
    Contribution
    Investigation
    Competing interests
    No competing interests declared
  37. Yasuaki Yamanaka

    Department of Cell Biology, Graduate School of Medicine, Kyoto University, Kyoto, Japan
    Contribution
    Investigation
    Competing interests
    No competing interests declared
  38. Shigeki Owada

    1. RIKEN SPring-8 Center, Hyogo, Japan
    2. Japan Synchrotron Radiation Research Institute, Hyogo, Japan
    Contribution
    Software, Supervision
    Competing interests
    No competing interests declared
  39. Yasumasa Joti

    1. RIKEN SPring-8 Center, Hyogo, Japan
    2. Japan Synchrotron Radiation Research Institute, Hyogo, Japan
    Contribution
    Software, Supervision, Investigation
    Competing interests
    No competing interests declared
  40. Kensuke Tono

    1. RIKEN SPring-8 Center, Hyogo, Japan
    2. Japan Synchrotron Radiation Research Institute, Hyogo, Japan
    Contribution
    Software, Supervision
    Competing interests
    No competing interests declared
  41. Ryuichiro Ishitani

    Department of Biological Sciences, Graduate School of Science, The University of Tokyo, Tokyo, Japan
    Contribution
    Supervision, Project administration
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-4136-5685
  42. Shigehiko Hayashi

    Department of Chemistry, Graduate School of Science, Kyoto University, Kyoto, Japan
    Contribution
    Supervision
    Competing interests
    No competing interests declared
  43. Hideki Kandori

    Graduate School of Engineering, Nagoya Institute of Technology, Nagoya, Japan
    Contribution
    Supervision, Writing - review and editing
    Competing interests
    No competing interests declared
  44. Peter Hegemann

    Institute of Biology, Experimental Biophysics, Humboldt-Universität zu Berlin, Berlin, Germany
    Contribution
    Supervision, Validation, Visualization, Writing - original draft, Writing - review and editing
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0003-3589-6452
  45. So Iwata

    1. RIKEN SPring-8 Center, Hyogo, Japan
    2. Department of Cell Biology, Graduate School of Medicine, Kyoto University, Kyoto, Japan
    Contribution
    Supervision, Project administration
    Competing interests
    No competing interests declared
  46. Minoru Kubo

    Graduate School of Life Science, University of Hyogo, Hyogo, Japan
    Contribution
    Data curation, Supervision, Validation, Investigation, Visualization, Writing - original draft, Project administration, Writing - review and editing
    For correspondence
    minoru@sci.u-hyogo.ac.jp
    Competing interests
    No competing interests declared
  47. Tomohiro Nishizawa

    1. Department of Biological Sciences, Graduate School of Science, The University of Tokyo, Tokyo, Japan
    2. Precursory Research for Embryonic Science and Technology (PRESTO), Japan Science and Technology Agency, Kawaguchi, Japan
    Contribution
    Conceptualization, Data curation, Supervision, Validation, Investigation, Visualization, Writing - original draft, Project administration, Writing - review and editing
    For correspondence
    t-2438@bs.s.u-tokyo.ac.jp
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0001-7463-8398
  48. Osamu Nureki

    Department of Biological Sciences, Graduate School of Science, The University of Tokyo, Tokyo, Japan
    Contribution
    Conceptualization, Funding acquisition, Project administration, Writing - review and editing
    For correspondence
    nureki@biochem.s.u-tokyo.ac.jp
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0003-1813-7008

Funding

Japan Society for the Promotion of Science (16H06294)

  • Osamu Nureki

Japan Society for the Promotion of Science (18J21256)

  • Kazumasa Oda

Japan Society for the Promotion of Science (17H05000)

  • Tomohiro Nishizawa

Japan Science and Technology Agency (JPMJPR14L9)

  • Kunio Hirata

Japan Science and Technology Agency (JPMJPR14L8)

  • Tomohiro Nishizawa

Japan Agency for Medical Research and Development (Platform Project for Supporting Drug Discovery andLife Science Research (Basis for Supporting Innovative Drug Discovery and Life Science Research(BINDS)))

  • Kunio Hirata

Deutsche Forschungsgemeinschaft (SFB1078 (B2))

  • Johannes Vierock

Cluster of Excellence in Inflammation Research (UniCat)

  • Johannes Vierock

Cluster of Excellence in Inflammation Research (BIG-NSE)

  • Johannes Vierock

Cluster of Excellence in Inflammation Research (E4)

  • Peter Hegemann

Cluster of Excellence in Inflammation Research (SPP 1926)

  • Johannes Vierock

European Research Council (ERC-2016-StG 714762)

  • Johannes Vierock

Hertie Foundation (Senior Research Professor)

  • Peter Hegemann

Ministry of Education, Culture, Sports, Science and Technology (X-ray Free-Electron Laser Priority Strategy Program)

  • So Iwata

The funders had no role in study design, data collection and interpretation, or the decision to submit the work for publication.

Acknowledgements

We thank Sae Okazaki and Masae Miyazaki for assistance in obtaining the C1C2 crystals, and Rie Tanaka, Toshi Arima, Ayumi Yamashita, and Jun Kobayashi for assistance in the collection of the X-ray data diffraction data at SACLA. We also thank the beamline staff scientists at SACLA and BL32XU of SPring-8 (Hyogo, Japan). The XFEL experiments were performed at the BL2 of SACLA, with the approval of the Japan Synchrotron Radiation Research Institute (JASRI) (proposal nos. 2015B8053, 2016A8054, 2016B8067, 2017A8022, and 2017B8035), and SPring-8 BL32XU (20170511, 20170721, and 20170728), with the approval of RIKEN. We acknowledge computational support from the SACLA HPC system and Mini-K super computer system.

Senior Editor

  1. Kenton J Swartz, National Institute of Neurological Disorders and Stroke, National Institutes of Health, United States

Reviewing Editor

  1. Andrew C Kruse, Harvard Medical School, United States

Reviewer

  1. Doeke Hekstra

Publication history

  1. Received: August 23, 2020
  2. Accepted: February 15, 2021
  3. Version of Record published: March 23, 2021 (version 1)
  4. Version of Record updated: March 31, 2021 (version 2)

Copyright

© 2021, Oda et al.

This article is distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use and redistribution provided that the original author and source are credited.

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