Feature Article pubs.acs.org/JPCB
The Grateful Infrared: Sequential Protein Structural Changes Resolved by Infrared Difference Spectroscopy Tilman Kottke,† Víctor A. Lórenz-Fonfría,‡,§ and Joachim Heberle*,∥ †
Department of Chemistry, Physical and Biophysical Chemistry, Bielefeld University, Universitätsstraße 25, 33615 Bielefeld, Germany Department of Biochemistry and Molecular Biology and §Interdisciplinary Research Structure for Biotechnology and Biomedicine (ERI BIOTECMED), Universitat de València, Carrer Doctor Moliner 50, 46100 Burjassot, Spain ∥ Experimental Molecular Biophysics, Freie Universität Berlin, Arnimalle 14, 14195 Berlin, Germany ‡
ABSTRACT: The catalytic activity of proteins is a function of structural changes. Very often these are as minute as protonation changes, hydrogen bonding changes, and amino acid side chain reorientations. To resolve these, a methodology is afforded that not only provides the molecular sensitivity but allows for tracing the sequence of these hierarchical reactions at the same time. This feature article showcases results from time-resolved IR spectroscopy on channelrhodopsin (ChR), light-oxygen-voltage (LOV) domain protein, and cryptochrome (CRY). All three proteins are activated by blue light, but their biological role is drastically different. Channelrhodopsin is a transmembrane retinylidene protein which represents the first light-activated ion channel of its kind and which is involved in primitive vision (phototaxis) of algae. LOV and CRY are flavin-binding proteins acting as photoreceptors in a variety of signal transduction mechanisms in all kingdoms of life. Beyond their biological relevance, these proteins are employed in exciting optogenetic applications. We show here how IR difference absorption resolves crucial structural changes of the protein after photonic activation of the chromophore. Time-resolved techniques are introduced that cover the time range from nanoseconds to minutes along with some technical considerations. Finally, we provide an outlook toward novel experimental approaches that are currently developed in our laboratories or are just in our minds (“Gedankenexperimente”). We believe that some of them have the potential to provide new science.
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Difference and Double Difference Spectroscopy. IR absorption spectroscopy responds to a vast number of vibrational signals from the protein and other biomolecules, leading to extremely crowded absorption spectra (Figure 1). Therefore, a filter is needed to enable the selection of relevant signals. Such discrimination is achieved by dif ference spectroscopy,7 where absorption changes associated with the reaction of a protein are selectively extracted from the constant absorption background. This approach is not merely a trivial subtraction of absorption spectra of the sample before and after a reaction step has taken place. More accurately, the respective transmitted infrared intensities Iequilibrium and Iperturbed are directly transformed into a
INTRODUCTION
Vibrational spectroscopy applied to proteins yields information on the structure and oxidation state of cofactors, on the protonation state and hydrogen bonding status of amino acids, and on the secondary structure elements via the analysis of the amide I and amide II vibrational bands of the peptide backbone.1 In comparison to other biophysical techniques, IR spectroscopy is particularly rewarding when subtle but crucial structural changes are of interest. This is the field of molecular biophysics with powerful methodologies as X-ray scattering and crystallography, electron microscopy, nuclear and electron magnetic resonance and other spectroscopies. IR spectroscopy, however, suffers from structural resolution unless the vibrational sibling of NMR, 2D-IR spectroscopy, is applied.2−4 Although great methodological progress has been achieved over the recent decade,5,6 the application of this fascinating technique to proteins is still in its infancy. The paramount advantage of IR spectroscopy is the inherent high temporal resolution attainable which is in the femtosecond time range. As proteins can be considered nanomachines whose structural changes cover the enormous time range from 10−15 to 100 seconds, reaction dynamics are traced by IR spectroscopy at utmost temporal resolution and fine structural sensitivity. © 2016 American Chemical Society
difference absorbance ΔA according to ΔA = −log
Iperturbed Iequilibrium
. In
the experiment, this procedure is equivalent to the sampling of the identical proteins before and after the reaction. Therefore, great care needs to be taken to maintain the sampled volume, concentration, temperature, pH, etc., during the reaction. This requirement favors the application of difference spectroscopy to reactions induced by light or a change in potential over others Received: September 12, 2016 Revised: November 6, 2016 Published: November 14, 2016 335
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The assignment of vibrational bands to specific molecular modes is achieved by a combination of mutations, isotope labeling, and computational chemistry. Mutations of proteins, if carefully selected, are invaluable to assign signals to specific side chains as will be showcased in the following sections. It is noted, however, that mutations may interfere indirectly and even unpredictably with the mechanism. Difference spectroscopy relies on functional proteins, and the replacement of an amino acid side chain critical to the studied molecular reaction may lead to an impaired or even nonfunctional protein from which it is hardly possible to draw meaningful conclusions. As an alternative, the rapid development of both computational power and inexpensive methods enables theoreticians to achieve better agreement between experimental and theoretical spectra by including larger parts of active sites of proteins into the quantum chemical calculations. Most demanding is the production of difference14,15 or even double difference8 spectra after folding of the line spectra with Lorentzians or Gaussians, because little deviations in the calculated frequencies lead to larger variations in the characteristic signal pattern. In the future, computational assignments may render experiments on mutants or isotope-labeled samples redundant. Time-Resolved IR Spectroscopy. The capability of monitoring and identifying biological processes continuously over the whole time range of femtoseconds to hours is a key advantage of IR spectroscopy as compared to many other techniques. In this review, the focus will be on the monitoring of processes in the time range from tens of nanoseconds to hundreds of seconds, a range central for understanding the chemistry and physics of protein function. The most commonly applied technique involves the broadband detection of the mid-infrared spectrum as delivered by the source, a blackbody radiator. The necessary sensitivity for detecting and analyzing such a comparatively weak light source is achieved by modulation of the electromagnetic field with a Michelson interferometer and subsequent demodulation by Fourier transformation (FT). Fast modulation by the mirror of the interferometer leads to the rapid-scan technique, by which a time resolution of around 10 ms is routinely achieved by commercial FT-IR spectrometers. More demanding is the stepscan technique,16,17 suitable to cover the time region from tens of nanoseconds to hundreds of milliseconds. There, the modulating mirror is moved in a stepwise manner, and time traces are recorded as changes in the integrated intensity at each interferogram position. The sorting of the resulting data matrix with respect to each time point then allows the researcher to perform the FT and finally yields continuous time-dependent spectra. Such gapless monitoring of processes in time is a decisive advantage to the pump−probe approach. Step-scan spectroscopy requires a strictly repetitive process to take place, or else the exchange for a spectroscopically identical fresh sample after each step. More straightforward is the complementary application of a monochromatic probe light as supplied by semiconductor laser diodes or, more recently, by quantum cascade lasers (QCL).18,19 Thereby, kinetics in proteins at single frequencies are directly monitored, bypassing the need of an interferometer.20−23 As these QCLs are tunable in their emission wavelength, spectral information can be retrieved over a limited frequency range 150−200 cm−1 per laser head. The collected time-resolved data represents the averaged spectral properties of all the intermediates weighted by their time-variable fraction. The decomposition of the recorded data
Figure 1. Concept of difference and double difference spectroscopy. The IR absorption spectrum is composed of a manifold of vibrational bands from protein and water, among them the most intense amide I (CO stretching vibration) and amide II (coupled CN stretching and NH bending vibration) modes of the protein backbone. The broad sum spectrum contains little extractable information except for the secondary structure derived by analysis of the amide I mode. If the protein is sampled before and after a (photo)reaction, the difference spectrum shows only bands of those vibrations that are changed in the process and are assigned to functional groups involved in the reaction. Two difference spectra (a and b) of proteins differing by the presence and absence of a segment (here: a C-terminal Jα helix11) are subtracted to produce a double difference spectrum (c), which ideally contains only signals from the segment of interest.
that depend on a mixing of two reaction partners or a change in the buffer conditions. The focus of the experiment and the isolation of the respective signals can be enhanced by applying double dif ference spectroscopy. Two different samples are processed in precisely the same manner to produce a set of two difference spectra that differ in only one aspect. A common example would be the direct comparison of an isotope-labeled sample with its counterpart at natural abundance. The double difference then shows ideally only two pairs of difference signals reflecting the changes by the increase in mass of this amino acid, which simplifies interpretation and assignment.8,9 The application can be extended to two proteins that differ by part of its fold or a whole domain, i.e., a segment-resolved double dif ference spectrum.10,11 The resulting spectrum contains only those signals that depend on the presence of the particular element or domain, which often allows spectroscopists to localize the spatial origin of a detected vibration.11−13 336
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Figure 2. All-trans and 13-cis photocycles of channelrhodopsin-2. (A) Surface plot of the experimental transient absorption changes of the E123T variant in the retinal ethylenic (CC stretching) region. Contour lines correspond to global fitting to eight exponentials. The position of the most intense negative band is indicated for the experimental (dots) and fitted (red trace) data. (B) Difference spectra extracted at selected times, band-narrowed by MaxEntD.50 The negative bands at 1551 cm−1 and at 1558 cm−1 are from bleached dark-state all-trans and 13-cis retinal, respectively. (C) Simplified version of the proposed 13-cis (top) and all-trans (bottom) retinal photocycles. Time constants for the all-trans photocycle correspond to wild-type CrChR2. Panel C has been adapted with permission from ref 22. Copyright 2015 American Chemical Society.
(SB).38 The function of microbial rhodopsins is driven by the photoisomerization of the retinal chromophore around the C13C14 bond.38 The protein follows a complex thermal relaxation process, lasting for tens of seconds in ChRs. The photocyclic reaction of CrChR2 displays at least four spectroscopically different intermediates: P1500, P2390, P3520, and P4480.39,40 The conductive state of the channel has been mostly ascribed to the P3520 intermediate39 and to a late P2390 substate.41 Practical Aspects of Time-Resolved IR Experiments on CrChR2. Two major challenges arise when performing timeresolved IR studies on CrChR2. The first challenge is the small IR absorption changes following a short laser flash in the order of 10−4 or less for most bands.42 Optimizing the optical throughput of the FT-IR setup43 and the averaging of ∼105 photoreactions yielded sufficient signal-to-noise ratio in our time-resolved stepscan experiments. Further reduction in noise was achieved by logarithmic averaging and by data reconstruction using singular value decomposition.43 The second challenge is the slow recovery after photoexcitation to the dark state of CrChR2, with a slower time constant of τ ∼ 20 s at room temperature.42,44 Due to the biphasic dark-state recovery kinetics,42 time-resolved step-scan experiments could be performed with minor distortions exciting the sample every 4 s. Even at this accelerated excitation rate, 5 days of uninterrupted data recording are required. Refilling of the liquid-nitrogen-cooled Dewar of the IR detector and sample replacement due to photodamage extend the duration of the step-scan experiment to 2 weeks in practice. All-trans and 13-cis Photocycles and Their Functionality. The retinal chromophore comes as a mixture of all-trans (70%) and 13-cis (30%) isomers in the dark state of CrChR2.45
into spectra and time evolution for each intermediate is notably ambiguous, 24,25 even though some progress has been achieved.26−29 One common compromise is to perform a phenomenological analysis. There, the time-resolved data is fitted to a discrete number of exponentials, and a sequential, unidirectional kinetic model is assumed to obtain the time evolution and spectra for pseudointermediates.30 Alternatively, if intermediate-specific bands are known, their time evolution can be selectively traced to derive dynamic information about given intermediates and their time for maximal contribution without the need of a kinetic model.31,32 We will showcase in the following recent results and advances in our laboratories based on applications of these techniques to contribute to a fundamental understanding of the reaction mechanism of the blue light receptor proteins channelrhodopsin, cryptochrome, and LOV. Finally, a perspective will be provided with novel experiments and new ideas for resolving the functional mechanism of other classes of membrane proteins by time-resolved IR spectroscopy.
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CHANNELRHODOPSIN Channelrhodopsins (ChRs) are transmembrane photoreceptor proteins located in the eyespot of unicellular green algae.33 When illuminated, they render the biomembrane transiently permeable to protons and other cations.34,35 Remarkably, blue light illumination of ChR2 from Chlamydomonas reinhardtii (CrChR2) expressed in neuronal cells was able to trigger action potentials,36,37 a breakthrough that led to the development of the vibrant field of optogenetics. ChRs are microbial rhodopsins with seven transmembrane helices and a retinal as a chromophore covalently bound to a lysine through a protonated Schiff base 337
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Figure 3. Proton transfer reactions in channelrhodopsin. (A) Illustration of spectral changes induced by protonation/deprotonation of a carboxylic group (left) and H-bonding changes (right). (B) Surface plot of the transient absorption changes of wild-type CrChR2 in the CO carboxylic region, and the global fit to a sum of seven exponentials (dashed contour lines). (C) Difference IR spectra extracted at selected times at which given intermediates reach maximal accumulation according to UV−vis experiments performed in parallel (brown trace), and after band-narrowing by Fourier self-deconvolution (black trace). The assignment of bands to E90, D156, and D253 is indicated. (D) Proposed protonation changes of SB, E90, D156, and D253 residues using the dark-state structure of the CrChR1−CrChR2 chimera.57 (E) Deprotonation and reprotonation kinetics of D156 (1737 cm−1, red trace), compared with the formation and decay of the P3520 intermediate (540 nm, gray trace). (F) H/D exchange of the hydroxyl group of D156 and the thiol group of C128 induce an 8 cm−1 down-shift (uncoupling) and 9 cm−1 up-shift (weaker H-bond) of the CO stretch of D156, respectively, accounting for its observed 1 cm−1 upshift in D2O. Panels C−E adapted with permission from ref 42. Copyright 2013 National Academy of Science.
the view that the main conductive state of CrChR2 is formed by photoisomerization of dark-state all-trans retinal (Figure 2C). Remarkably, a negative band at 1558 cm−1, assigned to the ethylenic band of dark-state 13-cis retinal, decayed with a half-life of t1/2 ≈ 1 ms (Figure 2B,C). Recovery of 13-cis retinal is 40times faster than the decay of the open to the dark-state in the secondary functional photocycle as predicted from the kinetic analysis of the photocurrents (O2 → D2),51,52 reasonably ruling out that the 13-cis photocycle could be its source as has been proposed.48,49 Although a conductive state could be still formed in the 13-cis photocycle, our results show that it is too short-lived to significantly contribute to the channel activity. Protonation Changes and Proton Transfer Reactions. The retinal SB deprotonates upon the formation of the P2390 intermediate as was clearly shown by resonance Raman spectroscopy.48,53 However, the identity of the SB proton acceptor remained unknown as well as the group that later reprotonates the SB. In retinal proteins, the SB proton acceptor and donor groups are mostly the carboxylic side chains of aspartates and glutamates and their corresponding acids, respectively.54 Protonation changes in Asp and Glu can be identified and characterized from the carbonyl stretching vibration of the carboxylic group, vCO, which typically appears in the frequency range between 1780 and 1690 cm−1. In a deprotonation reaction, the vCO of a carboxylic group is present in the starting but not in the intermediate state, leading to a negative band in the IR difference spectrum. Vice versa, a positive difference band is observed in a protonation reaction (Figure 3A). Hydrogen bonding changes at the CO or OH groups of protonated Asp or Glu affect the frequency of the vC
The photocycle starting from all-trans retinal is evidently associated with channel opening,40 but the analysis of the photocurrents of CrChR2 highlighted the presence of an additional photocycle.46,47 Since then, the functional relevance of the 13-cis photocycle has been a matter of a vivid debate.41,48,49 To address this issue, we characterized the recovery kinetics of dark-state 13-cis retinal after photoexcitation, taking advantage of the fact that the dark-state 13-cis retinal of CrChR2 shows a major band at ∼1558 cm−1 (λmax ≈ 460 nm), while the dark-state alltrans retinal does it at ∼1551 cm−1 (λmax ≈ 470 nm).45,48 Both ac-coupled and dc-coupled time-resolved step-scan FTIR experiments were conducted and merged to cover the time range from the nanoseconds to nearly 1 s (Figure 2A).22 The ac signal from the IR detector at a given optical retardation of the interferometer was digitalized by a 14-bit external transient recorder in 10 ns steps until 200 μs, with the effective resolution limited to 60 ns by the response time of the detector/ preamplifier. The dc signal was digitized by the internal ADC (24-bit), both at 6.25 μs and at 40 μs steps, allowing for experiments from 6.25 μs until 800 ms without exceeding the ADC memory. The faster cycling but fully functional E123T variant51,52 of CrChR2 was used with a 1 Hz excitation rate to reduce the total recording time.22 Time-resolved IR difference spectra showed a negative band at 1551 cm−1 after mathematical band-narrowing (Figure 2B), close to the expected frequency for dark-state all-trans retinal. The absorption changes of this band decayed with t1/2 ≈ 8 ms, similar to the decay of the open to the dark-state in the main photocycle (O1 → D1) as determined from photocurrents of CrChR2E123T in HEK cells.51,52 The good agreement is coherent with 338
DOI: 10.1021/acs.jpcb.6b09222 J. Phys. Chem. B 2017, 121, 335−350
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Figure 4. Hydration of transmembrane helices and its correlation with ion conductivity. (A) Surface plot of the transient absorption changes of wild-type CrChR2 in the amide I region, globally fitted to a sum of seven exponentials (dashed contour lines). Spectral changes reach a maximum at 1 ms (vertical dashed line). (B) Difference spectrum at 1 ms after photoexcitation, color coded by the cross-correlation of the transient absorption changes with the time-resolved photocurrents. (C) The maximum of the band at 1648 cm−1 (bottom left) but not the band at 1664 cm−1 (top left) is sensitive to H218O, suggesting the formation of H-bonds between water molecules and α-helical backbone carbonyls (right). (D) Lifetime distribution of the absorption changes in the amide I region. The bands at 1665 and 1648 cm−1 rise with time constants of ∼9 and ∼180 μs, and decay with a time constant of ∼9 ms. (E) Cartoon summarizing the correlation between the hydration of transmembrane regions and the formation of a transmembrane pore allowing for ion permeation. Adapted with permission from ref 69. Copyright 2015 National Academy of Science.
nonconservative (E90A, E123T, D156A, and D253N) and conservative (D156E and D253E) mutations for a complete band assignment (Figure 3C) and for a robust discrimination between protonation and H-bonding changes. Finally, to identify the SB proton acceptor (D253) and donor (D156) groups, the kinetics of their protonation changes were compared to the kinetics of the SB deprotonation and protonation.42 As an example, the bands at 1760 (+), 1745 (+), and 1736(−) cm−1 were assigned to D156 on the basis of their disappearance in the D156A mutant.42 The deprotonation of D156 in the P3520 intermediate was confirmed by the selective shift of the band from 1736 cm−1 in wild-type ChR2 to 1764 cm−1 in the D156E variant.42 Others have assigned the positive band at 1760 cm−1 to E123,60 despite the fact that this band is present in the E123T variant.22 We concluded that D156 is the SB proton donor based on the fact that the reprotonation kinetics of the retinal SB (rise of the P3520 state) tally the deprotonation kinetics of D156 (Figure 3E, blue arrow), and because the lifetime of the P2390 state is 104-fold retarded61 when a deprotonable residue is missing at position 156 of the polypeptide chain (e.g., in the D156A variant). Combining the determined protonation states of the retinal SB, D253, D156, and E90 during the photocycle (Figure 3C) with pH changes in the bulk measured using a pH-sensitive dye,62 we arrived at a self-consistent scheme for proton transfer reactions in CrChR2 (Figure 3D).42 Because D253 appears to remain protonated in P3520, the presence of a still unresolved group is invoked which released the proton to the medium
O bond, and generate a differential-shaped pair of positive/ negative bands in the IR difference spectrum (Figure 3A). Initially, vibrational changes in the carboxylic region have been recorded under continuous illumination at cryogenic and at room temperatures. Inasmuch as these static experiments55,56 formed a firm basis of understanding structural changes in CrChR2, it is evident that kinetic experiments performed under ambient conditions are superior to reflect on the functional mechanism. Therefore, we performed the first time-resolved IR experiments at microsecond resolution on a channelrhodopsin. Step-scan and rapid-scan FT-IR experiments were combined to monitor spectral changes from 6 μs to 100 s (Figure 3B).42 Parallel experiments using time-resolved UV−vis spectroscopy assisted identification of the contribution of the intermediates to the extracted IR difference spectra (Figure 3C). More recently, we have also used time-resolved QCL spectroscopy to study the carboxylic region with even higher temporal and spectral resolution.22,23 As a first step to understand the chemical changes encoded in the 1800−1690 cm−1 region, we applied Fourier selfdeconvolution (FSD), 58 a band-narrowing method that improves band separation and kinetic selectivity (Figure 3C).31 Thereby, several positive (1760, 1745, 1728, 1695 cm−1) and negative (1736 and 1717 cm−1) bands were resolved (Figure 3B,C). The negative bands at 1736 and 1717 cm−1 were previously assigned to Asp156 and Glu90, respectively, by static FT-IR difference spectroscopy.55,59 We performed complementary time-resolved FT-IR experiments on several variants with 339
DOI: 10.1021/acs.jpcb.6b09222 J. Phys. Chem. B 2017, 121, 335−350
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The Journal of Physical Chemistry B during the P2390 to P3520 transition. CrChR2 displays two seemingly separated proton transfer pathways (Figure 3D). The primary proton transfer pathway explains the proton pumping activity of CrChR2 as observed in electric recordings of CrChR2 in black lipid membranes.63 The secondary proton pathway is incompetent for proton pumping but colocalizes with the putative ion permeation pathway.57 An unexpected observation from low-temperature studies was that the CO stretch frequency of D156 was not down-shifted in D2O,64,65 suggesting a resistance to H/D exchange of the OH of the carboxylic group, seemingly inconsistent with its proposed role of D156 as the SB proton donor (Figure 3D). Yet, the CO stretch of D156 does shift, from 1736 to 1737 cm−1 in D2O at room temperature.23 The unusual 1 cm−1 upshift of the CO of D156 can be ascribed to its H-bond to C128. We expect the dipole moment of the SD group to be only half that of the S H group, as deduced from the 4 times smaller integrated extinction coefficient for the SD than for the SH stretching vibration.66,67 Upon deuteration of the thiol group of C128, its dipole moment is reduced, and as a consequence, the strength of its D-bond is weakened, with the resulting upshift of the CO stretching frequency of D156. This up-shift compensates the expected 8 cm−1 down-shift of the CO stretch upon deuteration of the hydroxyl group caused by vibrational uncoupling (Figure 3F).23 In the D156E variant, where E156 is too long to form a H-bond with C128, the CO frequency of E156 exhibits a normal down-shift in D2O by 8 cm−1 (from 1764 to 1756 cm−123) lending further support to the solvent accessibility of the residue at position 156. Ion Permeation and Hydration of Transmembrane Regions. Ion conductivity after retinal isomerization arises with t1/2 ≈ 200 μs, i.e., when the P2390 intermediate has already formed but prior to formation of the P3520 state.39,41 Conductivity decays with t1/2 ≈ 10 ms, simultaneously with the decay of the P3520 intermediate.39,41 What happens at the molecular level for ion conductivity to rise? Or in other terms, what does it take for the channel to open? The X-ray crystallographic structure of a CrChR1−CrChR2 chimera showed a pore intruding halfway into the membrane, framed by four helices: A, B, C, and G.57 Distance measurements by pulsed electron double resonance (PELDOR) spectroscopy on the cryo-trapped P3520 intermediate showed that helix B moved outward in the conductive state of the channel,68 presumably enlarging the half-pore in the dark state to form a continuous transmembrane pore. However, the static nature of the experiments did not allow for a conclusion regarding whether the structural changes in helix B occur simultaneously with the formation of the open state or if they precede the onset of ion permeation. A recent work featuring nonequilibrium MD simulations concluded that helix B tilts and forms a pore filled with water in less than 100 ns after retinal isomerization,60 3 orders of magnitude before the onset of ion permeation. We investigated which structural changes are temporally correlated to the onset of ion permeation.69 Two intense bands characteristic for α-helices at 1665 (−) and 1648 (+) cm−1 reach a maximum absorption change at 1 ms (Figure 4A), at the same time as the photocurrents of CrChR2 expressed in HEK cells.69 A cross-correlation analysis, quantitatively comparing the transients of the photocurrent and of the absorption changes in the IR, further indicates that ion permeation is coupled to a conformational change affecting mostly bands characteristic for helical structures (Figure 4B). The correlation of ion permeation and the spectral changes in the amide I region was corroborated
by experiments on fast and slow functional variants of CrChR2 (E123T and D156E), respectively.69 IR spectroscopic studies on peptides indicated that hydration of the carbonyl of the backbone amide down-shifts the amide I vibration.2 To check if the spectral changes correlating with ion permeation are caused by the hydration of helices, we set out on detecting vibrational coupling. Briefly, chemical groups with similar vibrational frequency can couple, most effectively through bonds but also through dipole−dipole interactions. Vibrational coupling is directly detected by 2D-IR spectroscopy experiments,4 but can also be indirectly detected in conventional linear IR spectroscopy by changes in the frequency of one group when the vibrational frequency of a second group is modified by isotope labeling. We recorded a steady-state IR difference spectrum of CrChR2 hydrated with either H216O or with H218O. Only amide I vibrations coupled to water molecules will be affected by exchanging H216O to H218O as a solvent. The frequency of the positive amide I band at 1648 cm−1 down-shifted by 2.5 cm−1 in H218O (Figure 4C, bottom left), whereas the negative band at 1665 cm−1 was not affected (Figure 4C, top left). The above experimental observations are consistent with the band at 1665 cm−1 reporting on the amide I vibration of transmembrane helices devoid of water, while the band at 1648 cm−1 corresponds to the amide I vibration of transmembrane helices where water is H-bonded to amide CO groups (Figure 4C, right). Therefore, ion permeation in ChR2 correlates with the hydration of transmembrane helices. Further details were obtained by lifetime distributions analysis by the maximum entropy method.70,71 Bands at 1665 (+) and 1648 (−) cm−1 indicated transient hydration of helices with time constants of 9 and 180 μs (Figure 4D). The rise of ion conductivity with τ ≈ 200 μs corresponded well with the second and final helix-hydration step (Figure 4E).69 Bands at 1665 (−) and 1648 (+) cm−1 indicated that the dehydration of helices occurs with τ = 10 ms (Figure 4D), simultaneously with the decay of the photocurrents.68,69,72 The influx of water to transmembrane regions to form a continuously water-filled pore is arguably the consequence of the outward tilt of helix B as detected by PELDOR experiments,68,72 reversed upon closing. The waterfilled pore provides a thermodynamically favorable pathway for the permeation of cations (Figure 4E), while ion selectivity will be dictated by the geometry and electrostatics of the formed pore.
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CRYPTOCHROME Flavin-containing proteins constitute a major group of biological photoreceptors. Among them, the cryptochromes are a large family of soluble receptors that regulate central processes in insects, plants, fungi, and bacteria such as the setting of the biological clock and plant growth.73 Time-resolved UV−vis spectroscopy has unraveled fundamental chemical steps in the photoreaction of cryptochromes in recent years. The flavin adenine dinucleotide (FAD) is photoreduced by an ultrafast electron transfer from tryptophan74,75 and then protonated on the time scale of a few microseconds.76,77 However, chemical reactions of nonaromatic amino acids or changes in the fold of the protein remain spectrally silent in the UV−vis region. Here, step-scan FT-IR spectroscopy provides the necessary sensitivity and time resolution. Sub-Microsecond Time Resolution to Resolve Proton Transfer. One central step in the photoreaction of plant cryptochrome is the protonation of the flavin anion radical FAD•−. Previous studies by steady-state infrared difference 340
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Figure 5. (A) Trigger scheme applied for obtaining a time resolution of 500 ns in the step-scan experiment with a given integration time of 5 μs for each data point. (B) IR difference spectrum of plant cryptochrome at 500 ns shows the presence of the flavin anion radical FAD•− (marker bands at 1621, 1518, and 1490 cm−1) as evidenced by comparison to the steady-state spectrum of FAD•− in the D396C mutant. (C) The intermediate CRYα is formed within 3.5 μs upon flavin protonation to FADH• (marker band at 1530 cm−1) and decays with an apparent lifetime of 290 μs. (D) Aspartic acid D396 is deprotonated (highlighted in orange at 1733 cm−1) concomitant with flavin protonation, thereby acting as the proton donor. Panels B and C adapted with permission from ref 89. Copyright 2015 American Chemical Society.
spectroscopy have allowed us to assign a band at 1733 cm−1 to the deprotonation of aspartic acid 396 (D396, according to the numbering of Arabidopsis cryptochrome 1),78−81 most likely acting as proton donor to flavin. Theoretical studies have challenged its connection to the formation of the flavin neutral radical FADH• because of the strict separation in time from the electron transfer.82,83 The identification of the proton donor by infrared spectroscopy is challenging for a process with τ = 1.7 μs. Only few studies on proteins have been performed with step-scan in the submicrosecond regime to date22,60,84−87 after the initial proof-ofprinciple on bacteriorhodopsin.88 One reason for this small number is that the low flux of the blackbody emitter leads to excessive noise in this early time region. Moreover, conventional spectrometers are limited in resolution to ∼2.5 μs (time interval of recording 0−5 μs). This limit in the dc mode can be overcome by separate, fast recording in the ac mode using a suitable set of electronics and data treatment (see above). We have instead increased the time resolution of the conventional spectrometers by precisely synchronizing externally the laser pulse with the recording interval of the spectrometer (Figure 5A). With such an approach, the time resolution is limited only by the rise time of the detector and the bandwidth of the ADC. It should be noted that these limitations
only affect the amplitude and therefore the sub-microsecond kinetics of the signal. The much more valuable information in the frequency domain is preserved. The resolution in the conventional dc mode is then improved up to 500 ns (time interval 0−1 μs), well into the sub-microsecond region. This approach necessitates only an external pulse generator for synchronization and is thereby easily accessible to other users. The resulting 500 ns spectrum of plant cryptochrome clearly shows major contributions from FAD•− such as the positive marker bands at 1621, 1518, and 1490 cm−1, in accordance with the sub-microsecond time resolution achieved (Figure 5B).89 More importantly, the band at 1733 cm−1 shows a very low amplitude, which increases strongly in the subsequent spectra at t > 1 μs (Figure 5C). This increase is observed concomitant with the formation of FADH• as evidenced by marker bands such as 1641 and 1530 cm−1.89 These findings confirm that deprotonation of D396 tallies flavin protonation to FADH• (Figure 5D). Changes in Secondary Structure Far from the Reaction Site. A question of general biophysical relevance is the transformation from a localized chemical reaction within the protein shell to a biochemical cascade in the cellular environment. A common scenario for photoreceptors is a change in the secondary and/or tertiary structure of the protein during the photoreaction. In cryptochromes, very little has been reported 341
DOI: 10.1021/acs.jpcb.6b09222 J. Phys. Chem. B 2017, 121, 335−350
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The Journal of Physical Chemistry B about such changes in the protein fold, which have been considered to be confined to the C-terminal extension outside of the sensory domain.90−92 Therefore, we have searched for structural changes in the photoreaction of the sensory domain of plant cryptochrome by inspecting the amide I and amide II region using step-scan and rapid-scan IR spectroscopy. To achieve a continuous data set from rapid- and step-scan time-resolved spectroscopy, we aimed for a maximal overlap in the early millisecond time range covered by both techniques. It should be noted that such overlap is limited for rapid-scan by the dead time of the experiment, the time lost between each change in direction of the moving mirror and the actual recording. We have minimized this dead time by exciting the sample well after the spectrometer indicated the start of the experiment. To this end, we synchronized the independent clocks of the spectrometer and laser with an external home-built device.89 We gained several milliseconds at the starting point of the experiment (from 8.6 ms to finally (4.3 ± 1.2) ms) and thereby optimized the overlap with the step-scan experiment. A demonstration of the actual limits of this approach is currently under way in our laboratory. The combined experiment finally covered six orders of time from 3.5 μs to 3.6 s (Figure 6A).89 An analysis of the data set ΔAij by a global fit routine was performed using a sequential model of r species represented by a sum of exponential functions. The least-squares fit included a weighting by a diagonal matrix containing the normalized inverse of the standard deviation of the noise wij in dependence of the wavenumber according to n
m
min{∑ ∑ wij[ΔAij − j=1 i=1
N
∑ ar (νĩ)exp(−krt j)]2 } r=1
The fit to the experiment produced three spectrally independent intermediates (CRYα, CRYβ, CRYγ) which all contained the same contributions by the FADH• albeit with different amplitudes. In contrast, the amide I and amide II region differed significantly between the intermediates evidencing changes in secondary structure on different time scales. The most significant finding was a pronounced negative band at 1632 cm−1 in CRYβ formed with τ = 500 μs (Figure 6B), which was assigned to a loss in β-sheet structure. Strikingly, the only β-sheet in the sensory domain is found in a distance of 25 Å to flavin (Figure 6C), which implies a signal transmission like a protein quake93 from the chromophore to the β-sheet. We postulate that this change in the fold of the sensory domain is key to triggering further functionally relevant structural changes within the Cterminal extension of plant cryptochrome.
Figure 6. (A) Global fit to the time-resolved spectra of plant cryptochrome covering the time range from 3.5 μs to 3.6 s. (B) Three intermediates are detected in this time range, which differ in the secondary structure of the domain. CRYβ is characterized by a loss in βsheet (highlighted at 1632 cm−1) and is formed with a 500 μs time constant from CRYα. (C) Structure of the sensory domain of plant cryptochrome (PDB 1U3D). The only β-sheet is found in the α/βsubdomain in a 25 Å distance to the photoactivated FADH•. Adapted with permission from ref 89. Copyright 2015 American Chemical Society.
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LOV PROTEIN A second large group of flavin-based photoreceptors besides the cryptochromes are the light-, oxygen-, and voltage-sensitive (LOV) proteins. In plants, algae, fungi, and bacteria, the LOV domain acts as a sensory module and is coupled with diverse output domains, rendering them sensitive to blue light.94 Among the LOV proteins studied so far, the aureochrome found in algae95 is exceptional, because it is coupled inversely to an Nterminal instead of C-terminal output domain, a DNA-binding basic region leucine zipper (bZIP) domain (Figure 7A). Moreover, the LOV domain undergoes light-dependent dimerization, which has already been implemented for optogenetic applications.96,97 Such dimerization has not been observed for the homologous LOV domains from plant phototropins, raising the question of the special molecular
mechanism of aureochrome-LOV and motivating an infrared spectroscopic investigation. Localization and Sequence of Structural Changes in LOV Proteins. We have investigated the role of two flanking helices, A′α and Jα (Figure 7B), found in the structures of aureochrome-LOV98−100 by applying the segment-resolved approach.13,101 The difference spectrum of Phaeodactylum tricornutum aureochrome1a A′α-LOV-Jα was resolved in direct comparison to those of A′α-LOV and LOV-Jα, which lack either one of the flanking helices. The calculation of the double 342
DOI: 10.1021/acs.jpcb.6b09222 J. Phys. Chem. B 2017, 121, 335−350
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The Journal of Physical Chemistry B
Figure 7. (A) Domain arrangement in the conventional LOV protein phototropin with a C-terminal effector domain (kinase) as opposed to the inversed arrangement in the aureochrome with an N-terminal bZIP domain. (B) Structure of the LOV domain of aureochrome1a binding the chromophore flavin mononucleotide (FMN) (PDB code 5A8B). The helices A′α and Jα flank the conserved LOV core domain, but do not come into direct contact. (C) Double difference spectrum of the light-induced difference of LOV with both flanking helices minus that of a Jα-deficient domain. Signals of the unfolding of both Jα and A′α are evident (1643 and 1656 cm−1, respectively). Therefore, both helices unfold in A′α-LOV-Jα but do not unfold in the Jαdeficient domain. (D) Double difference spectrum of the light-induced difference of LOV with both flanking helices minus that of an A′α-deficient domain. Only the unfolding of A′α is observed, which implies that, in LOV-Jα, the Jα helix can unfold even in the absence of A′α. This asymmetry in the response implies a temporal sequence, in which Jα needs to unfold prior to the response of A′α. Adapted with permission from refs 13 and 101. Copyright 2013 and 2015 American Chemical Society.
The question remained whether these findings on the LOV domain play a role for the function of the full-length receptor or if they are a consequence of the truncation. We demonstrated that Jα unfolding takes place in the full-length aureochrome as well, despite being located at the C-terminus of the receptor without a direct linkage to bZIP.12 More importantly, in the absence of the Jα helix, formation of the light state LOV dimer is inhibited, and thereby, the signal propagation to bZIP is lost. Activation of the LOV domains leads to changes in structure in bZIP,12,99,106 but the resulting rearrangement of the domains is still a matter of debate.12,99,107 The response of full-length aureochrome has even been investigated in the presence of DNA by IR difference spectroscopy. As a result, lysine residues were modified by the binding of DNA108 and an extension of helical elements with concomitant turn structural changes were detected after illumination.12 Interestingly, the latter response was highly specific for the DNA sequence, an effect that could not be resolved by DNA-binding studies with EMSA.
difference spectra revealed the (at least partial) unfolding of both helices upon illumination in A′α-LOV-Jα, as had been demonstrated before for phototropin-LOV (Figure 7C).102,103 However, our analysis revealed that, in LOV-Jα, unfolding occurred, whereas in A′α-LOV, no unfolding was observed (Figure 7C,D). Negative marker bands for the unfolding events were detected at 1643 cm−1 for the Jα helix similar to previous results on phototropin-LOV11,104,105 and at 1656 cm−1 for the A′α helix assigned here for the first time. This mutual dependence directly points to a time sequence of structural changes in the LOV domain. Only after the Jα helix unfolds can the A′α respond. Therefore, the sequence of events was clarified for aureochrome-LOV without applying any time-resolved experiment. Such asymmetry in the response implies an allosteric regulation, i.e., that unfolding of the C-terminal Jα regulates the unfolding of the N-terminal A′α. In conclusion, this allosteric regulation might allow for the strict light-dependent regulation of dimerization, because the A′α helix has been demonstrated to cover the dimerization site.99,101 In contrast, these events occur independently in other LOV domains and do not lead to dimerization. 343
DOI: 10.1021/acs.jpcb.6b09222 J. Phys. Chem. B 2017, 121, 335−350
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lished.109,110 This caveat is less of a problem using rapid-scan spectroscopy, where the best commercial FT-IR spectrometers reach time resolutions on the order of 10 ms. Yet, this time resolution is usually too poor to observe many relevant protein reactions. A novel ultrarapid scanning FT-IR spectrometer was recently introduced to overcome such limitations, and proof-ofprinciple IR difference spectra of bacteriorhodopsin were recorded with a time-resolution of 13 μs.111 An alternate approach to step-scan and ultrarapid-scan FT-IR is the application of tunable laser sources. The recent years have witnessed substantial progress in emission power and tunability of quantum cascade lasers (QCLs). For transient spectroscopy, continuous wave QCLs are preferred over pulsed sources. The high emission power (>200 mW) leads to paramount signal-tonoise ratio in time-resolved experiments.23 Although the range of tunability of many QCLs is commonly around 150−200 cm−1, a series of laser heads may be combined to further extend the spectral range to cover the mid-IR range. Transient spectroscopy using QCLs reaches a time resolution of 15 ns,23 making it attractive to complete and extend pump−probe spectroscopy studies which cover the time range from femtoseconds to a few nanoseconds. Thus, dynamics can be recorded over the entire “chemical” time range. Although the microsecond time range is also accessible in pump−probe spectroscopy by electronically synchronizing two femtosecond laser systems,112,113 the financial investment favors the cheaper QCL for such studies. The application of advanced time-resolved IR spectroscopies is mostly restricted to light-activated proteins such as those presented here. Evidently, other triggers are required for proteins activated by electron transfer or by a transmembrane potential. As electron transfer is critically distance-dependent and the transmembrane potential drops off very rapidly across the biomembrane, such experiments must be performed on the level of a single monolayer. The IR absorption of a monolayer is weak but can be enhanced by applying surface-enhanced IR absorption spectroscopy (SEIRAS).114−116 Varying the potential across a protein monolayer yielded redox-induced difference spectra of membrane proteins that are sensitive to such triggers, like the electron-driven respiratory protein cytochrome c oxidase.117 The influence of the membrane potential on the photoreaction of a membrane protein was elucidated on sensory rhodopsin II where it was demonstrated that a particular proton transfer reaction within SR II is halted by a membrane potential of opposite direction and sufficient field strength.118,119 It became evident from these experiments that the technique is applicable to the biomedically relevant family of voltage-gated ion channels. SEIRAS studies to resolve the functionally relevant structural changes in these ion channels are currently underway. Inasmuch as the past decade has witnessed an enormous progress in resolving the atomic structure of membrane proteins (see http://blanco.biomol.uci.edu/mpstruc/), the ways and means regarding how the 3D structure forms are far from being understood. The monolayer sensitivity of SEIRAS was exploited in a recent study of membrane protein folding during cell-free expression. Lipid nanodiscs were tethered to the plasmonic gold surface to provide a surrogate of the biomembrane, into which the nascent polypeptide inserts and folds after (or during) synthesis by the ribosome. Using SEIRAS, the folding pathway of bacteriorhodopsin via various secondary and tertiary structures was observed over time until the functional state of the protein structure was reached.120 Thus, a novel methodology has been introduced to trace the folding reaction in situ with the molecular resolution of vibrational
Figure 8. Artistic view of single molecule scanning near-field IR spectroscopy. The protein, bacteriorhodopsin (in purple), is attached to a gold (bow tie) nanoantenna. The tip (gray cone) scatters the IR light (from left), and the near-field absorption of the IR intensity by the protein is recorded in the optical far-field (to right).
Figure 9. Conceptual approach for investigating molecular prearrangements in voltage-gated ion channels on ultrafast time scales. Ultrashort THz pulses (red) are applied which provide an electric field strength comparable to the membrane potential Δψ of living cells.
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PERSPECTIVES We have showcased the power of mid-IR spectroscopy to contribute to the elucidation of the reaction mechanism of proteins. Even minute structural changes in a protein are resolved by IR spectroscopy while adding the sequence of events by performing time-resolved IR spectroscopy. Most of the experiments presented here rely on FT-IR spectroscopy which provides broad spectral information. High temporal resolution is achieved with the step-scan technique which requires, though, a strictly repetitive process to be studied. To overcome this limitation, a continuous sample exchange has been estab344
DOI: 10.1021/acs.jpcb.6b09222 J. Phys. Chem. B 2017, 121, 335−350
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The Journal of Physical Chemistry B Notes
spectroscopy. This pioneering methodology will be applied to other more complex membrane proteins to unravel different folding pathways. Originally, SEIRAS was performed using FT-IR spectroscopy, and time-resolved experiments on protein monolayers have been conducted using rapid-scan117,118,121,122 and step-scan techniques.123 Recent experiments demonstrated that femtosecond IR lasers can be used as well,124−127 opening an avenue toward ultrafast experiments on monolayers. The field of observation is drastically reduced from about 10 mm2 in typical SEIRAS experiments to 100 nm2 by applying scanning near-field infrared microscopy (SNIM).128,129 Here, the photonic field interacting with the molecules is efficiently condensed by the near-field interaction with a sharp tip that is used as a local scatterer of the incident radiation.130 A “chemical” image is generated by raster-scanning the surface while recording IR spectra at each point. The spatial resolution of SNIM is limited by the tip apex and can be
