Skip to main content
NCBI home page
ACS AuthorChoice logoLink to ACS AuthorChoice
. 2022 Nov 18;61(23):2698–2708. doi: 10.1021/acs.biochem.2c00483

Early Proton Transfer Reaction in a Primate Blue-Sensitive Visual Pigment

Yosuke Mizuno †, Kota Katayama †,‡,§,*, Hiroo Imai ∥, Hideki Kandori †,‡,*
PMCID: PMC9730847  PMID: 36399519

Abstract

graphic file with name bi2c00483_0008.jpg

The proton transfer reaction belongs to one of the key triggers for the functional expression of membrane proteins. Rod and cone opsins are light-sensitive G-protein-coupled receptors (GPCRs) that undergo the cis–trans isomerization of the retinal chromophore in response to light. The isomerization event initiates a conformational change in the opsin protein moiety, which propagates the downstream effector signaling. The final step of receptor activation is the deprotonation of the retinal Schiff base, a proton transfer reaction which has been believed to be identical among the cone opsins. Here, we report an unexpected proton transfer reaction occurring in the early photoreaction process of primate blue-sensitive pigment (MB). By using low-temperature UV–visible spectroscopy, we found that the Lumi intermediate of MB formed in transition from the BL intermediate shows an absorption maximum in the UV region, indicating the deprotonation of the retinal Schiff base. Comparison of the light-induced difference FTIR spectra of Batho, BL, and Lumi showed significant α-helical backbone C=O stretching and protonated carboxylate C=O stretching vibrations only in the Lumi intermediate. The transition from BL to Lumi thus involves dramatic changes in protein environment with a proton transfer reaction between the Schiff base and the counterion resulting in an absorption maximum in the UV region.

Rhodopsin is a light-sensitive G-protein-coupled receptor (GPCR) present in the rod photoreceptor cells of the retina.1,2 To date, rhodopsin has been extensively studied as the prototypical GPCR, and insights derived from its comprehensive spectroscopic and biophysical studies have significantly improved our understanding of the GPCR activation and signaling mechanism.3,4 One of the unusual features of rhodopsin, compared to other GPCRs, is its light-sensitive ligand, 11-cis-retinal. 11-cis-Retinal is covalently linked to the opsin protein (ε-amino group of Lys296 in the transmembrane helix7 (TM7)) via the protonated Schiff base.5 Another distinguishing feature of rhodopsin is its activation mechanism. The activation of rhodopsin involves two key chemical reactions: (i) retinal photoisomerization and (ii) proton transfer reaction from retinal Schiff base.

Retinal photoisomerization is the primary reaction in rhodopsin,6 which occurs within 200 fs after light absorption.6,7 The ultrafast photoisomerization results in the deposition of light energy in the distorted retinal conformation along its polyene chain8−11 and altered hydrogen bond network.12 This energy is then utilized for further structural changes in the protein moiety on slower time scales.12,13

Proton transfer from the retinal Schiff base is a well-characterized reaction that modulates the activation of rhodopsin. In the ground state of rhodopsin, the retinal Schiff base is protonated, with a glutamate side chain carboxylate serving as the counterion.14,15 Upon photoisomerization, the retinal Schiff base releases its proton to the counterion to form the signaling-competent Meta-II intermediate, with an >100 nm blue-shifted absorbance spectrum in the ultraviolet (UV) region.1 This indicates that the retinal Schiff base pKa is extraordinarily high in the ground state with 11-cis-retinal, but the pKa drops significantly in the Meta-II state with all-trans-retinal (pKa < 7).16,17 Unlike vertebrate rhodopsins, the drop in pKa is much smaller for invertebrates. In fact, as pKa is located at neutral pH, the active intermediates contain both protonated and deprotonated retinal Schiff base, which are often called acid-Meta and alkali-Meta, respectively.18,19

The photoreceptive GPCRs responsible for color vision, cone pigments, are assumed to undergo a similar photoactivation process as rhodopsin, since they share a common structural signature of seven transmembrane helices and 11-cis-retinal as the chromophore.20,21 However, variations exist between the pigments with respect to the lifetime of the photoactivated state, photosensitivity, and regeneration speed. Previous low-temperature time-resolved spectroscopic studies of chicken cone pigments show that the deprotonation of the retinal Schiff base only took place upon formation of Meta-II intermediate, and all other intermediates remained protonated.22−24 Difficulties in the isolation and preparation of cone pigments have hampered structural studies that could reveal the differences in their physicochemical properties.

To date, we have performed structural studies on primate cone pigments recombinantly expressed in a mammalian system using light-induced difference FTIR spectroscopy.25−30 Light-induced difference spectroscopic studies between Batho and ground states at 77 K revealed that primate blue (MB), green (MG), and red (MR) pigments, as well as rhodopsin, exhibit a common retinal structure and cis–trans isomerization, with a slightly different torsion pattern along the polyene chain.25,30 Furthermore, the Batho intermediate of all three cone pigments could be reverted back to the ground state by light, displaying a common photochromic property like rhodopsin. This photochromic property has been highly advantageous to improving the signal-to-noise ratio of FTIR spectra.25−30 This photochromic property has been hypothesized to be unique at 77 K. As a matter of fact, illumination of the Lumi intermediate (Lumi) of bovine rhodopsin, that is stable at 200 K, does not result in its reversal to the ground state due to the large structural changes in the opsin moiety.31 Therefore, difference FTIR spectroscopy is an effective method to study cone pigments; however, such studies have been limited to their Batho intermediate.

We recently reported that MB exhibits a photochromic property at 77 K as well as 163 K where the BL intermediate is in photoequilibrium with the ground state of MB.32 FTIR spectra of BL state clearly showed that the distorted retinal structure in Batho is relaxed during the transition to BL, which is accompanied by local structural relaxation. Thus, the structural features of both the retinal and local protein environment can be attributed to the photochromic property of MB at 163 K.

In this study, we present the structural changes in the Lumi state of MB. We show that Lumi is formed by light illumination at 223 K with an absorption maximum in the UV region, indicating deprotonation of the retinal Schiff base. Furthermore, this early proton transfer reaction during photoreaction is accompanied by large conformational changes in the protein moiety. In addition, we observed that Lumi can be photoconverted to ground state MB by light at 223 K. Thus, we conclude that MB exhibits photochromic properties in all its intermediates produced in the early photoreaction process, Batho, BL, and Lumi, strongly indicating the existence of MB-specific photoreaction dynamics. The unique structural changes in the retinal chromophore and the opsin moiety coupled with the proton transfer reaction and photochromic property are discussed on the basis of these data.

Materials and Methods

Sample Preparation

The cDNA of primate blue pigment (Uniprot ID: F7GQA6) was Rho1D4 tagged and cloned into the pFastBac HT expression vector (Bac-to-Bac, Thermo Fischer Scientific) using Infusion method. This construct was expressed in Sf9 cells that were cultured in ESF921 (Expression systems LLC) at 28 °C. After 3 days of incubation, the expressed cells were harvested by centrifugation and stored at −80 °C until further use. Cell pellets were resuspended in low-salt buffer (25 mM HEPES, 20 mM KCl, 10 mM MgCl2, pH 7.5) containing protease inhibitor (Sigma-Aldrich, Japan). The cells were disrupted twice using a Dounce homogenizer and centrifuged for 20 min. The disrupted cells were homogenized with high-salt buffer (25 mM HEPES, 20 mM KCl, 10 mM MgCl2, 1 M NaCl, pH 7.5) containing protease inhibitor (Sigma-Aldrich, Japan) and benzonase nuclease (Novagen) for increasing the solubilization efficiency to remove nucleotides, followed by centrifugation at 142 414g for 25 min. This process was repeated thrice.

The pellets were resuspended in Pm buffer (50 mM HEPES, 140 mM NaCl, pH 7.0), regenerated with 11-cis-retinal by stirring it overnight at 4 °C. The regenerated sample was solubilized with a buffer containing 1% (w/v) n-dodecyl-β-D-maltoside (DDM), 50 mM HEPES, 140 mM NaCl, pH 7.0. This mixture was centrifuged at 235 418g in a fixed-angle rotor for 40 min. The solubilized protein was mixed with 1D4 antibody conjugated resin, and the resin with bound MB was washed with buffer containing 50 mM HEPES, 140 mM NaCl, 0.02% DDM, pH 7.0, and then treated with buffer E (0.40 mg/mL 1D4 peptide, 50 mM HEPES, 140 mM NaCl, 0.03% DDM, pH 7.0).

Low-Temperature UV–Visible and FTIR Spectroscopy

Protein samples were reconstituted into phosphatidylcholine (PC) liposomes by using a protein:lipid molar ratio of 1:30 and dialyzed to remove DDM. The dialyzed sample was suspended in buffer containing 2 mM phosphate and 5 mM NaCl (pH 7.25). The resuspension was placed onto a BaF2 window and dried with an aspirator. Low-temperature UV–visible spectroscopy was performed on films hydrated with H2O at 223 K. Low-temperature FTIR spectroscopy was performed on films hydrated with H2O, D2O, or D218O at 223 K.

MB samples were illuminated with >430 nm light for 5 min at 223 K, followed by illumination with 360 nm light (by using an interference filter) for 5 min (UV–visible) and 10 min (FTIR) at 223 K. The former and latter illuminations convert MB to Lumi and Lumi to MB, respectively, from which Lumi minus MB difference FTIR spectra were obtained. To study the BL intermediate at 223 K, samples were light-activated with 380 nm light through an interference filter for 5 min at 223 K. This was followed by light activation with >500 nm light for 5 min at 223 K. For each FTIR measurement, a total of 128 interferograms were recorded and 40 H2O, 51 D2O, and 100 D218O recordings were averaged. FTIR spectra were recorded at 2 cm–1 resolution.

Results

Photochromism between MB and the Lumi Intermediate at 223 K

Figure S1 shows the UV–visible absorption spectra of before (black curve) and after illumination with 420 nm light (red curve) of MB at 223 K. By calculating the difference between black and red curves in Figure S1, the light-induced difference UV–visible spectra of MB were calculated as shown in Figure 1a. The two positive and one negative absorbances at 344, 469, and 406 nm correspond to photointermediates and the ground state of MB, respectively. Previously, light illumination at 163 K has been shown to form the BL intermediate with a red-shifted absorbance from the ground state.32 In fact, at 223 K, a red-shifted photointermediate with a peak at 466 nm was observed upon illumination by 380 nm light (Figure 1b, black solid curve). Subsequent illumination with >500 nm light recovered the ground state absorption spectrum, which is evident from the mirror-image difference spectra in Figure 1b (black dotted curve). The formation of the BL intermediate and its photochromic property at 223 K are fully identical with those at 163 K (Figure S2).

Figure 1.

Figure 1

Open in a new tab

(a) Light-induced difference UV–visible spectra at 223 K upon illumination at 420 nm. (b) Light-induced difference UV–visible spectra with 380 nm illumination (solid line) and with >500 nm illumination (dash line) showing the photoequilibrium between the ground and intermediate states (c) Light-induced difference UV–visible spectra with >430 nm illumination (solid line) and 360 nm illumination (dash line) showing photoequilibrium between the ground and intermediate states. Each gridline of the y-axis corresponds to 0.02 absorbance.

Unlike other early photointermediates such as Batho (77 K) and BL (163 K), another photointermediate formed by 420 nm light illumination in Figure 1a was blue-shifted to 344 nm. This result strongly suggests that the retinal Schiff base is deprotonated even at 223 K because of its absorption in the UV region. This is a surprising observation, as the proton transfer reaction never occurs at around 200 K for other visual animal rhodopsins. Unlike the observations with 420 nm illumination (Figure 1a), the light-induced difference UV–visible spectra of MB with >430 nm illumination show the accumulation of only blue-shifted photointermediate (Figure 1c, black curve). Subsequent illumination with 360 nm light reverted the blue-shifted photointermediate to the ground state. This can be clearly deduced as the solid and dotted curves are almost mirror images of each other. Bovine rhodopsin has been shown to convert to its Lumi intermediate upon illumination at 200 K with an absorption in the visible region (492 nm).33 Thus, the observed intermediate of MB with an absorption in the UV region formed at 223 K can be defined as the Lumi intermediate of MB. The present study showed that MB exhibits photochromic properties in all its intermediates produced in the early photoreaction process, Batho (77 K), BL (163 and 223 K), and Lumi (223 K).

Comparison of Light-Induced Difference FTIR Spectra of MB at 223, 163, and 77 K at the Chromophore Level

Next, we used low-temperature FTIR spectroscopy to determine the light-induced structural changes in the Lumi intermediate of MB, especially how the proton transfer reaction occurs at temperatures under 223 K. Figure S3 shows the light-induced difference FTIR spectra upon illumination with >430 nm (black curve) and 360 nm (red curve) light, respectively. Clearly, the spectral features look almost like mirror images, similar to what was observed in UV–visible spectra under the same illumination condition (Figure 1c). This suggests that the photochromic property of the Lumi intermediate of MB is observed as a light-induced conformational change in both the retinal chromophore and the protein moiety, which enabled us to obtain highly accurate spectra with a high signal-to-noise ratio. In fact, we observed significant peaks for the retinal chromophore at the C=C stretch frequency region (1600–1550 cm–1), C–C stretch (1250–1150 cm–1), hydrogen out-of-plane (HOOP) vibrations (1000–800 cm–1), and C=O stretch vibrations (1750–1600 cm–1) of the peptide carbonyl and protonated carboxylate as shown in Figure S3. On the other hand, low-temperature UV–visible spectroscopy revealed a red-shifted photoproduct at 223 K by illumination with 380 nm light in Figure 1b. Since this photoproduct has the same absorption maximum as the BL intermediate formed at 163 K in Figure S2, we assigned it to be the BL intermediate. In fact, the BL intermediate formed at 223 K also exhibited photochromic properties, reverting to the ground state MB upon light illumination. Then, we compared the difference FTIR spectra between the BL intermediates formed at 163 and 223 K to elucidate the structural differences between them (Figure S3). The C–C stretching vibrations and HOOP bands originating from the retinal chromophore are identical between them, indicating that their retinal structure is the same. In contrast, vibrational band differences were observed in the 1700–1500 cm–1 region corresponding to conformational changes in the protein moiety. The pair bands at 1641 (−)/1633 (+) cm–1 could be originated from amide-I C=O stretching vibration, and a positive 1559 cm–1 band could be originated from amide-II band. These differences in the vibrational bands may reflect differences in the local conformation of the BL intermediates formed at 163 and 223 K. However, from Figure S2, there is no difference in the visible absorption spectra between them, so we infer that those local structural differences do not affect the electronic state of the retinal. Therefore, as follows, the details of the structural changes of the Lumi intermediate formed by couples with proton transfer will be discussed in comparison with the Batho (77 K) and BL intermediates (163 K).

Figure 2 shows a comparison of light-induced difference FTIR spectra of MB at 223, 163, and 77 K measured in H2O (black curve) and D2O (red curve). It is well-known that the vibrational amplitudes of the retinal chromophore are diminished by weakening of IR absorption. IR absorption is diminished as a result of the retinal Schiff base deprotonation, while it is enhanced by the Schiff base protonation.34 Based on this, most of the bands in the 1300–800 cm–1 region, where C–C stretching and HOOP vibrations of retinal chromophore appear, are observed only on the negative side in the spectra of the Lumi intermediate. The negative peaks at 1229 and 1172 cm–1 at 223 K are commonly observed at 77 and 163 K, which would originate from C12–C13 stretching and C8–C9, C12–C13 group stretching vibrations, respectively, in the ground state MB as previously reported in the chicken red pigment.35 A fewer number of positive peaks is consistent with the deprotonation of the retinal Schiff base. The retinal chromophore exhibits HOOP wagging modes in the 1000–800 cm–1 frequency region, which reflect a structurally perturbed and/or distorted chromophore. At 223 K, the HOOP band is barely observed in the spectra, and this spectral feature is similar to that at 163 K. In general, the retinal structure is highly perturbed upon the formation of Batho intermediate by the cis–trans isomerization, resulting in many HOOP bands being observed at 77 K. Thus, the disappearance of the HOOP band at 163 and 223 K suggests that retinal relaxation took place with the formation of the BL and Lumi intermediates.

Figure 2.

Figure 2

Open in a new tab

Light-minus-dark difference FTIR spectra of MB at 223, 163, and 77 K in the 1800–800 cm–1 region. Spectra were measured in H2O (black) and D2O (red). The spectra at 163 and 77 K were reproduced from refs (32) and (30), respectively. While the ground state MB generates the negative bands, positive bands originate from the Lumi, BL, and Batho at 223, 163, and 77 K, respectively. The obtained spectra of Lumi, BL, and Batho are scaled by 5.4, 1.0, and 2.7, respectively. Each gridline of the y-axis corresponds to 0.004 absorbance units.

Although both the BL and Lumi intermediates were expected to undergo retinal relaxation, differences were observed in their retinal structure. The negative sharp 961 cm–1 band observed at 77 K corresponds to 967 cm–1 in bovine rhodopsin, which is attributed to the C11=C12 HOOP mode. Although, with reduced peak intensity, this band is also observed at 163 K. On the other hand, at 223 K, a band was observed at 972 cm–1 along with the disappearance of the 961 cm–1 band. Previous FTIR spectroscopy study has reported that two negative bands at 974 and 960 cm–1 were observed in chicken red pigment, and both bands were assigned to the C11=C12 HOOP mode, suggesting that the C11=C12 position is more distorted in chicken red in comparison to rhodopsin, which has only one band at 966 cm–1.35 Similar peak features are also observed in MG and MR.25,30 Taken together, these results indicate that the C11=C12 position is further relaxed during the transition from the BL to Lumi intermediates. In addition, two positive bands are observed at 964 and 957 cm–1 in the BL intermediate, but the peak intensities of these bands are considerably reduced in the Lumi intermediate. This results also indicate the relaxation of retinal polyene chain upon the formation of Lumi.

The 1600–1500 cm–1 frequency region exhibits the C=C stretching vibrations of the retinal chromophore. Unexpectedly, at 223 K, no C=C stretching vibration is observed on the negative side. In case of bovine rhodopsin, a strong negative C=C stretching vibration is observed in the difference FTIR spectra of the active Meta-II intermediate due to deprotonation of the retinal Schiff base,36 but this is not the case for Lumi intermediate of MB. This is presumably because the amide-II band, which is a mixture of a C–N stretch and H–N–C bend mode of the protein backbone, overlaps this frequency region and compensates for the retinal C=C stretching vibration.

Comparative Analyses of Light-Induced Difference FTIR Spectra of MB at 223, 163, and 77 K at the Protein Level

Figure 3a shows the difference FTIR spectra in the 1800–1600 cm–1 region that mainly monitor vibrations in the protein moiety. These vibrations include the C=O stretch of protonated carboxylate and peptide backbone of α-helix (amide-I band) at 1800–1700 and 1700–1600 cm–1, respectively. As is clearly seen in the difference FTIR spectra at 223 K (black curve), a prominent positive band was observed at 1714 cm–1. Furthermore, this band is slightly down-shifted by 2 cm–1 in the D2O spectra (red curve), suggesting a C=O stretching vibration of the protonated carboxylate. Note that the positive 1713 cm–1 band observed in the Meta-II spectra of bovine rhodopsin has been attributed to the protonated C=O stretching vibration of the counterion, Glu113.36,37 Therefore, since the 1714 cm–1 band observed in the Lumi of MB measured at 223 K appeared in a similar frequency region, this band is considered to originate from the counterion as well. Amino acids within 5 Å from the retinal chromophore are represented in Figure 4. MB has two carboxylic acids in the vicinity of the retinal chromophore, Glu113 and Glu181, respectively. The former plays role as the counterion for vertebrate rhodopsin and the latter for invertebrate rhodopsin.39 Since MB belongs to vertebrate rhodopsin, it is reasonable to assume that its counterion, Glu113, receives a proton directly from the retinal Schiff base. However, considering that the counterion switches from Glu113 to Glu181 during Meta-I formation in bovine rhodopsin (counterion switch model)40,41 and that MB shows photochromism like bistable invertebrate rhodopsin with photochromism,42 the proton acceptor in Lumi might be Glu181. In fact, we infer that Glu181 is deprotonated because none of the C=O stretching vibration of the protonated carboxylate was observed in the >1700 cm–1 region for both 77 and 163 K spectra in MB.32 Thus, based on these results, both Glu113 and Glu181 are potential candidates for being the proton acceptor that leads to the origin of the positive 1714 cm–1 band in the Lumi intermediate.

Figure 3.

Figure 3

Open in a new tab

(a) 1780–1600 cm–1 region spectra from Figure 2. Spectra were measured in H2O (black curve) and D2O (red curve), respectively. (b) Light-minus-dark difference FTIR spectra of MB hydrated with D2O at 223, 163, and 77 K in the 3590–3430 cm–1 region. The spectra at 163 and 77 K were reproduced from refs (32) and (30), respectively. Each gridline of the y-axis corresponds to 0.003 (a) and 0.001 (b) absorbance units, respectively.

Figure 4.

Figure 4

Open in a new tab

Amino acid residues surrounding the retinal chromophore in bovine rhodopsin (<5 Å, Protein Data Bank entry 1U19(38)) and the corresponding residues in MB, MG, and MR protein models.

For MB, the formation of the primary intermediate Batho has been characterized by a large conformational change in the α-helix, which corresponds to 1658 (−)/1651 (+) cm–1 pair amide-I bands in the spectra at 77 K. On the other hand, the amide-I band in the spectra at 163 K decreased dramatically with the transition into the BL intermediate. This probably suggests that the α-helical structure changes upon the formation of BL intermediate along with the relaxation of C11=C12 torsion of the retinal chromophore. Interestingly, an amide-I band corresponding to a large α-helical conformational change was again observed at 1658 (−)/1652 (+) cm–1 upon the formation of the Lumi intermediate. Although the spectral down-shift in the amide-I band at 223 K seems to be similar to what was observed in the spectra at 77 K, the amide-I band intensity at 223 K is much higher than at 77 K. Notably, the spectral down-shift of the amide-I pair band appearing at 1656 (−)/1644 (+) cm–1 is a hallmark in the Meta-II spectra of bovine rhodopsin.36,43 It has been suggested that this α-helical conformational change in rhodopsin corresponds to an outward tilt in TM6 that is coupled with the transfer of proton of the retinal Schiff base to the counterion.44,45 On the other hand, the crystal structure of Lumi rhodopsin shows a flipping motion of the β-ionone ring of the retinal chromophore.46 Local perturbation of amino acid side chains in the retinal-binding pocket is also increased in Lumi as compared with Batho, but global structural changes in the protein moiety are still limited. This suggest that only the retinal-binding pocket is affected and not the cytoplasmic surface of opsin. Thus, the conformational change of the Lumi intermediate of MB, while combining aspects of both the Lumi and Meta-II intermediates of rhodopsin, exhibits a unique state in which the proton transfer reaction from the retinal Schiff base to Glu113 or Glu181 occurs early during the photoreaction.

In rhodopsin, rearrangement of interhelical hydrogen bond networks plays a crucial role in the large α-helical conformational change that occurs during the transition to the Meta-II intermediate. Previous X-ray crystallographic47 and FTIR37,48 studies suggest that two carboxylates (Asp83 in TM2 and Glu122 in TM3) and water molecules reorganize key internal hydrogen bond networks which are in the middle of the TM region, connecting the retinal-binding extracellular side to the G-protein binding area. In contrast, Asp83 and Glu122 are not conserved in MB, suggesting a different mechanism for the large conformational change in the α-helix.

An intrahelical hydrogen bond network is another key factor that plays a critical role in the activation of rhodopsin. Thr118 located in TM3 of rhodopsin forms a hydrogen bond with the backbone oxygen of Gly114. Gly114 is located right before TM3 and interacts with the C9 methyl group of the chromophore.47 This indirect interaction between Thr118 and the C9 methyl group of the chromophore plays a crucial role in rhodopsin activation.49 Previous FTIR study has showed that the intense pair bands at 3488 (+)/3464 (−) cm–1 in bovine rhodopsin originate from the O–H stretching vibration of Thr118,50 which is only deuterated at high temperatures by the effect of protein fluctuations.51 Thus, the O–H stretching vibration of Thr118 can be used for monitoring the local environmental changes in the protein moiety. In the case of MB, 3552 (+)/3468 (−) cm–1 at 77 K and 3513 (−)/3479 (+) cm–1 at 163 K would originate from the O–H stretching vibration of Thr118 (Figure 3b). Interestingly, the change in the hydrogen bond strength of Thr118 exhibits an opposite direction between Batho and BL intermediates. Thus, during Batho formation, the Thr118 hydrogen bond strength weakens due to the up-shift of O–H stretching vibration, whereas the hydrogen bond strengthens due to the down-shift of the corresponding vibrational band with the formation of the BL intermediate. This hydrogen bond strength change is probably the result of perturbation of the protein environment mainly on TM3 where Thr118 is located upon formation of Batho intermediate. The partial relaxation of TM3 upon transition to the BL intermediate would allow Thr118 to reform hydrogen bonds. Our results show that the O–H stretching vibration in spectra of the Lumi intermediate, which is possibly derived from Thr118, completely disappeared. This result could be explained by two possibilities. One possibility is that the hydrogen bond strength of Thr118 is identical between the ground state and Lumi intermediate, and this would result in further relaxation changes due to the large conformational change in the α-helix that couples with the deprotonation of the retinal Schiff base. Another possibility is that the hydrophobic environment around Thr118 is altered into a hydrophilic environment because of the large α-helical conformational change, resulting in hydrogen/deuterium exchange.

S–H stretching vibration of the cysteine residue, whose stretching vibration reflects its protein environment, also serves as an exceptional vibrational probe because the frequency region where S–H stretching vibration appears is well-isolated from vibrations of other functional groups inside the protein.52,53 Pair bands at 2554 (+)/2640 (−) cm–1 and 2556 (+)/2537 (−) cm–1 were observed at 163 and 223 K, respectively, which could be originated from the S–H stretching vibration of cysteine, though no similar band was observed at 77 K in Figure 5a. Both pair bands were up-shifted upon light illumination, indicating that the hydrogen bonding of cysteine weakened with the formation of the BL and Lumi intermediates, respectively. In particular, the positive band at 223 K showed a 2 cm–1 up-shift with increased band intensity as compared to that at 163 K. Furthermore, judging from the clear down-shift of both pair bands to 1856 (+)/1845 (−) cm–1 and 1857 (+)/1844 (−) cm–1 in D2O (Figure 5b), the observed cysteine is expected to be in a more hydrophilic protein environment. There are a total of 13 cysteine residues in MB, of which only two, Cys187 and Cys211, are placed within 5 Å from the retinal chromophore. Among the two, Cys187 is an unlikely candidate because it forms a disulfide bond with Cys110 at the end of TM2. Interestingly, Cys211 corresponds to His211 in bovine rhodopsin. Previously, NMR spectroscopy of rhodopsin has shown that His211 on TM5 and Glu122 on TM3 form an interhelical hydrogen bond in the ground state. However, the hydrogen bond between His211 and Glu122 is reorganized during Meta-II formation.37 Thus, assuming that the observed S–H stretching vibration of cysteine is originated from Cys211, it suggests that interhelical hydrogen bond alteration occurs in MB in the early photoreaction intermediates such as BL and Lumi. This contrasts with rhodopsin where interhelical hydrogen bond alteration is observed in the late intermediate. While the hydrogen bond acceptors of S–H group were not identified in this study, these are likely to be the backbone carbonyl oxygen situated on TM3 or the internal water molecules.

Figure 5.

Figure 5

Open in a new tab

Light-minus-dark difference FTIR spectra of MB measured at 223, 163, and 77 K in the 2580–2521 cm–1 (a) and 1880–1810 cm–1 (b) regions. Spectra were measured in H2O (black curve) (a) and D2O (red curve) (b). The spectra at 163 and 77 K were reproduced from refs (32) and (30), respectively. Each gridline of the y-axis corresponds to 0.005 absorbance units. These frequency regions are typical S–H (a) and S–D (b) stretching vibrational mode regions.

Figure 6 shows the X–D stretching vibration region in D2O that comprises the structural information on the water hydrogen bonding network. As reported previously and identified by the isotope effect of 18O, MB and its Batho intermediate possess three positive and four negative water O–D stretching vibrations (Figure 6, green). Among these frequencies, bands at 2567 (−) cm–1 and 2527 (+) cm–1 correspond to deuterated tetrahedral ice suggesting that these frequencies originate from the water cluster. During the transition from Batho to the BL intermediate at 163 K, these water vibrational signals are altered with frequency shift and reducing band intensity, indicating that the formation of the BL intermediate is accompanied by the rearrangement of water molecules in the cluster. Here, difference spectra of Lumi intermediate at 223 K clearly showed a lack of a water O–D stretching vibration originating from the water cluster at 2600–2500 cm–1. In contrast, four negative and three positive water O–D stretching vibrations appeared distributed at 2698 (−), 2667 (−), 2616 (−), 2590 (−) cm–1 and 2682 (+), 2641 (+), 2600 (+) cm–1, respectively. The disappearance of water clusters can be interpreted as the water clusters perturbed by the BL intermediate reassemble with the formation of the Lumi intermediate to form water clusters that are identical to the ground state. On the other hand, among the distributed water O–D stretching vibrational bands in the 2700–2600 cm–1 region, the 2667 (−), 2641 (+), 2616 (−), 2600 (+), and 2590 (−) cm–1 bands would correspond to 2674 (−), 2652 (+), 2633 (−), 2608 (+), and 2567 (−) cm–1 bands in the 77 K spectrum (2567 cm–1 band also shares frequency with the water cluster) and 2674 (−), 2633 (−), 2606 (+), and 2590 (−) cm–1 bands at 163 K, respectively. In contrast, 2698 (−)/2682 (+) cm–1 pair bands would be the newly observed water O–D stretching vibrational change. This indicates that a reorganization of the water hydrogen bonding network occurred due to a major conformational change of the α-helix in the Lumi intermediate.

Figure 6.

Figure 6

Open in a new tab

Light-minus-dark difference FTIR spectra of MB in the 2800–1800 cm–1 region measured at 223, 163, and 77 K. Spectra were measured in D2O (red curve) and D218O (blue curve). Frequencies labeled in green correspond to those identified as water O–D stretching vibrations. The spectra at 163 and 77 K were reproduced from refs (32) and (30), respectively. The gray curve in the 2700–2000 cm–1 region represents O–D stretching vibrations of D2O at room temperature. The scaling of obtained spectra of Lumi, BL, and Batho are the same with Figure 2. Each gridline of the y-axis corresponds to 0.001 absorbance units. The typical frequency of the CO2 vibrational mode is shown by the thick bar at the top.

The 2576 (−)/2555 (+) cm–1 pair band in the Lumi intermediate shows no isotopic effect due to 18O, so this pair band could be an O–H or N–H stretching vibrational band of a hydrophilic amino acid. Harmonic oscillator calculations suggest that this band has been isotopically shifted from 3540 (−)/3511 (+) cm–1 by D2O hydration. In this frequency region, an O–H stretching vibration, which can be originated from Thr118, was observed in the 77 and 163 K spectra but was missing at 223 K (Figure 3b). This result strongly suggests that Thr118 is hydrogen/deuterium exchanged at 223 K due to a large conformational change associated with the formation of the Lumi intermediate.

Discussion

In this study, we found that the Lumi intermediate is formed at 223 K, coupled with the deprotonation of the retinal Schiff base specifically in MB. The UV–vis difference spectra of the ground state MB illuminated with >430 nm light at 223 K revealed an intermediate state with absorption at 349 nm. In bovine rhodopsin, the Lumi intermediate is formed at 200 K with absorption at 492 nm, but the retinal Schiff base remains protonated.33 Similarly, the Lumi intermediates of cone opsins are formed at 163–233, 153–223, and 193–233 K for chicken blue,24 green,22 and red pigments,23 respectively, with absorption peaks in the visible region. Based on these results, the intermediate with absorption in the UV region at 223 K is considered to correspond to the Lumi intermediate of MB. Furthermore, like the BL intermediate, the Lumi intermediate also exhibited photochromic properties. These results strongly indicate that the protein environment surrounding the retinal chromophore of MB is highly specific as compared with other animal rhodopsins.

We performed FTIR spectroscopy analysis to explore the structural elements that enable the deprotonation reaction of the retinal Schiff base and photochromic properties associated with the formation of the Lumi intermediate. As a result, we found structural differences specific to MB that could provide clues into its atypical behavior (Figure 7a). First, the retinal structure in the Lumi intermediate was relaxed similarly to the BL intermediate (Figure 7a; upper panel). The relaxation of the retinal structure upon the formation of the Lumi intermediate has also been observed in bovine rhodopsin (Figure 7a; under panel).46 On the other hand, previous low-temperature FTIR spectroscopy has reported that the Lumi intermediate of bovine rhodopsin formed at 200 K could only be photoconverted to rhodopsin at 77 K. It could be argued that after a small local rearrangement in the retinal-binding pocket of rhodopsin at 200 K, cooling of the Lumi intermediate from 200 to 77 K displaces the α-helical peptide backbone, which results in the reisomerization of an all-trans to an 11-cis retinal chromophore without a global change in the protein structure. In contrast, the Lumi intermediate of MB formed at 223 K induces a large conformational change in the α-helix. Nevertheless, the fact that photoconversion to MB was observed at the same temperature strongly suggests that the protein environment actively assisted in the reisomerization of retinal and the presence of a specific mechanism for reverting photoreaction in the protein moiety.

Figure 7.

Figure 7

Open in a new tab

Schematic model showing the early photoreaction of MB (a; upper panel) and its photoreaction scheme (b; upper panel) based on this study in comparison with bovine rhodopsin (a,b; under panel). In the Batho intermediate, all-trans-retinal is highly distorted especially at the C11=C12 portion by cis–trans photoisomerization. A water cluster constituted by at least three water molecules is situated along the polyene chain on the retinal chromophore. During the transition from Batho to BL, all-trans-retinal is relaxed along the polyene chain, which is accompanied by relaxation of the protein moiety around the retinal chromophore. Consequently, the water cluster is slightly disrupted, resulting in the partial distribution of water molecules. After formation of the Lumi intermediate from BL, further relaxation of both the retinal chromophore and protein moieties (mainly at TM3 and TM5) occur with the flipping of the β-ionone ring as observed in the crystal structure of Lumi in bovine rhodopsin.45 This results in the deprotonation of the retinal Schiff base. The water cluster is completely disrupted, and the water molecules are distributed within the retinal-binding pocket. The proton acceptor from the retinal Schiff base can be either Glu113 or Glu181.

Second, the rearrangement of an interhelical/intrahelical hydrogen bond network including protein-bound water molecules occurs during the formation of Lumi intermediate. The pair bands possibly originated from the Thr118 O–H stretch could be affected by hydrogen/deuterium exchange in the transition from BL to Lumi, strongly indicative of TM3 distortion. This was not observed in rhodopsin (Figure S5).54 Especially in early intermediates such as Batho and BL, Thr118 was embedded in a hydrophobic environment and was D2O unexchangeable. The fact that it became D2O exchangeable in Lumi can be explained by the large structural change of the α-helix with the alteration of the protein-bound water molecules. In addition, the intensity of S–H stretching vibration, which is thought to be originated from Cys211 on TM5, was slightly enhanced with the transition from BL to Lumi. In rhodopsin, the corresponding amino acid is His211, and its interaction with Glu122 on TM3 is caused by Meta-II formation. Thus, it is possible that the helical distortion of TM3 is propagated to TM5 in Lumi of MB, resulting in the local perturbation of Cys211. As a result of the movement of TM3 associated with the formation of the Lumi intermediate, the distance between the counterion Glu113 and the retinal Schiff base increased, which lowered the pKa value of the retinal Schiff base and may have resulted in deprotonation (Figure 7a).

Based on our comprehensive spectroscopy results, here we propose an early photoreaction scheme of MB (Figure 7b). MB transitions to a red-shifted Batho intermediate with a distorted all-trans-retinal by the cis–trans isomerization of the retinal chromophore upon blue light absorption. Subsequent local conformational relaxation of the retinal and protein moiety induces the formation of the BL intermediate with a slightly blue-shifted absorption maximum from Batho. Then, further retinal relaxation and a large α-helical conformational change resulting from deprotonation of the retinal Schiff base lead to the formation of a Lumi intermediate with absorption in the UV region. The BL and Lumi intermediates are in equilibrium.

In this study, the deprotonation of the retinal Schiff base during the early photoreaction process of MB, i.e., the characterization that the pKa of retinal Schiff base tends to drop, might be closely related to the evolution of cone pigments. Cone pigments are classified into four classes based on their primary sequence and spectral properties (λmax): S (SWS1, 355–450 nm), M1 (SWS2, 400–490 nm), M2 (Rh2, 450–535 nm), and L (LWS, 490–570 nm).55,56 Among these, SWS1 is the only class of cone pigments that has a deprotonated retinal Schiff base in its ground state. In a previous study, transient absorption spectroscopy of Siberian hamster ultraviolet cone pigment (SHUV) had revealed a photointermediate with absorption in the visible region due to the protonation of the retinal Schiff base in the early photoreaction process.57 Similar findings have been also observed for mouse ultraviolet cone pigment by using low-temperature UV–visible spectroscopy.58 These would be commonly achieved by controlling the pKa of the retinal Schiff base through the protein environment. Interestingly, MB has a protonated retinal Schiff base, but evolutionarily, it belongs to the SWS1 group. Although the protonation state of the retinal Schiff base is different in the ground state of SHUV/mouse UV and MB, there might be a common molecular mechanism controlling the pKa of the retinal Schiff base. The structural changes in the Lumi intermediate of MB observed in this study may contribute to the elucidation of this molecular mechanism. Further structural analysis of MB mutants and UV-sensitive cone pigments will provide insights into the subtle differences in the photochemical processes of cone pigments.

Conclusion

We have found that at 223 K, the Lumi intermediate of MB with an absorption maximum in the UV region was formed, which is strongly indicative of the deprotonated retinal Schiff base. Comparison of the light-induced difference FTIR spectra of Batho, BL, and Lumi intermediates shows that the HOOP and fingerprint bands between BL and Lumi are very similar. Thus, the retinal chromophore remains in a relaxed all-trans configuration in both Lumi and BL intermediates. However, significant α-helical backbone C=O stretching and protonated carboxylate C=O stretching vibrations were observed only in the Lumi intermediate, which are indicative of very different photoreaction dynamics. The transition from BL to Lumi involves dramatic changes in the protein environment during the proton transfer reaction between the retinal Schiff base and the counterion, thus resulting in an absorption maximum in the UV region. These Lumi-specific structural changes are accompanied by the alteration of interhelical/intrahelical hydrogen bond network along with protein-bound waters. This study characterizes the unique photoreaction dynamics that occurs in the early photointermediate in MB. Future work such as mutational and structural studies will focus on the photoreaction of the Meta-I intermediate to explore how the early proton transfer reaction between the retinal Schiff base and the counterion leads to the formation of the next active intermediate, Meta-II.

Acknowledgments

We thank the Kandori laboratory for helpful comments on this manuscript. We thank Dr. S Gulati (Gatan Inc.) for helpful comments on this manuscript. This research was financially supported by grants from the Japanese Ministry of Education, Culture, Sports, Science and Technology to K.K. (18K14662), H.I. (22H02674), and H.K. (21H04969), Grant-in-Aid for Scientific Research on Innovative Areas “Non-equilibrium-state molecular movies and their applications (Molecular Movies)” from MEXT, Japan (KAKENHI grant Nos. 20H05440 to K.K.), JSPS Joint Research Project (120229921) to H.I., and from Japan Science and Technology Agency (JST), PRESTO to K.K. (JPMJPR19G4).

Supporting Information Available

The Supporting Information is available free of charge at https://pubs.acs.org/doi/10.1021/acs.biochem.2c00483.

  • UV–visible absorption spectra of MB at 223 K (Figure S1), light-induced difference UV–visible spectra at 223 and 163 K in MB (Figure S2), light-induced difference FTIR spectra of MB at 223 K (Figure S3), light-induced difference FTIR spectra of MB at 223 and 163 K (Figure S4), and light-induced difference FTIR spectra of bovine rhodopsin in the Batho, Lumi, and Meta-I intermediates (Figure S5) (PDF)

Accession Codes

Uniprot: F7GQA6.

Author Contributions

K.K. and H.K. conceived and designed the experiments. Y.M. prepared samples and conducted all experiments with the help from K.K. Y.M. analyzed the data with the help of K.K. K.K. and H.K. wrote the manuscript. K.K., H.I., and H.K. coordinated and supervised the research. All authors have discussed the results and commented on the manuscript.

The authors declare no competing financial interest.

Supplementary Material

bi2c00483_si_001.pdf (252.3KB, pdf)

References

  1. Wald G. Molecular basis of visual excitation. Nature 1968, 219, 800–807. 10.1038/219800a0. [DOI] [PubMed] [Google Scholar]
  2. Palczewski K. Chemistry and biology of the initial steps in vision: The Friedenwald lecture. Invest. Opthalmol. Vis. Sci. 2014, 55, 6651–6672. 10.1167/iovs.14-15502. [DOI] [PMC free article] [PubMed] [Google Scholar]
  3. Palczewski K. G protein-coupled receptor rhodopsin. Annu. Rev. Biochem. 2006, 75, 743–767. 10.1146/annurev.biochem.75.103004.142743. [DOI] [PMC free article] [PubMed] [Google Scholar]
  4. Ernst O. P.; Lodowski D. T.; Elstner M.; Hegemann P.; Brown L. S.; Kandori H. Microbial and animal rhodopsins: structures, functions, and molecular mechanisms. Chem. Rev. 2014, 114, 126–163. 10.1021/cr4003769. [DOI] [PMC free article] [PubMed] [Google Scholar]
  5. Nakanishi K. Why 11-cis-retinal?. Am. Zool. 1991, 31, 479–489. 10.1093/icb/31.3.479. [DOI] [Google Scholar]
  6. Schoenlein R. W.; Peteanu L. A.; Mathies R. A.; Shank C. V. The first step in vision: Femtosecond isomerization of rhodopsin. Science 1991, 254, 412–415. 10.1126/science.1925597. [DOI] [PubMed] [Google Scholar]
  7. Peteanu L. A.; Schoenlein R. W.; Wang Q.; Mathies R. A.; Shank C. V. The first step in vision occurs in femtoseconds: complete blue and red spectral studies. Proc. Natl. Acad. Sci. U. S. A. 1993, 90, 11762–11766. 10.1073/pnas.90.24.11762. [DOI] [PMC free article] [PubMed] [Google Scholar]
  8. Kim J. E.; McCamant D. W.; Zhu L.; Mathies R. A. Resonance Raman structural evidence that the cis-to-trans isomerization in rhodopsin occurs in femtoseconds. J. Phys. Chem. B 2001, 105, 1240–1249. 10.1021/jp001236s. [DOI] [PMC free article] [PubMed] [Google Scholar]
  9. Kandori H.; Shichida Y.; Yoshizawa T. Resonance Raman structural evidence that the cis-to-trans isomerization in rhodopsin occurs in femtoseconds. Biophys. J. 1989, 56, 453–457. 10.1016/S0006-3495(89)82692-X. [DOI] [PMC free article] [PubMed] [Google Scholar]
  10. Palings I.; Pardoen J. A.; van den Berg E.; Winkel C.; Lugtenburg J.; Mathies R. A. Assignment of fingerprint vibrations in the resonance Raman spectra of rhodopsin, isorhodopsin and bathorhodopsin: implications for the chromophore structure and environment. Biochemistry 1987, 26, 2544–2556. 10.1021/bi00383a021. [DOI] [PubMed] [Google Scholar]
  11. Eyring G.; Curry B.; Broek A.; Lugtenburg J.; Mathies R. A. Assignment and interpretation of hydrogen out-of-plane vibrations in the Resonance Raman spectra of rhodopsin and bathorhodopsin. Biochemistry 1982, 21, 384–393. 10.1021/bi00531a028. [DOI] [PubMed] [Google Scholar]
  12. Warshel A.; Barboy N. Energy storage and reaction pathways in the first step of the vision process. J. Am. Chem. Soc. 1982, 104, 1469–1476. 10.1021/ja00370a003. [DOI] [Google Scholar]
  13. Birge R. R. Nature of the primary photochemical events in rhodopsin and bacteriorhodopsin. Biochem. Biophys. Acta. 1990, 1016, 293–327. 10.1016/0005-2728(90)90163-X. [DOI] [PubMed] [Google Scholar]
  14. Nathans J. Determinants of visual pigment absorbance: identification of the retinylidene Schiff’s base counterion in bovine rhodopsin. Biochemistry 1990, 29, 9746–9752. 10.1021/bi00493a034. [DOI] [PubMed] [Google Scholar]
  15. Sakmar T. P.; Franke R. R.; Khorana H. G. Glutamic acid-113 serves as the retinylidene Schiff base counterion in bovine rhodopsin. Proc. Natl. Acad. Sci. U. S. A. 1989, 86, 8309–8313. 10.1073/pnas.86.21.8309. [DOI] [PMC free article] [PubMed] [Google Scholar]
  16. Steinberg G.; Ottolenghi M.; Sheves M. pKa of the protonated Schiff base of bovine rhodopsin. A study with artificial pigments. Biophys. J. 1993, 64, 1499–1502. 10.1016/S0006-3495(93)81518-2. [DOI] [PMC free article] [PubMed] [Google Scholar]
  17. Lüdeke S.; Mahalingam M.; Vogel R. Rhodopsin Activation Switches in a Native Membrane Environment. Photochem. Photobiol. 2009, 85, 437–441. 10.1111/j.1751-1097.2008.00490.x. [DOI] [PubMed] [Google Scholar]
  18. Koutalos Y.; Ebrey T. G.; Gilson H. R.; Honig B. Octopus photoreceptor membranes. Surface charge density and pK of the Schiff base of the pigments. Biophys. J. 1990, 58, 493–501. 10.1016/S0006-3495(90)82394-8. [DOI] [PMC free article] [PubMed] [Google Scholar]
  19. Nakagawa M.; Iwasa T.; Kikkawa S.; Tsuda M.; Ebrey T. G. How vertebrate and invertebrate visual pigments differ in their mechanism of photoactivation. Proc. Natl. Acad. Sci. U. S. A. 1999, 96, 6189–6192. 10.1073/pnas.96.11.6189. [DOI] [PMC free article] [PubMed] [Google Scholar]
  20. Shichida Y.; Matsuyama T. Evolution of opsins and phototransduction. Philos. Trans. R. R. Soc., B 2009, 364, 2881–2895. 10.1098/rstb.2009.0051. [DOI] [PMC free article] [PubMed] [Google Scholar]
  21. Kefalov V. J. Rod and Cone Visual Pigments and Phototransduction through Pharmacological, Genetic, and Physiological Approaches. J. Biol. Chem. 2012, 287, 1635–1641. 10.1074/jbc.R111.303008. [DOI] [PMC free article] [PubMed] [Google Scholar]
  22. Imai H.; Imamoto Y.; Yoshizawa T.; Shichida Y. Difference in molecular properties between chicken green and rhodopsin as related to the function difference between cone and rod photoreceptor cells. Biochemistry 1995, 34, 10525–10531. 10.1021/bi00033a026. [DOI] [PubMed] [Google Scholar]
  23. Imamoto Y.; Yoshizawa T.; Shichida Y. Chromophore configuration of iodopsin and its photoproducts formed at low temperatures. Biochemistry 1996, 35, 14599–14607. 10.1021/bi9614850. [DOI] [PubMed] [Google Scholar]
  24. Imai H.; Terakita A.; Tachibanaki S.; Imamoto Y.; Yoshizawa T.; Shichida Y. Photochemical and biochemical properties of chicken blue-sensitive cone visual pigment. Biochemistry 1997, 36, 12773–12779. 10.1021/bi970809x. [DOI] [PubMed] [Google Scholar]
  25. Katayama K.; Furutani Y.; Imai H.; Kandori H. An FTIR study of monkey green- and red-sensitive visual pigments. Angew. Chem., Int. Ed. Engl. 2010, 49, 891–894. 10.1002/anie.200903837. [DOI] [PubMed] [Google Scholar]
  26. Katayama K.; Furutani Y.; Imai H.; Kandori H. Protein-bound water molecules in primate red- and green-sensitive visual pigments. Biochemistry 2012, 51, 1126–1133. 10.1021/bi201676y. [DOI] [PubMed] [Google Scholar]
  27. Katayama K.; Okitsu T.; Imai H.; Wada A.; Kandori H. Identical hydrogen-bonding strength of the retinal Schiff base between primate green- and red-sensitive pigments: new insight into color tuning mechanism. J. Phys. Chem. Lett. 2015, 6, 1130–1133. 10.1021/acs.jpclett.5b00291. [DOI] [PubMed] [Google Scholar]
  28. Katayama K.; Nakamura S.; Sasaki T.; Imai H.; Kandori H. Role of Gln114 in spectral tuning of a long-wavelength sensitive visual pigment. Biochemistry 2019, 58, 2944–2952. 10.1021/acs.biochem.9b00340. [DOI] [PubMed] [Google Scholar]
  29. Katayama K.; Imai H.; Kandori H. FTIR Study of S180A mutant of primate red-sensitive pigment. Chem. Lett. 2019, 48, 1142–1144. 10.1246/cl.190458. [DOI] [Google Scholar]
  30. Katayama K.; Nonaka Y.; Tsutsui K.; Imai H.; Kandori H. Spectral tuning mechanism of primate blue-sensitive visual pigment elucidated by FTIR spectroscopy. Sci. Rep. 2017, 7, 4904. 10.1038/s41598-017-05177-4. [DOI] [PMC free article] [PubMed] [Google Scholar]
  31. Furutani Y.; Kandori H.; Shichida Y. Structural changes in lumirhodopsin and metarhodopsin I studied by their photoreactions at 77 K. Biochemistry 2003, 42, 8494–8500. 10.1021/bi034438y. [DOI] [PubMed] [Google Scholar]
  32. Hanai S.; Katayama K.; Imai H.; Kandori H. Light-induced difference FTIR spectroscopy of primate blue-sensitive visual pigment at 163 K. Biophys. Physicobiol. 2021, 18, 40–49. 10.2142/biophysico.bppb-v18.005. [DOI] [PMC free article] [PubMed] [Google Scholar]
  33. Yoshizawa T.; Shichida Y. Low-temperature spectrophotometry of intermediates of rhodopsin. Methods Enzymol. 1982, 81, 333–354. 10.1016/S0076-6879(82)81051-3. [DOI] [PubMed] [Google Scholar]
  34. Rothschild K. J.; Zagaeski M.; Cantore W. A. Conformational changes of bacteriorhodopsin detected by Fourier transform infrared difference spectroscopy. Biochem. Biophys. Res. Commun. 1981, 103, 483–489. 10.1016/0006-291X(81)90478-2. [DOI] [PubMed] [Google Scholar]
  35. Hirano T.; Fujioka N.; Imai H.; Kandori H.; Wada A.; Ito M.; Shichida Y. Assignment of the vibrational modes of the chromophore of iodopsin and bathoiodopsin: Low-temperature Fourier transform infrared spectroscopy of 13C-and 2H-labeled iodopsins. Biochemistry 2006, 45, 1285–1294. 10.1021/bi0517077. [DOI] [PubMed] [Google Scholar]
  36. Mahalingam M.; Martinez-Mayorga K.; Brown M. F.; Vogel R. Two protonation switches control rhodopsin activation in membranes. Proc. Natl. Acad. Sci. U.S.A. 2008, 105, 17795–17800. 10.1073/pnas.0804541105. [DOI] [PMC free article] [PubMed] [Google Scholar]
  37. Zaitseva E.; Brown M. F.; Vogel R. Sequential rearrangement of interhelical networks upon rhodopsin activation in membranes: The Meta IIa conformational substate. J. Am. Chem. Soc. 2010, 132, 4815–4821. 10.1021/ja910317a. [DOI] [PMC free article] [PubMed] [Google Scholar]
  38. Okada T.; Fujiyoshi Y.; Silow M.; Navarro J.; Landau E. M.; Shichida Y. Functional role of internal water molecules in rhodopsin revealed by X-ray crystallography. Proc. Natl. Acad. Sci. U. S. A. 2002, 99, 5982–5987. 10.1073/pnas.082666399. [DOI] [PMC free article] [PubMed] [Google Scholar]
  39. Terakita A.; Koyanagi M.; Tsukamoto H.; Yamashita T.; Miyata T.; Shichida Y. Counterion displacement in the molecular evolution of the rhodopsin family. Nat. Struct. Mol. Biol. 2004, 11, 284–289. 10.1038/nsmb731. [DOI] [PubMed] [Google Scholar]
  40. Lüdeke S.; Beck M.; Yan E. C.; Sakmar T. P.; Siebert F.; Vogel R. The role of Glu181 in the photoactivation of rhodopsin. J. Mol. Biol. 2005, 353, 345–356. 10.1016/j.jmb.2005.08.039. [DOI] [PubMed] [Google Scholar]
  41. Vogel R.; Siebert F.; Yan E. C.; Sakmar T. P.; Hirshfeld A.; Sheves M. Modulating rhodopsin receptor activation by altering the pKa of the retinal Schiff base. J. Am. Chem. Soc. 2006, 128, 10503–10512. 10.1021/ja0627848. [DOI] [PubMed] [Google Scholar]
  42. Tsuda M. Kinetic study of photoregeneration process of digitonin-solubilized squid rhodopsin. Biochim. Biophys. Acta 1978, 502, 495–506. 10.1016/0005-2728(78)90082-8. [DOI] [PubMed] [Google Scholar]
  43. Ye S.; Zaitseva E.; Caltabiano G.; Schertler G. F.; Sakmar T. P.; Deupi X.; Vogel R. Tracking G-protein-coupled receptor activation using genetically encoded infrared probes. Nature 2010, 464, 1386–1389. 10.1038/nature08948. [DOI] [PubMed] [Google Scholar]
  44. Choe H.-W.; Kim Y. J.; Park J. H.; Morizumi T.; Pai E. F.; Krauss N.; Hofmann K. P.; Scheerer P.; Ernst O. P. Crystal structure of metarhodopsin II. Nature 2011, 471, 651–655. 10.1038/nature09789. [DOI] [PubMed] [Google Scholar]
  45. Deupi X.; Edwards P.; Singhal A.; Nickle B.; Oprian D.; Schertler G.; Standfuss J. Stabilized G protein binding site in the structure of constitutively active metarhodopsin-II. Proc. Natl. Acad. Sci. U.S.A. 2012, 109, 119–124. 10.1073/pnas.1114089108. [DOI] [PMC free article] [PubMed] [Google Scholar]
  46. Nakamichi H.; Okada T. Local peptide movement in the photoreaction intermediate of rhodopsin. Proc. Natl. Acad. Sci. U.S.A. 2006, 103, 12729–12734. 10.1073/pnas.0601765103. [DOI] [PMC free article] [PubMed] [Google Scholar]
  47. Palczewski K.; Kumasaka T.; Hori T.; Behnke C. A.; Motoshima H.; Fox B. A.; Le Trong I.; Teller D. C.; Okada T.; Stenkamp R. E.; Yamamoto M.; Miyano M. Crystal structure of rhodopsin: A G protein-coupled receptor. Science 2000, 289, 739–745. 10.1126/science.289.5480.739. [DOI] [PubMed] [Google Scholar]
  48. Nagata T.; Terakita A.; Kandori H.; Shichida Y.; Maeda A. The hydrogen-bonding network of water molecules and the peptide backbone in the region connecting Asp83, Gly120, and Glu113 in bovine rhodopsin. Biochemistry 1998, 37, 17216–17122. 10.1021/bi9810149. [DOI] [PubMed] [Google Scholar]
  49. Vogel R.; Fan G. B.; Sheves M.; Siebert F. The molecular origin of the inhibition of transducin activation in rhodopsin lacking the 9-methyl group of the retinal chromophore: a UV-Vis and FTIR spectroscopic study. Biochemistry 2000, 39, 8895–8908. 10.1021/bi000852b. [DOI] [PubMed] [Google Scholar]
  50. Nagata T.; Oura T.; Terakita A.; Kandori H.; Shichida Y. Isomer-specific interaction of the retinal chromophore with threonine-118 in rhodopsin. J. Phys. Chem. A 2002, 106, 1969–1975. 10.1021/jp0124488. [DOI] [Google Scholar]
  51. Lórenz-Fonfría V. A.; Furutani Y.; Ota T.; Ido K.; Kandori H. Protein fluctuations as the possible origin of the thermal activation of rod photoreceptors in the dark. J. Am. Chem. Soc. 2010, 132, 5693–5703. 10.1021/ja907756e. [DOI] [PubMed] [Google Scholar]
  52. Yamazaki Y.; Nagata T.; Terakita A.; Kandori H.; Shichida Y.; Imamoto Y. Intramolecular interactions that induce helical rearrangement upon rhodopsin activation: light-induced structural changes in metarhodopsin IIa probed by cysteine S-H stretching vibrations. J. Biol. Chem. 2014, 289, 13792–13800. 10.1074/jbc.M113.527606. [DOI] [PMC free article] [PubMed] [Google Scholar]
  53. Kandori H.; Kinoshita N.; Shichida Y.; Maeda A.; Needleman R.; Lanyi J. K. Cysteine S-H as a hydrogen-bonding probe in proteins. J. Am. Chem. Soc. 1998, 120, 5828–5829. 10.1021/ja980837i. [DOI] [Google Scholar]
  54. Furutani Y.; Shichida Y.; Kandori H. Structural changes of water molecules during the photoactivation processes in bovine rhodopsin. Biochemistry 2003, 42, 9619–9625. 10.1021/bi034592k. [DOI] [PubMed] [Google Scholar]
  55. Okano T; Kojima D; Fukada Y; Shichida Y; Yoshizawa T Primary structures of chicken cone visual pigments: vertebrate rhodopsins have evolved out of cone visual pigments. Proc. Natl. Acad. Sci. U. S. A. 1992, 89, 5932–5936. 10.1073/pnas.89.13.5932. [DOI] [PMC free article] [PubMed] [Google Scholar]
  56. Davies W. L.; Collin S. P.; Hunt D. M. Molecular ecology and adaptation of visual photopigments in craniates. Mol. Ecol. 2012, 21, 3121–3158. 10.1111/j.1365-294X.2012.05617.x. [DOI] [PubMed] [Google Scholar]
  57. Mooney V. L.; Szundi I.; Lewis J. W.; Yan E. C. Y.; Kliger D. S. Schiff base protonation changes in Siberian hamster ultraviolet cone pigment photointermediates. Biochemistry 2012, 51, 2630–2637. 10.1021/bi300157r. [DOI] [PMC free article] [PubMed] [Google Scholar]
  58. Dukkipati A.; Kusnetzow A.; Babu K. R.; Ramos L.; Singh D.; Knox B. E.; Birge R. R. Phototransduction by vertebrate ultraviolet visual pigments: Protonation of the retinylidene Schiff base following photobleaching. Biochemistry 2002, 41, 9842–9851. 10.1021/bi025883g. [DOI] [PubMed] [Google Scholar]

Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Supplementary Materials

bi2c00483_si_001.pdf (252.3KB, pdf)

Articles from Biochemistry are provided here courtesy of American Chemical Society

RESOURCES