3.E.1 The Ion-translocating Microbial Rhodopsin (MR) Family
Members of the MR family catalyze light-driven ion translocation across microbial cytoplasmic membranes or serve as light receptors. Among the high resolution structures for members of the MR family are the archaeal proteins, bacteriorhodopsin (Luecke et al., 1999), sensory rhodopsin II (Royant et al., 2001) and halorhodopsin (Kolbe et al., 2000) as well as an Anabaena cyanobacterial sensory rhodopsin (3.E.1.8.a) (Vogeley et al., 2004). Homologues include putative fungal chaparone proteins, a retinal-containing rhodopsin from Neurospora crassa (Maturana et al., 2001), a H+-pumping rhodopsin from Leptosphaeria maculans (Waschuk et al., 2005), retinal-containing proton pumps isolated from marine bacteria (Béjà et al., 2000), a green light-activated photoreceptor in cyanobacteria that does not pump ions and interacts with a small (14 kDa) soluble transducer protein (Jung et al., 2003; Vogeley et al., 2004) and light-gated H+ channels from the green alga, Chlamydomonas reinhardtii (Nagel et al., 2002). The N. crassa NOP-1 protein exhibits a photocycle and conserved H+ translocation residues that suggest that this putative photoreceptor is a slow H+ pump (Brown et al., 2001; see also Brown, 2004 and Waschuk et al., 2005). Allosteric structural changes in the photocycle are mediated by a sliding movement of a transmembrane helix (Takeda et al. 2004). MR proteins such as SRII exhibit fast internal motion and residual conformational entropy (O'Brien et al. 2020). Procedures for the formation of thin (mono-) and thick (multi-) layers from materials containing BR and BR/nanoparticle hybrids have been reviewed (Oleinikov et al. 2020).
The Anabaena sensory rhodopsin exhibits light-induced interconversion between 13-cis and all trans states (Vogeley et al., 2004). The ratio of its cis and trans chromophore forms depends on the wavelength of illumination, thus providing a mechanism for a single protein to signal the color of light, for example, to regulate color-sensitive processes such as chromatic adaptation in photosynthesis. Its cytoplasmic half channel, highly hydrophobic in the archaeal rhodopsins, contains numerous hydrophilic residues networked by water molecules, providing a connection from the photoactive site to the cytoplasmic surface believed to interact with the receptor's soluble 14-kilodalton transducer.
Most proteins of the MR family are all of about the same size (250-350 amino acyl residues) and possess seven transmembrane helical spanners with their N-termini on the outside and their C-termini on the inside. There are 8 subfamilies in the MR family: (1) bacteriorhodopsins pump protons out of the cell; (2) halorhodopsins pump chloride (and other anions such as bromide, iodide and nitrate) into the cell; (3) sensory rhodopsins, which normally function as receptors for phototactic behavior, are capable of pumping protons out of the cell if dissociated from their transducer proteins; (4) the fungal chaparones are stress-induced proteins of ill-defined biochemical function, but this subfamily also includes a H+-pumping rhodopsin (Waschuk et al., 2005); (5) the bacterial rhodopsin, called proteorhodopsin, is a light-driven proton pump that functions as does bacteriorhodopsins; (6) the N. crassa retinal-containing receptor serves as a photoreceptor (Zhai et al., 2001); (7) the green algal light-gated proton channel, channelrhodpsin-1, (8) sensory rhodopsins from cyanobacteria and (9) light-activated rhodopsin guanylyl cyclases. A phylogenetic analysis of microbial rhodopsins and a detailed analysis of potential examples of horizontal gene transfer have been published (Sharma et al., 2006).
Bacterio- and halorhodopsins pump 1 H+ and 1 Cl- per photon absorbed, respectively. Specific transport mechanisms and pathways have been proposed (see Kolbe et al., 2000; Lanyi and Schobert, 2003; Schobert et al., 2003). The mechanism involves (1) photo-isomerization of the retinal and its initial configurational changes, (2) deprotonation of the retinal Schiff base and the coupled release of a proton to the extracellular membrane surface, and (3) the switch event that allows reprotonation of the Schiff base from the cytoplasmic side. Six structural models describe the transformations of the retinal and its interaction with water 402, Asp85, and Asp212 in atomic detail, as well as the displacements of functional residues farther from the Schiff base. The changes provide rationales for how relaxation of the distorted retinal causes movements of water and protein atoms that result in vectorial proton transfers to and from the Schiff base (Lanyi and Schobert, 2003). Helix deformation is coupled to vectorial proton transport in the photocycle of bacteriorhodopsin (Royant et al., 2000).
The marine bacterial rhodopsin has been reported to function as a proton pump. However, it most closely resembles sensory rhodopsin II of archaea as well as an Orf from the fungus Leptosphaeria maculans (AF290180). These proteins exhibit 20-30% identity with each other. Sensory rhodopsins are widespread in the microbial world, but they exhibit different modes of signaling in different organisms, including interaction with other membrane proteins, interaction with cytoplasmic transducers and light-controlled Ca2+ channel activity. Work on cyanobacteria, algae, fungi and marine proteobacteria has shown that the common design of these proteins allows rich diversity in their signaling mechanisms (Spudich 2006).
The association of sensory rhodopsins with their transducer proteins appears to determine whether they function as transporters or receptors. Association of a sensory rhodopsin receptor with its transducer occurs via the transmembrane helical domains of the two interacting proteins. There are two sensory rhodopsins in any one halophilic archaeon, one (SRI) that responds positively to orange light but negatively to blue light, the other (SRII) that responds only negatively to blue light. Each transducer is specific for its cognate receptor. An x-ray structure of SRII complexed with its transducer (HtrII) at 1.94 Å resolution is available (Gordelly et al., 2002). Molecular and evolutionary aspects of the light-signal transduction by microbial sensory receptors have been reviewed (Inoue et al. 2014).
Sol-gel immobilization of proteins in transparent inorganic matrices provide a liposomal system in which the liposome provides membrane structure. Two transmembrane proteins, bacteriorhodopsin (bR) and F0F1-ATP synthase have been incorporated into such a matrix called proteogels; if containing only bRho, a stable proton gradient forms when irradiated with visible light, whereas proteogels containing proteoliposomes with both bRho and an F0F1-ATP synthase couple the photo-induced proton gradient to the production of ATP (Luo et al. 2005). Thus, the liposome/sol-gel architecture can harness the properties of transmembrane proteins and enable a variety of applications, from power generation and energy storage to the powering of molecular motors.
Channelrhodopsin-1 (ChR1) or channelopsin-1 (Chop1; Cop3; CSOA) of C. reinhardtii is most closely related to the archaeal sensory rhodopsins. It has 712 aas with a signal peptide, followed by a short amphipathic region, and then a hydrophobic N-terminal domain with seven probable TMSs (residues 76-309) followed by a long hydrophilic C-terminal domain of about 400 residues. Part of the C-terminal hydrophilic domain is homologous to intersectin (EH and SH3 domain protein 1A) of animals (AAD30271).
Chop1 serves as a light-gated proton channel and mediates phototaxis and photophobic responses in green algae (Nagel et al., 2002). Based on this phenotype, Chop1 could be assigned to TC category #1.A, but because it belongs to a family in which well-characterized homologues catalyze active ion transport, it is assigned to the MR family. Expression of the chop1 gene, or a truncated form of this gene encoding only the hydrophobic core (residues 1-346 or 1-517) in frog oocytes in the presence of all-trans retinal produces a light-gated conductance that shows characteristics of a channel, passively but selectively permeable to protons. This channel activity may generate bioelectric currents (Nagel et al., 2002).
A homologue of ChR1 in C. reinhardtii is channelrhodopsin-2 (ChR2; Chop2; Cop4; CSOB). This protein is 57% identical, 10% similar to ChR1. It forms a cation-selective ion channel activated by light absorption. It transports both monovalent and divalent cations. It desensitizes to a small conductance in continuous light. Recovery from desensitization is accelerated by extracellular H+ and a negative membrane potential. It may be a photoreceptor for dark adapted cells (Nagel et al., 2003). A transient increase in hydration of transmembrane α-helices with a t(1/2) = 60 μs tallies with the onset of cation permeation. Aspartate 253 accepts the proton released by the Schiff base (t(1/2) = 10 μs), with the latter being reprotonated by aspartic acid 156 (t(1/2) = 2 ms). The internal proton acceptor and donor groups, corresponding to D212 and D115 in bacteriorhodopsin, are clearly different from other microbial rhodopsins, indicating that their spatial position in the protein was relocated during evolution. E90 deprotonates exclusively in the nonconductive state. The observed proton transfer reactions and the protein conformational changes relate to the gating of the cation channel (Lórenz-Fonfría et al. 2013).
Most of the MR family homologues in yeast and fungi are of about the same size and topology as the archaeal proteins (283-344 amino acyl residues; 7 putative transmembrane α-helical segments), but they are heat shock- and toxic solvent-induced proteins of unknown biochemical function. They have been suggested to function as pmf-driven chaperones that fold extracellular proteins (Zhai et al., 2001), but only indirect evidence supports this postulate. The MR family is distantly related to the 7 TMS LCT family (TC #2.A.43) (Zhai et al., 2001). It is a part of the TOG superfamily which includes G-protein coupled receptors (GPCRs) (Yee et al. 2013), and the conclusioin of homology between MRs and GPCRs has been extensively confirmed (Shalaeva et al. 2015).
Archaerhodopsin-2 (aR2), a retinal protein-carotenoid complex found in the claret membrane of Halorubrum sp. aus-2, functions as a light-driven proton pump. Trigonal and hexagonal crystals revealed that trimers are arranged on a honeycomb lattice (Yoshimura and Kouyama, 2008). In these crystals, the carotenoid bacterioruberin binds to crevices between the subunits of the trimer. Its polyene chain is inclined from the membrane normal by an angle of about 20 degrees and, on the cytoplasmic side, it is surrounded by helices AB and DE of neighbouring subunits. This peculiar binding mode suggests that bacterioruberin plays a structural role for the trimerization of aR2. When compared with the aR2 structure in another crystal form containing no bacterioruberin, the proton release channel takes a more closed conformation in the P321 or P6(3) crystal; i.e., the native conformation of protein is stabilized in the trimeric protein-bacterioruberin complex.
A crystallographic structure of xanthorhodopsin at 1.9 Å resolution revealed a dual chromophore, the geometry of the carotenoid and the retinal (Luecke et al., 2008). The close approach of the 2 polyenes at their ring ends explains why the efficiency of the excited-state energy transfer is as high as approximately 45%, and the 46 degrees angle between them suggests that the chromophore location is a compromise between optimal capture of light of all polarization angles and excited-state energy transfer. At 1.9 Å resolution, the structure revealed a light-driven proton pump with a dual chromophore. Ion-transporting rhodopsins of marine bacteria have been reviewed (Inoue et al. 2014).
Most residues participating in the trimerization are not conserved in bacteriorhodopsin, a homologous protein capable of forming a trimeric structure in the absence of bacterioruberin. Despite a large alteration in the amino acid sequence, the shape of the intratrimer hydrophobic space filled by lipids is highly conserved between aR2 and bacteriorhodopsin. Since a transmembrane helix facing this space undergoes a large conformational change during the proton pumping cycle, it is feasible that trimerization is an important strategy to capture special lipid components that are relevant to the protein activity (Yoshimura and Kouyama, 2008).
Ion-pumping bacterial rhodopsins functioning as outward H+ or Na+ and inward Cl- pumps convert light energy into transmembrane electrochemical potential differences. The H+, Na+, and Cl- pumps possess conserved respective DTE, NDQ, and NTQ motifs in helices C, which likely serve as their functional determinants, and this has been verified (Inoue et al. 2016). Phylogenetic analyses suggested that a H+ pump was the common ancestor from which Cl- pumps emerged followed by Na+ pumps. Inoue et al. 2016 proposed that successful functional conversion was achieved when these amino acid sequences changed, possibly accompanied by other changes.
Nango et al. 2016 used time-resolved serial femtosecond crystallography at an x-ray free electron laser to visualize conformational changes in bRho from nanoseconds to milliseconds following photoactivation. An initially twisted retinal chromophore displaces a conserved tryptophan residue of transmembrane helix F on the cytoplasmic side of the protein while dislodging a key water molecule on the extracellular side. The resulting cascade of structural changes throughout the protein shows how motions are choreographed as bRho transports protons uphill against a transmembrane concentration gradient.
Brho (BR) has light-independent lipid scramblase activity (Verchère et al. 2017). This activity occurs at a rate >10,000 per trimer per second, comparable to that of other scramblases including bovine rhodopsin and fungal TMEM16 proteins. BR scrambles fluorescent analogues of common phospholipids but does not transport a glycosylated diphosphate isoprenoid lipid. In silico analyses suggested that membrane-exposed polar residues in transmembrane helices 1 and 2 of BR may provide the molecular basis for lipid translocation by coordinating the polar head-groups of transiting phospholipids. Consistent with this possibility, molecular dynamics simulations of a BR trimer in a phospholipid membrane revealed water penetration along transmembrane helix 1 with the cooperation of a polar residue (Y147 in transmembrane helix 5) in the adjacent protomer. These findings suggest that the lipid translocation pathway may lie at or near the interface of the protomers of the BR trimer (Verchère et al. 2017).
Electronic current passes through bR-containing artificial lipid bilayers in solid 'electrode-bilayer-electrode' structures. The current through the protein is more than four orders of magnitude higher than would be estimated for direct tunneling through 5-nm water-free peptides. Jin et al. 2006 found that electron transport (ET) occurs only if retinal or a close analogue is present in the protein. As long as the retinal can isomerize after light absorption, there is a photo-ET effect. The contribution of light-driven proton pumping to the steady-state photocurrents is negligible. Possibly this is relevant to the early evolutionary origin of halobacteria (Jin et al. 2006).
Parvularcula oceani xenorhodopsin (PoXeR) was the first light-driven inward proton pump with a brho topology and structure, binding retinal to TMS 7. Ultrafast pump-probe spectroscopy revealed that the isomerization time of retinal is 1.2 ps, considerably slower than those of other microbial rhodopsins (180-770 fs). Following the production of J, the K intermediate was formed at 4 ps. Proton transfer occurred on a slower time-scale. While a proton was released from Asp216 into the cytoplasm, no proton-donating residue was identified on the extracellular side. A branched retinal isomerization (from 13-cis-15-anti to 13-cis-15-syn and all-trans-15-anti) occurred simultaneously with proton uptake. Thus, retinal isomerization is the rate-limiting process in proton uptake, and the regulation of pKa of the retinal Schiff base by thermal isomerization enables uptake from the extracellular medium (Inoue et al. 2018).
The generalized transport reaction for bacterio- (and some sensory) rhodopsins is:
H+ (in) + hν → H+ (out)
That for halorhodopsin is:
Cl- (out) + hν → Cl- (in)
That for xenorhodopsin is:
H+ (out) + hν → H+ (in)
Bacteriorhodopsin. Proton efflux occurs via a transient linear water-molecule chain in a hydrophobic section of the Brho channel between Asp96 and Asp85 (Freier et al., 2011). It can be converted to a chloride uptake pump by a single amino acid substitution at position 85. However, halorhodopsin (3.E.1.2.1), which pumps chloride ions (Cl-) into the cell, apparently does not use hydrogen-bonded water molecules for Cl- transport (Muroda et al. 2012). Nango et al. 2016 used time-resolved serial femtosecond crystallography and an x-ray free electron laser to visualize conformational changes in bRho from nanoseconds to milliseconds following photoactivation. An initially twisted retinal chromophore displaces a conserved tryptophan residue of transmembrane helix F on the cytoplasmic side of the protein while dislodging a key water molecule on the extracellular side. The resulting cascade of structural changes throughout the protein shows how motions are choreographed as bRho transports protons uphill against a transmembrane concentration gradient. Nango et al. 2016 have created a 3-d movie of structural changes in the protein showing that an initially twisted retinal chromophore displaces a conserved tryptophan residue of transmembrane helix F on the cytoplasmic side of the protein while dislodging a key water molecule on the extracellular side. Brho has light-independent lipid scramblase activity (Verchère et al. 2017). This activity occurs at a rate >10,000 per trimer per second, comparable to that of other scramblases including bovine rhodopsin and fungal TMEM16 proteins. BR scrambles fluorescent analogues of common phospholipids but does not transport a glycosylated diphosphate isoprenoid lipid. In silico analyses suggested that membrane-exposed polar residues in transmembrane helices 1 and 2 of BR may provide the molecular basis for lipid translocation by coordinating the polar head-groups of transiting phospholipids. Consistent with this possibility, extensive coarse-grained molecular dynamics simulations of a BR trimer in a phospholipid membrane revealed water penetration along transmembrane helix 1 with the cooperation of a polar residue (Y147 in transmembrane helix 5) in the adjacent protomer. These findings suggest that the lipid translocation pathway may lie at or near the interface of the protomers of the BR trimer (Verchère et al. 2017). Retinal isomerization has been observed in the using a femtosecond x-ray laser (Nogly et al. 2018). S-TGA-1, a halobacterium-derived glycolipid, has the highest specificity to bRho, with a nanomolar dissociation constant (Inada et al. 2019). Weinert et al. 2019 recorded the structural changes in bacteriorhodopsin over 200 milliseconds in time. The snapshot from the first 5 milliseconds after photoactivation shows structural changes associated with proton release. From 10 to 15 milliseconds onwards, large additional structural rearrangements, up to 9 Å on the cytoplasmic side. Rotation of leucine-93 and phenylalanine-219 opens a hydrophobic barrier, leading to the formation of a water chain connecting the intracellular aspartic acid-96 with the retinal Schiff base. The formation of this proton wire recharges the membrane pump with a proton for the next cycle (Weinert et al. 2019). The effect of membrane composition on the orientation and activity of bR has been reported (Palanco et al. 2017). Efficient transfer of bRho from native membranes to covalently circularized nanodiscs has been accomplished (Yeh et al. 2018).
Bacteriorhodopsin of Halobacterium salinarum
"Middle" rhodopsin or Brhol; has 11-cis-retinal and shows intermediate properties between Brho and sensory rhodopsin II (Sudo et al., 2011). Its structure is known to 2.0 Å resolution following crystalization using polymer-bounded lipid nanodiscs (Broecker et al. 2017).
Middle rhodopsin of Haloquadratum walsbyi (G0LFX8)
Archaerhodopsin 3, AR3. Pumps protons in response to light absorption (Saint Clair et al. 2012). 86% identical to 3.E.1.1.2.
AR3 of Halorubrum sodomense
Bacteriorhodopsin I (HmBRI) of 250 aas and 7 TMSs. The structure is known to 2.5 Å resolution, revealing the usual BRI fold but with several modifications (Shevchenko et al. 2014). Expression in E. coli membranes does not affect the overall structure.
Bacteriorhodopsin I of Haloarcula marismortui
Sensory rhodopsin (green-light-activated photoreceptor; does not transport ions) (Jung et al., 2003). Has all-trans-retinal when dark adapted, but 11-cis-retinal when light adapted due to reversible interconversion (Sineshchekov et al., 2005). Anabaena sensory rhodopsin, a photochromic sensor that interacts with a soluble 14-kDa cytoplasmic transducer that is encoded on the same operon, interconverts between all-trans-15-anti and 13-cis-15-syn retinal forms depending on the wavelength of illumination, although only the former participates in a photocycle with a signaling M intermediate (Dong et al. 2016). A mutation in the cytoplasmic half-channel of the protein, replacing Asp217 with Glu (D217E), results in the creation of a light-driven, single- photon, inward proton transporter. Dong et al. 2016 presented the 2.3 A structure of dark-adapted D217E ASR, which reveals changes in the water network surrounding Glu217, as well as a shift in the carbon backbone near retinal-binding Lys210, illustrating a possible pathway leading to the protonation of Glu217 in the cytoplasmic half-channel, located 15 A from the Schiff base.
Sensory rhodopsin of Anabaena (Nostoc) sp. PCC7120
H+-pumping electrogenic bacteriorhodopsin of 250 aas and 7 TMSs (Kamo et al. 2006).
Brho of Haloterrigena turkmenica
Inward H+ pumping xenorhodopsin (bacteriorhodopsin) of 228 aas and 7 TMSs.
Xenorhodopsin of Nanosalina sp. (strain J07AB43)
Rhodopsin of 239 aas and 9 TMSs. This rhodopsin is from the thermophilic eubacterium Rubrobacter xylanophilus DSM 9941(T) and was isolated from thermally polluted water. Although R. xylanophilus rhodopsin (RxR) is from an Actinobacterium, it is located between eukaryotic and archaeal rhodopsins in the phylogenetic tree (Kanehara et al. 2017). E. coli cells expressing RxR showed a light-induced decrease in environmental pH and inhibition by a protonophore, indicating that it works as a light-driven outward proton pump. Purified RxR has an absorption maximum at 541 nm and binds all-trans retinal. The pKa values for the protonated retinal Schiff base and its counterion were 10.7 and 1.3, respectively. Of note, RxR showed an extremely high thermal stability in comparison with other proton pumping rhodopsins such as thermophilic rhodopsin TR (by 16-times) and bacteriorhodopsin from Halobacterium salinarum (HsBR, by 4-times) (Kanehara et al. 2017).
Rhodoopsin of Rubrobacter xylanophilus
Halorhodopsin Cl- uptake pump; homologous to bacteriorhodopsin (3.E.1.1.1) which can be converted from a proton pump with outwardly directed polarity into a chloride pump with inwardly directed polarity via a single amino acid substitution at position 85. Cl- transport does not depend on water hydrogen bonded to the chromophore as in the case of bacteriorhodopsin (Muroda et al. 2012). However, inter-helical hydrogen bonds, mediated by a key arginine residue, largely govern the dynamics of the protein and water groups coordinating the chloride ion (Jardón-Valadez et al. 2014). Helices E and F probably move considerable during chloride binding and ion transport (Schreiner et al. 2016).
Halorhodopsin of Halobacterium salinarum
Halorhodopsin (a trimer with the carotenoid, bacterioruberin, binding to crevices between adjacent protein subunits in the trimeric assembly; Sasaki et al., 2012). Structure known to 2.0 Å resolution (Kouyama et al., 2010) (PDB# 3A7K)).
Halorhodopsin of Natronomonas pharaonis (P15647)
Sensory rhodopsin II or photoreceptor phoborhodopsin (ppR). The 3-d structure has been solved by NMR (Gautier and Nietlispach 2012). The dynamics of light induced conformational changes have been studied (Taniguchi et al. 2007).
Sensory rhodopsin II (phoborhodopsin) of Halobacterium salinarum
Sensory rhodopsin II, SR2 (Sop2; ppR), also called phoborhodopsin. The NMR solution structure of the detergent solubilized protein is in good agreement with the x-ray structure (Gautier et al. 2010). The onformational dynamics of Sensory Rhodopsin II in nanolipoprotein and styrene-maleic acid lipid particles has been studied (Mosslehy et al. 2019). The retinal configuration of ppR intermediates have been studied (Makino et al. 2018).
SRII of Natronomonas pharaonis (P42196)
Sensory rhodopsin III, SRIII, of 232 aas and 7 TMSs (Fu et al. 2010).
SRIII of Haloarcula marismortui
Phoborhodopsin (sensory rhodopsin II) of 249 aas and 7 TMSs. The photochemistry and proton transport have been reviewed (Kamo et al. 2001) and the crystal structure is known (Kandori and Kamo 2002).
Phoborhodopsin of Halorubrum chaoviator
Retinal binding protein, Neurospora Opsin-1, NOP-1 (Bieszke et al. 1999; Bieszke et al. 2007).
NOP-1 of Neurospora crassa
H+ pumping rhodopsin (Idnurm and Howlett 2001; Waschuk et al., 2005)
Rhodopsin of Leptosphaeria maculans (AAG01180)
Acetaularia rhodopsin I, ARI or c102333 of 246 aas. It exhibits outward H+ pumping activity, and D89 and D100 are essential for pumping activity (Lee et al. 2015). Blue-light causes a shunt of the photocycle under H+ reuptake from the extracellular side (Tsunoda et al. 2006). Similarities and differences of AR with BR have been revealed by detailed electrophysiological studies, revealing among other things, the voltage dependencies of the pump (Tsunoda et al. 2006).
c102333 of Acetabularia acetabulum (Q1AJZ3)
Opsin 1, Bacteriorhodopsin-like protein
Opsin 1 of Guillardia theta (Q2QCJ4)
Possible chaperone membrane protein related to Hsp30, Mrh1 (320 aas; 33% identical to Hsp30p). This protein and its two paralogues, Hsp30 and YR02, are induced by heat shock and are present primarily in the plasma membrane (Wu et al. 2000). It plays a role in acetic acid tolerance and may be an acetic acid exporter (Takabatake et al. 2015).
Mrh1p of Saccharomyces cerevisiae (Q12117)
Cyanorhodopsin of 334 aas and 7 TMSs, Ops1 (Frassanito et al. 2010).
Cyanorhodopsin of Cyanophora paradoxa
Yro2 of 344 aas and 7 TMSs. Plays a role in acetic acid tolereance and is induced by acetic acid stress and by entry into the stationary phase. It is 72% identical to Mrh1 (TC# 3.E.1.4.6) which is also believed to be involved in the acetic acid stress response (Takabatake et al. 2015).
Yro2 of Saccharomyces cerevisiae
Bacterio-rhodopsin/guanylyl cyclase 1 fusion protein of 626 aas; light-activated enzyme, RhCG, Gc1 or Cyc1Op. The central bacteriorhodopsin domain with 7 TMSs is linked via an additional TMS to the C-terminal adenylate/guanylate cyclase catalytic domain. CyclOp enables precise and rapid optogenetic manipulation of cGMP levels in cells and
animals (Gao et al. 2015).
Gc1 of Blastocladiella emersonii (Aquatic fungus)
Learning/memory process protein of 704 aas and 7 N-terminal TMSs as the rhodopsin (Rh) domain with a C-terminal cyclic nucleotide phosphodiesterase (PDE) domain. The Rh-PDE enzyme light-dependently decreases the concentrations of cyclic nucleotides such as cGMP and cAMP. Photoexcitation of purified full-length Rh-PDE yields an "M" intermediate with a deprotonated Schiff base; its recovery is much faster than that of the enzyme domain (Watari et al. 2019). Mechanistic insights into rhodopsin-mediated, light-dependent regulation of second-messenger levels have thus been revealed (Watari et al. 2019).
Rh-PDE fusion protein of Salpingoeca rosetta
Green-light-absorbing H+ pumping proteorhopdopsin of 249 aas and 8 TMSs. It exhibits variable vectorality: H+ is pumped out at basic pH but not at acidic pH; see Friedrich et al., 2002). It presents a fast proton release and an alkaliphilic photocycle, consistent with its marine origin and the near-surface environment where this bacterium was collected. This proteorhodopsin has been used to measure membrane potentials and electrical spiking in E. coli (Kralj et al., 2011; Ward et al., 2011). 3-d structures of three proteorhodopsins show that they can exist as pentamers or hexamers, depending on the protein (Ran et al. 2013). Protonation states of several carboxylic acids, the boundaries and distortions of transmembrane α-helices, and secondary structural elements in the loops have been identified (Shi et al. 2009). Proteorhodopsin molecules incorporated into mesostructured silica films exhibit native-like function, as well as enhanced thermal stability compared to surfactant or lipid environments (Jahnke et al. 2018).
Green-light-absorbing proteorhodopsin from an uncultured γ-proteobacterium EOAC 31A08
Uncharaacterized bacteriorhodopsin of 289 aas and 7 TMSs.
UP of Parvularcula oceani
Uncharacterized bacteriorhodopsin of 321 aas and 7 TMSs.
UP of Parvularcula oceani
Blue-light absorbing proteorhodopsin (BPR) of 251 aas and 8 TMSs including a cleavable N-terminal TMS. BPR does not rely on the Sec pathway for inner membrane integration (Soto-Rodríguez and Baneyx 2018). The BPR signal sequence is recognized by the signal recognition particle (SRP; a protein that orchestrates the cotranslational biogenesis of inner membrane proteins) and serves as a beneficial "pro" domain rather than a traditional secretory peptide. It is a light-driven proton pump that may have a regulatory rather than energy harvesting function, based on light-induced opening of proton channels to modulate cell physiology depending on light intensity variations. It could therefore be a sensory rhodopsin, potentially associated with a transducer component. It presents a much slower photocycle than that of the green-absorbing proteorhodopsin, probably an adaptation to the intensity of solar illumination at a depth of 75m, where this bacterium was collected. Transport occurs only at pHs above 7 and is unidirectional.
BPR of Gamma-proteobacterium Hot 75m4
Xanthorhodopsin, a proton pump with a carotenoid antenna, salinixanthin (Lanyi and Balashov 2008). A crystal structure (1.9 Å resolution) is available (Luecke et al., 2008).
Xanthorhodopsin with a salinixanthin chromophore of Salinibacter ruber (Q2S2F8)
Rhodopsin of 298 aas and 7 TMSs (associates with salinixanthin, the light-harvesting carotenoid antenna of xanthorhodopsin) (Imasheva et al., 2009; Hashimoto et al., 2010). Expression in a chemotrophic E. coli enabled light-driven phototrophic energy generation (Kim et al. 2017).
Rhodopsin of Gloeobacter violaceus (Q7NP59)
Bacteriorhodopsin-like circadian clock related protein (Okamoto and Hastings, 2003)
BacRhodopsin of Pyrocystis lunula (Q8GZE7)
H+-pumping bacteriorhodopsin, Brho or ESR (Petrovskaya et al. 2010). Photoelectric potential generation correlates with the ESR structure and proposed mechanism of proton transfer (Siletsky et al. 2016).
Brho of Exiguobacterium sibiricum (B1YFV8)
Proton pumping proteorhodopsin of 253 aas (Kimura et al. 2011).
Proteorhodopsin of Nonlabens dokdonensis (Donghaeana dokdonensis)
Na+ or H+ pumping bacteriorhodopsin, NaR, Kr2 or KR2. It uses light to pump protons or sodium ions from the cell depending on the ionic composition of the medium. In cells suspended in a KCl solution, NaR functions as a light-driven proton pump, whereas in a NaCl solution, it exhibits light-driven sodium ion pumping, a novel activity within the rhodopsin family (da Silva et al. 2015). A cation switch controls its conformations, and specific interactions of Na+ with the half-channels open an appropriate path for ion translocation (da Silva et al. 2015). Several high resolution x-ray structures have been solved (4XTO, Kato et al. 2015). Putative Na+ binding sites have been identified, and it was shown how protonation and conformational changes gate the ion through these sites toward the extracellular side (Suomivuori et al. 2017). Evidence for homology of this and other microbial rhodopsin with GPCR receptors including mamalian rhodopsins has been presented (Yee et al. 2013; Shalaeva et al. 2015).
NaR of Dokdonia
eikasta (Krokinobacter eikastus)
Proteorhodopsin of 246 aas and 7 TMSs, Pro. A light-driven Na+ pump (Bertsova et al. 2015).
Proteorhodopsin of Dokdonia sp. PRO95
Bacteriorhodopsin (thermophilic rhodopsin; TR) of 260 aas and 7 TMSs. 53% identical to xanthorhodopsin (TC# 3.E.1.6.2). It is a photoreceptor protein with extremely high thermal stability and a light-driven electrogenic proton pump. The x-ray crystal structure revealed the presence of a putative binding site for a carotenoid antenna and a larger number of hydrophobic residues and aromatic-aromatic interactions than in most microbial rhodopsins (Tsukamoto et al. 2016). The structural changes upon thermal stimulation involved a thermally induced structure in which an increase of hydrophobic interactions in the extracellular domain, the movement of extracellular domains, the formation of a hydrogen bond, and the tilting of transmembrane helices were observed. An extracellular LPGG motif between helices F and G may play an important role in thermal stability, acting as a "thermal sensor" (Tsukamoto et al. 2016).
Bacteriorhodopsin of Thermus thermophilus
Channelrhodopsin-1 (chlamyrhodopsin-3) (ChR1; Cop3; CSOA) (light-gated proton channel) (Nagel et al., 2003). TMSs 1 and 2 are the main structures involved in desensitization involving the stabilization of the protein's conformation and the alteration of the charge distribution around the retinal-Schiff base (Zamani et al. 2017). Replacing the glutamate located at the central gate of the ion channel with positively charged amino acyl residues reverses the ion selectivity and allows anion conduction (Zhang et al. 2019).
Channelrhodopsin-1 of Chlamydomonas reinhardtii
Channelrhodopsin-2 (chlamyrhodopsin-4; ChR2; CR2; Cop4; CSOB) (light-gated cation-selective ion channel (both monovalent and divalent cations are transported)) (Nagel et al., 2003). Berndt et al. (2010) showed that ChR2 has two open states with differing ion selectivities. The channel is fairly nonspecific at the beginning of a light pulse, and becomes more specific for protons during longer periods of light exposure. Residues involved in channel closure have been identified (Bamann et al. 2010). ChR2 is 712 aas long; the MR domain is N-terminal (Lee et al. 2015). The free energy profiles computed for proton transfer to the counterion, either via a direct jump or mediated by a water molecule, demonstrate that, when retinal is all-trans, water and protein electrostatic interactions largely favour the protonated retinal Schiff base state (Adam and Bondar 2018).
Blue light illumination of ChR2 activates an intrinsic leak channel conductive for cations. Sequence comparison of ChR2 with the related ChR1 protein revealed a cluster of charged amino acids within the predicted transmembrane domain 2 (TM2), which includes glutamates E90, E97 and E101. Charge inversion substitutions altered ChR2 function, replacement of E90 by lysine or alanine resulted in differential effects on H+- and Na+-mediated currents. These results are consistent with this glutamate side chain within TMS2 contributing to ion flux through and the cation selectivity of ChR2 (Ruffert et al., 2011). Glutamate residue-97 lies in the outer pore where it interacts with a cation to facilitate dehydration. This residue is also the primary binding target of Gd3+(Tanimoto et al., 2012). Channelrhodopsin has been converted into a light-gated chloride channel (Wietek et al. 2014). TMSs 2, 6 and 7 reorient or rearrange during the photocycle with no major differences near TMSs 3 and 4 at the dimer interface. TMS2 plays a key role in light-induced channel opening and closing in ChR2 (Müller et al. 2015). Negative charges at the extracellular side of transmembrane domain 7 funnel cations into the pore (Richards and Dempski 2015). CrChR2, is the most widely used optogenetic tool in neuroscience. Water efflux and the cessation of the ion conductance are synchronized (Lórenz-Fonfría et al. 2015). light and pH induce changes in the structure and accessibility of TMSB (Volz et al. 2016). Residues V86, K93 and N258 form a putative barrier to ion translocation. These residues contribute to cation selectivity (V86 and N258), the transition between the two open states (V86), open channel stability, and the hydrogen-bonding network (K93I and K93N) (Richards and Dempski 2017). The x-ray structure is available and reveals much about the mechanism of channel regulation (Gerwert 2017; Volkov et al. 2017). The mechanism of formation of the ion channel of ChR2 has been studed by molecular dynamics simulation and steering (Yang et al. 2019).
Channelrhodopsin-2, CR2, of Chlamydomonas reinhardtii (Q8RUT8)
Channelrhodopsin-2 light-gated ion channel. A 6Å projection map is available (Müller et al., 2011). Glutamate residue-97 lies in the outer pore where it interacts with a cation to facilitate dehydration. This residue is also the primary binding target of Gd3+ (Tanimoto et al., 2012).
Channelrhodopsin-2 of Volvox carteri (B4Y105)
Channelopsin, MChR1 (Govorunova et al., 2011). In another channelrhodopsin (CrChR2) of this family, an E97A mutation in TMS 2 prevents high affiinity binding of the inhibitor, Gd3+ and interfers with photocurrent, but this ChR1 with an alanine at this position, has low affininty for Gd3+ but normal photocurrent (Watanabe et al. 2016).
MChR1 of Mesostigma viride (F8UVI5)
Anion-specific light-gated channel rhodopsin of 438 aas, Acr1, lacking measurable cation transport capability (Govorunova et al. 2015).
Arc1 of Guillardia theta
Anion-specific light-gated channel rhodopsin of 438 aas, Acr2, lacking measurable cation transport capability (Govorunova et al. 2015). Two conserved carboxylates, E159 and D230, play roles in the anion transport activity of ACR2 (Kojima et al. 2018).
Acr2 of Guillardia theta
Homologue of anion-specific light-gated channel rhodopsin of 461 aas and 7 putative TMSs, lacking apparent channel activity (Govorunova et al. 2015).
Acr homologue of Guillardia theta
Synthetic anion-specific channelrhodopsin of 307 aas and 7 TMSs, derived from an anion channel of Guillardia theta (Govorunova et al. 2018).
Channelrhodopsin of Guillardia theta