Article

ISSN 0006-2979, Biochemistry (Moscow), 2026, Vol. 91, No. 8, pp. 1359-1375 © Pleiades Publishing, Ltd., 2026.

1359

REVIEW

Photoreversible Reactions

of Visual and Microbial Rhodopsins

Mikhail A. Ostrovsky

1,2

, Olga A. Smitienko

1,a

*, and Tatiana B. Feldman

1,2

Emanuel Institute of Biochemical Physics, Russian Academy of Sciences, 119334 Moscow, Russia

Faculty of Biology, Lomonosov Moscow State University, 119991 Moscow, Russia

e-mail: djolia@gmail.com

Received April 19, 2026

Revised May 26, 2026

Accepted May 30, 2026

Abstract— The review discusses experimental findings and theoretical studies of photoreversible transforma-

tions of retinal-containing proteins (rhodopsins). Understanding these processes is essential for elucidating

the mechanisms of photochemical and conformational transformations of these proteins. Particular emphasis

is placed on visual rhodopsins, as photoreversibility plays a crucial role in its physiological regeneration

in the rhabdoms of invertebrates and the cone photoreceptors of the vertebrate retina. In recent years,

interest in the mechanisms of photoreversible reactions in retinal-containing proteins has grown substan-

tially, driven by their potential applications as active components in optoelectronic devices and biosensors.

DOI: 10.1134/S0006297926601267

Keywords: retinal-containing proteins, visual rhodopsin, chromophore, photoreversible reactions, photoisom-

erization, photoregeneration, dark adaptation

* To whom correspondence should be addressed.

INTRODUCTION

The diversity of retinal-containing proteins (rho-

dopsins) is extremely large [1-12]. They are found

in all domains of life. Type  I (microbial) rhodopsins

are characteristic of bacteria, archaea, lower eukary-

otes, and giant viruses, while type  II rhodopsins are

characteristic of higher animals. Type  I rhodopsins

perform photoenergetic (ion pumps), photoreceptor

(sensory rhodopsins and ion channels), and photoen-

zymatic functions, while type  II rhodopsins perform

primarily photoreceptor (visual and non-visual rho-

dopsins) and photoenzymatic functions. These types

appear to have independent origins.

Each of the diverse functions of retinal-contain-

ing proteins is initiated by photoisomerization of

the chromophore group of retinal – from all-trans

to 13-cis in microbial rhodopsins and from 11-cis to

all-trans in animal rhodopsins  [1,  2]. The character-

istics of this photochemical reaction were studied in

detail after its discovery in the laboratory of J.  Wald

in the late 1950s in bovine visual rhodopsin  [13].

It has now been established that this is a uniquely

rapid and selective reaction, occurring with a high

quantum yield in an electronically excited state [1-4,

8, 9]. Interactions between retinal and its protein en-

vironment (protein catalysis) play a major role in this

reaction. The dynamics and sequence of formation

of photoreaction products have been characterized

in detail in both microbial and visual rhodopsins.

The reverse photoreactions, initiated by the absorp-

tion of a second quantum of light by these products,

have been described less thoroughly. However, they

are of significant interest.

Reverse photoreactions of rhodopsins, initiated

from the early products of the forward photoreac-

tion, can serve as an important tool for studying the

interactions of retinal with its immediate protein en-

vironment and conformational changes in the protein

part of the molecule (opsin). Reverse photoreactions

of some microbial rhodopsins, initiated at later stag-

es, are used by the cell to more precisely discriminate

between the wavelengths of incident light, which is

important for its adaptation. Inthe case of visual rho-

dopsins, reverse photoisomerization of all-trans reti-

nal to the 11-cis form is of fundamental physiological

OSTROVSKY et al.1360

BIOCHEMISTRY (MOSCOW) Vol. 91 No. 8 2026

importance for photoreceptor cells – the rhabdoms

of invertebrates and the cones of vertebrate retinas.

This involves the return (regeneration) of rhodopsin

to its original state, which is necessary for restoring

normal physiological activity in the photoreceptor

cell.

This review examines the forward and reverse

photoreactions of visual rhodopsins, initiated at dif-

ferent stages of their photoconversion. A comparison

is made of the reverse photoreactions of bovine vi-

sual rhodopsin and bacteriorhodopsin, initiated from

the early products of their forward photoreactions.

The review also discusses the light-dependent and

light-independent pathways for regenerating visual

rhodopsins.

PHOTOCHEMISTRY

OF RETINAL-CONTAINING PROTEINS

Visual rhodopsins. Visual rhodopsins (type  II

rhodopsins) are transmembrane proteins that utilize

light energy to initiate the phototransduction cascade.

Light absorption is mediated by a chromophore group,

11-cis retinal, located in the chromophore-binding

pocket of the apoprotein opsin. The chromophore

is covalently bound to a lysine residue, forming a

protonated Schiff base linkage (Fig.  1). Visual rho-

dopsins are members of the G protein-coupled recep-

tor (GPCR) superfamily. The fundamental functional

event in these proteins is retinal photoisomerization

(Fig.  1), which occurs within 50-80  fs and proceeds

with a quantum yield of 0.6-0.7. These exceptional

parameters result from both intrinsic chemical prop-

erties of retinal and the influence of the surround-

ing protein environment. In contrast, when retinal

is not embedded within a protein matrix, such as

in the gas phase [14] or in a solution [15, 16], the

photoisomerization reaction is approximately an or-

der of magnitude slower and exhibits a substantially

lower quantum yield of only 0.1-0.3[17]. Theprimary

role of the photoreaction is to store the energy of

the absorbed photon in a form of a strained, highly

twisted all-trans retinal configuration. Subsequent re-

laxation of such distorted chromophore releases the

stored energy and induces structural rearrangements

in its immediate protein environment, which triggers

global structural changes in the rhodopsin molecule

necessary for its functioning.

The most extensively studied visual pigments in

vertebrates and invertebrates are bovine and octopus

rhodopsins, respectively. Bovine rhodopsin is located

in the outer segments of rod photoreceptor cells and

is responsible for scotopic (twilight and low-light) vi-

sion. Octopus rhodopsin is found in the rhabdomeres

of photoreceptor cells and serves as a primary visual

pigment mediating visual perception in this species.

Visual rhodopsins of vertebrates are monostable pro-

teins, meaning that under physiological temperatures

in the dark, they exist in a single thermally stable

state. Upon light absorption, these proteins undergo

a sequence of mainly irreversible transformations,

collectively referred to as photolysis (Fig.  2a). In con-

trast, invertebrate visual rhodopsins are bistable.

When exposed to light, they form an alternative ther-

mally stable state and can absorb a second photon,

which reverts them to the original state (Fig.  2b). This

mechanism enables physiological photoregeneration

of visual rhodopsin in invertebrates.

Vertebrate visual rhodopsin. Let’s consider the

stages of photolysis of vertebrate (bovine) visual rho-

dopsin in more detail. Following the photoinduced

11-cis

  →  all-trans isomerization of the retinal chro-

mophore, the protein progresses through a series of

intermediate conformational states. In the final stage

of photolysis, the Schiff base linkage between retinal

and opsin is hydrolyzed, leading to the release of

all-trans retinal from the chromophore-binding pock-

et and its subsequent removal from the photorecep-

tor cell. Rhodopsin intermediate states or products of

photoconversion have been characterized using a va-

riety of spectroscopic techniques, including low-tem-

perature absorption spectrophotometry, time-resolved

absorption and fluorescence spectroscopy (Fig.

  2a).

The cis → trans photoisomerization of retinal in

bovine rhodopsin proceeds with a quantum yield of

0.65-0.67 [27,  28] and is completed within 50-80  fs

[29-33]. This process leads to the formation of the

first photoproduct, photorhodopsin, through a tran-

sition from the first excited electronic state  (S

)

Fig.  1. Structures of the chromophore group in type  II rho-

dopsins: protonated Schiff base of 11-cis and all-trans reti-

nal in the rhodopsin dark state  (1) and after photon absorp-

tion  (2), respectively. Reactive C

double bond is shown

in bold; photochemical transitions are indicated by arrows.

PHOTOREVERSIBLE REACTIONS OF RHODOPSINS 1361

BIOCHEMISTRY (MOSCOW) Vol. 91 No. 8 2026

Fig.  2. Photoactivated processes in visual rhodopsins: a)photolysis of monostable bovine rhodopsin [2, 7, 18-20]; b)  photocy-

cle of bistable octopus rhodopsin [21-26]. The schemes show the sequence of intermediate states formed during rhodopsin

photoactivation, including the room temperature maxima of their α-absorption bands, characteristic formation times, and

minimum temperatures at which these intermediates were detected by low-temperature spectrophotometry. In panel  (b),

the minimum temperature values are given for squid visual rhodopsin  [18]. Photochemical transitions are indicated by

hν symbols together with respective quantum yields (φ). Rhodopsin, dark state of the visual pigment; Photo, photorho-

dopsin; Batho, bathorhodopsin; BSI, blue shifted intermediate (intermediate with the absorption spectrum shifted to the

blue region); Lumi, lumirhodopsin; Meta, metarhodopsin; Prime, primerhodopsin; Meso, mesorhodopsin; Meta’, short-lived

metarhodopsin; I

and I

, intermediate products of phototransition from acid metarhodopsin to octopus rhodopsin. States

stable at physiological temperatures are framed. See the text for details.

OSTROVSKY et al.1362

BIOCHEMISTRY (MOSCOW) Vol. 91 No. 8 2026

to the ground state (S

). This extremely fast transi-

tion is enabled by a conical intersection between the

and S

potential energy surfaces, which provides

an efficient pathway for the ultrafast conversion of

photon energy into chemical energy [3, 34,35]. Inre-

actions of this type, the product is formed as a result

of coordinated (coherent) movement of nuclei in the

chromophore. The dynamics of coherent reactions

is highly sensitive to the chromophore structure, its

local environment, and initial conditions of photoex-

citation. The protein environment of retinal actively

modulates this process by tuning the absorption max-

imum as well as the selectivity, rate, and quantum

yield of photoisomerization. In this way, the protein

environment effectively catalyzes such ultrafast and

highly efficient photochemical reaction. Steric con-

straints within the chromophore-binding site and

electrostatic interactions between retinal and sur-

rounding amino acid residues, particularly the nega-

tively charged counterion Glu113, play an important

role in this regulation [2, 7].

Photorhodopsin is finally formed within 200 fs

[36]. It contains vibrationally excited all-trans retinal

that remains highly distorted within the confined

volume of the binding pocket [37, 38]. Subsequent

intermediates, bathorhodopsin [30, 33, 36, 39] and

the blue-shifted intermediate (BSI) [19, 40, 41], arise

during vibrational relaxation of the chromophore and

its “straightening” in the β-ionone ring region. Later

stages of rhodopsin photolysis involve conformational

rearrangements of opsin and include the formation

of lumirhodopsin and metarhodopsins I, II, and III

(Fig.  2a) [2, 7, 20]. Metarhodopsin II is the key signal-

ing state, as it is capable of activating the G protein

transducin. Its formation is accompanied by depro-

tonation of the retinal Schiff base and proton transfer

to the acceptor residue Glu113, which triggers major

conformational changes in rhodopsin. On a minute

timescale, metarhodopsin I can reversibly convert

into metarhodopsinIII through the thermal anti → syn

isomerization of the C

=N Schiff base linkage (Fig.  1)

[42]. Metarhodopsins II and III subsequently decay

slowly through hydrolysis of the Schiff base linkage,

yielding the apoprotein opsin and all-trans retinal.

Invertebrate visual rhodopsins. The photoreac-

tions of invertebrate visual rhodopsins have been in-

vestigated primarily in cephalopod mollusks [21-26,

43-46]. The photocycle of octopus rhodopsin is il-

lustrated in Fig.  2b. Its early photochemical events

closely resemble those observed in vertebrate rho-

dopsins. Following photon absorption, chromophore

photoisomerization proceeds coherently within ~80  fs

[26] with a quantum yield of 0.69  [22]. This process

generates the first photoproduct, primerhodopsin,

which is analogous to the vertebrate photorhodopsin

and converts to bathorhodopsin within 2-4  ps [46].

Subsequent intermediates arise from progressive ret-

inal planarization and accompanying conformational

changes in opsin. These transitions lead to the for-

mation of lumirhodopsin, mesorhodopsin, short-lived

and ultimately thermally stable acid metarhodopsins

[21, 23-25]. Acid metarhodopsin represents the sig-

naling state of the photocycle; however, unlike verte-

brate metarhodopsin II, it retains a protonated Schiff

base chromophore, which likely contributes to its

enhanced stability and resistance to photolysis  [47].

Atelevated pH, acid metarhodopsin can convert to its

alkaline form through the Schiff base deprotonation.

Photon absorption by acid metarhodopsin initiates

the reverse photoreaction, regenerating the original

rhodopsin state. Importantly, this reverse transition

does not trigger photoreceptor cell excitation (see the

section “Photoreversibility at late stages of rhodopsin

conversion”).

Microbial rhodopsins. The primary photore-

action of microbial rhodopsins (all-trans → 13-cis

isomerization of retinal) occurs on a slightly longer

timescale than in visual rhodopsins, typically within

300-800  fs, and proceeds with a quantum yields of

0.3-0.7 [2, 3, 8]. Subsequent processes are closed in

a cycle, with one of the last stages involving thermal

retinal re-isomerization. This photocycle has been

characterized in greatest detail for bacteriorhodop-

sin from the haloarchaeon Halobacterium salinarum

[48]. The early intermediates J and K of this bacteri-

orhodopsin are considered analogs of photorhodopsin

and bathorhodopsin, respectively [9, 49-52]. A central

intermediate of the bacteriorhodopsin photocycle is

the long-lived M state, which is essential for the pro-

tein’s photoenergetic function as a light-driven pro-

ton pump. The created proton gradient supplies the

motive force for ATP synthesis [53]. Formation of the

M intermediate is accompanied by deprotonation of

the retinal Schiff base and substantial conformation-

al rearrangements within the seven-transmembrane

α-helical protein scaffold, reminiscent of structural

changes associated with the formation of metarho-

dopsin II in vertebrate visual rhodopsins.

In some microbial rhodopsins, the reverse pho-

toconversion from 13-cis to all-trans retinal is also

physiologically important. It enables these proteins to

function as photochromic sensors for regulating pho-

totactic responses and gene expression [54-58].

PHOTOREVERSIBILITY OFRHODOPSIN

PHOTOCONVERSION PRODUCTS

Retinal-containing proteins exhibit photorevers-

ibility, i.e., the ability to return to their original state

upon light absorption by the products of their pho-

toconversion. This reverse photoreaction is known

PHOTOREVERSIBLE REACTIONS OF RHODOPSINS 1363

BIOCHEMISTRY (MOSCOW) Vol. 91 No. 8 2026

as photoregeneration. Photoreversibility is observed

both in  vitro [9, 33, 59-61] and in  vivo [62-66].

Photoreversibility at early photochemical stages

of rhodopsin conversion. Photoreversibility at early

photochemical stages has been investigated experi-

mentally mainly in bovine visual rhodopsin [9, 33,

66-77] and bacteriorhodopsin [33, 60, 78-84], as well

as studied using computational modeling methods

[82, 85-87].

Low-temperature studies of rhodopsin reverse

photoreactions. The first studies of the early stages

of rhodopsin photolysis employed low-temperature

spectrophotometry, infrared  (IR) spectroscopy, and

Raman spectroscopy. These approaches are based

on the ability to stabilize individually each product

of rhodopsin photolysis (except for photorhodopsin

and BSI) at cryogenic temperatures, when the ther-

mal energy is insufficient to overcome the activation

barriers required for progression to subsequent states

(Fig.  2, a  and  b). Under such conditions, rhodopsin is

typically studied in detergent extracts. Irradiation

generates a photostationary mixture containing the

original state and the products of both forward and

reverse photoreactions, with their relative proportions

determined by the temperature and illumination con-

ditions. For example, bovine rhodopsin irradiated at

liquid nitrogen temperature (−196°C) with light in

the 450-500  nm range yields a mixture of rhodopsin

(λ

max

  =  504nm), bathorhodopsin (λ

max

  =  543  nm), and

isorhodopsin (λ

max

  =  490  nm), which contain 11-cis,

all-trans, and 9-cis retinal, respectively [67-69, 71-73,

75, 88] (Scheme  1). Illumination with longer-wave-

length light (550-600  nm) shifts the equilibrium to-

ward rhodopsin and isorhodopsin, whose relative

abundance (about 90  :  10) reflects the quantum yields

of their formation from bathorhodopsin (0.5 and 0.05,

respectively) [71]. These low-temperature experiments

on forward and reverse photoreactions provided

some of the earliest evidence that the primary event

in vision involves retinal isomerization and that ba-

thorhodopsin already contains all-trans retinal  [67].

These observations were later corroborated by Raman

spectroscopy [89], as well as by spectrophotometric

and picosecond spectroscopic studies of rhodopsin re-

constituted with retinal analogs [90, 91].

When lumirhodopsin was excited at low tem-

peratures, formation of rhodopsin was observed to-

gether with a minor fraction of isorhodopsin [67, 70,

72], as confirmed by IR spectroscopy [75] and HPLC

analysis [69]. However, the photoregeneration of rho-

dopsin from lumirhodopsin is less efficient than from

bathorhodopsin [69, 72], despite a reported quantum

yield of 0.5[70]. Reverse photoreactions at later stag-

es of rhodopsin photolysis will be discussed in the

section “Photoreversibility at late stages of rhodopsin

conversion.”

Investigation of reverse photoreactions of rhodop-

sin and bacteriorhodopsin using time-resolved spec-

troscopy. The photoreversibility of rhodopsin from

bathorhodopsin was first demonstrated in real time

using the nanosecond flash photolysis method [92].

Excitation with laser pulses (532 and 605  nm) sepa-

rated by 93  ns led to a significant decrease in bathor-

hodopsin absorption, indicating reverse phototransi-

tion to rhodopsin. Subsequently, femtosecond studies

with a time resolution of 300  fs[74] demonstrated the

possibility of reverse phototransition not only from

bathorhodopsin, but also from photorhodopsin. More

detailed investigations using femtosecond absorption

spectroscopy with a 30-fs time resolution [33, 76, 77]

employed two excitation pulses at 500 and 620  nm

with delays of 0.2-3.8  ps. It was shown that the pho-

totransition from bathorhodopsin or photorhodopsin

to rhodopsin occurred without formation of isorho-

dopsin or other side products (Scheme  2). The quan-

tum yields of reverse transitions from photorhodopsin

and bathorhodopsin were similar (0.14 and 0.16, re-

spectively) and significantly lower than those demon-

strated by low-temperature methods (Scheme  1). The

time of the photoreaction initiated from photorho-

dopsin was comparable to the time of forward pho-

toreaction, consistent with theoretical calculations

[85, 86]. Finally, the possibility of in  vitro ultra-fast

reverse phototransitions in the visual rhodopsin mol-

ecule [76, 77] was confirmed by elegant experiments

conducted at a femtosecond time resolution in  vivo on

dark-adapted mouse eyes in [66].

Scheme  1. Phototransitions occurring in the bovine visual

rhodopsin at −196°C under steady-state illumination. Rh

504

dark state of the visual rhodopsin; Batho

543

, bathorhodop-

sin; isoRh

490

, isorhodopsin. Subscripts indicate the low-tem-

perature absorption maxima; the quantum yields of the

corresponding phototransitions are indicated next to the

arrows [68, 71].

Scheme  2. Reverse photoreactions of bovine visual rho-

dopsin initiated from primary products at room tempera-

ture [29, 76, 77, 86]. Rh

498

, dark state of visual rhodopsin;

Photo

570

, photorhodopsin; Batho

535

, bathorhodopsin. Sub-

scripts indicate the absorption maxima; solid arrows show

phototransitions; dashed arrow indicates thermal transition.

Thecorresponding transition times and quantum yields are

shown next to the arrows.

OSTROVSKY et al.1364

BIOCHEMISTRY (MOSCOW) Vol. 91 No. 8 2026

Table 1. Quantum yields(φ) and characteristic times for cis → trans and trans → cis photoisomerization of ret-

inal with protonated Schiff base in a free form and as a component of bovine and octopus visual rhodopsins

and bacteriorhodopsin at room temperature

Retinal in various environments Forward photoreaction Reverse photoreaction

φ t, ps φ t, ps

11-cis → all-trans all-trans → 11-cis

Free form 0.18-0.24*

[17, 96]

0.4**

[14]

0.14-0.25*

[17]

3**

[14]

Bovine rhodopsin 0.65-0.67

[27, 28]

0.05-0.08

[29, 31-33]

0.16

[77]

< 0.1

[77]

Octopus rhodopsin 0.69

[22]

0.08

[26]

0.36***

[71]

–

all-trans → 13-cis 13-cis → all-trans

Bacteriorhodopsin 0.64

[79]

0.45-0.5

[33, 49-51]

0.81-1

[33, 83, 84]

0.19-0.3, 1.1-1.7,

11-16

[83, 84]

Note. Parameters of reverse photoreactions of rhodopsin and bacteriorhodopsin are given for reactions were initiated from

bathorhodopsin and K products, respectively.

*, in solvents; **, in vacuum; ***, for squid rhodopsin at 77  K.

Experimental studies of reverse photoreaction

were also carried out on bacteriorhodopsin, starting

from the first products of forward photoreaction J

andK [33, 83,84]. Reverse phototransition from K was

demonstrated with a substantially higher quantum

yield than that observed in visual rhodopsin, reach-

ing 0.81 in [33] and ~1 in [79, 80, 83, 84, 93]. In [84],

the dynamics of reverse photoreaction from K was

investigated following excitation conducted 5  ps after

the first laser pulse. The excited state K* was found

to decay through three channels with characteristic

lifetimes of 0.19, 1.1, and 16  ps, ultimately regener-

ating the original state of bacteriorhodopsin. Compa-

rable kinetic behavior was observed upon excitation

of K at a time delay of 60 ps [83]. The heterogeneity

of the excited state K* might be determined by the

heterogeneity of its chromophore-binding site, arising

from the presence of multiple configurations with

different nature of retinal–protein interactions, as it

was shown for the microbial rhodopsin KR2 from the

bacterium Krokinobacter eikastus [94, 95]. Since the

forward photoreaction is dominated by a single ultra-

fast component (~0.5  ps) [33, 49-51] and has a lower

quantum yield of 0.64 [79], it was concluded that the

reverse photoreaction initiated from the product K is

more efficient, albeit not necessarily faster in kinetics.

The quantum yields and the formation times for

the primary products of forward and reverse photo-

reactions in visual rhodopsin and bacteriorhodopsin

are presented in Table  1.

The dynamics of forward and reverse photoreac-

tions of visual rhodopsin and bacteriorhodopsin from

their primary products have been investigated in de-

tail using quantum mechanical/molecular mechanical

(QM/MM) calculations [82, 86]. In [86], it was demon-

strated that the reverse photoreaction initiated from

bathorhodopsin involves a reversal motion of the ret-

inal chromophore, including torsional deformations

along the C

, C

, and C

bonds in direc-

tions opposite to those observed during the forward

photoreaction. The quantum yields of both forward

and reverse photoreactions were found to depend on

the phase and amplitude of some vibrations of the

retinal at the moment of transition from the S

ex-

cited electronic state to the ground state. These are

hydrogen-out-of-plane (HOOP) vibrations at the car-

bon atoms C

and C

The authors of [82] compared the photoexcited

states of visual rhodopsin, bacteriorhodopsin, and

their primary photoproducts – bathorhodopsin and K

intermediate. Using available X-ray structures, they

calculated the absorption spectra of these species

and identified the retinal vibrational modes active in

the S

 → S

transition, and therefore, governing the

earliest stages of the excited state dynamics. Analy-

sis of the obtained data revealed that photoexcitation

of rhodopsin, bacteriorhodopsin, and the K interme-

diate selectively enhances the activity of the vibra-

tional modes directly involved in photoisomerization.

These modes are associated with the C

bond

PHOTOREVERSIBLE REACTIONS OF RHODOPSINS 1365

BIOCHEMISTRY (MOSCOW) Vol. 91 No. 8 2026

Fig.  3. Comparison of the vibrational activity of the protonated Schiff base of retinal in the initial state of rhodopsin and

in its primary photoproduct immediately after excitation to the S

electronic state  [82]: a)  bovine visual rhodopsin (left)

and its product bathorhodopsin (right); b)  bacteriorhodopsin (left) and its product K (right). The arrows in the structures

indicate the most active high-frequency valence C=C vibrations of the retinal polyene chain and HOOP vibrations; the

photoisomerizing C

  (a) and C

  (b) bonds are shown with an oval; the quantum yields of forward and reverse

photoreactions are given next to the arrows indicating the phototransitions.

in rhodopsin and the C

bond in bacteriorho-

dopsin and the K intermediate [87]. Simultaneous in-

phase excitation of these reactive vibrational modes

may facilitate phase synchronization as the molecule

passes through the S

conical intersection, there-

by contributing to the high quantum yields of the

forward photoreactions of rhodopsin and bacterior-

hodopsin, as well as of the reverse photoreaction of

bacteriorhodopsin originating from the K intermedi-

ate (Table  1) [3, 35]. This effect arises from specific

interactions between retinal and its immediate pro-

tein environment in the chromophore-binding pock-

et of opsin. The vibrational modes activated upon

excitation of rhodopsin are shown in Fig.  3a (left)

and include high-frequency valence vibrations of the

bond together with HOOP vibrations of hydro-

gen atoms attached to carbons C

and C

. The cor-

responding modes for bacteriorhodopsin and its K

intermediate are presented in Fig.  3b. Notably, in the

K intermediate, excitation induces not only reactive

vibrations associated with the C

bond but also

HOOP vibrations at the C

bond (Fig.  3b, right).

This additional vibrational activity may underlie a

complex mechanism of reverse photoreaction in bac-

teriorhodopsin, as mentioned above [83, 84].

In bathorhodopsin, photoexcitation non-selec-

tively activates vibrations of the C

and C

bonds and HOOP vibrations of hydrogen atoms at the

carbon atoms C

and C

(Fig. 3a, right panel; see

also [82]). Bathorhodopsin does not exhibit a dom-

inant high-frequency vibrational mode; instead, its

excited state is characterized by the HOOP vibrations

at two different double bonds, which may reduce the

coherence of retinal intramolecular motions during

the reverse photoreaction, thereby contributing to a

low quantum yield of bathorhodopsin phototransition

to rhodopsin (Table 1).

Thus, both experimental studies and QM/MM

simulations of forward and reverse photoreactions

indicate a more finely tuned interaction between ret-

inal and its protein environment in visual rhodopsin

than in bacteriorhodopsin [9, 82]. One manifestation

of this optimization is a substantially lower probabil-

ity of reverse photoreactions (which are functionally

unfavorable for phototransduction) in visual rhodop-

sin compared to bacteriorhodopsin. These differenc-

es in the photophysical behavior of retinal isomers

in visual (type II) rhodopsin and bacteriorhodopsin

(type I rhodopsin) are likely related to their distinct

biological functions. In bacteriorhodopsin, an efficient

reverse photoreaction from the K intermediate is

not essential for the proton-pumping activity during

the photocycle. In contrast, suppression of reverse

photoisomerization in the primary photoproducts

of visual rhodopsin – photorhodopsin and bathor-

hodopsin – enhances the reliability of the forward

OSTROVSKY et al.1366

BIOCHEMISTRY (MOSCOW) Vol. 91 No. 8 2026

photoreaction that initiates the phototransduction

cascade.

Photoreversibility at late stages of rhodop-

sin conversion. Reverse photoreactions of vertebrate

rhodopsins. The products of photoconversion of ver-

tebrate visual rhodopsin formed within the time

ranges from microseconds to seconds also retain

photoactivity [42, 64, 65, 72, 75, 88, 97-106]. Howev-

er, photoregeneration of rhodopsin from these later

intermediates is considerably less efficient than from

earlier photoproducts because it requires selective

all-trans → 11-cis isomerization in the altered chro-

mophore-binding site, followed by substantial con-

formational rearrangements of opsin. For example,

bathorhodopsin generated by irradiation of bovine

rhodopsin at −196°C can be converted back to rho-

dopsin almost quantitatively upon re-irradiation with

long-wavelength light (see “Photoreversibility at ear-

ly photochemical stages of rhodopsin conversion”).

Incontrast, photoregeneration from metarhodopsin  II

in bovine and frog rhodopsins reaches no more than

50%, as demonstrated both in  vitro at −20°C [97] and

in single photoreceptor cells at physiological tempera-

tures [105]. The efficiency of rhodopsin regeneration

from metarhodopsin III is even lower (less than 37%

depending on temperature) [42]. Studies examining

the effects of duration and intensity of irradiation

light on rhodopsin bleaching, conducted both in  vitro

[107] and in  vivo [62, 63], have shown that a brief

light flash (≤1  ms) never bleaches more than 50-75%

of rhodopsin, irrespective of light intensity. Based on

these observations, it was concluded that the pho-

tolysis products formed before metarhodopsin  II are

completely photoreversible, whereas later interme-

diates exhibit only partial photoreversibility. Impor-

tantly, regenerated rhodopsin remains physiologically

functional. Furthermore, experiments in bovine rod

outer segments [72] demonstrated that recovery of

a functional rhodopsin, not amenable to phosphor-

ylation, occurs much less efficiently from metarho-

dopsins I and II than from the earlier intermediates

bathorhodopsin and lumirhodopsin.

Using low-temperature IR spectroscopy of rod

outer segments [75] and flash photolysis of detergent

extracts of bovine rhodopsin [100], metarhodopsin I

was shown to be isochromic, i.e., existing in two spec-

trally indistinguishable forms. One of these forms

is capable of regenerating into rhodopsin with the

formation of isorhodopsin and several minor pho-

toproducts, whereas the other lacks photoreactivity.

Subsequent studies of photoreactions of frog metar-

hodopsin I by absorption spectrophotometry at −25°C

further demonstrated the formation of two isochro-

mic rhodopsin species, one of which remained stable

upon warming to room temperature, while the other

underwent decay [101, 102].

Photon absorption by metarhodopsin II results

predominantly in the formation of metarhodopsinIII,

together with smaller amounts of rhodopsin and isor-

hodopsin, as has been observed in both detergent ex-

tracts of rhodopsin and suspensions of photoreceptor

membranes [64, 98, 104, 106, 108]. Similar data have

been reported for albino rat eyes[64] and intact frog

rod cells[105]. Irradiation of metarhodopsinIII in bo-

vine photoreceptor membranes primarily drives its

conversion to metarhodopsin II via metarhodopsin I,

while producing lesser amounts of rhodopsin and

possibly isorhodopsin [42, 88, 104]. These findings

indicate that, during reverse photoreactions initiated

at the later stages of rhodopsin photolysis (beginning

with metarhodopsin  II), the anti ↔ syn photoisomer-

ization of the C

=N Schiff base becomes the domi-

nant photochemical process, whereas the trans → cis

photoisomerization at the C

and/or C

bonds

plays a comparatively minor role (Fig.  1).

It should be noted that the data on photorevers-

ible reactions of metarhodopsins remain contradic-

tory. For example, the restoration of the absorption

band in the 500  nm region is not necessarily associ-

ated with the return of rhodopsin to a physiological-

ly active state capable of initiating phototransduction

[20,  72], contrary to the assumptions made in the

earliest studies of rhodopsin reverse photoreactions.

Resolving this issue requires the application of vari-

ous methods, including HPLC analysis, IR spectrosco-

py, and time-resolved laser spectroscopy (for details,

see [7]).

Photoreversible reactions occurring at different

stages of rhodopsin photolysis provide a valuable

tool for investigating light-induced conformational

changes in the protein. For example, in our studies

of the rhodopsin activation mechanism using satura-

tion-transfer electron paramagnetic resonance (EPR)

spectroscopy with spin labels covalently attached

to the hydrophilic sulfhydryl groups of Cys140 and

Cys316 located in the second and third cytoplasmic

loops, respectively, we were the first to observe pho-

toreversible changes in the conformational mobility of

opsin [109-111]. The mobility of the labels increased

during rhodopsin transition to metarhodopsin II

and decreased during the reverse metarhodopsin II

phototransition to a mixture of products, including

photoregenerated rhodopsin and metarhodopsin III.

These observations indicated that light activation ex-

poses opsin regions involved in G protein binding.

Subsequent EPR studies, particularly those employ-

ing pulsed EPR techniques, together with numerous

complementary approaches, have revealed not only

rearrangements of the cytoplasmic loops but also

movements of the transmembrane α-helices, which

are critical elements of signal transmission from ac-

tivated rhodopsin to the G protein (see review [7]).

PHOTOREVERSIBLE REACTIONS OF RHODOPSINS 1367

BIOCHEMISTRY (MOSCOW) Vol. 91 No. 8 2026

Reverse photoreactions of invertebrate rhodop-

sins. Bistable rhodopsins of invertebrates are capa-

ble of efficient photoregeneration, which is initiated

when a second photon is absorbed by the thermally

stable acid metarhodopsin (Fig.  2b). In invertebrates,

photoreversibility has been extensively characterized

in the rhodopsins of the octopus, squid, and jumping

spider (JSR1) [22, 25, 112-116]. In octopus rhodopsin,

sequential blue- and orange-light irradiation at room

temperature initiated multiple rounds of rhodopsin

photoconversion to acid metarhodopsin and back on

both minute [112] and second [113] timescales. The

quantum yield of the reverse photoreaction from

acid metarhodopsin to rhodopsin (0.43 for octopus

rhodopsin) was lower than that of the forward pho-

toreaction [22].

Higher time-resolution experiments have re-

vealed that photoregeneration of invertebrate rho-

dopsins involves one photochemical step followed by

two thermal (dark) reactions [25, 115, 116]. The ini-

tial photochemical event rapidly reisomerizes retinal

to the 11-cis configuration, generating an intermedi-

ate with the absorption spectrum closely resembling

that of the dark-state rhodopsin[25]. Twosubsequent

thermal transitions (formation of the second product

and of the original state of rhodopsin) are thought

to reflect conformational rearrangements in protein

regions distant from the chromophore. In octopus

rhodopsin, these reactions proceed within 490  μs and

2.6  ms [25], whereas in JSR1 they are markedly fast-

er, occurring within 2  μs and 60  μs, respectively  [116].

In squid rhodopsin, these thermal transitions are ac-

companied by deprotonation and subsequent repro-

tonation of the retinal Schiff base  [115].

It was shown that if octopus rhodopsin undergoes

a significant increase in volume during the forward

photoreaction, reflecting formation of the function-

al metarhodopsin state that initiates phototransduc-

tion, the reverse photoreaction is accompanied by a

decrease in the protein volume back to its original

value  [25]. Importantly, the reverse photoreaction does

not trigger the phototransduction cascade [117, 118].

The photoreactions of squid alkaline metarhodop-

sin have also been investigated [114]. Upon illumina-

tion, this protein forms a photoproduct that slowly

reverts to rhodopsin in the dark. However, prolonged

repeated irradiation can drive the protein back to the

metarhodopsin state.

Reverse photoreactions of microbial rhodopsins.

Photoreversibility (13-cis  →  all-trans transition) on the

microsecond-to-millisecond timescale has also been

studied in type  I rhodopsins, including bacteriorho-

dopsin [60, 61], as well as in photochromic sensory

rhodopsins (SRIs) from halophilic archaea and bacte-

ria [56, 57], cyanobacterial sensory rhodopsin from

Anabaena  sp. (ASR) [54, 55], and channelrhodopsins

from the green alga Chlamydomonas reinhardtii and

related species (ChR1 and ChR2) [56, 58]. In bacteri-

orhodopsin, reverse photoreactions initiated from the

M, N, and O intermediates were found to be highly

complex, proceeding through the formation of multi-

ple transient states [60, 61].

PHOTOREVERSIBILITY OFRHODOPSIN

INVIVO ANDREGENERATION

OFVISUAL RHODOPSINS

Phototransduction in both invertebrates and

vertebrates is initiated by the photoisomerization of

11-cis retinal to its all-trans configuration. This pho-

tochemical event induces conformational changes in

opsin, triggering a signaling cascade that transmits

and amplifies the light signal and ultimately leads

to the generation of the photoreceptor potential. Be-

cause phototransduction relies on this forward photo-

reaction, all-trans retinal must subsequently be con-

verted back to the 11-cis form, and rhodopsin must

be restored to its original functional state through a

process known as regeneration.

Two distinct strategies have evolved to regener-

ate the thermodynamically less favorable 11-cis reti-

nal of visual pigments. The first, and evolutionarily

more ancient, strategy utilizes light energy to drive

retinal re-isomerization, as observed in many inver-

tebrate rhodopsins. A critical requirement of this

mechanism is preservation of the covalent linkage

between all-trans retinal and opsin following photo-

activation. Absorption of the second photon converts

the chromophore back to the 11-cis configuration,

thereby promoting restoration of the native opsin

conformation (see “Photoreversibility at late stages

of rhodopsin conversion”).

The second strategy for restoring 11-cis retinal

has evolved in response to the loss of retinal from

rhodopsin as a result of photolysis, which is charac-

teristic of vertebrate visual pigments. After hydroly-

sis of the covalent bond between opsin and all-trans

retinal, the chromophore dissociates from opsin and

is ultimately removed from the photoreceptor cell.

The key innovation of this evolutionarily more re-

cent strategy is the development of a light-indepen-

dent biochemical pathway – the visual (retinoid) cy-

cle, which involves retinal pigment epithelium (RPE)

cells and, potentially, Müller glial cells. This cycle me-

diates the multistep regeneration of the visual pig-

ment dark-adapted state by converting retinal back

to its 11-cis configuration required for the chromo-

phore function. Because rhodopsin-containing photo-

receptor discs in rod outer segments have a lifespan

of approximately 10-12 days, individual rhodopsin

molecules can repeatedly absorb photons, undergo

OSTROVSKY et al.1368

BIOCHEMISTRY (MOSCOW) Vol. 91 No. 8 2026

photolysis, and be regenerated. The evolution of this

complex and highly reliable regeneration mechanism

has enabled vertebrates to minimize the accumula-

tion of “errors” in structures of both retinal and opsin

that could arise during photocycles in invertebrates.

Indeed, in many invertebrates, retinal undergoes re-

peated photoisomerization while remaining associat-

ed with the same rhodopsin molecule. Such cycling

may promote the formation of side products contain-

ing alternative retinal isomers, potentially inducing

nonphysiological conformational changes in opsin.

In vertebrates, dissociation of all-trans retinal from

opsin permits chromophore regeneration in the visu-

al (retinoid) cycle, ensuring that retinal is re-isomer-

ized to the 11-cis form required for proper assem-

bly and function of the chromophore-binding site

in opsin.

It should be noted that both strategies can co-

exist within the same organism. Reverse trans → cis

photoisomerization is also present in vertebrates,

where it contributes to the regeneration of cone visu-

al pigments (photoreceptors of daylight vision) [119].

However, this is indirect photoregeneration, as it

involves a specialized non-visual protein, retinal

G protein-coupled receptor (RGR), which acts as a

photoisomerase in Müller glial cells of the retina and

RPE cells [120]. The involvement of non-visual reti-

nal proteins in rhodopsin photoregeneration has long

been recognized. Aclassic example is retinochrome in

cephalopod mollusks, which functions as a photoisom-

erase [121, 122]. Notably, retinochrome is phylogenet-

ically related to the vertebrate RGR protein [119].

Thus, the evolutionary landscape of light-depen-

dent and dark regeneration of visual pigments can be

summarized as follows:

a) direct photoregeneration of invertebrate rhodop-

sin, in which a second photon is absorbed by the

stable photoproduct (acid metarhodopsin), restor-

ing the original state;

b) indirect photoregeneration of invertebrate rho-

dopsin mediated by retinochrome, a non-visual

retinal protein that catalyzes the light-driven

conversion of all-trans retinal back to the 11-cis

configuration;

c) dark regeneration of vertebrate visual pigments

via the enzymatic visual (retinoid) cycle (see re-

view [123]);

d) indirect photoregeneration of vertebrate cone

visual pigments catalyzed by RGR (see re-

view [123]).

The physiological significance of indirect pho-

toregeneration of cone visual pigments lies in the

requirement for rapid restoration of their native

state, enabling sustained phototransduction during

the daytime and under continuously bright lighting

conditions. In this process, 11-cis retinal is supplied

to cones from two principal sources: Müller glia of

the retina and RPE cells. However, in both rods and

cones, the primary pathway responsible for slow but

complete dark adaptation is the classical visual (ret-

inoid) cycle, in which the RPE plays a central role.

Notably, in  vivo and in  situ experiments con-

ducted in rabbits and albino rats have demonstrat-

ed direct photoregeneration of rhodopsin from early

photointermediates, including metarhodopsin I (see

“Photoreversibility at late stages of rhodopsin con-

version”) [62-65]. These findings show that follow-

ing a sufficiently brief light flash, irrespective of its

intensity, approximately 25-50% of rhodopsin in the

retinas of the studied animals remained in the dark

state. This indicates the existence of an additional,

rapid mechanism for restoring the physiologically

active dark state of the visual pigment, which can

operate under conditions of high light intensity in

living organisms. However, the extent to which this

photoregenerative mechanism contributes to rhodop-

sin recovery under natural lighting conditions in ver-

tebrates remains unclear.

CONCLUSION

This review examines reverse photoreactions

in visual rhodopsins (type II), as well as in selected

type I rhodopsins, as “tools” for investigating both

forward and reverse photochemical pathways and

associated conformational rearrangements of these

proteins.

Although reverse photoreactions initiated from

the earliest photointermediates of forward rhodop-

sin conversion occurring on the femtosecond to pico-

second timescales are not considered physiologically

relevant, their experimental and theoretical analysis

provides important insights into the mechanism of

retinal photoisomerization and chromophore–protein

interactions that enable the protein to “catalyze” this

exceptionally rapid and efficient process. These stud-

ies are therefore crucial for understanding the fun-

damental photobiological principles governing light

energy conversion in retinal-binding proteins that

mediate both photoinformational and photoenergetic

functions.

Reverse photoreactions originating from later

stages of the rhodopsin photocycle can serve as

markers of changes in chromophore–protein inter-

actions and of protein (opsin) conformational rear-

rangements that ultimately lead to the formation of

the signaling state.

The physiological significance of photorevers-

ible reactions in restoring the native state of visual

pigments in both vertebrates and invertebrates is

evident. While the mechanism of rhodopsin photo-

PHOTOREVERSIBLE REACTIONS OF RHODOPSINS 1369

BIOCHEMISTRY (MOSCOW) Vol. 91 No. 8 2026

regeneration in invertebrates is relatively well char-

acterized, several key questions remain unresolved

regarding the photoregeneration of vertebrate cone

photoreceptors (see review [123]). In particular, the

relationship between the classical dark visual (reti-

noid) cycle of rhodopsin regeneration occurring in

the RPE and the distinct light-dependent and dark

regeneration pathways operating within the neural

part of the retina remains poorly understood.

From an applied perspective, retinal-containing

proteins are regarded as promising biomaterials for

molecular electronics and optoelectronics, owing to

their chemical stability, functional photocycles, and

photoreversible reactions. In particular, they can

function as molecular photoswitches, based on the re-

versible light-induced isomerization between the cis

and all-trans forms of retinal. For instance, bacteri-

orhodopsin has been used as a holographic medium

for processing optical information in high-speed bio-

chips. The development of logic elements and mem-

ory devices modeled on rhodopsin-based systems is

currently actively discussed. In this context, the ul-

trafast photoreversibility of both visual and microbial

rhodopsins, occurring on the femtosecond to picosec-

ond timescale, is of particular interest.

In conclusion, studying photoreversible reactions

in rhodopsins remains important not only for funda-

mental understanding in photochemistry, biophysics,

and visual physiology, but also for applied research

in molecular electronics and optoelectronic tech-

nologies.

Abbreviations

BSI blue shifted intermediate of visual

rhodopsin

HOOP hydrogen-out-of-plane

MM/QM hybrid method of molecular

mechanics and quantum mechanics

RPE retinal pigment epithelium

Acknowledgments

The authors express their gratitude to A.  V.  Bochenkova

and P.  A.  Kusochek for their assistance in creating the

Figure  3 and discussing review sections on the theo-

retical calculations.

Contributions

M.A.O. developed the study concept; M.A.O., O.A.S.,

and T.B.F. wrote and edited the manuscript.

Funding

The work was supported by the Ministry of Science

and Higher Education of the Russian Federation (proj-

ect no. 122041400102-9) and the State Assignment to

the Lomonosov Moscow State University.

Ethics approval and consent to participate

This work does not contain any studies involving hu-

man or animal subjects.

Conflict of interest

The authors of this work declare that they have no

conflicts of interest.

REFERENCES

1. Kandori, H., Shichida, Y., and Yoshizawa, T. (2001) Photoisomerization in rhodopsin, Biochemistry (Moscow), 66,

1197-1209, https://doi.org/10.1023/A:1013123016803.

2. Ernst, O.P., Lodowski, D.T., Elstner, M., Hegemann, P., Brown, L.S., and Kandori, H. (2014) Microbial and animal

rhodopsins: structures, functions, and molecular mechanisms, Chem. Rev., 114, 126-163, https://doi.org/10.1021/

cr4003769.

3. Gozem,S., Luk, H.L., Schapiro,I., and Olivucci,M. (2017) Theory and simulation of the ultrafast double-bond isom-

erization of biological chromophores, Chem. Rev., 117, 13502-13565, https://doi.org/10.1021/acs.chemrev.7b00177.

4. Agathangelou, D., Roy, P. P., del Carmen Marín, M., Ferré, N., Olivucci, M., Buckup, T., Léonard, J., and Haacke, S.

(2021) Sub-picosecond C=C bond photo-isomerization: evidence for the role of excited state mixing, C. R. Phys.,

22, 111-138, https://doi.org/10.5802/CRPHYS.41.

5. Nagata,T., and Inoue,K. (2021) Rhodopsins at a glance, J. Cell Sci., 134, jcs258989, https://doi.org/10.1242/jcs.258989.

6. De Grip, W. J., and Ganapathy, S. (2022) Rhodopsins: an excitingly versatile protein species for research, devel-

opment and creative engineering, Front. Chem., 10, 879609, https://doi.org/10.3389/fchem.2022.879609.

7. Hofmann, K.P., and Lamb, T.D. (2023) Rhodopsin, light-sensor of vision, Prog. Retin. Eye Res., 93, 101116, https://

doi.org/10.1016/j.preteyeres.2022.101116.

8. Inoue, K. (2023) Photochemistry of the retinal chromophore in microbial rhodopsins, J. Phys. Chem. B, 127, 9215-

9222, https://doi.org/10.1021/acs.jpcb.3c05467.

9. Ostrovsky, M. A., Smitienko, O. A., Bochenkova, A. V., and Feldman, T. B. (2023) Similarities and differences in

photochemistry of type I and type II rhodopsins, Biochemistry (Moscow), 88, 1528-1543, https://doi.org/10.1134/

S0006297923100097.

10. Rozenberg, A., Inoue, K., Kandori, H., and Béjà, O. (2021) Microbial rhodopsins: the last two decades, Annu. Rev.

Microbiol., 75, 427-447, https://doi.org/10.1146/annurev-micro-031721-020452.

OSTROVSKY et al.1370

BIOCHEMISTRY (MOSCOW) Vol. 91 No. 8 2026

11. Gordeliy, V., Kovalev, K., Bamberg, E., Rodriguez-Valera, F., Zinovev, E., Zabelskii, D., Alekseev, A., Rosselli, R.,

Gushchin, I., and Okhrimenko, I. (2022) Microbial rhodopsins, Methods Mol. Biol., 2501, 1-52, https://

doi.org/10.1007/978-1-0716-2329-9_1.

12. Engelhard, M. (2022) Molecular biology of microbial rhodopsins, Methods Mol. Biol., 2501, 53-69, https://

doi.org/10.1007/978-1-0716-2329-9_2.

13. Wald, G., Durell, J., and St. George, R. C. C. (1950) The light reaction in the bleaching of rhodopsin, Science, 111,

179-181, https://doi.org/10.1126/science.111.2877.179.

14. Kiefer, H. V., Gruber, E., Langeland, J., Kusochek, P. A., Bochenkova, A. V., and Andersen, L. H. (2019) Intrin-

sic photoisomerization dynamics of protonated Schiff-base retinal, Nat. Commun., 10, 1210, https://doi.org/

10.1038/s41467-019-09225-7.

15. Kandori, H., Sasabe, H., Katsuta, Y., and Ito, M. (1995) Femtosecond fluorescence study of the rhodopsin chro-

mophore in solution, J. Am. Chem. Soc., 117, 2669-2670, https://doi.org/10.1021/ja00114a040.

16. Zgrablić, G., Novello, A. M., and Parmigiani, F. (2012) Population branching in the conical intersection of the

retinal chromophore revealed by multipulse ultrafast optical spectroscopy, J. Am. Chem. Soc., 134, 955-961, https://

doi.org/10.1021/ja205763x.

17. Koyama, Y., Kubo, K., Komori, M., Yasuda, H., and Mukai, Y. (1991) Effect of protonation on the isomerization

properties of n‐butylamine Schiff base of isomeric retinal as revealed by direct HPLC analyses: selection of

isomerization pathways by retinal proteins, Photochem. Photobiol., 54, 433-443, https://doi.org/10.1111/j.1751-1097.

1991.tb02038.x.

18. Yoshizawa,T., and Shichida,Y. (1982) Low-temperature spectrophotometry of intermediates of rhodopsin, Methods

Enzymol., 81, 333-354, https://doi.org/10.1016/S0076-6879(82)81051-3.

19. Kliger, D. S., and Lewis, J. W. (1995) Spectral and kinetic characterization of visual pigment photointermediates,

Isr. J. Chem., 35, 289-307, https://doi.org/10.1002/ijch.199500032.

20. Ernst, O. P., and Bartl, F. J. (2002) Active states of rhodopsin, ChemBioChem, 3, 968-974, https://doi.org/10.1002/

1439-7633(20021004)3:10<968::AID-CBIC968>3.0.CO;2-Q.

21. Tsuda, M. (1979) Transient spectra of intermediates in the photolytic sequence of octopus rhodopsin, Biochim.

Biophys. Acta Bioenerg., 545, 537-546, https://doi.org/10.1016/0005-2728(79)90162-2.

22. Dixon, S. F., and Cooper, A. (1987) Quantum efficiencies of the reversible photoreaction of octopus rhodopsin,

Photochem. Photobiol., 46, 115-119, https://doi.org/10.1111/j.1751-1097.1987.tb04744.x.

23. Nakagawa, M., Kikkawa, S., Tominaga, K., Tsugi, N., and Tsuda, M. (1998) A novel photointermediate of octopus

rhodopsin activates its G-protein, FEBS Lett., 436, 259-262, https://doi.org/10.1016/S0014-5793(98)01138-7.

24. Nishioku, Y., Nakagawa, M., Tsuda, M., and Terazima, M. (2001) A spectrally silent transformation in the photol-

ysis of octopus rhodopsin: a protein conformational change without any accompanying change of the chromo-

phore’s absorption, Biophys. J., 80, 2922-2927, https://doi.org/10.1016/S0006-3495(01)76257-1.

25. Inoue, K., Tsuda, M., and Terazima, M. (2007) Photoreverse reaction dynamics of octopus rhodopsin, Biophys. J.,

92, 3643-3651, https://doi.org/10.1529/biophysj.106.101741.

26. Yabushita, A., Kobayashi, T., and Tsuda, M. (2012) Time-resolved spectroscopy of ultrafast photoisomer-

ization of octopus rhodopsin under photoexcitation, J.Phys. Chem. B, 116, 1920-1926, https://doi.org/10.1021/

jp209356s.

27. Dartnall, H. J. A. (1968) The photosensitivities of visual pigments in the presence of hydroxylamine, Vision Res.,

8, 339-358, https://doi.org/10.1016/0042-6989(68)90104-1.

28. Kim, J. E., Tauber, M. J., and Mathies, R. A. (2001) Wavelength dependent cis-trans isomerization in vision, Bio-

chemistry, 40, 13774-13778, https://doi.org/10.1021/bi0116137.

29. Polli, D., Altoè, P., Weingart, O., Spillane, K. M., Manzoni, C., Brida, D., Tomasello, G., Orlandi, G., Kukura, P.,

Mathies, R. A., Garavelli, M., and Cerullo, G. (2010) Conical intersection dynamics of the primary photoisomeri-

zation event in vision, Nature, 467, 440-443, https://doi.org/10.1038/nature09346.

30. Smitienko, O. A., Mozgovaya, M. N., Shelaev, I. V., Gostev, F. E., Feldman, T. B., Nadtochenko, V. A., Sarkisov,

O.M., and Ostrovsky, M. A. (2010) Femtosecond formation dynamics of primary photoproducts of visual pigment

rhodopsin, Biochemistry (Moscow), 75, 25-35, https://doi.org/10.1134/S0006297910010049.

31. Nadtochenko, V.A., Smitienko, O.A., Feldman, T.B., Mozgovaya, M.N., Shelaev, I. V., Gostev, F.E., Sarkisov, O. M.,

and Ostrovsky, M. A. (2012) Conical intersection participation in femtosecond dynamics of visual pigment

rhodopsin chromophore cis-trans photoisomerization, Dokl. Biochem. Biophys., 446, 242-246, https://doi.org/

10.1134/S1607672912050080.

32. Johnson, P. J. M., Halpin, A., Morizumi, T., Prokhorenko, V. I., Ernst, O. P., and Miller, R. J. D. (2015) Local vi-

brational coherences drive the primary photochemistry of vision, Nat. Chem., 7, 980-986, https://doi.org/10.1038/

nchem.2398.

PHOTOREVERSIBLE REACTIONS OF RHODOPSINS 1371

BIOCHEMISTRY (MOSCOW) Vol. 91 No. 8 2026

33. Feldman, T. B., Smitienko, O. A., Shelaev, I. V., Gostev, F. E., Nekrasova, O. V., Dolgikh, D. A., Nadtochenko,

V. A., Kirpichnikov, M. P., and Ostrovsky, M. A. (2016) Femtosecond spectroscopic study of photochromic reac-

tions of bacteriorhodopsin and visual rhodopsin, J. Photochem. Photobiol. B, 164, 296-305, https://doi.org/10.1016/

j.jphotobiol.2016.09.041.

34. Klessinger, M. (1995) Conical intersections and the mechanism of singlet photoreactions, Angew. Chem. Int. Ed.,

34, 549-551, https://doi.org/10.1002/anie.199505491.

35. Schapiro, I., Melaccio, F., Laricheva, E. N., and Olivucci, M. (2011) Using the computer to understand

the chemistry of conical intersections, Photochem. Photobiol. Sci., 10, 867-886, https://doi.org/10.1039/

c0pp00290a.

36. Schoenlein, R. W., Peteanu, L. A., Mathies, R. A., and Shank, C. V. (1991) The first step in vision: femtosecond

isomerization of rhodopsin, Science, 254, 412-415, https://doi.org/10.1126/science.1925597.

37. Kim, J. E., and Mathies, R. A. (2002) Anti-Stokes Raman study of vibrational cooling dynamics in the primary

photochemistry of rhodopsin, J. Phys. Chem. A, 106, 8508-8515, https://doi.org/10.1021/jp021069r.

38. Gruhl, T., Weinert, T., Rodrigues, M. J., Milne, C. J., Ortolani, G., Nass, K., Nango, E., Sen, S., Johnson, P. J. M.,

Cirelli,C., Furrer,A., Mous, S., Skopintsev,P., James,D., Dworkowski,F., Båth,P., Kekilli,D., Ozerov, D., Tanaka,R.,

et al. (2023) Ultrafast structural changes direct the first molecular events of vision, Nature, 615, 939-944,

https://doi.org/10.1038/s41586-023-05863-6.

39. Yan,M., Manor, D., Weng, G., Chao,H., Rothberg,L., Jedju, T.M., Alfano, R.R., and Callender, R.H. (1991) Ultrafast

spectroscopy of the visual pigment rhodopsin, Proc. Natl. Acad. Sci. USA, 88, 9809-9812, https://doi.org/10.1073/

pnas.88.21.9809.

40. Hug, S. J., Lewis, J. W., Einterz, C. M., Thorgeirsson, T. E., and Kliger, D. S. (1990) Nanosecond photolysis of

rhodopsin: Evidence for a new, blue-shifted intermediate, Biochemistry, 29, 1475-1485, https://doi.org/10.1021/

bi00458a019.

41. Fel’dman, T. B., Fedorovich, I. B., and Ostrovskii, M. A. (2004) Characteristics of the photoconversion of

rhodopsin in the early stages of photolysis, Neurosci Behav Physiol., 34, 735-742, https://doi.org/10.1023/

B:NEAB.0000036015.85880.9b.

42. Vogel,R., Lüdeke,S., Radu,I., Siebert,F., and Sheves,M. (2004) Photoreactions of metarhodopsin III, Biochemistry,

43, 10255-10264, https://doi.org/10.1021/bi049182q.

43. Doukas, A. G., Junnarkar, M. R., Alfano, R. R., Callender, R. H., Kakitani, T., and Honig, B. (1984) Fluorescence

quantum yield of visual pigments: evidence for subpicosecond isomerization rates, Proc. Natl. Acad. Sci. USA,

81, 4790-4794, https://doi.org/10.1073/pnas.81.15.4790.

44. Ohtani,H., Kobayashi,T., Tsuda,M., and Ebrey, T.G. (1988) Primary processes in photolysis of octopus rhodopsin,

Biophys. J., 53, 17-24, https://doi.org/10.1016/S0006-3495(88)83061-3.

45. Taiji, M., Bryl, K., Nakagawa, M., Tsuda, M., and Kobayashi, T. (1992) Femtosecond studies of primary

photoprocesses in octopus rhodopsin, Photochem. Photobiol., 56, 1003-1011, https://doi.org/10.1111/j.1751-1097.

1992.tb09723.x.

46. Kobayashi, T., Kim, M., Taiji, M., Iwasa, T., Nakagawa, M., and Tsuda, M. (1998) Femtosecond spectroscopy of

halorhodopsin and rhodopsin in a broad spectral range of 400-1000 nm, J. Phys. Chem. B, 102, 272-280, https://

doi.org/10.1021/jp970705w.

47. Kojima, K., and Sudo, Y. (2023) Convergent evolution of animal and microbial rhodopsins, RSC Adv., 13, 5367-

5381, https://doi.org/10.1039/d2ra07073a.

48. Lanyi, J. K. (2004) Bacteriorhodopsin, Annu. Rev. Physiol., 66, 665-688, https://doi.org/10.1146/annurev.physi-

ol.66.032102.150049.

49. Mathies, R. A., Brito Cruz, C. H., Pollard, W. T., and Shank, C. V. (1988) Direct observation of the femtosecond

excited-state cis-trans isomerization in bacteriorhodopsin, Science, 240, 777-779, https://doi.org/10.1126/science.

3363359.

50. Smitienko, O. A., Nekrasova, O. V., Kudriavtsev, A. V., Yakovleva, M. A., Shelaev, I. V., Gostev, F. E., Dolgikh, D. A.,

Kolchugina, I. B., Nadtochenko, V. A., Kirpichnikov, M. P., Feldman, T. B., and Ostrovsky, M. A. (2017) Femto-

second and picosecond dynamics of recombinant bacteriorhodopsin primary reactions compared to the na-

tive protein in trimeric and monomeric forms, Biochemistry (Moscow), 82, 490-500, https://doi.org/10.1134/

S0006297917040113.

51. Smitienko, O.A., Feldman, T. B., Petrovskaya, L.E., Nekrasova, O.V., Yakovleva, M. A., Shelaev, I. V., Gostev, F. E.,

Cherepanov, D. A., Kolchugina, I. B., Dolgikh, D. A., Nadtochenko, V. A., Kirpichnikov, M. P., and Ostrovsky, M. A.

(2021) Comparative femtosecond spectroscopy of primary photoreactions of Exiguobacterium sibiricum rho-

dopsin and Halobacterium salinarum bacteriorhodopsin, J. Phys. Chem. B, 125, 995-1008, https://doi.org/

10.1021/acs.jpcb.0c07763.

OSTROVSKY et al.1372

BIOCHEMISTRY (MOSCOW) Vol. 91 No. 8 2026

52. Chang, C., Kuramochi, H., Singh, M., Abe-Yoshizumi, R., Tsukuda, T., Kandori, H., and Tahara, T. (2022) A unified

view on varied ultrafast dynamics of the primary process in microbial rhodopsins, Angew. Chem. Int. Ed., 61,

e202111930, https://doi.org/10.1002/anie.202111930.

53. Balashov, S. P. (2000) Protonation reactions and their coupling in bacteriorhodopsin, Biochim. Biophys. Acta

Bioenerg., 1460, 75-94, https://doi.org/10.1016/S0005-2728(00)00131-6.

54. Sineshchekov, O. A., Trivedi, V. D., Sasaki, J., and Spudich, J. L. (2005) Photochromicity of Anabaena sensory

rhodopsin, an atypical microbial receptor with a cis-retinal light-adapted form, J. Biol. Chem., 280, 14663-14668,

https://doi.org/10.1074/jbc.M501416200.

55. Kawanabe, A., and Kandori, H. (2009) Photoreactions and structural changes of Anabaena sensory rhodopsin,

Sensors, 9, 9741-9804, https://doi.org/10.3390/s91209741.

56. Inoue,K., Tsukamoto,T., and Sudo,Y. (2014) Molecular and evolutionary aspects of microbial sensory rhodopsins,

Biochim. Biophys. Acta Bioenerg., 1837, 562-577, https://doi.org/10.1016/j.bbabio.2013.05.005.

57. Sudo, Y., Mizuno, M., Wei, Z., Takeuchi, S., Tahara, T., and Mizutani, Y. (2014) The early steps in the photocy-

cle of a photosensor protein sensory rhodopsin I from Salinibacter ruber, J. Phys. Chem. B, 118, 1510-1518,

https://doi.org/10.1021/jp4112662.

58. Bruun, S., Stoeppler, D., Keidel, A., Kuhlmann, U., Luck, M., Diehl, A., Geiger, M.-A., Woodmansee, D., Trauner, D.,

Hegemann, P., Oschkinat, H., Hildebrandt, P., and Stehfest, K. (2015) Light-dark adaptation of channelrhodopsin

involves photoconversion between the all-trans and 13-cis retinal isomers, Biochemistry, 54, 5389-5400, https://

doi.org/10.1021/acs.biochem.5b00597.

59. Ostroy, S. E. (1977) Rhodopsin and the visual process, Biochim. Biophys. Acta Rev. Bioenerg., 463, 91-125, https://

doi.org/10.1016/0304-4173(77)90004-0.

60. Balashov, S. P. (1995) Photoreactions of the photointermediates of bacteriorhodopsin, Isr. J. Chem., 35, 415-428,

https://doi.org/10.1002/ijch.199500040.

61. Hampp, N. (2000) Bacteriorhodopsin as a photochromic retinal protein for optical memories, Chem. Rev., 100,

1755-1776, https://doi.org/10.1021/cr980072x.

62. Hagins, W. A. (1955) The quantum efficiency of bleaching in situ, J. Physiol. (Lond.), 129, 22.

63. Dowling, J. E., and Hubbard,R. (1963) Effect of instantaneous flashes on adaptation of the eye: effects of brilliant

flashes on light and dark adaptation, Nature, 199, 972-975, https://doi.org/10.1038/199972a0.

64. Cone, R. A. (1967) Early receptor potential: photoreversible charge displacement in rhodopsin, Science, 155, 1128-

1131, https://doi.org/10.1126/science.155.3766.1128.

65. Grimm, C., Reme, C. E., Rol, P. O., and Williams, T. P. (2000) Blue light’s effects on rhodopsin: Photoreversal of

bleaching in living rat eyes, Invest. Ophthalmol. Vis. Sci., 41, 3984-3990.

66. Gaulier, G., Dietschi, Q., Bhattacharyya, S., Schmidt, C., Montagnese, M., Chauvet, A., Hermelin, S., Chiodini, F.,

Bonacina, L., Herrera, P. L., Rothlisberger, U., Rodriguez, I., and Wolf, J.-P. (2021) Ultrafast pulse shaping

modulates perceived visual brightness in living animals, Sci. Adv., 7, eabe1911, https://doi.org/10.1126/sciadv.

abe1911.

67. Yoshizawa, T., and Wald, G. (1963) Pre-lumirhodopsin and the bleaching of visual pigments, Nature, 197, 1279-

1286, https://doi.org/10.1038/1971279a0.

68. Hurley, J. B., Ebrey, T. G., Honig, B., and Ottolenghi, M. (1977) Temperature and wavelength effects on the

photochemistry of rhodopsin, isorhodopsin, bacteriorhodopsin and their photoproducts, Nature, 270, 540-542,

https://doi.org/10.1038/270540a0.

69. Maeda, A., Ogurusu, T., Shichida, Y., Tokunaga, F., and Yoshizawa, T. (1978) Formation of a 7-cis retinal pig-

ment by irradiating cattle rhodopsin at low temperatures, FEBS Lett., 92, 77-80, https://doi.org/10.1016/

0014-5793(78)80725-X.

70. Becher, B. (1980) The photoconversion of lumirhodopsin at 77 degrees K. Estimation of the quantum efficiency,

Biophys. J., 30, 1-8, https://doi.org/10.1016/S0006-3495(80)85072-7.

71. Suzuki, T., and Callender, R. H. (1981) Primary photochemistry and photoisomerization of retinal at 77 degrees

K in cattle and squid rhodopsins, Biophys. J., 34, 261-265, https://doi.org/10.1016/S0006-3495(81)84848-5.

72. Paulsen, R., and Bentrop, J. (1983) Activation of rhodopsin phosphorylation is triggered by the lumirhodopsin-

metarhodopsin I transition, Nature, 302, 417-419, https://doi.org/10.1038/302417a0.

73. Protasova, T. B., Tarasov, V. F., and Fedorovich, I. B. (1989) The retinal isomeric composition in the rhodopsin

illuminated at 77 K [in Russian], Sens. Syst., 3, 19-24.

74. Yan, M., Rothberg, L., and Callender, R. (2001) Femtosecond dynamics of rhodopsin photochemistry probed by a

double pump spectroscopic approach, J. Phys. Chem. B, 105, 856-859, https://doi.org/10.1021/jp002036j.

75. Furutani, Y., Kandori, H., and Shichida, Y. (2003) Structural changes in lumirhodopsin and metarhodopsin I

studied by their photoreactions at 77 K, Biochemistry, 42, 8494-8500, https://doi.org/10.1021/bi034438y.

PHOTOREVERSIBLE REACTIONS OF RHODOPSINS 1373

BIOCHEMISTRY (MOSCOW) Vol. 91 No. 8 2026

76. Mozgovaya, M. N., Smitienko, O. A., Shelaev, I. V., Gostev, F. E., Feldman, T. B., Nadtochenko, V. A.,

Sarkisov, O. M., and Ostrovsky, M. A. (2010) Photochromism of the visual pigment rhodopsin on the femtosec-

ond time scale: coherent control of retinal chromophore photoisomerization [in Russian], Dokl. Akad. Nauk,

435, 262-266.

77. Smitienko, O., Nadtochenko, V., Feldman, T., Balatskaya, M., Shelaev, I., Gostev, F., Sarkisov, O., and Ostrovsky, M.

(2014) Femtosecond laser spectroscopy of the rhodopsin photochromic reaction: a concept for ultrafast opti-

cal molecular switch creation (ultrafast reversible photoreaction of rhodopsin), Molecules, 19, 18351-18366,

https://doi.org/10.3390/molecules191118351.

78. Birge, R. R., Cooper, T. M., Lawrence, A. F., Masthay, M. B., Vasilakis, C., Zhang, C. F., and Zidovetzki, R. (1989)

A spectroscopic, photocalorimetric, and theoretical investigation of the quantum efficiency of the primary event

in bacteriorhodopsin, J.Am. Chem. Soc., 111, 4063-4074, https://doi.org/10.1021/ja00193a044.

79. Govindjee, R., Balashov, S. P., and Ebrey, T. G. (1990) Quantum efficiency of the photochemical cycle of bacteri-

orhodopsin, Biophys. J., 58, 597-608, https://doi.org/10.1016/S0006-3495(90)82403-6.

80. Bazhenov, V., Schmidt, P., and Atkinson, G. H. (1992) Nanosecond photolytic interruption of bacteriorhodopsin

photocycle: K-590 → BR-570 reaction, Biophys. J., 61, 1630-1637, https://doi.org/10.1016/S0006-3495(92)81966-5.

81. Delaney, J. K., Schmidt, P. K., Brack, T. L., and Atkinson, G. H. (2000) Photochemistry of K-590 in the room-tem-

perature bacteriorhodppsin photocycle, J. Phys. Chem. B, 104, 10827-10834, https://doi.org/10.1021/jp000374e.

82. Kusochek, P. A., Smitienko, O.A., and Bochenkova, A.V. (2024) Mode-specific photoresponse of retinal protonated

Schiff base isomers in the reversible photochromic reactions of microbial and animal rhodopsins, J. Phys. Chem.

B, 128, 12471-12482, https://doi.org/10.1021/ACS.JPCB.4C06832.

83. Malakar, P., Gholami, S., Aarabi, M., Rivalta, I., Sheves, M., Garavelli, M., and Ruhman, S. (2024) Retinal pho-

toisomerization versus counterion protonation in light and dark-adapted bacteriorhodopsin and its primary

photoproduct, Nat. Commun., 15, 2136, https://doi.org/10.1038/s41467-024-46061-w.

84. Smitienko, O., Feldman, T., Shelaev, I., Gostev, F., Aybush, A., Cherepanov, D., Nadtochenko, V., and Ostrovsky, M.

(2024) Reversible photochromic reactions of bacteriorhodopsin from Halobacterium salinarum at femto- and

picosecond times, Molecules, 29, 4847, https://doi.org/10.3390/molecules29204847.

85. Birge, R.R., and Hubbard, L.M. (1981) Molecular dynamics of trans-cis isomerization in bathorhodopsin, Biophys.

J., 34, 517-534, https://doi.org/10.1016/S0006-3495(81)84865-5.

86. Schapiro, I., Ryazantsev, M. N., Frutos, L. M., Ferré, N., Lindh, R., and Olivucci, M. (2011) The ultrafast pho-

toisomerizations of rhodopsin and bathorhodopsin are modulated by bond length alternation and HOOP driven

electronic effects, J. Am. Chem. Soc., 133, 3354-3364, https://doi.org/10.1021/ja1056196.

87. Kusochek, P. A., Logvinov, V. V., and Bochenkova, A. V. (2021) Role of the protein environment in photoisomeri-

zation of type I and type II rhodopsins: a theoretical perspective, Moscow Univ. Chem. Bull., 76, 407-416, https://

doi.org/10.3103/S0027131421060110.

88. Ritter, E., Zimmennann, K., Heck, M., Hofmann, K. P., and Bartl, F. J. (2004) Transition of rhodopsin into the

active metarhodopsin II state opens a new light-induced pathway linked to Schiff base isomerization, J. Biol.

Chem., 279, 48102-48111, https://doi.org/10.1074/jbc.M406857200.

89. Eyring, G., Curry, B., Mathies, R., Fransen, R., Palings, I., and Lugtenburg, J. (1980) Interpretation of the reso-

nance Raman spectrum of bathorhodopsin based on visual pigment analogs, Biochemistry, 19, 2410-2418, https://

doi.org/10.1021/bi00552a020.

90. Fukada, Y., Shichida, Y., Yoshizawa, T., Ito, M., Kodama, A., and Tsukida, K. (1984) Studies on structure and func-

tion of rhodopsin by use of cyclopentatrienylidene 11-cis-locked-rhodopsin, Biochemistry, 23, 5826-5832, https://

doi.org/10.1021/bi00319a023.

91. Kandori, H., Matuoka, S., Shichida, Y., Yoshizawa, T., Ito, M., Tsukida,K., Balogh-Nair, V., and Nakanishi, K. (1989)

Mechanism of isomerization of rhodopsin studied by use of 11-cis-locked rhodopsin analogues excited with a

picosecond laser pulse, Biochemistry, 28, 6460-6467, https://doi.org/10.1021/bi00441a045.

92. Lewis, J. W., Hug, S. J., Wallace-Williams, S. E., and Kliger, D. S. (1990) Direct evidence for an equilibrium be-

tween early photolysis intermediates of rhodopsin, J. Am. Chem. Soc., 112, 6711-6712, https://doi.org/10.1021/

ja00174a040.

93. Balashov, S. P., Imasheva, E. S., Govindjee, R., and Ebrey, T. G. (1991) Quantum yield ratio of the forward and

back light reactions of bacteriorhodopsin at low temperature and photosteady‐state concentration of the batho-

product K, Photochem. Photobiol., 54, 955-961, https://doi.org/10.1111/j.1751-1097.1991.tb02116.x.

94. Tahara, S., Takeuchi, S., Abe-Yoshizumi, R., Inoue, K., Ohtani, H., Kandori, H., and Tahara, T. (2018) Origin of

the reactive and nonreactive excited states in the primary reaction of rhodopsins: pH dependence of femto-

second absorption of light-driven sodium ion pump rhodopsin KR2, J. Phys. Chem. B, 122, 4784-4792, https://

doi.org/10.1021/acs.jpcb.8b01934.

OSTROVSKY et al.1374

BIOCHEMISTRY (MOSCOW) Vol. 91 No. 8 2026

95. Kusochek, P. A., Scherbinin, A. V., and Bochenkova, A. V. (2021) Insights into the early-time excited-state dy-

namics of structurally inhomogeneous rhodopsin KR2, J. Phys. Chem. Lett., 12, 8664-8671, https://doi.org/10.1021/

acs.jpclett.1c02312.

96. Becker, R. S., and Freedman, K. (1985) A comprehensive investigation of the mechanism and photophysics of

isomerization of a protonated and unprotonated Schiff base of 11-cis-retinal, J. Am. Chem. Soc., 107, 1477-1485,

https://doi.org/10.1021/ja00292a005.

97. Hubbard, R., and Kropf, A. (1958) The action of light on rhodopsin, Proc. Natl. Acad. Sci. USA, 44, 130-139,

https://doi.org/10.1073/pnas.44.2.130.

98. Matthews, R. G., Hubbard, R., Brown, P. K., and Wald, G. (1963) Tautomeric forms of metarhodopsin, J. Gen.

Physiol., 47, 215-240, https://doi.org/10.1085/jgp.47.2.215.

99. Williams, T. P. (1968) Photolysis of metarhodopsin II: rates of production of P470 and rhodopsin, Vision Res., 8,

1457-1465, https://doi.org/10.1016/0042-6989(68)90120-X.

100. Baker, B.N., and Williams, T.P. (1971) Photolysis of metarhodopsin I: rate and extent of conversion to rhodopsin,

Vision Res., 11, 449-458, https://doi.org/10.1016/0042-6989(71)90086-1.

101. Krongauz, V. A., Shifrina, R. R., Federovich, I. B., and Ostrovsky, M. A. (1975) Photochromism of visual pigments.

1. Formation of isochromic products in reversible transformations of frog rhodopsin [in Russian], Biophysics, 20,

219-224.

102. Krongauz, V. A., Shifrina, R. R., Federovich, I. B., and Ostrovsky, M. A. (1975) Photochromism of visual pigments.

2. Kinetics of photoconversions of frog rhodopsin rhodopsin [in Russian], Biophysics, 20, 419-424.

103. Reuter, T. (1976) Photoregeneration of rhodopsin and isorhodopsin from metarhodopsin III in the frog retina,

Vision Res., 16, 909-917, https://doi.org/10.1016/0042-6989(76)90220-0.

104. Bartl, F. J., Ritter, E., and Hofmann, K. P. (2001) Signaling states of rhodopsin. Absorption of light in active

metarhodopsin II generates an all-trans-retinal bound inactive state, J. Biol. Chem., 276, 30161-30166, https://

doi.org/10.1074/jbc.M101506200.

105. Kolesnikov, A. V., Korenyak, D. A., Shukolyukov, S. A., and Govardovskii, V. I. (2011) Photoreactions of metarho-

dopsin II [in Russian], Sens. Syst., 25, 55-64.

106. Arnis,S., and Hofmann, K.P. (1995) Photoregeneration of bovine rhodopsin from its signaling state, Biochemistry,

34, 9333-9340, https://doi.org/10.1021/bi00029a008.

107. Williams, T. P. (1975) Dynamics of opsin, a visual protein, Acc. Chem. Res., 8, 107-112, https://doi.org/10.1021/

ar50087a005.

108. Ritter, E., Elgeti, M., Hofmann, K. P., and Bartl, F. J. (2007) Deactivation and proton transfer in light-induced

metarhodopsin II/metarhodopsin III conversion: A time-resolved Fourier transform infrared spectroscopic study,

J. Biol. Chem., 282, 10720-10730, https://doi.org/10.1074/jbc.M610658200.

109. Pogozheva, I. D., Kuznetsov, V. A., Livshits, V. A., Federovich, I. B., and Ostrovsky, M. A. (1985) Photoinduced

changes in the hydrophilic region of the rhodopsin molecule. A study by saturation transfer EPR spectroscopy

[in Russian], Biol. Membr., 2, 880-896.

110. Pogozheva, I. D., Kuznetsov, V. A., Livshits, V. A., and Ostrovsky, M. A. (1985) Conformational mobility and inter-

action of rhodopsin domains [in Russian], Biol. Membr., 2, 897-905.

111. Shelyakin, P. V., Kovarskii, A. L., Kasparov, V. V., Feldman, T. B., and Ostrovsky, M. A. (2012) A study of the

photoinduced conformational mobility of spin-labeled regenerated rhodopsin by ESR spectroscopy, Russ. J. Phys.

Chem. B, 6, 694-698, https://doi.org/10.1134/S1990793112060127.

112. Ostrovsky, M. A., and Weetall, H. H. (1998) Octopus rhodopsin photoreversibility of a crude extract from

whole retina over several weeks’ duration, Biosens. Bioelectron., 13, 61-65, https://doi.org/10.1016/S0956-

5663(97)00078-X.

113. Paternolli, C., Neebe, M., Stura,E., Barbieri, F., Ghisellini, P., Hampp, N., and Nicolini, C. (2009) Photoreversibility

and photostability in films of octopus rhodopsin isolated from octopus photoreceptor membranes, J. Biomed.

Mater. Res. A, 88, 947-951, https://doi.org/10.1002/jbm.a.31925.

114. Naito,T., Nashima-Hayama,K., Ohtsu,K., and Kito, Y. (1981) Photoreactions of cephalopod rhodopsin, Vision Res.,

21, 935-941, https://doi.org/10.1016/0042-6989(81)90195-4.

115. Tsuda, M. (1978) Kinetic study of photoregeneration process of digitonin-solubilized squid rhodopsin, Biochim.

Biophys. Acta Bioenerg., 502, 495-506, https://doi.org/10.1016/0005-2728(78)90082-8.

116. Ehrenberg, D., Varma, N., Deupi, X., Koyanagi, M., Terakita, A., Schertler, G. F. X., Heberle, J., and Lesca, E.

(2019) The two-photon reversible reaction of the bistable jumping spider rhodopsin-1, Biophys. J., 116, 1248-1258,

https://doi.org/10.1016/j.bpj.2019.02.025.

117. Atzmon, Z., Hochstein, S., and Hillman, P. (1979) Transduction in photoreceptors: Determination of the pigment

transition or state coupled to excitation, Biophys. Struct. Mech., 5, 249-253, https://doi.org/10.1007/BF00535454.

PHOTOREVERSIBLE REACTIONS OF RHODOPSINS 1375

BIOCHEMISTRY (MOSCOW) Vol. 91 No. 8 2026

118. Hillman, P., Hochstein, S., and Minke, B. (1983) Transduction in invertebrate photoreceptors: role of pigment

bistability, Physiol. Rev., 63, 668-772, https://doi.org/10.1152/physrev.1983.63.2.668.

119. Chen, P., Hao, W., Rife, L., Wang, X. P., Shen, D., Chen, J., Ogden, T., van Boemel, G. B., Wu, L., Yang, M., and

Fong, H. K. W. (2001) A photic visual cycle of rhodopsin regeneration is dependent on RGR, Nat. Genet., 28,

256-260, https://doi.org/10.1038/90089.

120. Palczewski, K., and Kiser, P. D. (2020) Shedding new light on the generation of the visual chromophore, Proc.

Natl. Acad. Sci. USA, 117, 19629-19638, https://doi.org/10.1073/PNAS.2008211117.

121. Hara,T., and Hara,R. (1965) New photosensitive pigment found in the retina of the squid Ommastrephes, Nature,

206, 1331-1334, https://doi.org/10.1038/2061331a0.

122. Hara, T., and Hara, R. (1967) Vision in octopus and squid: rhodopsin and retinochrome in the squid retina,

Nature, 214, 573-575, https://doi.org/10.1038/214573a0.

123. Sato, S., and Kefalov, V. J. (2024) The retina-based visual cycle, Annu. Rev. Vis. Sci., 10, 293-321, https://

doi.org/10.1146/annurev-vision-100820-083937.

Publisher’s Note. Pleiades Publishing remains neutral with regard to jurisdictional claims in published

maps and institutional affiliations. AI tools may have been used in the translation or editing of this article.