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Spectroscopic and chemical properties of isomeric retinals andvisual pigment analogs

Zhu, Yun, Ph.D.

University of Hawaii, 1993

V·M·I300 N. ZeebRd.Ann Arbor.MI48106

SPECTROSCOPIC AND CHEMICAL PROPERTIES OF

ISOMERIC RETINALS AND VISUAL PIGMENT ANALOGS

A DISSERTATION SUBMITTED TO THE GRADUATE DIVISION OF THEUNIVERSITY OF HAWAII IN PARTIAL FULFILLMENT OF THE

REQUIREMENTS FOR THE DEGREE OF

DOCTOR OF PHILOSOPHY

IN

CHEMISTRY

AUGUST 1993

By

Yun Zhu

Dissertation Committee:

Robert S. H. Liu, ChairpersonRoger E. Cramer

Bradley S. DavidsonRandy W. Larsen

David Jameson

ACKNOWLEDGEl\1ENTS

I would like to express my deepest appreciation to my research supervisor,

Professor Robert S. H. Liu, for his guidance and support throughout my Ph.D. program.

His encouragement beyond science will always be with me.

I would like to express my sincerest thanks to Drs. Al Asato and Leticia

Colmenares for providing the substituted retinal analogs and their kind advice, and Drs.

S. Ganapathy and Achla Trehan for providing the geometric retinal isomers, and Dr.

Erik Krogh for his help in the early stage of construction of low temperature set up.

I would also like to thank Professor John Head for his instruction in the

calculation of retinal isomers, and Professor Randy Larsen for his help in the vibrational

calculation. Their time and thought shared with me are deeply appreciated. Also

. Professor Yoshinori Shichida, at Kyoto University for the valuable suggestions on the

low temperature spectroscopy of rhodopsin.

Special thanks to Xiaoyuan Li, Letty Colmenares, Coran Watanabe, Karen

Nishimura, Rongliang Chen and J. R. Thiel for their friendship and companionship.

Finally, I would like to thank my husband George and my parents for their

selfless loving support and encouragement for my accomplishments.

III

ABSTRACT

Complementing the recent interest in utilizing vibrational spectroscopies to probe

for structural information of protein bound retinyl chromophores, we have recorded FT­

IR spectra of all sixteen isomers of retinal. Characteristic trends of the poly-cis isomers

including C=C double bond stretching modes, C-C single bond stretching modes and

hydrogen out of plane bends (HOOP) have been discussed and compared with those of

the mono-cis and all-trans isomers. The normal modes of C-C stretching character of

two sterically hindered 7-cis and 7,9-dicis isomers and their Schiff bases (SB) and

protonated Schiff bases (PSB) have been assigned by using their isotopically labelled

analogs. These assignments are further substantiated through normal mode calculations.

These vibrational, data will provide a probe for studying the specific changes in the

chromophore-opsin interactions during the photobleaching processes of synthetic isomeric

rhodopsins.

Temperature-dependent isomerization of retinal isomers have been examined,

including the photoisomerization of mono-cis (7-cis, 9-cis, l l-cis) and 7,9-dicis-retinal

isomers and the thermal isomerization of. four unstable retinal isomers (11, 13-dicis,

7, 11,13-tricis, 9, 11-13-tricis and all-cis). The initial product distributions of the

photoisomerizations have been determined as a function of temperature. Relative

quantum yields for photoisomerization of 7-cis and 7,9-dicis-retinal have also been

determined at various temperatures. The results show a general trend of higher torsional

energy barrier values for those double bonds near the electron withdrawing carbonyl

group. The experimental trend is consisted with the bond orders of the conjugated

polyene chain. The four unstable isomers were known to undergo facile stereospecific

rearrangements to their corresponding Ll-transisomers (13-cis, 7, 13-dicis, 9, 13-dicis and

7,9,13-tricis). The enthalpy and entropy values of activation of these thermal

rearrangement reactions, which were monitored by UV/Vis absorption spectroscopy,

iv

have been determined. The data are in agreement with the postulated Kluge-Lillya

mechanism of isomerization, involving consecutive 6e-electrocyclization reactions for the

stereospecific reactions.

Two dicis-retinal isomers, 7,9-dicis-retinal and 9, l l-dicis-l z-fluoro-retinal, have

been incorporated into cattle opsin to yield stable pigments. Therefore, they are available

to study the mechanism of photobleaching of dicis isomeric rhodopsin. The

photochemical reactions of these two pigments have been investigated by low temperature

UV/Vis spectroscopy and HPLC analysis of extracted chromophores. Low temperature

spectroscopies revealed that the batho-intermediates from dicis-rhodopsin analogs are

blue-shifted than that from ll-cis-rhodopsin. HPLC extraction analysis results revealed

that two consecutive steps of one-photon-one-bond isomerization from 9, ll-dicis-12­

fluororhodopsin to 12-fluoro-batho rhodopsin (all-trans) via its 9-cis-intermediate, while

7,9-dicis-rhodopsin demonstrated exclusive two-bond-isomerization to the all-trans

isomer. Possible steric interactions between the isomeric chromophores and the

hydrophobic protein pocket are discussed.

v

TABLE OF CONTENTS

Acknowledgements............................................. III

Abstract............................................................... IV

Table of Contents......................................................................................................... VI

List of Tables......... IX

List of Figures............................................................................................................... X

List of Abbreviations................................................................................................... xv

1. Introduction ..

1.1 Molecular biology of rhodopsin....... 11.2 Spectroscopic properties of rhodopsin................................................................ 81.3 Photob1eaching processes of visual pigments.................................................... 111.4 Visual excitation and cycle................................................................................. 141.5 Raman and infrared studies of visual pigment structures........... 18

1.5.1 Vibrational spectra of retinal isomers......... 191.5.2 Vibrational studies of rhodopsin chromophores....................................... 211.5.3 FfIR difference studies on retinal proteins.............................................. 23

1.6 Studies of chromophore-protein interactions...................................................... 261.6.1 Specific interaction of the substrate-opsin................................................ 261.6.2 Binding studies on isomeric rhodopsin analogs....................................... 271.6.3 Secondary interaction of chromophore and protein................................. 321.6.4 Conformational and configurational properties

of rhodopsin chromophore.......................................... 331.7 Goal of this dissertation............. 34

2. FTIR Studies of Retinal Isomers, Their Schiff Bases andProtonated Schiff Bases................................................................................... 37

2.1 Introduction............................................ 372.1.1 Previous studies......................................................................................... 372.1.2 Raman and infrared spectroscopy... 392.1.3 Vibrational structure of retinaL..... 40

2.2 Materials and methods........................................................................................ 432.2.1 Materials.......... 43

2.2.1.1 Sixteen retinal isomers....... 432.2.1.2 Synthesis of deuterioretinals...................................................... 442.2.1.3 Preparation of 7-cis and 7,9-dicis

retinylidene SB and PSB............................................................ 482.2.2 Methods..................................................................................................... 48

2.2.2.1 FfIR spectroscopy............ 482.2.2.2 Normal mode calculations.......................................................... 49

2.3 Results................................................................................................................. 51

vi

2.3.1 FTIR spectra of sixteen retinal isomers......................... 512.3.2 FTIR spectra of 7-cis and 7,9-dicis-retinal

and their deuterio-analogs.......... 602.3.3 FTIR spectra of 7-cis and 7,9-dicis retinylidene SB

and their deuterio-analogs...................................... 652.3.4 FTIR spectra of 7-cis and 7,9-dicis retinylidene PSB

and their deuterio analogs.. 692.3.5 Normal mode calculation results.............................................................. 74

2.4 Discussion.. 772.4.1 C=O stretches..... 772.4.2 C=C stretches .. 772.4.3 C-C stretches............... 79

2.4.3.1 C-C stretches of retinal isomers................................................ 792.4.3.2 Comparison of retinyl aldehyde, SB

and PSB of 7-cis and 7,9-dicis isomers..................................... 882.4.4 Out-of-plane chain vibrations................................................................... 94

3. Temperature Dependent Isomerization of Retinal Isomers............................... 95

3.1 Introduction......................................................................................................... 953.2 Experiments......................................................................................................... 98

3.2.1 Sample preparation ~............ 983.2.2 Methods.i., 98

3.2.2.1 Irradiation procedure.................................................................. 993.2.2.2 Quantum yield measurements..................................................... 993.2.2.3 Calculation methods 100

3.3 Results 1043.3.1 Photoisomerization of retinal isomers 104

3.3.1.1 Direct irradiation of retinal isomersat different temperatures.. 104

3.3.1.2 Calculations of bond order andenergy barrier of retinal isomers......... 111

3.3.2 Mechanism of the thermal isomerization reactionof retinal isomers with the 11,13-dicis geometry..................... 115

3.3.2.1 Thermal isomerization reaction ofunstable retinal isomers 115

3.3.2.2 FTIR spectra of four thermally unstableretinal isomers 124

3.4 Discussion 1273.4.1 Ground state properties of retinal isomers 1273.4.2 Photochemical properties of retinal isomers... 132

3.4.2.1 Temperature effects on the photoisomerizationof retinal isomers 132

3.4.2.2 Wavelength dependent studies of photoisomerization 135

Vll

4. Photochemical Studies of Rhodopsin Analogs at Low Temperature 137

4.1 Introduction 1374.2 Experiments 142

4.2.1 Procedure for analogs of bovine rhodopsin 1424.2.1.1 Preparation of bovine opsin. 1424.2.1.2 Preparation of 7,9-dicis-rhodopsin 1454.2.1.3 Preparation of 9,II-dicis-12-fluororhodopsin 145

4.2.2 Low-temperature UV/Vis spectrometry 1464.2.3 HPLC chromophore extraction analysis 149

4.3 Results.. 1504.3.1 Effects of NHzOH on dicis-rhodopsins 1504.3.2 Photochemical reactions of dicis-rhodopsins

at liquid nitrogen temperature 1554.3.3 HPLC extraction analysis of chromophores

of dicis-rhodopsins 1634.3.4 Photosensitivity of dicis-rhodopsins

comparing to monocis-rhodopsins 1714.4 Discussion 175

4.4.1 Spectral properties of rhodopsin analogs 1754.4.2 Models to account for absorption properties of pigments...................... 1774.4.3 Mechanism of primary photoreaction of rhodopsin analogs......... 1814.4.4 Postulated models of mechanism of

photoisomerization of rhodopsin ,........... 187

5. Conclustion........................................................................................................... .... 193

References... 195

viii

.~------

LIST OF TABLES

Table

2.1 The C-C stretching frequencies in the region of 1250-1000 em'! ofsixteen retinal isomers......................................................................................... 53

2.2 HOOP band region, 1000-700 em'I, of the sixteen retinal isomers 59

2.3 C-C stretching frequencies of 7-eis- and 7,9-dicis-retinal andtheir deuterio-analogs in the fingerprint region of 1250-1000 crn' 61

2.4 C-C stretching frequencies of retinylidene SB and their deuterio-analogs in the fingerprint region of 1300-1100 em-I........................................ 66

2.5 C-C stretching frequencies of the 1300-1100 cm' region of 7-cis-and 7,9-dicis-retinylidene PSB and their deuterio-analogs............................... 71

2.6 Calculated retinal C-C normal modes and their assignments.......................... 75

2.7 Calculated C-C normal modes of protonated Schiff basesand their assignments......................................................................................... 76

3.1 Experimental and calculated bond distances (A) of all-trans,and cis isomers of retinal '" 102

3.2 Experimental and calculated dihedral angles of retinal isomers...................... 103

3.3 Total and component quantum yields of photoisomerizationof retinal isomers at 298 K and 193 K.......... 111

3.4 Calculated bond order of ground state and first excited stateof retinal isomers using the MNDO method........ 113

3.5 Rate constants of the isomerization of the four labileisomers at various temperatures in methylcyclohexane and thecalculated enthalpy and entropy of activation.................................................. 117

3.6 FfIR frequencies of the double bond stretching modes,single bond stretching modes and HOOP bands of the fourretinal isomers containing the 11,13-dicis geometry...................... 126

IX

LIST OF FIGURES

Figure

1.1 Cross section of the vertebrate eye.................................................................... 5

1.2 Structure of the rod cell..................................................................................... 5

1.3 A modified Hargrave 2-D model of rhodopsin based on thenew knowledge on the lengths of the helices................................................... 6

1.-.+ Helical bundle model of rhodopsin................................................................... 7

1.5 Absorption and circular dichroism spectra of cattle rhodopsin........................ 8

1.6 Photobleaching processes of rhodopsin and iodopsin...................................... 12

1.7 Flow of information in visual excitation and recovery............ 15

1.8 Model of the visual cycle....... 16

1.9 The cycle of rhodopsin and retinochrome......................................................... 17

1.10 Raman and infrared spectra of all-trans-retinal................................................ 20

1.11 Sixteen geometric isomers of retinal... 28

1.12 A 2D map of the binding site of opsin............................................................. 31

1.13 The external point charge model of rhodopsin......................... 32

2.1 A scheme for the synthesis of 7-deuterioretinal............................................... 45

2.2 A scheme for the synthesis of 19,19,19-trideuterioretinal................................ 46

1.3 A scheme for synthesis of 8,19,19, 19-terradeuterioretinal.............................. . 47

2.4 FTIR spectra of all sixteen isomers of retinal in thefingerprint region of 1700-500 cm-I

. .. .. . . . . . .. .. .. .. .. .. . . . .. . .. .. . . .. . .. . . .. .. .. . .. .. .. .. . .. .. .. . .. 54

2.5 Expanded FTIR spectra of all sixteen isomers of retinalin the region of 1250-1000 cm·1

.. . . . .. . .. .. . . . .. . . .. .. .. . . . . .. . .. .. .. .. .. . .. .. . . .. .. .. .. .. .. . . .. .. .. .. . 55

x

2.6 Expanded FfIR spectra of all sixteen isomers of retinalin the region of 1000-700 em·I . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 58

2.7 FfIR spectra (1700-600 em") of 7-cis and 7,9-dicis-retinaland their labelled analogs.................................................................................. 62

2.8 Expanded (1250-990 ern") FfIR spectra of 7-cis and7,9-dicis-retinal and their deuterated analogs................................................... 63

2.9 Correlation diagrams for isotopic shifts of 7-cis and 7,9-dicis-retinal............ 64

2.10 FTIR spectra (1800-600 ern") of 7-cis and 7,9-dieis-retinylideneSB and their deuterated analogs........................................................................ 67

2.11 Expanded FfIR spectra of fingerprint region of 1300-1100 cm' of7,9-dicis-retinylidene SB and its deuterated analogs........................................ 68

2.12 FfIR spectra (1800-600 em:') of 7-cis and 7,9-dieis-retinylidenePSB and their deuterated analogs........... 72

2.13 The FfIR spectra in fingerprint region, 1300-1000 cm', of 7,9-dicis­retinylidene PSB and its deuterated analogs..................................................... 73

2.14 Correlation diagram of the fingerprint frequencies forthe trans and mono-cis retinal isomers...... 80

2.15 Correlation diagram of C-C stretching frequencies of 9-cis,7,9-dieis and 7-eis-retinal................................................................................... 83

2.16 Correlation diagram of C-C stretching frequencies of 7-cis,7,13-dicis and 13-cis isomers of retinal.... 84

2.17 Correlation diagram of C-C stretching frequencies of 9-cis,9,13-dicis and 13-cis isomers of retinal. 84

2.18 Correlation diagram of C-C stretching frequencies of 7-cis,7, l l-dicis and l l-cis isomers of retinal........ 85

2.19 Correlation diagram of C-C stretching frequencies of 9-cis,9,II-dicis and l l-cis isomers of retinal...... 86

2.20 Correlation diagram of C-C stretching frequencies of l l-cis,7, l l-dicis and 7,9,l l-tricis-retinal isomers...... 87

Xl

2.21 Correlation diagram of C-C stretching frequencies of 13-cis,9,13-dicis and 7,9,13-tricis-retinal isomers....................................................... 88

2.22 Correlation diagram of C-C stretching frequencies of all-trans-retinal, its unprotonated and protonated Schiff bases and BRs68••• •• •••••• •• •••••••• 89

2.23 Correlation diagram of the fingerprint vibrations forretinal pronated Schiff base isomers.. 90

2.24 Correlation diagram of C-C stretching frequencies for 7-cis and7,9-dicis-retinal and their unprotonated and protonated Schiff bases.............. 93

3.1 Progress of products formation during direct irradiation ofl l-cis-retinal in hexane at four different temperatures.................................... 106

3.2 Progress of products formation during direct irradiation of9-cis-retinal in hexane at four different temperatures...................................... 107

3.3 Progress of products formation during direct irradiation of7-cis-retinal in hexane at four different temperatures...................................... 108

3.4 Progress of products formation during direct irradiation of7,9-dicis~retinal in hexane at four different temperatures................................ 109

3.5 Plots of product ratio of retinal isomers at different temperatures · 110

3.6 Calculated potential energy barrier of cisisomers in the first excited state............. 114

3.7 UVNis absorption changes during conversion of 1l,13-dicis-retina1to 13-cis-retina1 in methy1cyclohexane 118

3.8 Change of absorbance, 1n(A_-At ) versus time, at 363 nm duringthermal reactions of 11,13-dicis-retinal............................................................. 119

3.9 Change of absorbance, 1n(A_-At) versus time, at 357 nm duringthermal reactions of 7,11,13-tricis-retinal... 120

3.10 Change of absorbance, 1n(A_-At ) versus time, at 359 nm duringthermal reactions of 9,11,13-tricis-retinal......................................................... 121

3.11 Change of absorbance, 1n(A_-AI ) versus time, at 346 nm duringthermal reactions of all-cis-retinal 122

XlI

3.12 Plots of In(k{T) versus 1fT for determination of enthalpy andentropy of activation for isomerization....... 123

3.13 FTIR spectra of the four labile isomers of retinalin the 1700-440 cm' region 125

3.14 The equilibrium of 11-cis-12-s and 11-cis-12-s-trans-retinal 128

3.15 Mechanism for isomerization of dicis-dienals or dienonesto the cis,trans isomers 130

3.16 Scheme of all-cis-retinal isomerization to 7,9, 13-tricis-retina1........................ 131

4. 1 The flow chart of preparation of rhodopsin analogs.... 144

4.2 Diagram of optical cryostat for measuring absorption spectrum atliquid nitrogen temperature or above 147

4.3 Absorption spectrum of 7,9-dicis-rhodopsin in 2% digitonin solution 151

4.4 7,9-Dicis-rhodopsin in 100 mM NH20 H solution at room temperature 152

4.5 Degradation of 7,9-dicis-rhodopsin in 100 mM NH20H at roomtemperature as revealed by a decreased of absorption at 440 nm 153

4.6 Absorption spectrum of 9, 11-dicis-12-fluororhodopsinin 2% digitonin solution 154

4.7 Photochemistry of 7,9-dicis-rhodopsin at liquid nitrogen temperature 158

4.8 Photochemistry of isomeric rhodopsin pigments 159

4.9 Photochemistry of 9,1l-dicis-12-fluororhodopsinat liquid nitrogen temperature.. 161

4.10 Photochemistry of isomeric 12-fluororhodopsin pigments 162

4.11 HPLC chromatograms of chromophore mixture extracted from7,9-dicis-rhodopsin after varying periods of irradiationwith 437 nm light.. 165

4. 12 HPLC chromatograms of retinal isomers obtained from denaturingthe batho intermediate of 7,9-dicis-rhodopsin 166

xiii

4.13 Time plot showing changes in isomeric composition of thechromophore during irradiation of 7,9-dicis-rhodopsin at -190°C......... 167

4.14 HPLC chromatograms of the chromophore extracts obtainedduring the irradiation of a sample of 9,II-dicis-12-fluororhodopsin with 437 nm light 168

4.15 HPLC chromatograms of extracted chromophore of irradiatedsample of 9,II-dicis 12-fluororhodopsin 169

4.16 Progress of irradiation of 9, ll-dicis-12-fluororhodopsin at -190°Cas revealed by HPLC analysis of chromophore extracts......... 170

4.17 Relative photosensitivity of 7,9-dicis-rhodopsin and7-cis-rhodopsin......... 173

4.18 Relative photosensitivity of 9,II-dicis-12-fluororhodopsinand 9-cis-12-fluororhodopsin 174

4.19 Structure of chromophore of 7,9-dicis-rhodopsin comparingwith 9-cis and l l-cis isomers 176

4.20 Schematic showing interconversion among rhodopsin andisomeric rhodopsin analogs and the photoproducts by lightat liquid nitrogen temperature 183

4.21 Computer simulated chromophores of rhodopsin and its analogs.Overlaid chromophore structures of rhodopsin, 9-cis-rhodopsinand 7,9-dicis-rhodopsin............................................................. 188

4.22 Computer simulated chromophores of 12-fluororhodopsin and its analogs.Overlaid chromophore structures of 12-fluororhodopsin,its 9-cis and 7,9-dicis isomers........................................................................... 189

4.23 A possible "bicycle-pedal" path for 7,9-dicis-rhodopsinto bathorhodopsin (all-trans) 192

XIV

BR:c:CD:C-NMR:d:DIBAH:FTIR:GDP:GMP:GTP:HEPES:HOOP:HPLC:HRMS:IR:kDa:LSC:Mt.W:NMR:lH-NMR:PDE:PSB:·R*:Rh:ROS:RR:SB:str:THF:UV:UV/Vis:

LIST OF ABBREVIATIONS

bacteriorhodopsinciscircular dichroismcarbon-nuclear magnetic resonancedicisdiisobutylaluminum hydrideFourier transform infrared spectroscopyguanosine diphosphatecyclic guanosine monophosphateguanosine triphosphate4-(2-hydroxy ethyl)-l-piperazine ethanesulfuric acidhydrogen out-of-planehigh pressure liquid chromatographyhigh resolution mass spectroscopyinfraredKilodaltonlocal symmetry coordinatesmolecular weightnuclear magnetic resonanceproton NMRretinal cGMP phosphodiesteraseprotonated Schiff basephotoexcited rhodopsinrhodopsinrod outer segmentresonance RamanSchiff basestretching modetetrahydrofuranultravioletultraviolet and visible

xv

Chapter 1

Introduction

The structural and chemical changes of prosthetic groups in macromolecules play

an essential role in many biological processes such as the interaction of hormones with

receptors, the catalytic action of enzymes, and photochemical reactions. Among chromo­

proteins, rhodopsin is a major class of biological visual pigment. Rhodopsin consists of

the apoprotein-opsin and l l-cis-retinal, which serves as the chromophore (Wald, 1953).

Upon absorption of light, the chromophore of rhodopsin photoisomerizes to the all-trans

isomer forming bathorhodopsin, an energy rich photoproduct. The photobleaching process

yields all-trans-retinal and opsin as final products (Kropf & Hubbard, 1958).

1.1 Molecular biology of rhodopsin

A vertebrate eye (shown in Figure 1.1) is composed of the cornea, lens, iris, retina,

and epithelium (Shichi, 1983). The cornea is a transparent tissue in the front part of the

eye, which is a permanently fixed lens cover. The lens focuses an image on the retina

by regulating its thickness by using the surrounding ciliary muscle. The space between

lens and retina is filled with a viscous transparent substance called the vitreous humor or

vitreous body which is important for the eye to maintain its shape. The retina contains

visual pigments which absorb the light and then initiate the vision events. The pigmented

1

epithelium, which is required for the nutrition and metabolism of photoreceptor cells, is

also present in the retina but not involved in the transmission of photo signals.

There are two kinds of photoreceptor cells in the retina: rod photoreceptors and

cone photoreceptors (Shichi, 1983). The rod visual cells, which are primarily distributed

in the peripheral region of the retina, function as photoreceptors for dim light (night)

vision. In contrast, the cone visual cells are concentrated in the central region of the

retina and serve as photoreceptors for color vision. Both photoreceptor cells consist of

an inner segment, an outer segment, and the synaptic terminus (O'Brien, 1982), shown

in Figure 1.2. The inner segment is metabolically active. The rod outer segment (ROS)

contains visual pigments and enzymes required for the metabolism of visual pigments.

The ROS is composed of a stack of a few thousand disks enclosed by a plasma

membrane. The disk membrane and plasma membrane have distinct functions in the

visual transduction events. The disks absorb light and transduce light signals into

amplified chemical signals, which can be transduced into electrical signals by the plasma

membrane. Rhodopsin, the photoreceptor molecule in the rod outer segment, is localized

in the disk membranes. It has been found that 95% of the protein in the discs of the rod

photoreceptor cell is rhodopsin (Amis et al., 1981). There are also many other proteins

in cytoplasm, among them G-protein, cyclic GMP phosphodiesterase, a 48 KDa protein

and rhodopsin kinase, are known to be involved in visual transduction (Chabre & Deterre,

1989).

Rhodopsin consists of 348 amino acids with a molecular weight of about 40,000

daltons. Its complete amino acid sequence has been determined (Ovchinnikov et al.,

2

1982; Hargrave et al., 1983; Nathans & Hogness, 1983). However, since the crystal

structure of rhodopsin at high resolution is not available, the 3-dimensional structure of

the protein has not been established.

In order to describe how rhodopsin functions as a photoreceptor, knowledge of the

3-dimensiona1 structure of the protein is essential. Based on the structure of another

membrane protein, bacteriorhodopsin (BR), whose high resolution (3.5 A) crystal

structural data is available (Henderson et aI., 1990), a model structure of vertebrate

rhodopsin has been constructed (Hargrave et al., 1984). This model was later revised

(Mirzadegan & Liu, 1992, shown in Figure 1.3) based on new experimental data

(Zhukovsky & Oprian, 1989; Sakmar et al., 1990). This postulated model has been

partially confirmed by the crystal structural data at low resolution (Schertler et al., 1993).

In this model of rhodopsin, there are 7 helical bundles connected by loops of polypeptide,

see Figure 1.4. The inner surfaces of the helices define the binding pocket for the retinal

chromophore.

The disc membrane faces an external cytoplasm and an aqueous intradiscal

interior. The discal interior acts as a boundary enclosing a lipid layer approximately 28

A thick (Saibel et al., 1976). The total bilayer membrane width is approximately 40 A.

The center-to-center repeat spacing of the discs is approximately 300 A and the

cytoplasmic layer between disc membrane borders is less than 150 A thick (Chabre,

1985). The distance between rhodopsin molecules aligned along the axis perpendicular

to the dies is 55 A (Chabre, 1975) indicating the monomeric nature of rhodopsin.

3

Based on linear dichroism data, the l l-cis retiny I chromophoric portion of

rhodopsin is believed to be approximately parallel to the disc membrane with a small

angle of 16-20° between the chromophore and the membrane plane. It has generally been

accepted that the retinyl chromophore is located close to the cytoplasmic side of the

membrane in the core of the seven helices (Abdulaev, 1986).

Based on chemical and enzymatic modification studies, it has been found that the

nature of the binding site is hydrophobic (Hargrave, 1982). In this predominantly

hydrophobic binding pocket, only a few charged amino acids are present in the helices.

In the model shown in Figure 1.3, there are three positively charge amino acids (LyS296'

Arg13s, His 211) , which are shown as shaded heavy circles and three negatively charged

amino acids (Glu 134, GlUm, ASPs3)' which are shown as shaded squares. The helices also

contain a total of twelve polar amino acids, serine and threonine, which are required for

hydrogen-bonding. Most of the charged amino acids are present in the loops and on the

internal aqueous exposed surface of rhodopsin (Hargrave et al., 1983).

4

-~-~-~

Rctill<J-~~

Pigmcntcd/""cpithcliurn

Lioht--,...

Vitreous humor

-,~_._-- .-,-.---,--- ---

Figure 1.1 Cross section of the vertebrate eye. From Shichi, H. Biochemistry of Vision.

In "Structure of the eye", Academic Press, New York, 1983.

Oulersegment

Innersegmenl

Discs

Plasmamembrane

Cyloplasmicspace

1==1-- tntradlscatspncc

---"""'''--Cilium

Milochondrlon

Nucleus

Synapticlermlnnl

Figure 1.2 Structure of the rod cell (O'Brien, 1982).

5

0\

Figure 1.3 A modified Hargrave 2-D model of rhodopsin based on the new knowledge on the lengths of the helices (fromMirzadegan & Liu, 1992). The amino acid residues that have been displaced from those in the originalHargrave's model (Hargrave et al., 1984) are marked by those circles with the bottom half darkened.

oDisc - r- 300A Repeat Spacing

Membrane

oC Jj_ Cytoplasmic Space - < 150AC :>C ::>c :>C 311- Intradiscal SpaceC :>c :>

~J~~ -30X ~

..............................

/..... . .

Cytoplasm Sido I / \

TnR_7.J \8

28A 2 i •.

I. \~6A \v \

"-Olll k tnterlor I \ ..•.

.......••.......-::::::-.....

~..>­OJ

lD

"t:I

.~

...J-..l

Figure 1.4 Helical bundle model of rhodopsin (Hargrave et al., 1984).

1.2 Spectroscopic properties of rhodopsin

Visual pigments show three major absorption bands (Figure 1.5), which are termed

the a, ~ and y bands from the longer to the shorter wavelength (Honig et al., 1973). The

a (500 nm) and ~ (350 nm) bands of rhodopsin are due to the l l-cis-retinyl

chromophore, where the y (280 nm) band is attributed to aromatic amino acid residues

of the opsin.

..Io

:::E 8tnoa::z:<.>o

~ 4.J::><.>a: 2o

OI-----,.---------o;::IL--.~

0.4>-l-f/)

z0.3 YUI

0

.JoCtoi= 0.2Q.0

0.1

300 400 500 600WAVELENGTH (nm)

Figure 1.5 Absorption (lower) and circular dichroism (above) spectra of cattle rhodopsin

(Honig et al., 1973).

8

The protonated Schiff base (PSB) of retinal with n-butylamine, which is the model

compound of rhodopsin, absorbs at 445 nm in the organic solvent MeOH, whereas

rhodopsin, the chromophore of which is also a PSB, absorbs at 500 nm. The extent of

the red shift, expressed in crn', has been defined as the opsin shift (Nakanishi et al.,

1980). It represents the overall environmental effect of the protein binding site on the

absorption maxima of the visual pigment. Specifically, the effect must be due to

interactions of the retinyl chromophore with specific amino acid side chains of opsin.

Thus, the varying spectral properties of rhodopsins must be due to the different amino

acid sequences or different interactions between chromophore and protein in each

pigment.

The visual pigment chromophore exhibits optical activity which is shown in the

circular dichroism spectrum in Figure 1.5. However, retinal isomers in solution do not

show any optical activity. There are two possible sources for the optical activity of the,

visual pigments. An asymmetric environment in the region of the protein near the

chromophore would induce optical asymmetry on the part of retinyl chromophore.

Alternatively, the opsin protein may bind selectively only one enantiomeric form of 11-

cis-retinal due to twisting induced by the surrounding protein (Balogh-Nair & Nakanishi,

1990).

Circular dichroism spectra provide useful structural information for

macromolecules such as proteins and nucleic acids. The retinylidene chromophore of

rhodopsin has two distinct circular dichroism bands at 487 and 335 nm corresponding to

the a and ~ absorption bands, respectively (Shaw, 1972). But their intensity ratios differ

9

considerably among the rhodopsin analogs (Kropf et al., 1973). Comparison of the

intensities of the a. and ~-bands in the CD spectra of rhodopsin and rhodopsin analogs

suggested that the a.-band arises principally as a consequence of the interaction with opsin

on the portion of the polyene chain near the Schiff base linkage. The magnitude of the

~-band in the CD appears to be influenced by both the structure of the substituted

cyclohexenyl ring and its interaction with the protein (Balogh-Nair & Nakanishi, 1990).

10

1.3 Photobleaching of visual pigments

There are two kinds of visual pigments; one is rhodopsin, a rod visual pigment

responsible for the scotopic vision in the nightlight environment and the other is iodopsin,

a cone pigment responsible for the photopic vision in daylight. The photochemical

bleaching schemes of rhodopsin and iodopsin are shown in Figure 1.6, the main path of

the enzyme cascade reactions presumably are common to both rod and cone visual

pigments (Yoshizawa, 1993). The critical difference in physiological function between

the rod and cone pigment is the difference in their photosensitivities. Electrophysiological

studies have shown that the cone pigment is about one hundred times less sensitive than

the rod pigment (Schnapf and Baylor, 1986).

The retinal opsin interaction changes in discrete steps which are spectrally

identified as intermediates. These have been widely investigated by means of low

temperature spectrophotometry and room temperature laser photolysis, resulting in the

identification of several intermediates appearing in the bleaching processes (Shichida,

1986).

The first intermediate observed during the photobleaching of rhodopsin is called

photorhodopsin with ~ax =570 nm, detected when rhodopsin is irradiated with yellow

light (540 nm) at liquid helium temperature (-268°C). When the temperature goes to ­

250°C, this intermediate is converted to the next intermediate, bathorhodopsin with ~ax

= 535 nm. Application of pico second laser spectroscopy clearly demonstrated that

bathorhodopsin was produced at room temperature with a time constant of 45 pico

seconds

11

Rod visual pigment Cone visual pigment

Iodopsin

UhvExcited state

~ 200 IsPhotolodopsin

~ • ps

Bamoiodoosfn

cr-i 130 ns

Lumiiodopsin

t 67 I1sMetaiodopsin I

+6 ms

Metaiodopsin II

J~5 s

All-trans retinal+ Photopsin

conerod

GMP cGMP

~

Transducin·GDP

=====~:- f .~:=====Transducin·GTP

/Phosphodiesterase _ Phosphodiesterase(inactive form) (active form)

cGMP ...t. 5'-GMP

cGMP-Na-channel (open state) L Na-channel (closed state)

tGeneration of receptorpotential

Rhodopsin

UhVExcited state

1 200 Is

Photofhodopsin

l 45 psBathorhodopsinl 250 ns

Lumirhodopsin

l 120 I1s

Metarhodopsin IJ12 ms

Metarhodopsin II

l 520 sMetarhodopsin III

J> 1 h

AII·trans retinal+ Scotopsin

Figure 1.6 Photobleaching processes of rhodopsin and iodopsin, and the enzyme

cascade system common to both rods and cones (Yoshizawa, 1993).

12

(Shichidaet al., 1984). Recently, it was confirmed using a femto second laser photolysis

that photorhodopsin is produced directly from the excited state with a time constant of

about 200 femto seconds (Schoenlein et al., 1991).

Conversion of the batho-intermediate to the rest of the intermediates is completed

by thermal decay reactions. Compared to the early intermediate formation from

rhodopsin, the transition of bathorhodopsin to lumirhodopsin at room temperature is a

slow process (l0-100 nsec at room temperature). The meta I intermediate is formed from

lumirhodopsin within a few microseconds at physiological temperature. Then,

metarhodopsin I decays to metarhodopsin II with a lifetime of a few milliseconds. Time

constants for the formation of these intermediates are very important for the study of the

visual transduction processes. The rate of meta I to meta II conversion is markedly

influenced by the environment (Hofmann, 1986),

Similar photobleaching processes were detected in the iodopsin system using time

resolved laser photolyses (Kandori et al., 1990). Thus, photoisomerization of the

retinylidene chromophore from the l l-cis form to the all-trans form, is applicable to not

only scotopic vision, but also photopic vision as the primary process of vision.

Unlike rhodopsin, iodopsin has a chloride binding site which is responsible for

color regulation. The absorption maximum of iodopsin is at 571 nm when in chloride

binding form and at 512 nm when in the chloride depleted form (Shichida et al., 1990).

The later intermediates of the photobleaching process of iodopsin are approximately

identical to those of rhodopsin (Imamoto et al., 1991). In the rhodopsin system, one

molecule of meta II can catalyze about 500 molecules of transducin from the inactive

13

form to the active form. Since the extent of amplification of the photoresponse of the

photoreceptor cells is directly correlated with the amount of transducins which is activated

by meta II in its life time, the longer life time of meta II in the rhodopsin system

compared to that of iodopsin (Figure 1.6), accounts for the higher sensitivity of the rod

visual pigment.

1.4 Visual excitation and cycle

The visual transduction process is that visual pigments absorb light and convert

the light signal to an electronic signal which is transmitted to the brain via the optic

nerve. Visual perception is initiated by the absorption of photons by rhodopsin. Figure

1.7 shows the flow of information during the visual excitation and recovery (for reviews

see Stryer, 199t", Pugh &. Lamb, 1990, Chabre & Oeterre, 1989, Liebman et al., 1987).

When rhodopsin absorbs a photon of light, it photobleaches to all-trans retinal and opsin

through a series of intermediares. The photoisomerization of the l l-cis retinal

chromophore of rhodopsin (R) to the all-trans form generates photoexcited rhodopsin (R").

It is believed that the metarhodopsin II is the key intermediate which activates a series

of enzymes (Stryer, 1985) in a "cyclic nucleotide cascade". Accordingly, R" then

activates transducin (T), a member of the G-protein family, by catalyzing the exchange

of GTP for bound GOP. The GTP form of transducin (specifically, Ta-GTP, the

activated a-subunit), in turn, switches on a potent phosphodiesterase (POE) that rapidly

hydrolyses cOMPo cGMP regulates the opening and closure of the gate of plasma

membrane channels. In the dark, Na' and Ca2+ enter the outer segment through cation-

14

specific channels, which are kept open by cGMP. The light-induced decrease in the level

of cGMP closes these channels, which hyperpolarize the plasma membrane. The visual

process is a rapid and amplified transduction process (Liebman & Pugh, 1980; Yee &

Liebman, 1978). As a result, a single photoexcited rhodopsin can lead to the hydrolyses

of more than lOS molecules of cGMP. The resulting hyperpolarization is conveyed to the

synapse in the inner segment of the rod cell, which communicates with other neural cells

in the retina.

R hv • •- R -Ta-GTP- POE - cGMPl.

•GC - Recoverin - Ca.J.I -

jChannelClosure

jcGMPi - Channel

Opening

jMembrane

Hyperpolarization

Figure 1.7 Flow of information in visual excitation and recovery. Photoisomerization

of rhodopsin triggers a cascade leading to cGMP hydrolysis and the closure of

membrane channels. Channel closure cause the cytosolic calcium level decrease,

which leads to the activation of guanylate cyclase and the reopening of channels.

oc', activated guanylate cyclase (adapted from Stryer, 1991).

Recovery of the dark state is mediated by deactivation of PDE and activation of

guanylate cyclase by the closure of membrane channels (Lamb & Pugh, 1992).

15

----'----------- ---_.... -------- . ------- ------ ----

Stimulation of guanylated cyclase by recoverin increases the cGMP level, which reopens

channels and restores the dark state.

Photoactivated rhodopsin decays thermally to the apoprotein-opsin and all-trans-

retinal. The isomerization of all-trans-retinal to l l-cis-retinal in the pigmented epithelium

membranes closes the visual cycle. The model of the visual cycle is shown in Figure 1.8.

The all-trans-retinal is released from opsin after photobleaching, reduced to retinol, and

transported to the pigmented epithelium for storage. This process is rather slow

(Matthews et al., 1963). It has been shown that pigment epithelium membranes can

process added all-trans-retinol to produce an all-trans-retinyl ester that is directly

transformed into 11-cis-retinol by an isomerhydrolase enzyme (Rando, 1992). Retinol

returns to the retina, is oxidized to l l-cis-retinal, and used for regeneration of rhodopsin.

Rhodopsin

o~ ~11--'.Js-retlnal all-trans-retlnal

~ pro 5 t~ pro R

11-..cls-retlnol all-1r..ilDs-retlnol

~ ~11-cls-retlnyl esters all-trans-retinyl esters

Figure 1.8 Model of the visual cycle (Rando, 1992).

16

In cephalopods, there is another retinal binding pigment, retinochrome. which is

stored in the inner segments whereas rhodopsin is located in the outer segment in the

invertebrate eye (Hara-Nashimura, et al .. 1990). The configuration of the bound retinal

molecule is assigned to be all-trans on the basis of extraction of the chromophore by

HPLC analysis and NMR spectrometry (Hara-Nishimura et al., 1990). The relationship

between retinochrome and rhodopsin for the configuration of the retinal molecule is

shown in the scheme in Figure 1.9 (Tsujimoto et al., 1993). Photobleaching of

retinochrome, constituted by aporetinochrome and all-trans-retinal, results in l l-cis-retinal.

Then, l l-cis-retinal can be incorporated with opsin to regenerate rhodopsin. The function

of retinochrome is complementary to rhodopsin.

.-- ......A'- -.r ,

all-tran s-reti nal

Rhodopsin

11-cis-retinal Opsin'"-------.. .-------,)v

1'-- -1

Figure 1.9 The cycle of rhodopsin and retinochrome (Tsujirnoto et al., 1993).

17

1.5 Raman and infrared studies of visual pigment structures

To understand the molecular basis for the function of rhodopsin, it is necessary

to determine the structure of the chromophore, and its interaction with amino acid

residues in visual pigments and in their photochemical intermediates. For such

investigations, resonance Raman and infrared spectroscopy have proven to be important

tools.

Information from vibrational spectra of retinal proteins can be classified into two

categories: the chromophore and the protein. Vibrational information about the retinyl

chromophore has been obtained from resonance Raman and FfIR spectra while protein

vibrations are mainly obtained from FfIR techniques. The resonance Raman technique

takes advantage of intensity enhancement of Raman bands of a chromophore when the

excitation wavelength approaches the absorption maximum of the species to be examined.

Since different photoproducts usually have absorption maxima at different wavelengths,

tuning of the excitation wavelength enables selective detection of a particular intermediate

among several coexisting photointermediates (see Rothschild, 1988; Kitagawa & Maeda,

1989 for reviews). In contrast with RR spectroscopy, many vibrations of the

chromophore, protein and lipids are, in principle, infrared active; therefore IR spectra are

more complicated. In order to circumvent this complexity, a light induced difference

spectrum at low temperature is used to reveal changes in protein structure; e.g.

protonation/deprotonation states of carboxylic acid side chains (Engelhard et al., 1985;

Eisenstein et al., 1987; Roepe et al., 1987), of tyrosine residues (Rothschild et aI., 1986;

18

Dollinger et al., 1986; Lin et al., 1987) and membrane structures (Rath et al., 1991;

Fahmy, 1992).

1.5.1 Vibrational spectra of retinal isomers

The geometric isomers of retinal are biologically important as the chromophore

in visual receptors. The interpretation of the visual pigment spectra requires a more

detailed assignment of the vibrational normal modes. To facilitate vibrational studies of

the chromophore in visual pigments, it became necessary to make complete assignments

of the retinal isomers as model compounds. The assigned vibrational spectra of all-trans­

retinal is shown in Figure 1.10, as a representative example. The assignments are based

on the spectra of over one hundred D- and l3C-substituted derivatives (Curry et al., 1982).

The vibrational spectrum of all-trans-retinal in the region of 1700 - 500 ern:'

mainly includes the C=O stretching, C=C stretching, C-C stretching, hydrogen out-of­

plane bending (HOOP) and hydrogen in-plane wagging modes. Among these, the C-C

and hydrogen out-of-plane bending (HOOP) vibrational modes are most sensitive to

conformational changes and environment changes and can be used as a probe to study the

chromophore geometry and the interaction between chromophore and protein. For

example, the Vee frequencies are sensitive to the torsion around the C-C bond through

coupling with the C-C=C bending vibration. Removing this coupling in the cis

conformation lowers the Vc.c frequency by about 100 cm' (Smith et al., 1986). The fact

that the Yc.c frequency of the s-cis conformation is 100 cm' lower than that of the s-trans

conformation can be utilized to obtain the conformational information about single bonds.

19

~ ... Absorbance tntensuy ~riQ"C.,I'D- m- gr ::J

\r- 596

.oQ ..... ccc ~er.d -< -6510

~

3:;:0 0~ ~ 03 co ::J~ o,::3

~ co{3- -onone

::3792P- O

5° 0 Meslr ~ - 849::t' 14 HOOP - 866 Pe; ------ \0 HOOP - 8-:,0 --(l)

IP- __________ ·11 + 12 HOOP~ /959 ---..(fl - 966 7+8 HOOP -367 "'"Cl 0 ~(l)

Me r ocx 1008 ~() 0....

In'"10 -1045~ ...,

Iv 0~CI4-CI5~ co...... I II I

~0

~ 1123-~CIO-CII~

-1134- =:l,..!.. - -1163 0""1 N 1164-

1198 ~C8-C9~~

1197::30 1215 C12-C13

(flI

0""1

1270-IIH P1270

(l)....5" 1281- 1279~ 7H-BH ~= 1302e:. 1333

~ ~ 1334,.-.. - 1360 ~ 12H(Jt:: ~ 1385 14H 1389~ 0

1446 Me del r~ 1446t'1:> 0....l::l:-

mL 1550~ ----CI3=CI4 ------......\0

-157700 1576 -- ~ --C7=C8+C9=CI0+CII=C12--Vl 0'-"

0'1660 C==O :> 1663

1.5.2 Vibrational studies of rhodopsin chromophores

To maximize the information obtained from the resonance Raman and FTIR,

specific vibrations have to be assigned to extract the detailed structural information

inherent in the spectra (Mathies et al., 1989). These problems can be surmounted by

regenerating the rhodopsin with isotopically labelled retinyl chromophores prepared by

total organic synthesis. The method is time consuming but has the obvious advantage of

not leading to changes in steric interactions and electronic properties of the molecule

(Curry et al., 1984). Through the effort of Lugtenberg, Mathies and coworkers (Mathies

et al., 1989), specific isotopic labelling of the retinal prosthetic group of the chromophore

of the visual pigment has made it possible to study the structure and function of these

pigments in exquisite detail using laser resonance Raman scattering and infrared

spectroscopy.

With the vibrational spectra of the model retinals, the following structural features

of the retinyl chromophore can determined (Mathies et al., 1989). The procedure involves

using isotopically labelled derivatives to assign the vibrational lines to specific normal

modes and comparing these results with model compound assignments and the predictions

of normal coordinate calculations.

1. Geometry of isomers of chromophore

2. Linkage of the chromophore to the protein

3. Conformational distortions of any single or double bonds.

4. Secondary interactions between protein and chromophore.

21

Geometrical isomers of the retinyl chromophore can be identified by comparison

of the vibrational spectra of the retinyl chromophore in rhodopsin with the model

compound PSB (Mathies et al., 1977). The frequencies of the normal modes of rhodopsin

in the fingerprint region are very close to those of l l-cis PSB. Thus, the in situ

configuration of retinal in rhodopsin is l l-cis, Similarly, other geometrical retinal

isomers of other rhodopsin analogs and their intermediates during the photobleaching

process can be determined in the same manner. The linkage between the retinal

chromophore and the protein in rhodopsin is a protonated Schiff base bond with a lysine

residue. The evidence for this is the 1655 ern" line which has been assigned as a C=NH+

stretch which shifts to 1630 cm' in D20 (Mathies et al., 1976; Narva & Callender, 1980).

In contrast, the C=N stretch in the Schiff base reveals difficulty in this shift upon

deuteration (Narva & Callender, 1980).

The chrornophore-protein interaction in rhodopsin also can be determined using

vibrational spectra. The shift of the absorption maximum of the l l-cis PSB from about

440 to 498 nm upon binding is accompanied by two other significant changes in the

vibrational spectrum (Mathies et al., 1977). The ethylenic band shifts from 1556 cm' in

the ll-cis PSB to 1545 em:' in rhodopsin, and the hydrogen-out-of-plane (HOOP) band

at 971 ern" significantly increases in intensity. In isorhodopsin, similar changes have

been observed (Mathies et aI., 1977). The frequency decrease of the ethylenic band is

the result of a more delocalized electronic structure, with a lower C=C bond order, and

a higher adjacent C-C bond order. HOOP modes are weak in the Raman spectra of

polyenes in the planar form but have significant intensity if the polyene experiences the

22

out-of-plane distortion ch as C=C or C-C twists (Eyring et al., 1980, 1982). In

rhodopsin, the intensi., of the HOOP mode indicates that there is a conformational

distortion of the CIO-Cll=CI2-C13 moiety. In bathorhodopsin, the unusually intense HOOP

mode was also rationalized as due to the conformational distortion in the CIO to Cl4 region

(Eyring et al., 1982). The perturbation of the protein was presumed to arise from a

negatively charged protein residue that acts as a "point charge" in rhodopsin (Eyring et

al., 1982; Honig et al., 1979a).

A large number of 0- and l3C-isotopic derivatives have been used to complete a

detailed analysis of the vibrational normal modes of other retinal binding visual pigments,

e.g. BR568 and BR548 (Tsuda et al., 1980; Pettei et al., 1977). The important result is that

the vibrational lines in the 1100-1300 ern" fingerprint region can be readily assigned to

specific C-C and C-H normal modes which can .be used to probe structural features at

specific locations.

1.5.3 FTIR difference studies on retinal proteins

FfIR spectroscopy is an independent technique for looking at chromophore

vibrations with the advantage that the energy of the ir photon is too small to disturb the

system. More importantly, it reveals changes in the apoprotein which could not be

detected by resonance Raman. However, since there are too many vibrational modes, due

to the chromophore (MW = 294) or a single amino acid residue are often buried under

those due to the apoprotein (Rh MW = 40,000, BR MW = 28,000). This problem can

be solved by the FfIR difference technique.

23

The analysis of such a spectrum and comparison with RR results could provide

the following structural information:

1) the protein chromophore linkage, namely, whether the Schiff base is

protonated or not.

2) changes in the chromophore.

3) changes in the protein moiety, particularly any changes of carboxylic acid

residue.

Carboxylic acid residues are found dispersed throughout rhodopsin. Examination

of the sequence map of rhodopsin shows that the carboxylic acid residues lie close to the

imine linkage regions. Carboxylic acid residues have been implicated as the possible

perturbation in the point charge model (Honig et al., 1979). Difference FTIR

spectroscopy at low temperature for photosensitive pigments has been found to be a very

useful technique for detecting small changes in the protein (Braiman & Rothschild, 1988;

Kitagawa & Maeda, 1989).

Protonated carboxylic side chains of amino acid residues show the C=O stretching

mode around 1720-1760 cm', and the CO2' antisymmetric and symmetric stretching

modes around 1610-1550 and 1450-1390 cm', respectively. By using specifically labelled

amino acid(s) incorporated into the purple membrane, FTIR bands can be assigned to

certain amino acid side chains (Eisenstein et al., 1987).

In the FTIR study of rhodopsin, changes in two carboxylic acids with vc=o bands

at 1772 and 1734 cm' are observed for upon conversion of rhodopsin to bathorhodopsin

24

and from rhodopsin to lumirhodopsin (De Grip et al., 1988; Ganter et al., 1988). Upon

conversion from rhodopsin to metarhodopsin I, deprotonation of one carboxylic acid

(1737 ern") and concomitant protonation of another carboxylic acid (1701 cm') take

place. Upon formation of meta II, the deprotonated carboxylic acid (1737 cm') becomes

protonated and another unprotonated carboxylic acid in rhodopsin becomes protonated

(1746 em:') (Rothschild et al., 1986; Ganter et al., 1989).

Recently, functionally important amino acid residues have been identified through

selective mutagenesis. The combination of site-directed mutagenesis with FTIR difference

spectroscopy appears especially powerful for studying the functional role of specific

amino acids (Gerwart et al., 1989, Rothschild et al., 1990, Fahmy et al., 1992).

25

1.6 Studies of chromophore-protein interactions

There have been a number of considerations of the interaction between the

chromophore and the protein. Studies with visual pigment analogs began with the

geometric isomers of retinal (all-trans, 13-cis, ll-cis, 9-cis, 9,13-dicis and 1l,13-dicis

(Hubbard & Wald, 1952; Wald et al., 1956). This was followed by a series of

structurally diverse analogs to reveal conformational properties of the chromophore and

protein-substrate interaction (review see Balogh-Nair & Nakanishi, 1986). The specific

protein-substrate interactions are expected to have an influence on chemical processes of

the chromophore.

1.6.1 Specific interaction of the substrate and opsin

As early as 1967, it was shown that the retinyl chromophore was attached 'to a

lysine residue through a Schiff base linkage (Bownds, 1967), which the latter was shown

to be lys-296 (Wang et aI., 1980). Based on vibrational spectroscopic studies (Paling et

al., 1987; Bagley et al., 1985), the linkage was unambiguously shown to be a protonated

Schiff base. Site-specific mutation results show that Glu-113 in bovine rhodopsin appears

to be the counterion to the protonated Schiff base (Zhukovsky & Oprian, 1989; Sakmar

et al., 1990). However, Be NMR studies indicated that the proton of the Schiff base is

hydrogen-bonded (Smith et al., 1985).

Analog binding studies suggest a strong protein substrate interaction near the ring

portion of the molecule by the evidence that modification of the chromophore near the

26

ring drastically changes the nature of the photochemical reaction. A specific hydrophobic

interaction of the apoprotein of rhodopsin with the trimethylcyclohexenyl ring is involved

(Kropf et al., 1973). Recognition of each of the methyl groups contributes to pigment

formation because removal of anyone of the methyl groups on the ring resulted in greatly

reduced rates of pigment formation, e.g. 5-demethylretinal formed a pigment with opsin

at a very slow rate (Kropf et al., 1973; Kropf, 1976). Furthermore, pigment formation

of acyclic substrates of open-ring structure with opsin indicated that the binding site does

not recognize the ring itself (Rao et al., 1985). Instead, it is the methyl groups on the

ring which are recognized by the protein during pigment formation.

Modification of the 5-methyl group has been conducted in a series of ex. and ~­

retinal analogs (see review Liu & Asato, 1990). The results indicate that there is

substantial free space available in this region of the binding site of rhodopsin.

Recognition of the 9-methyl group is reflected in another unique way. Studies of

9-demethyl rhodopsin analogs have shown a different interaction between the

chromophore and opsin by evidence of the absence of the hydrogen-out-of-plane (HOOP)

band of Hll, Hl2 of the bathorhodopsin intermediate (Ganter et al., 1988). Binding

studies showed that removal of the 13-methyl group has a relatively small effect on the

substrate-opsin interaction (Shichida et al., 1981).

1.6.2 Binding studies on isomeric rhodopsin analogs

All sixteen possible geometric isomers of retinal are known (shown in Figure

1.11). Among these sixteen isomers, the four containing 11,13-dicis geometry are not

27

stable at room temperature (Wald et al.. 1955; Knudsen et al., 1980; Trehan & Liu, 1988;

Trehan et al., 1990b), hence are unsuitable for binding and photoisomerization studies.

Of the remaining twelve isomers, ten were shown to react with opsin to give isomeric

rhodopsin analogs (Liu et al., 1984b). The two exceptions are the all-trans and 13-cis

isomers.

CHO

~ CHO

9-c is~~ CHO ~I -.;:: ~ I ~ ....

all-trans - ....~11-C1S CHO

~' - ACHO

11.13-dicis 9.13-dicis

~ CHO

~CHO' 7-~i: - CHO~~CHO7.lJ-dicis 7.9-dicis 7.9.13-tricis

~13 -cis CHO

~ '_/CHO

lvll-~ 7.11-dicis ~ 7.11,13-tricisCHO

~ I 9.11,13-tricis~~

CHO~I -.;:: .... CHO 9.11-dicis

e-, :-0..

CHO

~ all-cis~ 7.9.11-tricisCHO

Figure 1.11 Sixteen geometric isomers of retinal.

Verification of retention of configuration via chromophore extraction experiments

has so far been conducted on seven isomers: 7-cis, 9-cis, l l-cis, 7,9-dicis, 7, 13-dicis,

28

9,13-dicis, and 9,11-dicis (Groenenduk et al., 1979; Maeda et al., 1979, Shichida et al.,

1988b; Trehan et al., 1990a). These results appear to suggest that not only the singly

bent isomers are capable of forming isomeric pigments with bovine opsin but so are the

doubly and triply bent isomers. However, the rates of pigment formation were found to

be quite different. The structurally similar l l-cis and 9-cis isomers yielded pigments

most readily, while the others at considerably reduced rates (Liu et al., 1984a). The

binding cavity of rhodopsin appears to be able to readjust itself in order to accommodate

substrates of different shapes. The failure of binding all-trans and 13-cis to bovine opsin

was first explained by a longitudinal restriction of binding pocket (Matsumoto et al.,

1978).

Based on binding studies of retinal isomers, a 3D model of the binding site of

opsin has been constructed (Mirzadegan, 1989). In this model, all the binding isomers

of retinal were overlaid with their two ends (Co; of lysine 296 and the cyclohexenyl ring)

fixed and the van der Waals radii of carbon atoms were outlined (shown in Figure 1.12).

It was aiso shown that the failure of the 13-cis isomer to bind with opsin is not due to

a longitudinal restriction but the interference with a forbidden region probably occupied

by the counterion.

The analyses of chromophore extraction of isomeric rhodopsin analogs have

revealed that the catalyzed chromophore isomerization by opsin takes place concurrently

with pigment formation (Trehan et al., 1990a). Double bonds near the carbonyl group

are more sensitive to the catalyzed isomerization. A mechanism of catalyzed

isomerization by opsin has been proposed by Sack and Seltzer (Sack & Seltzer, 1978).

29

This mechanism involves a reversible Michael addition of a nucleophile from the protein

to an electrophilic carbon of the polyene. Preferential isomerization at the 13,14 double

bond is a reflection of the more electrophilic character near the carbonyl group. Hence

only the 7,9-dicis isomer is unaffected by this process. All other dicis pigments are

affected by this catalyzed isomerization. 9,1l-Dicis pigment retains about 40% of its

original geometry accompanied by 60% isomerization to the 9-cis product; 9, 13-dicis

retains 0-28% giving a large amount of the 9-cis product; 7,13-dicis is nearly completely

isomerized to the 7-cis product.

30

oQ

'0•

...,

..,r

,

,~ • Y :\7

• • ,'\7"", • • :. v 11~. ~~.. ~.. "~'-, 12. , ", "Kt... ~-, •• ".-:--0 " '_,'_."· :" u "", v ":":-",'- \7 -'-:'0;' r7 ':.() , '., , .... V ,

~' ,(; U \7 '. .:' .".. '. • \7 nI 'f..J.... o •. .¥"

• I " " .. ,','_ \I '.. Vo _ -, ,

.It-'\I n.. n-- ":' .• __1.)4\. (\VO',_ '. o. ..... ... .. ~

.. ". ,,'"#, "- \ • • .0 . ,(\..>...,. • ;\ '. • • 0~.. ~ o. • • 0 :

r... •• • 0 "

Y" • '.. \ ~ -, 0. . ' ............. 'N.' ,. ..

'..": \7 ,

,.. n.. ...._-"' __ ': v·., .. .

I

• : , 0

J!..---.'1fV1 0o •.

\I.

V.>

Figure 1.12 A 2D map of the binding site of opsin: Overlay of carbon atoms of all ten binding isomers of tetheredchromophores of isomeric rhodopsin (solid dots). Superimposed are carbon atoms of the non binding isomers: I3-cis(triangles) and all-trans (circle). The dashed straight lines are the carbon framework of the l l-cis isomer. Themaximum perimeter of the binding site is defined by the van der Waals radii of the outer carbons of the bindingisomers. The shaded area is occupied by the non binding I3-cis isomers (Mirzadegan, 1989).

1.6.3 Secondary interaction of chromophore and protein

The spectral difference between the protonated Schiff base (440 nm) and rhodopsin

(500 nm) has been proposed to be due to non-covalent interactions between the

chromophore and the protein, in addition to the covalent Schiff base linkage. The

external point charge model (shown in Figure 1.13) was proposed based on studies of

synthetic dihydroretinals (Honig et al., 1979a: Kakitani et al., 1985). According to this

model, there are two negative charges near the chromophore; a counterion at 3 A from

the protonated Schiff base nitrogen and a second negative charge at 3 A from C-12 and

C-14. Recent l3C NMR studies support the presence of protein-induced perturbations in

the C12-C14 region (Smith et al., 1991).

Figure 1.13 The external point charge model of rhodopsin (Honig et al, 1979b). The

existence of a counterion near the protonated nitrogen is assumed. A second

negative charge is located at - 3 A from C-12 and C-14.

32

The source of the point charges could be charged or polar amino acid(s) residues

near the retinyl chromophore. In rhodopsin, most of the polar amino acid residues are

found as expected outside the membrane region, within the protruding loops. There are

three positively charged residues (Arg-l35, His-I22 and Lys-296) in helices 3, 5 and 7,

and four negatively charges residues (Asp-83, Glu-Il3, Glu-I22 and Glu-I34) in helices

2 and 3 of the transmembrane segments, as mentioned in section 1.1 in Figure 1.3. There

are additional charged residues in the loop and end segments (Hargrave et aI., 1984).

Site specific mutation studies have been carried out to elucidate the role of these

amino acid residues (Zhukovsky & Oprian, 1989; Sakmar et aI., 1990; Nathans, 1990).

Upon mutation of each of the charged residues (Asp-83, Glu-Il3, Glu-I22, Glu-I34, Arg­

135, and His-I22) it was shown that only replacement of Glu-1I3 by GIn caused a drastic

shift in the absorption maximum of rhodopsin (from 500 to 380 nm) (Zhukovsky &

Oprian, 1989; Sakmar et al., 1990). This gave experimental evidence that Glu-II3 is the

likely counter-ion to the protonated Schiff base. Also, Glu-I22 and Asp-83 are thought

to be involved in the receptor activation process, and Glu-I34 and Arg-l35 are proposed

to interact with transducin (Zhukovsky & Oprian, 1989; Sakmar et al., 1990; Nathans,

1990).

1.6.4 Conformational and configurational properties of the rhodopsin chromophore

Another aspect of interaction is the conformational and configurational nature of

the various single and double bonds of the retinyl moiety as well as the C=N portion of

rhodopsin. Raman data indicated an anti geometry for the C=N in both rhodopsin and

33

isorhodopsin (also bacteriorhodopsin) (Paling et al., 1987; Bagley et al., 1985). Similar

studies have provided strong evidences that the CIO-C l l' C12-C13 and C14-C15 single bonds

are s-trans (Paling et aI., 1987). The same situation was observed for isorhodopsin and

bacteriorhodopsin (Paling et aI., 1989).

NMR data of 13C-labeled analogs indicated a twist of the C6-~ bond, induced by

the 5-methyl group on the ring (Mollevanger et al., 1987). The conformation of C6-C7

was not perturbed by the protein. CD studies by using ring-locked rhodopsin analogs,

have shown that twists induced by the protein on C6-~ and C\2-Cl3 correspond to the 0.­

and 13-bands of the CD spectrum (Balogh-Nair & Nakanishi, 1990; Ito et al., 1990).

A twisted ring-chain conformation in the retinyl chromophore was found to be

necessary for facile pigment formation based on studies of phenyl analogs (Matsumoto

et al., 1980). It has been shown that the phenyl and o-tolyl analogs gave low pigment

yields, while the bulkier mesityl analogs formed with high yield, which is not compatible

with a simple steric model. Thus, ring-chain planarity, as a result of a lack of steric

effects in the phenyl and tolyl derivatives, probably causes the unfavorable pigment

formation. Similar difficulties probably account for the low yields of pigments from the

5-demethyl and l,l-didemethyl analogs (Kropf et al., 1973; Kropf, 1976).

1.7 Goal of this dissertation

In order to understand the molecular basis of the function of rhodopsin in the

visual tranduction process, it is necessary to know the structure of the chromophore and

its interaction with the protein. Due to the lack of high quality crystallographic data of

34

rhodopsin, the 3-dimensional structure of the chromophore binding site has not been

established. Spectroscopic techniques using synthetic rhodopsin analogs have been an

important tool to reveal the structural information and interactions between the

chromophore and the protein.

The primal)' goal of this dissertation is to contribute to current understanding of

properties of the visual chromophore by studying the vibrational spectroscopic properties

(Chapter 2), chemical properties of retinal isomers (Chapter 3) and the photobleaching

mechanism of two dicis-isomeric rhodopsin analogs (Chapter 4).

In chapter 2, characteristic trends of all sixteen isomers of retinal, including C=C

double bond stretching modes, C-C single bond stretching modes, and hydrogen out of

plane bends (HOOP), will be discussed. In the case of the 7-cis and 7,9-dicis isomers,

FTIR studies have been extended to their Schiff bases and protonated Schiff bases using

isotopically labelled analogs. This vibrational information provides a probe for studying

specific changes in the chromophore-opsin interaction during the photobleaching process.

In order to understand the mechanism of photobleaching of rhodopsin, the

isomerization reaction of retinal isomers in organic solvent has been studied. In chapter

3, the temperature dependence of the photoisomerization of several retinal isomers has

been examined. Also the mechanism of the thermal reaction of four labile hindered

retinal isomers has been studied.

Photobleaching studies have been carried out on two dicis rhodopsin analogs at

low temperature to reveal the mechanism of photochemistry (Chapter 4). Through low

temperature photochemistry and UV/Vis spectroscopy, as well as chromophore extraction

35

experiments, the isomerization mechanisms of 7,9-dicis-rhodopsin and 9, ll-dicis-12­

fluororhodopsin have been established. The difference in the photochemical properties

of these two dicis pigment analogs was rationalized by possible local protein

perturbations.

36

Chapter 2

FTIR Studies of Retinal Isomers, Their Schiff Bases and Their

Protonated Schiff Bases

2.1 Introduction

2.1.1 Previous studies

Detailed information about the chromophore structure and the chromophore-protein

interaction is in principle obtainable from vibrational spectroscopy including Raman and

infrared data as mentioned in section 1.5. Vibrational analysis of retinal isomers provides

a framework for the analysis of the pigment spectra (Curry et al., 1984; Saito et al., 1983;

Curry et al., 1982). The first attempt to assign the retinal spectra was performed by

Rimai (Rimai et al., 1973), and their assignments were largely supported by later studies

(Cookingham et al., 1978). A major step forward was made by Curry et al. (Curry et al.,

1982), who used an extensive series of isotopic derivatives to assign the vibrational

features of all-trans-retinal to specific normal modes.

The vibrational spectra of retinal, its Schiff base (SB), and its protonated Schiff

base (PSB) have been extensively utilized as models for elucidation of the molecular

configuration of the chromophore in the photoreaction intermediates of rhodopsin and

bacteriorhodopsin (Mathies et al., 1987). The vibrational spectra of rhodopsin,

isorhodopsin, metarhodopsin I, and metarhodopsin II have been compared with those of

37

ll-cis-PSB, 9-cis-PSB, all-trans-PSB, and all-trans-SB, respectively (Smith et at., 1985).

The vibrational spectra of the bacteriorhodopins as well as its intermediates have been

compared with those of all-trans-PSB, 13-cis-PSB, and 13-cis-SB (Baasove et al., 1987;

Gilson et al., 1988). Thus, vibrational studies of isomeric rhodopsin chromophores have

been limited to those of the natural occurring system: all-trans, l l-cis and the 9-cis, 13­

cis, and 9,13-dicis isomers. The configurations of the chromophores of other isomeric

rhodopsins, e.g. 7-cis and 7,9-dicis isomers, have not yet been established, although stable

pigments have been prepared (deGrip et at 1973; Liu et al., 1984a). Thus, the vibrational

spectra of 7-cis and 7,9-dicis isomers of retinal, their Schiff bases and their protonated

Schiff bases may provide additional information on these new isomeric chromophores

during their bleaching processes. Especially important is the expected effect of the larger

ring-chain dihedral angles of these compounds .(Liu et al., 1983b) upon the rates of

interconversion of the later bleaching intermediates (Shichida et al., 1991).

The goal of the project in this chapter is to obtain the vibrational information for

model compounds containing the 7-cisor 7,9-dicis geometry, using known retinal isomers,

their Schiff bases and protonated Schiff bases supplemented by several isotopically

labelled analogs. The vibrational assignments of these two isomers can be further

substantiated through normal mode calculations. This vibrational information will be

reserved for future studies of specific changes in the chromophore-opsin interaction during

the photobleaching process of the isomeric rhodopsins.

In fact, this effort has been broadened to assign key vibrations of all sixteen retinal

isomers for a complete correlation of key signals which result from geometric and

38

electronic perturbations. The emphasis has been placed on the fingerprint region of C-C

stretches and hydrogen out of plane bends (HOOP band). Comparison of observed

frequencies, intensities, and isotopic shifts of the aldehydes with those of the visual

pigments has pointed to systematic discrepancies which can be attributed to protein

perturbations (Eyring et al., 1982).

2.1.2 Raman and infrared spectroscopy

When a molecule having a highly polar group is irradiated by infrared light, it

absorbs light and changes its dipole moment resulting in an infrared spectrum. In Raman

spectroscopy, we do not observe transmitted light but light scattered by the sample. If

the molecule is symmetrically substituted and does not change its dipole moment, the

Raman scattering is observed. Because Raman spectroscopy usually uses a strong laser

beam, application of this method to rhodopsin requires appropriate precautions to

minimize photodecomposition of rhodopsin and avoid measurement of false Raman bands

due to photoproducts. This problem can be overcome by the molecular flow resonance

Raman technique (Mathies et al., 1976; Callender et al., 1976), in which a rhodopsin

sample is circulated through the sample chamber so that each rhodopsin molecule is

exposed to light for less than 10 usee. IR is an independent technique for looking at

chromophore vibrations with the advantage that the energy of the IR photon is too small

to disturb the rhodopsin system. More importantly, it reveals changes in the chromophore

and protein which are not detected by RR (Nakanishi, 1987).

39

In retinal, n to n" excitation shortens the single bonds and lengthens the double

bonds of the conjugated chain. Thus only the stretching vibrations of the chain carbon­

carbon bonds carry significant intrinsic RR intensity, and the observed intensity of most

of the normal modes between 500 and 1700 ern" depends on their stretching character.

Stretches near the center of the chain have the greatest intensity because this is where the

excitation-induced bond-order changes are largest. Significant intrinsic infrared amplitude

is expected for C=C and C-C stretches, as well as for the C=O stretch, and the out-of­

plane wags of the vinyl protons. This is presumably due to parallel charge polarization

of all the bonds, imposed by the electronegative end group. This is consistent with the

observation that the intrinsic infrared intensities are greatest for stretches near the

aldehyde group rather than for those near the center of the chain.

2.1.3 Vibrational structure of retinal

Retinal has four double bonds in the isoprenoid chain. Since each double bond

can be either cis or trans configuration, a total of 16 cis/trans isomers are possible, as

shown in Figure 1.11. The synthesis of all these isomers were successfully completed

recently (Trehan et al., 1990b).

The electrons of the isoprenoid chain are delocalized over the entire polyene

system stabilized by resonance. Therefore, the single bonds acquire some double bond

character and the double bonds become somewhat closer to single bonds. In the absence

of steric hindrance, the n electron system assumes a planar form but becomes twisted

around a single bond where steric hindrance exists (Mathies et al., 1989).

40

Retinal consists of a ~-cyclohexenyl ring, a methylated polyene chain made up of

repeating isoprenoid units and a conjugated aldehyde group. IR intensity of the ring

vibrations is weak and also generally insensitive to the chain configuration with the end

group modifications. The most diagnostic information about configuration is from the

chain vibrations in the 500-1700 cm' range, which include CC stretching vibrations, in­

plane and out-of plane hydrogen bends, and most methyl group vibrations (Curry et al.,

1985).

The most useful way of looking at retinal vibrations is in terms of individual

localized basis coordinates, with specific interactions among them. These local symmetry

coordinates are linear combinations of the geometrically internal coordinates and are

referred to as C=C, C-C, and C-CH3 stretches, CCH vinyl or methyl rocks, and C-H or

C-CH3 out of plane wags.

Each local symmetry coordinate (LSC) is considered to have an "intrinsic "

frequency. Potential and kinetic coupling among a small number of such coordinates

produces normal modes with frequencies that are shifted from the intrinsic frequencies

of their constituents. These normal modes take on some of the character of each of the

constituent basis coordinates (Curry et ai., 1985). In general, strongly coupled

coordinates will also be strongly mixed. When coordinates are coupled, the resulting

normal modes have a significant contribution from both coordinates and these normal

modes appear shifted in frequency from the intrinsic frequency.

The patterns of coupling and mixing of basis coordinates in the normal modes are

most easily investigated by isotopic substitution. The substitution alters the reduced mass

41

of three or more LSC's and may also change the kinetic coupling between LSC's. If the

perturbation is small, such as a 13C_ substitution, the normal modes are left unchanged and

only their reduced mass and thus their frequencies change (Curry et al., 1985). When

deuterium is substituted for hydrogen, the resulting perturbation is much larger. The

intrinsic wavenumbers of the substituted LSC's and their coupling with other coordinates

are altered so strongly that it is most useful to think of these interactions as being entirely

removed (Curry et al., 1985).

In summary, the isotopic frequency shifts observed in the retinal derivatives enable

us to understand the patterns of coupling among the basis coordinates, and the intensity

patterns in the derivatives can be explained by variations in the mixing of basis

coordinates in the normal modes. Such an understanding of the molecular basis for the

observed frequencies and intensity patterns provides a useful physical model for the

normal modes of retinal chromophores.

42

---~---

2.2 Materials and methods

2.2.1 Materials

2.2.1.1 Sixteen retinal isomers

All-trans and 13-cis isomers of retinal were purchased from Sigma, and 9-cis­

retinal from Eastman Kodak. 11-Cis-retinal was provided as a gift sample from

Hoffmann-La Roche. 7-Cis, 7,9-dicis, and 7, l l-dicis-retinal, obtained as by products from

synthesis of poly-cis isomers of retinal, were provided by Dr. A. Trehan. 9,11-Dicis,

obtained from the photostationary state mixture resulted from irradiation of all-trans­

retinal in acetonitrile at >340 nm (Coming 0-52 filter), was provided by Dr. S.

Ganapathy. 7,9,11-Tricis-retinal was obtained from the DIBAH reduction of 7,9,11-tricis

retinonitrile provided by Dr. A. Trehan. 11,13-Dicis, 7,11, 13-tricis, 9,11,13-tricis or all­

cis-retinal was obtained from the corresponding quasi photostationary state mixture

resulted from direct irradiation of l l-cis, 7,II-dicis, 9,11-dicis or 7,9,II-tricis-retinals,

respectively in hexane at >340 nm (CS 0-52 filter) at OOC. The yields were 25% for

1l,13-dicis, 24% for 7,1l,13-tricis, 26% for 9,1l,13-tricis and 42% for all-cis-retinal.

7,13-Dicis, 9,13-dicis and 7,9,13-tricis were obtained from thermal isomerization of

7,11,13-tricis, 9,11,13-tricis and all-cis-retinals in hexane at 400C for 10-12 hours. All

compounds were purified by preparative HPLC before taking FTIR spectra. The four

thermally unstable isomers (11, 13-dicis, 7,11, 13-tricis, 9,11, 13-tricis and all-cis) were

collected at 2°C.

43

2.2.1.2 Synthesis of deuterioretinals

7-Deuterioretinal: A mixture of methyl geranate and nerate was reduced with

lithium aluminum deuteride. Oxidation with active Mn02 afforded I-deuteriocitral which

was condensed with acetone in the presence of KOH to provide an isomeric mixture of

pseudoionone. Ring closure by acid catalyzed cyclization gave a mixture of 7-deuterio-rx­

and 13-ionones from which the requisite 13-isomer was isolated by preparative HPLC. The

above synthesis steps were done by Dr. A. Asato. Then, the subsequent C2-chain

extension, selective sensitization, and Cs-chain extension were accomplished, based on

established procedures for retinal synthesis (Liu & Asato, 1982). HRMS for 7-cis:

285.218, 7,9-dicis: 285.219 (Calculated data for C2oH270D =285.220). For both isomers,

IH-NMR (CDCI3) signals were identical to those of 7-cis-retinal (Zhu et al., 1990)

excepting the absence of the H-7 doublet and the appearance of a singlet for H-8. The

product mixture was separated by HPLC and was found to contain 7,13-dicis, 9-cis, 7,9­

dicis, 7-cis and all-trans isomers.

19,19,19-Trideuterioretinal: The base-catalyzed (NaOD in D20-pyridine

deuterium exchange of 13-ionone, effected twice to afford trideuterio-ji-ionone of isotopic

purity in excess of 95% (Fransen et al., 1980), was done by Dr. A. Asato. Thereafter,

chain elaboration followed the same procedures mentioned above (Liu & Asato, 1982)

giving the desired trideuterio-retinal. All isomers exhibited expected spectral data.

HRMS for 7-cis: 287.232; 7,9-dicis: 287.233 (Calculated data for C2oH250D3 =287.233).

44

8,19,19,19-Tetradeuterioretinal: Condensation of l3-cyclocitral with

perdeuterated acetone provided tetradeuterated ~-ionone. Following the same sequence

of reaction (Liu & Asato, 1982), tetradeuterioretinal was obtained. All isomers exhibited

expected spectral data. HRMS for 7-cis: 288.238; 7,9-dicis: 288.237 (Calculated data for

Ci a ce~Me b ceO'OH• ..

c ceOOd ~o e... • ..

CXUof ..

1

~ eNIrradiation..sensitizer

7-cis and 7,9-dicis-l g ..~ CHO

hII

g .. ~ CHO

a = (CH3CH20hPOCH2C02CH3/NaOCHy'CH30Hb = LiAID4, C = Mn02, d =CH 3COCHy'KOH, e =H2SO/HOAcf =(CH3CH20hPOCH2CN/n-BuLi/THF, g = DIBAHh = (CH3CH20hPOCHiCH3)C=CHCN/LDA/THF

Figure 2.1 A scheme for the synthesis of 7-deuterioretinal.

45

a .. b ...

CD3

"- CN Irradiation'-cis and ',9-dicis-l c

sensitiz;;'...

1

CD3 CD3

"- '- "~ CN

CHO d ..

c ...

a = Py/D20/NaOD, b = (CH3CH20hPOCH2CN/n-BuLiffHFc = DIBAH, d = (CH3CH20hPOCH2(CH3)C=CHCN/LDA/THF

Figure 2.2 A scheme for the synthesis of 19,19,19-trideuterioretinal.

46

a o b

1

Irradiation. . . 7-cis and 7,9-dicis-l

~O

d c

~ CHO

a = CD3COCD/NaOD, b = (CH3CH20hPOCH2CN/n-BuLi/THFc = DIBAH, d = (CH3CH20hPOCH2(CH3)C=CHCN/LDAITHF

Figure 2.3 A scheme for synthesis of 8,19,19, 19-tetradeuterioretinal.

47

2.2.1.3 Preparation of 7·cis· and 7,9·dicis.retinylidene SB and PSB

The general procedure to prepare retinylidene SB and PSB are same as those

reported (Colmenares et al., 1992) with a slight modification, as described below:

At about aoc, to a 10 ml flask containing about 1 mg of a freshly purified retinal

or deuterioretinal analog, 3 ml dry hexane containing 10 ul of n-butylamine were added

along with 2-3 pieces of molecular sieves (3 A). After reaction was kept for about 10 h

at aoc, an aliquot was taken to check the completion of the reaction by UV-Vis

spectrophotometry. The reaction solution was filtered through a cotton plug in a small

pipet, then evaporated under vacuo. Addition of cyclohexane facilitated the removal of

excess n-butylamine. After further evacuation using a vacuum pump, the retinylidene SB

was redissolved into a small amount of dry hexane for FfIR study. Protonation was

accomplished by addition of hexane saturated with HCI gas.

UV- VIS data for 7-cis SB in methanol: Amax =356 nm; 7,9-dicis SB: Amax = 347

nm. 7-cis PSB: Amax =436 nm; 7,9-dicis PSB: Amax = 422 nm.

2.2.2 Methods

2.2.2.1 FTIR spectroscopy

FrIR spectra were recorded on a Nicolet-740 spectrometer and ratioed against a

clean window. A freshly prepared sample in a hexane solution was deposited dropwise

on a KBr plate followed by evaporation of the hexane by a stream of argon. One

hundred scans of interferograms were accumulated, then Fourier transformed to yield a

48

spectral resolution of better than I cm'. The data were smoothed using a three-point

sliding average and the peak positions are accurate to 2 ern". For the aldehyde

compounds, after completion of recording each spectrum, the sample was redissolved in

hexane and analyzed by HPLC to assess the extent of sample degradation from

photochemical or thermal isomerization (the latter being particularly worrisome for the

less stable 11,13-dicis isomers). In no instance were rearranged products found to

constitute more than 5% of the sample. For the retinylidene SB and PSB compounds,

special care was taken to keep the compounds at low temperature and to minimize

manipulation time. Absence of any significant irreproducible results in the spectrum

during the successive scans was the criteria for a satisfactory spectrum.

2.2.2.2 Normal mode calculations

Normal mode calculations were performed using the FG matrix method of Wilson,

Decius and Cross (Wilson et al., 1955). The program was obtained from Quantum

Chemistry Program Exchange in Indiana University. This package of four Fortran

programs has been designed to allow the user a simple and reliable method for generating

a complete vibrational analysis. The geometries of retinal and the protonated Schiff base

were calculated using PCMODEL (molecular modelling software from Indiana

University). All geometric parameters (bond angles, bond lengths, and dihedral angles)

were minimized. The n-butyl group of the protonated Schiff base was replaced by an R

group, where R is a dump atom having mass 15. Carbons 1,4, and 18 of the ring were

also replaced by R atoms. The initial force constants for retinal and the protonated Schiff

49

base were set to be the same as the most recent version used for all-trans retinal (Curry

et al., 1985) and the all-trans protonated Schiff base (Smith et al., 1985). The force

constants for the conjugated chain were then refined to fit the observed frequencies and

frequency shifts of the 7-cis or 7,9-dicis-retinal and protonated Schiff base derivatives.

50

2.3 Results

2.3.1 FfIR spectra of the sixteen retinal isomers

We have recorded the FfIR spectra of all sixteen isomers of retinal. Their

fingerprint region, 1700-600 em:' are shown in Figure 2.4. These isomers of retinal

provide an excellent example of the effect of double bond geometry on vibrational

characteristics (Mathies et al., 1985). The expanded region 1250 cm-l-lOOO cm', shown

in Figure 2.5, is highly sensitive to deviation of the polyene geometry. Complete

assignments of the vibrational spectra of five retinal isomers (all-trans, 13-cis, l l-cis, 9­

cis and 9,13-dicis) have been achieved by use of an extensive 2H and l3C-isotopic

derivatives (Curry et al., 1982; 1984; Mathies et al., 1989). It was shown (Curry et al.,

1984) that normal modes are fairly localized C-Cstretches but that they also contain

significant in-plane C-H rocking character. The localization of the C-C stretching

character makes the description of the normal modes as individual C-C stretches

reasonably accurate.

Analyses of the vibrational spectra of retinal isomers revealed that the major

effects of isomerization on the vibrational spectra of the C-C stretching region are

restricted to normal modes localized on these carbon atoms forming and immediately

adjacent to the cis bond. The normal modes which are not immediately adjacent are

essentially unaffected by the cis band (Curry et al., 1985). The penurbations caused by

these isomerizations are vinually independent of each other. The spectral differences of

poly-cis isomers exhibited the same features as the sum of the differences exhibited by

51

the mono-cis isomers (Mathies et al., 1989). Therefore, the assignments of the observed

bands of the poly-cis isomers of retinal were made by comparison with the spectra of the

all-trans and mono-cis isomers (to be elaborated in the discussion section). A summary

of the assignments of the C-C stretching modes are listed in Table 2.1.

52

Table 2.1 The C-C stretching frequencies in the region of 1250-1000 ern" of sixteen

retinal isomers.

ISOMERS C14-CI 5 CI2-C13 CIO-C11 Cg-C9

all trans• 1111 1215 1163 1197

13-cis' 1115 1222 1162 1191

l l-cis' 1126 1204 1084 1214

9-cis' 1113 1216 1148 1200

9, 13-dicis' 1115 1224 1145 1194

7-cis 1112 1211 1154 1188

7,9-dicis 1113 1213 1148 1189

7,11-dicis 1128 1204 1105 1186

7,13-dicis 1122 1212 1156 1186

9,11-dicis 1124 1202 1088 1187

7,9,11-tricis 1125 1204 1104 1169

7,9, 13-tricis 1115 1216..

1188w

1172

11,13-dicis 1126 1202..

1171w

7, 11,13-tricis 1122 1212 1156 1186

9,11,13-tricis 1128.. •• 1171w w

all-cis 1127..

1095 1170w

• assignments are from literature (Mathies et al., 1989; Curry et al., 1985).•• weak signals or degenerate with others.

53

All-trans 9,11-dicis

I3-cis

9-cis

II-cis

9,13-dicis

7,9,13-tricis

7, 9- dicis 7,11,13-tricis

7,11-dicis

7,13-dicis

500800

9,11,13-tricis

wavenumber

I

1400500 1700i i

1100 800wavenumber

I

1400I

1700

Figure 2.4 Fr-IR spectra of all sixteen isomers of retinal in the fingerprint region of

1700-500 cm', obtained after depositing each sample on a KBr plate.

54

(11 (5) (91 (13)

~~.~~(2) (6) ( 10) (14)

~~~~

~~~~VIVI

(3)

(4)

(7)

(6)

(11)

( 12)

(15 )

(16)

~~~~• I I I I , • I I I I I I I Iii i I I i I I I

1250 1000 1250 1000 1250 1000 1250 1000wavenumber wavenumber wavenumber wavenumber

Figure 2.5 Expanded Fl'-IR spectra of all sixteen isomers of retinal in the region of 1250-1000 cm': (1) all-trans, (2) 13-cis,(3) 9-cis, (4) l l-cis, (5) 7-cis, (6) 7,9-dicis, (7) 7,11-dicis, (8) 7,13-dicis, (9) 9,11-dicis, (10) 9,13-dicis, (II) 7,9,11­tricis, (12) 7,9,13-tricis, (13) 11,13-dicis, (14) 7,11,13-tricis, (15) 9,ll,13-tricis, (16) all-cis.

The out-of-plane vibrations of the retinal chain can be classified as CCCC torsions

or as wags of the hydrogen substituents and methyl groups. The CCCC torsional modes

and the out of plane wags of the methyl carbons appear below 500 em'), in a region of

the spectrum masked by KBr absorption in the IR. These modes, therefore are not

observed and will not be discussed.

Hydrogen out-of-plane vibrations of linear polyenes are observed between looo

and 700 em:'. The 1000-900 em') and 800-700 em') regions of FfIR spectra of the

sixteen isomers of retinal are presented in Figure 2.6. The most striking feature in this

region of the all-trans-retinal FfIR spectrum (Figure. 2.6(1)) is the intense line at 966 em'

l, which corresponds to a well-known IR group frequency associated with the in-phase

(Au in the local ~h point group) HOOP vibration of trans ethylenic protons (Curry et al.,

1985). There are two such groups in retinal: H~=C8H and HCl1=Cl2H. According to

the shifts observed upon deuteration, the 967 and 955 em" peaks have been, assigned to

be the H~=C8H and HCll=C12H Au HOOP bands respectively (Curry et al., 1982). This

is a characteristic frequency for the in-phase wag combination of trans ethylenic hydrogen

(Curry et al., 1985).

The HOOP bands of 13-cis-retinal, 9-cis retinal and 9, l3-dicis-retinal (Curry et al.,

1985) are observed in the same region.

11-Cis and 7-cis retinal HOOP bands are very different from the above isomers.

The 761 ern" infrared band of l l-cis retinal can be assigned to the cis HCl1=C I2H HOOP

mode. Similarly the 743 ern" infrared band of 7-cis retinal can be assigned to the cis

H~=C8H HOOP mode.

56

The remaining out-of-plane wags can be assigned to bands appearing close to their

values observed for the all-trans isomer. Thus the 966 cm' infrared band of the l l-cis

isomer is assigned to the H~=C8H trans HOOP mode and the 961 crn' infrared band of

7-cis retinal is assigned to the HCl1=C12H trans HOOP mode.

Other isomers which contain either 7-cis (i.e. 7,9-dicis, 7,13-dicis, 7,9,13-tricis)

or II-cis (i.e, 9,Il-dicis, I I, 13-dicis, 9,II,I3-tricis) geometries show single peak at -740

or -760 ern", characteristic of the HOOP bands of the 7-cis or II-cis double bonds. The

remaining HOOP bands fall in the 900-1000 cm' region expected for the l l-trans bonds

of the 7-cis isomers or 7-trans bands of the l l-cis isomers.

The isomers which contain both 7-cis and l l-cis geometries (i.e. 7, ll-dicis,

7,11,13-tricis, 7,9,1I-tricis, and all-cis) are expected to exhibit weak absorption in the

region of 900-1000 cm' because no trans HOOP bands exist in such isomers. In the

meantime, the cis HOOP region contain two modes at -760 and -740 em" which are

associated with the cis HCl1=C12H and the cis H~=C8H respectively. The experimental

data which are summarized in Table 2.2 are consistent with these expected features.

57

(9)(5)(1 )(13)

~~~~(2) (6) (10) (14)

~~--A-~~(3) (7 ) (11) (15)

VI00 ~~~~

(4) (6)(12) ( 16)

~~~~I I I --, I I -.--~. I I I .--------1------------.-- •

1000 700 1000 700 1000 700 1000 700wavenumber wavenumber wavenumber wavenumber

Figure 2.6 Expanded FT-IR spectra of all sixteen isomers of retinal in the region of 1000-700 cm': (I) all-trans, (2) 13-cis,(3) 9-eis, (4) l l-cis, (5) 7-eis, (6) 7,9-dicis, (7) 7,II-dicis, (8) 7,I3-dicis, (9) 9,II-dicis, (10) 9,I3-dicis, (11) 7,9,11­trieis, (12) 7,9,13-trieis, (13) II,I3-dicis, (14) 7,1 1,13-tricis, (15) 9,11,13-tricis, (16) all-cis.

Table 2.2 The HOOP band region, 1000-700 ern", of the sixteen retinal isomers.

Isomers 11+12 HOOP 7+8 HOOP 11+12 HOOP 7+8 HOOP

trans trans cis cis

all trans 967 967 - -

13-cis 954 966 757(w) -

l l-cis - 965 761 -

9-cis 962 962 760(w) -

9,13-dicis 962 962 w -

7-cis 961 - - 743

7,9-dicis 961 - - 746

7,II-dicis - - 758 740

7,13-dicis 951 - - 741

9,II-dicis - 961 758 -

7,9,II-tric - - 760(w) 743

7,9,13-tric 957 - 760(w) 746

Il,13-dicis - 966 761 -

7,11,13-tric - - 760 740

9,11,13-tric - 964 759 -

all-cis - - 765 743

59

2.3.2 FfIR spectra of 7·cis- and 7,9·dicis·retinal and their deuterio-analogs

In section 2.3.1, assignments of major signals in the FTIR spectra of all sixteen

retinal isomers were made. These assignments rely heavily on information gleaned from

the assignments of the all-trans and the common mono-cis isomers. In this section, the

FTIR spectra of three deuterioretinals (7-0, 19,19,19-03, and 8,19,19, 19-D4) are described.

These data will be used to confirm the assignments for the 7-cis and 7,9-dicis retinal

isomers in the C-C stretching region.

The 1700-600 ern" region of the FTIR spectra of 7-cis and 7,9-dicis retinal and

their labelled analogs are shown in Figure 2.7.

In Figure 2.8 are the expanded fingerprint regions from 1250 to 1000 ern" of 7-cis ­

and 7,9-dicis retinal and their deuterated analogs. The peaks of the 7,9-dicis retinal at

1113, 1213 and i 148 cm' can be firmly assigned to CI4-CIS' CI2-C13 and CIO-Cll because

they are consistently close to the corresponding modes of 9-cis-retinal (Curry et al.,

1985). Two peaks remain unassigned (1189 and 1171 ern") which are most likely due

to C6-~ and Cg-C9 stretching. The 1189 cm' band is sensitive to deuteration shifting to

1204 ern" (Figure 2.8) in 7-D-7,9-dicis-retinal, to 1195 in 9-CD3-7,9-dicis-retinal, and to

1156 em:' in 8-D-9-CD3-7,9-dicis-retinal. On the other hand, the 1171 em:' peak is not

affected by the isotopic labels at similar positions. This pattern of behavior upon isotopic

substitution suggests that the 1189 cm' band is more consistent with Cg-~ stretching.

Similar isotopic shifts are observed in 7-cis-retinal in which the 1189 cm' band shifts to

1203 crn' in 7-D-7-cis-retinal, 1193 cm' in 9-C03-7-cis-retinal, and 1156 em'! in 8-0-9­

CD3-7-cis-retinal. In Figure 2.9 are correlation diagrams for the isotopic shifts of 7-cis-

60

and 7,9-dicis-retinal. Table 2.3 summarizes the assignments made for the single bond

stretching modes for 7-cis and 7,9-dicis-retinal.

Table 2.3 C-C stretching frequencies of 7-cis and 7,9-dicis-retinal and their deuterio­

analogs in the fingerprint region of 1250-1000 cm', The number in parentheses are

the shifts from the parent compound.

Isomers C14-C15 C12-C13 CIO-Cl1 Cg-C9

7-Cis- 1112 1211 1154 1189

7-0- 1120(+8) 1211(0) 1160(+6) 1203(+14)

9-CD3- 1117(+5) 1217(+6) 1148(-6) 1193(+4)

8-0-9-C03- 1127(+15) 1217(+6) 1156(+2) 1144(-49)

8-F 1111(-1) 1211(0) 1159(+5) 1140(-49)

7,9-Dicis- 1113 1213 1148 1189

7-D- 1113(0) 1215(+2) 1158(+10) 1204(+15)

9-CD3- 1114(+1) 1213(0) 1155(+7) 1195(+6)

8-0-9-CD3- 1112(-1) 1213(0) 1148(0) 1156(-33)

61

0­N

7-cis retinal

7-0-

9-CO,-

7,9-dicis retinal

7-0-

9-CO,-

8-0-9-CO,-

_ L1~~80--1~60 -10-;;-0-820---600 1-70Jo -- -lTeo - -125J - l(')i±O.--- -t§"2J-~---'---~GO

WAVENUMSER WAVE~UMa~R

Figure 2.7 FTIR spectra (1700-600 cm') of 7-cis and 7,9-dicis-retinal and their labelled analogs.

7 -cis- 7,9-dicis-

7·0

1250 1120WAVENUMBER

1250 1120 990WAVENUMBER

Figure 2.8 Expanded (1250-990 cm') FTIR spectra of 7-cis and 7,9-dicis-retinal and

their deuterated analogs.

63

7-cis-retinal

1211 1211 1217 1217- _-- ......._._-

1188 ....·····1203 ···· 1194.

1154 ~.i-:: 1160

<,.1 148 ~~.:::\l156

1144

1120 ..... 1117 .....···11271112.- .. -» .. ---- "._-'

7-D

7,9-dicis-retinal

1215

1189 ··'i204···· 1195 .

-' 1158 1155 '\"'\,11561148 -_ _- '----_..- .._-

1148

1113 1113 1114 _11_1_2

7-D

Figure 2.9 Correlation diagrams of isotopic shift of 7-cis and 7,9-dicis-retinal.

64

2.3.3 FTIR spectra of 7-cis- and 7,9-dicis-retinylidene SB and their deuterio-analogs

The 1800-600 cm' region of the FTIR spectra of 7-cis and 7,9-dicis-retinylidene

SB and their deuterated analogs are shown in Figure 2.10.

The vibrational structure of the retinyl Schiff base is significantly altered at the

substituted end of the chromophore. The oxygen atom is replaced by a less

electronegative nitrogen, and an n-butyl group is added, which changes the C-C stretches

from those of the aldehyde (Mathies et al., 1987).

The C-C stretching character of all-trans retinylidene SB in the fingerprint modes

has been assigned using isotopic substitution (Smith et al., 1985). Conversion of all­

trans-retinal to its Schiff base causes a-50 ern" increase in the C14-C15 stretching

frequency, placing it nearly degenerate with the CIO-CII stretch at 1163 em", These two

internal coordinates then interact strongly, forming two nearly equally mixed normal

modes at 1175 and 1153 em:'. The Cg-C9 normal mode is unchanged in frequency, while

the C12-C13 mode moves up 6 cm' as a consequence of increased interaction with the

nearby C14-C15 stretch.

The fingerprint region, 1300-1100 cm', of 7,9-dicis-retinylidene SB and its

deuterioanalogs are shown in Figure 2.11. The frequency assignments of these

compounds, based on the trends noted in the all-trans isomers, are listed in Table 2.4.

The 1222 em:' band of 7-cis-retinylidene SB with its frequency and intensity similar to

the corresponding band in all-trans-retinylidene SB, was assigned to the CI2-C13 stretch.

The C14-C15 and ClO-CIl stretches are degenerate at 1167 em" in 7-cis retinylidene SB.

The 1189 cm' band of 7-cis-SB which is downshifted from 1196 crn' of all-trans-SB was

65

assigned to Cg-C9• From Table 2.4 we can see the isotopic shifts for these retinylidene

SB isomers are relatively small. A similar observation was applied to the 7,9-dieis-

retinylidene SB (see Table 2.4).

Table 2.4 C-C stretching frequencies of retinylidene SB and their deuterio-analog in

the fingerprint region of 1300-1100 ern"

Isomers C14-C15 C IZ-CI 3 CIO-CI I Cg-C9

(ern") (crn') (cm') (ern")

'-Cis- 1167 1218 1167 1189

7-D- 1164 1220 1164 1187

9-CD3- 1164 1219 1164 1187

8-D-9-CD3- 1166 1219 1166 1184

',9-Dicis- 1172 1218 1163 1191

7-D- 1171 1220 1171 1190

9-CD3- 1172 1218 1166 1191

8-D-9-CD3- 1167 1218 1167 1190

66

7-cis Schiff base 7,9-dicis Schiff base

8-0-9-CDJ -

7-0-

"'J,'N,~

7-0-

8-0-9-COJ -

1800 ! 600 --Ifoo---- 1200 - 1600---800~600I eoo---i60·o---1TOo-- .-1200---jocio--eoo--sooWAVF":-:IIMR;:-1=l WAV=NIIMAFR

Figure 2.10 FTIR spectra (1800-600 crn') of 7-cis and 7,9-dicis-retinylidene SB and their deuterated analogs.

~

9-cis-

7,9-dicis-

7-D-

Figure 2.11 Expanded FTIR spectra of fingerprint region of 1300-1100 crn' of 7,9-dicis­

retiny1idene SB and its deuterated analogs.

68

2.3.4 FTIR spectra of7-cis and 7,9-dicis-retinylidene PSB and their deuterio-analogs

The FTIR spectra of 7-cis- and 7,9-dicis retinylidene PSB and their deuterio­

analogs in the 1800-600 ern:' region are shown in Figure 2.12.

The dominant effect of Schiff base protonation is a significant increase in the

delocalization of the n-electrons as evidenced by the shift of the absorption maximum

from 360 to 440 nm. The obvious expectation is that the single bonds would increase in

frequency as a result of reduced bond-order alternation.

The fingerprint region, 1300-1000 cm', of 7,9-dicis retinylidene PSB and its

deuterioanalogs is shown in Figure 2.13. In order to compare with 9-cis retinylidene

PSB, its FrIR spectrum of 9-cis retinylidene PSB is also shown. The single-bond

stretching modes in 7,9-dicis-retinylidene PSB (see Figure 2.13) can be assigned on the

basis of assignments previously made for 9-cis-retinylidene PSB and 9-cis-rhodopsin. In

9-cis-retinylidene PSB, the CIO-CII , Cg-C9, C14-C15 and C12-C13 single bond stretches are

assigned to the 1139, 1194, 1206 and 1238 crn' Raman bands. In the FTIR spectrum

these bands are at 1136, 1190, 1202 and 1237 ern", slightly different from the Raman

bands (Palings et al., 1987). In 7,9-dicis-PSB, the major difference in the structure is the

change from a 7-trans to a 7-cis configuration. A 15-20 cm' downshift is expected in

those C-C stretches of bonds adjacent to a cis double bond (Curry et al., 1985). Thus,

in the 7,9-dicis isomer, one would expect most signals should be identical with those of

9-cis-PSB except that the Cg-C9 stretch should shift down about 15-20 ern" from that in

9-cis-retinylidene PSB. The 1140, 1196 and 1234 ern:' FTIR bands of 7,9-dicis­

retinylidene PSB exhibit frequencies and intensities similar to those of 9-cis-retinylidene

69

PSB and thus have been assigned to the CIO-C l l' C14-C15 and C1Z-C13 stretching modes

respectively. The remaining band at 1179 cm' is 15 cm' downshifted from a similar

band (1194 em:') in 9-cis-PSB, and is thus assigned to the Cg-C9 band.

Isotopic derivatives have been used to confirm the assignments of 7,9-dicis-PSB.

Upon deuteration, the 1179 band in 7,9-dicis retinylidene PSB shifts to 1207 cm' in 7-0­

7,9-dicis retinylidene PSB, to 1199 ern" in 9-C03-7,9-dicis PSB, and to 1191 cm' in 8-D­

9-C03-7,9-dicis PSB respectively. The isotopic shifts of 7-cis- and 7,9-dicis retinylidene­

PSB and their deuterated analogs are summarized in Table 2.5.

From Table 2.5 we notice that the isotopic shifts of 7-cis-retinylidene PSB upon

deuteration are relatively small. The reason for this may be due to insignificant coupling

between the C-C single bond and the C-H in plane rocking modes.

70

Table 2.5 C-C stretching frequencies of the 1300-1100 em" region of 7-cis- and 7,9­

dicis-retinylidene PSB and their deuterio-analogs. The number in parentheses are the

shifts from the parent compound.

Isomers C14-C15 C12-C13 CIO-ClI Cg-C9

7-Cis- 1195 1236 1157 1184

7-D- 1196(+1) 1237(+1) 1154(-3) 1180(-4)

9-CD3- 1196(+1) 1237(+1) 1153(-4) 1181(-3)

8-D-9-CD3- 1193(-2) 1237(+1) 1153(-4) 1180(-4)

7,9-Dicis· 1196 1234 1147 1179

7-D- 1184(-12) 1234(0) - 1207(+28)

9-CD3- 1181(-15) 1233(-1) - 1199(+20)

8-D-9-CD3- 1191(-5) 1233(-1) - 1191(+12)

71

7-cis PSB 7,9-dicis PSB

--.JIv

7-0-

8-0-9-COJ -

l§OQ ----'600 l80o'16Oo--iToo---1200----1OO0··- --'--soo ----600

WAVENUMBER

FfIR spectra (1800-600 cm') of 7-cis and 7,9-dicis-retinylidene PSB and their deuterated analogs.

1880- 1600 --i ~ DO -12OO-~1 DODWAVENUMOfR

Figure 2.12

9-cis-

7,9-dicis-

7-0-

1300 -----i"2oc)"---i--1 00

WAVENUMBERFigure 2.13 Fingerprint region of 1300-1000 ern" of FfIR spectra of 7,9-dicis-

retinylidene PSB and its deuterated analogs (9-cis isomer spectrum is shown for

comparison).

73

2.3.5 Normal mode calculation results

Normal mode calculations for 7-cis and 7,9-dicis-retinal and their protonated Schiff

bases have been performed to confirm the experimental data and the results are listed in

Tables 2.6 and 2.7. Normal mode analysis is very useful for studying the coupling

pattern for 7-cis and 7,9-dicis isomers.

The 1300-1100 ern" region of 7-cis, 7,9-dicis aldehyde, Schiff base (SB) and

protonated Schiff base (PSB) have been assigned based on isotopic shifts and

calculational results. These results of the model systems are expected to facilitate future

vibrational spectroscopic studies of the 7-cis and 7,9-dicis chromophores in visual pigment

analogs.

74

Table 2.6 Calculated retinal C-C normal modes and their assignments

obsd calc description

7-cis

1211 1215 0.16(12-13)-0.08(10-11)-0.06(8-9)+O.27(9-CH3)

+O.06(13-CH3)-0.46( 1OH)-0.28(11H)

1189 1186 0.18(8-9)+0.06( 10-11)-0.06( 12-13)-0.11 (9-CH3)

+0.42(7H)+0.15(8H)

1154 1158 0.16( 10-11 )-0.09(6-7)+O.08(8-9)+0.35( IIH)

+0. 16(lOH)

1112 1116 0.38(14-15)-0.11(12-13)-0.43(l5H)+0.21( lOH)

7,9-dicis

1213 1217 0.19( 12-13)+0.07 (8-9)-0.01(14-15)+0.31 (13-CH3)

+0.24(l5H)-0.19(lIH)+O.21(12H)

1189 1184 0.13(8-9)+0.07( 10-11)-0.08(7=8)-0.11 (C1S=O)-

0.12(9-CH3)+O.43( 15H)+0.14(7H)-0.34(8H)

1148 1147 0.29(10-11 )-0.15(13-CH3)-0.42(lIH) +O.l8(l2H)

1113 1118 0.11 (14-15)-0.12(13-CH3)+O.11(11=12)

+O.18(13=14)+0.50( 15H)+O.13(14H)

Coefficients (asfc)Q) of internal coordinates S in the normal modes Q. Symbols used: H,

in-plane hydrogen rock

75

Table 2.7 Calculated protonated Schiff base C-C normal modes and their assignments

obsd calc description

7·cis

1236 1239 0.20(12-13)-0.13(10-11 )-0.12(8-9)-0.14(13-CH3)

+O.28(14H)

1195 1191 0.18(8-9)+O.03(14-15)-0.09(9-CH3)+O.09(9=1 0)

+0.06(7=8)+0.41 (10H)-0.43(8H)-0.50(7H)

1184 1180 0.16(14-15)+O.12(12-13)-0.1O(10-1l)+0.02(15=N)-

0.51(12H)-0.33(14H)-0.24(15H)

1157 1152 0.27(10-11 )+0.09(8-9)-0.17(13-CH3)-0.44(11H)-

0.31(14H)-0.16(IOH)

7,9-dicis

1234 1239 0.14(12-13)-0.12(8-9)-O.12( 10-11)+O.21(13-CH3) -

0.11(11=12)-0.11 (12H)-0.12(14H)

1196 1193 0.16(14-15)+0.11 (12-13)+0.14(15=N)-0.14(13-CH3) -

0.17(8H)+O.14(14H)+O.21(11H)

1179 1174 0.11(8-9)-0.07(12-13)-0.22(9-CH3)-0.58( 10H)-

0.56(11H)+0.19(8H)-0.1O(7H)

1147 1143 0.16(10-11)+0.06(12-13)+0.07(11=12)+O.12(13-CH3)

+0.15(8H)+0.38(ISH)

Coefficients (CJS/CJQ) of internal coordinates S in the normal modes Q. Symbols used: H,

in-plane hydrogen rock

76

------

2.4 Discussion

2.4.1 C=O stretches

The C=O stretches in the spectra of all of the retinal isomers fall in a narrow

range of 1660-1670 ern" and are practically constant in intensity, thus not useful for

structural assignment. This agrees with the highly localized character of this normal

mode.

2.4.2 C=C stretches

In the retinal isomers, except for the severely hindered isomers, only a single'

intense "ethylenic line" is observed, even though there are five conjugated double bonds

and thus five C=C fundamentals. The vibrational spectra of the five common retinal

isomers have been analysized in detail. In all-trans-retinal, the ethylenic band appears at

1577 cm'. Because the C=C stretches are close together in energy and significantly

coupled through the n-electron system, they mix considerably in the normal modes.

Furthermore, the 1577 em" band, based on the peak shift of the deuterium and 13C

derivatives, has been shown to contain in-phase contributions from the Ci=C g, C9=CIO' and

Cll=C12 stretches, and the shoulder at 1550 ern:' is due to C14=C 15 stretch (Curry et al.,

1982).

The frequencies and isotopic shifts of the C=C stretching modes of 13-cis-retinal

are quite similar to those of all-trans isomer. The difference in this region is the 7 ern:'

higher wavenumber of the intense ethylenic stretching mode at 1584 ern" (Curry et al.,

77

1984). The 6 nm blue shift and reduced extinction coefficient of the n to n" transition

of 13-cis-retinal (Sperling, 1973) suggest that the cis bond shortens the effective region

of conjugation. The resulting decrease in n-electron delocalization along the chain is

reflected by a drop in the magnitude of the long range coupling constant between

conjugated stretches. This coupling change is responsible for the higher wavenumber of

the main ethylenic band for the cis isomer.

As was observed for the 13-cis isomer, the 9-cis bond decreases the effective

conjugation length and thus the long-range C=C, C=C potential coupling (Mathies et al.,

1987). The intense in-phase C=C stretch is therefore observed at higher wavenumber than

for all-trans-retinal. The 1587 cm' band of 9-cis-retinal is even higher than that of the

13-cis isomer, both because of the more central position of the cis bond and because of

a increased intrinsic Cl1=CI2 stretching frequencies.

The assignments of the observed bands of 9,13-dicis-retinal were made by

comparison with the spectra of the 9- and 13-cis isomers. The intense C=C stretching

band, which shifts from 1557 cm' for all-trans-retinal to 1584 em" for 13-cis and 1587

cm' for 9-cis retinal. increases even further to 1592 cm -I for the dicis isomer. This trend

that the increase in wavenumbers of this stretch in cis isomers is nearly additive (Curry

et al., 1985) was noted previously.

In contrast to other cis isomers, the ethylenic band of Ll-cis retinal does not

appear at higher wavenumber than that for the all-trans form. Such a reduced wavelength

could result from a small twist about the Cl1=C12 bond, which causes a decrease in the

n-bond order.

78

We note that in the C=C stretching region, most of the other isomers behaves in

similar manner as the above mono-cis isomers. The most noticeable changes are the

peaks for the unstable isomers (11,13-dicis, 7,1 1.13-tricis, 9,11, 13-tricis and all-cis) both

in terms of their relative intensities and a simultaneous appearance of a second peak at

slightly higher frequency. These trends have been noted and will be elaborated along

with a discussion of the mechanism of the thermal isomerization of these compounds,

presented in chapter 3. Briefly, these IR characteristics were attributed to the highly

twisted conformation of these isomers, making the Cl1=C12 and C13=C14 double bonds not

coplanar with the carbonyl group.

2.4.3 C-C stretches

2.4.3.1 C-C stretches of retinal isomers

The C-C stretching region is highly sensitive to variation of polyene geometry

(Mathies et al., 1989). In this fingerprint region, the normal modes are fairly localized

C-C stretches. Our analyses of the new isomers are again based on the frequencies and

assignments of these fingerprint modes in trans and the common mono-cis isomers, shown

in Figure 2.14. Effects of changes which result from the presence of the cis double-bond

are often additive.

The strong 1111 cm' fingerprint line is a highly localized C14-C15 stretch. Its

intensity in the IR spectrum arises from polarization of the charge distribution in this

bond by the nearby electronegative carbonyl oxygen (Mathies et al., 1985). The

79

significantly lower frequency of this mode compared to the other C-C stretches is a

consequence of the reduced C\4-C\5 bond order caused by its close proximity to the

electron-withdrawing oxygen. The 1163 cm' mode is a localized CIO-CII stretch. This

mode is at a frequency characteristic for an unsubstituted C-C stretch in a trans-polyene

chain. The normal modes having Cg-C9 and C12-C13 stretch character are significantly

higher in frequency at 1197 cm' and 1215 cm', respectively. This frequency increase

is characteristic of single-bond stretches with a methyl substituent on one carbon.

Coupling with the C\3-CH3 (or Cg-C9) stretch increases the frequency by -35 em:' (Curry

et al., 1985).

12221215 -- 1216 '" 12Il 1214

.......-.... 1204

Il97 ~ ~ 1188 :; -...._#.

1163 1162IlS4

Il131111 IllS-_ ..

-_ _--.......... 1148 .i->:

..._-\ ~1112 :-:

........_ ...._....__.... \\

\1084-----TRANS 13-CIS 9·CIS 7-CIS ll-CIS

Figure 2.14 Correlation diagram of the fingerprint frequencies for the trans and

mono-cis-retinal isomers.

80

The following effects of a cis C=C bond on positions of peaks in the fingerprint

region in 9-ds were noted (Curry et al., 1985). The frequency of a C-C stretch depends

on its interaction with the skeletal bends. The CCC bends associated with a cis double

bond couple less strongly with the adjacent C-C stretches, resulting in a 15-20 em:'

frequency decrease in the C-C stretch as mentioned earlier. Consequently, the CIQ-C11

stretch drops from 1163 cm' in all-trans-retinal to 1148 cm' in the 9-ds isomer as a

result of its proximity to the cis bond. The other C-C normal modes have the same

frequencies as observed in all-trans-retinal. Thus a -15 em" reduction in the CIQ-Cll

stretch frequency is characteristic of a 9-ds chromophore.

The fingerprint vibrational frequencies in 13-cis-retinal are very similar to those

seen in all-trans-retinal. The most noticeable changes are the 7 ern" increase in the C1Z­

C13 mode and the 6 em" decrease in the Cg-C9 and ClZ-C13 stretches in l3-ds-retinal

resulting from increased long-range coupling between the C-C stretches.. The largest

frequency change in l3-ds-retinal is expected in the Cl4-C l5 stretch, which should be

lowered because it is adjacent to the cis double bond. But such a change is not observed

in l3-cis retinal. This is probably due to the fact that the cis bond in l3-ds-retinal also

induces changes in the ground-state electronic structure. The Cl4-C15 bond order increases

as a result of an increased contribution from delocalized resonance structures such as

(Curry et al., 1984)

C=C-C=o t--+ C+-C=C·O'

which counteracts the direct geometric effect of the cis bond.

81

Isomerization about the Cl l-Cl2 bond has even more dramatic effects on this

fingerprint region. Altered coupling with the bends of the cis Cll=C12 moiety causes a

79 em:' decrease of the C IO-Cll stretch and an 11 em" decrease of the Cl2-Cl 3 stretch.

The Cl4-Cl 5 stretch is pushed up to 1126 ern" as a result of coupling with the CIO-CIl

mode, which is now below it. The Cg-C9 stretch is similarly pushed up by the lowered

C12-C13 stretch. Secondly, a new line appears at 1270 em", which is assigned to the

coupled in-plane rocking of the Cl1H and Cl 2H hydrogens. This is a characteristic

vibration of cis-vinyl rocks that is also observed in 7-cis-retinal.

For new isomers, we note the following trends. The effects of the cis ~=Cg bond

of 7-cis-retinal are the 8 cm' decrease in the Cg-C9 mode and the 9 ern" decrease in the

CIO-Cl1 mode. This is due to the same cis double bond effects for 9-cis retinal. The CCC

bends associated with a cis double bond couple less strongly with the adjacent C-C

stretches, resulting in frequency decreases in the Cg-C9 and C10-Cll stretches.

The single-bond stretching modes in 7,9-dicis retinal have been assigned on the

basis of 9-cis (Palings et al., 1987) and the 7-cis retinal assignments. These results have

been confirmed using isotopically labelled analogs, see section 2.4.2.2. A correlation

diagram for 9-cis, 7,9-dicis and 7-cis retinal isomers is shown in Figure 2.15.

Comparison of the 7,13-dicis-retinal spectrum with 7-cis and 13-ds-retinal spectra

is shown in Figure 2.16. Because the two cis double bonds are not adjacent to each

other, their effects on the C-C stretching frequencies are additive. Therefore, the results

are straight forward with the Cg-C9, CIQ-C11and Cl2-Cl 3 frequencies close to those of 7-cis

retinal and the Cl4-Cl 5 higher than both mono-cis isomers. Another dicis-retinal which

82

12111216

1200

1213-_ __ _-1188-- 1189--_ _-

1148 1148 1154----_ _._--_ _----_ ...

-- _--_ _.._- - __ .._--1113 1113 1112

9·CIS 7,9·DICIS 7·CIS

Figure 2.15 Correlation diagram of C-C stretching frequencies of 9-cis, 7,9-dicis and 7­

cis-retinal.

exhibits a similar additive relationship in the vibrational structure with its monO-CIS

isomer is 9,13-dicis-retinal. Correlation of results for 9-cis, 9,13-dicis and B-cis are

shown in Figure 2.17. In 9,13-dicis-retinal, the CIO-Ctl stretching frequency (1145 crn')

is very close to that (1148 em") of 9-cis and show a 17 cm' shift from that (1162 crn')

of 13-cis-retinal. This is due to the fact that the cis double bond C9-CIO causes a slight

downshift of its adjacent single bond stretching frequency. The C14-C15 stretch in the

9,13·dicis isomer is again insensitive to the 13-cis geometry, as noted before.

83

12221211 1212 ---_ .. _- ---- __ ...

11911188 1186--- ..- - __ -_ --

11621154 1156 ---_........ -_ ......... -_ .......__...-

~ .1122

. ~

7·CIS 7,13·DICIS 13·CIS

Figure 2.16 Correlation diagram of C-C stretch frequencies of 7-cis, 7, 13-dicis and 13­

cis isomers of retinal.

122212241216 ..-_ --_ .

1200CS,C 9 __••••••••••••••• 1194 1191-_ _-

1162

1148 1145.................-

-_ __ -1113 1115 1115

9·CIS 9,13·DICIS 13·CIS

Figure 2.17 Correlation diagram of C-C stretch frequencies of 9-cis, 9,13-dicis and 13­

cis isomers of retinal.

84

The l l-cis geometry in 7, l l-dicis-retinal makes its bond order of CIO-C11lower

than that of the 7-cis isomer. Consequently, the CIO-C11 frequency decreases from 1154

em:' in 7-cis-retinal to 1105 crn' in 7,11-dicis-retinal. Thus, the 1105 cm' band is the

CIO-CIl stretching frequency of the CIl=C 12 cis geometry in 7,11-dicis-retinal. Figure 2.18

shows the correlation diagram of the C-C stretching frequencies among the 7, 11-dicis and

its corresponding mono-cis-retina1 (7-cis and l l-cis-retinal). It shows that C!4-Cl5 and

C12-Cn frequencies are very close to each other while that of Cg-C9 of 7,11-dicis is about

28 em'! lower than that of l l-cis, as expected. Similar analyses of 9, l l-dicis-retinal have

been made by comparing with the 9-cis and l l-cis isomers, and the correlation diagram

for the observed C-C stretches is shown in Figure 2.19.

1211 1214--..... 1204 ...•.. 1204..._-_ -_ -1188 1186 ....•....-_ _ __»'

1154

11261128

- "............

..- _- _-....:'.. ~,..-

1112 .> · 1105--....•..••...... 1084

"--

ClO,C 11

C 14-C 1S

7·CIS 7.11.DICIS ll·CIS

Figure 2.18 Correlation diagram of C-C stretching frequencies of 7-cis, 7,11-dicis and

l l-cis isomers of retinal.

85

-------

12141216

1200 1202 .<.:::_-__.........:::.. 1204

-- 1187 .

1148

1126

1084

1124., --_ _-

~.>\::.:::.:

'\ 1088"--- ................. - .._-

9·CIS 9,1l-DICIS H·eIS

Figure 2.19 Correlation diagram of C-C stretching frequencies of 9-cis, 9,lI-dicis and

l l-cis isomers of retinal.

In tricis-retinal isomers, the polyene chain becomes more twisted because of the

presence of multiple cis double bonds. By comparing the spectrum of 7,9, l l-tricis retinal

with those of l l-cis and 7, l l-dicis-retinal, approximate assignments can be made (Figure

2.20). It shows that, the CI2-Cn and C14-C l5 stretches are approximately the same in the

three isomers. Expectedly the cis geometry of either ~=C8 and C9=CIO has no effect on

the vibrational frequencies of a single bond at the end of the polyene. However, the C8-

C9 stretch of 7,9,l l-tricis-retinal has decreased 17 em'! from 7,11-dicis due to the change

of trans to cis geometry at C9=C lO' and 45 em" from that of l l-cis retinal due to the

alternation at both C9=C lO and C7=Cg double bonds.

86

1214

1204 •... 1204 1204-,--........................ .. _-......... 1186

-,

- .•................. 1169

-_ _-_ _-1126 1128

1105

1125

1104

.... ..... ..... ...-_ _._-

1084_ .

ll-CIS 7,I1·DICIS 7,9,11·TRICIS

Figure 2.20 Correlation diagram of C-C stretching frequencies of l l-cis, 7,11-dicis and

7,9, l l-tricis-retinal isomers.

The fingerprint region of 7,9, 13-tricis retinal is expected to be similar to those of

13-cis retinal and 9, 13-dicis retinal. Therefore its assignment is based on comparison of

its spectrum with those of 13-cis-retinal and 9,13-dicis-retinal. The prominent peak at

1115 ern" is clearly that of CI4-C1S' characteristic of the 13-cis geometry. Since Cr=Cs

eis to trans isomerization generally has no effeets on CIO-Cl1 and C12-C13, the C12-C13 and

CIO-Ct I of 7,9-13-tricis are expected to be similar to those of 9, 13-dicis, i.e. around 1224

cm' and 1145 cm', the peaks at 1216 cm' and 1146 em" in the spectrum of 7,9,13-tricis

retinal are assigned to be C12-C13 and CIO-C 11 respectively. The assignments of the C-C

stretch modes of 7,9,13-tricis retinal are shown in Figure 2.21. .

87

121612241222--_ -- ......... ..........._-

118811941191-_ -- _-

1162

--...••.......•.. 1145 1146.._ -.._-

Cl4"C 151115 1115 IllS

13·CIS 9,13·DICIS 7,9,13·TRICIS

Figure 2.21 Correlation diagram of C-C stretching frequencies of l3-cis, 9, l3-dicis and

7,9,13-tricis-retinal isomers.

The vibrational spectra of the four sterically hindered isomers (11, l3-dicis,

7,11,13-tricis, 9,11,13-tricis and all-cis-retinal), are different from those of the other

isomers. Detailed discussions of their fingerprint region are presented in chapter 3 along

with the chemical propenies of these isomers.

2.4.3.2 Comparison of the aldehydes, Schiff bases and protonated Schiff bases of 7-cis

and 7,9.dicis isomers

Now that the C-C stretching vibrational modes of 7-cis and 7,9-dicis retinal, SB

and PSB have been assigned, it is possible to compare them and discuss the effects of

Schiff base formation and protonation on the fingerprint region of C-C stretches.

88

Figure 2.22 shows that the largest increase in all-trans retinylidene PSB appears

in the C14-C15 stretch, which shifts about 28 ern from 1163 to 1191 crn'. The C\2-Cn

stretch shifts up by 21 cm', whereas the CIO-C" and CS-C9 stretches experience smaller

shifts of approximately -4 and +7 em", respectively. Increased IT-electron delocalization

leading to a red-shifted absorption and higher C-C stretch frequencies is also observed

when the all-trans PSB binds to bacteriorhodopsin (Mathies et ai., 1987).

IIII

Figure 2.22 Correlation diagram of C-C stretching frequencies of all-trans retinal, its

unprotonated and protonated Schiff bases and BR56S (Smith et al., 1985).

89

It is evident that the increased complexity of the fingerprint region in the PSB

occurs because the C14-C15 normal mode shifts up into near degeneracy with the other

stretches. This new vibration must be identified in the fingerprint region of the various

PSB isomers. Also, the other C-C frequencies will be altered as a result of coupling with

the C14-C15 stretch. In Figure 2.23, a correlation diagram of the fingerprint vibrational

frequencies for the PSB isomers (Mathies et al.. 1987) is reproduced. For the all-trans

and 13-cis isomers, the assignments are based on isotopic derivative studies (Smith et al.,

1985). For the 9-cis and l l-cis isomers, the bands have been assigned based on the

trends observed in the aldehydes (Mathies et al., 1987). The frequencies of the 9-cis

isomer are very similar to those of all-trans except for the CIO-ClI stretch, which is 20 ern

1 lower in 9-cis. This is exactly the pattern observed in the aldehydes.

TRANS 13-CIS 9-CIS II-CIS1250

1237 1236 1237C1Z-C I3

1226~C8-C9

1200 1204 1202 1206

/1194 C14-C151191 ~ " 761192

1/66

II 50 1159

" 39

1100 -.1100 C,O-CII

Figure 2.23 Correlation diagram of the fingerprint vibrations for retinal protonated

Schiff base isomers (Mathies et al., 1987).

90

In Figure 2.24 is a correlation diagram for the C-C stretching frequencies of 7-cis

and 7,9-dicis retinal and their unprotonated and protonated Schiff bases. The data show

that the frequencies of the C-C stretches in the fingerprint region change dramatically in

response to Schiff base formation and protonation. The frequency ordering of the C-C

stretches in 7-cis and 7,9-dicis retinal are the same as that found in other isomers. The

C14-CISstretches are at 1112 and 1113 cm', the CIO-Ctl stretches at 1154 and 1148 ern"

and the methyl substituted Cg-C9 and C12-C13 stretches are higher at 1188, 1211 cm' for

7-cis and 1189, 1213 ern", respectively. The most dramatic results of Schiff base

formation are 55 and 59 cm' shifts of the C14-C15 stretches in 7-cis and 7,9-dicis Schiff

bases. The other C-C stretches that shift appreciably are the CIO-ClI stretches which rise

from 1154 to 1167 em" in 7-cis SB, from 1148 to 1163 em" in 7,9-dicis PSB.

Protonation results in a further increase for the Ct4-CISstretches. The other C-C stretches

shift relatively small due to protonation of the Schiff bases

The extent of increase in n-electron delocalization as a result of protonation of

the Schiff base nitrogen should be greatest near the Schiff base and decrease toward the

cyclohexenyl ring (Smith et al., 1985). The shift in frequencies of the C-C stretches upon

protonation is in agreement with this prediction. The frequency increase of the C!4-ClS'

C12-C13 and CIO-C lI Cg-~ stretches are 22, 18, -10 and -5 cm' in 7-cis PSB and 24, 16, ­

16 and -12 em:' in 7,9-dicis PSB, respectively.

Resonance Raman spectra of 7,9-dicis-rhodopsin in H20 and D20 at room

temperature (Loppnow et al., 1990) have been obtained. The shift of the 1654 cm' C=N

stretch to 1627 cm' in D20 demonstrates that the Schiff base nitrogen is protonated. The

91

absence of any shift of the 1201 em:' mode, which is assigned as the C14-C15 stretch, or

of any other C-C stretching modes in D20 indicates that the Schiff base C=N

configuration is trans (anti). The Schiff base C=NH+ stretching frequency and its D20

shift in 7,9-dicis-rhodopsin are very similar to those in 11-cis and 9-cis-rhodopsin,

indicating that the electrostatic or hydrogen-bonding environments of these Schiff bases

are effectively the same. The Raman bands of 1149, 1173, 1201 and 1236 cm' of 7,9­

dicis rhodopsin (Loppnow et al., 1990) are similar to those of the fingerprint bands (1147,

1179, 1196 and 1234 ern") of 7,9-dicis protonated Schiff base and have been assigned to

the c;.c, CIO-Cl\, CI2-C\3 and CI4-CI5. This fingerprint region of the 7,9-dicis rhodopsin

may be indicative of protein-chromophore interactions that are different from other

pigments. In 9-cis rhodopsin, the CIO-C1I and C14-C15 stretching frequencies are at 1154

and 1206 cm', shifted from 9-cis protonated Schiff base frequencies observed at 1137 and

1189 em" respectively (Palings et al., 1987). These shifts were interpreted as evidence

for a protein perturbation near C\3, a charge perturbation near C\3 may be expected to

produce similar shifts. However, much smaller shifts have been observed in the CIQ-C11

and CI4-C!5 stretching frequencies in 7,9-dicis rhodopsin. This suggests that the

postulated interaction between the charged group and the chromophore near C\3 is absent

in the 7,9-dicis rhodopsin, which is probably related to the blue shifted absorption, a

characteristic of the dicis pigment that might be common to all 7-cis pigment analogs.

7-Cis and 7,9-dicis are known to be sterically crowded with a C5-Cg dihedral angle

estimated to be about 40° for molecules in solution (Liu et al., 1983b). Recent

investigations on 7-cis-rhodopsin by low-temperature spectrophotometry and laser

92

photolysis (Shichida et al., 1991) indicated that the chromophore configuration plays an

important role in the interaction of chromophore and protein (This discussion will be

expanded in chapter 4).

...........-1211 . 1218 ......•.._ ..

C12-CI3

PSB

1236

SBCHO

1195

1188 1189 ./l'i'84_ -- - _-1167 •..•........

___--4.. 11571154 . ___ ,/

.".'

1112 ....., ....""

--"

7·CIS

7,9·DICIS

CIrCl3

12341213 1218 ...............•.--_ -.

119611911189 .'-_........................ <-; -»

1172 .....~::><::.... 1179,-'

../ 1163

: ::: :.:::~.::.:;.:/ __._ _ 1147

---J'

Figure 2.24 Correlation diagram of C-C stretching frequencies for 7-cis and 7,9-dicis­

retinal and their unprotonated and protonated Schiff bases.

93

2.4.4 Out-of-plane chain vibrations

Hydrogen out of plane bending (HOOP) bands have been utilized to interpret the

interaction between the chromophore and protein. In rhodopsin, the increased intensity

in the HOOP bands indicates that there is conformational distortion of the ClO-Cll=CI2-CI3

moiety (Eyring et al., 1980). In isorhodopsin the 960 cm' mode is not eliminated by

deuteration at CII and C12, suggesting that it is due to the ~=C8 HOOP mode, which is

similarly enhanced by conformational distortion. In the bathointermediate of rhodopsin,

the unusually intense HOOP bands have been interpreted as an indication of a

perturbation near C12 (Eyring et al., 1980). This perturbation was presumed to arise from

the same negatively charged protein residue that acts as the point charge in rhodopsin

(Honig et al., 1979). The intensities of these HOOP modes are due to conformational

distortions of the bathorhodopsin chromophore in the CIO to C14 region, and explicit

models have been proposed (Eyring et al., 1980; Honig et al., 1979).

In summary, isotopic substitution combined with normal mode calculations has

been the classical method for vibrational analysis. It has been long recognized that

vibrational spectra of the pigments are sensitive to the configuration and conformation of

the chromophore. In this chapter, the key vibrational modes for all sixteen retinal isomers

along with SB and PSB of 7-cis and 7,9-dicis isomers have been identified and

characterized. These results are necessary stepping stones to future studies of molecular

actions of isomeric retinal containing pigments.

94

Chapter 3

Temperature Dependent Isomerization of Retinal Isomers

3.1 Introduction

To understand the molecular mechanism of the photobleaching event of isomeric

retinyl bound proteins, extensive studies of solution photochemistry of retinal isomers

have been earned out either by direct photolysis (Kropf & Hubbard, 1970; Ganapathy &

Liu, 1992a,b) or triplet sensitization (Jensen et al., 1989; Ganapathy & Liu, 1992b). In

addition, a number of theoretical studies concerning photoisomerization of isomeric

retinals have been conducted (Schaffer et al., 1974; Birge & Hubbard, 1980). Those

studies preceding 1988 are covered in the review article by Becker (1988).. Since then,

Jensen et al. (1989) reexamined the triplet-state reaction redetermining quantum yields

of isomerization and photostationary-state compositions. Recently the synthesis of all

sixteen isomers of retinal has been completed (Trehan et al., 1990), following by

characterization of their thermal and photochemical properties (Zhu et al., 1992;

Ganapathy & Liu, 1992a,b). The isomerization process was found to be concentration

dependent giving, in some cases, quantum yields of isomerization greater than unity. In

these cases, a quantum chain process was believed to be involved. Under these

conditions, one-photon-two-bond isomerization has been observed to be present during

the photoisomerization of some of the retinal isomers.

95

Isomerization of retinals can be induced photochemically or thermally. Of the

sixteen possible isomers, four (11, 13-dicis, 7,11, 13-tricis, 9,11, 13-tricis and all-cis) contain

the doubly hindered 1l,13-dicis geometry. Wald & Hubbard (1955) first showed that

ll,13-dicis retinal is thermally unstable, isomerizing at room temperature to the 13-cis

isomer. Following their preparation, the 7,1 1,13-tricis, 9,1 1,13-tricis, and all-cis isomers

were also shown to be unstable at room temperature isomerizing about the 11-12 double

bond to 7,13-dicis, 9,13-dicis and 7,9,13-tricis-retinal respectively (Trehan et al., 1990b).

A rearrangement mechanism has been suggested, in analogy with related simple molecules

(Kini et al., 1979; Kluge & Lillya, 1971). However, accurate kinetic parameters for these

hindered retinal isomers in support of the. proposed mechanism were lacking. For

7,11,13-tricis and all-cis-retinal, preliminary data on activation energies of their

arrangement were reported (Trehan et al., 1990b), but for ll,13-dicis and 9,11,13-tricis

only the half lives at a limited number of temperatures were available (Wald & Hubbard,

1955; Knudsen et al., 1980). None of these data were sufficiently accurate for

determination of reaction parameters such as activation enthalpy and entropy.

In spite of the extensive effort made in studies of isomerization of retinals (Becker,

1988; Jensen et al., 1989; Ganapathy & Liu, 1992a,b), there are several interesting

chemical aspects of retinals that remain to be explained. Studies of the temperature

effects on the isomerization of retinal have been limited to the all-trans isomer (Waddell

et al., 1981a) and the data of their energy barrier which are in support of their

isomerization mechanism for any cis isomer are lacking. In an attempt to clarify these

points, we have made measurements of the solution photoisomerization of the isomeric

96

retinals involving ll-cis, 9-cis, 7-cis and 7,9-dicis isomers, and thermal isomerization

including four doubly hindered retinal isomers.

For photoisomerization of polyenes, the initial product distribution (proportional

to the quantum yield of product formation) reflects the relative ease of isomerization at

each double bond in a given isomer. In this chapter, the initial product distributions as

a function of temperature have been determined, The study involved the use of HPLC

as an analytical tool in the determination of the photoisomerization quantum yields of

retinals in solution at low temperature upon direct excitation. Calculations on excited

state properties have been performed to substantiate the mechanism of photoisomerization

of the retinal isomers. Also, an improved procedure for obtaining and analyzing the

UVNis data used in monitoring the thermal rearrangement reaction provided more

accurate data which is sufficient for calculation of all of the activation parameters of the

hindered isomers.

It should be pointed out that the protonated Schiff base (PSB) is generally

recognized as the most appropriate model system for the visual chromophore. However,

an accurate analytical method for PSB isomeric mixtures is not available. For example,

the recent report on HPLC separation of PSB isomers (Mukai et al., 1992; Koyama et ai.,

1991) provided rather poor peak resolutions giving results not substantially improved over

the traditionally more qualitative NMR method (Childs & Shaw, 1988). On the other

hand, excellent conditions for separating retinal isomers are known (Zhu et al., 1992).

Therefore, in this photochemical study, only the retinals have been examined.

97

----_._-

3.2 Experiments

3.2.1 Sample preparation

Procedures for preparation of retinal isomers were discussed in the previous

chapter. They were purified by preparative HPLC.

3.2.2 Methods

Infrared spectra were recorded on a Nicolet 740 FTIR spectrometer (resolution,

I cm': scan, 100). The possibility of thermal isomerization of each sample used for the

FTIR study was examined by HPLC. The purity of these four isomers was found to .

exceed 95%.

HPLC analysis were conducted on Rainin Dynamax system using a 25 em x 8 mm

i.d. column packed with 5 urn silica gel. The eluent solvents are 1% methyl t-butyl ether

(MTBE) in trichlorotrifluoroethane or 4-5% ether in hexane. Extinction coefficients of

the retinal isomers at 360 nm were used to correct the HPLC peaks.

The thermal isomerization reactions were carried out in an UVNis cell situated

in a thermostated cuvette cell holder of a Perkin-Elmer Lambda 5 absorption

spectrometer. The temperature of the solution was determined by an immersion

thermocouple thermometer. The progress of the reaction was monitored by the increase

in absorbance at the absorption maximum of the product isomer. Four runs at

temperatures ranging from 20 to 500 C were conducted for each hindered isomer, reaching

conversions for about three half-lives. The reaction mixture was then allowed to convert

98

completely to the product isomers in order to determine final absorbance, A_. For lower

temperature runs, this involved warming the solution to 40°C for 10 h before cooling to

the original temperature to record the final spectrum. The changes in absorbance,

expressed as In (A_-At ) is a measurement of the fraction of conversion (in x). They were

used for calculation of rates of isomerization which are shown in Table 3.5.

3.2.2.1 Irradiation procedure

All samples were freshly purified by preparative HPLC. Retinal solutions in

hexane with concentration of 1 x 10-3 M was irradiated with 366 ± 4 nm light from a 150

W Hanovia Xe-Hg arc lamp isolated with a combination of Corning 0-52 and 7-60 filter

plates. The temperature of the solution was determined by a thermocouple thermometer.

The products of irradiated samples were analyzed by HPLC. The absorption coefficient

at the detection wavelength were used to correct the chromatography diagrams. The

photoisomerization reaction was followed up to 10% completion.

3.2.2.2 Quantum yield measurements

Quantum yield measurements were carried out on a small merry-go-round

apparatus (Applied Photophysics). All-trans-retinal was used as a reference to monitor

the light intensity. Its concentration dependent quantum yield data are in the literature

(Ganapathy & Liu, 1992). The irradiation products were analyzed by HPLC. The

products were identified by retention time and UV spectra. Product analyses were

99

conducted at 360 nm where correction factors for each isomer are available (Zhu et al.,

1992).

3.2.2.3 Calculation methods

The calculation of bond orders of the retinal isomers has been preformed using a

semi-empirical quantum mechanical program. The starting geometries of the retinal

isomers were calculated using PCMODEL (Molecular Modelling software from Indiana

University). All geometric parameters (bond angles, bond lengths and dihedral angles)

were minimized via the MOPAC program using the MNDO Hamiltonian method. To

calculate the bond order of the excited state molecule, the geometry of molecule was

optimized for the first singlet state. The structural parameters (bond lengths and dihedral

angles) comparing, whatever available; all-trans (Hamanaka et al., 1972); l l-cis (Drikos

et aI., 1981); 13-cis (Simmons et al., 1981) and 9-cis (Simmons et al., 1986), to crystal

structural data are listed in Table 3.1-3.2. A bond order is defined as the sum of the

squares of the density matrix elements connecting any two atoms. Then the density

matrix is decomposed into sigma, pi and delta components. This is very useful

information on the pi electron character of the conjugated system.

The potential energies of the excited state of the retinal molecule have been

calculated using the MOPAC program. First the geometry of each isomeric retinal in the

excited state were optimized. With the aid of the PCMODEL program, the dihedral angle

of the each double bond was rotated while the rest of molecular geometry remained fixed.

100

Then the energies of excited state molecules were calculated based on the twisted

geometry.

101

-oN

Table 3.1 Experimental and calculated bond distances (A) of all-trans and cis isomers of retinal

Bond Experimental Calculated Ground State Calculated Excited State

all·t· 9·cis· l l-cls' B-cis' all-t 7-cis 9-cis l l-cis 7,9·cis all-t 7·cis 9·cis l l-cis 7,9·cis

1,2 1.534 1.529 1.523 1.523 1.567 1.570 1.56X 1.569 1.544 1.566 1.569 1.56X 1.564 1.567

1,6 1.540 1.539 1.528 1.534 1.547 1.553 1.550 1.549 1.524 1.549 1.549 1.551 1.554 1.563

2,3 1.421 1.489 1.498 1.424 1.534 1.531 1.530 1.569 1.529 1.533 1.536 1.534 1.533 1.534

3,4 1.493 1.511 1.532 1.540 1.535 1.534 ISB 1.532 1.530 1.540 1.534 1.536 1.533 1.533

4,5 1.507 1.516 1.521 1.515 1.51X 1.516 1.515 1.51X 1.50X 1.519 1.518 1.518 1.518 1.515

5,6 1.329 1.330 1.333 1.331 1.365 1.367 1.365 1.368 1.35\ 1.373 1.370 1.377 1.36X 1.373

6,7 1.483 10489 10486 10483 IAXX IAX7 10488 10489 1.502 1.480 1.4711 10472 10486 1.473

7,8 1.317 1.312 1.339 1.311 I.W) 1.349 1.350 1.348 1.353 1.365 1.362 1.376 1.358 1.371

8,9 10469 1.451 10461 1.463 1.486 104M \.483 1.484 1.474 1.457 1.462 1.446 1.470 1.446

9,10 1.346 1.336 1.347 1.364 1.356 1.360 1.358 1.359 1.367 1.410 1.405 10407 1.385 1.416

10,11 1.433 1.439 1.454 1.456 1.470 1.467 1.470 1.484 1.464 1.437 1.436 10436 1.436 1.441

11,12 1.339 1.353 1.339 1.354 1.349 1.351 1.349 1.349 1.362 1.377 1.381 1.376 1.392 1.166

12,13 1.455 1.424 1.472 1.443 1.485 1.486 1.485 1.484 1.471 1.462 1.462 1.470 1.447 1.474

13,14 1.346 1.364 1.358 0.313 1.354 1.356 1.355 1.157 1.366 1.376 1.373 1.369 1.385 1.366

14,15 1.458 1.429 1.467 1.416 1.491 1.491 1.490 1.489 1.468 1.479 1.479 \.479 1.475 1.482

15,16 1.201 1.221 1.213 1.201 1.222 1.223 1.222 1.223 1.221 1.227 1.227 1.228 1.229 1.226

• references see text.

-oU.)

Table 3.2 Experimental and calculated dihedral angles of retinal isomers

Bonds Experimental" Calculated Ground State Calculated Excited State

all-l 9-cis l l-cis . 13-cis all-t 7-cis 9-eis l I-cis 7,9-cis all-l 7-cis 9-cis l l-cis 7,9-cis

C-C 6,7 -54 -76 41 65 -55 62 -54 75 76 -5S 60 54 SO 59

H,9 -177 -177 171 IllO -162 -171 -15K 139 16K -161 -146 161 144 -145

10,11 IllO -175 -179 IHO IXO 177 -17X 161 175 -17X ISO 173 166 -17X

12,13 170 -179 39 179 167 170 16S 123 167 -154 153 149 139 145

14,15 179 -I XO -175 175 177 176 -175 -17K -176 162 -171 1711 176 -174

C=C 5,6 17K 176 179 175 IT! -175 173 -179 -175 174 -174 -172 -179 -174

7,8 -177 -178 180 177 -178 4 180 -179 -6 180 I 17X 180 4

9,10 17S 0.9 -176 IllO 178 -180 7 -179 -7 167 177 -15 -177 22

11,12 -17X 179 2 177 -179 180 ISO -3 17S -176 176 177 -22 174

13,14 179 -179 180 1.2 177 177 -177 179 -177 17S IXO 179 IXO -178

•• references see text

3.3 Results

3.3.1 Photoisomerization of retinal isomers

3.3.1.1 Direct irradiation of retinal isomers at different temperatures

Photoisomerization of retinal isomers in hexane solution at various temperatures

has been carried out. The isomeric compositions of the irradiated retinal mixtures were

analyzed by HPLC. The results are shown in Figure 3.1-3.4 in the form of plots of the

progress of photoisomerization of l Lcis, 9-cis, 7-cis and 7,9-dicis-retinal isomers.

Irradiation of l l-cis-retinal with 366 nm light at room temperature gave all-trans

and 11,13-dicis in the ratio of -2: 1. When the irradiation temperature was lowered, the·

ratio of all-trans and 11,13-dicis increased (Figure 3.1). The results suggested the

possibility that the energy barrier of rotating the Cll=CI2 double bond is smaller than that

'of C13=CI4 double bond. Similar trends have been observed in the 9-cis-retinal (Figure

3.2) and 7-cis-retinal (Figure 3.3), in the respective product ratios of all-trans to 9,13-dicis

and all-trans to 7,13-dicis. Similarly, we suspect that the energy barrier of rotating the

cis double bond at ~=C8 or C9=CIO is lower compared to that of rotating the

corresponding C13=CI4 double bond.

Photoisomerization of 7,9-dicis-retinal gave initially all-trans, 7,9,13-tricis, 9-cis

and a small amount of the 13-cis isomer upon direct irradiation with 366 nm light at room

temperature. As the irradiation temperature is decreased, the product distribution was

found to be altered with the 7,9, 13-tricis isomer decreasing, while 9-cis increased. The

amount of all-trans-retinal, a one-photon-two-bond isomerization product, remained the

104

same at different temperatures. The results for 7,9-dicis-retinal suggested that the energy

barrier for rotating the ~=C8 double bond is lower than that of C13=CI4 , and a different

pathway was possibly involved for formation of all-trans.

The product ratio of one bond isomerization plotted against the reciprocal of

temperature for l I-cis, 9-cis, 7-cis and 7,9-dicis-retinal are shown in Figure 3.5. The

slope of each curve is a measure of relative energy barriers of isomerization taking place

at different double bonds.

105

11,13-dicis

2 4 6 8 10% Conversion

y = 0.10 + 0.933x R'2 = 1.000

Y= - 0.10 + 6.67e·2x R~2 = 0.97812FT:-80 C

10Ul all-trans...IIIE 80Ul

0 6-0s:0. 4~

:k::0 11,13-dicis

J• • t • ~I i I I i i

2 4 6 8 10 12% Conversion

'if!. 2

UlG; 6EoUl

oo 4s:0.

y = 0.12+ 0.73x R~2 = 0.999

Y = - 0.11 + 0.27x R~2 = 0.991

8~T:23 C

Ul... 6IIIE0Ul

0

0 4s:0.

~0 2

00 2 4 6 8 10

% Conversion

y = . 2.32e-2 + 0.804x RA2 = 1.000-0 sF+ O." . '

R'2 = 0.9990'

T:-25 C

G; 6E0Ul

0

o 4.c0.

~02

00 2 4 6 8 10

% Conversion

Figure 3.1 Progress of products formation during direct irradiation of ll-cis-retinal in hexane at four different temperatures.

o r . I -\L - • •••• Iii' i • I iii 0 iii Iii iii

o 2 4 6 8 10 0 2 4 6 8 10% Conversion % Conversion

Figure 3.2 Progress of products formation during direct irradiation of 9-cis-retinal in hexane at four different temperatures.

y = - 0.11 + 0.844x R'2 = 1.000 Y = - 8.1e·3 + 0.874x R'2 = 0.999

Y = 6.7e-2 + 0.162x R'2 = 0.977 Y = 1.0ge-2 + 0.125x R'2 = 0.94910

T=23" C 10pT=0.5 C

III 8 8.. ...Gl GlE E0

60 6.~ .!!!

0 0-0 au-trans 0s:

4s: 40. 0.

::!! ::!!09,13-dicis

0

2

:1«9,13-dicis

• ~

O~ i i II I I i I i i I i

0 2 4 6 8 10 0 2 4 6 8 10% Conversion °/0 Conversion

y = - 3.34e-3 + 0.891x RA2 = 1.000 Y = 3.1e-3 + 0.955x R'2 = 1.000

- Y = 3.3e-3 + 0.109x R'2 = 0.998 - 9.4e-3 + 4.61e-2x R'2 = 0.9990 Y =....... 10

101 t>T=-25 C

.. 8 .. 8

T=-80 C

Gl Gl

E E0 0

6.l all-transIII 6 III

s 0

0 0s: s:0. 4 0. 4

::!! ::!!0 0

2 9,13-dicis 2s.f a-drcts

1082

~ .' JI' •o Iii i i : I i

° 4 6 8 10 0 2 4 6'Yo Conversion % Conversion

Figure 3.3 Progress of products formation during direct irradiation of 7-cis-retinal in hexane at four different temperatures.

y = - 2.8ge-2 + 0.852x R'2 = 0.999 Y = 1.06e-2 + 087x R~2 = 1.000Y :: 2.8ge-2 + 0.148x W2 :: 0.982 y = - 1.06e-2 + 0.13x RA 2 :: 0.995

B1 T=23 " c :7 I lap1::0.5 C

~ 6 ~ 8ell Cl

E E0 0III III 60 4 0

0 -0s: s:0. 0. 4

~ 2 7,13-dicis ~0 0

21 -: 7,13-dicis

-aa 2 4 6 8 10 0 2 4 6 8 10% Conversion 0/0 Conversion

y = - 5.4e-3 + 0.903x R~2 = 1.000 Y :: - 2.73e-2 + 0.947x RA 2 = 1.000-0 y :: 5.4e-3 + 9.74e-2x RA2 :: 0.996 y :: 2.73e-2 + 5.2ge-2x R~2 = 0.956oc8 11=-25" C l°F7 I 1::-80 C

~8

~ 6ell ell

E E0 0

6III III

04 s

0 0.I: .I:

40. 0.

~ ~0 2 0

7,13-dicis 2 . ~ 7,13-dicis

y = 0.23 + 0.829x R'2 = 0996Y = - 2.22e-2 + 7.04e-2x R'2 = 0.992Y = - 8.11 e-2 + 6.35e-2x R'2 = 0.987

1:0.5 DC10..-

y = 0.366 + 0.829x R'2 = 0.999Y = 6.40e·2 + 7.31e-2x R'2 = 0.997Y = - 2.2e-2 + 2.96e-2x R'2 = 0998

10 I " 7 I1=23 C

4

2 7.9.13-lricis 9-cis

a~

0 2 4 6 8 10% Conversion

o!!.

~ 8ell

Eo:~ 6o-os:Q.4

2J /9.13-lriCiS 9-cis

~a-~ I i ei i U i

0 2 4 6 8 10 12% Conversion

(/) 8...QI

E~ 6o-oL:

a.~o

7.9.13-tricis

all-trans

T:-80"C

y = - 1.le-2 + 0.688x R'2 = 1.000Y = 2.95e-2 + 0.305x R'2 = 0.997Y = 2.6ge-2 + 6.78e-3x R'2 = 0.990

8

e 6QIE0~

£ 40L:a.,,!! 20

7.9.13-tr.icis

all-trans

1=-25 ()C

y = - 0.18 + 0.894x R'2 = 0.999Y = 0.21 + 8.36e-2x R'2 = 0.985Y = - 365e-2 + 3.43e-2x R'2 = 0.980

~ 12

10...ell

E 80(/)

0

0 6.cQ.

4~0

2

0 f'FT ••• p. -I ~ • II IJ J ,M. Ii' ' »i 0' i i. , f' • iii;

o 2 4 6 8 10 12 0 2 4 6 8 10 12% Conversion % Conversion

Figure 3.4 Progress of products formation during direct irradiation of 7,9-dicis-retinal in hexane at four different temperatures.

trans/9,13-dicis

trans/7,13-di is

trans/11 ,13-dicis9-cis/7,9,13-tricis

1-f--~~r------.----...----------...------1

100 -r---------------_-_

0-ca~

-10o~

"C0~

a.

3 4 5

(1/T)x1000

6

Figure 3.5 Plots of product ratio of retinal isomers at different temperatures.

110

The initial product distributions resulting from irradiation of 7,9-dicis and l l-cis

retinal were determined to be independent on the excitation wavelength (from 340 nm to

420 nm).

The quantum yields of isomerization for 7-cis and 7,9-dicis-retinal at two different

temperatures have been measured, using all-trans-retinal as a reference of which the

quantum yield values at room temperature and at 193 K have been previously determined

(Ganapathy & Liu, 1992b; Waddell & Chikara, 1981a). The resulting quantum yield

values of total and component isomerization processes of 7-cis and 7,9-dicis-retinal at 298

K and 193 K are listed in Table 3.3.

Table 3.3 Total and component quantum yields of photoisomerization of retinal

isomers at 298 K and 193 K

all-trans 7-cis 7,9-dicis

Total T-13c T-9c Total T-13c 7c-T Total Trans 13c-T 7c-T

298K 0.12 0.10 0.015 0.71 0.09 0.62 0.57 0.51 0.04 0.02

193K 0.0194 0.018 0.0014 0.27 0.02 0.25 0.20 0.13 0.01 0.06

3.3.1.2 Calculations of bond order and energy barrier of retinal isomers

The calculated results of bond order for selected retinal isomers at the ground state

and the first excited state are listed in Table 3.4. We notice that the bond orders of

111

C13=CI4 and ClI=C I2 of the excited ll-cis-retinal are 0.5658 and 0.2114. The higher value

for the C13=CI4 is consistent with the experimental data that the energy barrier of rotating

C13=CI4 is larger than that of rotating ClI=CI2 in 11-cis-retinal. In 9-cis-retinal, the bond

order of C13=CI4 (0.6739) is also higher than that of C9=CIO (0.1774), thus also consistent

with the isomerization data (Figure 3.2). However, results of 7-cis-retinal are not in

agreement with experiment. The experimental result suggests that rotation of ~=C8 bond

of 7-cis-retinal is easier than that of C13=CI4 bond, while the calculated bond order for

~=C8 is 0.7018 and for C13=C14 0.5847. This indicates that excited state bond order is

not the only factor controlling relative ease of isomerization. Environment effects are

likely to play roles in affecting the kinetic factor. The calculation results of 7,9-dicis­

retinal with bond order 0.6244 for ~=C8 and 0.7034 for C13=CI4 double bonds are

consistent with the experiment data.

Another perhaps more meaningful way to examine the ease of isomerization is to

determine relative slope of torsional potential energy in twisting different double bonds.

This calculation is more difficult because minimization of energy required as each of the

bonds is twisted. Results of such a calculated study are shown in Figure 3.6. The slopes

of the plots in Figure 3.6 indicate that rotation of ClI=C12 in l l-cis-retinal is relatively

easy to take place than that of C13=C14 • The resulting curves of potential energy can be

used to explain the photochemical results of these retinal isomers. For 7-cis, 9-cis-retinal

isomers, the rotation of cis double bonds is labile to isomerize by comparing the slope

of potential energy versus dihedral angles, which is exactly what was observed from the

photoisomerization reaction.

112

v.J

Table 3.4 Calculated Bond Orders of Ground State and First Excited State of Retinal Isomers Using the MNDO Method.

all 5=6 6·7 7=H H·9 9=10 10-1i 11=12 12·13 13=14 14-15 15=()

So 0.9309 0.0385 0.9767 0.0150 0.9372 0.0532 0.9442 0.0387 0.9582 0.03K9 0.9981

S\ 0.8904 0.0754 0.6245 0.3165 0.IX94 0.5666 0.2X26 O.2X64 O.621X (l.(1845 0.9398

7·c

So 0.9268 O.(142X O.973X 0.0375 0.9092 0.0737 0.933X 0.035X 0.9353 (1.04XO 0.9862

S\ 0.K890 0.0681 0.7018 0.2555 0.2224 0.5664 0.24K5 0.3162 0.5K47 0.0920 0.9306

9-c

So 0.9302 O,{1396 0.9678 0.(1418 05.1282 0.0552 0.9400 0.0350 0.9599 0.()444 0.9911

S\ 0.8606 0.1023 0.5458 0.3745 0.1774 0.5451 0.3263 0.2400 0.6739 O.083X 0.9390

ll-c

So 0.9329 0.0362 0.9789 0.0363 0.91X2 0.0677 0.9277 0.0365 0.9454 0.0518 O.9K IK

SI 0.9305 0.0420 0.7792 0.1951 0.2936 0.5425 0.2114 0.3323 0.5658 0.0988 0.9224

7,9·d

So 0.9332 0.0373 0.9747 0.0427 0.8996 0.0779 0.9310 0.0349 0.9414 0.(1439 0.9914

S\ 0.8816 0.0840 0.6244 0.3222 0.1637 0.5404 0.3372 0.2152 0.7034 O.07K5 0.9463

130 13011-cis ~ 9-cis

125~

0 0 125E 13=14 E- 120 -III

~III

U U.:i: .:i: 120

115~....

1101 ~ I....

CIl CIlW 115

M w-tC

11=12105 I , i i I , i , I , I I 110

0 10 20 30 40 50 0 10 20 30 40 50Double Bond Rotation Double Bond Rotation

1301 130

- 7·els ~ I 17,9-dicis+:-

- 125 ~

0 0 125E E- -III IIIU U.:i: 120 .:i: 120.....~ ~.... ....CIl 115 CIl..... ..... 115w w

9=10

110 1100 10 20 30 40 50 0 10 20 30 40 50 60

Double Bond Rotation Double Bond Rotation

Figure 3.6 Calculated potential energy barrier of cis isomers at first excited state.

3.3.2 Mechanism of the thermal isomerization reaction of the retinal isomers with

the 1l,13·dicis geometry

3.3.2.1 Thermal isomerization reaction of thermally unstable retinal isomers

The thermal isomerization reaction of 11,13-dicis-retinal was monitored at different

temperatures (20°C, 30°C, 40 DC, 50°C). In these experiments, as time progressed, a

decrease in the absorption at 302 nm and an increase in the absorption at 357 nm was

observed. The time dependent plots of the UVNis absorption spectra are shown in

Figure 3.7, each containing an isosbestic point, indicating direct conversion to the product.

The rates of the reaction at various ternperatures were determined form increase

of absorbance at absorption maxima of the l l-trans product in plots of log of increase of

absorption of product versus time at different temperatures. Standard treatment of rate

constants versus temperature yielded energy and entropy of activation for the

transformations.

Specifically, the rate constants of isomerization of 11,13-dicis-retina1 to the

product 13-cis-retinal at 50.3, 39.8, 30.5 and 21.2 °c were calculated from the plots in

Figure 3.8, which are listed in Table 3.5. The different rate constants (k) of isomerization

were plotted against temperature in the form of lntk/T) versus Iff (Figure 3.11a) yielding

a straight line. From the slope and y-intercept, energies of activation and entropy of

activation of the thermal isomerization reaction from 11,13-dicis-retinal to 13-cis-retinal

were calculated.

115

The products of isomerization of 7,11,13-tricis, 9,11,13-tricis and all-cis-retinal

were found to be 7,13-dicis, 9, 13-dicis, and 7,9,13-tricis-retinal. The progress of7,11,13­

tricis, 9,1l,13-tricis or all-cis-retinal to 7,13-dicis, 9, 13-dicis, or 7,9,13-tricis-retinal

respectively was followed in a similar manner using UV/Vis absorption spectroscopy. For

7,11,13-tricis-retinal, the corresponding absorption versus time plots at different

temperature, and Ink versus reciprocal temperature are shown in Figure 3.8 and Figure

3.12b. And the corresponding plots for 9,11,13-tricis to 9,13-dicis-retinal (measured at

359 nm) are shown in Figure 3.9 and Figure 3.12c, and for all-cis to 7,9,13-tricis-retinal

(measured at 346 nm) are shown in Figure 3.10 and Figure 3.12d.

116

Table 3.5 Rate constants of isomerization of the four labile isomers at various

temperatures in methylcyclohexane and the calculated enthalpy and entropy of

activation

ISOMERS T k L}H* L}S* L}E*a(oC) (mol) (kcal) (cal/mol)

11,13-c 50.3 0.027 22.4±O.7 -4.7±1.7 23.0

39.8 0.00868

30.5 0.00255

21.2 0.000784

9,11,13-c 49.8 0.0293 21.2±O.6 -8.5±1.0 21.8

39.8 0.00884

30.6 0.00345

21.1 0.00108

7,11,13-c 49.3 0.0338 21.5±O.4 -6.8±1.3 22.1

39.5 0.0115

30.6 0.00442

21.1 0.00119

all-c 49.8 0.0287 23.1±O.2 -2.3±O.5 23.6

39.5 0.00844

30.6 0.00302

21.3 0.000789

a: 6H* + RT

117

0uj:l

(a)

III

o I

.QJ.l

U

/ \

0

j:l

15

CII

III

(0)

~

.QJ.l0CII

~

300 400 500 300 400 500

Wavelength (run) wavelength (run)

(e) 0 1 / \ .. (d)

H~,IIi~'uj:lIII

00 .ClJ.l0CII

~

300 400 soo 300 400 500

Wavelength (run) Wavelength (run)

Figure 3.7 UVNis absorption changes during conversion of 11,13-dicis-retinal to lS-cis-retinal in methylcyclohexane at (a)50.3°e, spectra taken at the following intervals: 0 (curve 1), 5, 8, 12, 15, 18, 21, 27, 31, 37, 46, 60, 72 m and 00 (curve 14);(b) 39.8°e, at 0 (curve 1), 10, 16, 22, 30, 38,46,60, 76, 90, 110, 140, 170, 210 m and 00 (curve 15); (c) 30.5°C, at 0 (curve1),60,90,120,160,180,210,240,270,300,375,450,540 m and 00 (curve 14); and (d) 2 I.2°C, at 0 (curve 1),60, 120, 180,270, 385,490,610, 720, 840, 1020, 1200, 1320 m and 00 (curve 14). The last entry of each after warming overnight at 40°C.

~::K I0.0

(a)-0.5

-10

I.

5 -1.5 1 ~""'- I -loS~ri

-2.0 j~I

-2.0

. . -2.5-2.5 . I I0 20 40 60 80 0 50 100 150 200

time (min) time (min)

(d)

500 1000 1500

time (min)

0.0 I I 0.0-\C-0.51~ I -0.5

IiI -1.01 '""h.... I -1.0•l'l;~

~ri

.

, 5

1 =:J-1.5

-2.0 , I , I , I . I -2.00 100 200 300 400 500 0

time (min)

Figure3.8 Change of absorbance, In(A_-AJ versus time, at 363 nm during thermal reactions of II, 13-dicis-retinal at (a) 50.3,(b) 39.8, (c) 30.5 and (d) 21.20 C.

250200100 150

time (min)

50

--4.0 I i I • iii • I i I • I • I • Ii> I -4.0 Iii I • I iii

o 10 20 304050 60708090 0

time (min)

-1.0 ........ . - . I -1.0

,; -2.0 ~ ~ I ~~(b)

I-2.0

ttt.'r::r-i -3.0, ~ I -3.0

-0.5 I •

1500

(d)

1000

time (min)

500

-2.0

-1.0

-1.5

-2.5

-3.0 I ' I ' I ' I ......, I

100 200 300 400 500 600 0

time (min)

-1.0

~

,;J -2.0

~

~r-i

-3.0

-4.00

No

Figure 3.9 Change of absorbance, In(A_-AI) versus time, at 357 nm during thermal reactions of 7,11,13-tricis-retinal at (a)49.3, (b) 39.5, (c) 30.6 and (d) 21.2°C.

time (min)

-1.0 I I

200

(b)

150100

time (min)

50

-2.0

-3.0 I i I I I i I • "Ia

-2.5

-1.5

125100755025

-1

-2

-~

-3I,

~~r4 -4

-1.0 I I

1500500 1000

time (min)

-1.0 I I

-3.5 I I I I I I I I Ia

-2.0

-3.0

-2.5

-1.5(e)

100 200 300 400 500 600

time (min)

-3.5 I i I • I ' I ' I ' I i Io

-3.0

-1.5

-~ -2.0~

I,~ -2.5

I=:r4

tv

Figure 3.10 Change of absorbance. In(A~-At) versus time, at 359 nm during thermal reactions of 9,11,13-tricis-retinal at (a)49.8, (b) 39.8, (c) 30.6 and (d) 2I.l°C.

(b)

100 200 300 400

time (min)

-2.0 - I -2.0(a)

I "b-...~ -3.0,( J a I -3.0I

tt,' -4.0-J::rl I <, I -4.0

-5.0

-6.0 I "" -5.0. I . I ,0 50 100 1 5 0 0

time (min)

1500

(d)

1000

time (min)

500

-1.6I-Jt-J

-2.0 -i -.., \CI I -t.a

-2.0tt, 1 <, II -2.2,~ -3.0

~J ~-2.4

-2.6

. I . I -2.60 200 400 600 0

time (min)

Figure 3.11 Change of absorbance, In(A~-AI) versus time, at 346 nm during thermal reactions of all-cis-retinal at (a) 49.8,(b) 39.5, (c) 30.6 and (d) 21.3°C.

-8 I I

3.4

(b)

3.33.23.1-13 I . I ' I • I ' I

3.0

·8 to:: I

-12

-10

-9

-;; -11

....:;

3.4

(a)

3.33_23.1-13 I ' I ' I ' I ' 'I'

3.0

-9

-12

,y;

-;; -11

_ -10....

(11T)x1000 (11T)x1000

3.4

(d)

3.33.23.1

-8 [ I

-12

·9

-10

,y;

-;; -11

....

-13 I ' I ' I • I ' 'I'3.4 3.0

(e)

3.33.23.1-13 I ' I ' I ' I • I

3.0

-9

-10

-12

·8 I I

t--,y;

-;; -11

N'..#J

(1/T)xl000 (1/T)xl000

Figure 3.12 Plots of In (kIT) versus Iff for determination of enthalpy and entropy of activation for isomerization of (a) 11,13­dicis-retinal, (b) 7, II, 13-tricis-retinaI, (c) 9,11, 13-tricis-retinal and (d) all-cis-retinal.

3.3.2.2 FTIR spectra of four thermally unstable retinal isomers

The FTIR spectra in the 1650-1550 cm' region (in Figure 3.13) of four labile

hindered retinal isomers are different from the other retinal isomers. Most other retinal

isomers have an intense peak at 1580-1590 cm' with a shoulder at lower wavenumbers.

To the intense peak was assigned the conjugated ~=Cg, C9=C IO and CIl=C12 and to the

shoulder the Cl3=C14 double bond stretch modes (Curry et al., 1985). However, in retinal

isomers containing 11,13-dicis geometry, the steric effect made CI2-C13 single bond more

twisted so that the double bonds of polyene chain become less conjugated. The CIl=C12

and Cl3=C14 double bonds are more localized than those of other isomers so that

frequencies of these two double bond stretching modes expectedly increase. Therefore,

the region of 1616-1612 cm' are assigned to be mainly CIl=Cl2 and C13=Cl4 mixed

double bond stretching modes and the remainder signals at 1580-1590 ern" are C7=Cgand

C9=C IO conjugated double bond stretching modes. Also the frequencies of C=O stretching

mode appearing at higher wavenumbers (1676-1679 crn' instead of the common values

of 1660-1670 cm') reflect the highly distorted shape of the polyene chain. The C-C

single bond region, on the other hand, is characterized by the appearance of a single

medium intensity peak near 1170-1171 ern", which is based on the assignment of the all­

trans isomer (see chapter 2), corresponding to overlapping signals of Cg-C9, CIO-CIl and

C12-C13•

The HOOP region has been assigned based on the knowledge from other known

retinal isomers (see chapter 2). The 960 cm' peak due to the H7 and H, for isomers with

the 7-trans geometry appears only in the 11,13-dicis and 9,11,13-tricis isomers. In

124

7, l Ll S-tricis and all-cis, these signals are replaced by a doubled cis HOOP bands near

740-760 cm'. Table 3.6 shows the frequencies of the vibrational normal modes of the

polyene ehain of these four isomers.

11,13-dicis =etinal

7,11,13-tricis retinal

9,11,13-tricis retinal

all-cis retinal

Figure 3.13 FrIR spectra of the four labile isomers of retinal in the 1700-440 em"

region.

125

Table 3.6 FTIR frequencies of the double bond stretching modes, single bond

stretching modes and HOOP bands of the four retinal isomers containing the 11,13­

dicis geometry.

Descriptions 11,13- 7,11,13- 9,11,13- all-cis

dicis tricis tricis

C=O str. (cm') 1676 1679 1676 1677

C I I =CI2+CI3=C14 str. 1613 1615 1612 1616

(cm')

~=C8+C9=CIO str. 1580 1582 1595 1593

(cm')

C-C str. 1171 1170 1171 1170

(cm')

7+8 trans HOOP 966 - 964 -

(cm')

11+12 trans HOOP - - - -

(em")

7+8 cis HOOP - 740 - 743

(cm')

11+12 cis HOOP 761 760 759 965

(em:')

126

3.4 Discussion

3.4.1 Ground state properties of retinal isomers

The effect of temperature on the absorption spectrum of l l-cis-retinal was first

described by Wald and co-workers in 1959. The intensity of the Amax absorption band

increased markedly on cooling in contrast to the little or no band intensity changes of the

all-trans, 9-cis and 13-cis-retinals. More detailed measurements by Sperling and Rafferty

(Sperling et al., 1969; 1973) and by Becker and coworkers (Schaffer et al., 1974)

demonstrated that, in l l-cis retinal, the cis band decreased in intensity in proportion to

the increase in the Amax intensity, and that the temperature effect was more pronounced

in polar solvents.

An explaination for the temperature dependent phenomenon came from theoretical

calculations coupled with experimental absorption studies on l l-cis-retinal as a function

of temperature in that there was an equilibrium between l l-cis-Ll-s-cis and 11-cis-12-s­

trans type conformers (shown in Figure 3.13) at room temperature (Schaffer et al., 1974;

Becker et al., 1976). At low temperature (77 K) the 12-s-trans conformer predominated.

Other types of calculations and NMR studies also indicated that l l-cis retinal could exist

as these two conformation (Rowan et al., 1974). In the case of the 9-cis and 13-cis

retinals there is only a single isomer in each case, apparently twisting about 20° around

the adjacent single bonds (10-11 and 14-15 respectively) (Schaffer et al., 1974). So no

temperature effects of these unhindered isomers were observed.

127

.. ...

12-S-trans, 11-cis-retinal

~o 0#

12-S-cis, 11-cis-retinal

Figure 3.14 The equilibrium of 12-s-cis and 12-s-trans conformers of l l-cis-retinal.

The presence of an equilibrium between the 12-s-cis and 12-s-trans conformers,

with the latter conformer more populated at lower temperatures, provides the most

reasonable explanation for the temperature effect on the absorption spectrum of l l-cis

retinal.

Temperature effects on other retinal isomers and 12-methyl substituted analogs

have also been examined. For the highly hindered isomers (e.g. 7, 11, 13-tricis and all-cis­

retinal), their UVNis spectra at liquid nitrogen temperature showed no significant changes

of absorption bands (Trehan et al., 1990b). Hence, for these isomers, there are probably

no spectrally distinct conformers that equilibrate readily within the temperature range

128

studied. Possible role of conformers in photoisomerization will be discussed in a later

section.

For the unstable retinal isomers containing the twisted 11,13-dicis geometry, it

should be noted that no detectable temperature dependent absorption properties (Trehan

et al., 1990b) does not rule out the possibility that conformational equilibrium may play

a role in their thermal stability.

Based on a study of simple dienones and dienals, Lillya and Kluge (Kluge &

Lillya, 1977) postulated the consecutive 6e electrocyclization pathway for isomerization

of some of the dicis dienals and dienones. At room temperature, these compounds

isomerized from cis, cis to trans, cis-dienones, losing the cis configuration at the y,'fJ

bond. The isomerization is proposed to occur via an n-pyran intermediate which ring

opened to give the dienone with an isomerized, more stable double bond.

Stable ce-pyrans exist only in cases where their valence isomeric a, ~-cis-dienones

cannot exist in a planar form, thus forced into a severely twisted conformation (Kluge &

Lillya, 1977). Marvell et al. (1966) showed that cis-Ii-ionone exists as a minor form in

equilibrium with the corresponding e-pyran.

The diagram below (Figure 3.13) shows the proposed mechanism of 6-electron

cyclization, in which the stereospecific isomerization of cis, cis-dienones to trans, cis­

dienones occurs via an a-pyran intermediate by two consecutive 6e electrocyclization

reactions:

129

Figure 3.15 Mechanism for isomerization of dicis-dienals or dienones to the cis, trans

isomers (Kluge & Lillya, 1976).

The isomerization of those retinal isomers containing the 11,13-dicis geometry has

been proposed to proceed via a similar pathway (Kini et al., 1979; Knudsen et al., 1983).

The twisted 12-s-cis conformation of the chromophore orients the molecule in a way to

facilitate the 6e electrocyclization process. The resulting o-pyran intermediate opens

through another 6e electrocyclization process giving the more stable l l-trans, I3-cis

isomer. The disrotatory nature of the ring opening and ring closing procedures cause the

substituents at C-I1 to go through a 1800 rotation, changing the configuration of the

C11=C12 double bond from cis to trans. The complete conversion of the l l-cis to the 11-

trans geometry merely reflects the high energy content associated with the l l-cis

geometry.

130

Their UVNis absorption spectra reflect the highly sterically crowded nature of

these retinal isomers in the following manner: the absorption maxima are blue shifted to

287 nm for all-cis, 289 nm for 7,11, 13-tricis and 302 nm for both 9,11, 13-tricis and

11,13-dicis-retinal (Trehan et al., 1990). The chromophore must be highly twisted

chromophore, in agreement with the current FTIR data shown in section 3.3.2.2.

The activation parameters determined in this work for rearrangement of the four

unstable retinal isomers are consistent with the involvement of the Lillya-Kluge

mechanism for these rearrangements. In the case of all-cis-retinal, the detailed processes

can be formulated as below (Figure 3.14).

~_F\ /CHO

l.)lFFall-cis

7,9,13-tricis

t

~-ryl.)l r:o::-Jbis-S-cis

...bis-S-cis

Figure 3.16 Scheme of all-cis-retinal isomerization to 7,9,13-tricis-retinal.

131

The 6.Ht values (Table 3.5), a reflection of relative ease of bond formation

function and bond breaking show little difference between the four isomers. The negative

6.S~ values for all cases (Table 3.5) are consistent with the involvement of the highly

organized reaction intermediate (bis-S-cis conformer) necessary for the cyclization, which

must therefore be the rate determine step. All-cis is the most twisted isomer of these four

isomers so it should have the highest strain energy, making the conversion from s-trans

to bis-s-cis easier than the others; so its 6 st is less negative than those of the other

isomers. This is in agreement with the expected more twisted starting material making

the bis-Svcis conformer more accessible.

3.4.2 Photochemical properties of retinal isomers

3.4.2.1 Temperature effects on photoisomerization of retinal isomers

The photochemistry of retinal isomers under direct irradiation were investigated

only as a function of temperature. The purpose is to understand the low temperature

photochemical properties of the pigments to be discussed in the following chapter.

The initial product distributions of retinal isomers (ll-cis, 9-cis, 7-cis and 7,9­

dicis) were found to be temperature dependent in addition to that of all-trans-retinal

already in the literature (Waddell et al., 1977). In the cases of 7-cis, 9-cis and l l-cis­

retinal, isomerization of 13-trans to 13-cis versus isomerization of cis to trans (from 7-cis

to all-trans; 9-cis to all-trans; l l-cis to all-trans) decreased when temperature was

decreased. This variation could be derived from the fact that the excited-state energy

132

barriers (Eo) of different double bonds of retinal isomers are different. These

isomerization results show that energy barriers of excited-state trans to cis isomerization

at the C13=CI4 double bond are higher than those of cis to trans at C7=Cg, C9=C IO and

Cl1=C I2 double bonds. The slopes of the curves in Figure 3.5 indicate that energy barriers

of 7-cis to trans is the smallest, 9-cis to trans relatively large while l l-cis to trans ranks

in the middle.

Excited state bond order could be an important kinetic parameter in determining

the direction of isomerization. Calculated results (Table 3.4) show the correct trend,

namely the lowest excited state bond order was found to be the C13=CI4 bond. However,

we suspect that relief of steric crowding probably plays an independent important role,

making the 7-cis bond the most labile and l l-cis the second, because, both 7-cis and 11­

cis-retinal are hindered isomers which are sterically crowded at C7=Cg or Cl1=C I2 double

bond. Steric relief provides another driving force to facilitate isomerization at these

double bonds.

The experimental results of 7,9-dicis-retinal show that the product of one-photon­

two-bond isomerization (i.e. all-trans) is temperature independent, which suggests a

different mechanism is involved in formation of this product. One-bond isomerization

is a volume demanding process while two-bond isomerization is more concerted process

which is favored in the restricted environment, for example, in the binding cavity of the

visual pigment (see chapter 4) In solution, there two processes are competitive favoring

the one bond process at the room temperature (Ganapathy & Liu, I992a).

133

A more quantitative way of expressing initial product ratio is quantum yield of

isomerization. Temperature effects on the quantum yield values of all-trans-retinal are

in the literature (Waddell et al., 1981b). In a nonpolar solvent, 13-cis and 9-cis-retinal

are the two primary photoproducts. The 13-trans to 13-cis isomerization process has a

higher quantum yield value than that to 9-cis and their temperature behavior gave a lower

energy barrier for 13-cis (2.1 kcal/mol) than that (3.2 kcal/mol) for 9-cis.

The quantum yield values for photoisomerization of 7-cis and 7,9-dicis-retinal

determined at 193 K and 298 K (shown in Table 3.3) showed a similar temperature

behavior, i.e. decreasing as temperature is lowered. In contrast, for l l-cis retinal,

anomalous quantum efficiencies of photoisomerization have been reported: i.e. quantum

yield increasing at low temperature (Kropf & Hubbard, 1970). This has been interpreted

as the noted conformational effect, shifting toward the 12-s-trans conformer as

temperature decreased (Birge et al., 1975; 1976). Thus, the temperature dependent

quantum yield data for the crowded 7-cis and 7,9-dicis isomers are in agreement with

their UVNis absorption spectra (absence of a temperature effect, see above) again may

suggesting predominantly the presence of one conformer in solution.

It is interesting to compare the above conformational effects with those observed

for 9-cis and l l-cis-retinyl chromophore when they are combined with opsin. The cis to

trans photoisomerization quantum efficiency of 9-cis-retinal is 0.5 and that of 9-cis­

rhodopsin is 0.2 (Kropf & HUbbard, 1970). This decrease is to be expected since the

pigment matrix could readily increase the barrier to isomerization of the chromophore

now trapped in a highly specific binding pocket. The quantum efficiency for l l-cis

134

retinal, instead increases form 0.2 to 0.67 (Dartnall, 1968) upon incorporation into

rhodopsin. Many suggestions have been made to account for the increased efficiency.

Among these is the suggestion that the 12-s-trans geometry is the conformation of the

visual chromophore in rhodopsin since this conformer is expected to have a lower barrier

to photoisomerization (Birge et al., 1975; 1976). Consequently, the increase in l l-cis­

retinal quantum efficiency upon either lowering solution temperature or incorporation into

the pigment system was suggested to be associated with similar conformational effects.

3.4.2.2 Wavelength dependent studies of photoisomerization

The quantum yields and ratio of photo products of all-trans-retinal in both polar'

and nonpolar solvents have been determined to be independent of the excitation

wavelength (Waddell et al., 1977).

Experimental data on l l-cis and 7,9-dicis-retinal also demonstrate that the initial

photochemical product ratios upon excitation at 340 to 420 nm are indistinguishable from

those on the excitation wavelength 360 nm. In the case of 7,9-dicis-retinal, this result

reflects the fact that the compound exists predominantly as a single conformer. For 11­

cis-retinal, the result is probably reflecting that its two conformers have similar

photochemical properties.

Wavelength dependent photoisomerization, however, can be a rather complicated

phenomenon. Thus, the quantum efficiency for photoisomerization of l l-cis and all-trans­

retinylidene compounds varies depending on the wavelength of the exciting light (Becker

& Freedman, 1985; 1986). Higher Isomerization yields from the l l-cis to the all-trans

135

configuration occurs at longer wavelengths, whereas the isomerization of the all-trans

isomer prefers shorter wavelengths. Singlet to triplet state crossover from certain

vibrational levels has been proposed to cause the wavelength dependency (Becker, 1988).

But the wavelength dependence remains to be explained more quantitatively. It is to be

noted that the quantum efficiency for isomerization of the rhodopsin chromophore is

independent of the wavelength of irradiating light.

In summary, the temperature and excitation wavelength effects on the

photochemistry of several retinal isomers have been discussed in this chapter. Also

spectroscopic properties of the ground state of hindered retinal isomers have been

examined and their labile isomerization mechanism has been confirmed with more

accurate kinetic parameters.

136

Chapter 4

Photochemical Studies of Rhodopsin Analogs at Low Temperatures

4.1 Introduction

The photoreaction of rhodopsin is very fast at physiological temperature. The

conversion of rhodopsin to bathorhodopsin takes place in a few picoseconds. Because

such a fast photoreaction is very difficult to study at room temperature, many

investigations have been carried out at low temperatures, where the thermal reaction rate

of intermediates ofrhodopsin is remarkably reduced (Yoshizawa, 1972). Low temperature

spectrophotometry has been used to investigate the photochemical reactions of rhodopsin

since the 1950s (Wald et al., 1950; Hubbard & Kropf, 1958; Yoshizawa & Wald, 1963).

The method has a big advantage in being able to measure the spectra precisely and being

able to manipulate with conventional spectrometry. But there are some disadvantages in

which irradiation of rhodopsin at very low temperature may produce different

photochemistry from those initiated at physiological temperature.

Time-resolved laser photolysis had been used to study the photochemical reaction

of rhodopsin since 1972, when Busch et al. first applied picosecond laser photolysis for

the detection of bathorhodopsin at physiological temperature. Since then several groups

have investigated whether any intermediates are present prior to bathorhodopsin.

However, it has recognized that intense laser excitation of rhodopsin easily produces a

137

multi-photon reaction because of extraordinarily high photosensitivity of rhodopsin.

Shichida et at. (1984) observed photorhodopsin upon excitation of cattle rhodopsin with

a weak laser pulse. It decayed to bathorhodopsin at 45 ps. Recently, the femto second

laser photolysis has been used to study the photochemistry of rhodopsin (Schoenlein et

al., 1991).

Bathorhodopsin with an all-trans chromophore was the first intermediate observed

from rhodopsin (Ll-cis) and isorhodopsin (9-cis) shown by both low temperature and time

resolved spectroscopic studies (Yoshizawa, 1972; Shichida et al., 1984). The

photoisomerization of the chromophore is completed within a pico-second (Shichida et

al., 1984), a process too fast to be accompanied by conformation changes of opsin. Thus,

relaxation of the initially formed trans species is likely to be inhibited by a steric

interaction with the surrounding protein residues. This would result in the formation of

a distorted all-trans form. This has been confirmed by a direct measurement of distortion

of the chromophore of bathorhodopsin by the use of resonance Raman spectroscopy

(Oseroff & Callender, 1974; Eyring et al., 1982). In addition, resonance Raman

spectroscopy proved the chromophore of bathorhodopsin to be a protonated retinylidene

Schiff base like that of rhodopsin (Oseroff & Callender, 1974; Eyring et al., 1979; Aton

et al.. 1980). Bathorhodopsin decays through a series of intermediates that lead to the

eventual release of all-trans-retinal from the protein.

Investigations using various spectroscopies led to the conclusion that relaxation

of the strained chromophore of bathorhodopsin is through changes in the chromophore­

opsin interaction near the cyclohexenyl ring binding site (Shichida, 1986). Recently

138

research by Yoshizawa and his coworkers (Shichida et al., 1991) indicate that the

difference in the configuration of the original chromophore between 7-cis and l l-cis

rhodopsin is sufficient to cause different chromophore-opsin interactions in the

bathorhodopsin and lumirhodopsin stages of the two pigments. At the meta I stage, the

difference has disappeared probably following relaxation of the protein near the

chromophores.

The interest in the photochemistry of rhodopsin analogues containing the dicis

geometry was prompted by two postulated models for the photochemical process of the

visual chromophore: "Bicycle pedal" (Warshel, 1976) and "Hula twist" model (Liu &

Asato, 1985). Highly hindered 7,9-dicis-rhodopsin has a greatly blue-shifted absorption

maximum ( Amax=445 nm shown in Figure 4.2) from that of rhodopsin (498 nm) and close

to the other hindered 7-cis-rhodopsin (450 nm). Furthermore, it was shown that the final

photobleaching product of 7,9-dicis-rhodopsin is also all-trans-retinal and opsin (Trehan

et al., 1990). 9, l l-Dicis-rhodopsin pigment retained only 40% original geometry with

60% the 9-cis isomerization product, according to the HPLC extraction result (Trehan et

ai., 1990). However, 9,I1-dicis geometry can be retained by adding 12-F-substituent on

the chromophore (Colmenares & Liu, 1992). Photobleaching of 9,11-dicis-12­

fluororhodopsin also gives all-trans as its photoproduct. The intriguing question is that

within the restricted binding pocket what will be the altered mode (if any) of

photoisomerization for these isomeric pigment analogs. It will be of interest to determine

in each case whether the photobleaching process undergoes two steps isomerization

process via mono-cis geometry, the latter by way of rhodopsin or 9-cis-rhodopsin analogs.

139

We have therefore carried out the studies of the photochemical reaction of 7,9-dicis­

rhodopsin and 9,II-dicis-12-fluororhodopsin.

To carry out photochemical studies of pigment analysis, a variety of synthetic

detergents are available for extraction and purification of pigments (Fong et al., 1982;

Kropf, 1982), containing either hydrophobic or hydrophilic structures. Detergent

molecules in aqueous medium associate to form a micelle. The binding force for a

micelle is interactions between the hydrophobic structures. Rhodopsin-detergent micelles

are hydrated by water molecules and remain "solubilized". The outer segment of a rod

visual cell, in which rhodopsin is localized, is readily detached from the inner segment

by shaking of the retinas in buffer and can be purified by centrifugation in a gradient of

sucrose (Knowles & Dartnall, 1977). If necessary, rhodopsin can be further purified by

various chromatographic methods (review see Shichi, 1983). Rhodopsin analogs can be

regenerated in detergent by incubating the bleached pigment with retinal analogs.

To assist in determining the polyene configuration in a pigment analogs and/or the

extent of isomerization of the chromophore during binding interaction, the analytical

method combining extraction of retinyl chromophores and analysis of the resulting retinal

isomers (Groenendijk et aI., 1979; 1980) will be used. This method has been successfully

used for the identification of the chromophore from some retinal bound pigments, i.e.

porphyropsins (Suzuki & Makino-Tasaka, 1983), bacteriorhodopsin (Groenendijk et al.,

1980), retinochrome (Hara et al., 1977) and insect visual pigments (Seki et al., 1987).

Also this method has been applied to study the extent of configuration retention of

rhodopsin isomers derived from dicis-retinal isomers (Trehan et al., 1990a).

140

The goal of this study is to characterize the mechanism of photobleaching reaction

of two dicis-rhodopsin systems (7,9-dicis and 9, 11-dicis-12-fluororhodopsin analogs) and

study the interaction of isomeric retinylidene chromophore and opsin during their

photobleaching processes.

141

4.2 Experiments

4.2.1 Procedure for analogs of bovine rhodopsin

4.2.1.1 Preparation of bovine opsin

The procedure for isolation of bovine opsin and preparation of rhodopsin analogs

is shown in Figure 4.1. The isolation of rod outer segments (ROS) from bovine retinas

was carried out at 4°C in the cold room and with dim light. The thawed retinas (4 vials

of 50 each obtained from Hormel Company) were homogenized in 200 mL chilled 34%

sucrose containing 10 mM HEPES (pH 7.0), 65 mM NaCl and 2 mM MgCI2. The

supernatant fraction was diluted with 3 volumes of HEPES buffer and centrifuged under'

vacuum at 7,000 rpm using a Beckman rotor type 27 for 15 min to yield a pellet (crude

ROS).

The pellets are homogenized with 90 mL HEPES buffer using a homogenizer

(speed 60) for I min. The suspension was divided into six 15 mL fractions and floated

by syringe on chilled 34% sucrose solution. The tubes were centrifuged using rotor

buckets under vacuum at 25,000 rpm for 45 min. The middle orange band containing

ROS was collected. ROS was bleached in the presence of 1 mL of 1 M NH20 H by

illumination with an orange light (>540 nm) at OoC (3-73 cut-off filter) for -45 min.

Excess hydroxylamine was removed by repeated washings with HEPES buffer (5 times)

and with distilled water (one time) by centrifugation (15,000 rpm under vacuum for 15

min). The ROS were lyophilized, followed by washing 8-10 times with cold hexane until

retinal oxime was completely removed (monitored by UV). The purified ROS thus

142

obtained were suspended in 10 mM HEPES buffer (pH 7.0) and divided into several

fractions and stored in the freezer until use.

143

Pelletdiscard

tSupernatant

1. Dilute 3 times with buffer2. Centrifuge 7,000rpm, 15 min

200 frozen and thawed retinasin 34 % buffered sucrose

IShake/strain through stocking filter,....------"--------.,.Filtrate

ICentrifuge 3,000rpm,

•

tMembrane, etc.discard

Pellets (crude ROS)

1

1. Resuspend & homogenize in buffer2. Density gradient. 34 % sucrose3. Centrifuge, 25,000 rpm, 45 min

Middle layer (ROS)

1.Add 1 ml 1 M NH20H, photobleach for 30 min2. Wash w/Hepes buffer 5 times, w/H 20 1 time3. Centrifuge, 10,000 rpm, 15 min

Supernatantdiscard

Supernatantdiscard

Pellet (opsin)

j1. Lyophilizate2. Wash with cold hexane 6 times3. Vacuum pump to dry

Dry pellet (purified opsin)

1. Resuspend & homogenize in buffer

2. Add retinal analogs in ethanol solution

3. Incubate in the dark at 5° C for 20 h

4. Centrifuge, 10,000 rpm, 20 min

tPellet (Rhodopsin analogs)

j1. Remove excess retinal by hexane washings .2. Iyophilizate .3. Extract with 2% digitonin in buffer, stir In cold for 2 h.4. Centrifuge, 25,000 rpm, 25 min.

Supernatant (Rhodopsin analogs in 2% digtonin)

tSupernatantdiscard

Figure 4.2 The flow chart of preparation of rhodopsin analogs.

144

4.2.1.2 Preparation of 7,9-dicis-rhodopsin

7,9-Dicis-rhodopsin was prepared by incubation of an opsin suspension with a 2­

fold molar excess of 7,9-dicis-retinal dissolved in a small amount of ethanol at SoC for

about 20 h. Because 7,9-dicis-rhodopsin is very unstable in the presence of the

hydroxylamine at room temperature (see Figure 4.4), hydroxylamine was not added to the

7,9-dicis-rhodopsin preparation. After incubation in the dark for 20 h, the sample was

washed 6 times with 10 mM HEPES buffer (pH 7.0), followed by lyophilization and

washing lO-times with cold hexane by centrifugation to remove excess retinal. The pellet

obtained was dried under a stream of nitrogen gas, and 7,9-dicis-rhodopsin contained in

the pellet was extracted with 2% digitonin dissolved in 10 mM HEPES buffer (pH 7.0), .

stirring in an ice bath for about 2 h.

4.2.1.3 Preparation of 9,1l-dicis-12-fluororhodopsin

9, l l-Dicis-Iz-fluororhodopsin was prepared by incubation of an opsin suspension

for about 20 h with a I.5-fold molar excess of 9,1l-dicis-12-fluororetinal which was

dissolved in a small amount of ethanol. The sample was kept at 5°C because this

rhodopsin analog is unstable at room temperature and thermally isomerized to its 9-cis

isomer. After formation of 9,II-dicis-12-fluororhodopsin, a neutralized solution of

hydroxylamine (I M) was added to the rhodopsin preparation to a final concentration of

10 mM to change the unreacted retinal into its oxime. Then the sample was washed 6

times with 10 mM HEPES buffer (pH 7.0), followed by lyophilization and washing six

145

times with cold hexane by centrifugation to remove retinal oximes. The pellet obtained

was dried under a stream of nitrogen gas. 9, ll-Dicis-12-fluororhodopsin contained in the

pellet was extracted with Z % digitonin dissolved in 10 mM HEPES buffer (pH 7.0) at

about OoC for 2 h and stored at low temperature until further use.

4.2.2 Low-temperature UV/Vis spectrometry

The following is a general procedure for measuring the absorption spectra at low

temperatures which was adapted from that described in the literature (Shichida &

Yoshizawa, 1972). A cryostat with quartz windows from Oxford Instrumentation

Company was used (shown in Figure 4.2). A visual pigment sample was introduced into'

a detachable sample cell, fitted within sample cell holder. The latter was attached to the

copper tube at the bottom of the cold finger by a screw. The cold finger filled with liquid

nitrogen was placed in the metal jacket with the sample cell holder. The cryostat

chamber was then evacuated by a rotary pump. The temperature of the sample was

monitored with a copper-constantan thermocouple attached to the sample cell holder. The

sample temperature could be kept at a constant temperature within ± z'c.

For photochemical reactions at low temperature, the sample was irradiated with

a light introduced by fiber optics from a projector lamp (750 W) with an interference

filter or a cut-off filter. A 5-cm water layer was placed in front of the lamp as a heat

filter. To minimize complication from light scattering when working with glass samples,

opal glass (as suggested by Yoshizawa, 1972) was placed in the path of both the sample

and reference beams of the spectrophotometer.

146

TOP PlATE ALSO INCLUDES

[NNEH EVACU,\TION ~VAl.VE ,\Nn IO-PINSEAL I

couur xtn EVi\CU,\TIONi\ND SAfETY VALVE

-++4--- LIQUID NITROGENflESEHVOIR

GAS EXIIAUST VALVE

u

lI,! IiiI:ii

III

l Ji

!O,'lmm d 1 a. i-------'1

SA,\lI'LE .HTI';SS I'()I{'I'

NI THOGEN EN'I'I{YI'OIlTS

SOHU PUMP

EEo<l'

I=.l--+l,----- liEAT EXC IIi\NGERTEHPERATUnE SENSORAND HEATER

EXCIIANGE

DEMOUNTAULE OUTERVACUUM CASE WINDOWS

Figure 4.2 Diagram of optical cryostat for measuring absorption spectrum at liquid

nitrogen temperature or above (from Oxford Instruments).

147

For low-temperature spectrophotometry, stock solutions of neutralized

hydroxylamine (l M) and glycerol were added to the extracts to a final concentration of

10 mM and 66 %, respectively. This extract mixed with glycerol in a final concentration

of 66% can be frozen to form a clear glass without cracks above about -lODGe, but below

this temperature some cracks which cause multiple reflection of the measuring light are

usually formed in the mixture (Tokunaga et al., 1976; Yoshizawa et al., 1963). This

method has been improved by applying a rapid cooling technique (Horiuchi et al., 1980),

with which a clear glass without any cracks can be prepared at liquid nitrogen

temperature or below. In our case, a quartz optical cell in a cell holder was attached to

the cold finger and then immersed in liquid nitrogen. Immediately after removal from

the liquid nitrogen, the optical cell was filled with the rhodopsin preparation by rapid

transfer with a syringe. The cold finger with the sample was again immersed into liquid

nitrogen. Quickly the cryostat was assembled and the chamber evacuated. The UV/Vis

spectrum was recorded at liquid nitrogen temperature. For 9, 11-dicis-12-fluororhodopsin,

hydroxylamine was added to the sample so as to diminish any absorption contributed by

the random Schiff bases, which may be produced from irradiation of 9,11-dicis-12­

fluororhodopsin in the sample. After adding hydroxylamine, 9,11-dicis-12­

fluororhodopsin was carefully kept at DoC. 7,9-Dicis-rhodopsin was found to be very

unstable against the hydroxylamine, so hydroxylamine was not added.

148

4.2.3 HPLC chromophore extraction analysis

Procedures for denaturation of the pigment analogs and extraction of the

chromophores in the form of retinal were the same as those described in the literature

(Groenendijk et al., 1979; 1980) with a slight modification. Namely, to a 1 mL solution

of rhodopsin or rhodopsin analogs (concentration around 1 x 10-4 M) in 2% digitonin was

added 100 ul EtOH (absolute) to denature the pigment. Cold hexane solvent was then

added to extract the retinal from the aqueous layer. The organic layer which contain most

retinal was separated from aqueous layer by centrifugation and collected. The exacted

solution was passed through a silica gel column and dried under vacuum. The final

solutions were analyzed by HPLC (Rainin Instrument Co. HPLC system) using a 25 x .

0.46 em DynamaxMicrosorb Si 60 (Sum) column and 4-5% ethyl ether in hexane as

solvent. The UV detector wavelength was set at 360 nm. The different extinction

coefficients of the various retinal isomers (Zhu et al., 1992) were taken into consideration

for calculation of isomer compositions at 360 nm.

The extraction yields ranged between 50 to 60%. Completed extraction of the

chromophoric group requires prolonged or repeated treatments at higher temperatures but

the extent of isomerization becomes unacceptable.

149

4.3 Results

4.3.1 Effects of NH20H on dicis-rhodopsins

The UVNis spectrum of 7,9-dicis-rhodopsin (Amax = 444 nm) in 2% digitonin

solution is shown in Figure 4.3. 7,9-Dicis-rhodopsin was found to be unstable in NH20H

solution at room temperature. In Figure 4.4 is shown the UVNis spectrum of

decomposition of 7,9-dicis-rhodopsin in the 100 mM NH20H solution, presumably into

7,9-dicis-retinal oxime and opsin. Figure 4.5 shows a time plot of the degradation of 7,9­

dicis-rhodopsin in hydroxylamine (lOa mM) at room temperature. The absorbance loss,

due to the decomposition of 7,9-dicis-rhodopsin by NH20H was monitored at 440 nm.

The rate of decomposition at room temperature is very fast with a half-life approximate

by 18 min. Hence, for the low-temperature spectroscopic experiments described here,

NH20H was not added during the preparation sample of 7,9-dicis-rhodopsin.

Absorption spectrum of 9, I1-dicis-12-fluororhodopsin in 2% digitonin extracted

solution is shown in Figure 4.6 with an absorption maximum at 484 nm. This rhodopsin

analog was found to be relatively stable in NH20H solution at room temperature, so for

low-temperature spectrophotometry, both neutralized hydroxylamine and glycerol were

added to the extracts to make final concentrations of 10 mM and 66%, respectively.

150

1.9430

1.1515415

1.115152

0.7778

0.3894

0.0010

t8

I..2150.0 300.0 3150.0 400.0 400.0 1500.0 15150.0 600.0 6150.0 700.0

nil

Figure 4.3 Absorption spectrum of 7,9-dicis-rhodopsin in 2% digitonin solution

(An.ax =444 nm).

151

.80

.60

.20

.00

300 400WAVELENGTH (nm)

500

Figure 4.4 7,9-Dicis-rhodopsin in 100 mM NH20H solution at room temperature,

at following time delays: 0, 5, 10, 15, 20, 25, 35, 60, 120 min.

152

E 100c

0 R"2 = 0.998'<:t'<:t-C'O

Q)oeC'O.c...0en.cC'O

Q)

>-C'OQ) 10ex:

0 1 0 20 30 40 50 60

Incubation time (min)

Figure 4.5 Degradation of 7,9-dicis-rhodopsin in 100 mM NH20H at room temperature

as revealed by a decrease of absorbance at 440 om.

153

0.liU80

0.7323

0.e48e

A

0.3648

0.1810

-0.0027

aso.0 300. 0 3150.0 400.0 4eO. 0 1500. 0 eec.0 600.0 6eO. 0 700.0n.

Figure 4.6 Absorption spectrum (!..max =484 nrn) of 9,11-dicis-12-fluororhodopsin in 2%

digitonin solution.

154

4.3.2 Photochemical reactions of dicis-rhodopsins at liquid nitrogen temperature

(-190°C)

7,9-Dicis-rhodopsin: A 7,9-dicis-rhodopsin-66% glycerol sample was cooled to

liquid nitrogen temperature by the rapid cooling technique to minimize cracks in the

sample. Its absorption spectrum corresponds to curve 1 in Figure 4.7. Irradiation of 7,9-

dicis-rhodopsin with blue light at 437 nm at liquid nitrogen temperature causes a

photochemical reaction which produced a red shift in the pigment absorption. Spectral

changes as a result of the photochemical reactions of 7,9-dicis-rhodopsin at liquid nitrogen

temperature are shown in Figure 4.7. Irradiation was continued for 8 h, resulting in

formation of a mixture containing mainly the bathointermediate of 7,9-dicis-rhodopsin .

(curve 14 in Figure 4.7). The relative absorbance of 7,9-dicis-rhodopsin at 440 nm versus

irradiation time is plotted in the insert of Figurea.Z.

In principle, absorption spectrum of the bathoproduct can be obtained from the

spectra before and after warming of an irradiated sample (Yoshizawa & Wald, 1963).

But reproducible spectra are difficult to obtain during thawing and refreezing of the

rhodopsin sample (probably due to varying degree of glass cracking). A much simpler

and more accurate method which does not involve warming of the samples has been used

to obtain at least a partial spectrum of the bathoproduct of rhodopsin (Shichida et al.,

1991). This entails measuring the difference spectrum between a sample containing the

bathoproduct and other rhodopsin isomers which is the same sample after irradiation with

light absorbed only by the bathoproduct.

155

The mixture (curve 14 in Figure 4.7, replotted as curve 2 in Figure 4.8) was then

irradiated with red light (>610 nm) to convert the batho intermediate in the mixture to

presumably l l-cis- and 9-cis-rhodopsin (curve 3 in Figure 4.8 and curve 1 in Figure 4.7

replot as curve I in Figure 4.8). The spectra shifted to shorter wavelengths with an

isosbestic point at 513 nm (Figure 4.8) and it reached another stationary state (curve 3

in Figure 4.8). The difference spectrum before and after irradiation with the red light was

obtained (curve 1 in the insert of Figure 4.8) and compared with that from similar

experiments using l l-cis-rhodopsin (curve 2 in the insert of Figure 4.8). The difference

absorption maximum of the batho intermediate produced from 7,9-dicis-rhodopsin is blue

shifted from that of bathorhodopsin produced from l l-cis-rhodopsin.

The components of photoproducts of 7,9-dicis-rhodopsin were readily

interconvertible by light at -190°C. When the mixture (curve 3 in Figure 4.8) was again

irradiated with the 437 nm light, the spectrum (curve 4 in figure 4.8) red-shifted to be

identical with that of curve 2 in Figure 4.8.

Irradiation of 7,9-dicis-rhodopsin with 437 nm light resulted in photoreversible

reactions among the batho intermediate and l l-cis and 9-cis pigments produced from 7,9­

dicis-rhodopsin via the batho intermediate. This was shown by results of the following

experiments. The mixture of l l-cis and 9-cis pigments was reirradiated with 437 nm

light to form the mixture containing mainly the bathointennediate whose spectrum was

identical with that produced earlier after irradiation in a similar manner. Then the batho

intermediate in the mixture was irradiated with red light to convert it to l l-cis and 9-cis

pigments. The difference spectrum from before and after red light irradiation was

156

identical to that of curve 2 in the insert of Figure 4.8. These facts suggested that the 11­

cis and 9-cis pigments produced from 7,9-dicis-rhodopsin are different from those in the

regenerated l l-cis- and 9-cis-rhodopsin samples. If II-cis and 9-cis pigments produced

form the batho intermediate of 7-cis-rhodopsin were identical with those contained in the

l l-cis-rhodopsin sample, the same difference spectrum as that from the l l-cis-rhodopsin

would be obtained.

157

600500WAVELENGTH (om)

400.00 -+-----r-----,r-----r------,...-----,r------t

100

.36

10

.24

CI'i 1.D 0 100 200m~

.12

Figure 4.7 Photochemistry of 7,9-dicis-rhodopsin at liquid nitrogen temperature. A

sample of the pigment was irradiated with 437 nm light with the spectra recorded after

the following intervals of irradiation: 0, 4, 8, 16, 32,64, 90, 110, 130, 160,220, 300 min.

Insert. A log plot of decrease of 7,9-dicis pigment absorption at 440 nm upon 437 nm

irradiation versus time.

158

600 nm500

.68

-.70 +--,---.----.------,.------,---11

-.02

.24

.36

.12

.00400 500

WA VELENGTH (om)600

Figure 4.8 Photochemistry of isomeric rhodopsin pigments. The 7,9-dicis rhodopsin

(curve 1, also curve 1 in Figure 4.7) at liquid nitrogen temperature, after irradiation with

437 nm light (curve 2, also last curve in Figure 4.7), followed by prolonged irradiation

with red light (>610 nm) (curve 3) and then prolonged irradiation with 437 nm light

(curve 4). Insert: difference spectra. Curve 1: The curve 3 in Figure 4.8 minus the curve

4 in the same figure; curve 2: from the corresponding spectra of a rhodopsin sample.

159

9,1 l·dicis· 12.fluororhodopsin: Upon irradiation of 9, 11-dicis-12-fluororhodopsin

with light at 437 nm at -190°C, a red shift of the absorption spectrum was immediately

detected (Figure 4.9), indicating the formation of a bathoproduct. Prolonged irradiation

yielded a photostationary state with the absorption maximum at 505 nm (curve 8 in

Figure 4.9). The relative absorbance of 9,11-dicis~12-fluororhodopsinat 500 nm versus

time is plotted in the insert of Figure 4.9. Then this photostationary state mixture

(replotted as curve 2 in Figure 4.10) was irradiated with light at wavelengths longer than

610 nm. The spectrum shifted to a shorter wavelength to reach another photostationary

state with the absorption maximum at 495 nm (curve 3 in Figure 4.10). The process was

found to be completely reversible in that reirradiation of the later photostationary state

mixture (curve 3 in Figure 4.10) with 437 nm light yielded the identical spectrum (curve

4 in Figure 4.10) with that of curve 2 in Figure 4.10. Such spectral changes are

reminiscent of the curves recorded in the study of interconversions between

bathorhodopsin, rhodopsin and isorhodopsin (Yoshizawa & Wald, 1963). While the

interconversion of photostationary mixtures shown in Figure 4.10 were found completely

reversible, there was no evidence that the mixture could be converted back to the dicis

isomers.

160

100

•.108

·072

.036

10 ­o 5 10 15m

.000 ..J-......----.,r------r-----r~--_r_---,._--_;

450 550WAVELENGTH (om)

650

Figure 4.9 Photochemistry of 9,l1-dicis-12-fluororhodopsin at liquid nitrogen

temperature. A sample of 9, ll-dicis-12-fluororhodopsin was irradiated with blue light

(437 nm) with the spectra recorded after the following intervals of irradiation: 0, 1, 3, 5,

8, 10, 15, 20 min. Insert: a log plot of disappearance of pigment absorption at 500 nm

versus time.

161

.120

.096

.072en

,Q

<.048

.024

.000

400

1.00

500 600WAVELENGTH (nm)

600 nm

Figure 4.10 Photochemistry of isomeric l2-fluororhodopsin pigments. The 9,1l-dicis­

l2-fluororhodopsin (curve 1 and also curve 1 in Figure 4.9) at liquid nitrogen temperature

was irradiated with 437 nm light (curve 2 and also last curve in Figure 4.9), followed by

prolonged irradiation with >610 nm light (curve 3) and then prolonged irradiation with

437 nm light (curve 4). Insert: difference spectra of l2-fluoro-bathointennediate and its

isomer. Curve 1, curve 3 in Figure 4.10 minus curve 4 in the same figure; curve 2, the

corresponding difference spectrum from irradiation of 9-cis-l2-fluororhodopsin.

162

4.3.3 HPLC extraction analysis of chromophores of dicis-rhodopsins

The HPLC chromatograms of the extracted chromophore from the 7,9-dicis­

rhodopsin sample irradiated with 437 nm light for different irradiation times at liquid

nitrogen temperature are shown in Figure 4.11. The peaks have been identified as 7,9­

dicis-, all-trans-, l l-cis- and 9-cis-retinal by coinjection with the corresponding isomers

(Figure 4.12). 13-Cis isomer of the original chromophore is the product of a dark

reaction during the extraction procedure. Progress of the isomerization during the

irradiation process was monitored by HPLC extraction analysis. The data were plotted

and shown in Figure 4.13. The absence of substantial amounts of the two mono-cis

products suggests that direct isomerization from 7,9-dicis to all-trans took place during .

irradiation.

In order to elucidate the photochemical reaction mechanism of 12-F-9,II-dicis­

rhodopsin during the photobleaching process, analyses of the extracted chromophore of

9,II-dicis-12-fluororhodopsin by HPLC have been performed. Figure 4.14 shows the

HPLC patterns of retinal isomers extracted from the irradiation samples of 9, l l-dicis-l Z­

fluororhodopsin at different irradiation times at liquid nitrogen temperature. The peaks

corresponding to 9,II-dicis, all-trans, 9-cis, and l l-cis-l Zsfluororetinal were identified by

coinjection of corresponding isomers (Figure 4.15) and by their UVNis absorption

spectra. The small peak between the 9-cis and all-trans isomers during the latter stage

of irradiation is suspected to be the 13-cis isomer based on its retention time. The

amounts available were too small for further characterization. The percentage of retinal

isomers in these extracts are plotted in Figure 4.16 versus irradiation time. Particularly

163

noteworthy is the amount of the 9-cis isomer. It was larger at early period of irradiation

than latter stages. This indicates that the process of conversion of 9,II-dicis-12­

fluororhodopsin to its all-trans batho-intermediate takes place via its 9-cis isomer.

164

a) 1=0 min

JI--J7,9-dicis

b) 1=10 min

11-cis s-etsJI---J ~ -..J'OJ

all-trans

c) t=40 min7,9-dicis

all-trans

11-cis s-ols

JJall-trans

d) 1=120 min

7,9-dicis

11-cis s-ets

Jf

Figure 4.11 HPLC chromatograms of chromophore mixture extracted from 7,9-dicis­

rhodopsin after varying periods of irradiation with 437 nm light. The minor peak at t=O

corresponds to that of a 13-cis isomer. However, all isomers with the 13-cis geometry

have nearly identical retention times for unambiguous assignment.

165

a)7,9-dicis

all-trans

13-cis /\11-cis 9-cis

If .

7-cisb)

9-cis 7,9-dicis all-trans13-cis

11-cis

Ifc)

11-cis

7,9-dicis all-trans

13-cis /\If

Figure 4.12 HPLCchromatograms of retinal isomers obtained from denaturing the batho­

intermediate of7,9-dicis rhodopsin. a) original extraction, b) coinjected with 7-cis

and 9-cis isomers, c) coinjected with l l-cis isomer.

166

100

7,9-dicis80...

0:J"'C all-trans0 60~

a.0...0

40J:a.~0 20

9-ci s 11 -cis

00 50 100

Irradiation Time (min)

Figure 4.13 Time plot showing changes in isomeric composition of the chromophore

during irradiation of 7,9-dicis-rhodopsin at -190°C, data from Figure 4.11.

167

a) t=O 9, t t-dicis

e) t=120 min all-trans

t t-crs s-ets

Figure 4.14 HPLC chromatograms of the chromophore extracts obtained during the

irradiation of a sample of 9, ll-dicis-12-fluororhodopsin with 437 nm light. The

major peaks were verified through coinjection with authentic samples.

168

a) 9.11-dicis

9-cis

all-trans

t i-cis

IJ~

b)

11-cis 9-cis

9,11-dicisall-trans

If 1..--_..... -..1 \"----

Figure 4.15 HPLC chromatograms of extracted chromophore of irradiated sample 9,11­

dicis-12-fluororhodopsin. a) original extract, b) coinjected with ll-cis and 9-cis

isomers.

169

11-cis

all-trans

200

Irradiation Time (min)

1009,11-dicis

80-CJ~

"C 600I-

a.0

40-0.ca..

20~0

0

-200 100

Figure 4.16 Progress of irradiation of 9,II-dicis-12-fluororhodopsin at -19<fC as

revealed by HPLC analysis of chromophore extracts in Figure 4.14.

170

4.3.4 Photosensitivity of dicis-rhodopsins relative to mono-cis rhodopsins

The procedure for measurement of the photosensitivities of visual pigments was

established by Dartnall et al (1968). The bleaching of visual pigments can be expressed

as a unimolecular reaction. Therefore, when the proportions of the residual pigment in

a sample after successive irradiations are plotted on a logarithmic scale against the

numbers of photons of irradiating light, the points yield a straight line, the slope of which

represents the relative photosensitivity of the sample.

By following the rates of disappearance of pigments to form bathoproducts in

samples containing 7,9-dicis-rhodopsin and 7-cis-rhodopsin, we have obtained qualitative

information relating the photosensitivity (relative quantum yield) of 7,9-dicis pigments to .

those of 7-cis-rhodopsin. 7,9-Dicis-rhodopsin was irradiated at -190°C (liquid nitrogen

temperature) with 437 nm light, and progress of the reaction was followed by the decrease

of the absorbance at 440 nm which corresponds to the 7,9-dicis-rhodopsin (similar spectra

shown in Figure 4.4). In Figure 4.17 are shown plots corresponding to the disappearance

of the 7,9-dicis-rhodopsin and 7-cis-rhodopsin. They clearly show that 7,9-dicis­

rhodopsin disappears at a slower rate than that of 7-cis-rhodopsin (ratio -2: 1), indicating

a lower quantum yield for the photoreaction of 7,9-dicis-rhodopsin at -190°C than that of

7-cis rhodopsin.

Photosensitivity of 9, II-dicis-12-fluororhodopsin was measured relative to that of

9-cis-12-fluororhodopsin. The photobleaching process conducted after irradiation with

437 nm light was followed by determination of the decrease of absorbance maxima as a

function of time. The initial slope in a plot of absorbance versus irradiation time was

171

used for calculation of relative photosensitivity. A 9-cis-12-fluororhodopsin sample of

equal absorbance was used as the reference in the photosensitivity determination. The

results are shown in Figure 4.18. From the initial slope of the plot in Figure 4.18, the

photosensitivity of 9-cis-12-fluororhodopsin is about two times larger than that of 9,11­

dicis-12-fluororhodopsin.

172

0.16 -----------------------,

0.15

<l> 7,9-diciso 0.14c::('\3.Q~

00.13C/)

.Q<t

7-cis0.12

0.11 +--...-------r---.,..----,~-__r--.,_--~

o 1 0 20 30 40

Irradiation Time (min)

Figure 4.17 Relative photosensitivity of 7,9-dicis-rhodopsin and 7-cis-rhodopsin during

parallel irradiation with 437 nm light as reflected in changes in absorbance at 440 nm.

173

0.4~------------------------,

0.3

(1)(J

ccu.c 0.2"-0 9,11-dicis(/)

.c~

0.19-cis

86

Time (min)

4

Irradiation

2o.0 +--r---"""'--~---'---~-"""T"--~-....,..----i

o

Figure 4.18 Relative photosensitivity of 9,11-dicis-12-fluororhodopsin and 9-cis-12­

fluororhodopsin during parallel irradiation with 437 nm light as indicated by changes in

absorbance at 500 nm.

174

4.4 Discussion

4.4.1 Spectral properties of rhodopsin analogs

In addition to ll-cis-retinal, 9-cis, 7-cis and 7,9-dicis isomers also bind the opsin

to form photosensitive isomeric pigments. These results show that the structure of the

binding site is relatively flexible. A variety of visual pigment analogs have been formed

between opsin and chemically synthesized retinal analogs (see review Derguini &

Nakanishi, 1986; Liu & Asato, 1990).

The UVNis absorption maximum of 7-cis (450 nm) and 7,9-dicis rhodopsin (444

nm) are blue shifted from those of II-cis (498 nm) and 9-cis (485 nm). Opsin shifts, as

defined as the red shift of chromophores upon binding with opsin (Nakanishi et al., 1980),

for l l-cis, 9-cis,.7 -cis and 7,9-dicis pigments are 2620, 2430 (Honig et al., 1979a, Sheves

et al., 1979), 980 and 1120 cm', respectively. This trend is likely due to combined

effects of distorted chromophore from a twisted 6,7 bond (Liu et al., 1983b) and modified

binding sites for 7-cis and 7,9-dicis pigments. Hence the blue shifts of the 7-cis and 7,9­

dicis pigment could be either due to a weakened protonation as a result of a misplaced

counterion (Blatz et al., 1972) or due to a relocated second point charge (Honig et al.,

1979b).

On the absorption of light, 7,9-dicis-rhodopsin converts to its bathointermediate

in the all-trans form. Experimental results show that the absorption spectrum of the

bathointermediate from 7,9-dicis-rhodopsin is different from that of native rhodopsin in

its absorption maximum is shifted by 8 nm to the blue as shown in the bathointermediate

175

difference spectra in the insert of Figure 4.8. On the other hand, the difference between

the batho from 9,1l-dicis-12-fluororhodopsin and that from 9-cis-I2-tluororhodopsin is

relatively small (2 nm blue shifted as shown in the insert of Figure 4.10).

CHO

7,9-dicis

Ll-cisCHO 9-cis

CHO

Figure 4.19 Structure of chromophore of 7,9-dicis-rhodopsin comparing with 9-cis and

l l-cis isomers.

It is important to know the differences in the chromophore-opsin interactions

between 7,9-dicis-rhodopsin and ll-cis-rhodopsin in order to elucidate the difference in

photobleaching process and the spectroscopic properties. Inhibition studies (Matsumoto

& Yoshizawa, 1975) and recent molecular modelling analysis (Liu & Mirzadegan, 1988)

showed the presence of a hydrophobic pocket interacting with the cyclohexenyl ring.

These features are implicit in subsequent analyses of the binding site of opsin. Figure

4.19 shows the structure of chromophore of 7,9-ditis rhodopsin comparing with those of

176

the 9-cis and II-cis isomers. l l-Cis- and 9-cis-chromophores are bent near the center of

the polyene chain and have a slightly twisted structure between the cyclohexenyl ring and

the chain, while 7,9-dicis-chromophore is doubly bent near the ring and also has a highly

twisted structure between the cyclohexenyl ring and the chain due to high steric hindrance

between 5-methyl and 9-methyl. Therefore, the interaction between the ~-cyclohexenyl

ring of the chromophore in 7,9-dicis-rhodopsin and opsin must be different from that of

II-cis. The difference may be the cause for the blue shift of the absorption maxima of

7,9-dicis-rhodopsin.

4.4.2 Models to account for absorption properties of pigments

As mentioned earlier, red shifts in absorption spectra are observed for rhodopsin,

which absorbs in the range of 440-650 nm, while the retinyl protonated Schiff base (PSB)

in MeOH absorbs at 445 nm. The bathochromic shift, expressed in em:', has been

defined as the opsin shift (Nakanishi et aI., 1980) and represents the overall

environmental effect of the protein binding site on the absorption maxima of the

pigments.

The visible absorption band of retinal results from electronic excitation of the

conjugated it electrons in the retinal polyene chain. Several mechanisms for delocalizing

these electrons and inducing red-shifts in retinal pigments have been discussed (Blatz et

al., 1972; Honig et al., 1976; Kakitani et al., 1985) including:

(1) charge separation between the protonated Schiff base and its protein

177

counterion,

(2) electrostatic interactions between the chromophore and a charged amino

acid residue, and

(3) twisting about double bonds or about single bonds with significant double

bond character.

Varying the distance between the Schiff base nitrogen and its protein counterion

represents an effective mechanism for modulating the visible absorption band of retinal

pigments (Blatz et al., 1972). The A.nax for rhodopsin is at 498 nm, and the absorption

band of PSB model compounds with large and easily polarizable counterions has been

observed to red-shift to 480 nm (Blatz et al., 1972; Childs et al., 1987).

In fact, the charge separation between the positively charged nitrogen of the Schiff

base and its negative counterion has been suggested as an energy storage mechanism

(Honig et al., 1979a; Birge et al., 1983), which induces a series of thermal relaxations of

the pigment through a series of intermediates.

The second mechanism for red-shifting the absorption band in retinal pigments

involves stabilizing delocalized electronic structures by placing protein charges near the

conjugated retinal chain, a mechanism proposed originally by Kropf and Hubbard (1958).

Nakanishi, Honig and coworkers found evidence for a negative protein charge, based on

rhodopsin regenerated with dihydroretinal derivatives (Nakanishi et al., 1979; Honig et

al., 1979a). The model postulates the existence of a counterion at 3 A from the

protonated Schiff base nitrogen and a second negative charge at 3 A from the C12 and C14

to account for the opsin shift for rhodopsin. The models for rhodopsin and

178

bacteriorhodopsin have contrasting charge distributions; in the case of rhodopsin the

second charge which assists in delocalizing the distribution of positive charge along the

polyene chromophore is near the C-12 to C-14 region, whereas it is near the ring in

bacteriorhodopsin.

The source of a point charge could in principle be a dipolar amino acid or a

carboxylic residue positioned so that the negative electron density is near the

chromophore. Displacement of the counterion to a longer distance from the iminium

nitrogen could, in principle, leads to large wavelength shifts (Blatz & Mohler, 1975;

Honig, et ai., 1976).

Conformational distortions of the planar retinal chain may also generate red-shifts

in the retinal absorption band. In general, in conjugated polyenes, twisting about double

bonds induces red-shifts, while twisting about single bonds induces blue-shifts (Kakitani

et ai., 1985).

In the resonance Raman spectra of rhodopsin, strong hydrogen out-of-plane

wagging vibrations of the Cll-H and C12-H protons indicate conformational distortion

in this region of the chromophore (Eyring et al., 1980). These vibrations most likely are

due to the distortion in the CIO-Cll=C1Z-Cl3 bonds. Thus, in the presence of the external

charge believed to be located between C-12 and C-14 (Honig et al. 1979b), a twist in the

C1Z-Cl3 single bond may contribute to the opsin shift. l3C NMR data also provide support

for the model of the opsin shift which relies on a protein perturbation in the vicinity of

C-13 (Smith et al., 1991).

179

In 7,9-dicis-rhodopsin, the blue shifted absorption is probably due to the relocation

of the second point charge or increased single-double bond alternation. The chromophore

of 7,9-dicis-rhodopsin could have its binding site deviated from the native binding pocket.

So it is reasonable to assume that the second point charge would now be located at a

different position relative to the polyene chain. Since the second point charge has been

considered a cause of red shift of pigment absorption, a diminished interaction could

cause the chromophore in a dicis-chromophore blue shifted. Alternatively, a distortion

of the 7,9-dicis-retinylidene chromophore is another contributing cause. Because of the

ring-chain interactions near the C5-~' the 7,9-dicis-retinylidene chromophore is highly

twisted in the protein pocket, and the conjugation of single-double bonds should decrease.

Thus causing a blue-shift in the 7,9-dicis-rhodopsin and its bathointermediate.

9,Il-Dicjs-12-fluororhodopsin is also similarly blue-shifted in its absorption.

Since the chromophore shape of 9, Il-dicis isomer is close to that of the original binding

site, the protein perturbation of 9, ll-dicis-rhodopsin is expected to be similar with that

of rhodopsin. This is the possibly the reason why the difference between bathorhodopsin

from 9, ll-dicis-12-fluororhodopsin and bathorhodopsin from 9-cis-12-fluororhodopsin is

relatively small.

Thus, the spectral properties of the dicis-pigments appear to follow the general

principle that long wavelength absorption is correlated with increased electron

delocalization and decreased single-double bond alternation. The spectroscopic properties

of conjugated hydrocarbon chains are related to the extent of 1t electron delocalization

180

(Labhart, . 1957). Any mechanism that increases delocalization and reduces bond

alternation should induce bathochromic shifts in the absorption maxima.

4.4.3 Mechanism of primary photoreaction of rhodopsin analogs

It has been established that rhodopsin, isorhodopsin, 7-cis-rhodopsin and

bathorhodopsin all undergo one-photon-one-bond isomerization reaction. One-photon­

two-bond isomerization reactions have been observed for photochemical process of retinal

isomers in organic solution. This raises the question of the isomerization mechanism in

the dicis-rhodopsin system.

The proposed mechanism for photochemistry of 7,9-dicis-rhodopsin is based on

the combining results of the irradiation of 7,9-dicis-rhodopsin at liquid nitrogen

temperature (Figure 4.7) and HPLC analysis of extracted chromophores (Figure 4.11).

The spectra shifted to longer wavelengths with an isosbestic point at 517 nm during the

early stage of the irradiation (Figure 4.11) indicating the formation of a bathoproduct.

HPLC analysis of chromophore extracts (Figure 4.11) revealed that 7,9-dicis-rhodopsin

photoisomerized to the all-trans form. The presence of the sharp isosbestic point in the

UV/Vis spectra in the photochemical process (Figure 4.7) is supporting evidence that only

two components are involved in the photoirradiation process of 7,9-dicis-rhodopsin. Also

plots (insert in Figure 4.7) of relative absorption at 440 nm on a semilogarithmic scale

against the time of irradiation yield a straight line at the early stages of irradiation.

Prolonged irradiation yielded a photosteady-state mixture composed of rhodopsin,

181

isorhodopsin and bathorhodopsin. These components were readily interconvertible by

light at liquid nitrogen temperature (Figure 4.8).

On the other hand, the photochemical behavior of 9,II-dicis-12-fluororhodopsin

was different from that of 7,9-dicis-rhodopsin. Irradiation of 9,11-dicis-12­

fluororhodopsin also produced the bathoproduct by blue light (437 nm) at liquid nitrogen

temperature (Figure 4.9). The HPLC patterns of the chromophore extracted from different

irradiated samples indicated that the 9-cis pigment is formed from 9,1l-dicis-rhodopsin

(Figure 4.14). Therefore, one-photon-one-bond isomerization at liquid nitrogen

temperature took place in 9,1l-dicis-12-fluororhodopsin to the all-trans bathoproduct by

way of the 9-cis intermediate.

In Figure 4.20 are shown the photochemical processes identified for rhodopsin, 7­

cis-rhodopsin, 7,9-dicis-rhodopsin and 9, l l-dicis-rhodopsin at liquid nitrogen temperature.

Based on the experimental results cited above, the retinyl chromophore is anchored

at both ends within the hydrophobic region of the opsin (Matsumoto & Yoshizawa, 1975):

the protonated Schiff base linkage with lys-296 (Bownds, 1967; DeGrip et al, 1973;

Oseroff et al., 1974; Mathies et al, 1976; Callender et al, 1976) and the specific

hydrophobic pocket for the trimethylcyclohexenyl ring. Furthermore, the remaining

polyene chain is sandwiched between two layers of protein helices including the chiral

cavity (Matsumoto et aI., 1978). Thus, photoisomerization of the chromophore of

rhodopsin takes place in a restricted cavity of opsin.

On absorption of light, rhodopsin is converted first to photorhodopsin and then the

more stable bathorhodopsin (which is stable at liquid nitrogen temperature). During this

182

Rhodopsin "::Q;::=:======="~ Bathorhodopsin ..:::..;====-:::- Isorhodopsin

(11-cis) (all-trans) (9-cis)

7-cis-Rhodopsin

(7-cis)

-----,I... Bathorhodopsin

(all-trans)

~ Rhodopsin

~ (11-cis)

~ Isorhodopsin

(9-cis)

7,9-dicis-Rhodopsin '---l... Bathorhodopsin

(7,9-dicis) (all-trans)

~Rhodopsin

~ (11-cis)

~ Isorhodopsin

(9-cis)

9,11-dicis-Rhodopsin . Bathorhodopsin ...:;;;,;===-~ Rhodopsin

(9,11-dicis) (all-trans) (11-cis)

<:>Isorhodopsin

(9-cis)

Figure 4.20 Schematic showing interconversion among rhodopsin and isomeric rhodopsinanalogs and their photoproducts by light at liquid nitrogen temperature.

183

process the Cl1=CI2 bond is isomerized from cis to trans. The steric potential energy

induced by the cis-trans photoisomerization of the chromophore in the restricted cavity

cause a twisted all-trans chromophore in the bathorhodopsin stage. It has been implied

that electrostatic and steric potential energies stored in the bathorhodopsin may induce a

conformational change near the p-cyclohexenyl ring binding site of opsin to form

lumirhodopsin (Shichida, 1986).

The distribution of products from the photoisomerization of a polyene is

determined by regioselective twisting of the double bonds in the planar excited state (Liu

et al., 1983a). Because of the extremely short life time of these intermediates, usually

minor factors, such as relative ease in displacing surrounding molecules, could become.

a important factors to determine the direction of the photoreaction. For a molecule in a

confined medium, such as a retinyl chromophore inside the binding site of opsin, the

medium effect is expected to be very dramatic. That the major 13-cis isomer in solution

is not an observed photoproduct from rhodopsin and bathorhodopsin suggests that the

binding site is highly restrictive near the Schiff base portion of the chromophore, possibly

due to the association of the counter-ion with the protonated nitrogen.

The cyclohexenyl ring of the chromophore is constrained through hydrophobic

interaction with the protein (Matsumoto et al., 1978, 1975). That 7-cis isomers are not

observed in photoreaction of visual pigments could lead to the impression that the binding

site is also tightly packed near this end. However, since even in solution the 7-cis isomer

is either absent or formed in trace amounts (Ganapathy & Liu, 1992a), the ring terminus

need not be as tightly congested as that near the Schiff base. That bathorhodopsin readily

184

gives rhodopsin and isorhodopsin (9-cis) suggests that the protein-induced constraint near

the middle portion of the chromophore is relatively small. Considering that geometric

isomerization of any double bond requires twisting one end of a double bond, one may

further rationalize from the observed photochemistry that the binding site is such specific

that the region surrounding ClO and C11 of the chromophore is most likely to induce

twisting of the double bonds.

The study of 9,13-dicis-rhodopsin showed that irradiation of the 9,13-dicis­

rhodopsin mainly produced the 13-cis isomer, a one-photon-one-bond isomerization

process (Shichida et al., 1988b). Thus, it is a regioselective process isomerizing

preferentially at the 9,1O-bond instead of the 13,14-bond. This result again reflects the

highly confined nature of the binding site near the iminium center of the chromophore in

rhodopsin, which becomes relatively more open near the central portion of the polyene

chromophore (Liu & Asato, 1985).

The photochemical reactions of 7-cis-rhodopsin by low temperature

spectrophotometry and laser photolysis have shown that the bathorhodopsin from 7-cis

exhibits different absorption properties from that of rhodopsin. A difference in

configuration of the chromophore leading to different chromophore-opsin interactions, is

a likely cause for the difference in the two bathoproducts. For example, a different

interaction between the 9-methyl group of the chromophore and the neighboring protein

in 7-cis rhodopsin has been attributed to the different absorption properties (Shichida et

al., 1992). In rhodopsin and 9-cis rhodopsin, the conversion of the chromophore does not

require any significant relocation of the 9-methyl group. On the other hand, the

185

conversion of 7-cis-rhodopsin to the all-trans chromophore requires reorientation of the

9-methyl group, possibly causing a local protein perturbation manifested in spectral

properties of the primary photoproduct different from those of bathorhodopsin.

In 7,9-dicis-rhodopsin, the all-trans product results from isomerization near the

terminal ~=C8 and ~=ClO double bonds. For this more sterically crowded isomer with

a highly twisted ring-chain conformation (Ramamurthy et aI., 1972), a different protein

substrate interaction might be involved. There might be the possibility that newly

imposed protein restrictions have suppressed all one-bond isomerization processes, thus

with little recourse, the 7,9-dicis-chromophore isomerizes via an ordinarily high energy

pathway, i.e., the two-bond isomerization process.

Such a rationale seems to be supported by the modelling of chromophore

structures of these isomeric pigments. In Figure 4.21 are the overlaid structures of the

tethered chromophores of the l l-cis, 9-cis, 7,9-dicis pigments anchored at the a-carbons

of the lysine residue and the cyclohexenyl rings. Figure 4.22 are the overlaid structures

of the tethered chromophores of the l l-cis, 9-cis and 9, l l-dicis pigments with the same

anchors at the two ends of the chromophore. For the 7,9-dicis chromophore with the

established 15-anti configuration (Loppnow et aI., 1990), its ring chain conformation and

the positions of the ~-ClO atoms are seen to be substantially displaced from those of the

9-cis and l l-cis isomers. It is reasonable to expect that the 7,8 and 9,10 bonds are now

in a more crowded environment of the protein pocket. On the other hand, it is also

evident from the figure of 9,II-dicis pigment (Figure 4.21) that the 11,12 bond is less

186

displaced from that of rhodopsin, thus likely to face the same perturbing force that

enhances isomerization of rhodopsin.

A different selectivity in photoisomerization is also exhibited by the retinyl

chromophore trapped in the binding cavity of bacteriorhodopsin (Ottolenghi and Sheves,

1986), halobacteriorhodopsin (Lanyi, 1986), or phobobacteriorhodopsin (Shichida et al.,

1988a). In all these cases the protonated all-trans-retinyl Schiff base undergoes

regiospecific photoisomerization at the C13=CI4 bond. The apparent difference from the

preferred isomerization at the Cll =C12 bond for the free chromophore and visual

pigments must be due to the unique shape of the binding site, which is known to be more

steroselective. Bacteriorhodopsin reacts only with the all-trans and 13-cis isomers and

not with 7-cis and the centrally bent 9-cis and ll-cis isomers of retinal (Stoeckenius &

Bogomolni, 1982).

4.4.4 Postulated models of mechanism of photoisomerization of rhodopsin

Seeking for a minimal change of the moment of inertia during the isomerization

process, Warshel proposed a so-called "bicycle pedal" model, in which two double bonds

are rotated in a concerted manner after absorption of a photon (Warshel, 1976). This

model successfully explained the ultrafast isomerization, with high quantum yields.

However this model predicted a stepwise conversion from isorhodopsin (9-cis) to

rhodopsin (ll-cis), 13-cis, 15-cis and eventually to bathorhodopsin (all-trans isomer).

187

-0000

Figure 4.21 Computer simulated chromophores of rhodopsin and its analogs. Overlaid chromophore structures of rhodopsin,9-cis-rhodopsin, 7,9-dicis-rhodopsin. The energy minimized isomeric retinal chromophores were replaced with inimiumnitrogen and appended with the butyl tether and an extra carbon for the a-carbon of the anchored lysine residue.

-00'-0 F

~N

/

H

Figure 4.22 Computer simulated chromophores of 12-fluororhodopsin and its analogs. Overlaid chromophore str-uctures of12-fluororhodopsin, 9-cis-12-fluororhodopsin and 9, Il-dicis-12-fluororhodopsin. The energy minimized isomeric retinalchromophores were replaced with inimium nitrogen and appended with the butyl tether and an extra carbon for the 0.­

carbon of the anchored lysine residue.

Experimentally, there was no evidence for such direct conversion of isorhodopsin to

rhodopsin (Sarai et ai., 1980) nor the existence of any intermediates. This model was

later revised (Warshel & Barboy, 1982) to include a concerted "extended" bicycle-peda!

model to allow direct conversion from a cis isomer to the trans. The second model, so­

called "Hula-twist" model was proposed (Liu & Asato, 1985). According to this model,

the isomerization proceeds with a concerted rotation of one double bond and its

neighbouring single bond, resulting in formation of bathorhodopsin having the 10-s-cis

conformation of its chromophore. This model can nicely explain the formation of a

photosteady state mixture containing only rhodopsin, isorhodopsin and bathorhodopsin

at liquid nitrogen temperature. However, vibrational spectroscopic results for

bathorhodopsin suggested that CIO-CII is in the s-trans conformation (Mathies et ai.,

1989).

The photoisomerization of 7,9-dicis-rhodopsin involves an one-photon-two-bond

process, a motion consistent with the volume conserving "bicycle pedal model". In

Figure 4.23 are presented the bicycle pedal model of 7,9-dicis-rhodopsin to

bathorhodopsin (all-trans). Since the binding site of 7,9-dicis-rhodopsin is dramatically

different from those of l l-cis and 9-cis-rhodopsin, in terms of the steric effects around

the C7-C IO atoms of the chromophores, the unique photoisomerization process of the 7,9­

dicis chromophore can be rationalized by specific protein perturbation for this

chromophore. It is reasonable to expect that the C7=Cg and C9=C IO bonds are in a more

crowded environment in the protein pocket, therefore the one-bond isomerization which

is a more volume demanding process would be hindered. Instead, the less volume

190

demanding "bicycle-pedal" two bond isomerization process could become more

favorable. The consequence is that two adjacent Cg=C9 and C9=CIO double bonds can

isomerize at the same time without a large shift in atomic coordinates. For the 9, l l-dicis

pigment, since the 11,12 bond is less displaced from that of rhodopsin and no additional

twist in the polyene chain is introduced, it is likely to have a similar protein perturbation

as that which enhances isomerization of rhodopsin and isorhodopsin. Therefore selective

isomerization at the more hindered l l-cis double bond leads via a two-step reaction to

the eventual batho product.

It will be of interest to design model systems to mimic and test the concept of

host restrictions being able to impede the normal process of one-bond isomerization to

the extent favoring exclusively a less common two-bond process.

Finally, it should be added that the current method for product analysis, whether

of the pigments or their photoproducts, does not allow detection of structures involving

rotation of a single bond or the imino double bond in the polyene chromophore during

the photoisomerization process. Future FTIR or resonance Raman studies on these dicis

rhodopsin analogs might provide additional information for elucidation of the specific

nature of the isomerization process.

191

\0tv

~N-"W~

)-""~

Figure 4.23 A possible "bicycle-pedal" path for 7,9-dicis-rhodopsin to bathorhodopsin (all-trans). The parallel doublebonds of molecule moves in a concerted rotation without large shift of the atomic coordinates.

Chapter 5

Conclusion

Following the completion of the syntheses of all sixteen geometric isomers of

retinal (Trehan et al., 1988; Liu & Asato, 1984a), studies of their photochemical

(Ganapathy & Liu, 1992a,b) and spectroscopic properties have been reported including

the UV/Vis, IH-NMR spectral data (Zhu et ai., 1992), and HPLC separation conditions.

In this work added to this list are the vibrational spectra of all of these isomers and the

corresponding SB's and PSB's. In this study, we have also undertaken a combined study

of low temperature photochemistry and chromophore extraction during the course of

irradiation of the only two stable dicis pigments: 7,9-dicis-rhodopsin and 9, ll-dicis-12­

fluororhodopsin. Unexpectedly, they have been shown to react photochemically by two

divergent pathways: one-photon-two-bond isomerization for 7,9-dicis-rhodopsin and one­

photon-one-bond isomerization for 9, l l-dicis-Iz-fluororhodopsin. The final bleaching

products are the all-trans isomers in both systems. During this study, we also showed

that the batho-intermediate of 7,9-dicis-rhodopsin has a blue-shifted absorption spectrum

compared to the native bathorhodopsin. These results reflect the protein effects on the

photoisomerization of dicis chromophores.

In view of these results, it will be of interest to examine the photochemical

properties of a stable tricis-rhodopsin analog. At issue will be the question whether a

one-photon-three-bond mechanism could take place in such a system. If so, clearly it

193

will occur by a pathway other than the bicycle pedal process. However, while two tricis­

retinal isomers (7,9,II-tricis or 7,9, 13-tricis) were believed to yield rhodopsin analogs,

the extent of configuration retention of the chromophore was not demonstrated in these

early studies (Liu et al., 1984b). In fact, based on the dicis isomers (Trehan et ai.,

199Gb), one can predict that the protein will likely catalyze isomerization at either the

Cll =C12 or C13=C14 double bond. Thus, for such a study, conditions for stable pigment

formation, e.g. adding a substituent to stabilize the labile l l-cis geometry, must first be

established.

Since the bathe-intermediate of 7,9-dicis rhodopsin has been shown to be different

from the batho-intermediate of l l-cis rhodopsin, it will be interesting to know whether

the following thermal decay intermediates still show differences from the corresponding

ones of the native rhodopsin. Time-resolved spectroscopic techniques, including

vibrational spectroscopy, could provide useful information on each intermediate during

the photobleaching process of these rhodopsin analogs.

194

---.. _---

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