Visual Pigment Function

Molecular evolution of zebrafish (Danio rerio) visual pigment function

James Michael Morrow

A thesis submitted in conformity with the requirements for the degree of Doctor of Philosophy Cell & Systems Biology University of Toronto

Molecular evolution of zebrafish (Danio rerio) visual pigment function

James Michael Morrow

Degree of Doctor of Philosophy Department of Cell & Systems Biology University of Toronto 2014

Abstract

Photoreception in vertebrates is mediated by opsins, members of the G protein-coupled receptor (GPCR) superfamily of proteins. In the dark, opsins are covalently bound to a light-sensitive chromophore, 11-cis-retinal, which acts as an inverse agonist to suppress dark state activation. When exposed to light, the chromophore isomerizes to its all-trans conformation, activating opsin and initiating a signaling cascade within the photoreceptor cell. Zebrafish (Danio rerio) has a large complement of visual opsins, and serves as a model for the vertebrate visual system on a developmental, physiological, and behavioural level, making it an ideal candidate for the study of opsin evolution and function. Following a general introduction, chapter 2 of my thesis presents the initial characterization of a novel rhodopsin-like gene that we identified in zebrafish, rh1-

2. Chapter 3 further investigates rh1-2, showing rh1-2 expression in photoreceptor cells of the zebrafish retina and using in vitro expression to identify functional characteristics of rh1-2 that are similar to rhodopsin, despite the fact that it is a much less stable pigment. Chapter 4 describes biochemical investigations of zebrafish rhodopsin and, in

ii particular, addresses the question of the key molecular mechanisms mediating retinal release rate differences among vertebrate rhodopsins. Finally, chapter 5 investigates duplicated rh2 cone opsin genes in teleost fish and notes that many with multiple rh2 genes have at least one copy with Q122 and one with E122, a discrepancy that is known to help in establishing the contrasting functional identities of rhodopsins and cone opsins. This thesis contributes significantly to the characterization of visual pigment function in zebrafish. These experiments have also increased the understanding of functional variation within rhodopsins and cone opsins, particularly by investigating the molecular mechanisms of retinal release in rhodopsin, and the role of duplicated cone opsin genes, and will serve as an important non-mammalian reference against which future studies of the vertebrate visual system will be compared and contrasted.

iii Acknowledgments

At this moment in time, I feel very lucky. From the age of 23 to 29, I was granted the opportunity to think for a living. While my time in graduate school did not allow for a frivolous lifestyle, this was an opportunity that very few people are afforded during this period of their lives, and I have done my very best to take advantage of my fortunate situation.

Throughout my studies, I have grown both as an academic and an individual. Since it won’t be possible to list the names of everyone that has contributed to this growth and supported me during this time, I want to quickly acknowledge my friends, extended family, fellow lab members, collaborators, CSB Faculty and staff. If you are reading this right now, chances are you are a part of this list, so thank you! I also want to extend this thanks to my enemies, critics, competitors and saboteurs; I am often motivated by the challenges presented to me, and am often able to reach new heights of accomplishment only when told they are beyond my capacities.

The most central figure to my academic success has without a question been my supervisor, Belinda Chang. I have been extremely fortunate to have a supervisor that always gave me a chance to both prove and improve my worth. To be able to openly discuss any topic or idea with my supervisor, academic or otherwise, and to be able to trust that her thoughts and actions are always in my best interest is a feeling that I greatly appreciate and have never taken for granted. Both Vince Tropepe and Melanie Woodin were also central in guiding my research goals as members of my advisory committee,

iv and I would like to thank them for their helpful insights and patience. I would also like to thank Dr. Shoji Kawamura, my external examiner, for a very thoughtful and thorough thesis appraisal.

Mom, dad, and Matt, I hope you all know how important your support has been throughout this process. I know that we have a very hands-off family dynamic, allowing each part to have its much needed independence, but I never forget nor take for granted the security that the three of you have provided and continue to provide for me. I feel that one of the reasons that I’ve been able to avoid falling down in life is because I have the confidence in knowing that the three of you would be there to catch me if I ever do.

Last but not least, my puppy dog Cody. You’ve only been at my side for 18 months, but you’re probably the main reason that I have remained in a healthy psychological state over that period of time, especially during the latter stages of this writing process.

You’re a very good boy and yes, it is time for a walkie.

v Table of Contents

Abstract ...... ii Acknowledgments ...... iv Table of Contents ...... vi List of Figures ...... ix List of Tables ...... xi List of Abbreviations ...... xii Publications and author contributions ...... xv

Chapter 1 – General introduction ...... 1

1.1 Visual pigments ...... 2 1.2 Phototransduction ...... 7 1.3 The retinoid cycle ...... 9 1.4 Cells of the retina ...... 11 1.5 Heterologous opsin expression ...... 15 1.6 Visual pigment structure and function ...... 18 1.7 Zebrafish ...... 23 1.8 Thesis objectives ...... 25 1.9 Thesis overview ...... 26

Figures...... 30 References ...... 34

Chapter 2 – A novel rhodopsin-like gene expressed in zebrafish retina ...... 51

2.1 Abstract ...... 52 2.2 Introduction ...... 53 2.3 Materials and methods ...... 56 2.3.1 Opsin sequences ...... 56 2.3.2 Protein expression ...... 57 2.3.3 Localizing expression of zebrafish rh1-2 ...... 58 2.3.4 Phylogenetic analyses ...... 58 2.4 Results ...... 60 2.4.1 Sequence analysis of rh1-2 ...... 60 2.4.2 In vitro expression ...... 62 2.4.3 Zebrafish rh1-2 expression pattern ...... 62 2.4.4 Phylogenetic analyses ...... 63 2.5 Discussion ...... 64

Figures...... 71 Tables ...... 77 References ...... 81

vi Chapter 3 – Cellular localization and functional characterization of zebrafish rh1-2: investigating its role in vision ...... 92

3.1 Abstract ...... 93 3.2 Introduction ...... 94 3.3 Materials and methods ...... 98 3.3.1 Opsin sequences ...... 98 3.3.2 In situ hybridization ...... 99 3.3.3 Expression and spectroscopy ...... 100 3.3.4 Phylogenetic and molecular evolutionary analyses ...... 102 3.4 Results ...... 104 3.4.1 Zebrafish rh1-2 is expressed in the outer nuclear layer of peripheral rod photoreceptor cells ...... 104 3.4.2 Zebrafish rh1-2 releases retinal at a rate similar to rhodopsin, but slower than exo-rhodopsin ...... 105 3.4.3 Co-transfection with rh1-2 alters thermal stability of rhodopsin ...... 106 3.4.4 The rh1-2 gene family is sister to Ostarioclupimorpha rh1 genes ...... 107 3.5 Discussion ...... 109

Figures...... 118 Tables ...... 124 References ...... 131

Chapter 4 – Investigating retinal release in light-activated zebrafish rhodopsin using fluorescence spectroscopy: the role of steric effects and Schiff base stability ...... 140

4.1 Abstract ...... 141 4.2 Introduction ...... 142 4.3 Materials and methods ...... 145 4.3.1 Visual pigment expression and purification ...... 145 4.3.2 Spectroscopy ...... 146 4.3.3 Homology modeling ...... 147 4.4 Results ...... 148 4.4.1 Biochemistry of zebrafish rhodopsin ...... 148 4.4.2 Effects of mutations at opening in metarhodopsin II ...... 149 4.4.3 Effects of mutations in the chromophore binding pocket ...... 150 4.4.4 Effects of temperature on retinal release ...... 152

4.4.5 Effects of mutations on λMAX ...... 153 4.5 Discussion ...... 154 4.6 Conclusions ...... 163

Figures...... 165 Tables ...... 173 References ...... 176

vii Chapter 5 – A functional dichotomy of rod-like cone opsin genes in teleost fish linked to a single amino acid at site 122 ...... 185

5.1 Abstract ...... 186 5.2 Introduction ...... 187 5.3 Materials and methods ...... 190 5.3.1 Visual pigment expression and purification ...... 190 5.3.2 Spectroscopy ...... 191 5.3.3 Phylogenetic Analyses ...... 192 5.4 Results ...... 192

5.4.1 Site 122 helps to establish the λMAX of zebrafish rh2 opsins ...... 192 5.4.2 Zebrafish rh2-1 and rh2-4 are both susceptible to hydroxylamine ...... 193 5.4.3 Site 122 can mediate retinal release rates in bovine rhodopsin ...... 194 5.4.4 Zebrafish rh2-4 releases retinal at least 240 times faster than rhodopsin...... 195 5.4.5 Phylogenetic distribution of teleost rh2 opsins ...... 195 5.5 Discussion ...... 196

Figures...... 201 References ...... 207

Chapter 6 – General Discussion ...... 211

6.1 GPCR Dimerization ...... 212 6.2 Influence of retinal release on in vivo function ...... 215 6.3 Genetic diversity in Actinopterygian fish ...... 217

References ...... 220

viii List of Figures

Chapter 1 Figure 1. A schematic of the phylogenetic relationships among the five classes of vertebrate visual pigments ...... 30 Figure 2. A schematic of the phototransduction cascade in a rod outer segment ..... 31 Figure 3. A schematic of the various steps of the retinoid cycle ...... 32 Figure 4. The major steps involved in generating and assaying visual pigment samples using a heterologous expression system ...... 33

Chapter 2 Figure 1. Amino acid alignments of zebrafish rh1-2 ...... 71 Figure 2. Structures of three key motifs of rh1 containing cone-like and unique substitutions in the rh1-2 gene family ...... 73 Figure 3. Absorbance spectra of zebrafish rh1 and rh1-2 after in vitro expression and purification ...... 74 Figure 4. RT-PCR analysis of zebrafish rh1-2 expression ...... 75 Figure 5. Phylogeny of teleost rh1 nucleotide sequences, including the rh1-2 gene family, generated using both maximum likelihood and Bayesian methods ...... 76

Chapter 3 Figure 1. In situ hybridization [ISH] of rh1-2 in the zebrafish retina at various developmental stages ...... 118 Figure 2. Retinal release rates of various zebrafish rhodopsins at 20oC ...... 119 Figure 3. Co-transfection with zebrafish rh1-2 alters thermal stability of rhodopsin ...... 120 Figure 4. A schematic of the position of rh1-2 gene family with other Vertebrate visual pigment genes based on a Bayesian phylogenetic analysis ...... 121 Figure S1. In situ hybridization of rh1 in 3 dpf zebrafish embryos ...... 122 Figure S2. Detailed Bayesian phylogeny of teleost rh1 and rh1-2 genes ...... 123

Chapter 4 Figure 1. Ultraviolet-visual absorbance spectra of bovine and zebrafish rhodopsin ...... 165 Figure 2. Increase in fluorescence intensity, representing release of all-trans-retinal from the chromophore binding pocket, in bovine and zebrafish rhodopsin following photoactivation...... 166 Figure 3. Crystal structure homology model of zebrafish rhodopsin, highlighting TM5 and 6 ...... 167 Figure 4. Crystal structure homology model of several motifs lining the retinal channel in zebrafish rhodopsin ...... 168

ix Figure 5. Arrhenius plot of the natural logarithm of the rates of fluorescence increase in photoactivated zebrafish rhodopsin between 5 and 24oC at pH 7 ...... 169 Figure 6. Increase in fluorescence intensity in bovine and zebrafish rhodopsin following photoactivation at respective physiological temperatures ...... 170 Figure S1. Absorbance of zebrafish rhodopsin is not altered by incubation with hydroxylamine ...... 171 Figure S2. Sequence logo representing variation among vertebrate rhodopsins of eight residues shown to mediate retinal release rates ...... 172

Chapter 5 Figure 1. Crystal structure of the chromophore binding pocket of dark state bovine rhodopsin ...... 201 Figure 2. Ultraviolet-visual absorbance spectra of zebrafish rh2 opsins ...... 202

Figure 3. Decrease in λMAX absorbance in bovine and zebrafish visual pigments following incubation in hydroxylamine at 25oC ...... 203 Figure 4. Increase in fluorescence intensity representing release of all-trans-retinal from bovine rhodopsin following photoactivation ...... 204 Figure 5. Increase in fluorescence intensity representing release of all-trans-retinal from zebrafish rhodopsin and rh2-4 following photoactivation ...... 205 Figure 6. Phylogeny of teleost rh2 nucleotide sequences generated using maximum likelihood methods ...... 206

x List of Tables

Chapter 2 Table 1. Dominant identities of key sites of rh1-2 in other opsin families ...... 77 Table S1. List of primer sequences used in this chapter ...... 78 Table S2. List of opsin sequences from GenBank used for amino acid alignments or phylogenetic analyses ...... 79

Chapter 3 Table S1. List of sequences used to generate rh1 gene phylogeny ...... 124 Table S2. Results of random sites (PAML) analyses on subsets of the vertebrate rh1 gene tree ...... 128 Table S3. Results of branch, branch-site, and clade model C (PAML) analyses with the branch leading to the rh1-2 clade place into the foreground ..... 129 Table S4. Results of branch, branch-site, and clade model C (PAML) analyses with the rh1-2 clade placed into the foreground ...... 130

Chapter 4

Table 1. Retinal release rate and λMAX values for wild type and mutant Rhodopsins measured at 20oC ...... 173 Table 2. Retinal release rates of wild type bovine and zebrafish rhodopsin at various temperatures...... 174 Table S1. List of rhodopsin sequences used to generate the sequence logo along with respective GenBank accession numbers ...... 175

xi List of Abbreviations

λMAX wavelength of maximal absorbance A1 11-cis-retinal A2 11-cis-3,4-didehyroretinal

A280 absorbance at 280 nanometers AMAX absorbance at wavelength of maximal absorbance AIC akaike information criterion AML adenovirus major late BLAST basic local alignment search tool bp base pairs Ca2+ calcium ion cDNA complementary deoxyribonucleic acid cGMP cyclic guanosine monophosphate CmC clade model C CMV cytomegalovirus COS CV-1 origin with SV40 DIG digoxygenin DM N-dodecyl-β-D-maltoside DNA deoxyribonucleic acid DOPE discrete optimized protein energy dpf days post-fertilization

Ea activation energy EL extracellular loop ER endoplasmic reticulum ERG electroretinogram ESS effective sample size exorh extraretinal rhodopsin FRET fluorescence resonance energy transfer GABA gamma-aminobutyric acid GDP guanosine diphosphate GFP green fluorescent protein GMP guanosine monophosphate GPCR G protein-coupled receptor GRK rhodopsin kinase GTP guanosine triphosphate GTR generalized time reversible model HEK human embryonic kidney HKY+I+G Hasegawa, Kishino and Yano model with estimated proportion of invariable sites and gamma-distributed rate heterogeneity IPM interphotoreceptor matrix IRBP interphotoreceptor retinoid-binding protein

xii ISH in situ hybridization kcal kilocalory kDa kilodalton Ln natural logarithm LRAT lecithin:retinol acyltransferase LRT likelihood ratio test lws long wavelength-sensitive visual pigment MCMC markov chain monte carlo MEGA molecular evolutionary genetic analysis melop melanopsin meta II metarhodopsin II ML maximum likelihood MSP microspectrophotometry Na+ sodium ion NaPhos sodium phosphate nm nanometer NCBI national center for biotechnology information ONL outer nuclear layer ORF open reading frame PAML phylogenetic analysis by maximum likelihood PBS phosphate-buffered saline PBT phosphate-buffered saline with Tween-20 PCR polymerase chain reaction PDE phosphodiesterase PSRF potential scale reduction factor R* activated rhodopsin visual pigment RDH retinol dehydrogenase RGS regulator of G protein signaling rh1 rhodopsin visual pigment rh1-2 rhodopsin visual pigment 2 rh1dso deep sea rhodopsin visual pigment rh1fwo fresh water rhodopsin visual pigment rh2 rhodopsin-like cone visual pigment RNA ribonucleic acid ROS rod outer segment RPE retinal pigment epithelium RT-PCR reverse transcription polymerase chain reaction S2 section 2 component of dark adaptation SV40 simian virus 40 sws1 short wavelength-sensitive visual pigment 1

xiii sws2 short wavelength-sensitive visual pigment 2 t1/2 half-life t1/2Rel relative half-life TM transmembrane helix tmtop teleost multiple tissue opsin VA opsin vertebrate ancient opsin VAL opsin vertebrate ancient long opsin valop vertebrate ancient opsin gene VCOP Xenopus short wavelength-sensitive visual pigment 1 VS violet-sensitive UVS ultraviolet-sensitive

xiv Publications and Author Contributions

Chapter 2

Contributors: James M Morrow, Savo Lazic, Belinda SW Chang

This study was conceived by JMM, SL, and BSWC, and has been published:

Morrow JM, Lazic S, Chang BS (2011) A novel rhodopsin-like gene expressed in zebrafish retina. Vis Neurosci 28, 325-335.

Amino acid alignment, structural modeling, opsin expression and absorption spectra were generated by JMM. DNA sequences were amplified by JMM and SL. Phylogenetic analysis was performed by BSWC. Manuscript was written by JMM, with edits and guidance from SL and BSWC.

Chapter 3

Contributors: James M Morrow, Savo Lazic, Monica Dixon, Ryan Schott, Claire Kuo, Vince Tropepe, and Belinda SW Chang.

This study was conceived by JMM, SL, VT, and BSWC. In situ hybridizations were performed by SL, MD, and CK. Opsin expression, retinal release measurements, and thermal stability measurements were performed by JMM. Phylogenetic analyses were performed by RS. Chapter was written by JMM, with guidance from BSWC, and text from RS and VT.

Chapter 4

Contributors: James M Morrow and Belinda SW Chang

This study was conceived by JMM and BSWC. All data was collected by JMM. Chapter was written by JMM, with guidance from BSWC.

Chapter 5

Contributors: James M Morrow and Belinda SW Chang

This study was conceived by JMM and BSWC. All data was collected by JMM. Chapter was written by JMM, with guidance from BSWC.

Chapter 1:

General introduction

1 1.1 Visual pigments

Vision provides organisms with the capacity to assess and understand their environment almost instantaneously. On a molecular level, vision is mediated by visual pigments, members of the G protein-coupled receptor [GPCR] superfamily that initiate the critical first step in the biochemical cascade of vision (Stryer, 1986). These molecules comprise an opsin apoprotein linked to a retinylidene chromophore via a protonated Schiff base linkage (Wald, 1968). Opsin is a 35 kDa integral membrane protein consisting of seven transmembrane α-helices, a common feature of GPCRs (Baldwin et al., 1997).

Meanwhile, the light-absorbing chromophore, either 11-cis-retinal [A1] or 11-cis-3,4- dehydroretinal [A2], acts as an inverse agonist when bound to opsin, increasing stability and suppressing activity of the dark state visual pigment (Han et al., 1997). The light- sensitivity of a visual pigment is largely due to the relationship between opsin and chromophore, which influences many key biochemical properties including the

characteristic wavelength of maximal absorption, referred to as λMAX (Nathans, 1999).

While many features of visual pigments have been conserved throughout vertebrate evolution, they have also evolved to perform specialized visual functions; cone opsins are responsible for photopic [bright light] and colour vision while rod opsins are responsible for scotopic [dim light] vision (Menon et al., 2001). Mutations that compromise this highly specialized function of visual pigments are the main cause of retinal diseases such as retinitis pigmentosa (Sung et al., 1993). Because of their unique role as the molecular liaison between organisms and their photic environment, many studies have focused on gaining a better understanding of the structure and function of visual pigments, which has led to a much greater comprehension of the vertebrate visual system.

2 Visual pigments resolve into five distinct phylogenetic classes, one class of rhodopsins

[RH1] and four classes of cone opsins: long wavelength-sensitive [LWS], short wavelength-sensitive 1 [SWS1], short wavelength-sensitive 2 [SWS2] and rhodopsin-like

[RH2] (Fig. 1). While the term ‘rhodopsin’ can sometimes be used as a generic term for either any visual pigment, or any retinal-based photopigment, for the purpose of this thesis rhodopsin will refer only to visual pigments coded by the RH1 phylogenetic group of opsins. These five opsin classes have arisen from a single ancestral opsin gene through a series of gene duplication events (Yokoyama, 2000; Bowmaker and Hunt,

2006). Phylogenetic evidence suggests that the divergence of opsins occurred prior to that of vertebrates, aided by several whole genome duplications, meaning that the ancestors of vertebrates likely possessed all five classes (Yokoyama and Yokoyama,

1996). However, many extant groups of vertebrates have either lost one or more opsins classes, or have experienced gene duplication within one or more opsin classes, which often exist as tandem gene arrays following local gene duplications (Chinen et al., 2003;

Matsumoto et al., 2006; Weadick and Chang, 2007).

The group of vertebrates with the most diverse complement of visual pigments are teleost fish [infraclass: Teleostei], where many families have duplicated at least one class of cone opsin genes to expand the range of peak spectral sensitivity for certain wavelengths of light (Bowmaker, 2008). These expansions were supported by a whole-genome duplication event that occurred early in Actinopterygian evolution, providing many duplicate opsin gene copies on which selection could act (Hoegg et al., 2004).

Expansions of the LWS and RH2 gene families are the most common, with some species

3 having up to 6 and 5 copies of each gene, respectively (Chinen et al., 2003; Hoffmann et al., 2007; Weadick and Chang, 2007; Nakamura et al., 2013). There are also spectrally distinct subfamilies of SWS2 genes (Fuller et al., 2004; Bowmaker et al., 2006), however

λMAX values for teleosts with only one SWS2 gene will not necessarily fall into one of these two spectral classes (Chinen et al., 2005a). The SWS1 gene is rarely duplicated in vertebrates, including teleosts, but so far one exception has been identified in the smelt,

Plecoglossus altivelis (Minamoto and Shimizu, 2005). Meanwhile, the only examples of vertebrates with duplicated RH1 genes are found in teleosts. Multiple species of eels express a different rhodopsin gene when migrating from freshwater to saltwater during maturation (Beatty, 1975; Hope et al., 1998; Zhang et al., 2000; Zhang et al., 2002). The short-fin pearleye [Scopelarchus analis], a deep-sea teleost, also expresses an additional rhodopsin gene in adult fish after descending to greater ocean depths (Pointer et al.,

2007). Other examples are the result of more recent duplication events (Lim et al.,

1997). There have not been any cases of vertebrates with duplicated rhodopsin genes found outside of teleost fish.

The amphibian visual system is most notorious for the presence of two spectral classes of rod photoreceptors, with the dominant class expressing rhodopsin, as expected, and a small percentage expressing the SWS2 cone pigment (Ma et al., 2001; Darden et al.,

2003). While SWS1 (Hisatomi et al., 1998) and LWS genes (Babu et al., 2006) have also been identified, the RH2 opsin appears to have been lost early in amphibian evolution. However, in some cases, the SWS2 gene has undergone a significant red-shift

4 to move its λMAX to a range similar to RH2 genes from other vertebrates (Takahashi and

Ebrey, 2003).

In reptiles, the well-studied anoline lizards have maintained a copy from each visual pigment class, suggesting that reptiles have the potential for tetrachromatic colour vision

(Kawamura and Yokoyama, 1998; Loew et al., 2002). While other groups of reptiles have had less extensive examinations of their visual pigments, microspectrophotometry

[MSP] has been used to verify that tetrachromacy is also likely in alligators (Sillman et al., 1991) and turtles (Loew and Govardovskii, 2001). The visual pigments of birds, which include all five opsin classes, are considered highly conserved, except when considering the divergence of violet-sensitive [VS] and ultraviolet-sensitive [UVS] sws1 genes. While the ancestral vertebrate SWS1 is believed to be UVS (Hunt et al., 2001), the ancestral avian sws1 is thought to be VS, with a single amino acid substitution from phenylalanine to serine at site 86 believed to be largely responsible (Hart and Hunt,

2007). Various avian lineages have re-evolved UVS sws1 genes many times independently, primarily through a substitution of serine to cysteine at site 90, as opposed to a simple reverse substitution to phenylalanine at site 86 (Wilkie et al., 2000; Hunt et al., 2004).

Each of the three major groups of mammals [monotremes, marsupials, and eutherians] has distinguishing characteristics in their visual systems. All three groups have an LWS opsin attuned to long-wavelength light, and either an SWS1 or SWS2 opsin.

Monotremes, represented only by the platypus and two genera of echidna, are the only

5 mammalian group that have maintained an SWS2 opsin for short-wavelength sensitivity in addition to their LWS gene, making them functionally dichromatic (Davies et al.,

2007a; Wakefield et al., 2008). In marsupials, three spectrally distinct classes of cone photoreceptors have been identified, despite the fact that only SWS1 and LWS cone opsin genes have been detected (Arrese et al., 2002; Arrese et al., 2006). While neither an RH2 or modified LWS being identified as a candidate gene to represent a 505 nm cone signal, and despite examples of cone opsins being expressed in some rod photoreceptors of amphibians (Ma et al., 2001; Darden et al., 2003), the possibility of RH1 expression being responsible for this cone signal is not supported (Ebeling et al., 2010). The majority of eutherian mammals are functional dichromats, expressing both an SWS1 and

LWS, each showing a wide range of spectral sensitivities (Jacobs, 1993). However, marine mammals, as well as some nocturnal terrestrial mammals, have lost their SWS1 gene, presumably because colour vision is not essential to nocturnal organisms, but the pattern is far from consistent and the reasons for this loss are currently unknown

(Bowmaker and Hunt, 2006). Conversely, trichromacy re-evolved in primates either through polymorphism or duplication of the LWS gene (Bowmaker, 2008). In some

New World monkeys, polymorphisms resulted in allelic variants of LWS with different spectral sensitivities (Hunt et al., 1993; Jacobs and Deegan, 2003). Meanwhile, trichromacy is uniform across all catarrhines, including humans, due to a gene duplication of LWS that produced a blue-shifted variant, often referred to as MWS, or middle wavelength-sensitive (Hunt et al., 1998). The main theory put forth to explain the maintenance of trichromacy in primates is related to foraging, where wavelength discrimination in the red-green part of the visual spectrum allows an organism to better

6 identify orange and red fruits against a background of green foliage (Sumner and Mollon,

2000).

1.2 Phototransduction

Phototransduction is the process by which a visual stimulus in the environment is converted into electrical signals processed by cells in the retina, and is one of the most well studied G protein signaling systems. Visual pigments, such as rhodopsin, are responsible for mediating the critical first step of this process by activating in response to light, and consequently initiating a series of biochemical reactions in the photoreceptor cell (Fig. 2). Within femtoseconds of absorbing a photon, the 11-cis-retinal chromophore of rhodopsin is converted to its stereoisomer, all-trans-retinal, which elevates the potential energy of the complex due to the presence of a highly twisted retinal in the restricted chromophore binding pocket (Schick et al., 1987). Rapid thermal reactions that take only milliseconds then guide rhodopsin through a series of conformational intermediates, including bathorhodopsin and lumirhodopsin, before settling into an equilibrium of three different metarhodopsin intermediates. Metarhodopsin II is the only intermediate to have a deprotonated Schiff base linkage and it is considered the biologically active intermediate due to its efficient activation of the G protein transducin

(Bennett et al., 1982). This is supported by evidence showing that the presence of transducin shifts the metarhodopsin equilibrium in favor of metarhodopsin II (Emeis et al., 1982).

7 A single metarhodopsin II can activate hundreds of molecules of transducin, a heterotrimeric, peripheral membrane protein that consists of α, β, and γ subunits (Bohm et al., 1997). This causes an exchange of bound GDP for GTP, and leads to dissociation of the α subunit from the complex along with GTP, and allowing it to subsequently bind and activate cGMP phosphodiesterase [PDE] (Wensel and Stryer, 1986). Prior to photoactivation, a cGMP-mediated transmembrane cation channel allows entry of Na+ and Ca2+ into the photoreceptor outer segment, depolarizing the membrane. When activated by transducin, PDE hydrolyzes cGMP to GMP, disabling the ability of the cGMP to maintain the cationic channel in an open conformation (Fesenko et al. 1985).

With the cationic channel closed, intracellular Ca2+ levels are reduced leading to a hyperpolarization of the photoreceptor cell membrane (Baylor, 1996). This reduces the rate of release of neurotransmitter from the synaptic terminal of the photoreceptor cell, eliciting a response from second-order horizontal and bipolar cells (Lagnado and Baylor

1992).

Timely recovery from phototransduction is key to maintaining sensitivity, especially in rod photoreceptors, where rhodopsin activation causes significantly more signal amplification upon activation compared to the cone opsins (Chen et al., 2010). This recovery depends on deactivation of rhodopsin, the transducin-PDE complex, and increased synthesis of cGMP (Burns and Pugh, 2010). Within milliseconds of formation, metarhodopsin II is subjected to a two-step deactivation process, comprising C-terminal phosphorylation by rhodopsin kinase [GRK1] followed by arrestin binding, which quenches catalytic activity (Wilden et al., 1986; Gross and Burns, 2010). PDE activity

8 attenuates when GTP bound to the α transducin subunit is hydrolyzed, a process that is catalyzed by a regulator of G protein signaling [RGS9-1] and considered a rate limiting step in recovery of the rod photoresponse (He et al., 1998; Krispel et al., 2006). Finally, when intercellular Ca2+ levels decrease due to closure of cation channels following activation, guanylate cyclase activating proteins [GCAPs] become active, promoting guanylate cyclase activity, which in turn increases the formation of cGMP (Palczewski et al., 1994; Haeseleer et al., 1999).

1.3 The retinoid cycle

For rhodopsin to regenerate in rod photoreceptors in vivo, a new 11-cis-retinal ligand must form a Schiff base linkage with rhodopsin following release of all-trans-retinal

(Hofmann et al., 1992; Pulvermüller et al., 1997). The process required to enzymatically isomerize all-trans-retinal back to 11-cis-retinal, the retinoid cycle, comprises a series of metabolic steps in both rod outer segments [ROS] and the retinal pigment epithelium

[RPE] (Fig.3; Kiser et al., 2012). These events are preceded by the release of all-trans- retinal from activated rhodopsin, involving hydrolysis of the Schiff base linkage and dissociation from opsin, which can occur following either photoactivation or thermal activation (Luo et al., 2011). Because retinal can be extremely toxic to photoreceptor cells (Maeda et al., 2008; Maeda et al., 2009), the critical first step of the retinoid cycle involves a reduction of all-trans-retinal to all-trans-retinol, which takes place in the ROS, and is catalyzed by a retinol dehydrogenase, RDH12 (Chrispell et al., 2009). Unlike all- trans-retinal, all-trans-retinol is able to diffuse across the interphotoreceptor matrix

[IPM] that separates the ROS and the RPE, facilitated by the interphotoreceptor retinoid-

9 binding protein [IRBP] (Pepperberg et al., 1993). Once in the RPE, lecithin:retinol acyltransferase [LRAT] catalyzes a deprotonation of the hydroxyl group of retinol, allowing for the decomposition of a thioester intermediate into a retinyl ester (Mondal et al., 2000). The retinoid isomerase, RPE65, is responsible for converting retinyl esters to

11-cis-retinol, with a lack of proper RPE65 activity causing an accumulation of all-trans- retinyl esters in the RPE (Redmond et al., 1998). RDH5, an 11-cis-retinol dehydrogenase that forms a complex with RPE65 (Golczak et al., 2010), then mediates a second redox reaction, producing 11-cis-retinal, which is transferred back to the ROS by the IRBP and made available to regenerate ligand-free opsin.

Despite the fact that the canonical retinoid cycle services both rod and cone photoreceptors, a supplementary cycle also exists specifically for cones and works independently of the RPE (Wang and Kefalov, 2009). This second retinoid cycle likely fulfills the sudden requirement for regenerated chromophore in cones following phototransduction, which is much more rapid than that in rods (Wang and Kefalov,

2011). To bypass the RPE, isomerization of all-trans-retinol to 11-cis-retinol occurs in

Müller cells, catalyzed by dihydroceramide desaturase-1 [DES1], bypassing RPE65 activity (Kaylor et al., 2013). Moreover, while rod photoreceptors cannot complete the final oxidation reaction converting 11-cis-retinol to 11-cis-retinal, this process can be accomplished in cones (Jones et al., 1989).

10 1.4 Cells of the retina

Visual perception in vertebrates relies on five classes of retinal cells. Briefly, activation of a photoreceptor cell in the outer nuclear layer of the retina generates a signal in the form of reduced neurotransmitter release, which is relayed in a forward pathway to bipolar cells in the inner nuclear layer, and then to retinal ganglion cells; this signal can be modified through lateral connections with horizontal and amacrine cells (Sung and

Chuang, 2010). Each of these five cell types make specific contributions to the processing of the visual signal before the retinal ganglion cell submits this information to various regions of the brain, including the thalamus, hypothalamus, and mesencephalon.

The main role of photoreceptors is to convert an environmental light stimulus into a signal that can be interpreted by other retinal cells [phototransduction, see section 1.2].

To achieve a dynamic range of sensitivity spanning several orders of magnitude of light intensities, vertebrates have two distinct classes of photoreceptors, rods and cones, which can be distinguished on the basis the molecular components of their phototransduction cascade, their cellular structure and their electrophysiology (Ebrey and Koutalos 2001).

Several phototransduction proteins have both cone and rod isoforms. These isoforms seem to have co-evolved as a coherent system (Hisatomi and Tokunaga 2002) and there is evidence to suggest that they can be used interchangeably with either rod or cone opsins (Imai et al., 1997; Ma et al., 2001; Deng et al., 2013). This implies that the components of the phototransduction cascade that interact with opsins are highly conserved, which agrees with evidence suggesting that the signal transducing domains of rod and cone opsins are not substantially different (Carleton et al., 2005). The most

11 notable morphological difference between rod and cone photoreceptors is that rods have larger outer segments, allowing for increased capacity of visual pigment packing to improve sensitivity to small amounts of light. The electrophysiological contributions of rod and cone photoreceptors can be measured using electroretinograms [ERGs], which measure electrical responses of cells after delivery of an electrical stimulus. Reponses from cone and rod photoreceptors are different and their contributions to specific ERG components can be measured using the double flash protocol (Robson et al., 2003; Pinilla et al., 2004), however this method can lack resolution. Single photoreceptor cell recordings provide an alternative method of taking electrophysiological measurements with improved resolution (Luo and Yao 2005).

Both horizontal and bipolar cells receive input from photoreceptors, and are located in the outer plexiform layer of the retina, which serves as a boundary between the inner and outer nuclear layers. There are two main types of horizontal cells, H1 and H2, which can differ in axon length, synaptic connections, or both, depending on the organism

(Dowling, 2012). The interactions of these cells with both photoreceptor and bipolar cells, is critical for early visual processing, including the generation of receptive fields for bipolar cells and, subsequently, retinal ganglion cells (Thoreson and Mangel, 2012).

However, while horizontal cells help to mediate lateral connections within the outer plexiform layer, their processes do not extend to any other parts of the retina (Dacey,

1999). Instead, bipolar cells act as the output neurons for the outer plexiform layer, vertically transmitting the signal received from photoreceptor cells to retinal ganglion cells. There are three major classes of bipolar cells: rod, ON cone, and OFF cone

12 (Wässle, 2004). Both rod and ON cone bipolar cells contain glutamate receptors that close specific channel proteins when activated, meaning that a light-evoked response that reduces glutamate signaling from photoreceptors will lead to depolarization (Shen et al.,

2009). Conversely, OFF cone bipolar cells contain glutamate receptors that open channels when activated, leading to hyperpolarization in response to light stimuli. Spatial and temporal components of the visual signal have also been attributed to different subtypes of ON and OFF cone bipolar cells (DeVries, 2000), as well as distinct glutamate receptor subtypes located at bipolar dendrites (Li and DeVries, 2006).

Amacrine cells are located in the inner plexiform layer of the retina and are responsible for mediating upstream feedback and signal output with bipolar cells as well as forming lateral connections with retinal ganglion cells (Dowling, 2012). In this sense, the functional constraints of amacrine cells are in line with those of horizontal cells in the outer plexiform layer, whose processes do not extend to other parts of the retina. Despite these restrictions, signal processing in amacrine cells contributes to a number of aspects of visual perception, including edge detection and motion encoding (Baccus, 2007;

Famiglietti, 2005). Significant diversity exists in amacrine cell morphology and physiology, which has made them a challenging class of cells to organize resulting in a variety of competing classification schemes (Grimes, 2012).

Finally, ganglion cells are the final output neurons of the retina, transmitting a processed visual signal to other parts of the brain through the optic nerve. Similar to amacrine cells, ganglion cells are quite diverse, with some vertebrates having as many as 15

13 morphological subtypes, however the two most common classes are referred to as the midget and parasol ganglion (Polyak, 1941). Midget ganglion cells are the most common, and were assigned their designation due to the small sizes of their cell bodies and dendritic trees. These cells are usually connected to bipolar cells that connect to single cone photoreceptors, where one midget ganglion cell can eventually signal for illumination of the photoreceptor, and another can signal for a lack of illumination, designating them ON and OFF ganglion cells, respectively (Dowling, 1991). Midget ganglion cells are also known to respond to changes in color, such as the red-green opponent pathway in primates (Lee, 1996). Parasol ganglion cells have larger cell bodies than midget cells, and have a very distinct dendritic morphology (Dacey and Petersen,

1992). Parasol cells usually receive input signals from many rod and/or cone photoreceptors, providing them larger receptive fields than midget cells (Lee et al.,

1997). Unlike midget cells, parasol cells are not as involved in color vision, but are instead critical to motion detection and object detection at low contrast (Dobkins and

Albright, 1994; Lee and Sun, 2009).

While the above description of phototransduction relates to the process of image formation through light stimuli, recent evidence also suggests that photoreception can initiate in cells other than rods and cones. While it had been suggested for years that ganglion cells transmitted both image and non-image forming information to the brain, it was always assumed that both of these signals originated in rod and cone photoreceptors

(Do and Yau, 2010). Only in the last 20 years or so was intrinsic photoreception in some retinal ganglion cells shown to initiate non-image forming light signals associated with

14 circadian photoentrainment; experiments in both mice and human with degenerated rod and cone photoreceptors were still capable of adjusting their circadian systems to external light-dark cycles (Provencio et al., 1994; Czeisler et al., 1995). Around the same time, a novel opsin was discovered in the photosensitive dermal melanophores of the frog,

Xenopus laevis, which was named melanopsin (Provencio et al., 1998), and expressed in a small subset of retinal ganglion cells in the mammalian eye (Hattar et al., 2002;

Provencio et al., 2002). These ganglion cells project to a variety of brain regions serving non-image vision (Hannibal and Fahrenkrug, 2004; Hattar et al., 2006), and are also intrinsically photosensitive (Berson et al., 2002). Another example of novel photoreception takes place in a subset of horizontal cells in the retina of teleost fish, which are characterized by a depolarizing response to light (Jenkins et al., 2003). The absorbance peak of this response conforms with a vertebrate ancient opsin [VA opsin], previously isolated from the eye of Atlantic salmon and localized to horizontal cells via in situ hybridization (Soni et al., 1998). This intrinsic light response is thought to play a role in modulating lateral inhibition mediated by these cells, which suggests that photoreception initiated in retinal cells other than rods and cones may at least indirectly contribute to image formation (Cheng et al., 2009).

1.5 Heterologous opsin expression

An issue that arises when attempting to express membrane proteins using heterologous systems is how to generate sufficient amounts of protein to perform functional analyses.

GPCRs, such as opsins, are notoriously difficult to express due to the complex folding, trafficking and modifying mechanisms associated with the translation of membrane

15 proteins. This leads to significant amounts of misfolded protein either stranded in the

ER, moved to unsuitable transport vesicles, or missing post-translational modifications required for proper function (Sarramegna et al., 2003). Bovine rhodopsin is still the only vertebrate visual pigment with a high-resolution crystal structure of multiple conformations (Palczewski et al., 2000; Okada et al., 2004; Park et al., 2008; Scheerer et al., 2008; Choe et al., 2011), as large amounts of the native protein can be obtained from cow retina; other GPCRs with resolved crystal structures include squid rhodopsin

(Murakami and Kouyama, 2008) as well as the β1- and β2-adrenergic receptors

(Rasmussen et al., 2007; Warne et al., 2008). Since the majority of visual pigments and

GPCRs in general cannot be extracted at such high levels in their native state, finding competent expression systems is a top priority for studies requiring functionally active proteins.

Over the past several decades, experiments probing the structure and function of visual pigments have resorted to heterologous expression systems in order to generate visual pigment samples (Ablonczy et al., 2006; Yokoyama et al., 2008; Fig. 4). Most systems involve an expression vector being transiently transfected into one of a number of different mammalian cell types, including COS-1 (Matsumoto et al., 2006), HEK293

(Kuwayama et al., 2005), and HEK293T (Parry et al., 2004), using lipid-based transfection reagents. While this method is usually successful, even large-scale transfections are sometimes unable to produce serviceable visual pigment samples

(Pointer et al., 2007; Davies et al., 2007b).

16 Heterologous expression systems are not just dependent on cell type, but also on the expression vector used to transfect the cells. Many early expression studies of visual pigments employed the pMT expression vector, featuring the adenovirus major late

(AML) promoter. This choice was likely made because various versions of pMT had successfully expressed large proteins with post-translational modifications (Bonthron et al., 1986; Kaufman et al., 1989), suggesting it could handle the expression of complex membrane proteins. As pMT use became more frequent for visual pigment studies, newer versions were engineered to include the 1D4 epitope, which consists of nine C- terminal residues of bovine rhodopsin [TETSQVAPA] recognized by the 1D4 monoclonal antibody (Molday and MacKenzie, 1983); this allowed the immunoaffinity purification process to be streamlined (Franke et al., 1988; Tsutsui et al., 2007). Other promoters have since been used for visual pigment expression, such as the combination of the SV40 early promoter containing the R segment and part of the U5 sequence of the long terminal repeat of human T-cell leukemia virus type 1 (Kayada et al., 1995; Kojima et al., 2008), and the cytomegalovirus (CMV) promoter (Reeves et al., 2002). Recently, the CMV promoter was shown to drive higher levels of visual pigment expression when compared to the AML promoter (Morrow and Chang, 2010).

Other heterologous systems that rely on non-mammalian cells, including E. coli, yeast and insect cells, have been used regularly to express other GPCRs (Sarramegna et al.,

2006), but only sparingly to express visual pigments (Mollaaghababa et al., 1996;

Klaassen et al., 1999). These systems often strive for large-scale production of GPCRs at the expense of protein function, which can suffer due to differences in membrane

17 composition and posttranslational mechanisms of the host cells (Stanasila et al., 1998;

Opekarova and Tanner, 2003). Ultimately, mammalian cells provide the most authentic cell environment for vertebrate GPCRs and are often the system of choice when attempting functional characterizations of a protein of interest, even though the yields produced with mammalian cells tend to be lower than those of other systems (Junge et al., 2008). This fact emphasizes the importance of continuing to improve the efficiency of mammalian cell-based heterologous expression systems.

1.6 Visual pigment structure and function

A number of key amino acid residues and motifs have been identified that are integral to various aspects of visual pigment structure and function. The site of the protonated

Schiff base linkage between opsin and chromophore is located at K296 [all amino acid numbering is based on the sequence of bovine rhodopsin], in transmembrane helix 7

[TM7] (Bownds, 1967). In dark state visual pigments, the Schiff base is protonated and stabilized by a negatively charged counterion supplied by E113 (Nathans, 1990a); upon activation, the counterion switches to E181 (Lüdeke et al., 2005). The interaction of

K296 and E113 is also important when opsin is not bound to chromophore, as a lack of counterion leads to a constitutively active visual pigment (Robinson et al., 1992). During activation, the salt bridge between protonated Schiff base and E113 is broken when a proton is transferred from the former to the latter (Jäger et al., 1994). This is followed by the uptake of a proton from the solvent by E134 of the highly conserved E(D)RY motif in

TM3 (Mahalingham et al., 2008). E134 protonation is able to occur when the salt bridge it forms with R135 and E247 in the dark state, known as the ‘ionic lock’, is broken

18 (Laricheva et al., 2013). The NPXXY motif is located in TM7 and helps to establish a water-mediated hydrogen-bonding network, with D83 and W265, in dark-state rhodopsin

(Fritze et al., 2003). During activation, Y306 of NPXXY is able to swing into the interior of the protein, helping to extend the hydrogen-bonding network to residues at the cytoplasmic surface critical to G protein activation (Standfuss et al., 2011). Finally, visual pigments also undergo several different post-translational modifications, including glycosylation at N2 and N15 that is important for proper folding and signal transduction

(Kaushal et al., 1994), as well as palmitoylation at C322 and C323, which fastens cytoplasmic helix 8 to the intercellular surface of the plasma membrane to maintain proper rhodopsin structure (Ovchinnikov et al., 1988; Maeda et al., 2010).

Each of the five main visual pigment classes has evolved to have λMAX values within a specific range of the light spectrum when bound to 11-cis-retinal: sws1, 360-430 nm; sws2, 440-460 nm; rh2, 470-510 nm; lws, 510-560 nm; and rh1, ~500 nm (Ebrey and

Koutalos, 2001; Hisatomi and Tokunaga, 2002). Because the absorption spectra of all classes of visual pigments have the same shape when employing an identical

chromophore, this λMAX value is often the lone property used to describe the spectral sensitivity (Lamb, 1995). Interactions in the chromophore binding pocket between the opsin apoprotein and retinal, often facilitated by structural water molecules, determine

the λMAX value of a given visual pigment, therefore amino acid substitutions at residues

involved in these interactions can lead to shifts in λMAX, a process referred to as spectral tuning (Sekharan, 2009; Katayama et al., 2012).

19 For each class of visual pigment, a number key residues have been identified that can

mediate significant shifts in λMAX through site-directed mutagenesis studies that are often meant to replicate the natural variation that exists across vertebrates (Yokoyama, 2008).

As rhodopsin is the most characterized visual pigment and exclusively mediates dim-light vision, when naturally occurring substitutions are replicated in a bovine rhodopsin

background, they cause significant shifts in λMAX [some examples are D83N (Nathans,

1990b), E122Q (Sakmar et al., 1989), and A292S (Janz and Farrens, 2001)]. It is interesting to note that mutations in the opposite direction or created in a background other than bovine rhodopsin do not always reproduce a similar spectral shift (Fasick and

Robinson, 2000; Sugawara et al., 2010; Bickelmann et al., 2012).

Among the four classes of cone visual pigments, spectral tuning research is most prominent for sws1, due to its role in differentiating between violet and ultraviolet vision in birds (Hunt et al., 2001), and lws, because of the role the duplicated mws/lws genes play in colour vision in primates, including humans (Yokoyama and Radlwimmer, 2001).

While the main determinant of ultraviolet vision in mammals is F86, with other identities of site 86 providing violet-sensitive visual pigments (Cowing et al., 2002), ultraviolet vision in birds evolved through a different mechanism, involving C90, which doesn’t

depend on F86 (Wilkie et al., 2000). Meanwhile, the 33 nm λMAX difference between the human lws and mws opsins has been shown in mutagenesis studies to be accounted for by the blue-shifting substitutions S164A, Y261F, and T269A (Asenjo et al., 1994). The

RH2 opsins seem to rely mostly on sites 122 and 292 for major spectral shifts, similar to rhodopsin, with combinations of multiple small effect substitutions also producing more

20 notable shifts (Yokoyama et al., 1999; Chinen et al., 2005b; Takenaka and Yokoyama,

2007). Finally, the main variable sites that can alter λMAX in SWS2 visual pigments are

118 and 269 (Cowing et al., 2002; Yokoyama and Tada, 2003).

While visual pigments have long been known to differ in terms of their most scrutinized

property, λMAX, a closer examination of other biochemical properties has also identified several significant differences between rhodopsins and cone opsins. For example, hydroxylamine is a small molecule that is thought to enter the chromophore binding pocket and react with the Schiff base linkage to form a retinal oxime compound with the chromophore, leaving the opsin protein without a ligand (Johnson et al. 1993); this retinal oxime absorbs light at a maximal wavelength of about 363 nm, therefore this reaction can be monitored via absorbance spectrophotometry (Wang et al. 1992). Because the binding pocket in rhodopsin is more highly structured and restrictive than in cone opsins, the former experiences little to no effect when incubated with hydroxylamine (Okano et al.,

1989), while the latter allows hydroxylamine to sequester retinal relatively quickly

(Starace and Knox 1998; Fasick et al., 1999).

Cone opsins are also known to have faster meta II decay rates and retinal regenerations rates compared to rod visual pigments, which is consistent with the idea that cone opsins are less photosensitive but have a faster photoresponse relative to rod opsins (Shichida et al., 1994; Kojima et al., 1995; Imai et al., 1997). Site-directed mutagenesis studies have implicated some of the sites that may be responsible for these differences. Site 122 located in TM3 is conserved in rod opsins as glutamate, but is not conserved in cone

21 opsins, having various identities including glutamine and isoleucine. Mutating site 122 to glutamate in cone opsins reduces meta II decay and retinal regeneration rates, while these rates are increased when mutating site 122 in rod opsins (Imai et al., 1997). Site 122 has also been examined in vivo; transgenic mice expressing E122Q rhodopsin showed lower photosensitivity and lower meta II lifetime than wild type mice (Imai et al., 2007).

Another site, 189, is part of a cytoplasmic, anti-parallel β-sheet in the 2nd extracellular loop that might help to shield the chromophore from the extracellular environment

(Palczewski et al., 2000; Okada et al., 2004). This site is highly conserved as a proline residue in cone opsins, but is usually represented by isoleucine in rod opsins. Mutating site 189 to proline in rod opsins increases meta II decay and retinal regeneration rates, while these rates are reduced when mutating site 189 to isoleucine in cone opsins

(Kuwayama et al., 2002). That these are the only two sites responsible for the differences between rod and cone opsins is highly improbable, but additional research has yet to identify other key sites.

While the drastic differences between molecular properties of rhodopsin and cone opsins have been thoroughly established, there have been relatively few studies that investigate

distinctions among rhodopsins other than shifts in λMAX. Earlier studies focusing on model vertebrate rhodopsins highlighted very similar metarhodopsin intermediate kinetics in bovine, human, and chicken rhodopsin, with a slightly faster meta III decay rate in humans (Imai et al., 1995; Lewis et al., 1997). More recently, the meta II decay of pufferfish rhodopsin was also measured to be similar to bovine and chicken rhodopsin

(Tarttelin et al., 2011), while meta II formation rates in African cichlids were shown to be

22 shorter in fish with N83 compared to those with D83 (Sugawara et al., 2010). A recent characterization of echidna rhodopsin revealed a faster retinal release rate than bovine rhodopsin, demonstrating that not all mammalian rhodopsins have similar molecular properties (Bickelmann et al., 2012). Additionally, while sensitivity to hydroxylamine has traditionally been used to determine whether a visual pigment was functionally rhodopsin- or cone opsin-like, there are some examples of rhodopsins that do react when exposed to hydroxylamine, including the anole lizard and the echidna (Kawamura and

Yokoyama, 1998; Bickelmann et al., 2012).

1.7 Zebrafish

Given that visual pigments can directly influence molecular, physiological, genetic, and behavioral aspects of vision, studying them provides a unique opportunity to explore a number of neuroscience disciplines. Therefore, an ideal system in which to study visual pigments should be well studied in all of these fields across a series of developmental stages. For this reason, zebrafish [Danio rerio] is an ideal experimental system to study visual pigment structure and function. Additionally the zebrafish visual system has many similarities with those of other vertebrates, and there is a wealth of existing information on its visual processing from a variety of biological fields (Bilotta and Saszik, 2001).

Zebrafish is a member of the family Cyprinidae, the most speciose family of the order

Cypriniformes, and belonging to the second largest superorder of fish, Ostariophysi.

Ostariophysi, along with Acanthomorpha, make up the two dominant components of the class Actinopterygii, representing ray-finned fish and comprising nearly 99% of all fish

23 species. Relationships within Actinopterygii have come into question with molecular data, with some evidence suggesting a sister relationship between Ostariophysi and

Clupeomorpha; the taxon Ostarioclupeomorpha, also called Otocephala, is used to describe this putative monophyletic group (Saitoh et al., 2003).

Zebrafish have an impressive array of visual pigment genes, including one rh1, four rh2, one sws2, one sws1 and two lws (Chinen et al., 2003). Additionally, zebrafish, along with other teleosts, are the only vertebrates known to have both a visual rhodopsin and a non-visual rhodopsin-like gene, exo-rhodopsin, expressed in the light-sensitive pineal gland (Mano et al., 1999). Curiously, exo-rhodopsin is orthologous to rhodopsins of non- teleost vertebrates, while rhodopsin in teleosts arose following a retrotransposition event, and contains no introns (Fitzgibbon et al., 1995). This duplication occurred early in the evolution of ray-finned fish, as basal Actinopterygians such as the sturgeon [Chondrostei] and gar [Holostei] also have intronless rhodopsin genes (Bellingham et al., 2003). That would place this duplication event no sooner than 284 million years ago, which marks the divergence of these fish from teleosts (Hurley et al., 2007). Another rhodopsin-like gene, rh1-2, was also recently identified in the retina of juvenile and adult zebrafish, along with several other cyprinid fish. While this novel pigment can be activated by light when

regenerated with 11-cis-retinal, and produces an absorption spectrum with a λMAX value of approximately 500 nm, limited functional characterization prevented its classification as either visual or non-visual (Morrow et al., 2011). Meanwhile, among cone opsins, a common feature between the expansion of both rh2 and lws gene families is the

diversification of λMAX, providing optimal sensitivity at a range of wavelengths. Along

24 with differential expression patterns, this allows duplicated rh2 and lws genes to provide distinct spectral sensitivities within the retina, both spatially and temporally (Takechi and

Kawamura, 2005).

1.8 Thesis objectives

The primary objective of this thesis is to investigate the evolution of zebrafish visual pigment structure and function, using mutagenesis, in vitro expression, and spectroscopic methods to assay pigment function. The specific goals of my research are:

1. Investigate a novel rhodopsin-like gene recently discovered in the zebrafish

genome, rh1-2, and characterize its expression pattern and function.

2. Explore the evolutionary history of rh1-2 to better understand its origin and to

determine how common this duplicate is among Actinopterygian fish.

3. Investigate the importance of rh1-2 to zebrafish photoreception, including the

possibility of heterodimerization with rhodopsin.

4. Functionally characterize zebrafish rhodopsin using a variety of spectroscopic

assays, as fish rhodopsins have yet to be investigated as thoroughly as

mammalian rhodopsins.

5. Investigate the mechanisms underlying differences in retinal release in

zebrafish and bovine rhodopsins through a comprehensive mutagenesis study.

6. Demonstrate the advantages of exploring natural sequence variation to better

understand structural and functional differences in visual pigments.

25 7. Explore hypotheses to explain the retention of numerous duplicated rh2 opsin

genes in teleost fish.

8. Investigate the degree to which the identity of site 122 can influence functional

differentiation in rh2 opsins

1.9 Thesis overview

A number of key studies of visual pigments helped to establish zebrafish as a model organism for the vertebrate visual system (Chinen et al., 2003; Takechi and Kawamura,

2005; Tsujimura et al., 2007). This thesis seeks to build on this foundation to further characterize visual pigment function in zebrafish, and to continue to establish zebrafish as a model for visual pigment study, similar to bovine and chicken. Some studies of non- model rhodopsins have reported more functional variation among vertebrate rhodopsins than previously anticipated (Kawamura and Yokoyama, 1998; Bickelmann et al., 2012).

This suggests a need for more thorough characterizations of visual pigments from a wider phylogenetic distribution of species, ideally non-mammalian model organisms that already serve as models of the visual system, like zebrafish. My thesis consists of a series of experiments designed to increase our understanding of the visual system of zebrafish. One of the key functional assays of my thesis measures the rate of the release of retinal following photoactivation using fluorescence spectroscopy (Farrens and

Khorana, 1995). This assay has been used in a number of key studies of visual pigment structure and function (Yan et al., 2002; Chen et al., 2002; Piechnick et al., 2012), and is associated with the rate of decay of the photo-activated metarhodopsin intermediates. I will utilize this assay as a means to perform comparative analyses of kinetic rates within different groups of opsins.

26 Chapter 2 of my thesis presents a novel rhodopsin-like gene that we identified in zebrafish, rh1-2. While zebrafish, like many other teleosts, possesses an impressive complement of both visual and non-visual opsins, it is quite rare for any organism to possess multiple visual rhodopsins. We performed an initial characterization of rh1-2 function to verify that it was a functional opsin, capable of binding 11-cis-retinal and activating in response to light. Like rhodopsin, rh1-2 is expressed in the retina and has a

λMAX value of approximately 500 nm, however rh1-2 is also much less stable and expressed later in development. While rh1-2 was sequenced from three other fish from the same family [Cyprinidae], phylogenetic analyses suggest a more ancient origin within the teleost lineage. The results of our initial characterization of rh1-2 were published in

Visual Neuroscience: Morrow JM, Lazic S, Chang BS (2011) A novel rhodopsin-like gene expressed in zebrafish retina. Vis Neurosci 28, 325-335.

After publishing this initial characterization of rh1-2, there were still several key questions that remained about the function and purpose of rh1-2. Chapter 3 sets out to address some of these questions, by attempting to elucidate key aspects of rh1-2 function that would help to establish its role as either a visual or non-visual opsin. In some cases, we also decided to assay either rhodopsin or exo-rhodopsin to establish both visual and non-visual rhodopsin controls to which we could compare our data for rh1-2. By establishing that rh1-2 is expressed in the photoreceptor cell layer of the zebrafish retina, and releases retinal at a rate similar to rhodopsin, but several times slower than exo- rhodopsin, we conclude that it is likely a second visual rhodopsin. Also, after noting that expression of rhodopsin and rh1-2 may overlap in a subset of peripheral photoreceptors,

27 we performed preliminary analyses that suggest the presence of rh1-2 may alter the thermal stability of rhodopsin in vitro, possibly through heterodimerization. This chapter provides evidence to form initial hypotheses surrounding the role of rh1-2 and provides some direction for future studies.

Following the resolution of the biologically active metarhodopsin II crystal structure

(Choe et al., 2011), several recent studies have investigated conserved amino acids that are important to establishing rates of retinal release in rhodopsin following photoactivation. In Chapter 4, we add to this field of research by investigating the contribution of variable sites to retinal release, using zebrafish rhodopsin as a model system. As a poikilotherm, zebrafish is likely to have variation in kinetic rates relative to homeothermic mammals. Therefore, we expected to find natural variation in kinetic rates, such as retinal release, between zebrafish and bovine rhodopsin. After determining that zebrafish rhodopsin does in fact release retinal more than twice as fast as bovine rhodopsin at 20oC, the phenotypic difference between the retinal release rates of these opsins was mapped to 8 variable sites found across several motifs. Based on the proximity of most of these sites to the β-ionone ring of retinal, we propose that the natural variation of retinal release among rhodopsins is primarily mediated through steric effects that influence dissociation of retinal from opsin. This complements other studies that attribute conserved residues as being mostly responsible for maintaining stability of the Schiff base (Piechnick et al., 2012), which must be hydrolyzed before retinal can dissociate from opsin.

28 While most groups of vertebrates possess at most a single copy of the rod-like opsin gene, rh2, this gene family has experienced a significant expansion in Actinopterygian fish. Chapter 5 investigates this expansion and notes that many species with multiple rh2 genes have at least one copy with Q122 and one with E122. This site is of particular interest because it has previously been highlighted as a determinant of rhodopsin and cone opsin function, and changing its identity can have significant consequences to opsin function. We have performed an initial characterization of zebrafish rh2-1 and rh2-4, which have Q122 and E122, respectively. Preliminary results suggest that site 122 can

alter the λMAX, retinal release, and hydroxylamine stability of rh2 opsins. This may imply that a functional dichotomy of rh2 genes exists in teleost fish, mediated by site 122, and may provide the basis of an explanation for why these fish have conserved so many rh2 gene duplicates.

This thesis contributes substantially to the characterization of visual pigment function in zebrafish. These experiments have also increased the understanding of functional variation within rhodopsins and cone opsins, particularly by investigating the molecular mechanisms of retinal release, and the roles of duplicated opsin genes. My hope is for this body of work to serve as an important non-mammalian reference against which future studies can be compared and contrasted, further cementing Danio rerio as an important model organism for the vertebrate visual system.

Figure 1. A schematic of the phylogenetic relationships among the five classes of vertebrate visual pigments: four cone opsin classes [RH2, SWS2, SWS1, M/LWS], and a single rod opsin class [RH1]. Wavelength values represent the range of λMAX values for each class of visual pigments.

Figure 2. A schematic of the phototransduction cascade in a rod outer segment. Briefly, rhodopsin [R] is activated [R*] by a photon of light, allowing it to bind the heterotrimeric G protein, transducin [G]. Once bound to R*, transducin replaces bound GDP with GTP, which dissociates the activated α subunit [G*]. Two G* bind a nearby phosphodiesterase [PDE], whose activity hydrolyzes cGMP. When levels of cytosolic cGMP are reduced, cGMP-gated ion channels close, preventing further influx of Na+ and Ca2+. This leads to hyperpolarization of the cell membrane, thus reducing levels of neurotransmitter release (adapted from Leskov et al., 2000).

Figure 3. A schematic of the various steps of the retinoid cycle. Briefly, in the rod outer segment [ROS], rhodopsin is activated by a photon of light, causing photoisomerization of 11-cis-retinal to all-trans-retinal and leading to its release from opsin. Reduction of all-trans-retinal to all-trans-retinol is catalyzed by RDH12, an all-trans-retinol dehydrogenase. Upon diffusion through the interphotoreceptor matrix [IPM] to the retinal pigment epithelium [RPE], facilitated by the interphotoreceptor retinoid-binding protein [IRBP], all-trans-retinol esterification is catalyzed by lecithin:retinol acyltransferase [LRAT]. The retinoid isomerase, RPE65, converts retinyl esters to 11-cis- retinol, which is further oxidized to 11-cis-retinal by a retinal dehydrogenase, RDH5. Newly formed 11-cis-retinal diffuses across the IPM, once again facilitated by the IRBP, and once back in the ROS is able to bind free opsin to form rhodopsin (adapted from Kiser et al., 2012).

Figure 4. The major steps involved in generating and assaying visual pigment samples using a heterologous expression system. An expression vector containing the full-length coding sequence of a visual pigment of interest is used to transiently transfect a mammalian cell culture. After several days of growth, visual pigment is harvested from cells, regenerated with 11-cis-retinal, and immunoaffinity purified. This visual pigment sample can now be assayed using spectroscopy.

33 References

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Chapter 2:

A novel rhodopsin-like gene expressed in

zebrafish retina

51 2.1 Abstract

The visual pigment rhodopsin [rh1] constitutes the first step in the sensory transduction cascade in the rod photoreceptors of the vertebrate eye, forming the basis of vision at low light levels. In most vertebrates, rhodopsin is a single copy gene whose function in rod photoreceptors is highly conserved. We found evidence for a second rhodopsin-like gene

[rh1-2] in the zebrafish genome. This novel gene was not the product of a zebrafish- specific gene duplication event, and contains a number of unique amino acid substitutions. Despite these differences, expression of rh1-2 in vitro yielded a protein that not only bound chromophore, producing an absorption spectrum in the visible range

[λmax ≈ 500 nm], but also activated in response to light. Unlike rh1, rh1-2 is not expressed during the first 4 days of embryonic development; it is expressed in the retina of adult fish, but not the brain or muscle. Similar rh1-2 sequences were found in two other Danio species, as well as a more distantly related cyprinid, Epalzeorhynchos bicolor. While sequences were only identified in cyprinid fish, phylogenetic analyses suggest an older origin for this gene family. Our study suggests that rh1-2 is a functional opsin gene that is expressed in the retina later in development. The discovery of a new, previously uncharacterized opsin gene in zebrafish retina is surprising given its status as a model system for studies of vertebrate vision and visual development.

52 2.2 Introduction

Opsins are members of the G protein-coupled receptor [GPCR] superfamily that is distinguished by a Schiff base linkage between a conserved lysine residue and a retinaldehyde chromophore (Ebrey and Koutalos, 2001). The photosensitive chromophore absorbs light to activate the opsin protein, initiating a signaling cascade in the photoreceptor cell (Burns and Baylor, 2001; Shichida and Morizumi, 2007). Gene duplication events throughout vertebrate evolution have given rise to several opsin families. There are five groups of visual opsins, including rhodopsin [rh1] and four cone opsin groups: rhodopsin-like [rh2], short wavelength-sensitive 1 [sws1], short wavelength-sensitive 2 [sws2], and long wavelength-sensitive [lws]. Visual opsins are expressed in photoreceptor cells of the retina and, while being primarily responsible for initiating the visual transduction cascade (Menon et al., 2001), are also involved in non- visual processes (Altimus et al., 2008; Lall et al., 2010). Non-visual opsins, such as pinopsin, melanopsin and exo-rhodopsin, are expressed in a wide variety of tissues and thought to only be involved in non-visual, light-dependant biological processes, such as the circadian system (Newman et al., 2003; Pierce et al., 2008; Peirson et al., 2009).

GPCRs comprise one of the largest known families of integral membrane proteins.

Rhodopsin [rh1], a highly characterized GPCR, is the visual opsin that initiates dim-light vision in vertebrates (Okawa and Sampath, 2007). Because of its stability and abundance in rod photoreceptor outer segments, rhodopsin is amenable to detailed studies of structure and function, often through site-directed mutagenesis and in vitro expression

(Imai et al., 2007; Knierim et al., 2007; Nickle & Robinson, 2007). Rhodopsin was also

53 the first opsin with a high-resolution crystal structure of its dark state (Palczewski et al.,

2000), with crystal structures of both its activated (Scheerer et al., 2008) and chromophore-free (Park et al., 2008) states also having been resolved. Due to high levels of sequence and structural conservation among GPCRs (Rosenbaum et al., 2009), data collected and methods developed on rhodopsin have also facilitated further studies on other visual and non-visual opsin families (Teller et al., 2003; Ramon et al., 2009).

It is rare for a vertebrate to have more than one rh1 gene. Despite two ancestral genome duplication events that are thought to predate the vertebrate radiation (Dehal and Boore,

2005), the majority of vertebrates maintain only a single copy of rh1. In contrast, gene numbers of some cone opsin groups can vary greatly in vertebrates. No mammalian rh2 opsins have been identified (Trezise and Collin, 2005; Bowmaker, 2008), while many other species have up to four rh2 genes (Chinen et al., 2003; Parry et al., 2005;

Matsumoto et al., 2006). Derived primates have experienced lws opsin duplication

(Dulai et al., 1999), as have teleost fish, where it is not uncommon to have at least four lws genes (Weadick and Chang, 2007; Owens et al., 2009). However, having multiple rh1 genes is a very unique trait. Comprising 96% of living fish species due to an adaptive radiation after a genome duplication event 150 million years ago (Taylor et al.,

2003), teleosts are the only known group of vertebrates containing species expressing multiple rh1 genes, although most are still believed to have only one. One example is the eel, having multiple species that express a second rh1 gene during maturation, coupled with its migration from freshwater to seawater (Beatty, 1975; Hope et al., 1998; Zhang et al., 2000; Zhang et al., 2002). Another example is the short fin pearleye [Scopelarchus

54 analis], a deep-sea teleost that expresses up to two rh1 genes depending on its lifecycle

(Pointer et al., 2007). Our investigation of the genome of zebrafish [Danio rerio], a well- studied teleost and model vertebrate, revealed an open reading frame for a previously uncharacterized rh1-like gene, suggesting yet another case of multiple rh1 genes in a species of teleost fish.

With version Zv9 of the zebrafish genome project having recently been made available, researchers are increasingly identifying and characterizing zebrafish genes and gene families of interest, including opsins. So far, nine visual opsins have been identified in zebrafish, including rh1, four rh2, sws2, sws1 and two lws (Chinen et al., 2003), a large complement even among teleosts. Additionally, many non-visual opsins have been discovered in zebrafish, including exo-rhodopsin (Mano et al., 1999), two melanopsins

(Bellingham et al., 2002; Bellingham et al., 2006), teleost multiple tissue [tmt] opsin

(Moutsaki et al., 2003), and two vertebrate ancient long [VAL] opsins (Kojima et al.,

2008); a non-annotated sequence for encephalopsin [opn3] has also been highlighted in the zebrafish genome. The advantage of studying opsin genes in the zebrafish is not only due to the availability of a genome sequence, but also to the considerable body of work assembled on the zebrafish visual system (Bilotta and Saszik, 2001; Neuhauss, 2003;

Fleisch and Neuhauss, 2006; Fadool and Dowling, 2008), and to the variety of existing and emerging tools available to study genes of interest in zebrafish (Amsterdam and

Becker, 2005; Amacher, 2008; Halpern et al., 2008).

55 In this study, we used zebrafish as a model to identify and characterize a novel rh1 gene.

Data mining of the zebrafish genome revealed an open reading frame encoding a novel rhodopsin gene, herein referred to as rh1-2. The amino acid sequence of rh1-2 contains several unique amino acid substitutions compared to normally conserved rh1 residues, and some substitutions more characteristic of cone opsins. Despite these substitutions, the rhodopsin-like protein was successfully expressed in vitro, and not only bound

chromophore, yielding an absorption spectra in the visible range [λmax ≈ 500 nm], but also activated in response to light, indicative of a functional opsin. In contrast to rh1, rh1-2 is not expressed during the first four days of embryonic development, but is expressed by day 21 and is expressed in the adult retina, but not in adult brain or muscle tissue. While rh1-2 sequences have only been identified in cyprinid fish, phylogenetic analyses suggest a much older origin. The novel gene is also intronless, similar to other rh1 sequences of teleosts. The discovery of a new, previously uncharacterized opsin gene in zebrafish retina is surprising given its status as a model system for studies of vertebrate vision and visual development.

2.3 Materials and methods

2.3.1 Opsin sequences

The zebrafish genome assembly, currently in version Zv9

(http://www.sanger.ac.uk/Projects/D_rerio/) was explored through a BLAST search using the full-length sequence of zebrafish rh1 [GenBank: AB087811] to identify putative rh1 homologs; various BLAST homology algorithms were employed to ensure the best possible coverage, including blastn, blastp, and tblastn. Genomic DNA was extracted

56 from fish tissues using the DNeasy Blood & Tissue Kit (QIAGEN). Primers were designed to amplify fragments of the coding region of the rh1-2 gene from cyprinid fish, including zebrafish, using genomic DNA as a template (Table S1). RNA was extracted from fish eyes using the TRIzol reagent (Invitrogen) and cDNA libraries were generated using the SMART cDNA Library Construction Kit (BD Biosciences). Teleost rh1 gene fragments were amplified from these cDNA libraries using either custom rh1 primers

(Table S1) or previously designed acanthomorph rh1 primers (Chen et al., 2003). PCR was performed using PfuTurbo (Stratagene) or FastStartTaq (Roche), with resulting bands being cloned into the pJET1.2 cloning vector (Fermentas) and sequenced using a

3730 DNA Analyzer (Applied Biosystems).

2.3.2 Protein expression

Complete coding sequences of zebrafish rh1 and rh1-2 were cloned into the p1D4-hrGFP

II expression vector (Morrow and Chang, 2010). This vector was used to transiently transfect cultured HEK293T cells using Lipofectamine 2000 (Invitrogen; 12 ug of DNA per 10-cm plate). Cells were harvested 48 hours post transfection and regenerated using

11-cis-retinal, generously provided by Dr. Rosalie Crouch (Medical University of South

Carolina). The pigments were then solubilized in 1% dodecyl maltoside, and immunoaffinity purified with the 1D4 monoclonal antibody (Molday & MacKenzie,

1983) as previously described (Chang et al., 2002; Morrow and Chang, 2010). The UV- visible absorption spectra of purified zebrafish rh1 and rh1-2 were recorded at 25oC using the Cary4000 double-beam spectrophotometer (Varian). For zebrafish rh1-2, a difference

57 spectrum was calculated as the difference between dark and light-bleached spectra after bleaching for 60 seconds.

2.3.3 Localizing expression of zebrafish rh1-2

RNA was extracted from zebrafish embryos [1, 2, 3, 4 dpf], juvenile heads [21 dpf] and adult tissues [retina, eye, brain & muscle] using TRIzol Reagent (Invitrogen) and cDNA libraries were generated using the SMART cDNA Library Construction Kit (BD

Biosciences), with identical amounts of RNA used as template in all reactions. Genomic

DNA was digested with DNAses during RNA extraction to avoid template contamination. Primers were designed to amplify ∼300 bp fragments of zebrafish rh1 and rh1-2; primers for rh1-2 were designed specifically to avoid amplification of rh1

(Table S1). Primers for β-actin, used as a positive control for RT-PCR experiments, were obtained from a previous study (Wang et al., 2003). PCR was performed using

PfuTurbo (Stratagene) under standard cycling conditions, with resulting bands being cloned into the pJET1.2 cloning vector using the CloneJET PCR cloning kit (Fermentas) and sequenced using a 3730 DNA Analyzer (Applied Biosystems).

2.3.4 Phylogenetic analyses

All rh1 and rh1-2 nucleotide sequences isolated from zebrafish, D. roseus [purple danio],

D. albolineatus [pearl danio] and E. bicolor [red tail shark] in this study were aligned with other teleost rh1 sequences obtained from NCBI (Table S2) using ClustalW (Larkin et al., 2007). Teleost exo-rhodopsin sequences were included as outgroups, in order to root the tree. Due to the short length of some of the teleost rh1 sequences obtained from

58 NCBI, this aligned data set was trimmed to include only nucleotides 94-912 and subjected to phylogenetic analyses using both maximum likelihood and Bayesian methods (Guindon and Gascuel, 2003; Ronquist and Huelsenbeck, 2003). Maximum likelihood phylogenetic methods were implemented in the program PHYML 3.0

(Guindon and Gascuel, 2003; Guindon et al., 2005), under the HKY+I+G model. For the likelihood analyses, bootstrapping methods were used to assess the degree of confidence in nodes of the phylogeny (Felsenstein, 1985). Bayesian inference was performed in

MrBayes 3.1.2 (Ronquist and Huelsenbeck, 2003) under the HKY+I+G model. To identify the most appropriate models for the molecular analyses, we used MrModeltest

2.3 (Nylander, 2004) and selected the best models favored under the Akaike Information

Criterion [AIC].

Two independent analyses were performed, each composed of 4 Markov chains with default heating values. Markov chains were run for 10 million generations, sampling trees (and parameters) every 1000 generations. Convergence was assessed using a number of methods. The average standard deviation of split frequencies, as calculated in

MrBayes, were all well below 0.01 at stationarity. Also implemented in MrBayes, a convergence diagnostic for branch length posterior probabilities, the potential scale reduction factor [PSRF], roughly approached 1 as the runs converged (Gelman and

Rubin, 1992). Convergence to stationarity was also assessed by plotting log-likelihood scores and other parameter values in the program Tracer 1.4.1 to ensure that there were no trends in the data post burn-in (Rambaut and Drummond, 2007). Finally, adequacy of mixing was assessed by examining acceptance rates for parameters in MrBayes, and by calculating in Tracer the effective sample sizes [ESS], the number of independent

59 samples from the marginal posterior distribution for each parameter; higher values being indicative of better sampling from the posterior distribution. These values were all well above 100. By these measures convergence was achieved within the first 25% of trees sampled, which were discarded as burn-in, and remaining trees were taken as representative of the posterior probability distribution.

2.4 Results

2.4.1 Sequence analysis of rh1-2

Zebrafish rh1-2 shares 73% nucleotide and 78% amino acid sequence similarity to zebrafish rh1, and, like other teleost rh1 genes, is intronless (Fitzgibbon et al., 1995). In general, the novel opsin gene shares much greater amino acid sequence similarity with the other visual opsins of zebrafish [38-78%] as opposed to its non-visual opsins [25-

32%]; the only exception is the non-visual exo-rhodopsin, paralog to mammalian Rh1 genes, which shares 70% amino acid sequence similarity with rh1-2 (Fig. 1a). Zebrafish rh1-2 is not a tandem duplicate of rh1, as the two genes are located on chromosomes 11 and 8 respectively. Additionally, rh1-2 is not likely the result of a whole genome duplication, due to a lack of synteny between chromosomes 8 and 11 (Catchen et al.,

2011). Full-length rh1-2 coding sequences were amplified from two other species from the Danio genus: Danio roseus and Danio albolineatus, while partial coding sequence was amplified from the cyprinid Epalzeorhynchos bicolor (Fig. 1b); partial rh1 coding sequences were also isolated for these three fish, with all rh1 and rh1-2 sequences amplified in this study being deposited in GenBank under accession nos. HQ286326-32.

60 The rh1-2 gene family contains a series of amino acid substitutions that suggest it might have properties different from other rhodopsins. The most unique substitutions are sites

C51, K145, Q150, L151, E199 and S316, all of which are conserved in all four rh1-2 sequences and not found in any other opsin gene family. There is also a group of residues, L45, N65, S/T149 and L183, whose identities are usually found in cone opsins or non-visual opsins, but rarely in rhodopsins (Table 1). All of these residues group into one of three distinct opsin motifs: helix 1/helix 8, cytoplasmic loop 2, or extracellular loop 2 (Fig. 2).

In addition to unique and cone-like substitutions, rh1-2 contains many of the sites and motifs required for visual opsin function. Both K296, the lysine residue that forms a

Schiff base linkage with the chromophore (Zhukovsky et al., 1991), and E113, the glutamate counter-ion to the Schiff base linkage (Sakmar et al., 1989) are present. Both

E134 and R135 of the D/ERY motif are present, being highly conserved in opsins and involved in light-dependant proton uptake (Arnis et al., 1994) and transducin activation

(Fanelli and Dell'Orco, 2008). Cysteine residues C110 and C187, required for the formation of a critical disulfide bond (Davidson et al., 1994; Hwa et al., 1999), are present as well as conserved glycosylation sites N2 and N15 (Kaushal et al., 1994;

Murray et al., 2009), and palmitoylation sites C322 and C323 (Traxler and Dewey,

1994).

61 2.4.2 In vitro expression

Complete coding sequences of zebrafish rh1 and rh1-2 were cloned into the p1D4-hrGFP

II expression vector. These genes were expressed in mammalian cell culture, regenerated with 11-cis-retinal, solubilized with dodecyl maltoside, and purified with the 1D4 monoclonal antibody as previously described (Chang et al., 2002; Morrow and Chang,

2010). Zebrafish rh1 was expressed and purified with relative ease (Fig. 3a) and produced spectra similar to previous studies (Chinen et al. 2003). Zebrafish rh1-2 was more difficult to express, but still bound 11-cis-retinal and produced a stable

photopigment with an absorption maximum [λmax] of approximately 500 nm (Fig. 3b), a

typical λmax value among fish rhodopsins (Johnson et al., 1993; Yokoyama et al., 1995;

Chang et al., 2009). When bleached with light, the absorption peak of zebrafish rh1-2 shifted to approximately 380 nm, characteristic of the biologically active meta II state of

visual opsins. Both expression level and absorbance ratio [A280/AMAX] of zebrafish rh1-2 were significantly lower than those of zebrafish rh1, suggesting that the former is less stable and more likely to misfold than the latter. These results imply that, while unique substitutions in zebrafish rh1-2 might reduce its stability, it can be expressed, has an

appropriate λmax for a teleost rhodopsin, and properly activates when bleached with light.

2.4.3 Zebrafish rh1-2 expression pattern

The temporal and spatial expression pattern of zebrafish rh1-2 was investigated using

RT-PCR. Expression of zebrafish rh1-2 was not detected during the first four days of embryonic development, but was detected in the heads of juveniles by 21 days post fertilization. Expression of rh1-2 was also detected in the adult retina, but not in the adult

62 brain or muscle (Fig. 4). Zebrafish rh1 expression was detected as early as 2 days post fertilization and strictly in the adult eye and retina, as previously reported (Raymond et al., 1995; Takechi and Kawamura, 2005). These results suggest that zebrafish rh1-2 is not expressed as early as rh1 in zebrafish development.

2.4.4 Phylogenetic analyses

Maximum likelihood and Bayesian phylogenetic analyses were performed in order to investigate the origins of the rh1-2 gene family, and its relationship to rh1 visual pigments. These analyses indicate that all four rh1-2 sequences isolated from cypriniform fish form a well-supported, monophyletic group, and that this novel gene was the result of a duplication that most likely occurred within the rh1 gene family during teleost evolution (Fig. 5). Although the rh1-2 genes form a well-defined group, due to the low support for many of the early teleost rh1 lineages, it remains difficult to pinpoint exactly when in teleost evolution this novel gene family may have emerged.

With respect to teleost rh1 sequences, this phylogeny recovers many of the commonly accepted systematic relationships among ray-finned fish (Forey et al., 1996; Hurley et al.,

2007; Wang et al., 2007). This includes support for both acanthopterygian and ostariophysian groups of fishes, although positions of the dory, cod and pearl eye rh1a sequences are somewhat unresolved. With respect to duplications within teleost rh1 genes, it is interesting that our analyses do not show strong support for a grouping of the rh1-2 sequences with the duplicate pearl eye and eels sequences. This might suggest multiple gene duplications within teleost rh1 genes, although due to the low support of many of these divergences, at the moment this is difficult to conclude with certainty.

63 Interestingly, in our analyses the pearl eye rh1b sequence does appear to group, albeit with variable support, with the duplicate copies of the eel rh1 sequences.

2.5 Discussion

In this study, we investigate rh1-2, a new family of opsin genes discovered in cyprinid fish. All four rh1-2 sequences form a monophyletic group within the rh1 clade of actinopterygian fish. While being most closely related to the rh1 family of visual opsins, rh1-2 has multiple substitutions that are either cone opsin-like in nature or unique to the rh1-2 gene family. Despite these unique substitutions at otherwise highly conserved sites, zebrafish rh1-2 was successfully expressed in vitro, and found to bind 11-cis-retinal with an absorption maximum around 500 nm; when bleached with light, it activated and changed conformation to its meta II intermediate. This activity coincides with the presence of conserved amino acids required for opsin function in the rh1-2 gene sequence, including lysine at site 296 that forms a Schiff base linkage with 11-cis-retinal

(Zhukovsky et al., 1991), and glutamate at site 113 that acts as a counterion to the protonated Schiff base linkage (Sakmar et al., 1989). Unlike rh1, rh1-2 is not expressed during the first four days of embryonic development, but is expressed by 21 days post fertilization. In the adult, rh1-2 is expressed in the retina, but not in the brain or muscle.

Although rare, zebrafish is not the only diploid teleost to express a second rh1 gene. The rod photoreceptors of eels start expressing an alternate rh1 gene during the transition from a freshwater habitat as a juvenile to a deep-sea habitat as an adult (Beatty, 1975;

Archer et al., 1995; Zhang et al., 2000). This switch in rh1 gene expression also results

64 in a blueshift of λmax of approximately 40 nm (Beatty, 1975; Hope et al., 1998). The short fin pearleye expresses an rh1a gene throughout its lifespan, but only expresses the

rh1b gene as an adult; the λmax of rh1b is also blueshifted by 6 nm compared to rh1a

(Pointer et al., 2007). Zebrafish rh1 is expressed as early as 2 days post fertilization and is expressed at high levels throughout adulthood (Raymond et al., 1995). Meanwhile, rh1-2 expression initiates sometime between 4 and 21 days post fertilization and also

remains at more modest levels throughout adulthood. The λmax values of zebrafish rh1

[501 nm] (Chinen et al., 2003) and rh1-2 [∼ 500 nm] are similar. Ultimately, the rh1 gene pair in zebrafish is similar to that of the pearleye, which has one gene expressed earlier in development at high levels, and a second gene expressed later in development at lower levels. However, unlike the pearleye, the two rh1 genes of zebrafish do not

seem to differ significantly in their λmax values.

Like most protein families, there are key residues and motifs present in opsins that are required for proper structure and function (Iannaccone et al., 2006). Alternatively, some sites in opsins act as determinants of rhodopsin and cone opsin function (Imai et al.,

1997; Carelton et al., 2005; Kuwayama et al., 2005), or differentiate between visual and non-visual opsins (Bellingham et al., 2002; Murakami and Kouyama, 2008). The rh1-2 gene family contains a number of substitutions at sites known to vary between rod, cone and non-visual opsins, as well as some that are entirely unique at otherwise highly conserved sites. Because opsins have been examined so extensively, many of these substitutions have been implicated in experimental opsin structure and function studies.

65 There are three motifs in rh1-2 that have substitutions suggesting altered function compared to rhodopsin: helix 1/helix 8, cytoplasmic loop 2, and extracellular loop 2. All three of these motifs are involved in meta II stability, transducin activation, or both, suggesting that rh1-2 does not behave as a traditional rhodopsin. Residues L45 and C51 are found in the first transmembrane helix of rh1-2. Helix 1 plays a role in assuring proper transmembrane packing of helices 2 and 7, with site 51 being of particular importance (Bosch et al., 2003). It is possible that interfering with helix 7 specifically could lead to disruption of helix 8, a key motif in both meta II stability and transducin activation (Ernst et al., 2000; Marin et al., 2000). Additionally, N65, a substitution only seen in melanopsin at the cytoplasmic surface of helix 1, is in close proximity to S316, an rh1-2-specific substitution in helix 8, and a residue involved in the meta I/meta II equilibrium (Weitz and Nathans, 1992).

There are also a number of substitutions in the second cytoplasmic loop [K145, S/T149,

Q150], as well as L151, the first residue of helix 4, which are either unique to rh1-2 or cone-like in nature. Cytoplasmic loop 2 is a key motif in transducin interaction and activation (König et al., 1989; Franke et al., 1992; Ernst et al., 1995; Natochin et al.,

2003), with some residues of the loop also being implicated in phosphorylation and glycosylation patterning (Shi et al., 1995; Zhu et al., 2006). Finally, L183 and E199 are part of extracellular loop 2, which folds into two short β-strands that form a shield over the chromophore-binding site (Palczewski et al., 2000; Okada et al., 2004); the movement of this motif is key to rhodopsin activation (Ahuja et al., 2009). Since other residues in extracellular loop 2 have been previously identified as key determinants of

66 differences between rhodopsins and cone opsins, including meta II decay rate

(Kuwayama et al., 2002), these substitutions could also be involved in altering rh1-2 function.

The λmax of an opsin gene is a property often used as an indicator of biological function,

with each group of visual opsins having a characteristic λmax range: rh1 ~ 500 nm, rh2 =

470-510 nm, sws2 = 440-460 nm, sws1 = 360-430 nm, and lws = 510-560 nm (Trezise

and Collin, 2005). The λmax values of non-visual opsins have not been analyzed to the same extent as those from visual opsins, often because they are more difficult to express.

The only non-visual opsins in fish that have been successfully expressed in vitro are the two isoforms of zebrafish VAL-opsin, both of which absorb maximally around 500 nm

(Kojima et al., 2008). Melanopsin, a non-visual opsin, has been investigated in mammals

and reported to have a λmax around either 480 nm (Panda et al., 2005; Dacey et al., 2005;

Torii et al., 2007; Terakita et al., 2008), or 420 nm (Newman et al., 2003; Melyan et al.,

2005). The λmax of zebrafish rh1-2, at approximately 500 nm, falls within the range of both rh1 and rh2 visual opsin groups, but is also similar to some non-visual opsins, such

as VAL-opsin. While the λmax value of rh1-2 might not help in distinguishing its functional role, the fact that it is much more difficult to express relative to zebrafish rh1 is indicative of a protein less stable than rhodopsin.

The spatial and temporal expression pattern of an opsin is an integral functional feature.

The onset of expression of the visual opsin repertoire in zebrafish varies from 2 days post fertilization [dpf] for rh1 and lws-2, to 7 dpf for rh2-2 (Takechi and Kawamura, 2005).

67 Meanwhile, the expression of both isoforms of VAL-opsin can be seen as early as 1 dpf

(Kojima et al., 2008). In adult vertebrates, visual opsins are expressed in the outer segments of photoreceptor cells found in the retina (Lamb et al., 2007) and, in rare circumstances, in the brain (Koyanagi et al., 2004). Non-visual opsins can be expressed in a wide variety of tissues, often simultaneously, including the retina (Hannibal et al.,

2002; Grone et al., 2007), brain (Mano et al., 1999; Philp et al., 2000), and even the testis

(Tarttelin et al., 2003) and ovaries (Kojima et al., 2008). Zebrafish rh1-2 expression initiates between 4 and 21 dpf and is localized in the retina. These results suggest that rh1-2 is more likely to serve as a visual opsin, as few non-visual opsins are expressed strictly in the retina. Furthermore, since the development of photoreceptors in vertebrates is dependent on rh1 expression (Lem et al., 1999), rh1-2 likely lacks a similar role in photoreceptor development.

The evolutionary origins of the rh1-2 gene family were investigated via phylogenetic analyses of rh1 and rh1-2 sequences. Both maximum likelihood and Bayesian methods recovered a phylogeny in which all rh1-2 sequences formed a well-supported, monophyletic group within the rh1 gene family. So far, we have only isolated rh1-2 sequences from cyprinids, a family of fish including zebrafish, goldfish, and barbs that originated between 32 and 39 million years ago (Zardoya and Doadrio, 1999; Wang et al., 2007). However, our phylogenetic analyses suggest a much older origin for this novel gene. As with rh1 genes of all actinopterygians, rh1-2 does not contain introns

(Fitzgibbon et al., 1995), suggesting that the duplication event leading to the birth of the rh1-2 family likely occurred after the retrotransposition event that split rh1 and exo-

68 rhodopsin in this group of fish (Bellingham et al., 2003). Although the exact time of actinopterygian divergence is a contentious issue, this would mean that the origin of the rh1-2 gene family is certainly no older than 350 million years ago (Hurley et al., 2007).

According to our analyses, there is some evidence that the origins of the rh1-2 group might be have occurred around the divergence of the eel rh1 sequences. The divergence of the elopomorphs [eels] from more derived teleosts occurred approximately 140 million years ago (Forey et al., 1996).

Such origins of the rh1-2 gene family suggest that its distribution among teleost fish may be much wider than our current experiments would indicate, particularly as we have not yet attempted to find rh1-2 genes outside of cyprinid fishes. This raises the possibility that rh1 gene duplicates in the pearl eye and eels might in fact be part of the same gene duplication that gave rise to the rh1-2 gene family. This would be rather surprising, as these rh1 duplicates are not generally thought to be widespread among fish. However, this possibility cannot be ruled out, as our analyses show low support at key divergences deep within the teleost rh1 phylogeny. Better sampling of basal teleost rh1 sequences, as well as a wider sampling of rh1-2 sequences would be needed to resolve this issue.

While rh1-2 is most closely related to the rh1 visual opsin group, this alone cannot classify it as a visual opsin, as the same can be said about the non-visual opsin, exo- rhodopsin (Mano et al., 1999). Despite expressing zebrafish rh1-2 in cell culture and characterizing its expression pattern via RT-PCR, it is still difficult to ascertain its function, whether within the visual system or serving in another system that relies on

69 photo entertainment. Regardless, the discovery of a new, previously uncharacterized opsin gene in the zebrafish retina is an important and surprising discovery, given its status as a model system for studying vertebrate vision and visual development.

Additional insights into the function of this novel gene could help to further elucidate how the vertebrate visual system functions. Localizing the specific cell types within the retina that express rh1-2 would help to determine its functional role. This could be done through extensive in situ hybridizations of adult zebrafish retina or by generating a transgenic zebrafish with expression of a GFP reporter driven by elements found upstream of the zebrafish rh1-2 ORF. Additional biochemical characterization of rh1-2 expressed in vitro would also help to determine the functional repercussions of its many unique amino acid substitutions. Obtaining additional rh1-2 nucleotide sequences from other teleosts would help to resolve the evolutionary origins of this novel gene family.

70 A Helix 1 Helix 2 Helix 3 45 51 65 rh1-2 FAFYCMAAYMLFLIVTCVPVNGLTLYVTIENKKLRTPLNYILLNLAVADLFMVFGGFTTTFYTSMHGYFVLGRAGCNLEGLFATVGGEIALWSLVVLAVE 134 rh1 WAYGLLAAYMFFLIITGFPVNFLTLYVTIEHKKLRTPLNYILLNLAIADLFMVFGGFTTTMYTSLHGYFVFGRLGCNLEGFFATLGGEMGLWSLVVLAIE exorh WQFSLLAAYMLFLILGSFPINALTLYVTVQHKKLRTPLNYILLNLAVADLFMVLGGFTVTLYTALHGYFLLGVTGCNIEGFFATLGGEIALWSLVVLAIE rh2-1 WKFKALAFYMFLLIIFGFPINVLTLVVTAQHKKLRQPLNYILVNLAFAGTIMVIFGFTVSFYCSLVGYMALGPLGCVMEGFFATLGGQVALWSLVVLAIE rh2-2 WQFKALAFYMFFLICFGLPINVLTLLVTAQHKKLRQPLNYILVNLAFAGTIMAFFGFTVTFYCSINGYMALGPTGCAIEGFFATLGGQVALWSLVVLAIE rh2-3 WQFKLLAVYMFFLMCFGFPINGLTLVVTAQHKKLRQPLNFILVNLAVAGTIMVCFGFTVTFYTAINGYFVLGPTGCAIEGFMATLGGQISLWSLVVLAIE rh2-4 WQFKLLAVYMFFLICLGFPINGLTLLVTAQHKKLRQPLNFILVNLAVAGTIMVCFGFTVTFYTAINGYFVLGPTGCAIEGFMATLGGEVALWSLVVLAVE sws2 GVFMGMSAFMLFLFIAGTAINVLTIVCTIQYKKLRSHLNYILVNLAISNLWVSVFGSSVAFYAFYKKYFVFGPIGCKIEGFTSTIGGMVSLWSLAVVALE sws1 WAFYLQAAFMGFVFIVGTPMNGIVLFVTMKYKKLRQPLNYILVNISLAGFIFDTFSVSQVFVCAARGYYFLGYTLCAMEAAMGSIAGLVTGWSLAVLAFE lws-1 WVYNVATVWMFFVVVASTFTNGLVLVATAKFKKLRHPLNWILVNLAIADLGETLFASTISVINQFFGYFILGHPMCIFEGYTVSVCGIAALWSLTVISWE lws-2 WVYNVATVWMFFVVVASTFTNGLVLVATAKFKKLRHPLNWILVNLAIADLGETLFASTISVINQVFGYFILGHPMCIFEGYTVSVCGIAGLWSLTVISWE valop-1 WNYSVLAALMFVVTALSLSENFTVMLVTFRFQQLRQPLNYIIVNLSLADFLVSLTGGSISFLTNYHGYFFLGKWACVLEGFAVTFFGIVALWSLAVLAFE valop-2 WNYTFLACLMFIVTSLSITENFTVMLVTYRFKQLRKPLNYIIVNLSVADFLVSMTGGTISFLTNARGFFFLGVWACVLEGFAVTFFGIVALWSLAILAFE tmtop TGHILVAVSLGFIGTFGFLNNLLVLVLFGRYKVLRSPINFLLVNICLSDLLVCVLGTPFSFAASTQGRWLIGDTGCVWYGFANSLLGIVSLISLAVLSYE melop-1 HAHYIIGSVILIVGITGVIGNALVVYVFCRSRTLRTAGNMFIVNLAVADFLMSVTQSPVFFAASLHRRWVFGERPCELYAFCGALFGICSMMTLTAIAAD melop-2 HAHYTIGAVILTVGITGMLGNFLVIYAFSRSRTLRTPANLFIINLAITDFLMCATQAPIFFTTSMHKRWIFGEKGCELYAFCGALFGICSMITLMVIAVD

Helix 4 Helix 5 145 149-151 183 199 rh1-2 RWVVVCKPFTKF-RFSQLHATLGVAFSWSMACSCAIPPLLGWSRYIPEGLQCSCGVDYYTPNPETENESFVIYMFVVHFSIPLTIISFCYGRLLCTVKVA 233 rh1 RWMVVCKPVSNF-RFGENHAIMGVAFTWVMACSCAVPPLVGWSRYIPEGMQCSCGVDYYTRTPGVNNESFVIYMFIVHFFIPLIVIFFCYGRLVCTVKEA exorh RYIVVCKPMSTF-RFGEKHAIIGVGFTWVMALTCAVPPLLGWSRYIPEGMQCSCGIDYYTPKPEVHNTSFVIYMFILHFSIPLLIIFFCYSRLLCTVRAA rh2-1 RYIVVCKPMGSF-KFSANHAMAGIAFTWFMACSCAVPPLFGWSRYLPEGMQTSCGPDYYTLNPEYNNESYVMYMFSCHFCIPVTTIFFTYGSLVCTVKAA rh2-2 RYIVVCKPMGSF-KFSSNHAMAGIAFTWVMASSCAVPPLFGWSRYIPEGMQTSCGPDYYTLNPEFNNESYVLYMFSCHFCVPVTTIFFTYGSLVCTVKAA rh2-3 RYIVVCKPMGSF-KFSSNHAFAGIGFTWIMALSCAAPPLVGWSRYIPEGMQCSCGPDYYTLNPDYNNESYVLYMFCCHFIFPVTTIFFTYGRLVCTVKAA rh2-4 RYIVVCKPMGSF-KFSASHAFAGCAFTWVMAMACAAPPLVGWSRYIPEGMQCSCGPDYYTLNPEYNNESYVLYMFICHFILPVTIIFFTYGRLVCTVKAA sws2 RWLVICKPLGNF-TFKTPHAIAGCILPWCMALAAGLPPLLGWSRYIPEGLQCSCGPDWYTTNNKFNNESYVMFLFCFCFAVPFSTIVFCYGQLLITLKLA sws1 RYVVICKPFGSF-KFGQGQAVGAVVFTWIIGTACATPPFFGWSRYIPEGLGTACGPDWYTKSEEYNSESYTYFLLITCFMMPMTIIIFSYSQLLGALRAV lws-1 RWVVVCKPFGNV-KFDAKWASAGIIFSWVWAAAWCAPPIFGWSRYWPHGLKTSCGPDVFSGSEDPGVQSYMVVLMITCCIIPLAIIILCYIAVYLAIHAV lws-2 RWVVVCKPFGNV-KFDGKWASAGIIFSWVWAAVWCAPPIFGWSRYWPHGLKTSCGPDVFGGNEDPGVQSYMLVLMITCCILPLAIIILCYIAVFLAIHAV valop-1 RFFVICRPLGNI-RLRGKHAALGLVFVWSFSFIWTVPPVLGWSSYTVSRIGTTCEPNWYSGN--FHDHTFIITLFSTCFIFPLGVIIVCYCKLIRKLRKV valop-2 RFFVICRPLKNV-RLGGKHAAMGLIFVWTFSFIWTIPPVLGWNSYTVSKIGTTCEPNWYSTN--YYDHTYIITFFVTCFILPLGVIIISYGKLMQKLRKV tmtop RYCTMMGSTEAD-ATNYKKVIGGVLMSWIYSLIWTLPPLFGWSRYGPEGPGTTCSVDWTTKT--ANNISYIICLFIFCLIVPFLVIIFCYGKLLHAIKQV melop-1 RCLAITQPLALVSRVSRRKAGAVFVVVWLYSLGWSLPPFFGWSAYVPEGLQTSCSWDYMTFTP--SVRAYTILLFVFVFFIPLGIIGSCYFAIFQTIRAA melop-2 RYFVITRPLASIGVLSQKRALLILLVAWVYSLGWSLPPFFGWSAYVPEGLLTSCTWDYMTFTP--SVRAYTMLLFIFVFFIPLIVIIYCYFFIFRSIRTT

Helix 6 Helix 7 Helix 8 316 rh1-2 AAQQQESETTQ------RAEREVTRMVILMVIAFLICWLPYASVAWYIFTHQGSQFGPVFMTVPAFFAKSSALYNPLIYVFMNKQFRHSMMMTV 321 rh1 AAQQQESETTQ------RAEREVTRMVIIMVIAFLICWVPYAGVAWYIFTHQGSEFGPVFMTLPAFFAKTSAVYNPCIYICMNKQFRHCMITTL exorh AAQQQESETTQ------RAEREVTRMVVVMVIAFLVCWVPYASVAWYIFANQGAEFGPVFMTVPAFFAKSAALYNPVIYIMLNRQFRNCMLSTV rh2-1 AAQQQESESTQ------KAEREVTRMVILMVLGFLFAWVPYASFAAWIFFNRGAAFSAQAMAVPAFFSKTSAVFNPIIYVLLNKQFRSCMLNTL rh2-2 AAQQQESESTQ------KAEREVTRMVILMVLGFLVAWVPYASFAAWIFFNRGAAFSAQAMAIPAFFSKASALFNPIIYVLLNKQFRSCMLNTL rh2-3 AAQQQESESTQ------KAEREVTRMVILMVLGFLVAWTPYASVAAWIFFNRGAAFSAQFMAVPAFFSKSSSIFNPIIYVLLNKQFRNCMLTTL rh2-4 AAQQQESESTQ------KAEREVTRMVILMVLGFLIAWTPYATVAAWIFFNKGAAFSAQFMAVPAFFSKTSALYNPVIYVLLNKQFRNCMLTTL sws2 AKAQADSASTQ------KAEREVTKMVVVMVFGFLICWGPYAIFAIWVVSNRGAPFDLRLATIPSCLCKASTVYNPVIYVLMNKQFRSCMMKMV sws1 AAQQAESESTQ------KAEREVSRMVVVMVGSFVLCYAPYAVTAMYFANSDEPNKDYRLVAIPAFFSKSSCVYNPLIYAFMNKQFNACIMETV lws-1 AQQQKDSESTQ------KAEKEVSRMVVVMIFAYCFCWGPYTFFACFAAANPGYAFHPLAAAMPAYFAKSATIYNPVIYVFMNRQFRVCIMQ-L lws-2 AQQQKDSESTQ------KAEKEVSRMVVVMILAFCLCWGPYTAFACFAAANPGYAFHPLAAAMPAYFAKSATIYNPIIYVFMNRQFRVCIMQ-L valop-1 SNTHGRLGNAR------KPERQVTRMVVVMIVAFMVAWTPYAAFSIIITAHPSMHVDPRLAAIPAFVAKTAAVYNPIIYVFMNKQFRKCLVQLL valop-2 SNTHGRLGNAR------KPDREVARMVVVMIVAFMVGWTPYAAFSITVTACPTIYIDPRLGSIPAFFSKTAAVYNPIIYVFMNKQFRKCLIQMF tmtop SSVN--TSVSR------KREHRVLLMVITMVVFYLLCWLPYGIMALLATFGAPGLVTAEASIVPSILAKSSTVINPVIYIFMNKQFYRCFRALL melop-1 GKEIRELDCG---ETHKVYERMQNEWKMAKVALVVILLFIISWSPYSVVALTATAGYSHFLTPYMNSVPAVIAKASAIHNPIIYAITHPKYRVAIARYI melop-2 NEAVGKINGDNKRDSMKRFQRLKNEWKMAKIALIVILMYVISWSPYSTVALTAFAGYSDFLTPYMNSVPAVIAKASAIHNPIIYAITHPKYRLAIAKYI

B Helix 1 Helix 2 45 51 65 D. rerio rh1-2 MNGTEGPDFYVPMSNESGVVRSPYEYPQYYLASPFAFYCMAAYMLFLIVTCVPVNGLTLYVTIENKKLRTPLNYILLNLAVADLFMVFGGFTTTFYTSMH 100 D. roseus rh1-2 MNGTEGPDFYVPMSNETGVVRSPYEYPQYYLASPLAFYCMAAYMLFLIVTCIPVNGLTLYVTIENKKLRTPLNYILLNLAVADLFMVFGGFTTTFYTSMH D. albolineatus rh1-2 MNGTEGPDFYVPMSNETGVVRSPYEYPQYYLASPLAFYCMAAYMLFLIVTCIPVNGLTLYVTIENKKLRTPLNYILLNLAVADLLMVFGGFTTTFYTSMH E. bicolor rh1-2 ------ASPVAFYCVAAYMLFLIVTCIPVNGLTLYVTINNKKLRTALNYILLNLAVADLFMVFGGFTTTFYTSLH D. rerio rh1 MNGTEGPAFYVPMSNATGVVRSPYEYPQYYLVAPWAYGLLAAYMFFLIITGFPVNFLTLYVTIEHKKLRTPLNYILLNLAIADLFMVFGGFTTTMYTSLH D. roseus rh1 ------VAPWAYGCLAAYMFFVILTGFPVNFLTLYVTIEHKKLRTPLNYILLNLAIADLFMVFGGFTTTMYTSLH D. albolineatus rh1 ------VAPWAYGCLAAYMFFVILTGFPVNFLTLYVTIEHKKLRTPLNYILLNLAIADLFMVFGGFTATMYTSLH E. bicolor rh1 ------VAPWAYACLAAYMFFLIITGFPINFLTLYVTIEHKKLRTPLNYILLNLAISDLFMVFGGFTTTMYTSLH B. taurus RH1 MNGTEGPNFYVPFSNKTGVVRSPFEAPQYYLAEPWQFSMLAAYMFLLIMLGFPINFLTLYVTVQHKKLRTPLNYILLNLAVADLFMVFGGFTTTLYTSLH

Helix 3 Helix 4 145 149-151 183 199 D. rerio rh1-2 GYFVLGRAGCNLEGLFATVGGEIALWSLVVLAVERWVVVCKPFTKFRFSQLHATLGVAFSWSMACSCAIPPLLGWSRYIPEGLQCSCGVDYYTPNPETEN 200 D. roseus rh1-2 GYFVLGRAGCNLEGLFATAGGEIALWSLVVLAVERWVVVCKPFTKFRFSQLHATMGVAFSWSMACSCALPPLLGWSRYIPEGLQCSCGVDYYTLNPETEN D. albolineatus rh1-2 GYFVLGRAGCNLEGLFATAGGEIALWSLVVLAVERWVVVCKPFTKFRFSQLHATMGVAFSWSMACSCALPPLLGWSRYIPEGLQCSCGVDYYTLNPETEN E. bicolor rh1-2 GYFVLGRPGCNLEGLFATLGGEIALWSLVVLAVERWVVVCKPFTKFRFTQLHATFGVAFSWAMACSCAIPPLLGWSRYIPEGLQCSCGVDYYTLNPETEN D. rerio rh1 GYFVFGRLGCNLEGFFATLGGEMGLWSLVVLAIERWMVVCKPVSNFRFGENHAIMGVAFTWVMACSCAVPPLVGWSRYIPEGMQCSCGVDYYTRTPGVNN D. roseus rh1 GYFVFGRLGCNLEGFFATLGGEMGLWSLVVLAIERWMVVCKPVSNFRFGENHAIMGVAFTWVMACSCAVPPLVGWSRYIPEGMQCSCGVDYYTRAPGVNN D. albolineatus rh1 GYFVFGRLGCNLEGFFATLGGEMGLWSLVVPAIERWMVVCKPVSNFRFGENHAIMGVAFTWVMACSCAVPPLVGWSRYIPEGMQCSCGVDYYTRAPGVNN E. bicolor rh1 GYFVFGRVGCNLEGFFATLGGEMGLWSLVVLAFERWMVVCKPVSNFRFGENHAIMGVAFTWVMACSCAVPPLVGWSRYIPEGMQCSCGVDYYTRVPGVNN B. taurus RH1 GYFVFGPTGCNLEGFFATLGGEIALWSLVVLAIERYVVVCKPMSNFRFGENHAIMGVAFTWVMALACAAPPLVGWSRYIPEGMQCSCGIDYYTPHEETNN

Helix 5 Helix 6 Helix 7

D. rerio rh1-2 ESFVIYMFVVHFSIPLTIISFCYGRLLCTVKVAAAQQQESETTQRAEREVTRMVILMVIAFLICWLPYASVAWYIFTHQGSQFGPVFMTVPAFFAKSSAL 300 D. roseus rh1-2 ESFVIYMFIVHFSIPLTVISFCYGRLLCTVKVAAAQQQESETTQRAEREVTRMVILMVIAFLICWLPYASVAWYIFTHQGSQFGPVFMTIPVFFAKSSAL D. albolineatus rh1-2 ESFVIYMFIVHFSIPLTVISFCYGRLLCTVKVAAAQQQESETTQRAEREVTRMVILMVIAFLICWLPYASVAWYIFTHQGSQFGPVFMTIPAFFAKSSAL E. bicolor rh1-2 ESFVIYMFVVHFSIPLTIISFCYGRLLCTVKVAAAQQQESETTQRAEREVTRMVVLMMIAFLICWLPYASVAWYIFTHQGSQFGPIFMTIPAFFAKSSAL D. rerio rh1 ESFVIYMFIVHFFIPLIVIFFCYGRLVCTVKEAAAQQQESETTQRAEREVTRMVIIMVIAFLICWVPYAGVAWYIFTHQGSEFGPVFMTLPAFFAKTSAV D. roseus rh1 ESFVIYMFIVHFFIPLVVIFFCYGRLVCTVKEAAAQQQESETTQRAEREVTRMVIIMVIAFLICWCPYAGVAWYIFTHQGTEFGPVFMTLPAFFAKTSAV D. albolineatus rh1 ESFVIYMFIVHFFIPLVVIFFCYGRLVCTVKEAAAQQQESETTQRAEREVTRMVIIMVIAFLICWCPYAGVAWYIFTHQGSEFGPVFMTLPAFFAKTAAV E. bicolor rh1 ESFVIYMFIVHFLIPFAVIFFCYGRLVCTVKEAAAQQQESETTQRAEREVTRMVVIMVIGFLICWLPYAGVAWYIFTHQGSEFGPVFMTLPAFFAKTAAV B. taurus RH1 ESFVIYMFVVHFIIPLIVIFFCYGQLVFTVKEAAAQQQESATTQKAEKEVTRMVIIMVIAFLICWLPYAGVAFYIFTHQGSDFGPIFMTIPAFFAKTSAV

Helix 8 316 D. rerio rh1-2 YNPLIYVFMNKQFRHSMMMTVCCGKDPFQDEEEGSSSSKSKTETSSVSSSSASSA 355 D. roseus rh1-2 YNPLIYVFLNKQFRHSMMMTVCCGKDPFQDEEEGSSSSKSKTETSSVSSSSASSA D. albolineatus rh1-2 YNPLIYVFMNKQFRHSMMMTVCCGKDPFQDEEEGSSGSKSKTETSSVSSSSASSA E. bicolor rh1-2 YNPL------D. rerio rh1 YNPCIYICMNKQFRHCMITTLCCGKNPF-EEEEGASTTASKTEASSVSSSSVSPA D. roseus rh1 YNPC------D. albolineatus rh1 YNPC------E. bicolor rh1 YNPC------B. taurus RH1 YNPVIYIMMNKQFRNCMVTTLCCGKNPLGDDE--ASTTVSKTETSQVAPA-----

71 Figure 1. Amino acid alignments of zebrafish rh1-2. (A) Zebrafish rh1-2 aligned with other visual and nonvisual opsin genes from zebrafish. N- and C-termini were excluded as they were too variable to align. (B) The rh1-2 gene family aligned with corresponding rh1 sequences and bovine rhodopsin. The seven transmembrane helices of rhodopsin, along with the eighth cytoplasmic helix, are labeled and shaded in light gray. Unique substitutions of the rh1-2 gene family are numbered and shaded in dark gray. Sequences obtained from GenBank are listed in Table S2.

72 ABC

Extracellular Loop 2 L45FL45F L183ML183M

Helix 1 E199NE199N

extracellular plasma membrane C51GC51G

plasma membrane cytoplasmic

Helix 8 S/T149GS/T149G plasma membrane Q150EQ150E cytoplasmic L151NL151N Cytoplasmic Loop 2 K145NK145N S316CS316C N65HN65H

Figure 2. Structures of three key motifs of rh1 containing cone-like and unique substitutions in the rh1-2 gene family. (A) The interaction between helix 1 and 8. (B) The second cytoplasmic loop. (C) The second extracellular loop. Unique substitutions of the rh1-2 gene family are highlighted in blue, with key motifs being colored green, and other residues colored gray. Approximate plasma membrane boundaries with either the extracellular or cytoplasmic environment are emphasized with a dashed line. The crystal structure of dark state bovine rhodopsin (1U19) and MacPyMOL (DeLano, 2008) were used for all images.

73 A

Figure 3. Absorbance spectra of zebrafish rh1 and rh1-2 after in vitro expression and purification. (A) Dark spectrum of zebrafish rh1 comparable to previous expressions (Chinen et al., 2003). (B) Dark spectrum of zebrafish rh1-2, whose expression level and absorbance ratio (A280/AMAX) are significantly lower than zebrafish rh1. The absorption maximum of zebrafish rh1-2 is approximately 500 nm, which is highlighted in the dark-light difference spectrum (inset).

74 1 dpf 2 dpf 3 dpf 4 dpf 21 dpf retina eye brain muscle

rh1

rh1-2

β-actin

Figure 4. RT-PCR analysis of zebrafish rh1-2 expression. Zebrafish rh1-2 is not expressed during the first 4 days of embryonic development but is expressed by the 21- day mark. In the adult, zebrafish rh1-2 is expressed in the retina but not in brain or muscle tissue. Zebrafish rh1 expression was similar to previous studies (Raymond et al., 1995; Takechi & Kawamura, 2005). b-actin serves as a positive control.

75 97/1.0 tetraodon 100/1.0 takifugu kingfish mullet 81/0.98 Acanthomorphs 76/1.0 58/0.98 medaka 96/1.0 guppy 48/0.92 dory 64/1.0 cod pearleye rh1a salmon 99/1.0 ayu 57/0.98 100/1.0 zebrafish purple danio 100/1.0 99/1.0 pearl danio 99/1.0 red tail shark 95/1.0 goldfish Ostariophysians 21/0.63 77/1.0 tetra sardine 62/0.97 milkfish 55/0.99 100/1.0 red tail shark rh1-2 zebrafish rh1-2 100/1.0 rh1-2 purple danio rh1-2 100/1.0 pearl danio rh1-2 53/0.97 pearleye rh1b 100/1.0 euro eel rh1fwo 16/0.96 conger eel rh1fwo Eels 100/1.0 52/- 100/1.0 euro eel rh1dso conger eel rh1dso butterflyfish 99/1.0 bowfin Basal Teleosts gar sturgeon 71/0.64 takifugu exorh Exo-rhodopsin 100/1.0 salmon exorh (outgroup) 50/0.80 ayu exorh zebrafish exorh

0.1

Figure 5. Phylogeny of teleost rh1 nucleotide sequences, including the rh1-2 gene family, generated using both maximum likelihood and Bayesian methods; rh1-2 sequences form a monophyletic group within the rh1 gene family. Sequences amplified in this study are indicated in bold. Numbers above nodes indicate ML bootstraps (100 replications) followed by posterior probabilities. Branches are proportional to ML estimated branch lengths under HKY+I+G model, the best model as determined by Modeltest AIC. Sequences obtained from GenBank are listed in Table S2.

Site Motif rh1-2 rh1 exorh rh2 sws2 sws1 lws valop tmtop melop 45 H1 L F L/I F F F/G I F G L 51 H1 C G A G G G S S G G 65 H1 N H H H Y Y F F Y N/S 145 CL2 K N N N/S N N N N A S/A 149 CL2 S/T G G S R/K S/N D R N S 150 CL2 Q E E S/A G/T S A G Y K 151 CL2 L N K T/S S/P K K K K R 183 EL2 L M M M L L L I P L 199 EL2 E N N/H N/H N N/K G N/H N R 316 H8 S C C C C C C C C A

Table 1. Dominant identities of key sites of rh1-2 in other opsin families. The following opsin gene families were sampled: rh1-2, rh1 [rhodopsin], exorh [exo-rhodopsin], rh2 [rod-like opsin], sws2 [short wavelength-sensitive 2], sws1 [short wavelength-sensitive 1], lws [long wavelength-sensitive], exorh [exo-rhodopsin], valop [vertebrate ancient opsin], tmtop [teleost multiple tissue opsin], and melop [melanopsin].

Primer Sequence (5' to 3') Purpose Reference RH1-2F ATGAACGGCACAGAGGGACCAGAC This study RH1-2R TCAGGCAGAAGAAGCTGAGCTAGAG Amplify rh1-2 This study RH1-2F91 TTGGCATCTCCRBTTGCVTTCTACTGCA from teleosts This study RH1-2R919 CGTAAATGAGYGGGTTGTAGAGAGCT This study RH1F93 GTATGAATATCCNCAGTAYTAYCT This study RH1R945 CAGCACARRGTGGTGATCATGCA Amplify rh1 from This study ChenRH1F CNTATGAATAYCCTCAGTACTACC teleosts Chen et al. 2003 ChenRH1R CCRCAGCACARCGTGGTGATCATG Chen et al. 2003 RH1-300F GTGCCGTGCCGCCACTGGTGGGCT Amplify zebrafish This study RH1-300R CCATGATGATACCATGCGGGTG rh1 (300 bp) This study RH1-2-300F TCCCATGTCGAATGAGAGTGGCGT Amplify zebrafish This study RH1-2-300R TGCAACCAGCCCGTCCCAGGACAA rh1-2 (300 bp) This study β-actinF ACAGTTGTAAGCAATGCCTCCTG Amplify zebrafish Wang et al. 2003 β-actinR CACGGAAGGCCATACCAGTAAGC β-actin (380 bp) Wang et al. 2003

Table S1. List of primer sequences used in this chapter.

Sequence Name Gene Species Accession # ayu rh1 Plecoglossus altivelis AB074484 ayu exorh exorh Plecoglossus altivelis AB089247 B. taurus RH1 RH1 Bos taurus NM_001014890 bowfin rh1 Amia calva AF137208 butterflyfish rh1 Pantodon buchholzi AF137210 cod rh1 Gadus morhua AF385832 conger eel rh1dso rh1 Conger myriaster AB043818 conger eel rh1fwo rh1 Conger myriaster AB043817 dory rh1 Zeus faber Y144484 euro eel rh1dso rh1 Anguilla anguilla L78008 euro eel rh1fwo rh1 Anguilla anguilla L78007 gar rh1 Lepisosteus osseus AF137207 goldfish rh1 Carassius auratus L11863 guppy rh1 Poecilia reticulata Y11147 kingfish rh1 Gasterochisma melampus DQ882021 medaka rh1 Oryzias latipes AB180742 milkfish rh1 Chanos chanos FJ197072 mullet rh1 Mugil cephalus Y18668 pearleye rh1a rh1 Scopelarchus analis EF517404 pearleye rh1b rh1 Scopelarchus analis EF517405 salmon rh1 Salmo salar AF201470 salmon exorh exorh Salmo salar AF201469 sardine rh1 Sardina pilchardus Y18677 sturgeon rh1 Acipenser sp. AF137206 takifugu rh1 Takifugu rubripes AF201471 takifugu exorh exorh Takifugu rubripes AF201472 tetra rh1 Astyanax mexicanus U12328 tetraodon rh1 Tetraodon nigroviridis AJ293018 zebrafish rh1 Danio rerio AB087811 zebrafish exorh exorh Danio rerio BC098524 zebrafish lws-1 lws Danio rerio AB087803 zebrafish lws-2 lws Danio rerio AB087804 zebrafish melop-1 melop Danio rerio AY078161 zebrafish melop-2 melop Danio rerio BC162681

79 zebrafish rh2-1 rh2 Danio rerio AB087805 zebrafish rh2-2 rh2 Danio rerio AB087806 zebrafish rh2-3 rh2 Danio rerio AB087807 zebrafish rh2-4 rh2 Danio rerio AB087808 zebrafish sws1 sws1 Danio rerio AB087810 zebrafish sws2 sws2 Danio rerio AB087809 zebrafish tmtop tmtop Danio rerio NM_01118899 zebrafish valop-1 valop Danio rerio BC128815 zebrafish valop-2 valop Danio rerio AY996588

Table S2. List of opsin sequences from GenBank used for amino acid alignments or phylogenetic analyses.

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Chapter 3:

Cellular localization and functional characterization of zebrafish rh1-2: investigating

its role in vision

92 3.1 Abstract

Rhodopsin is the visual pigment expressed in the retinal rod photoreceptors of vertebrates and is responsible for initiating the critical first step of dim-light vision. In most vertebrates, rhodopsin is a single-copy gene with highly conserved structure and function.

We recently discovered a novel rhodopsin-like gene expressed in the zebrafish retina, rh1-2, which we identified as a functional, photosensitive pigment. Here, we localize expression of rh1-2 in the zebrafish retina to a subset of peripheral photoreceptor cells, which partially overlaps the expression pattern of rhodopsin. Additionally, we measured the half-life of the rate of retinal release in rh1-2 to be 7.6 min at 20oC, which is more similar to rhodopsin [6.6 min] than the non-visual pigment exo-rhodopsin [1.6 min]. We also performed phylogenetic and molecular evolutionary analyses that suggest the presence of rh1-2 could be widespread in ostariophysian fish. Taken together, our data indicates that rh1-2 likely has a function similar to that of rhodopsin, and should be classified as a visual pigment. Finally, co-transfection with rh1-2 caused rhodopsin to have decreased thermal stability, possibly due to heterodimerization between the two visual pigments. Therefore, any photoreceptors that co-express these two visual pigments in vivo may have altered contributions from rhodopsin. These findings suggest that a second visual rhodopsin gene is present in a large phylogenetic group of vertebrates, which contradicts the well-established idea that rhodopsin is almost strictly a single-copy gene. The reasons for retention of this duplicate gene, as well as functional consequences for the visual system, are discussed.

93 3.2 Introduction

Photoreception in vertebrates is mediated by opsins, members of the G protein-coupled receptor (GPCR) superfamily of proteins (Terakita, 2005). In the dark, opsins are covalently bound to a light-sensitive chromophore, 11-cis-retinal, which acts as an inverse agonist to suppress dark state activation (Menon et al., 2001). When exposed to light, the chromophore isomerizes to its all-trans conformation, activating opsin and initiating a signaling cascade within the cell (Baylor, 1996). Visual opsins are responsible for initiating the visual transduction cascade, while non-visual opsins are likely involved in processes such as circadian entrainment (Doyle et al., 2008) and the metabolism of retinal (Bellingham et al., 2003a), with some possibly contributing indirectly to image formation (Cheng et al., 2009). Rhodopsin is the visual opsin expressed in rod photoreceptors responsible for mediating dim-light vision in vertebrates

(Nathans, 1992), and was the first GPCR to have its crystal structure resolved at high resolution (Palczewski et al., 2000).

Teleost fish are the only group of vertebrates known to have a non-visual rhodopsin-like gene, exorh, expressing exo-rhodopsin in the light-sensitive pineal gland (Mano et al.,

1999). Curiously, exorh is orthologous to rh1 of non-teleost vertebrates, while rh1 in teleosts is the product of a retrotransposition event, and contains no introns (Fitzgibbon et al., 1995). This duplication occurred no sooner than 284 million years ago, marking the onset of the radiation of ray-finned fish (Hurley et al., 2007), as basal Actinopterygians such as the sturgeon and gar also have intronless rh1 genes (Bellingham et al., 2003b).

Aside from melanopsin, exo-rhodopsin is one of the most highly characterized non-visual

94 opsins, as it has been implicated in the circadian clock system (Vuilleumier et al., 2006), positively regulates melatonin production (Pierce et al., 2008), and shows a 10-fold reduction in the lifetime of the biologically active meta II intermediate relative to rhodopsin (Tarttelin et al., 2011).

There are only a few vertebrates that express multiple visual rhodopsins. Multiple species of eels have both a freshwater [rh1fwo] and marine [rh1dso] rh1 gene, with expression changing from the former to the latter following migration during maturation

(Beatty, 1975; Hope et al., 1998; Zhang et al., 2000; Zhang et al., 2002). The short-fin pearleye [Scopelarchus analis], a deep-sea teleost, also expresses an additional rh1 gene, rh1B, in the accessory retina of adult fish after descending to greater ocean depths

(Pointer et al., 2007). Other examples of multiple rh1 genes are the result species- specific duplication events (Lim et al., 1997). Meanwhile, rhodopsin-like non-visual pigments seem restricted to the exorh gene family found in teleost fish (Philp et al.,

2000).

Another rhodopsin-like gene, rh1-2, was recently identified in juvenile and adult zebrafish, along with three other cyprinid fish, but limited functional characterization prevented its classification as either visual or non-visual (Morrow et al., 2011). Despite the overall scarcity of rh1 gene duplications, zebrafish is a logical candidate for such a rare event, considering its array of nine visual opsins is a large complement even among teleost fish (Chinen et al., 2003). Additionally, many non-visual opsins have been discovered in zebrafish, including exo-rhodopsin (Mano et al., 1999), two melanopsins

95 (Bellingham et al., 2002; Bellingham et al., 2006), teleost multiple tissue [tmt] opsin

(Moutsaki et al., 2003), and two vertebrate ancient long [VAL] opsins (Kojima et al.,

2008). This novel gene is expressed in the retina of adult zebrafish, but not in the brain.

When regenerated with 11-cis-retinal, rh1-2 produced an absorption spectrum with a

λMAX value of approximately 500 nm. Orthologous sequences were also found in three other cyprinid fish, suggesting that rh1-2 is not the result of a zebrafish-specific duplication event, and initial phylogenetic analyses hinted at a divergence from rh1 during the earlier stages of teleost evolution.

However, in order to clarify the function role of rh1-2, additional information relating to its cellular localization and biochemical characteristics, relative to other visual and non- visual opsins, will have to be assessed. Visual opsins are expressed primarily in the photoreceptor cells of the retina, where rhodopsin and cone opsins can usually be found in rod and cone photoreceptors, respectively. Meanwhile, non-visual opsins are expressed in other retinal cells, including horizontal cells (Cheng et al., 2009) and retinal ganglion cells (Dacey et al., 2005; Panda et al., 2005), and in neural tissues outside of the retina, including the pineal (Mano et al., 1999), cerebellum (Blackshaw and Snyder,

1999), and spinal chord (Tartellin et al., 2003). The five classes of visual opsins communally absorb light across the entire visual spectrum, while non-visual opsins seem to primarily absorb either violet (Newman et al., 2003), blue (Nakamura et al., 1999;

Torii et al., 2007), or green (Kojima et al., 2000) wavelengths of light. While there has been little evaluation of the kinetic process of non-visual opsins, exo-rhodopsin in teleost fish has a faster meta II decay rate than rhodopsin (Tarttelin et al., 2011). Furthermore,

96 identifying additional rh1-2 sequences, especially from fish outside of the family cyprinidae, will allow for a more accurate prediction of the evolutionary history of this gene family.

Here, we further characterize rh1-2 by drawing comparisons to both rhodopsin and exo- rhodopsin in order to gain a better understanding of its role in photoreception. We localized rh1-2 expression in the retina to a subset of peripheral photoreceptors, a pattern that partially overlaps rh1 expression, but that is distinct from exorh expression in the pineal gland of the brain. Additionally, rh1-2 releases retinal at a rate comparable to rhodopsin, and around five times faster than exo-rhodopsin. Considering these similarities to rhodopsin, we conclude that rh1-2 likely plays a role in image formation, and should be classified as a visual pigment. Moreover, preliminary evidence suggests that co-transfection with rh1-2 may alter the dark state stability of rhodopsin, possibly due to heterodimerization. Finally, rh1-2 was identified in three additional species, including one outside of the family cyprinidae, Misgurnus anguillicaudatus.

Phylogenetic analyses show the rh1-2 gene family is sister to ostariophysian rh1 genes, and was subjected to purifying selection following duplication and divergence, further solidifying the idea that it codes for a functional visual opsin. This study adds new insights to the already well-characterized visual system of zebrafish, while raising new questions about the functional role and implications of a second visual rhodopsin-like gene expressed in a taxonomically-rich group of teleost fish.

97 3.3 Materials and methods

3.3.1 Opsin sequences

RNA was extracted from adult eyes of teleost fish using the TRIzol reagent (Invitrogen) and cDNA libraries were generated using the SMART cDNA Library Construction Kit

(BD Biosciences). Genomic DNA was extracted from various tissues of teleost fish using the DNeasy Blood & Tissue Kit (QIAGEN). Gene fragments of rh1 and rh1-2 were amplified from cDNA libraries and genomic DNA, respectively, using either previously designed rh1-2 (Morrow et al., 2011) or acanthomorph rh1 primers (Chen et al., 2003), with resulting bands being cloned into the pJET1.2 cloning vector

(Fermentas). Full length sequences of rh1, rh1-2, and exorh, were amplified from these cDNA libraries generated with RNA from adult zebrafish eyes [rh1, rh1-2] or brain

[exorh]. PCR was performed using PfuTurbo (Stratagene) with resulting bands being cloned into the p1D4-hrGFP II expression vector (Morrow and Chang, 2010). All vectors were sequenced using a 3730 DNA Analyzer (Applied Biosystems). In addition to the rh1 and rh1-2 sequences obtained from the current study, vertebrate rhodopsin sequences, including rh1, exo-rhodopsin (exorh), and rh1-2 were obtained from

GenBank. Care was taken to maintain even sampling across vertebrates within the limits of the available data. A total of 144 rhodopsin sequences were obtained spanning lampreys, Chondrichthyes, lobe-finned fish, amphibians, mammals, reptiles [including birds], and all available major orders of ray-fined fish. Four vertebrate rh2 sequences were used as outgroups. Sequences were aligned using the webPRANK (Löytynoja and

Goldman, 2010) implementation of PRANK (Löytynoja and Goldman, 2005). Species

98 list and accession numbers for all sequences used in the study are provided in

Supplementary Table S1.

3.3.2 In situ hybridization

Eyes were dissected from 21 and 175 dpf zebrafish anesthetized with 160 mg/L of tricaine [ethyl 3-aminobenzoate methanesulfonate salt] (Fluka), then fixed in 4% paraformaldehye in phosphate-buffered saline [PBS] at 4oC overnight. Eyes were rinsed in PBS with 0.1% Tween-20 [PBT], then in methanol, before being stored in fresh methanol at -20oC. In situ hybridizations were performed on 3 dpf [control] and 5 dpf zebrafish embryos [whole mount], and eyes from 21 and 175 dpf zebrafish, as previously described (Jowett and Lettice, 1994). DIG-labeled RNA probes 700 bp in length, as well as unlabeled, full length blocking RNA from rh1-2 [control], were amplified from rh1 and rh1-2 sequences inserted into the pBluescript cloning vector using T3 RNA

Polymerase (Fermentas). A probe concentration of 1 ng/uL was using during the 70oC hybridization. Semi-thin plastic sections were made on 5 dpf embryos, where whole- mounts were rinsed with PBT, dehydrated using increasing concentrations of ethanol

(from 30% to 90% in PBT), followed by 100% ethanol, and then embedded with increasing concentrations of Spurr’s resin in ethanol (3:1, 1:1, 1:3). Samples were then left to polymerize at 65°C in 100% Spurr’s resin. Semithin coronal sections were cut with a glass knife using an ultramicrotome and dried onto glass slides. Sections were 1.5 mm thick without counterstaining to maximize visualization. Cryosections were performed on 21 dpf and 175 dpf zebrafish eyes, which were washed 3 times in PBS, then put through a sucrose gradient at room temperature, 30 minutes per step: 5% sucrose in PBS,

99 2:1 5%:30%, 1:1 5%:30%, 1:2 5%:30%, with a final step in 30% sucrose in PBS at 4oC overnight. Eyes were incubated in 2:1 30% sucrose in PBS:OCT compound (Tissue-Tek) for 4 hours at room temperature, then 4oC overnight. Cryosections were performed at 20

μm on a Leica CM3050S cryostat, and collected on Superfrost Plus slides (VWR) mounted in 90% glycerol/10% PBS. All images were taken on a Leica DM4500B compound microscope with a QIMAGING digital camera and OpenLab 4.0.2 software

(Improvision).

3.3.3 Expression and spectroscopy

The p1D4-hrGFP II expression vector constructs containing full coding sequences of zebrafish rh1, rh1-2, and exorh were used to transiently transfect cultured HEK293T cells using Lipofectamine 2000 (Invitrogen; 8 ug of DNA per 10-cm plate); for the co- transfection experiment, 4 ug of expression vector containing each of rh1 and rh1-2 were used, for a total of 8 ug of DNA per 10-cm plate. Cells were harvested 48 h post- transfection and opsins were regenerated using 11-cis-retinal, generously provided by Dr.

Rosalie Crouch (Medical University of South Carolina). Visual pigments were solubilized in 1% N-dodecyl-β-D-maltoside [DM] and immunoaffinity purified with the

1D4 monoclonal antibody (Molday and MacKenzie, 1983), as previously described

(Morrow and Chang, 2010). Purified visual pigment samples were eluted in sodium phosphate buffer [50 mM NaPhos, 0.1% DM, pH 7]. The ultraviolet-visible absorption spectra of purified opsin were recorded at 25oC using the Cary4000 double-beam

spectrophotometer (Varian) and quartz absorption cuvettes (Helma). All λMAX values

100 were calculated after fitting absorbance spectra to a standard template for A1 visual pigments (Govardovskii et al., 2000).

The protocol used to determine retinal release rates of visual pigments was modified from that of Farrens and Khorana (1995). Briefly, 0.05-0.20 μM visual pigment samples were incubated at 20oC and bleached for 30 seconds using a Fiber-Lite MI-152 Illuminator external light source (Dolan-Jenner), using a filter used to restrict wavelengths of light below 475 nm. Fluorescence measurements were integrated for 2 seconds at 20-second

[rh1-2] or 30-second [rhodopsin, exo-rhodopsin] intervals using a CaryEclipse fluorescence spectrophotometer (Varian). Thermal stability was measured by incubating

0.1-0.2 μM visual pigment samples at 56oC for 300 minutes, with temperature being maintained by a Cary Temperature Controller employing a Peltier Multicell Holder

(Varian) and monitored by a temperature probe. Both control [rhodopsin] and experimental [rhodopsin & rh1-2] data collected simultaneously to account for inter- experimental variation, with data points being integrated for 2 seconds, and collected every 2 to 6 minutes, increasing over the course of the experiment. For both experiments, samples were incubated in sodium phosphate buffer [50 mM NaPhos, 0.1% DM, pH 7] using submicro fluorometer cell cuvettes (Varian). The excitation wavelength was 295 nm [1.5 nm slit width] and the emission wavelength was 330 nm [10 nm slit width]; no noticeable pigment bleaching by the excitation beam was detected. Retinal release was demonstrated through a sharp initial rise in intrinsic tryptophan fluorescence, representing a decrease in fluorescent quenching of W265 by the retinal chromophore.

Data from the initial rise was fit to a three variable, first order exponential equation [y =

101 -bx yo + a(1-e )], with half-life values calculated based on the rate constant 'b' [t1/2 = ln2/b].

All curve fitting resulted in r2 values of greater than 0.9 for both light-activated retinal release and thermal stability assays.

3.3.4 Phylogenetic and molecular evolutionary analyses

An rh1 gene tree was estimated in MrBayes 3 (Ronquist and Huelsenbeck, 2003) using reversible jump MCMC with a gamma rate parameter (nst=mixed, rates=gamma), which explores the parameter space for the nucleotide model and the phylogenetic tree simultaneously. The analysis was run for five million generations with a 25% burn-in.

Convergence was confirmed by checking that the standard deviations of split frequencies approached zero and that there was no obvious trend in the log likelihood plot.

To estimate the strength and form of selection acting on rhodopsin, the alignment, along with the Bayesian gene tree, was analyzed with the codeml package of PAML 4 (Yang,

2007) using the random sites models (M0, M1a, M2a, M3, M7, M8a, and M8), branch, branch-site model, and clade model C (CmC). Analyses were run on the complete rh1 alignment and tree as well as two subsets, one pruned to only include ray-finned fish rh1

(including exorh as the outgroup) and the other pruned to constrain only rh1-2 (no outgroup).

Comparisons between the PAML random sites models were used to test for variation in ω

[M3 vs M0] and for the presence of a positively selected class of sites [M2a vs M1a and

M8 vs M7 and M8a]. All analyses were run starting with the branch lengths estimated by

102 MrBayes repeated at least three times with varying initial starting points of κ [transition to transversion ratio] and ω to avoid potential local optima. The model pairs were compared using a likelihood ratio test [LRT] with a χ2 distribution.

The branch, branch-site (Zhang et al., 2005) and clade models [CmC] (Bielawski and

Yang, 2004) were used to test for changes in selective constraint and positive selection on the branch leading to the rh1-2 clade and between the rh1-2 clade and other rhodopsins.

The branch model estimates a single omega value for each branch and/or clade type specified a priori. This model is useful for testing for overall changes in selective constraint between branches/clades. The branch-site and clade models allow ω to vary both among sites and between branches/clades. The branch-site model has four site

classes: 0) 0 < ω0 < 1 for all branches; 1) ω1 = 1 for all branches, 2a) ω2a = ω2b ≥ 1 in the

foreground and 0 < ω2a = ω0 < 1 in the background, and 2b) ω2b = ω2a ≥ 1 in the foreground and ω2b = ω1 = 1 in the background. This model provides a test for positive selection on specified branches/clades. CmC assumes that some sites evolve

conservatively across the phylogeny [two classes of sites where 0 < ω0 < 1 and ω1 = 1],

while a class of sites is free to evolve differently among two or more partitions [e.g., ωD1

> 0 and ωD1 ≠ ωD2 > 0], which can be branches, clades, or a mix of both. Rather than a test for positive selection this provide a test for divergent selective pressure, although a test for positive selection can be performed if desired (see Chang et al., 2012). These models were applied only to the dataset pruned to contain only ray-finned fish rhodopsins.

103 3.4 Results

3.4.1 Zebrafish rh1-2 is expressed in the outer nuclear layer of peripheral rod photoreceptor cells

In chapter 2, we established that rh1-2 expression did not occur during the first 4 days of embryonic development and was expressed in the retina as early as 21 days post fertilization. Here, we performed a series of in situ hybridizations to further pinpoint the onset of rh1-2 expression and to localize its expression in the retina. In situ hybridizations were performed using 700 bp coding sequence probes for both rh1 and rh1-2 in order to localize cellular expression in both whole mount embryos [5 dpf] and both juvenile [21 dpf] and adult [175 dpf] eyes. At 5 dpf, expression of rh1 was strongest in the peripheral retina, although some limited expression was also seen in the central retina (Fig. 1A), while rh1-2 was only detected in a limited portion of the peripheral retina (Fig. 1B). At 21 dpf, both rh1 and rh1-2 expression in the peripheral retina were more prominent relative to expression at 5 dpf (Fig. 1C and D), while rh1 expression was also more widespread in the central retina, where the absence of rh1-2 remained (Fig. 1E and F). At 175 dpf, expression of rh1 was stronger and more uniform throughout both the central (Fig. 1G) and peripheral retinal (data not shown). Meanwhile, rh1-2 expression was still absent in the central retina (Fig. 1H), and was present in a similar subset of peripheral cells compared to 21 dpf (data not shown).

Expression of rh1-2 was consistent with RT-PCR results from chapter 2, which showed expression in 21 dpf juvenile fish, and the adult retina, but at significantly lower levels than rh1 (Morrow et al., 2011). Furthermore, all expression of rh1 and rh1-2 was

104 confined to the outer nuclear layer [ONL], consisting of the cell bodies of rod and cone photoreceptors, suggesting that rh1-2 protein expression likely occurs in photoreceptor outer segments, as opposed to other retinal cells. Another interesting feature of rh1-2 expression is that it often overlaps rh1 expression in the peripheral ONL, which suggest expression of both genes is either proximal, or even concurrent in the same photoreceptor. Because of this overlap, and since the nucleotide sequences of rh1 and rh1-2 share approximately 75% similarity, a sense-strand probe control experiment was run to ensure that probes were not cross-hybridizing. The same 700 bp rh1 probes were used to stain 3 dpf embryos both with and without the addition of full-length rh1-2 blocking RNA, present at double the concentration of the rh1 probe (Fig. S1). The presence of the rh1-2 blocking RNA did not have a significant effect on rh1 staining, which suggests that there is likely no cross-hybridization occurring between rh1/rh1-2 probes and their respective target transcripts. Patterns of rh1 expression were similar to those presented in previous studies (Raymond et al., 1995; Robinson et al., 1995;

Takechi and Kawamura, 2005).

3.4.2 Zebrafish rh1-2 releases retinal at a rate similar to rhodopsin, but slower than exo-rhodopsin

In order to better classify rh1-2 as either a visual or non-visual opsin, we set out to identify a biochemical property that could differentiate between the two opsin groups.

The λMAX of most rhodopsins [~500 nm] is similar that of rh1-2 (Morrow et al., 2011), in addition to many non-visual opsins (Kojima et al., 2000; Tarttelin et al., 2011), making

λMAX a poor measure to differentiate between visual and non-visual opsins. However

105 kinetic properties of opsins following activation have been shown to differ between these groups (Nakamura et al., 1999; Tarttelin et al., 2011). Release of all-trans-retinal occurs after photoactivation, requiring both hydrolysis of the Schiff base linkage between opsin and retinal, as well as dissociation of retinal from opsin. In chapter 4, we report measurements of the retinal release half-life of zebrafish rhodopsin to be 6.6 min using fluorescence spectroscopy. Exo-rhodopsin released retinal just over four times faster than rhodopsin, with a half-life of 1.6 min, while rh1-2 had a half-life very similar to rhodopsin, 7.6 min (Fig. 2). This is the first time that retinal release has been measured in a non-visual opsin, and suggests that the kinetics of photoactivation in rh1-2 is more similar to rhodopsins than exo-rhodopsin, despite the fact that in vitro expression shows rh1-2 is much less stable than both rhodopsin and exo-rhodopsin.

3.4.3 Co-transfection with rh1-2 alters thermal stability of rhodopsin

Our in situ hybridization results show the expression pattern of rh1-2 overlaps that of rhodopsin in a subset of peripheral photoreceptor cells, suggesting the potential for

interaction in vivo. Since both λMAX and retinal release rates are similar between rhodopsin and rh1-2, but the stability of rh1-2 during expression was much lower, we carried out a co-transfection experiment to determine if rh1-2 had any influence on the thermal stability of rhodopsin. Fluorescence spectra were recorded for both rhodopsin alone, as well as rhodopsin co-transfected with rh1-2, upon incubation at 56oC, with an increased fluorescence signal accumulating upon release of all-trans-retinal from the chromophore binding pocket (Fig. 3). Data for rh1-2 alone was not collected, as it could not be expressed in high enough levels for this thermal stability assay. When rhodopsin

106 was expressed in the presence of rh1-2, the half-life of thermal activation measured was only 61.1% of the half-life of rhodopsin alone, suggesting that the presence of rh1-2 may reduce the thermal stability of rhodopsin. All fluorescence measurements were normalized to the maximum values recorded after the increased signal had plateaued in accordance with a first order exponential reaction.

3.4.4 The rh1-2 gene family is sister to Ostarioclupimorpha rh1 genes

In chapter 2, we identified rh1-2 as a monophyletic clade within the teleost rh1 clade that separated following the divergence of eel rh1 genes. However, bootstrap support for this topology was very low due to a lack of sequences, most notably those of basal teleost rh1 genes, and other rh1-2 genes. Here, we greatly increased our statitstical power by adding additional sequences to better understand the evolutionary history of rh1-2. In order to determine the phylogenetic placement of rh1-2 within the rh1 tree, we performed a

Bayesian phylogenetic analysis of vertebrate rh1 genes. The resulting topology largely recovered expected high-level relationships, including the placement of lampreys, chondrichthyes, lobe-finned fish, amphibians, amniotes, and ray-finned fish (Fig. 4, Fig.

S2). rh1-2 was recovered with some support [60% posterior probability] as the sister group to Clupemorpha plus Ostariophysi [Ostarioclupimorpha] rh1 (Fig. S2). rh1-2 was not found to be most-closely related to other duplicated ray-finned fish rh1, such as exorh, eel deep-sea rhodopsin [dso] and freshwater rhodopsin [fwo], or pearleye rh1a and rh1b. Together, this suggests that the duplication that led to rh1-2 occurred in the ancestor of Ostarioclupimorpha. This finding is corroborated by the absence of rh1-2 from euteleosts and basal ray-finned fish both in sequenced genomes [e.g., fugu, tilapia,

107 cod, stickleback] and our cDNA screening. However, it is also possible that duplicated copies of rh1 genes were only retained in one group and were ancestrally lost in others, and this could produce complicated phylogenetic patterns that would not be easily resolved. Previous phylogenetic analysis of rh1-2 found, with weak support, that rh1-2 was sister to Ostariophysians plus Acanthomorphs (Morrow et al., 2011), although the apparent absence of rh1-2 from Acanthomorphs may not support this hypothesis. The discrepancy is likely due to the vastly increased taxon sampling in the current study.

These results further suggest that additional copies of rh1-2 have yet to be identified from several groups of ray-finned fish.

Molecular evolutionary analyses were used to determine what changes in selective constraint occurred during and after the duplication that lead to rh1-2. The random site models of PAML revealed that vertebrate rhodopsins as a whole were under strong selective constraint [average ω = 0.073, M0] (Table S2) with no evidence of positive selection [M2a vs M1a; M8 vs M8a, p >> 0.5 in all cases] (Table S2). Significant rate variation among sites was found, as is likely to be the case for all functional protein coding genes [M3 vs M0, p < 0.00] (Table S2). This was also true when only ray-finned fish were considered [ω = 0.074, M0; M2a vs M1a; M8 vs M8a, p >> 0.5 in all cases; M3 vs M0, p < 0.00] (Table S2) and when only rh1-2 was considered [ω = 0.075, M0; M2a vs M1a; M8 vs M8a, p >> 0.5 in all cases; M3 vs M0, p < 0.00] (Table S2).

To further test for differences in selective constraint between rh1-2 and other rh1 genes we used the branch, branch-site, and clade [CmC, clade model C] models on the dataset

108 pruned to contain only ray-finned fish rhodopsins. The branch model was used to test for overall changes in selective constraint both on the branch leading to rh1-2, and on the entire clade. If rh1-2 were non-functional, we would expect a relaxation of selective constraint where ω would eventually reach one. We also might expect a relaxation of constraint following duplication [i.e., on the branch] before a return to constraint [i.e., in the clade]. Instead we found that the ω along the branch and for the whole clade did not significantly differ from background ω [p >> 0.5] (Table S3). This suggests that rh1-2 is a functional gene since it has been maintained under high levels of negative selection, corroborated by the M3 results showing significant rate variation. To test for positive selection and divergence in selective pressures we used the branch-site model and CmC.

We found significant evidence for positive selection on the branch leading to the rh1-2 clade using the branch-site model [p = 0.28] (Table S3). This suggests that rh1-2 may have experienced a burst of positive selection following duplication. We found no significant evidence for positive selection on the whole rh1-2 clade using the branch-site models or for divergent selective pressures on the rh1-2 branch or in the entire clade using CmC [p > 0.05 in all cases] (Table S4). Overall these results indicate that rh1-2 has been maintained under similar selective pressures as other vertebrate rhodopsins.

3.5 Discussion

Using in situ hybridization and fluorescence spectroscopy, we have shown that rh1-2 is a functional opsin gene that is expressed in peripheral photoreceptors of the zebrafish retina, releases retinal at a similar rate to rhodopsin, and may alter the thermal stability of rhodopsin when co-transfected. However, rh1-2 expression only starts around 5 dpf, and

109 is expressed weakly in vitro, both traits that are uncharacteristic of traditional rh1 genes.

Meanwhile, phylogenetic analyses place the rh1-2 clade sister to ostariophysian rh1 genes, suggesting a fairly widespread presence within this large group of teleost fish.

Here, we will discuss potential functional roles for rh1-2 considering our findings, and the implications of this opsin gene for the zebrafish visual system.

Considering how rare it is to for rhodopsin genes to duplicate relative to cone opsins genes (Bowmaker, 2008), an initial hypothesis concerning the role of rh1-2 is that it is a gene duplicate with no unique functional role in photoreception, experiencing leaky expression. Where traditional rhodopsin genes maintain a high level of expression in the retina, the duplication event that gave rise to rh1-2 failed to transfer the same regulatory elements that drive rh1 expression (Kennedy et al., 2001), resulting in much lower expression levels. Furthermore, only a subset of retinal photoreceptor cells express rh1-

2. While there is precedence for opsin gene duplicates to have differing spatial expression patterns in the retina (Takechi and Kawamura, 2005), it is hard to determine

why only these specific cells experience rh1-2 expression. Additionally, the λMAX values of rhodopsin and rh1-2 are very similar; therefore the spectral sensitivity provided by these two opsins is likely redundant. Finally, in vitro expression of zebrafish rh1-2 suggests it is considerably less stable than zebrafish rhodopsin, although this could be partially due to a bias in the expression protocol historically refined for expression of bovine rhodopsin. However, there is also evidence to suggest that rh1-2 is a functionally relevant opsin gene.

110 While rh1-2 does have some characteristics of a redundant gene duplicate, this classification is usually attributed to genes resulting from relatively recent gene duplication events, with the vast majority of gene duplicates being silenced within a few million years (Lynch et al., 2001). However, unlike a more recent gene duplication, which may generate species-specific duplicates (Lim et al., 1997), our analyses suggest a much more ancient origin for the rh1-2 gene family, sister to ostariophysian rh1 genes, which would place the duplication and divergence event leading to the birth of rh1-2 somewhere between 153 and 248 million years ago (Nakatani et al., 2011; Chen et al.,

2013). Moreover, we have direct evidence that seven ostariophysians [order: cypriniformes] have maintained a functional copy of rh1-2. All of these sequences contain the highly conserved residues required for proper opsin function, such as K296 to form a Schiff base linkage with retinal (Zhukovsky et al., 1991), and E113 to serve as a counterion to the protonated Schiff base (Sakmar et al., 1989). This function was confirmed when zebrafish rh1-2 responded to light by converting to its activated state,

meta II, characterized by a shift in absorbance from its λMAX to approximately 380 nm

(Morrow et al., 2011). Finally, PAML analyses indicate a low Ω value along the branch leading to the rh1-2 clade, implying characteristic purifying selection after divergence seen in genes that survive duplication and divergence events (Lynch and Conery, 2000).

This period of purifying selection may have allowed time for novel rh1-2 function to evolve or ancestral function to be stabilized, as opposed to the acquisition of negative function, or pseudogenization (Gayral et al., 2007).

111 The reason for proposed rh1-2 maintenance is likely different than those most commonly presented as the sources of opsin gene family expansion: differences in spectral sensitivity and subfunctionalization of expression pattern. One of the earliest opsin gene duplications to be characterized was the m/lws expansion in Old World primates responsible for trichromacy (Neitz et al., 1991; Hunt et al., 1998; Nathans, 1999), with the distribution of cone photoreceptors expressing either MWS or LWS being distributed in the retina in a single, stochastic mosaic (Roorda and Williams, 1999), and thought to be a visual adaptation to either frugivory and/or sexual selection (Osorio and Vorobyev,

1996; Sumner and Mollon, 2000; Smith et al., 2003). However, this duplication, along with polymorphic diversity of the lws locus in New World monkeys (Jacobs and Deegan,

2003), ended up being an evolutionary outlier in mammals, as the large majority of opsin gene family duplication and expansion has occurred in teleost fish (Bowmaker, 2008).

There are examples throughout the teleost lineage of duplications in every cone opsin category, including rh2 (Chinen et al., 2003; Nakamura et al., 2013), lws (Weadick and

Chang, 2007; Owens et al., 2009), sws2 (Matsumoto et al., 2006; Weadick et al., 2012), and, in one case, sws1 (Minamoto and Shimizu, 2005). A feature in some of these gene

duplications is diversification of λMAX within an opsin group, providing optimal sensitivity at a range of wavelengths. A previous study outlined differential expression patterns of duplicated rh2 and lws genes in zebrafish, allowing different parts of the retina to be maximally sensitive to different wavelengths of light (Takechi and

Kawamura, 2005), which may be an adaptation to light refracting at different angles and targeting different areas of the retina below the surface of an aquatic habitat. Other studies have shown that ontogenetic factors influence expression levels of opsin gene

112 duplicates, providing maximum sensitivity to specific wavelengths of light during different developmental stages (Spady et al., 2006; Carleton et al., 2008; Shand et al.,

2008; Temple et al., 2008). In fact, some species of cichlid fish display differential opsin expression based on sex, or even from individual to individual (Sabbah et al., 2010). In

zebrafish, there is less than 3 nm of difference between the λMAX of rhodopsin and rh1-2, much less than the 10 and 38 nm discrepancies in zebrafish lws and rh2 cone opsins respectively (Chinen et al., 2003). Additionally, rh1-2 expression does not replace that of rh1, but instead overlaps a portion of photoreceptor cells in the peripheral retina as of

5 dpf. This suggests that, while a functional rh1-2 does appear to have been maintained in zebrafish, it was not for the same reasons as most cone opsin gene duplicates.

Aside from rh1-2, only two other rh1 gene duplications have also been retained in

Actinopterigian fish, making rh1, along with sws1, the least common opsin gene to experience duplication. The first example is from eels, which express the rh1fwo gene with a 3,4-didehydroretinal [A2] chromophore in the early stages of life, where a red- shifted rhodopsin provides an advantage in the more long wavelength-shifted spectral environment of freshwater (Bridges, 1972; Loew, 1995). During maturation, eels migrate to a marine environment, with a more restricted and blue-shifted light spectrum, coupled with expression of a blue-shifted rh1dso gene, and 11-cis-retinal [A1] chormophore

(Hope et al., 1998; Zhang et al., 2000). This switch of both opsin and chromophore is a clear example of an adjustment of the visual system due to a change in photic environment. Another example is the deep-sea pearleye, S. analis, which has a more traditional rh1A gene, along with rh1B, expressed alongside rh1A in adult fish living over

113 900 m below the surface (Pointer et al., 2007). The pearleye has unique cylindrical eye morphology, containing both a main retina, used for image formation, and an accessory retina, likely only capable of gross light perception (Collin et al., 1998), with rh1B expression likely being localized in this accessory retina (Pointer et al., 2007). Zebrafish does not experience an ontogenetic migration between fresh and marine environments, possesses only A1 chromophore-based visual pigments (Allison et al., 2004), does not occupy deep-sea habitats, and does not possess a cylindrical eye, or an accessory retina, suggesting that it is unlikely for rh1-2 to serve a similar function as duplicated rhodopsin genes in either eels or the pearleye. This idea is supported by our phylogenetic analyses, which present rh1fwo, rh1dso, rh1B, and rh1-2 all as distinct clades resulting from separate duplication and divergence events. However, focusing on aspects of rh1-2 that are distinct from other rh1 and cone opsin gene duplications might provide the required insight to better understand the function of this unique gene family.

Two key results highlighted in this study are: 1) rh1 and rh1-2 expression overlaps in a subset of peripheral photoreceptors, and 2) co-transfection of rh1-2 may alter the thermal stability of rhodopsin. Taken together, these results suggest a possible functional consequence of rhodopsin and rh1-2 interaction in rod photoreceptors in vivo. When rhodopsin is packed into the outer segments of rod photoreceptors, it forms an array of dimers (Fotiadis et al., 2003). This arrangement could serve a structural benefit to maximize the capacity of the rod outer segments, but likely also serves a functional purpose, with higher order rhodopsin oligomers being a more active species than monomers (Fotiadis et al., 2006). This is not a unique characteristic of rhodopsin, as

114 dimerization is common in GPCRs (Babcock et al., 2003), with heterodimers sometimes allowing for differential ligand binding relative to respective homodimers (Waldhoer et al., 2005). For example, in the gamma-aminobutyric acid [GABA] receptor, heterodimers are formed between the B1 and B2 subunits, which bind ligand and G protein, respectively (Monnier et al., 2011). This functional dichotomy, where one unit responds to stimulus and the other bind the G protein, was also predicted for rhodopsin by molecular dynamics simulations, despite the presence of identical subunits (Neri et al.,

2010). This may be possible because each unit of the rhodopsin dimer exists in a different conformation following activation, promoting distinct functions from otherwise identical subunits (Jastrzebska et al., 2013). Considering the possible effect of rh1-2 on the thermal stability of rhodopsin in vitro, we hypothesize that these opsins may be co- expressed in a subset of peripheral rod photoreceptors and form heterodimers.

Despite a lack of direct evidence for heterodimerization among opsins, it is not uncommon for multiple opsins to be expressed in the same photoreceptor, which could allow for dimerization between compatible molecules. This has been observed in a range of organisms from invertebrates, such as Drosophila, expressing multiple rhodopsin subtypes in a single photoreceptor (Mazzoni et al., 2008), to humans that express short and long wavelength-sensitive opsins in some cone cells, most notably early in development

(Xiao et al., 2000). Therefore, it is reasonable to assume that both rhodopsin and rh1-2 could be expressed in a subset of photoreceptors, considering their overlapping expression patterns. Opsin dimerization can also be achieved in vitro, assuming the appropriate type and concentration of detergent is used (Jastrzebska et al., 2004); our

115 experimental conditions of 0.1% dodecyl maltoside should produce a mixture of both monomers and dimers (Jastrzebska et al., 2006). Additionally, even though rh1-2 is expressed at much lower levels compared to rhodopsin, its presence could still have a significant influence on rhodopsin function, similar to autosomal dominant rhodopsin mutants causing retinitis pigmentosa (Wilson and Wensel, 2003). When examining the hypothesized dimerization surface along TM4 and 5 (Liang et al., 2003; Guo et al.,

2005), it is interesting to note that amino acid sequence similarity between zebrafish rh1 and rh1-2 actually decreases in this region [75.5%, from 78.8% overall], while similarity between zebrafish rh1 and bovine RH1 increases significantly [87.8%, from 79.9% overall]. Despite these differences, several key residues thought to be critical to dimerization, such as W175 and Y206 are conserved in rh1-2 (Kota et al., 2006).

If rh1-2 does not share any direct association with rhodopsin in vivo, it is still most likely to function as a visual opsins gene similar to rhodopsin; this conclusion is drawn based

on a similar λMAX, spatial expression pattern, and retinal release rate to zebrafish rhodopsin. It is less likely that rh1-2 is a non-visual opsin, like exo-rhodopsin, due to the significantly faster rate of retinal release measured in exo-rhodopsin, and, more importantly, the fact that non-visual opsins have not been detected in rod and cone photoreceptors of vertebrates. Instead, non-visual opsins are found in other retinal cells, including horizontal cells (Cheng et al., 2009) and retinal ganglion cells (Dacey et al.,

2005; Panda et al., 2005), as well as in neural tissues outside of the retina, including the pineal (Mano et al., 1999), cerebellum (Blackshaw and Snyder, 1999), and spinal chord

(Tartellin et al., 2003). The two main discrepancies between rh1-2 and traditional

116 features of rh1 are the onset of expression, which is later in rh1-2 than would be expected of an rh1 gene, and lower stability of the expressed rh1-2 protein. Future studies that could help clarify whether rh1-2 heterodimerizes with rhodopsin could include crosslinking mutagenesis or fluorescence resonance energy transfer [FRET] in order to determine the proximity of the two opsins after expression. While immunohistochemistry is another possibility, generating antibodies that can distinguish between rhodopsin and rh1-2 may be challenging. However, regardless of this potential interaction, the presence of the rh1-2 gene family in ostariophysian fish should promote additional investigation into the influence of a second visual rhodopsin-like gene on the visual system.

117 rh1 rh1-2

Figure 1. In situ hybridization [ISH] of rh1-2 in the zebrafish retina at various developmental stages. (A-B) Whole mount ISH of 5 dpf embryos showing strong expression of rh1 and weak expression of rh1-2 in the peripheral retina. (C-F) ISH of juvenile eyes at 21 dpf showing strong expression of rh1 in both the central and peripheral retina, but weak only weak expression of rh1-2 in the peripheral retina. (G-H) ISH of adult eyes at 175 dpf showing strong expression of rh1 in the central retina, but no signal from rh1-2. Both rh1 and rh1-2 also showed expression in the peripheral retinal at 175 dpf (results not shown). All expression of rh1 and rh1-2 was confined to the outer nuclear layer of the retina, consisting of the cell bodies of rod and cone photoreceptors.

118

Figure 2. Retinal release rates of various zebrafish rhodopsins at 20oC. The release of all-trans-retinal is represented by an increase in fluorescence intensity following photoactivation of zebrafish rhodopsin [black], rh1-2 [blue], and exo-rhodopsin [red]. Half-life values are averages of at least three replicates, while data shown for comparative purposes are from experiments representative of these averages.

119

Figure 3. Co-transfection with zebrafish rh1-2 alters thermal stability of rhodopsin. The release of all-trans-retinal is represented by an increase in fluorescence intensity following the incubation of zebrafish rhodopsin alone [black], or with rh1-2 [red] at 56oC. Every assay consisted of both a control [rhodopsin] and experimental [rhodopsin & rh1-

2] sample being run simultaneously, with the relative half-life [t1/2Rel] being the ratio of a sample to the half-life of the control for that assay. Relative half-life values are averages of at least three replicates, while data shown for comparative purposes are from experiments representative of these averages. Data for rh1-2 alone was not collected because it could not be expressed at high enough levels to be assayed.

120 RH2 Lamprey RH1 Chondrichthyes RH1 Coelocanth RH1 Lungfsh RH1 Amphibian RH1 Non-avian Reptile RH1 Avian RH1 Mammal RH1

Exorhodopsin Chondrostei RH1 Holostei RH1 Elopiformes RH1 Anguilliformes DSO Osteoglossiformes RH1 Anguilliformes FWO RH1-2 Ostarioclupimorpha RH1 Euteleostei RH1

Figure 4. A schematic of the position of rh1-2 gene family with other vertebrate visual pigment genes based on a Bayesian phylogenetic analysis. This topology places the rh1- 2 gene family as the sister group to rh1 genes from Ostarioclupimorpha [Clupemorpha plus Ostariophysi].

121

Figure S1. In situ hybridization of rh1 in 3 dpf zebrafish embryos. Embryos were stained with the same 700 bp rh1 probe used in all other experiments in this study both (A) with and (B) without the addition of full-length rh1-2 blocking RNA, present at double the concentration of the rh1 probe. No significant difference in staining was detected.

122

Figure S2. Detailed Bayesian phylogeny of teleost rh1 and rh1-2 genes. The resulting topology mostly recovered expected relationships, including the positioning of lampreys, chondrichthyes, lobe-finned fish, amphibians, amniotes, and ray-finned fish. High support was recovered for the placement of the rh1-2 gene family as the sister group to Ostarioclupimorpha. Several rh2 opsin sequences were used as an outgroup.

123 Species Name Common Name Gene Accession/GI Number Amphiprion melanopus Fire clownfish exorh HM107820.1 Danio rerio Zebrafish exorh 71534272 Oncorhynchus masou Masu salmon exorh 478430872 Oreochromis niloticus Nile tilapia exorh XM003438995.1 Oryzias latipes Japanese medaka exorh XM004070540.1 Paralichthys olivaceus Olive flounder exorh HM107825.1 Phreatichthys andruzzii Cavefish exorh GQ404491.1 Plecoglossus altivelis Ayu exorh 28201134 Salmo salar Atlantic salmon exorh 185133057 Takifugu rubripes Pufferfish exorh 76362825 Anguilla japonica Japanese eel rh1dso 5931797 Conger myriaster Whitespotted conger rh1dso 12583666 Anguilla japonica Japanese eel rh1fwo 5931795 Conger myriaster Whitespotted conger rh1fwo 12583664 Abudefduf sexfasciatus Scissortail sergeant rh1 HQ286548.1 Acipenser sp. Sturgeon rh1 AF137206.1 Alepocephalus agassizii Slickhead rh1 JN544545.1 Alligator mississippiensis American alligator rh1 U23802.1 Ambystoma tigrinum Tiger salamander rh1 U36574.1 Amia calva Bowfin rh1 AF137208.1 Amphiprion melanopus Fire clownfish rh1 HM107824.1 Anchoa cayorum Key anchovy marine rh1 This study Anchoviella sp. Freshwater anchovy rh1 This study Anolis carolinensis Carlonina anole rh1 XM003224879.1 Apteronotus albifrons Black ghost knifefish rh1 JN230983.1 Arapaima gigas rh1 JN230972.1 Astyanax mexicanus Mexican tetra rh1 U12328.1 Atherina boyeri Big-scale sand smelt rh1 Y18676.1 Atractosteus tropicus Tropical gar rh1 JN230970.1 Barilius Sp. rh1 This study Bathysaurus ferox Deepsea lizardfish rh1 JN412585.1 Bathysaurus mollis Highfin lizardfish rh1 JN412586.1 Bos taurus Cow rh1 NM001014890.1 Bufo bufo Common toad rh1 U59921.1 Carassius auratus Golfish rh1 L11863.1 Chanos chanos Milfish rh1 JN230981.1 Chauliodus macouni Pacific viperfish rh1 EU407250.1 Chelon labrosus Thicklip grey mullet rh1 Y18669.1 Chologaster cornuta Swampfish rh1 HQ729684.1 Chrysemys picta Painted turtle rh1 Extracted from genome Columba livia Pigeon rh1 4887218-4887219 Cottus kessleri Kesslers sculpin rh1 L42953.1 Cynops pyrrhogaster Fire-belly newt rh1 14041791 Danio albolineatus Pearl danio rh1 JQ614122.1 Danio rerio Zebrafish rh1 18859316: Danio roseus Rose danio rh1 JQ614148.1 Dasypus novemcinctus Armadillo rh1 XM004477246.1 Diaphus metopoclampus Spothead lantern fish rh1 JN544536.1

124 Elops saurus Ladyfish rh1 JN230971.1 Engraulis japonicus Japanese anchovy rh1 AB731902.1 Epalzeorhynchos bicolor Redtailed black shark rh1 This study Eurypharynx pelecanoides Pelican eel rh1 JN544544.1 Gadus morhua Atlantic cod rh1 AF385832.1 Galeus melastomus Blackmouth catshark rh1 Y17586.1 Gallus gallus Chicken rh1 NM001030606.1 Gasterochisma melampus Butterfly kingfish rh1 DQ882021.1 Geotria australis Pouched lamprey rh1 AY366493.1 Gnathonemus petersii Elephantnose fish rh1 This study Grammicolepis brachiusculus Thorny tinselfish rh1 EU637964.1 Gymnothorax tile Moray eel rh1 This study Gymnotus tigre Tiger knifefish rh1 This study Halosauropsis macrochir Abyssal halosaur rh1 JN544541 Hippoglossus hippoglossus Atlantic halibut rh1 AF156265.1 Histiobranchus bathybius Deepwater arrowtooth eel rh1 JN544542.1 Homo sapiens Human rh1 169808383 Hoplostethus mediterraneus Mediterranean slimehead rh1 JN412583.1 Idiacanthus antrostomus Pacific blackdragon rh1 EU407249.1 Latimeria chalumnae Coelacanth rh1 4836673-4836677 Lepisosteus oculatus Spotted gar rh1 This study Lepisosteus osseus Longnose gar rh1 AF137207.1 Lethenteron camtschaticum Artic lamprey rh1 46917273 Leucoraja erinacea Little skate rh1 U81514.1 Loxodonta africana Elephant rh1 344275992 Lucania goodei Bluefin killifish rh1 AY296738.1 Megalops atlanticus Atlantic tarpon rh1 AY158050 Megalops cyprinoides Indo-Pacific tarpon rh1 This study Melanotaenia australis Western rainbowfish rh1 FJ940704.1 Monodelphis domestica Short-tailed opossum rh1 XM001366188.1 Mul cephalus Flathead mullet rh1 4210736 Mullus surmuletus Striped red mullet rh1 4210732 Muraena lentiginosa Moray eel rh1 This study Neoceratodus forsteri Lungfish rh1 EF526295.1 Notacanthus bonaparte rh1 JN544543 Oncorhynchus masou Masu salmon rh1 478430870 Orectolobus maculatus Spotted wobbegong rh1 JX534163.1 Orectolobus maculatus Spotted wobbegong rh1 JX534163.1 Oreochromis niloticus Niles tilapia rh1 348502996 Ornithorhynchus anatinus Platypus rh1 NM001127627.1 Oryzias latipes Japanese medaka rh1 66796119 Osmerus eperlanus Smelt rh1 JN230996.1 Osteoglossum bicirrhosum Silver arowana rh1 This study Pantodon buchholzi Freshwater butterflyfish rh1 AF137210.1 Paracheirodon innesi Neon tetra rh1 84095053 Paralichthys olivaceus Olive flounder rh1 HQ413772.1 Pelodiscus sinensis Chinese softshell turtle rh1 Extracted from genome Petromyzon marinus Sea lamprey rh1 1513320-1513324 Photoblepharon palpebratum Eyelightfish rh1 EU637993.1 Phreatichthys andruzzii Cavefish rh1 JQ413240.1

125 Phycis blennoides Greater forkbeard rh1 JN412579.1 Phyllophichthus xenodontus Flappy snake-eel rh1 This study Pimelodus blochim Blochs catfish rh1 This study Plecoglossus altivelis Ayu rh1 19912833 Poecilia reticulata Guppy rh1 DQ912024.1 Polyodon spathula American paddlefish rh1 AF369050.1 Protopterus sp. African lungfish rh1 AF369054.1 Pseudobagrus tokiensis Balgrid rh1 FJ197075.1 Pterigoplichthys sp. Armored catfish rh1 This study Rana temporaria Common frog rh1 U59920.1 rella punctata Largescale blackfish rh1 AB158262.3 Salmo salar Atlantic salmon rh1 185133085 Sarcophilus harrisii Tasmian devil rh1 395516643 Sardina pilchardus Sardine rh1 Y18677.1 Sargocentron diadema Crown squirrelfish rh1 U57537.1 Scleropages jardini Gulf saratoga rh1 This study Scyliorhinus canicula Small-spotted catshark rh1 3256019 Scyliorhinus canicula Smaller spotted catshark rh1 3256019 Sebastolobus altivelis Longspine thornyhead rh1 DQ490124.1 Serrivomer beanii Sawtooth eel rh1 This study Sewellia lineolata Tiger hillstream loach rh1 This study Solea solea Common sole rh1 4210866 Stegastes gascoynei Coral Sea gregory damselfish rh1 HQ286557.1 Stenobrachius leucopsarus Northern lampfish rh1 EU407251.1 Synodus foetens Inshore lizardfish rh1 JN231001.1 Tachyglossus aculeatus Echidna rh1 398018454 Taeniopya guttata Zebra finch rh1 115529257 Takifugu rubripes Pufferfish rh1 118344635 Tetraodon nigroviridis Green spotted puffer rh1 AJ293018.1 Typhlichthys subterraneus Southern cavefish rh1 HQ729699.1 Uta stansburiana Side-blotched lizard rh1 DQ100323.1 Xenopeltis unicolor Sunbeam snake rh1 FJ497233.1 Xenopus laevis African clawed frog rh1 93007319 Zeus faber John dory rh1 Y14484.1 Barilius Sp. rh1-2 This study Carassius auratus Goldfish rh1-2 This study Danio albolineatus Pearl danio rh1-2 HQ286328 Danio rerio Zebrafish rh1-2 HQ286326 Danio roseus Rose danio rh1-2 HQ286327 Epalzeorhynchos bicolor Redtailed black shark rh1-2 HQ286329 Misgurnus anguilicaudatus Dojo loach rh1-2 This study Scopelarchus analis Pearleye rh1A EF517404.1 Scopelarchus analis Pearleye rh1B EF517405.1 Gallus gallus Chicken rh2 45382766 Latimeria chalumnae Coelacanth rh2 4836680-4836684 Oryzias latipes Japanese medaka rh2 86198067 Danio rerio Zebrafish rh2-1 42476236 Anas platyrhynchos Mallard (duck) rh1 XM005012054.1 Corvus macrorhynchos Crow rh1 AB555651.1

126 Falco cherrug Falcon rh1 XM005443603.1

Table S1. List of sequences used in the phylogenetic and molecular evolutionary analyses of rh1 genes.

127 Parameters2 Tree1 Model np lnL К Null LRT df p ω0/p ω1/q ω2/ωp Vert M0 285 -51405.1 1.9 0.07 n/a M1a 286 -50828.3 2.0 0.06 (93.0%) 1 (7.0%) M0 1153.6 1 0.000 M2a 288 -50828.3 2.0 0.06 (93.0%) 1 (1.0%) 1 (6.0%) M1a 0.0 2 1.000 M3 289 -49434.4 1.9 0.01 (56.0%) 0.11 (33%) 0.34 (11%) M0 3941.4 4 0.000 M7 286 -49371.6 1.9 0.38 0.11 n/a M8a 287 -49363.0 1.9 0.42 4.27 1 (1.0%) n/a M7 17.1 2 0.000 M8 288 -49363.0 1.9 0.42 4.27 1 (1.0%) M8a 142.7 -1 1.000 Acti M0 203 -34366.8 2.0 0.07 n/a M1a 204 -33791.0 2.1 0.06 (93.0%) 1 (7.0%) M0 1151.6 1 0.000 M2a 206 -33791.0 2.1 0.06 (93.0%) 1 (5.0%) 1 (2.0%) M1a 0.0 2 1.000 M3 207 -32962.3 2.0 0.01 (56.0%) 0.10 (33%) 0.38 (11%) M0 2809.0 4 0.000 M7 204 -32909.1 2.0 0.31 2.40 n/a M8a 205 -32892.5 2.0 0.35 3.23 1 (1.0%) n/a M7 33.1 2 0.000 M8 205 -32892.5 2.0 0.35 3.23 1 (1.0%) M8a 0.0 1 1.000 rh1-2 M0 13 -2976.8 2.5 0.07 n/a M1a 14 -2952.9 2.5 0.04 (94.0%) 1 (6.0%) M0 47.7 1 0.000 M2a 16 -2952.9 2.5 0.04 (94.0%) 1 (6.0%) 11.15 (0.0%) M1a 0.1 2 0.956 M3 17 -2944.2 2.4 0.00 (68.0%) 0.21 (31.0%) 2.72 (1.0%) M0 65.1 4 0.000 M7 14 -2947.5 2.5 0.16 1.62 n/a M8a 15 -2946.1 2.4 0.22 3.03 1 (2%) n/a M7 5.0 2 0.081 M8 16 -2944.9 2.5 0.201 2.39 3.06 (1%) M8a 2.4 1 0.123

1The full vertebrate RH1 (Vert) and gene trees pruned to contain only actinopterygian rhodopsins (Acti) and only RH1-2. 2 ω values of each site class are shown for models M0-M3 (ω0– ω2) with the proportion of each site class in parentheses. For M7-M8, the shape parameters, p and q, which describe the beta distribution are listed. In addition, the ω value for the positively selected site class (ωp, with the proportion of sites in parentheses) is shown for M8a (where ωp is constrained to equal one) and M8. Abbreviations—np, number of parameters; lnL, ln Likelihood; К, transition/transversion ratio; LRT, likelihood ratio test statistic; df, degrees of freedom; p, p-value; n/a, not applicable.

Table S2. Results of random sites (PAML) analyses on subsets of the vertebrate rh1 gene tree.

128 Parameters2 Partition Model np lnL К LRT df p ω0 ω1 ω2a ω2b M0 Null 203 -34366.8 2.0 0.07 B: 0.07 0.57 1 0.452 Branch Alt 204 -34366.6 2.0 F: 0.06 B: 0.06 B: 1 B: 0.06 B: 1 Branch-site Null 205 -33784.8 2.1 F: 0.06 F: 1 F: 1 F: 1 (85.8%) (6.4%) (7.2%) (0.5%) 4.86 1 0.028 B: 0.06 B: 1 B: 0.06 B: 1 Branch-site Alt 206 -33782.4 2.1 F: 0.06 F: 1 F: 39.5 F: 39.5 (89.0%) (6.7%) (4.4%) (0.3%) 0.01 1 0.17 M2a_rel Null 207 -33050.5 2.0 (62.9%) (2.7%) (34.4%) B: 0.01 B: 1 B: 0.16 0.424 1 0.515 CmC Alt 207 -33050.5 2.0 F: 0.01 F: 1 F: 0.17 (62.9%) (2.7%) (34.4%) 2ω values of each site class are shown with the proportion of each site class in parentheses. B and F refer to the background and foreground partitions. Abbreviations—np, number of parameters; lnL, ln Likelihood; К, transition/transversion ratio; LRT, likelihood ratio test statistic; df, degrees of freedom; p, p-value; n/a, not applicable.

Table S3. Results of branch, branch-site, and clade model C (PAML) analyses with the branch leading to the rh1-2 clade placed into the foreground.

129 Parameters2 Partition Model np lnL К LRT df p ω0 ω1 ω2a ω2b M0 Null 203 -34366.8 2.0 0.07 B: 0.07 1.12 1 0.289 Branch Alt 204 -34366.3 2.0 F: 0.07 B: 0.06 B: 1 B: 0.06 B: 1 Branch-site Null 205 -33784.8 2.1 F: 0.06 F: 1 F: 1 F: 1 (88.7%) (6.9%) (4.2%) (0.3%) 0.00 1 0.911 B: 0.06 B: 1 B: 0.06 B: 1 Branch-site Alt 206 -33782.4 2.1 F: 0.06 F: 1 F: 1 F: 1 (88.7%) (6.9%) (4.2%) (0.3%) 0.01 1 0.17 M2a_rel Null 207 -33050.5 2.0 (62.9%) (2.7%) (34.4%) B: 0.01 B: 1 B: 0.17 0.023 1 0.880 CmC Alt 207 -33050.5 2.0 F: 0.01 F: 1 F: 0.15 (62.9%) (2.7%) (34.4%) 2ω values of each site class are shown with the proportion of each site class in parentheses. B and F refer to the background and foreground partitions. Abbreviations—np, number of parameters; lnL, ln Likelihood; К, transition/transversion ratio; LRT, likelihood ratio test statistic; df, degrees of freedom; p, p-value; n/a, not applicable

Table S4. Results of branch, branch-site, and clade model C (PAML) analyses with the rh1-2 clade placed into the foreground.

130 References

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139

Chapter 4:

Investigating retinal release in light-activated

zebrafish rhodopsin using fluorescence

spectroscopy: the role of steric effects and Schiff

base stability

140 4.1 Abstract

Rhodopsin is the visual pigment within vertebrate rod photoreceptor cells responsible for initiating scotopic [dim-light] vision. Once activated by light, decay of the active metarhodopsin II [meta II] state of rhodopsin involves hydrolysis of the Schiff base link, followed by dissociation of retinal from the protein moiety. This process of meta II decay followed by retinal release is known to be a key step in the cycle by which rhodopsin is reset to its native dark state configuration, so that it can again be activated by light. It has been well studied in model systems such as bovine rhodopsin, but not in cold-blooded animals in which physiological temperatures might vary considerably.

Here, we characterize retinal release of rhodopsin in a poikilotherm, the zebrafish [Danio rerio], demonstrating in a fluorescence assay that it is more than twice as fast as bovine rhodopsin. Using site-directed mutagenesis, we found that the differences in retinal release rates can be attributed to a series of variable residues lining the retinal channel in three key motifs: an opening in activated rhodopsin between transmembrane helix 5

[TM5] and TM6, TM3 near E122, and the ‘retinal plug’ formed by extracellular loop 2

[EL2]. The majority of these sites are more proximal to the β-ionone ring of retinal than the Schiff base, suggesting their influence on retinal release is based more on steric effects during retinal dissociation, rather than affecting Schiff base stability. An

Arrhenius plot of zebrafish rhodopsin was consistent with this model, indicating an activation energy for Schiff base hydrolysis very similar to that of bovine rhodopsin.

Variation at key sites identified in this study suggests that retinal release might be an adaptive property of rhodopsin in vertebrates. Our study is one of the few investigating the structure and function of non-mammalian rhodopsins, which will help to establish a

141 better understanding of the molecular mechanisms that contribute to in vivo processes such as light response recovery and dark adaptation.

4.2 Introduction

Rhodopsin, a member of the pharmacologically significant G protein-coupled receptor

[GPCR] superfamily, is the visual pigment responsible for mediating the critical first step of dim-light vision in vertebrates (Burns and Baylor, 2001). The 11-cis-retinal chromophore, bound through a protonated Schiff base linkage at K296 (Ebrey and

Koutalos, 2001), serves as a strong inverse agonist, maintaining stability and greatly reducing thermal activation of dark state rhodopsin (Palczewski et al., 2000). Upon activation, 11-cis-retinal is converted to its stereoisomer, all-trans-retinal, resulting in a series of conformational changes in rhodopsin, including the metarhodopsin II [meta II] state, which activates the G protein transducin (Pugh and Lamb, 1993). Within milliseconds of formation, meta II is subjected to a two-step deactivation process, comprising C-terminal phosphorylation followed by arrestin binding, quenching the catalytic activity of the activated state (Gross and Burns, 2010). For rhodopsin to regenerate in vivo, a new Schiff base linkage must form following release of all-trans- retinal (Hofmann et al., 1992; Pulvermüller et al., 1997), involving hydrolysis of the existing Schiff base linkage, and dissociation of retinal from opsin.

Retinal release has been examined in detail in recent years in part due to the resolution of crystal structures of not only the dark state (Palczewski et al., 2000), but of the open pocket (Park et al., 2008) and activated states as well (Choe et al., 2011). A notable but

142 previously unanticipated aspect of the dark state structure was a b-sheet consisting of strands from the N-terminal region and extracellular loop 2 [EL2], effectively plugging the extracellular opening in rhodopsin (Palczewski et al., 2000; Okada et al., 2004), and blocking what was believed to be one of the possible access sites of retinal. Further investigation suggested the possibility of a channel within the transmembrane domains that would facilitate the travel of retinal to and from the chromophore binding pocket, along with terminal binding sites for retinal on the surface of rhodopsin (Schädel et al.,

2003; Wang and Duan, 2007). Crystal structures for the chromophore-free (Park et al.,

2008), G protein-interacting (Scheerer et al., 2008), and meta II (Choe et al., 2011) states of rhodopsin, confirmed the existence of two transmembrane openings on opposite sides of rhodopsin that open into the hydrophobic membrane layer: one between TM1 and

TM7, and one between TM5 and TM6 (Hildebrand et al., 2009). The latter opening is a good candidate for retinal release, considering the outward rotation of TM6 following activation (Farrens et al., 1996; Altenbach et al., 2008), and molecular dynamics simulations that have shown the possibility of retinal exiting through this opening, led by its β–ionone ring (Wang and Duan, 2011). Recent work has attributed changes in retinal release rates to either Schiff base stability (Piechnick et al., 2012) or retinal dissociation

(Chen et al., 2012), suggesting conflicting ideas currently exist about the dominant mechanism mediating this kinetic rate. While these studies examined the contributions of conserved sites in rhodopsin, and differences between rod and cone opsins, the mechanism that mediates natural variation of retinal release among rhodopsins has yet to be investigated.

143 Despite being the one of the most extensively studied members of the GPCR superfamily, the vast majority of biochemical studies of rhodopsin have utilized bovine and other model mammalian rhodopsins (e.g., Liu et al., 2013), or chicken rhodopsin (e.g.,

Kuwayama et al., 2005). Moreover, an investigation of retinal release in the echidna suggests that variation in kinetic rates do exist even among mammalian rhodopsins

(Bickelmann et al., 2012), even if the differences are subtler than those between rhodopsin and cone opsins (Chen et al., 2012). This is not surprising considering that other functional differences between rhodopsins have also been measured, including hydroxylamine stability (Yokayama and Kawamura, 1998), and meta intermediate kinetics (Imai et al., 1995; Lewis et al., 1997). Further investigations of the natural variation of retinal release in rhodopsin should include organisms where kinetic rates would be expected to vary. A poikilotherm, like zebrafish, is an ideal candidate to highlight natural differences in rhodopsin relative to homeothermic mammals, as kinetic processes have likely adapted to function at variable physiological temperatures. This suggests that rates of kinetic processes, such as retinal release, are likely to vary between poikilotherms and homeotherms. Additionally, zebrafish is a well-studied model for the vertebrate visual system, where increased understanding of rhodopsin structure and function may reveal insight into visual development, as well as behavior and physiology

(Muto et al., 2005; Fadool and Dowling, 2008; Maurer et al., 2011), despite the fact that zebrafish is not a nocturnal organism. Since it has been suggested that retinal release rates influence dark adaptation at low bleaching levels (Ala-Laurila et al., 2006), additional insight into the variation of retinal release among rhodopsins could help contrast dim-light vision among vertebrates.

144 Here, we investigate the structural basis of the natural variation in retinal release rates in rhodopsin by characterizing zebrafish rhodopsin, along with mutants based on bovine rhodopsin residues at variable sites. We found that zebrafish rhodopsin releases retinal more than twice as fast bovine rhodopsin. This difference is mainly attributable to the rate of dissociation of retinal from opsin, not to differences in Schiff base stability, as variation at functional motifs that are proximal to the β-ionone ring of retinal are responsible for most of this difference. Additionally, an Arrhenius plot of zebrafish rhodopsin reveals a similar energy of activation required to hydrolyze the Schiff base compared to bovine rhodopsin. This study also reinforces the importance of sites 122 and 189 on the stability of metarhodopsin intermediates, where substitutions that influence the positioning of these sites in the chromophore binding pocket also affect retinal release. This is the first time that a fish rhodopsin has been investigated for any

function other than λmax. Implications of these findings for structure/function studies of rhodopsin, and for the molecular evolution of this gene as a whole are discussed.

4.3 Materials and methods

4.3.1 Visual pigment expression and purification

RNA was extracted from adult zebrafish eyes using the TRIzol reagent (Invitrogen), and cDNA libraries were generated using the SMART cDNA Library Construction Kit (BD

Biosciences). The complete coding sequence of zebrafish rhodopsin (GenBank:

AB087811) was amplified using PfuTurbo (Stratagene) and inserted into the pJET1.2 cloning vector (Fermentas). Site-directed mutagenesis primers were designed to induce single amino acid substitutions, with double and triple mutants being generated through

145 successive rounds of mutagenesis via PCR. The complete coding sequence of bovine rhodopsin was cloned into the pIRES-hrGFP II expression vector (Stratagene), while zebrafish rhodopsin and mutants were cloned into the p1D4-hrGFP II expression vector

(Morrow and Chang, 2010). These constructs were used to transiently transfect cultured

HEK293T cells using Lipofectamine 2000 (Invitrogen; 8 ug of DNA per 10-cm plate).

Cells were harvested 48 h post-transfection and opsins were regenerated using 11-cis- retinal, generously provided by Dr. Rosalie Crouch (Medical University of South

Carolina). Visual pigments were solubilized in 1% N-dodecyl--D-maltoside [DM] and immunoaffinity purified with the 1D4 monoclonal antibody (Molday and MacKenzie,

1983), as previously described (Morrow and Chang, 2010; Morrow et al., 2011). Purified visual pigment samples were eluted in sodium phosphate buffer [50 mM NaPhos, 0.1%

DM, pH 7].

4.3.2 Spectroscopy

The ultraviolet-visible absorption spectra of purified zebrafish, bovine, and mutant rhodopsins were recorded at 25oC using the Cary4000 double-beam spectrophotometer

(Varian) and quartz absorption cuvettes (Helma). Samples were incubated and assayed in

sodium phosphate buffer. All λMAX values were calculated after fitting absorbance spectra to a standard template for A1 visual pigments (Govardovskii et al., 2000). Zebrafish

rhodopsin activated in response to light, shifting its λMAX to ~380 nm, representing to active meta II state. Reactivity to hydroxylamine in the dark was determined by

o incubating zebrafish rhodopsin in 50 mM NH2OH (Sigma-Aldrich) at 25 C. The protocol used to determine retinal release rates of visual pigments was modified from that of

146 Farrens and Khorana (1995). Briefly, 0.1-0.2 μM visual pigment samples in submicro fluorometer cell cuvettes (Varian) were bleached for 30 seconds using a Fiber-Lite MI-

152 Illuminator external light source (Dolan-Jenner), using a filter used to restrict wavelengths of light below 475 nm. Fluorescence measurements were integrated for 2 seconds at 30-second intervals using a CaryEclipse fluorescence spectrophotometer

(Varian). The excitation wavelength was 295 nm [1.5 nm slit width] and the emission wavelength was 330 nm [10 nm slit width]; no noticeable pigment bleaching by the excitation beam was detected. Retinal release was demonstrated through a sharp initial rise in intrinsic tryptophan fluorescence, representing a decrease in fluorescent quenching of W265 by the retinal chromophore. Data from the initial rise was fit to a three variable,

-bx first order exponential equation [y = yo + a(1-e )], with half-life values calculated based

2 on the rate constant 'b' [t1/2 = ln2/b]. All curve fitting resulted in r values of greater than

0.95. Differences in retinal release rates were assessed using a two-tailed t test with unequal variance, where p values of < 0.05 were deemed significant. All data for zebrafish rhodopsin mutants was recorded at 20oC, while Arrhenius plot data for zebrafish rhodopsin was collected at 5oC, 18oC, 20oC, and 24oC, and bovine rhodopsin was also assayed at 37oC.

4.3.3 Homology Modeling

The 3D structures of zebrafish rhodopsin was inferred via homology modeling by

Modeller (Sali and Blundell, 1993; Eswar et al., 2006), using bovine meta II (PDB code:

3PQR; Choe et al., 2011) as template, including chromophore and water molecules.

Minimizing Modeller objective function generated twenty-five separate models, and the

147 run with the lowest DOPE score was assessed and visualized (Shen and Sali, 2006).

ProCheck was used to ensure bond angles and lengths were generally in high probability stereochemical conformations as indicated by positive overall G-factor (Laskowski et al.,

1993). Model quality was further examined by ProSA-web by comparing the model’s total energy to that expected by random chance (Wiederstein and Sippl, 2007). Our model and template structures had comparable z-scores, standardized for the number of residues. Structure images were generated using MacPyMol (DeLano, 2008).

4.4 Results

4.4.1 Biochemistry of zebrafish rhodopsin

Both bovine and zebrafish rhodopsin were expressed, regenerated with 11-cis-retinal, and

purified, producing characteristic dark spectra with λMAX of 499 and 501 nm, respectively

(Fig. 1). These values are similar to previously published results (Oprian et al., 1987;

Chinen et al., 2003), and are consistent with λMAX values from the majority of vertebrates, which are approximately 500 nm (Menon et al., 2001; Bowmaker et al., 2008). Zebrafish rhodopsin was not susceptible to hydroxylamine (Fig. S1), making it similar to bovine rhodopsin, which shows no significant reaction over an extended period of time

(Kawamura and Yokoyama, 1998). When light-bleached and monitored for release of all-trans-retinal, bovine rhodopsin had a half-life of 13.9 min, similar to previously recorded values of 12.5-15.5 min (Farrens and Khorana, 1995; Yan et al., 2002), while zebrafish rhodopsin produced a much shorter half-life of 6.6 min (Fig. 2; Table 1). To elucidate the cause of this discrepancy, we investigated sites lining the retinal channel of rhodopsin, as well as those surrounding terminal openings of this channel. We generated

148 mutations in zebrafish rhodopsin to bovine rhodopsin identities, in the hopes of bridging the phenotypic gap between these two visual pigments (Table 1). To avoid experimental bias, we also included substitutions in all other structural domains of rhodopsin that were not associated with the retinal channel.

4.4.2 Effects of mutations at openings in metarhodopsin II

We first investigated substitutions near an opening formed in meta II between TM5 and

TM6, which is near the β-ionone ring of retinal and hypothesized to be the site of retinal dissociation (Hildebrand et al., 2009; Wang and Duan, 2011; Piechnick et al., 2012). The largest difference was caused by W273F, also the substitution most proximal to the opening, which changed the retinal release rate half-life to 8.8 min, an increase of 2.2 min compared to wild type. The effects of other substitutions also seemed to be connected to their proximity to the opening, as three additional nearby substitutions had a minor influence on retinal release: I209V [5.3 min; -1.3], F213I [8.1 min; +1.5], and V266L

[8.3 min; +1.7]. Meanwhile, R225Q, E241A, and R248K were not as close to the opening, and did not significantly affect retinal release. A quadruple mutant [I209V,

F213I, V266L, W273F] was also generated to determine whether the influence of these residues was additive, and caused a much larger shift in retinal release relative to the individual substitutions [13.1 min; +6.5] (Fig. 3). This provides evidence that variable site substitutions can be responsible for shifts in retinal release large enough to encompass the difference in rates between zebrafish and bovine rhodopsin.

149 A second, smaller opening also formed in meta II between TM1 and TM7 at the other end of the retinal channel, closer to the Schiff base link with K296 of rhodopsin (Park et al., 2008; Choe et al., 2011). Overall, the effects of substitutions at this opening were less frequent, and smaller in magnitude compared to those at the TM5/TM6 opening.

Both V286I [5.7 min; -0.9] and L290I [5.3 min; -1.3] caused significant changes to retinal release rates, although they were only minor differences; meanwhile, both A36Q and C304V had no significant effect. These results follow the same trend as the substitutions near the TM5/TM6 opening, where increased proximity to the opening increases the chances of influencing retinal release. However, in this case, both V286I and L290I were also the substitutions closest to the Schiff base, suggesting that they might be able to influence retinal release by altering Schiff base stability.

4.4.3 Effects of mutations in the chromophore binding pocket

A series of residues from the tightly packed α-helices of TM3, TM5 and TM6 are responsible for keeping the β-ionone ring of retinal in its place in the chromophore binding pocket (Palczewksi et al., 2000; Okada et al., 2004). While these sites have the same identity in both zebrafish and bovine rhodopsin, there are other nearby residues that are variable. The single substitution with the most significant impact on retinal release in this study was M123I, which produced a half-life of 11.5 min, almost 5 minutes more than wild type zebrafish rhodopsin. The neighboring G124A also had an effect [8.4 min;

+1.8], although not as pronounced as that of M123I. A124 has previously been identified as a red-shifting counterpart to S124 in spectral tuning studies of deep-sea fish (Hunt et al., 2001). However, our results imply that the G124A substitution does not have a

150 significant effect on spectral tuning, despite the suggestion that G124 is responsible for a red-shift in rhodopsin relative to exo-rhodopsin in teleosts (Tarttelin et al., 2011).

Surprisingly, the double mutant M123I/G124A had an intermediate phenotype relative to the two individual substitutions [10.6 min; +4.0], which suggests that these substitutions are affecting the chromophore indirectly, by repositioning a nearby residue with a more direct influence. A likely candidate is the neighboring E122, a determinant of opsin function previously shown to mediate decay rates of metarhodopsin intermediates (Imai et al., 1997; Kuwayama et al., 2002). Homology models of zebrafish rhodopsin confirm the proximity of E122 to the β-ionone ring of the chromophore, similar to bovine rhodopsin (Fig. 4A). Other substitutions less proximal to E122 did not have significant effects on retinal release, including R107P and W136Y.

Forming the boundary between the chromophore binding pocket and the extracellular environment is a ‘retinal plug’ made up of B-strands from the N-terminus and EL2

(Palczewski et al., 2000; Sakai et al., 2010). Strand B4 contains site 189, a known determinant of rod and cone opsin function (Kuwayama et al., 2005) that also varies between valine and isoleucine among rhodopsins. The V189I substitution in zebrafish rhodopsin increased the retinal release half-life to 9.1 min [+2.3]. In bovine rhodopsin,

I189 is only a few angstroms away from the chromophore, and homology modeling suggests this holds true for V189 in zebrafish rhodopsin (Fig. 4B). Considering the tightly packed structure of chromophore binding pocket, the additional carbon atom of isoleucine relative to valine likely alters the position of the side chain relative to the chromophore, which could be responsible for the change in rate.

151 4.4.4 Effects of temperature on retinal release

Being a kinetic rate, another variable affecting retinal release is ambient temperature.

Additional retinal release measurements were collected for zebrafish at 5 and 18oC in order to construct an Arrhenius plot to measure the activation energy of hydrolysis of the

Schiff base in rhodopsin, which was calculated to be 18.2 kcal/mol at pH 7 (Fig. 5). This is comparable to previously measured activation energies of both bovine rhodopsin and

Xenopus violet (Chen et al., 2012). This suggests that the energy required to hydrolyze the Schiff base in all three of these visual pigments is similar, meaning Schiff base stability is likely only a minor contributor to any differences in retinal release. This result supports our mutagenesis data, which illustrated how the majority of variable site substitutions with significant effects on retinal release were located near the hypothesized exit site of retinal, or the β-ionone ring, as these sites are more likely to sterically hinder the dissociation of retinal from rhodopsin following Schiff base hydrolysis.

While the retinal release of zebrafish rhodopsin occurs more than twice as fast compared to bovine rhodopsin at uniform temperatures, zebrafish is poikilothermic, whose body temperature is dependent on its environment. This contrasts homeothermic mammals, whose body temperatures are usually regulated around 37oC. When measured at physiological temperatures, the half-life of bovine rhodopsin [2.0 min] was actually shorter than that of zebrafish [4.1 min] (Fig. 6). This implies that even though bovine rhodopsin has a slower retinal release rate than zebrafish rhodopsin at the molecular level, it releases retinal faster than zebrafish rhodopsin in vivo, and that these differences are driven by amino acid substitutions at non-conserved sites. These rates may also reflect

152 differences on a physiological level for properties such as dark adaptation. It also seems as though substitutions that have a larger effect on retinal release, such as 123 and 273, occur at more invariant sites, while those with more variability, such as 213 and 290, cause smaller changes (Fig. S2).

4.4.5 Effects of mutations on λMAX

Spectral tuning is the most scrutinized biochemical property of opsin function, with studies spanning all major phylogenetic groups of visual pigments, including a diverse taxonomical sampling (Fasick and Robsinson, 1998; Takahashi and Ebrey, 2003; Hunt et al., 2007; Takenaka and Yokoyama, 2007). However, compared to the vast base of literature addressing natural variation in spectral sensitivity, other important aspects of visual pigment function have received much less attention. Our study targeted variable sites lining the retinal channel and lining its terminal openings, and found that no single

substitutions changed λMAX by more than 2 nm (Table 1). This was not a surprising result, considering that substitutions were made between zebrafish and bovine rhodopsin

identities, and there is only a 2 nm difference between the λMAX values of these visual pigments (Oprian et al., 1987; Chinen et al., 2003). It should also be noted that mutants

with multiple substitutions were able to shift λMAX by more than this 2 nm difference, suggesting that the discrepancy between zebrafish and bovine rhodopsin is more likely the result of a combination of minor changes than a single substitution causing a 2 nm shift.

153 4.5 Discussion

Through heterologous opsin expression, fluorescence spectroscopy, and site-directed mutagenesis, we have shown that zebrafish rhodopsin releases retinal more than twice as fast as bovine rhodopsin, and have highlighted several structural motifs lining a previously identified retinal channel (Park et al., 2008; Hildebrand et al., 2009) where intrinsic sequence variation is capable of tuning retinal release rates. However, the energy of activation required to hydrolyze the Schiff base in zebrafish rhodopsin is comparable to that of bovine rhodopsin (Chen et al., 2012), suggesting that all-trans- retinal dissociation, and not Schiff base stability, is the molecular mechanism that likely mediates retinal release differences among rhodopsins. Additionally, at respective physiological temperatures, bovine rhodopsin releases retinal faster than zebrafish rhodopsin, implying that sequence evolution impacting retinal release may be tuning this kinetic rate as an adjustment to ambient temperature. Here, we discuss the importance of our findings to the understanding of molecular mechanisms mediating retinal release, their impact on the hypothesized unidirectional channeling of retinal passage, and their implications on the vertebrate visual system.

When the crystal structure of dark state bovine rhodopsin was first resolved, the Schiff base linkage with K296 was shown to be tightly constrained by parts of TM1, 2, and 7, while the β-ionone ring was similarly kept in place by residues in TMs 3, 5, and 6

(Palczewski et al., 2000; Okada et al., 2004). Perhaps more notably, an anti-parallel β- sheet from the N-terminus and EL2 was shown to form a 'retinal plug', blocking a suspected opening to the extracellular environment atop the α-helical barrel structure of

154 rhodopsin. Analyses identifying two putative hydrophobic binding sites on the surface of rhodopsin then lead to the development of a channeling hypothesis where retinal travels through opsin in a unidirectional manner (Schädel et al., 2003; Heck et al., 2003).

Resolved crystal structures of ligand-free and meta II rhodopsin supported this hypothesis by locating two openings that did not exist in the dark state, one located between TM5 and TM6, and another between TM1 and TM7, both caused by small backbone movements and different side-chain orientations during activation (Park et al., 2008;

Choe et al., 2011). Molecular dynamics simulations then highlighted how aromatic residues lining the TM5/TM6 opening could sterically hinder the release of retinal (Wang and Duan, 2011). However, this series of interesting results still required experimental evidence to confirm the unidirectional channeling theory, and to better understand the molecular mechanisms underlying retinal release.

This study explored the effects of substitutions at variable sites in zebrafish rhodopsin in the retinal channel to identify substitutions responsible for a faster retinal release compared to bovine rhodopsin. The opening located between TM5 and TM6 in meta II is hypothesized as the site of all-trans-retinal release following photoactivation (Schädel et al., 2003; Hildebrand et al., 2009). In bovine rhodopsin, key residues at this motif, including a triad of phenylalanines at 208, 273, and 276, form a ‘hydrophobic cage’ that could sterically hinder the release of all-trans-retinal (Choe et al., 2011; Wang and Duan,

2011). In zebrafish rhodopsin, site 273 is instead tryptophan, where a mutation to phenylalanine increased retinal release half-life by 2.2 min. This is also the only site in this study whose effects on retinal release had previously been characterized, with F273L

155 and F273Q being assayed in bovine rhodopsin and causing changes to retinal release

(Piechnick et al., 2012); however, neither L273 nor Q273 are naturally occurring variants.

Three other substitutions, I209V, F213I, and V266L also caused minor shifts of 1.3 to 1.7 min, while other substitutions less proximal to the opening, such as R225Q, E241A, and

R248K did not lead to significant differences. Meanwhile, a quadruple mutant [I209V,

F213I, V266L, W273F] had a more substantial effect on retinal release, increasing the half-life by 6.5 min and producing a phenotype similar to that of bovine rhodopsin. With the exception of W273, side chains of these residues might have little direct contact with retinal during release, but their identities could alter the position of surrounding residues.

These changes likely influence steric effects during all-trans-retinal dissociation, due to their proximity to the β-ionone ring and the hypothesized retinal exit site. Another terminal channel opening is located between TM1 and TM7. This opening is formed during activation with the backbone of F293 rotating 120o (Park et al., 2008; Choe et al.,

2011), and two nearby substitutions, V286I and L290I, altered retinal release half-life values by 0.9 and 1.3 min, respectively; two other substitutions near this opening, A36Q and C304V, did not affect retinal release. Despite not being found at the proposed site of retinal release, V286I and L290I were able to have minor effects on this rate, likely because of their proximity to either the Schiff base or to F293, and may have a minor influence on Schiff base stability.

Along with parts of TM5 and TM6, several residues in TM3, including E122, are responsible for stabilizing the β-ionone ring of retinal in dark state rhodopsin (Palczewski et al., 2000; Okada et al., 2004). Site 122 is a known functional determinant of

156 rhodopsin and cone opsin function, where mutations have previously shown to mediate kinetic properties of opsins such as meta II decay (Imai et al., 1995; Imai et al., 1997) and meta III decay (Kuwayama et al., 2005), along with physiological properties of the photoreceptor cell (Imai et al., 2007). While E122 is conserved in zebrafish rhodopsin,

M123I and G124A increased retinal release half-life values by 4.9 and 1.8 min, respectively. Surprisingly, a double mutant at these sites [M123I/G124A] had an intermediate effect, shifting the half-life by only 4.0 min. This suggests that M123 and

G124 are likely having an indirect effect on retinal release by altering the position of

E122 relative to the chromophore. Since E122 is a charged residue, it is possible that its repositioning could modify the hydrostatic environment of the binding pocket and its hydrogen bonding network (Yan et al., 2003; Liu et al., 2009). Other substitutions along

TM3, including R107P and W136Y, did not have a significant impact on retinal release.

The latter result was somewhat surprising, as Y136 is highly conserved in most rhodopsin-like GPCRs as the third residue of the D(E)RY motif, and this motif has previously been shown to be relevant to rhodopsin activation (Fahmy et al., 2000; Fritze et al., 2003).

Separating the retinal channel of rhodopsin from the extracellular environment is a β- sheet structure formed by β -strands from both the N-terminal region and EL2, where strand β4, consisting of sites 186-190 from EL2, lines the chromophore binding pocket

(Palczewski et al., 2000; Sakai et al., 2010). The V189I substitution in zebrafish rhodopsin adds an additional carbon atom to the tightly packed binding pocket, and increases the retinal release half-life by 2.3 min. Site 189 has also been identified as a

157 functional determinant for opsins, where changing between cone opsin and rhodopsin identities affects kinetic properties of meta intermediates (Kuwayama et al., 2002;

Kuwayama et al., 2005). However, this is the first study to reveal a biochemical impact of a substitution between V189 and I189, the two identities found in rhodopsin. One last substitution that caused a minor change in retinal release was C165L, found in TM4, shifting retinal release half-life by 0.9 min. Despite being located only 10 Å from retinal, the side chain of C165 is faces the hydrophobic environment outside of rhodopsin

(Palczewski et al., 2000). Because this substitution has such a minor effect on retinal release, it may instead be more relevant to the dimerization interface of rhodopsin, located along the extracellular surface of TMs 4 and 5 (Guo et al., 2005; Fotiadis et al.,

2006).

Recently, two key studies investigated the mechanism responsible for mediating retinal release. The first performed site-directed mutagenesis at highly conserved sites around both retinal channel openings in bovine rhodopsin (Piechnick et al., 2012). Significant differences were often shown for both retinal uptake and release in these mutants, leading to the conclusion that these substitutions were altering the stability of the Schiff base linkage. The authors argued that local steric effects of amino acid side chains alone could not explain why a single substitution would change rates of both retinal uptake and release. Another study used bovine rhodopsin and a short wavelength-sensitive cone opsin from Xenopus [VCOP] to explore the differences between retinal release in rod and cone opsins (Chen et al., 2012). While release from VCOP occurred 250 times faster than in bovine rhodopsin, Arrhenius plots revealed that activation energies in both visual

158 pigments were remarkably similar. With comparable energetics for Schiff base hydrolysis, the authors concluded that steric interactions of all-trans-retinal with side chains during release was the likely mechanism to explain differences in retinal release rates between rod and cone opsins.

Our mutagenesis results support the idea of retinal dissociation being the primary mechanism for mediating retinal release rates, as the majority of sites shown to alter release rates, as well as individual substitutions causing the most significant changes, were found closer to the β-ionone ring of retinal than to the Schiff base. It should be noted, however, that this conclusion is only viable when considering retinal release variation among rhodopsins, and that Schiff base stability is also an intrinsic component in determining retinal release rates, as was shown when Piechnick et al. (2012) mutated highly conserved residues responsible for establishing core structural stability in rhodopsin. In order to confirm the hypothesis derived from our mutagenesis results, we constructed an Arrhenius plot for zebrafish rhodopsin to measure the activation energy of

Schiff base hydrolysis. Previous measurements of activation energy for bovine rhodopsin are 18.5 and 15.8 kcal/mol at pH 6 and 8, respectively (Chen et al., 2012). We

calculated an Ea of 18.2 kcal/mol at pH 7 for zebrafish rhodopsin, suggesting a comparable activation energy to bovine rhodopsin, and, consequently, that differences

Schiff base stability are not largely responsible for the discrepancy between their retinal release rates, which supports the conclusions of our mutagenesis data.

Additionally, it was interesting to see that retinal release in zebrafish rhodopsin was slower than that in bovine rhodopsin when both were assayed at respective physiological

159 temperatures. This may hint at evolutionary adaptations in the rhodopsins of homeothermic vertebrates to slow down kinetic reactions in order to compensate for higher ambient temperatures. This concept also applies to other kinetic processes of visual pigments, including the lumirhodopsin (Szundi et al., 2010) as well as the metarhodopsin equilibrium, both in vitro (Lewis et al., 1997) and in vivo (Korenyak and

Govardovskii, 2012). Moreover, changes to the metarhodopsin equilibrium due to temperature can subsequently vary rates of G protein activation (Parkes et al., 1999).

Intriguingly, there is even evidence that rhodopsin in Drosophila is involved in thermotactic discrimination to help locate settings of ideal temperature (Shen et al., 2011), suggesting an even more fundamental connection between rhodopsin and temperature in its evolutionary history. Much of this association is likely rooted in the steep dependence observed between temperature and dark state Schiff base hydrolysis (Liu et al., 2011).

The hydrogen bond network surrounding the Schiff base mediates this relationship, where substitutions at residues involved in critical interactions can change the activation energy of hydrolysis (Janz and Farrens, 2004).

This study also builds upon the notion that functional differences among rhodopsins are significant and deserving of further study, contrasting the idea that thoroughly characterizing mammalian rhodopsins is sufficient to fully comprehend dim-light vision across vertebrates. We recently highlighted a different retinal release rate in echidna rhodopsin relative to bovine rhodopsin (Bickelmann et al., 2012), suggesting additional variation may exist outside of mammals. This is supported by the fact that other features of rhodopsin also differ across vertebrates. Hydroxylamine stability was a biochemical

160 property first used to distinguish between rhodopsins and cone opsins, with cone opsins experiencing Schiff base hydrolysis fairly rapidly upon exposure (Okano et al., 1989;

Johnson et al., 1993; Starace and Knox, 1998). However, unlike zebrafish rhodopsin, some rhodopsins have elicited a reaction to hydroxylamine, including the anole

(Kawamura and Yokoyama, 1998), and the echidna (Bickelmann et al., 2012).

Meanwhile, the kinetics of rhodopsin following photoactivation, specifically those of the meta-intermediate equilibrium, also has distinct rates. Meta II decay rates in human

(Lewis et al., 1997) and bovine rhodopsin (Yoshizawa and Shichida, 1982) are notably different than those in non-mammalian rhodopsins, such as chicken (Imai et al., 1995), salamander (Das et al., 2004), and cichlid fish (Sugawara et al., 2010), while meta III decay rates are even different among mammalian rhodopsins (Lewis et al., 1997). While these differences might not be of the same magnitude as those between rhodopsins and cone opsins, they reveal intriguing insights into the evolution of dim-light visual systems in vertebrates.

Visual pigment regeneration rates differ greatly between rod and cone opsins, and serve as a key adaptation of rhodopsin to mediate dim-light vision (Imai et al., 2005). Slower regeneration rates seem to have evolved in tandem with other key aspects of rod photoreceptor function, such as reduced thermal isomerization in the dark (Kefalov et al.,

2003; Fu et al., 2008; Luo et al., 2011), and increased stability of the biologically active meta II, which increases photoresponse amplitudes (Imai et al., 2007; Sakurai et al.,

2007). Previous studies have stated that release of all-trans-retinal may be required before 11-cis-retinal can form a new Schiff base linkage in the chromophore binding

161 pocket (Hofmann et al., 1992; Pulvermüller et al., 1997), suggesting natural variation in retinal release rates may contribute in some capacity to rates of visual pigment regeneration in vivo.

Under some circumstances, retinal release may also be connected to dark adaptation, described as the recovery of sensitivity of the visual system following exposure to light

(Hecht et al., 1937). At the molecular level, dark adaptation requires both release of all- trans-retinal from opsin following photoactivation, and regeneration of opsin with a new

11-cis-retinal chromophore. Under larger bleaching conditions, the rate limiting step of visual pigment regeneration is related to the availability of new 11-cis-retinal in rod outer segments [ROS], and has been linked to either retinal isomerase activity in the retinal pigment epithelium [RPE] (Lee et al., 2010), or transport of newly metabolized 11-cis- retinal from the RPE to the ROS (Mahroo and Lamb, 2004; Lamb and Pugh, 2006). For small bleaches, which are likely more relevant in the absence of artificial light, an existing pool of 11-cis-retinal in the ROS can regenerate 3-10% of visual pigments in the retina (Azuma et al., 1997; Cocozza and Ostroy, 1987), bypassing the rate-limiting activities involving the RPE and possibly leaving retinal release as the rate-limiting factor

(Ala-Laurila et al., 2006). This would mean changes in retinal release may be proportional to physiological rates, such as dark adaptation, and suggest that the 2-fold differences measured in this study could be physiologically significant. Additional in vitro measurements of retinal release may be able to further explore details of the connection between retinal release and physiological properties of vision such as dark adaptation and visual pigment regeneration.

162 4.6 Conclusions

Retinal release is a kinetic property that varies both among and within major groups of opsins. While highly conserved residues in rhodopsin influence retinal release by establishing a stable Schiff base link, these residues are invariant and do not contribute to the natural sequence variation among rhodopsins. Our experiments provide evidence that substitutions as variable sites lining several motifs of the retinal channel can mediate retinal release, including TM3 near E122, an opening in meta II between TM5 and 6, and the ‘retinal plug’ formed by EL2. Our most salient result is that the majority of these sites were situated closer to the β-ionone ring of retinal, with sites more proximal to the

Schiff base being more invariant. This suggests that variable sites among rhodopsins are more likely to alter retinal release via steric effects during dissociation, than by influencing the stability of the Schiff base, and is supported by an Arrhenius plot showing an activation energy of Schiff base hydrolysis for zebrafish rhodopsin similar to that of bovine rhodopsin. However, some effect on Schiff base stability cannot be ruled out, especially with rearrangement of the charged E122, or by the effects of V286I and L290I, which are somewhat proximal to the Schiff base. Our results also support the concept of retinal exiting opsin via the TM5/TM6 opening; however, we cannot confirm whether this opening is also used for retinal uptake. It should also be noted that these experiments were performed with visual pigments re-suspended in dodecyl maltoside solutions, not native rod outer segments, and that some molecular components that may influence retinal release rates, such as transducin and arrestin, were not present. When drawing conclusions from kinetic experiments in vitro, it is important to be able to place these results in context in vivo. This study is a perfect example, where zebrafish rhodopsin

163 releases retinal around twice as fast as bovine rhodopsin at uniform temperature, but around twice as slow at respective physiological temperatures. Based on these results, future studies in this field should include opsins from a wide variety of species that can not only address questions about visual pigment structure and function, but also contribute to the broader topic of the vertebrate visual system.

164

Figure 1. Ultraviolet-visual absorbance spectra of [A] bovine and [B] zebrafish rhodopsin. Visual pigments were regenerated with 11-cis-retinal and purified with the

1D4 monoclonal antibody in 0.1% dodecyl maltoside. The λMAX is indicated for each pigment.

165

Figure 2. Increase in fluorescence intensity, representing release of all-trans-retinal from the chromophore binding pocket, in bovine and zebrafish rhodopsin following photoactivation. A sharp increase in fluorescence at 330 nm that eventually plateaus is seen in both bovine [black] and zebrafish [grey] at t = 0, which followed a 30-second bleach with white light. Half-life values of the first-order kinetic reactions were 13.9 and 6.6 min, respectively, at 20oC, pH 7.0, in solution containing 0.1% dodecyl maltoside.

166

Figure 3. Crystal structure homology model of zebrafish rhodopsin [left], highlighting TM5 and 6. Side chain identities of highlighted sites alter retinal release rates, likely through steric effects during retinal dissociation from the depicted opening. Release of all-trans-retinal, as depicted by an increase in fluorescence intensity [right] is shown for both wild type and I209V, F213I, V266L, W273F, producing half-life values of 6.6 and 13.1 min, respectively. Amino acid numbering is relative to bovine rhodopsin.

167

Figure 4. Crystal structure homology model of several motifs lining the retinal channel in zebrafish rhodopsin. [A] The area surrounding E122, highlighting its proximity to retinal. [B] The ‘retinal plug’ formed by EL2, illustrating the proximity of V189 to retinal. Amino acid numbering is relative to bovine rhodopsin.

168

Figure 5. Arrhenius plot of the natural logarithm of the rates of fluorescence increase in o photoactivated zebrafish rhodopsin between 5 and 24 C at pH 7. Activation energy (Ea) was calculated from the negative reciprocal of the slope of a linear regression line that best fit the data (r2 = 0.979). Error bars indicate standard deviation.

169

Figure 6. Increase in fluorescence intensity in bovine and zebrafish rhodopsin following photoactivation at respective physiological temperatures. Results were collected at approximate physiological temperatures [bovine – 37oC; zebrafish – 24oC]. A sharp increase in fluorescence at 330 nm that eventually plateaus is seen in both bovine [black] and zebrafish [grey] at t = 0, which followed a 30-second bleach with white light. Half- life values of the first-order kinetic reactions were 2.0 and 4.1 min, respectively, at pH 7.0, in solution containing 0.1% dodecyl maltoside.

170

Figure S1. Absorbance of zebrafish rhodopsin is not altered by incubation with hydroxylamine. No significant difference in absorbance is detected before exposure to hydroxylamine [black], or after incubating in 50 mM hydroxylamine for two hours [red].

171

Figure S2. Sequence logo representing variation among vertebrate rhodopsins of eight residues shown to mediate retinal release rates. Generated using WebLogo (Crooks et al., 2004) with standard settings. Sequences used were obtained from GenBank (www.ncbi.nlm.nih.gov) and listed in Table S1.

172 Difference Opsin t (min) from wt λ MAX 1/2 (nm) (min)

Bovine Rhodopsin 13.9 +/- 1.6 (19) ---- 499

Zebrafish Rhodopsin 6.6 +/- 0.6 (54) ---- 501 A33E 6.5 +/- 0.1 (3) ---- 500 A36Q 7.0 +/- 0.6 (3) ---- 501 E64Q 7.0 +/- 0.4 (3) ---- 501 M95L 6.9 +/- 0.7 (3) ---- 501 R107P 5.9 +/- 1.1 (3) ---- 501 M123I 11.5 +/- 0.7 (5) +4.9** 501 G124A 8.4 +/- 0.5 (3) +1.8* 501 W136Y 7.1 +/- 0.3 (4) ---- 500 C165L 5.7 +/- 0.4 (3) -0.9* 501 V189I 9.1 +/- 0.9 (4) +2.3* 500 I209V 5.3 +/- 0.7 (3) -1.3* 500 F213I 8.1 +/- 0.6 (9) +1.5** 500 R225Q 5.8 +/- 0.7 (3) --- 502 E241A 6.0 +/- 0.5 (3) --- 500 R248K 7.1 +/-0.8 (4) ---- 500 V266L 8.3 +/- 0.3 (5) +1.7** 499 W273F 8.8 +/- 0.8 (7) +2.2** 499 V286I 5.7 +/- 0.3 (3) -0.9** 501 L290I 5.3 +/- 0.5 (3) -1.3* 500 C304V 6.6 +/- 0.4 (3) ---- 501 C308M 6.9 +/- 0.5 (3) ---- 500 H315N 6.9 +/- 0.6 (3) ---- 501 M123I, G124A 10.6 +/- 1.6 (6) +4.0** 502 F213I, V266L 11.1 +/- 0.4 (3) +4.5** 500 F213I, W273F 10.1 +/- 0.9 (5) +3.5** 499 V266L, W273F 10.0 +/- 1.0 (8) +3.4** 498 F213I, V266L, W273F 13.3 +/- 0.6 (4) +6.7** 498 I209V, F213I, V266L, W273F 13.1 +/- 1.0 (3) +6.5** 497

Table 1: Retinal release rates, λMAX values for wild type and mutant rhodopsins, measured at 20oC. For retinal release rates, standard deviations and number of replicates are listed. Significant differences from wild type zebrafish rhodopsin are listed, based on a two-tailed T-test with unequal variance, where p < 0.05 (*) or < 0.01 (**).

173 Visual Pigment Temp. t1/2 (min) (oC)

20 13.9 +/- 1.6 (19) Bovine Rhodopsin 37 2.0 +/- 0.2 (7) 5 35.7 +/- 2.6 (2) 18 10.5 +/- 1.6 (4) Zebrafish Rhodopsin 20 6.6 +/- 0.6 (54) 24 4.1 +/- 0.6 (4)

Table 2: Retinal release rates of wild type bovine and zebrafish rhodopsin at various temperatures. Standard deviations and number of replicates are listed.

174 Scientific Name Accession Number Plecoglossus altivelis AB074484 Chlamydera maculata JQ034385 Emberiza bruniceps JQ695942 Ailuroedus crassirostris JQ034381 Pseudobagrus tokiensis FJ197075 Gallus gallus NM_001030606 Bos Taurus NM_001014890 Corvus macrorhynchos AB555651 Tursiops truncates AF055456 Sminthopsis crassicaudata AY159786 Tachyglossus aculeatus JX103830 Anguilla Anguilla L78008 Anguilla Anguilla L78007 Xenopus tropicalis BC135234 Lepisosteus osseus AF137207 Carassius auratus L11863 Poecilia reticulate Y11147 Homo sapiens U49742 Lethenteron camtschaticum AB116382 Uta stansburiana DQ100323 Lefua costata EU409634 Macaca fascicularis S76579 Anas platyrhynchos AF021240 Trichechus manatus AF055319 Oryzias latipes AB180742 Chanos chanos FJ197072 Mus musculus BC031766 Cynops pyrrhogaster AB043890 Polyodon spathula AF369050 Scopelarchus analis EF517404 Esox Lucius AY158044 Takifugu rubripes AF201471 Python regius FJ497236 Oryctolagus cuniculus NM_001082349 Ambystoma tigrinum U36574 Salmo salar NM_001123537 Mirounga angustirostris AY228452 Xenopeltis unicolor FJ497233 Acipenser sp. AF137206 Astyanax mexicanus U12328 Bufo bufo U59921 Eubalaena glacials JQ730751 Taeniopygia guttata AF222329 Danio rerio AB087811

Table S1: List of rhodopsin sequences used to generate the sequence logo in Figure S1, along with respective GenBank accession numbers.

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184

Chapter 5:

A functional dichotomy of rod-like cone opsin

genes in teleost fish linked to a single amino acid

at site 122

185 5.1 Abstract

In order to achieve a dynamic range spanning several orders of magnitude of light intensities, vertebrates have two distinct classes of retinal photoreceptor cells: rods, responsible for dim-light and peripheral vision, and cones, responsible for bright light and

colour vision. Molecular properties such as λMAX, chromophore accessibility, and retinal release are all known to differ between rhodopsins and cone opsins, and key residues, such as site 122, have been identified that can induce shifts between rhodopsin and cone opsin phenotypes when mutated. Rod-like cone opsins (rh2), the class of cone opsins with the most sequence similarity to rod opsins, experienced several duplication events following the adaptive radiation of teleost fish over 150 million years ago. As a result, a significant number of teleosts containing at least two rod-like cone opsin (rh2) genes have one copy with Q122 and one with E122. We propose that a functional dichotomy exists among rh2 opsin genes in teleosts, contingent on the identity of site 122, and that this dichotomy may be involved in the maintenance of gene duplicates through functional diversification. We are investigating the implications of substitutions at this residue in zebrafish rh2 genes by characterizing several aspects of opsin function in both wild type and mutant pigments, following expression in mammalian cell culture. Preliminary results support the existence of two functional classes of rh2 genes in teleost fish based on the identity of site 122. This is the first study of molecular function study to

investigate aspects of function other than λMAX that may be linked to the expansion of the rh2 gene family in teleosts, and should help to elucidate some of the reasons for the functional differences within rh2 opsins.

186 5.2 Introduction

Visual pigments are members of the G protein-coupled receptor [GPCR] superfamily and initiate the critical first step in the biochemical cascade of vision (Stryer, 1986).

Vertebrate visual pigment genes resolve into five distinct phylogenetic classes; four comprise cone opsins [lws/mws, sws1, sws2 and rh2] while one comprises rhodopsin

[rh1] (Yokoyama, 2000). These five opsin classes have arisen from a single ancestral opsin gene through a series of gene duplication events, with some classes having duplicated further in certain groups of vertebrates (Bowmaker and Hunt, 2006). While many features of visual pigments have been conserved throughout evolution, they have also evolved to perform specialized visual functions. Cone opsins are responsible for photopic [bright light] and colour vision and are often characterized based on their

wavelength of maximal absorption, λMAX (Menon et al., 2001). The expansion of cone opsin groups through gene duplication and retention may be due to the evolution of

variation in molecular properties, such as λMAX.

The rod-like cone opsin [rh2] gene family expanded during the evolution of the

Actinopterigian lineage, with many species of teleost fish having upwards of four or five rh2 genes (Chinen et al., 2003; Nakamura et al., 2013). The expansion of this gene family followed by evolutionary changes to their amino acid sequences might reflect an adaptation to the diversity of aquatic light environments, or may confer a set of genes that can be expressed at various developmental stages (Archer et al., 1995; Carleton and

Kocher, 2001). However, while recent phylogenetic and molecular evolutionary analyses have helped to track the duplication and divergence events of opsin genes in ray-finned

187 fish (Rennison et al., 2012), there is still a dearth of studies focused on characterizing aspects of cone opsin function that may have contributed to the retention of gene duplicates.

Previous investigations of visual pigment function have implicated site 122 as an integral determinant of function in rhodopsins and cone opsins. Site 122, located in transmembrane helix 3, is conserved in most rhodopsins as glutamate [E122] and in most rh2 opsins as glutamine [Q122]. However, teleost fish that possess multiple rh2 opsins often have a combination of Q122- and E122-containing variants. In zebrafish, the identity of this site has been identified as a key determinant of spectral tuning during rh2 opsin evolution (Chinen et al., 2005). Site-directed mutagenesis studies that expressed chicken rh2 Q122E reduced meta II decay and retinal regeneration rates relative to wild type. Similarly, the E122Q mutant of chicken rhodopsin showed an increase in these kinetic rates relative to wild type (Imai et al., 1997). Site 122 has also been examined in vivo, where transgenic mice expressing E122Q rhodopsin showed lower photosensitivity and lower meta II lifetime than wild type mice (Imai et al., 2007). These studies suggest that a substitution at site 122 can lead to significant alterations to molecular properties of visual pigment function, as well as corresponding physiological properties in vivo.

Here, we investigate the functional discrepancies that exist between zebrafish rh2 opsins containing either Q122 or E122. Additionally, we look at the evolutionary history of teleost rh2 opsins to determine whether functional differences between Q122 and E122 identities may have supported expansion of the rh2 gene family. We expressed both

188 zebrafish rh2-1 [Q122] and rh2-4 [E122], confirming the 39 nm difference revealed in previous studies (Chinen et al., 2003). The Q122E mutation in rh2-1 caused a red-shift

of 19 nm, suggesting that site 122 contributes to the differences of λMAX values found in teleost rh2 opsins. Both zebrafish rh2-1 and rh2-4 were sensitive to hydroxylamine, unlike bovine rhodopsin, although rh2-4 reacted slightly faster, which is an interesting result considering rh2-4 contains the E122, commonly found in rhodopsins. The influence of site 122 on retinal release was also examined in a number of different ways.

First, the E122Q substitution in bovine rhodopsin increased the rate of retinal release 5- fold. However, this difference was much smaller than the 240-fold difference seen between zebrafish rhodopsin and zebrafish rh2-4, both of which contain E122. This suggests that, while the identity of site 122 does contribute to establishing retinal release rates, it alone has a less significant effect than other key differences between rhodopsins and cone opsins. Finally, a phylogenetic analysis of teleost rh2 opsin sequences showed several major groups of teleost fish [ostariophysians, acanthomorphs, etc.] have large clades of rh2 opsins with mostly either Q122 or E122, suggesting the possibility that functional differentiation caused by substitutions at site 122 may have led to a selective advantage to conserve both variants early in the evolutionary history of these major groups of fish. The implications of these findings on the function and evolution of teleost rh2 opsins, as well as potential further directions for this research, will be discussed.

189 5.3 Materials and methods

5.3.1 Visual pigment expression and purification

RNA was extracted from adult zebrafish eyes using the TRIzol reagent (Invitrogen), and cDNA libraries were generated using the SMART cDNA Library Construction Kit (BD

Biosciences). The complete coding sequence of zebrafish rh2-1 and rh2-4 (GenBank:

AB087805 and AB087808) were amplified using PfuTurbo (Stratagene) and inserted into the pJET1.2 cloning vector (Fermentas). Site-directed mutagenesis primers were designed to induce a single amino acid substitution [Q122E] in rh2-1. The complete coding sequence of bovine rhodopsin was cloned into the pIRES-hrGFP II expression vector (Stratagene), while wild type and mutant zebrafish visual pigments were cloned into the p1D4-hrGFP II expression vector (Morrow and Chang, 2010). These constructs were used to transiently transfect cultured HEK293T cells using Lipofectamine 2000

(Invitrogen; 8 ug of DNA per 10-cm plate). Cells were harvested 48 h post-transfection and opsins were regenerated using 11-cis-retinal, generously provided by Dr. Rosalie

Crouch (Medical University of South Carolina). Visual pigments were solubilized in 1%

N-dodecyl--D-maltoside [DM] and immunoaffinity purified with the 1D4 monoclonal antibody (Molday and MacKenzie, 1983), as previously described (Morrow and Chang,

2010; Morrow et al., 2011). Purified visual pigment samples were eluted in sodium phosphate buffer [50 mM NaPhos, 0.1% DM, pH 7]. Buffers used to regenerate and purify some samples of zebrafish rh2-4 that were used in retinal release assays also included 25% glycerol.

190 5.3.2 Spectroscopy

The ultraviolet-visible absorption spectra of purified wild type and mutant visual pigments were recorded at 25oC using the Cary4000 double-beam spectrophotometer

(Varian) and quartz absorption cuvettes (Helma). Samples were incubated and assayed in

sodium phosphate buffer [50 mM NaPhos, 0.1% DM, pH 7]. All λMAX values were calculated after fitting absorbance spectra to a standard template for A1 visual pigments

(Govardovskii et al., 2000). All visual pigments activated in response to light, shifting

their λMAX to ~380 nm, representing to active meta II state. Reactivity to hydroxylamine

in the dark was determined by incubating a visual pigment in 50 mM NH2OH (Sigma-

o Aldrich) at 25 C, then monitoring absorbance at the λMAX; absorption spectra were taken

every 3-15 minutes for 90 minutes to measure the rate at which λMAX decreased following

-bx incubation. Collected data was fit to a first order exponential decay curve [y = yo +ae ],

with half-life values being calculated based on rate constant ‘b’ [t1/2 = ln2/b]. All curve fitting resulted in r2 values of greater than 0.95. Bovine rhodopsin was used as a positive control, as it has previously been shown to have no significant reaction upon hydroxylamine incubation (Okano et al., 1989; Kawamura and Yokoyama, 1998).

The protocol used to determine retinal release rates of visual pigments was modified from that of Farrens and Khorana (1995). Briefly, 0.1-0.2 μM visual pigment samples in submicro fluorometer cell cuvettes (Varian) were bleached for 30 seconds using a Fiber-

Lite MI-152 Illuminator external light source (Dolan-Jenner), using a filter used to restrict wavelengths of light below 475 nm. Fluorescence measurements were integrated for 2 seconds at 30-second intervals using a CaryEclipse fluorescence spectrophotometer

191 (Varian). The excitation wavelength was 295 nm [1.5 nm slit width] and the emission wavelength was 330 nm [10 nm slit width]; no noticeable pigment bleaching by the excitation beam was detected. Retinal release was demonstrated through a sharp initial rise in intrinsic tryptophan fluorescence, representing a decrease in fluorescent quenching of W265 by the retinal chromophore. Data from the initial rise was fit to a three variable,

-bx first order exponential equation [y = yo + a(1-e )], with half-life values calculated based

2 on the rate constant 'b' [t1/2 = ln2/b]. All curve fitting resulted in r values of greater than

0.95. Data was recorded at -5oC, without glycerol to draw an initial comparison between zebrafish rhodopsin and rh2-4, and also collected for rh2-4 at -6.5oC in 25% glycerol.

5.3.3 Phylogenetic Analyses

All visual pigment nucleotide sequences in this study were retrieved from NCBI

(http://www.ncbi.nlm.nih.gov/) and aligned using ClustalW (Larkin et al., 2007). Several vertebrate rh1 sequences were added to the rh2 dataset to form an outgroup in order to properly root the tree. This alignment was subjected to a phylogenetic analysis using maximum likelihood methods (Guindon and Gascuel, 2003), employed by MEGA version 5 (Tamura et al., 2011). Bootstrapping methods were used to assess the degree of confidence in the nodes of the phylogeny (Felsenstein, 1985).

5.4 Results

5.4.1 Site 122 helps to establish the λMAX of zebrafish rh2 opsins

Previous mutagenesis experiments that altered the identity of site 122 between glutamate

[E] and glutamine [Q] caused significant shifts in λMAX in both chicken visual pigments

192 (Imai et al., 1995) and zebrafish (Chinen et al., 2005), is likely due to the proximity of site 122 to 11-cis-retinal in the chromophore binding pocket (Palczewski et al., 2000; Fig.

1). The most prevalent phenotypic difference among expressed rh2 opsins in zebrafish is

their λMAX values, with the most red-shifted opsin, rh2-4, also being the only opsin with

E122. Therefore, we predicted that the identity of site 122 is likely contributing to these

differences in λMAX. To investigate this hypothesis, zebrafish rh2-1 and rh2-4 were both expressed using a mammalian cell culture, along with the Q122E mutant in rh2-1. All three visual pigments were regenerated with 11-cis-retinal, producing characteristic dark

spectra (Fig. 2). Zebrafish rh2-1 and rh2-4 had λMAX values of 467 and 506 nm, respectively, similar to previously published results (Chinen et al., 2003). The Q122E

mutant of rh2-1 produced a λMAX of 487 nm, which is a 19 nm red-shift from the wild type.

5.4.2 Zebrafish rh2-1 and rh2-4 are both susceptible to hydroxylamine

Hydroxylamine stability is a biochemical property traditionally used to distinguish between traditional rhodopsin and cone opsin function. When exposed to the retinal chromophore, hydroxylamine is able to disrupt the Schiff base linkage with the opsin apoprotein and sequester the retinal in solution by forming a retinal oxime that absorbs at

363 nm. This process occurs swiftly for cone opsins, whereas rhodopsins usually do not experience a reaction (Okano et al., 1989; Johnson et al., 1993; Starace and Knox, 1998), likely because the chromophore binding pocket of rhodopsin is packed much more tightly than that of cone opsins, preventing the entry of small molecules, such as hydroxylamine.

This suggests that measuring the rate of hydroxylamine reactivity could be a means to

193 characterize the environment of the chromophore binding pocket between cone opsins, therefore, we measured the rate at which both rh2-1 and rh2-4 react upon exposure to

hydroxylamine, as determined by a reduction in λMAX absorbance due to retinal oxime formation (Fig. 3). While zebrafish rh2-1 reacted with a half-life of 21.5 min, rh2-4 reacted more quickly, with a half-life of 13.4 min. While rh2-4 has E122, the most common identity in rhodopsin, it reacted more quickly than rh2-1, which has the more prominent cone opsin identity, Q122. However, it is unlikely that a single amino acid identity, even for a functionally important site such as 122, could be a significant determinant for the accessibility of the chromophore binding pocket, which consists of dozens of residues. Bovine rhodopsin was also assayed as a control, and showed no significant reaction.

5.4.3 Site 122 can mediate retinal release in bovine rhodopsin

Site 122 has previously been identified as a critical functional determinant of meta II decay rates between rhodopsins and cone opsins (Imai et al., 1997). Since the stability of meta II, the biologically active intermediate, should relate to its retinal release rate, we investigated the influence of site 122 mutations on this kinetic property. We first performed mutagenesis on bovine rhodopsin in order to assess the affect of site 122 on a well-studied visual pigment that can be readily expressed. When wild-type bovine rhodopsin was light-bleached and monitored for release of all-trans-retinal, it had a half- life of 13.9 min, similar to previously recorded values of 12.5-15.5 min (Farrens and

Khorana, 1995; Yan et al., 2002). However, the E122Q mutation led to a shorter half-life of 3.0 min, a significant shift towards a more cone opsin-like phenotype (Fig. 4). Based

194 on these preliminary results, it is likely that the identity of site 122 may also be responsible for natural variation in retinal release rates in rh2 opsins.

5.4.4 Zebrafish rh2-4 releases retinal at least 240 times faster than rhodopsin

Despite the fact that the E122Q mutation in bovine rhodopsin causing approximately a 5- fold increase in the rate of retinal release, the magnitude of difference between kinetic rates in rhodopsins and cone opsins is usually significantly greater (Kuwayama et al.,

2002, 2005; Chen et al., 2012). We monitored retinal release in both zebrafish rhodopsin and rh2-4, both of which have E122, to quantify the difference between the background sequences of a rhodopsin and a cone opsin. Even when incubated at -5oC, the half-life of retinal release of rh2-4 was barely within the threshold of measurement of our fluorescence spectrophotometer at 0.57 min. Meanwhile, zebrafish rhodopsin had a half- life of 138.6 min at the same temperature, more than 240 times slower than rh2-4 (Fig.

5a). In order to distinguish differences among rh2 opsins, we had to further slow the retinal release reaction, by reducing the incubation temperature to -6.5oC, and adding

25% glycerol to buffers used to harvest and purify zebrafish rh2-4. These steps increased the half-life of retinal release for zebrafish rh2-4 to 5.17 min, slow enough to perform comparative analyses with other rh2 opsins in future experiments (Fig. 5b).

5.4.5 Phylogenetic distribution of teleost rh2 opsins

A maximum likelihood analysis was performed in order to investigate the evolutionary distribution of rh2 opsins in teleost fish, with particular focus being placed on the identity of site 122. This analysis also included several non-teleost rh2 opsins, all with Q122, as

195 well as several teleost rh1 opsins, all with E122, as outgroups. The resulting phylogeny recovers many of the commonly accepted systematic relationships among ray-finned fish

(Forey et al., 1996; Hurley et al., 2007; Wang et al., 2007). Additionally, it presents an interesting distribution of rh2 opsins, as major groups of teleost fish have large clades of opsins with mostly either Q122 or E122 (Fig. 6). This is most notable in ostariophysians

[zebrafish, goldfish, etc.] and acanthomorphs [guppy, tuna, cichlids, etc.], and suggests that functional differentiation caused by substitutions at site 122 may have led to a selective advantage to conserve both variants early in the evolutionary history of each of these major groups of fish independently. Additional rh2 opsins sequences and molecular evolutionary analyses would be required to further investigate this hypothesis.

5.5 Discussion

In this study, we have started to functionally characterize several rh2 opsins from zebrafish, using heterologous opsin expression, site-directed mutagenesis, and both absorbance and fluorescence spectroscopy, with the purpose of determining the

contribution of site 122 to various aspects of rh2 opsin function. We measured the λMAX values of zebrafish rh2-1 and rh2-4, which contain Q122 and E122 respectively, to be 39 nm apart, similar to previously published results (Chinen et al., 2003). However, the

Q122E mutant of rh2-1 caused a red-shift of 19 nm, confirming the results of previous mutagenesis at this site in zebrafish rh2 opsins (Chinen et al., 2005), and suggesting that site 122 could be responsible for a significant portion of this natural variation. While both rh2-1 and rh2-4 were sensitive to hydroxylamine, rh2-4 reacted more quickly, suggesting the possibility of differences in the accessibility of the chromophore binding

196 pocket between the two opsins. Finally, we also measured the retinal release rate of zebrafish rh2-4 to be over 240 times faster than that of zebrafish rhodopsin, while the retinal release rate of an E122Q mutant in bovine rhodopsin was only 5 times faster than wild type. This suggests that the background sequence of a visual pigment likely plays a more significant role in establishing retinal release rates than the identity of site 122 alone. The implications of these findings to the molecular evolution of rh2 genes in

Actinopterigian fish will be discussed, as will potential future directions for this project.

In recent years, additional sequence information has revealed expansions of the rh2 opsin family in several lineages of Actinopterygian fish, with some species having up to five duplicates (Nakamura et al., 2013). Not only was this an increase in rh2 opsin genes relative to other vertebrates, but while Q122 is invariant in other groups of vertebrates, many of these genes were also found to have E122. Because of prior knowledge of the influence of site 122 on opsin function, this newly discovered class of rh2 opsins with

E122 is ideal for studying functional differentiation among cone opsins. The most characteristic functional property of visual pigments is their wavelength of maximal

absorption, λMAX, with a specific range of λMAX values existing for each group of cone opsins (Yokoyama, 2000). The rh2 opsins usually absorb blue and green wavelengths of

light maximally, with teleost fish have been shown to have a wide variation of λMAX values among their duplicated rh2 opsins. For example, the medaka [O. latipes] has three

rh2 genes whose λMAX values differ by up to 64 nm (Matsumoto et al., 2006). While

zebrafish has an additional rh2 gene compared to medaka, the difference in λMAX among its pigments is a more modest 39 nm (Chinen et al., 2003). A commonality between the

197 opsin sequences in these two species and other ray-finned fish is that their most blue- shifted rh2 opsin gene has Q122, while the most red-shifted has E122. We expressed zebrafish rh2-1 and rh2-4, the most blue- and red-shifted rh2 genes respectively, and

reproduced the 39 nm different in λMAX between the two pigments. However, the rh2-1

Q122E mutant had a λMAX value of 487 nm, a 19 nm red-shift compared to wild type.

This suggests that site 122 is largely responsible for the wide range of λMAX values seen in the rh2 opsins of teleost fish. Future experiments should perform the reverse substitution,

E122Q, in rh2-4 to see of the result mirrors that of rh2-1 Q122E. Additionally, the effect

of a Q122E substitution on zebrafish rh2-2 and rh2-3, which have more red-shifted λMAX values than rh2-1, should be explored.

While the developmental expression pattern and gene regulation of rh2 opsins have been well characterized in zebrafish (Takechi and Kawamura, 2005; Tsujimura et al., 2007), most previous experiment that have characterized functional aspects of rh2 opsins were performed on chicken green, an rh2 opsin with Q122. When chicken green was first studied, it was called a rhodopsin-like visual pigment (Yokoyama, 1994), and were later shown to have a molecular extinction coefficient and photosensitivity similar to rhodopsin, along with a higher amino acid sequence similarity that other cone opsin groups (Shichida et al., 1994). However, further analyses of the kinetics of chicken green revealed faster regeneration rates when incubated with 11-cis-retinal and faster meta II formation and decay kinetics compared to rhodopsin (Shichida et al., 1994), which correlates with the physiological differences between cone and rod photoreceptor cells

(Schnapf and Baylor, 1987). Mutagenesis experiments then identified that the faster

198 kinetic properties in chicken green could largely be attributed to the amino acid identities of two key sites, Q122 and P189, where mutations to respective rhodopsin identities,

Q122E and P189I respectively, produced pigments with slower, more rhodopsin-like kinetics (Imai et al., 1997; Kuwayama et al., 2002; Kuwayama et al., 2005). Our results confirm that some of the differences measured between wild type chicken rhodopsin and chicken green are also found among zebrafish opsins, where zebrafish rh2-4 releases retinal over 240 times faster than rhodopsin. Additionally, the fact that the half-life values of zebrafish rh2-1 and rh2-4 reactivity when incubated with hydroxylamine differ by over 8 minutes, compared to a difference of just over 2 minutes between two E122 pigments in goldfish (Johnson et al., 1993), suggests that the functional differences

among rh2 opsins due to the identity of site 122 go beyond shifts in λMAX. However, the specific effect of mutations at site 122 on the kinetic properties of zebrafish rh2 opsins still needs to be investigated.

With Actinopterygian fish being the only group of vertebrates to experience considerable expansion of the rh2 gene family, it stands to reason that there is likely differentiation among these genes that provides novel utility to the host organism, which could lead to gene retention (Lynch and Conery, 2000). Based on our results, and previous experiments implicating site 122 as a key determinant of kinetic properties of visual pigment function (Imai et al., 1997; Kuwayama et al., 2005), we decided to examine the phylogenetic distribution of both Q122 and E122 rh2 opsins in teleost fish. Maximum likelihood methods were able to recover a phylogeny with well-supported clades for several major groups of teleost fish, including ostariophysians [e.g. zebrafish, goldfish]

199 and acanthomorphs [e.g. guppy, medaka]. The most basal Actinopterigian rh2 sequence included in our analysis was from the eel, A. anguilla, which has E122. Considering the large majority of vertebrate rh1 sequences have E122, this suggests that the common ancestor of all rh1 and rh2 genes may have also had E122. What is even more intriguing is how subgroups of predominantly Q122- and E122-containing rh2 genes seem to form within clades representing the two largest groups of teleosts: ostariophysians and acanthomorphs. This may imply that independent rh2 duplications occurred early in the evolution of both of these groups, and that these duplicate genes may have been retained partly due to the functional differentiation provided by an E122Q substitution in both cases. A larger dataset and additional characterization of different rh2 opsins would be required to further test this hypothesis.

While the reason for rh2 expansion and retention in many species of teleost fish has yet to be ascertained, this study has started to elucidate some of the aspects of opsin function

that may differ among this group of visual pigments, such as λMAX, hydroxylamine sensitivity, and retinal release. In addition to previous studies examining rh2 opsin evolution (Chinen et al., 2005) and those that explore regulatory and developmental differences in model systems, such as zebrafish (Takechi and Kawamura, 2005;

Tsujimura et al., 2007), a clearer image is now being crafted of the potential benefits of retaining multiple rh2 opsin genes. Additional insights into functional aspects of rh2 opsins, as well as molecular evolutionary analyses of rh2 opsins sequences could help to further elucidate how this gene family impacts the visual system of Actinopterygian fish.

200

Figure 1. Crystal structure of the chromophore binding pocket of dark state bovine rhodopsin. The side chain from site 122 [yellow] from transmembrane helix 3 [blue] is aimed toward the 11-cis-retinal chromophore [white] (Palczewski et al., 2000).

201

Figure 2. Ultraviolet-visual absorbance spectra of zebrafish rh2 opsins. The λMAX values of zebrafish rh2-1 [green], rh2-4 [red], and rh2-1 Q122E are 467, 506, and 487 nm respectively. All visual pigments activated in response to light, which shifted their λMAX to approximately 380 nm (data not shown).

202

Figure 3. Decrease in λMAX absorbance in bovine and zebrafish visual pigments following incubation in hydroxylamine at 25oC. A decrease in absorbance represents the accessibility of hydroxylamine to the chromophore binding pocket, and occurred with a half-life of 21.5 and 13.4 min for zebrafish rh2-1 and rh2-4, respectively. Bovine rhodopsin was used as a control to demonstrate that rhodopsin does not react to hydroxylamine.

203

Figure 4. Increase in fluorescence intensity representing release of all-trans-retinal from bovine rhodopsin following photoactivation. A sharp increase in fluorescence at 330 nm that eventually plateaus is seen in both bovine wild type [black] and E122Q [grey] at t = 0, which followed a 30-second bleach with white light. Half-life values of the first-order kinetic reactions were 13.9 and 3.0 min, respectively, at 20oC, pH 7.0, in solution containing 0.1% dodecyl maltoside.

204

Figure 5. Increase in fluorescence intensity representing release of all-trans-retinal from zebrafish rhodopsin and rh2-4 following photoactivation. An increase in fluorescence at 330 nm following illumination. (A) Zebrafish rhodopsin and rh2-4 produced half-life values of 138.6 and 0.57 min, respectively, at -5oC. (B) Zebrafish rh2-4 produced a half- life value of 5.17 min at -6.5oC in 25% glycerol. All experiments were run at pH 7.0 in 0.1% dodecyl maltoside.

205

Figure 6. Phylogeny of teleost rh2 nucleotide sequences generated using maximum likelihood [ML] methods. Numbers above nodes indicate ML bootstraps [100 replications]. Branches are proportional to ML estimated branch lengths under GTR model. Sequences were obtained from GenBank, with accession numbers listed next to sequence names.

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207 Imai H, Kojima D, Oura T, Tachibanaki S, Terakita A, Shichida Y (1997) Single amino acid residue as a functional determinant of rod and cone visual pigments. Proc Natl Acad Sci USA 94, 2322-2326 Imai H, Kefalov V, Sakurai K, Chisaka O, Ueda Y, Onishi A, Morizumi T, Fu Y, Ichikawa K, Nakatani K, Honda Y, Chen J, Yau KW, Shichida Y (2007) Molecular properties of rhodopsin and rod function. J Biol Chem 282, 6677-6684 Johnson RL, Grant KB, Zankel TC, Boehm MF, Merbs SL, Nathans J, Nakanishi K (1993) Cloning and expression of goldfish opsin sequences. Biochemistry 32, 208- 214 Kawamura S, Yokoyam S (1998) Functional characterization of visual and nonvisual pigments of American chameleon (Anolis carolinensis). Vision Res 38, 37-44 Kuwayama S, Imai H, Hirano T, Terakita A, Shichida Y (2002) Conserved proline residue at position 189 in cone visual pigments as a determinant of molecular properties different from rhodopsins. Biochemistry 41, 15245-15252 Kuwayama S, Imai H, Morizumi T, Shichida Y (2005) Amino acid residues responsible for the meta-III decay rates in rod and cone visual pigments. Biochemistry 44, 2208- 2215 Larkin MA, Blackshields G, Brown NP, Chenna R, McGettigan PA, McWilliam H, Valentin F, Wallace IM, Wilm A, Lopez R, Thompson JD, Gibson TJ, Higgins DG (2007) Clustal W and Clustal X version 2.0. Bioinformatics 23, 2947-2948 Lynch M, Conery JS (2000) The evolutionary fate and consequences of duplicate genes. Science 290, 1151-1155 Matsumoto Y, Fukamachi S, Mitani H, Kawamura S (2006) Functional characterization of visual opsin repertoire in Medaka (Oryzias latipes). Gene 371, 268-278 Menon ST, Han M, Sakmar TP (2001) Rhodopsin: structural basis of molecular physiology. Physiol Rev 81, 1659-1688 Molday RS, MacKenzie D (1983) Monoclonal antibodies to rhodopsin: characterization, cross-reactivity, and application as structural probes. Biochemistry 22, 653-660 Morrow JM, Chang BS (2010) The p1D4-hrGFP II expression vector: a tool for expressing and purifying visual pigments and other G protein-coupled receptors. Plasmid 64, 162-169

208 Morrow JM, Lazic S, Chang BS (2011) A novel rhodopsin-like gene expressed in zebrafish retina. Vis Neurosci 28, 325-335 Nakamura Y, Mori K, Saitoh K, Oshima K, Mekuchi M, Sugaya T, Shigenobu Y, Ojima N, Muta S, Fujiwara A, Yasuike M, Oohara I, Hirakawa H, Chowdhury VS, Kobayashi T, Nakajima K, Sano M, Wada T, Tashiro K, Ikeo K, Hattori M, Kuhara S, Gojobori T, Inouye K (2013) Evolutionary changes of multiple visual pigment genes in the complete genome of Pacific bluefin tuna. Proc Natl Acad Sci USA 110, 11061-11066 Okano T, Fukada Y, Artamonov ID, Yoshizawa T (1989) Purification of cone visual pigments from chicken retina. Biochemistry 28, 8848-8856 Palczewski K, Kumasaka T, Hori T, Behnke CA, Motoshima H, Fox BA, Le Trong I, Teller DC, Okada T, Stenkamp RE, Yamamoto M, Miyano M (2000) Crystal structure of rhodopsin: A G protein-coupled receptor. Science 289, 739-745 Rennison DJ, Owens GL, Taylor JS (2012) Opsin gene duplication and divergence in ray-finned fish. Mol Phylogenet Evol 62, 986-1008 Schnapf JL, Baylor DA (1987) How photoreceptor cells respond to light. Sci Am 256, 40-47 Shichida Y, Imai H, Imamoto Y, Fukada Y, Yoshizawa T (1994) Is chicken green- sensitive cone visual pigment a rhodopsin-like pigment? A comparative study of the molecular properties between chicken green and rhodopsin. Biochemistry 33, 9040- 9044 Starace DM, Knox BE (1998) Cloning and expression of a Xenopus short wavelength cone pigment. Exp Eye Res 67, 209-220 Stryer L (1986) Cyclic GMP cascade of vision. Annu Rev Neurosci 9, 87-119 Takechi M, Kawamura S (2005) Temporal and spatial changes in the expression pattern of multiple red and green subtype opsin genes during zebrafish development. J Exp Biol 208, 1337-1345 Tamura K, Peterson D, Peterson N, Stecher G, Nei M, Kumar S (2011) MEGA5: molecular evolutionary genetic analysis using maximum likelihood, evolutionary distance, and maximum parsimony methods. Mol Biol Evol 28, 2731-2739

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210

Chapter 6:

General Discussion

211 6.1 GPCR Dimerization

While performing analyses of rhodopsin in vitro has revealed a wealth of knowledge on the structure and function of both rhodopsin and GPCRs in general, there will always be concerns about whether an in vitro experimental system can appropriately relate information when on a single or handful of molecules are isolated from their natural, in vivo milieu. For example, one concern that is specific to visual pigments is how much the molecular properties of an opsin contribute to the cellular properties of the photoreceptor (Kefalov et al., 2003). These types of questions have led to further investigation of the natural state of rhodopsin and how it interacts with other molecules in vivo, studies that are often mirrored in other GPCRs. One aspect that is of particular interest to our research group is rhodopsin dimerization, because of our hypotheses of the functional role of zebrafish rh1-2.

Over the last decade, one key aspect of rhodopsin structure that has been revealed is how this visual pigment exists in higher order oligomeric states in their native rod outer segment membranes (Fotiadis et al., 2003). This consists both of intradimeric contacts that likely involve TM4 and 5, as well as contacts between TM1, TM2, and cytoplasmic loop 3 that facilitate the formation of rhodopsin dimer rows (Liang et al., 2003).

Rhodopsin is not the sole GPCR that exists as a dimer, as many other GPCRs that dimerize have been studied and evolutionarily conserved residues along TM4-6 exist in various GPCR families that could likely serve as dimerization surfaces (Soyer et al.,

2003). The identification of these potential dimerization surfaces coincides with evidence from GPCR crystal structures (Palczewski et al., 2000; Rasmussen et al., 2007;

212 Wu et al., 2010) as well as over 10 years of functional studies (Milligan, 2009). The two main aspects of dimerization that are often investigated in these functional analyses are:

1) conformational changes occurring to a monomer upon GPCR dimerization and/or ligand binding, and 2) interactions of GPCR dimers with G proteins and other GPCR- interacting proteins. It should be noted that the extent and effect of dimerization is still viewed as a contentious topic, with some studies questioning the ubiquity and stability of

GPCR dimers (Bouvier et al., 2007; Lambert, 2010).

There are two main ways that monomers have been shown to alter conformational properties of their complementary subunit. The first often occurs with homodimers, where a GPCR monomer that binds a certain ligand inherits the ability to influence the binding capacity of another subunit. For example, while a dopamine D2 receptor binding its antagonist negatively affects the ability for the monomer to interact with a G protein, it also allows for a dimer with only one antagonist-bound subunit be fully active (Han et al., 2009). There are also instances where having access to a variety of different ligands to bind will generate dimers that activate their G protein more efficiently, such as the oxytocin receptor (Albizu et al., 2010). Unlike homodimers, whose subunits can assume differential properties based on their bound ligand, heterodimers often have similar

distinctions built into their primary sequences. The gamma-aminobutyric acid [GABA]B receptor is an obligatory heterodimer, where GB1 is responsible for binding ligand and

GB2 can bind G protein, but neither subunit can perform its function without existing as a dimer (Galvez et al., 2001). While only homodimerization has been detected and investigate for rhodopsin, the introduction of a second visual rhodopsin gene in zebrafish,

213 rh1-2, that may be expressed alongside rhodopsin, may offer the opportunity for heterodimerization in rod photoreceptor outer segments.

With GPCR dimerization in vivo being an established concept, another group of studies began to explore how this dimer interacts with its G protein and other components of its signal transduction cascade. While some studies suggest that each GPCR monomer is paired with its own G protein (Bayburt et al., 2007; Whorton et al., 2007), others support the idea of a 2:1 stoichiometry, with two receptors binding a single heterotrimeric G protein (Hamm, 2001; Baneres and Perello, 2003). There are two distinct G protein

regions that interact with the receptor: the primary docking site of the Gα subunit that penetrates the cytoplasmic domain of the activated receptor, and a site in the C-terminal

region of the Gγ subunit (Oldham and Hamm, 2006; Hofmann et al., 2009). In rhodopsin, the distance between these two regions in the G protein transducin is 55 Å, larger than the 45 Å width of a rhodopsin monomer, supporting the idea that a rhodopsin dimer interacts with a single transducin and inspiring most modeling studies to employ the 2:1 stoichiometry (Liang et al., 2003; Johnston and Siderovski, 2007). This asymmetric model of rhodopsin dimers could also apply to other phototransduction proteins, such as the regulator of G protein signaling [RGS] (Maurice et al., 2010), rhodopsin kinase (Huang and Tesmer, 2011), and arrestin when there is a high density of photoactivated pigments (Sommer et al., 2011). This model suggests that there could be functional differences between the two rhodopsin subunits in a dimer if they are interacting with different regions of transducin and other phototransduction proteins,

214 which may support the concept of heterodimerization between rhodopsin and rh1-2 in zebrafish.

6.2 Influence of retinal release on in vivo function

2003; Fu et al., 2008; Luo et al., 2011), and increased stability of the biologically active meta II intermediate [R*], which increases photoresponse amplitudes (Imai et al., 2007;

Sakurai et al., 2007). Previous studies have suggested that release of all-trans-retinal is required before 11-cis-retinal can form a new Schiff base linkage in the chromophore binding pocket (Hofmann et al., 1992; Pulvermüller et al., 1997). Consequently, retinal release is thought to be the rate-limiting step of visual pigment regeneration at low light levels (Ala-Laurila et al., 2006). This is supported by the fact that a retina isolated from the retinal pigment epithelium [RPE] can still regenerate up to 10% of its rhodopsin complement (Donner and Hemilä, 1975; Cocozza and Ostroy, 1987), making the need for

11-cis-retinal replenishment less immediate under these conditions. Even at high bleach levels, constant signaling keeps the photoreceptor saturated and functionally insensitive to light (Firsov et al., 2005), making the decay of meta intermediates a key step to recover sensitivity. Thus, natural variation in retinal release rates will likely influence of visual pigment regeneration in vivo.

215 The S2 component is a kinetically distinct time-course of scotopic recovery that is remarkably consistent in human rod photoreceptors, regardless of bleaching exposure

(Pugh, 1975; Lamb, 1981). The S2 component has been reproduced as a first-order reaction with a time constant of approximately 1.8 min-1 in several physiological experiments, including retinal densitometry (Alpern, 1971; Rushton and Powell, 1972), psychophysical recovery (Lamb and Pugh, 2004), and ERG a-wave recovery (Thomas and Lamb, 1999). We have measured the half-life of retinal release for bovine rhodopsin to be 2.0 min at the physiological temperature of 37oC [data not shown], which is comparable to the extrapolation of a previous Arrhenius plot of bovine rhodopsin retinal release (Farrens and Khorana, 1995). This half-life value, along with retinal release being a first order kinetic reaction, make it a remarkably similar process to the S2 component of dark adaptation in humans, and is also elicited by a light bleach. With bovine and human rhodopsin having similar decay rates of metarhodopsin intermediates

(Lewis et al., 1997; Shichida and Imai, 1998), it is reasonable to assume that there may be a connection between retinal release and dark adaptation. This would also imply that measuring retinal release could be a means to approximate the S2 component of dark adaptation in vitro.

There are also indications from physiological studies that the meta III decay rate is linked to the rate of dark-current recovery post light bleach in isolated photoreceptors in the mouse (Imai et al., 2007). The meta III intermediate has also been referred to as the

'storage form', as it can convert back to the physiologically active meta II, potentially further activating transducin and sustaining signaling activity (Heck et al., 2003; Lamb

216 and Pugh, 2004). It has been suggested that having a longer signaling state increases the sensitivity of the photoreceptor cells (Imai et al., 1997; Kuwayama et al., 2002; Shichida and Matsuyama, 2009), and thus would be advantageous for scotopic vision by amplifying second messenger activation (Sugawara et al., 2010). This idea is supported by a recent study where rhodopsin mutants that have longer retinal release rates also tend to have an increased capacity to activate transducin (Piechnick et al., 2012), indicating that variation in retinal release rates may also affect G protein signal amplification.

6.3 Genetic diversity in Actinopterygian fish

The basic concept of gene duplications being a force for evolution provided a platform for whole genome duplications to facilitate the emergence of novel protein function and promote radiations of new species (Ohno, 1970). Following a duplication event, gene copies will generally experience one of several fates. Neofunctionalization results when one copy retains ancestral function and a second copy takes on a novel function, subfunctionalization has each duplicate inheriting a different component of ancestral function, and nonfunctionalization involves one duplicate accumulating mutations that make it a non-functional pseudogene (Otto and Yong, 2002). Teleost fish represent the first clear example of an ancient genome duplication in vertebrate evolution (Jaillon et al., 2004; Woods et al., 2005). This event was first suggested based on the fact that both medaka [O. latipes] and zebrafish [D. rerio] possess seven clusters of the Hox gene compared to four in mammals and one in most invertebrates (Amores et al., 1998;

Wittbrodt et al., 1998). Because genome-wide duplications allowed for the expansion and retention of a gene family so essential to proper development, it stands to reason that

217 other types of gene families may also have expanded to provide novel function. Since I proposed the concept of functional diversification as a mechanism for the retention of duplicated rh2 opsins in teleost fish, I will now look at other well-studied examples of this phenomenon to help support this hypothesis.

A perfect example of a trait that is relatively diverse in teleost fish compared to other vertebrates is sex determination, which is attributed to the dynamic genomes of teleosts as a result of a genome-wide duplication event (Mank and Avise, 2009). Sex is determined by the formation of either the male or female gonad, which relies on a series of proteins involved in a biochemical cascade typically referred to as sex determinants and often located on sex chromosomes (Charlesworth, 1991). Remarkable diversity of sex chromosomes exists in teleost fish, even in cases when sex determination seems highly conserved. For example, even though most salmonids have XY sex chromosomes, some evidence suggests they are not all orthologous, and have attained this state through several different independent mechanisms (Woram et al., 2003; van

Doorn and Kirkpatrick, 2007). Similar phenomena can bee seen in the genus Oryzias, including medaka, and the stickleback genera Gasterosteus and Pungitius (Peichel et al.,

2004; Tanaka et al., 2007). In addition the evolved structures of sex chromosomes, sex determination in teleosts can also be influenced by environmental triggers, often associated with inducible genetic promoters, often when an ecological factor can differentially effect male and female fitness (Charnov and Bull, 1977; Conover, 1984).

The fact that a biological aspect such as sex determination can be so plastic in teleost fish

218 relative to other vertebrates, suggests that both genetic and ecological diversity could have a similar influence on the visual system.

Phototransduction is the process by which light is converted to electrical signals in the eye and is initiated by visual pigments; however, many other proteins are involved in this signaling process (see Chapter 1.2). In some cases, the same phototransduction proteins are used by both rod and cone photoreceptors, but there are also many examples where each cell type employs a different member of the same protein family. One example is the heterotrimeric G protein transducin, where rods and cones express distinct variants of all three subunits (Nordström et al., 2004). These variants are thought to have been generated as the result of two ancestral genome duplication events occurring prior to the radiation of jawed vertebrates (Miyata and Suga, 2001). Teleost fish have also experienced additional duplications of several phototransduction proteins, including cyclic nucleotide-gated ion channels, opsin kinases, and arrestins (Larhammar et al.,

2009). This suggests the potential for further functional diversification in the teleost visual system beyond changes in visual opsin complements.

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