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New understanding of rhodopsin in retinal degeneration and high gain phosphorylation

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New understanding of rhodopsin in retinal degeneration and high gain phosphorylation

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NEW UNDERSTANDING OF RHODOPSIN IN RETINAL DEGENERATION
AND HIGH GAIN PHOSPHORYLATION

by
Jiayan Chen

A Dissertation Presented to the
FACULTY OF THE GRADUATE SCHOOL
UNIVERSITY OF SOUTHERN CALIFORNIA
In Partial Fulfillment of the
Requirements for the Degree
DOCTOR OF PHILOSOPHY
(NEUROSCIENCE)

December 2006

Copyright 2006 Jiayan Chen

Dedication

To My Dearest Parents and Husband

献给我最亲爱的父母和先生

ii

Acknowledgements

I would like to send my words of gratitude to all those who helped to make
this thesis possible. First and foremost, I want to address my heartily profound
gratitude and appreciation to my Ph.D. mentor Dr. Jeannie Chen for being a very
patient and supporting advisor in my Ph.D. work with ideas, criticism and
encouragement. Her invaluable guidance, editorial advice and suggestions are strong
support to complete this dissertation. I am very grateful to my committee members
Dr. Robert H. Chow and Dr. Ralf Langen for their invaluable and continuous
support from experiment materials, techniques, criticism to spiritual influences as
passionate scientists. I want to thank Dr. Michael Quick, Dr. Emily Liman and Dr.
Chien-ping Ko for both practical and spiritual support for years. I want to thank
Francis A. Concepcion, Dr. Guang Shi and Dr. Ana Mendez for being congenial
colleagues and collaborators to work with and numerous enlightening discussions. I
want to thank all laboratory members for providing me with an amiable environment
and support for my work. I also really have to thank my friends Lan Wang,
Haijiang Cai and other friends for standing by me in both good and bad times in all
five years. Last but not least, I want to thank my dearest parents and husband for
their endless love and support in all my life.
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Table of Contents

Dedication
Acknowledgements
List of Figures
Abstract
vii
Chapter 1. Overview of Phototransduction Cascade and Possible
Mechanisms of Retinal Degeneration
1.1. Retina and Photoreceptor Cells
1.1.1. Retina and Photoreptor Cells in Vertebrates
1.1.2. Invertebrate Photoreceptors
1.2. Phototransduction Signaling in Rod Photoreceptors
1.3. Visual Cycle
1.4. Dark/Light Adaptation
1.5. Rhodopsin and Arrestin Interaction
1.6. Retinitis Pigmentosa and Retinal Degeneration
1.7. Possible Mechanisms in Retinal Degeneration
1.7.1. Visual G-protein Dependent Retinal Degeneration:
Constitutively Activation of Phototransduction Cascade.
1.7.2. Visual G-protein Independent Retinal Degeneration:
Toxic Complexes of Rhodopsin/Arrestin
1.7.3. Rhodopsin Misfolding
1.7.4. Rhodopsin Mistrafficking
1.8. Thesis Outline

Chapter 2. Stable Rhodopsin/Arrestin Complex Leads to Retinal
Degeneration in a Transgenic Mouse Model of Autosomal
Dominant Retinitis Pigmentosa
Abstract
2.1. Introduction
2.2. Meterials and Methods
2.2.1. Generation of Mouse Lines
2.2.2. Retinal Morphometry
2.2.3. Immunocytochemistry
2.2.4. Western Blot Analysis
2.2.5. GTP
γ
S Assay
2.2.6. Electroretinogram (ERG)

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v

2.2.7. Sample Preparation for Liquid Chromatography Mass
Spectrometry (LC-MS) Analysis
2.3. Results
2.3.1. K296E-induced Retinal Degeneration Is Transducin-
independent
2.3.2. K296E Activates Transducin and Causes Retinal
Degeneration in the arr1-/- Background
2.3.3. K296E-induced Retinal Degeneration Is Slowed in the
Absence of Arrestin and Transducin
2.3.4. Constitutive Activity of K296E Causes Retinal
Degeneration in arr1-/-Tr-/- Retinas
2.3.5. K296E Is Colocalized with Arrestin in the Retina.
2.3.6. K296E Is Retained in the Inner Segment and ONL
through Its Interaction with Arrestin
2.3.7. Hyperphosphorylation of K296E May Contribute to
Stable Complex Formation with Arrestin
2.4. Discussion
Acknowledgements

Chapter 3 Light Causes Phosphorylation of Non-Activated Visual
Pigments in Intact Mouse Rod Photoreceptor Cells
Abstract
3.1. Introduction
3.2. Meterials and Methods
3.2.1. S-opsin and K296E Transgenic Mouse Lines 63
3.2.2. Standard Peptides 64
3.2.3. Light Stimulation and Sample Preparation for LC-MS 64
3.2.4. LC-MS 65
3.2.5. Isoelectric Focusing 66
3.2.6. Mathematical Simulations of Trans-phosphorylation
3.3. Results
3.3.1. Detection of Rhodopsin and S-opsin Phosphorylation.
3.3.2. S-opsin Becomes Trans-phosphorylated Following
Activation of Rhodopsin by Long (515-620 nm)
Wavelength Light
3.3.3. Trans-phosphorylation Results in Multiply
Phosphorylated S-opsin
3.3.4. Demonstration of Trans-phosphorylation in
Transgenic Mice that Express Human K296E Opsin
3.4. Discussion
Acknowledgements

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Chapter 4 Rhodopsin Turnover and Trafficking in Rhodopsin-Timer
Transgenic Mice (Timer)
4.1. Introduction
4.2. Meterials and Methods
4.2.1. Generation of Non-aggregated Timer
4.2.2. Generation of Rhodopsin-Timer Transgenic Mice
(Timer) and Other Mutant Rhodopsin Constructs
4.2.3. Genotype Analysis
4.2.4. Retinal Whole Mount
4.2.5. Vibrotome and Confocol Imaging
4.2.6. EPON Morphometry
4.2.7. Cryosection
4.2.8. Western Blot
4.2.9. Fluorospectrometry
4.2.10. Cell Culture and Immunocytochemistry (ICC)
4.3. Results 100
4.3.1. Generation of Timer Transgenic Mice
4.3.2. Rhodopsin-Timer Fluorescence Expression
4.3.3. Morphology of Timer Transgenic Mice Was Normal in
rho+/+ and rho+/- Background But Not in rho-/-
Background
4.3.4. Fluorescence in Timer Transgenic Mouse Was Largely
from Membrane Fraction
4.3.5. Rhodopsin-Timer Fusion Protein Did Not Traffic to
ROS in rho-/- Background
4.3.6. Rhodopsin-Timer Was Elusive in the Western Blot
4.4. Discussion 109
4.4.1. Rhodopsin Trafficking
4.4.2. Rhodopsin Turnover Pattern in ROS 112

Bibliography 117

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List of Figures
Figure 1.1. Retina structure catoon and morphology.
Figure 1.2. Vertebrates and invertebrates photoreceptors structures.
Figure 1.3. Rod phototransduction cascade in vertebrates.
Figure 2.1. Experimental scheme leading to rescue of K296E-
induced retinal degeneration.

Figure 2.2. Retinal morphology of K296E-induced retinal
degeneration.

Figure 2.3. Retinal degeneration was more rapid when the proportion
of K296E was increased in the rhodopsin +/- background.

Figure 2.4. In vitro and in vivo analyses of K296E catalytic activity
in the arr1-/- background.

Figure 2.5. Quantification of K296E-induced retinal degeneration.

Figure 2.6. Pattern of retinal degeneration in dark-reared K296E
arr1-/-
Tr-/-
mice is recapitulated in light exposure of arr1-/-Tr-/- mice.

Figure 2.7. K296E was phosphorylated in a light-independent
manner and colocalized with arrestin in the photoreceptor cell
layer in darkness and in light.

Figure 2.8. K296E was retained in the inner segment and ONL via its
interaction with arrestin.

Figure 2.9. Tandem liquid chromatography-mass spectrometric
quantification of K296E expression in mouse retinal
homogenates.

Figure 2.10. K296E was highly phosphorylated and formed a stable
complex with arrestin.

Figure 3.1. Ion Chromatograms of eluted synthetic peptides
corresponding to C-terminal sequences of rhodopsin and S-
opsin.

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Figure 3.2. Phosphorylation of rhodopsin and S-opsin following
exposure to 360-420 nm Light.

Figure 3.3. Trans-phosphorylation of S-opsins following generation
of R* by 515-620 nm Light.

Figure 3.4. Rhodopsin phosphorylation and S-opsin trans-
phosphorylation level as a function of dark incubation time
after exposure to different intensities of light.

Figure 3.5. Detection of phosphorylated rhodopsin and S-opsin
species by isoelectric focusing.

Figure 3.6. Trans-phosphorylation of endogenous mouse rhodopsin
in dark-adapted K296E transgenic mouse retina.

Figure 3.7. Simulation of trans-phosphorylation by light-activated
rhodopsin.

Figure 4.1 Three-piece-ligation to generate non-aggregated form of
Timer construct.

Figure 4.2. Timer expression in rhodopsin+/+ and +/- background
seen in wholemount of freshly dissected retinas.

Figure 4.3. Timer fusion proteins did not completely traffic to ROS,
instead some of them were retained in ONL.

Figure 4.4. The morphology of Timer transgenic mice in different
background rho+/+, +/- and -/-.

Figure 4.5. Fluorescence in Timer transgenic mouse in rho+/- was
from membrane fraction.

Figure 4.6. Rhodopsin-Timer showed mislocalization in rho-/- mice
and COS cells.

Figure 4.7. Timer expression was not even in whole retina but in
selected region in cryosections.

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Abstract

Through the experiments described in this thesis, I strived to obtain a better
understanding the function of rhodopsin in retinal degeneration and light adaptation.
Over 100 rhodopsin mutation alleles have been associated with autosomal dominant
retinitis pigmentosa (ADRP), a blindness disorder that affects one in 3000 people
globally. These mutations appear to cause photoreceptor cell death through diverse
molecular mechanisms. We show that Lys296Glu (K296E), a rhodopsin mutation
associated with ADRP, forms a stable complex with arrestin that is toxic to mouse
rod photoreceptors. This cell death pathway appears to be conserved from flies to
mammals. Accumulation of stable rhodopsin/arrestin complexes in the inner segment
may be an important mechanism for triggering cell death in the mammalian
photoreceptor cells. Abnormal turnover of rhodopsin mutants could also underlie a
mechanism leading to cell death. In order to investigate rhodopsin turnover rate,
rhodopsin was tagged with a special fluorescent reporter Timer, which changes color
with the function of time, to report rhodopsin turnover activity.
Phosphorylation of rhodopsin is a required step in signal deactivation.
Rhodopsin exhibits high-gain phosphorylation in vitro whereby hundred-fold molar
excess of phosphates are incorporated into the rhodopsin pool per mole of activated
rhodopsin. The extent, by which high-gain phosphorylation occurs in the intact
mammalian photoreceptor cell, and the molecular mechanism underlying this
reaction in vivo, are not known. Trans-phosphorylation is a proposed mechanism for
high-gain phosphorylation whereby rhodopsin kinase, upon phosphorylating the
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activated receptor, continues to phosphorylate nearby non-activated rhodopsin. We
utilized two different transgenic mouse models: K296E and cone short-wavelength
opsin (S-opsin), and found that trans-phosphorylation occurs in the intact
photoreceptor cell. Trans-phosphorylation may play an important role in light
adaptation by decreasing transduction gain and thereby extending the range of the
rod response under certain steady-state lighting conditions.

Chapter 1
Overview of Phototransduction Cascade and
Possible Mechanisms of Retinal Degeneration

1.1. Retina and Photoreceptor Cells
1.1.1. Retina and Photoreptor Cells in Vertebrates
G-protein coupled receptors (GPCR) also known as seven-transmembrane
domain receptors, represent 1-5% of the invertebrate and vertebrate genomes
(Bockaert et al., 2003). They are the most numberous and diverse type of membrane-
bound proteins in a large family of membrane receptors. They transduce signaling
elicited by diverse stimili, including light, odorants, taste, hormones and neural
transmitters. In photoreceptors, the light pigments, rhodopsin and cone opsins, are
the prototypical G-protein-coupled receptors that mediate light signal transduction.
Vision starts in retina photoreceptor cells. The basic laminar organization and
physiological function are similar in all vertebrates’ retinas. The retina, around 0.4
millimeters thick, is a highly ordered, multilayered neural tissue that is composed of
five classes of neurons: photoreceptors, bipolar cells, ganglion cells, horizontal cells
and amacrine cells (Figure 1.1.A).
Photoreceptors, lying in the outer region of the retina, absorb photons by
specialized visual pigment molecules, and then convert this information via a
signaling cascade into a change in current which is passed onto the next layer of cells
and ends in the visual cortex to produce a visual image. There are two types of
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bottom
top
top
B
A
bottom

Figure 1.1. Retina structure catoon and morphology. (A) cartoon of retina cells. (B) Transverse
section of human retina. The retina can be recognized as 10 layers. They are RPE: retinal
pigment epithelium; OS: outer segment; IS: inner segment; ONL: outer nuclear layer; fiber layer;
OSL: outer synaptic layer; INL: inner nuclear layer; ISL: inner synaptic layer; GCL: ganglion
cell layer, and optic fiber layer. Measurement of the thickness of outer nuclear layer (ONL) can
be used to quantify the degree of retinal degeneration. (from “The First Step in Seeing”)

photoreceptors in vertebrates, rods and cones, named after their outer segment shapes
(Figure 1.2.A-B). Rods and cones are morphologically and functionally distinct.
Both rods and cones have outer segments connected to inner segments by a cilium.
The rod outer segment, located at the distal surface of the retina, consists of a stack
of membranous discs formed by internalization of the plasma membrane. These discs
contain the light-absorbing photopigments, rhodopsin. The membrane pinch off to
form stacked discs in rods, but is continuous with the plasma membrane in cones.
The outer segment is the specialized site for phototransduction in both rods and
cones. Rods and cones have different spectral sensitivity because of the different
light pigments. Visible light ranges from short- (blue), medium- (green), and long-
wavelength (red) radiation. Rhodopsin in rods has maximal spectra ( λm) sensitivity
at 500 nm, whereas human blue, green and red opsins are most sensitive to short-
(430 nm), medium- (535 nm) and long-wavelength (560 nm) light, respectively. The
murine cones have two opsins: short-wavelength opsin (S-opsin) with λm = 360 nm
and medium-wavelength opsin with λm = 510 nm. Rods with rhodopsin are
specialized for function in dim light, providing a scotopic vision up to 10
-2
lux
illumination; whereas cones function well in bright light and provide high temporal
and spatial resolution. Cones mediate a so-called photopic vision that ranges between
10
-1
– 10
5
lux light. 10
-2
- 10
-1
lux is known as mesopic vision, which operates under
moon light. This function is mediated by both rod and cones. Hence we use rods to
see starlight, use both rods and cones to see moonlight, and use cones to see any light
brighter than moon light. Most vertebrates are rod dominant, with rods comprising as
3

Figure 1.2. Vertebrates and invertebrates photoreceptors structures. (A-B) vertebrate
photoreceptors cells: A. rod, B. cone. In rods and cones, the outer segment is separated from
inner segment (cytosol) by a connecting cilium. Rhodopsin turnover is through ROS
shedding and phagocytosis by retinal epithelial cells. (C-E) invertebrate single eye and
photoreceptor cell. C. a photoreceptor cell composed of cytosol and rhabdomere where
phototransduction takes place. D. the sagital section of ommatidium (single eye) structure,
including rod-like R1-6 cells, cone-like R7-8 cells. E. the horizontal section of a
photoreceptor cell. In invertebrate photoreceptors, rhodopsin turnover through endocytosis
from rhabdomere to cytosol. (This picture is adapted from Dr. G. Shi and Dr. C. Montell)

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much as ~95% of photoreceptors. In adult human retina, there are ~91 million rods
and ~5 million cones (Oyster, 1999). Because of their prevalence, rods not only
mediate phototransduction, but also play a significant role in maintaining the overall
structural integrity of the retina, and in retinal degeneration. In this study, I will focus
on the rod system to investigate mechanism of retinal degeneration,
transphosphorylation and rhodopsin turnover.
The retina can be recognized as 10 layers under light microscope, four of
which are photoreceptor layers including outer segment (OS), inner segment (IS),
outer nuclear layer (ONL) and fiber layer (synapse layer) (Figure 1.1B). Retinal
degeneration can be easily detected by changes in morphology of these layers, e.g.
disorganized outer segment and inner segment, and cell loss in the ONL. The
thickness of ONL is the classical index of the severity of retinal degeneration. By
this parameter, we can compare and quantify the degree of retinal degeneration in
different genetic backgrounds in the experiments. One layer of cells outside of
photoreceptor cells are retinal pigment epithelium (RPE), a single layer of polygonal,
polarized epithelial cells which are very critical in maintaining photoreceptor cell
functions.
The rod outer segment is the site of phototransduction. It is composed of
hundreds of flattened membranous discs that pinched off from the plasma
membrane. Embedded in the discs are a large number of visual pigments rhodopsin,
as many as 10
8
per cell. Rhodopsin is a prototypical seven transmembrane protein in
the large G-protein coupled receptor (GPCR) family. It is composed of two parts:
5

chromophore and apoprotein opsin. The 11-cis-retinal chromophore is covalently
attached to opsin through a protonated Schiff base linkage to the Lys-296 (K296)
(Robinson et al., 1992). The rod outer segment contains high levels of
phototransduciton proteins. Large amount of proteins that maintain house-keeping
functions are contained in rod inner segment.
Photoreceptor cells depend on the RPE to support their survival and functions.
RPE have long microvilli on the apical surface interdigitating with photoreceptor
outer segment, whereas their basal surfaces are adjacent to Bruch’s membrane. RPE
controls the volume and composition of fluid in the subretinal space through the
active transport of ions, fluid and metabolites, which are necessary for photoreceptor
survival. They also play a major role in regeneration of the chromophore, with which
opsin binds to become rhodopsin. RPE cells are also the most active phagocytes in
the body, digesting ~2000-4000 discs of photoreceptor outer segment per day.
Photoreceptor cells use 3-4 times more oxygen than other retinal cells and CNS
neurons because of their high turnover rate of ROS. They have the highest rate of
oxidative metabolism in the whole body (Wu et al., 2006). With this fast and highly
efficient turnover, rod outer segment can maintain fully functional light pigments.
ROS turnover also may underlie a mechanism by which the retina adapts to new
environmental light (Young, 1971a; Schremser and Williams, 1995a, b). We will
further discuss rhodopsin turnover in Chapter 4. In general, the function of RPE cells
is very critical in maintaining and supporting photoreceptor survival and function. If
RPE cells die, photoreceptors cells will also succumb to death.
6

Rhodopsin and opsin are constrained in an inactive state by a salt bridge
between the protonated ε-amino group of K296 and its counterion E113 (Robinson et
al., 1992; Kim et al., 2004). Upon light illumination, isomerization of 11-cis-retinal
to the all-trans form breaks the salt bridge of rhodopsin, and induces rhodopsin to
convert into an active conformation metarhodopsin II (MII), which in turn activates
the phototransduction cascade. Similarly, the opsin can switch to the active state
when the salt bridge in opsin is broken which can occur in the opsin point mutations,
K296E, G90D, A292E. In these mutations, the salt bridge between K296 and E113 is
interrupted by mutation, which results in constitutively active form of opsin. The
constitutively activated opsin mutant K296E can cause retinal degeneration in both
mice and human (Keen et al., 1991; Li et al., 1995).

1.1.2. Invertebrate Photoreceptors
During the long history of animal evolution, two major classes of
photoreceptors have emerged: ciliary photoreceptors typified by vertebrate rods and
cones, and microvillar or rhabdomeric photoreceptors typical of arthropods and
most mollusks (Hardie, 2001). Mouse and Drosophila are classical animal models
of each type. In contrast to mammals, Drosophila has compound eyes, composed of
~800 simple eyes named ommatidia. Each ommatidium has 20 cells, among which 8
are photoreceptors R1-8 (Figure 1.2.C-E). Phototransduction takes place in the
specialized rhabdomere structure. The rhabdomere of R7 and R8 occupy the central
area of the ommatidium, while that of R1-6 are around them, forming the circle of
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outer rhabdomere. R1-6 cells functionally resemble rods. They are very sensitive to
light, express a single visual pigment, and make up the majority of photoreceptor
cells; whereas R7-8 cells resemble cones, which are less sensitive to light, express
multiple visual pigments, and comprise a high-acuity system (Montell, 1999). In
Drosophila photoreceptors, rhodopsin turnover is through endocytosis, which occurs
from rhabdomere to cytosol. As mentioned previously, rhodopsin turnover is
through phagocytosis by RPE in vertebrate. In general, invertebrates photoreceptors
function similarly as vertebrates although structures are different.

1.2. Phototransduction Signaling in Rod Photoreceptors
Phototransduction is the process by which a photon of light (or photons)
generates an electrical response in a photoreceptor cell through a signaling cascade.
The ability of rods to mediate dim light vision is largely attributed to the sensitivity
of rod pigment, rhodopsin, that signals single photon absorption. Phototransduction
begins with the absorption of a single photon by rhodopsin (Figure 1.3). Upon
photon absorption, chromophore is photoisomerized from 11-cis-retinal to all-trans-
retinal within subpicosecond time scale, which induces the conversion of rhodopsin
(R) through a sequence of thermal intermediates – photorhodopsin, bathorhodopsin,
lumirhodopsin and metarhodopsin I – III rhodopsin within milliseconds (Hayward et
al., 1981; Schoenlein et al., 1991). The metarhodopsin II conformation catalyses the
activation of heterotrimeric G-protein (G
t
, transducin) by GTP-GDP exchange
(Baumann, 1976; Dickopf et al., 1998), which leads to G
α
subunit dissociation from
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G
βγ
subunits. Transducin G
α
subsequently binds to γ-subunit of the effector enzyme
phospodiesterase (PDE) which releases the inhibition to the catalytic α and β
susbunits. PDE
αβ
then hydrolyses 3’-5’ cyclic guanosine monophosphate (cGMP) to
5’GMP. In the dark, ~3% of the cGMP-gated channels are open (Yau and Baylor,
1989) to form the “dark current” carried by influx of Na
+
, Ca
2+
and small amounts of
Mg
2+
. The permeability of the CNG channels to cations enables a steady inward
current carried by Na
+
and Ca
2+
driven by the concentration gradients of these ions
across plasma membrane in darkness (Yau and Nakatani, 1984; Nakatani and Yau,
1988). The influx of photocurrent is balanced by current efflux by Na
2+
/K
+

exchanger from inner segment, generating a loop of current flow known as
circulating current. In light, decrease of cGMP level results in closure of cGMP-
gated channels thus decreases or terminates Na
+
and Ca
2+
influx, and the
photoreceptor cells are hyperpolarized and Ca
2+
level decreases. When GTP is
hydrolyzed to GDP by GTPase activating protein (GAP) composed of regulator of G
protein signaling type 9 and G-protein β subunit 5 (RGS9/G β5) complex anchored
by RGS9 anchoring protein (R9AP), PDE
γ
will rebind with αβ subunit which
terminates hydrolysis of cGMP and returns to the basal level activity.
When cGMP-gated channels close, guanylate cyclase activating proteins
(GCAPs) are activated by decreased Ca
2+
level early after flash of light, and start
activating guanylate cyclase which synthesizes cGMP. The increased cGMP level
causes the reopening of cGMP-gated channels. After activating transducin, the
photolysed rhodopsin (R*) is deactivated by multiple phosphorylations at the C-
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terminus by rhodopsin kinase (RK), followed by binding of arrestin to
phosphorylated, light-activated rhodopsin (R*-P). Subsequently, the chromophore is
released and arrestin dissociates from R*-P in the outer segment. The opsin is then
dephosphorylated by a yet to be identified rhodopsin phosphatase. After all-trans-
retinal is regenerated to11-cis-retinal in RPE cells, it will reassociate with opsin and
regenerate rhodopsin to be ready for another photon absorption.
Upon photoactivation of rhodopsin, the deactivation and adaptation systems
are activated. The guanylate cyclase activity begins to increase about 60 ms after the
flash, rhodopsin kinase activity commences about 100 ms after a flash of light and
lasts about another 30-50 ms upon arrestin binding, and GAP begins to shut-off
transducin at the same time as arrestin binding (Makino et al., 2003). It was reported
recently that RGS9-mediated inactivation of transducin is the rate-limiting step for
mammalian rod response recovery, and it is directly proportional to the expression
level (Krispel et al., 2006). Metarhodopsin II can also achieve inactivation via
thermal decay, however, the speed is ~1000 fold slower than seen in the real
photoresponse (Hofmann, 2000). Interruption of either phototransduction cascade or
inactivation of rhodopsin can result in dysfunction and retinal degeneration.

1.3. Visual Cycle
After inactivation by rhodopsin kinase and arrestin binding, R* becomes
opsin with the release of all-trans retinal. The opsin must be regenerated to form
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Figure 1.3. Rod phototransduction cascade in vertebrates (Curtesey of F.A.
Concepcion). Upon photoexcitation, 11-cis retinal of rhodopsin is isomerized to all-
trans retinal, leading to the formation of catalytic active rhodopsin (R*). R* catalyzes
GTP–GDP exchange on transducin α subunit (G
α
). GTP-G
α
then binds with PDE γ
subnuit on disk membrane which leads to exposure of the catalytic sites on PDE αβ
subunits and subsequently to catalyze cGMP hydrolysis to GMP. As a result of a fall
of intracellular cGMP concentration, the cGMP is removed from cGMP-gated
channel, causing channel to close. The inward “dark current” is suppressed and
photoreceptor cell hyperpolizes.
After activation of transducin, R* is phosphorylated by rhodopsin kinase.
Phosphorylation reduces the catalytic activity of rhodopsin and promotes arrestin
binding, after which rhodopsin activity is completely quenched. All-trans retinal
dissociates from opsin following Schiff base hydrolysis. All-trans retinal is converted
back to 11-cis retinal by retinoid cycle reactions in RPE. The opsin is
dephosphorylated by protein phasphase and reconstituted with 11-cis retinal to form
regenerated rhodopsin.

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rhodopsin by binding with 11-cis retinal before another photon absorption can occur.
This regeneration process is called “the visual cycle”.
The visual cycle in invertebrates is a rather simple process. For some
invertebrates like insects and cephalopods, the conversion from all-trans retinal to
11-cis retinal can take place within photoreceptors by photoisomerization without
chromophore release (Hardie and Raghu, 2001; Lamb and Pugh, 2004). However it
is a much more complicated, lengthy and enzymatic process in vertebrates which
requires both photoreceptors and retinal pigment epithelial cells (RPE). It is called
the retinoid cycle. Cone pigment regeneration is much faster than rod pigment. For
example in humans, the cone regenerates in less than 10 minutes, whereas rod
pigments needs at least 20-30 minutes (Fain et al., 2001).
The retinoid cycle has four steps: photochemistry, removal of retinoid,
reconversion of retinoid and delivery of retinoid (Lamb and Pugh, 2004). 1)
Photochemistry: Upon photon absorption, the 11-cis retinal is photoisomerized to all-
trans retinal, which converts rhodopsin into R*. 2). Removal of retinoid: The all-
trans retinal can be flipped into the cytoplasm by the ATP-binding cassette, retina
(ABCR) transporter and reduced by all-trans retinol dehydrogenase (RDH) after
hydrolysis. It was also suggested by a group that the all-trans retinal is reduced into
all-trans retinol (vitamin A) by RDH when still non-covalently bound to
metarhodopsin, and then released (Schadel et al., 2003). The all-trans retinol is then
carried by inter-photoreceptor retinol binding protein (IRBP) to cross cytoplasm and
plasma membrane and reaches RPE. 3) Reconversion of retinoid: In RPE cytoplasm,
12

the all-trans retinol is carried by cellular retinol binding protein (CRBP) and
esterified by lecithin retinol acyl transferase (LRAT). The all-trans retinyl ester is
then carried by RPE65, and isomerized to 11-cis retinol by retinyl ester
isomerohydrolase. It is further oxidized to 11-cis retinal by 11-cis RDH and bound to
cellular retinaldehyde binding protein (CRALBP). 4) Delivery of retinoid: The 11-cis
retinal then diffuses to photoreceptor outer segment disc membrane and covalently
binds to opsin to form a visual pigment. This process is probably facilitated by IRBP.
In cones, there is an alternative, novel Müller cell retinoid cycle which is 20
fold faster than the RPE cycle (Mata et al., 2002). In this pathway, all-trans retinol is
absorbed by Müller cell instead of RPE. Here all-trans retinol is converted directly to
11-cis retinol by a novel all-trans retinol isomerase. 11-cis-retinol is then esterified
by a novel enzyme 11-cis retinyl ester synthase into 11-cis retinyle ester which is
hydrolyzed by retinyl ester hydrolase to yield 11-cis retinol. This product is absorbed
directly and only by cones and oxidized by 11-cis retinol dehydrogenase into 11-cis
retinal. This new pathway is very efficient, enabling cones to keep up with the high
demand of cone pigment regeneration under bright light which may play a
significant role in the fast cone light adaptation.

1.4. Dark/Light Adaptation
Our visual system can function over an enormously wide range of light
intensities, from weak starlight to very bright light that spans 10 log units of light
intensity. Visual threshold is the dimmest light that transduction cascade is able to
13

amplify and produce a reliable photoresponse that is clearly distinguished from dark
noise events.
The ability of the visual system continue to function in increasing
illumination is called light adaptation. Light adaptation prevents saturation at
increasing light levels and enables photoreceptors to continually respond to light.
The responsiveness of saturated photoreceptors recovers slowly in a subsequent dark
period. This recovery process is termed dark adaptation (Fain et al., 1996). Dark
adaptation restores rod function by regenerating rod pigment rhodopsin. Both light
adaptation and dark adaptation allow photoreceptors to “see” as soon as possible.
Two significant hallmarks of photoreceptor light adaptation are: sensitivity
reduction and accelerated response kinetics in the presence of background
illumination (Baylor and Hodgkin, 1974). The flash sensitivity is defined as S
F
which
is peak amplitude of the flash response divided by the flash intensity (Pugh et al.,
1999). There are several mechanisms contributing to light adaptation, including Ca
2+

dependent feedback by GCAP, recoverin and CNG-gated channels, and Ca
2+

independent mechanism like protein translocation including transducin and arrestin
(Pugh et al., 1999; Burns and Baylor, 2001; Fain et al., 2001; Sokolov et al., 2002;
Burns and Arshavsky, 2005; Strissel et al., 2006). Trans-phosphorylation is possibly
one mechanism in light adaptation. Different independent groups have found that,
several hundred-fold molar excess of phosphates are incorporated into the rhodopsin
pool per mole of R* in isolated rod outer segments, which is named “high gain
phosphorylation” (Aton, 1986; Binder et al., 1990; Chen et al., 1995a; Binder et al.,
14

1996). The nearby unbleached rhodopsin could be phosphorylated by rhodopsin
kinase activated by a nearby photoexcited rhodopsin. This putative mechanism is
named trans-phosphorylation. The trans-phosphorylation could possibly decrease
transduction gain and contribute to less saturation and therefore extend the rod
response range. The transphosphorylation mechanism was investigated and proved
in our transgenic system. The details will be discussed in Chapter 3.
In general, light adaptation plays an important role in photoreceptor
physiology. It can also be used as a parameter to analyze visual function of mouse
model of retinal degeneration as described in Chapter 2.

1.5. Rhodopsin and Arrestin Interaction
Phosphorylation of rhodopsin C-terminus and arrestin binding play a key role
in rhodopsin shut-off. Phosphorylation of rhodopsin’s C-terminal serine and
threonine residues partially deactivated R*, and arrestin binding completely shuts
down the phototransduction signal (Chen et al., 1995b; Chen et al., 1999a; Chen et
al., 1999b). In mammals, there are four arrestin proteins, arrestin 1 or visual arrestin,
arrestin 4 or cone arrestin, arrestin 2 and arrestin 3. Among them, arrestin 2 and 3 are
expressed globally to regulate over 1000 different GPCRs. Receptor-bound β-
arrestins can serve as an adaptor to internalize through coated pit via binding to
clathrin and AP2, and serve as a scaffold protein to promote the formation of
multiprotein signaling complexes (Ferguson et al., 1996; Perry and Lefkowitz, 2002).
Visual arrestin and cone arrestin are specifically expressed in rods or cones,
15

respectively, regulating rhodopsin or cone opsin activity (Luttrell and Lefkowitz,
2002; Vishnivetskiy et al., 2004). It was found that residues 49-90 ( β-strands V and
VI and adjacent loops in the N-domain) and residues 237-268 ( β-strands XV and
XVI in the C-domain) in visual arrestin and their homologous regions in arrestin2 are
necessary and sufficient to determine their preferred receptor targets (Vishnivetskiy
et al., 2004).
Visual arrestin binding with rhodopsin can shut down R* activity and prevent
further G-protein signaling, which is significant to phototransduction. Visual arrestin
shows 10-20 times higher binding affinity to light-activated phosphorylated
rhodopsin (R*-P, K
D
= 20 nM) than to non-activated, phosphorylated (P-R) and
activated unphosphorylated rhodopsin (R*) (Gurevich and Benovic, 1993). A model
of sequential multisite binding of arrestin to rhodopsin was proposed to explain the
binding selectivity (Gurevich and Gurevich, 2004). In this model, arrestin has two
“sensor” sites, one for phosphorylation and one for activation of rhodopsin.
Simultaneous engagement of both sensors triggers a global conformation change of
arrestin and transits it from inactive state into the high-affinity receptor-binding state.
To date, only the phosphate sensor has been characterized (Gurevich and
Gurevich, 2004). A network of five interacting charged residues connecting two
arrestin domains constitutes the arrestin phosphate sensor. Lys14 and Lys15 are the
primany phosphate-binding residues and deliver receptor-attached phosphates to the
polar pore. Phosphates binding to Arg175 and its partner Asp296 also neutralize the
charge and help to break up the structural constraint, therefore triggering arrestin into
16

the active state. The activation sensor is less clear than the phosphorylation sensor
and still awaits more investigation. It was shown recently that the active receptor
preferentially engage the C-domain of arrestin (Hanson and Gurevich, 2006).
After arrestin binds R*-P, metarhodopsin II (MII) decays to opsin and the
arrestin dissociates from opsin. Therefore the R*-P/arrestin complex forms
transiently. However it is different for mutants in pathological conditions. We will
further discuss the abnormal rhodopsin/arrestin complex which can cause retinal
degeneration in section 1.7.

1.6. Retinitis Pigmentosa and Retinal Degeneration
Many diseases, like retinitis pigmentosa, age-related macular degeneration,
congenital stationary night blindness, and cone dystrophy, can cause blindness by
photoreceptor cell death. Although associated with mutations in different
components of the visual signaling cascade, they all share a common final outcome:
the apoptotic death of photoreceptor cells (Reme et al., 2000; Ranganathan, 2003).
Retinitis pigmentosa (RP) is one of the most common diseases resulting in retinal
degeneration, affecting about 15 million people in the world (van Soest et al., 1999).
Such diagnosed patients are clinically characterized as progressive night blindness,
narrowing of the visual field, reduced central vision, and increasing sensitivity of
glare followed by complete vision loss in later life. As a heterogeneous group of
genetic disorders, it can be inherited as autosomal dominant RP (ADRP), autosomal
recessive RP, sex-linked RP and mitochondrial RP (Wenzel et al., 2005). Currently,
17

over 100 mutations in the rhodopsin/opsin gene have been described that account for
30-40% of ADRP (RetNet at http://www.sph.uth.tmc.edu/RetNet/). Thus rhodopsin
plays a significant role in the retinal degeneration of Retinitis Pigmentosa.
It has been widely accepted that apoptosis is the final cell death pathway in
Retinitis Pigmentosa and other retinal degeneration diseases (Reme et al., 2000). The
apoptotic cells are characterized by condensation of chromatin, membrane blebbing
and disintegration of dying cells into apoptotic bodies which are finally engulfed by
phagocytic cells (Kerr et al., 1972). Apoptosis can also include internucleosomal
DNA cleavage (Nagata, 2000), exposure of phosphatidylserine to the outside of
cellular membranes (Schlegel and Williamson, 2001; Williamson et al., 2001).
Endopeptidase caspase were considered as the central executioners of the apoptotic
programmable cell death by cleaving a variety of intracellular substrates at aspartic
acid sites. However it was found in recent years that there are also caspase-
independent apoptosis including cathepsins, calpains, granzymes A and B, serine
proteases like AP24 and proteasomes (Wenzel et al., 2005). Apoptosis was further
found to also involve various cellular compartments including mitochondria,
lysosomes, proteasomes or autophagic vacuoles. It was even noticed that some dying
cells show morphological features of both apoptosis and necrosis. In general, retinal
cell death involves many different cellular compartment and molecules.
There are a number of mouse models of Retinitis Pigmentosa. The
Lys296Glu (K296E) transgenic mouse is one of them. K296E is a naturally
occurring opsin mutation found in human ADRP patients. The patients present a
18

severe phenotype with early onset of blindness and development of cataracts (Keen
et al., 1991). This mutation renders the opsin incapable of binding to chromophore
and abolishes the salt bridge between K296 and E113, thus results in a constitutively
active form of opsin (Robinson et al., 1992). It was further shown that the K296E
mutant can constitutively activate transducin in vitro (Rim and Oprian, 1995), and
subject to be phosphorylated by rhodopsin kinase and binding of arrestin. So it was
commonly thought K296E and other constitutively active opsin mutants cause retinal
degeneration by constitutive signaling of phototransduction. K296E transgenic mice
show light-independent retinal degeneration in transgenic mouse, and transducin was
not activated by K296E until arrestin removal and dephosphorylation treatment (Li et
al., 1995). This suggested that constitutive activation of phototransduction is not the
mechanism of K296E retinal degeneration as traditionally thought. We further
discussed whether the mechanism of retinal degeneration caused by K296E is
through formation of toxic rhodopsin/arrestin complex. This topic is discussed in
Chapter 2.

1.7. Possible Mechanisms in Retinal Degeneration
Numerous genes have been shown to be responsible for retinal degeneration,
including genes of transcription factors, genes involved in photoreceptor metabolism,
structural support, and almost all the genes participating in phototransduction
cascade (Travis, 1998; Lev, 2001). However, there is a large gap between gene
mutations to the final cell death signaling. Since apoptosis is still so far considered
19

the major final pathway of cell death, prevention of apoptosis is conceptually
feasible for all types of mutations and therapeutic targeting. Nevertheless, successful
prevention of apoptosis requires good understanding of signaling and death
mechanisms.
In ontogenetic terms, photoreceptor cells are post-mitotic neuron cells that
can survive and function for the full lifetime of the animal, as is the case for all other
central nervous system neurons (Wu et al., 2006). However, that photoreceptor cells
are more vulnerable to small mutations in their proteins and environmental damage
makes them a lot more fragile than other central nervous system neurons (Lavail,
1989). In general, the dominant rhodopsin-mediated degeneration mechanisms can
include the following.

1.7.1. Visual G-protein Dependent Retinal Degeneration: Constitutively
Activation of Phototransduction Cascade.
Some rhodopsin mutants are constitutively active like G90D, A292E, and
cause night blindness (Rao et al., 1994). Some mutations affecting rhodopsin like
RPE65
-/-
, which is defective in producing chromophore 11-cis-retinal, can produce
weak but constitutively active opsin and lead to retinal degeneration (Grimm et al.,
2000; Wenzel et al., 2001; Woodruff et al., 2003). Similarly, arrestin or rhodopsin
kinase (RK) knockout can lead to retinal degeneration in light-exposed mice (Chen
et al., 1999a; Chen et al., 1999b). Because all these degeneration can be blocked in
the transducin null background, it indicates that the retinal degeneration results from
20

constitutively activation of phototransduction cascade. It is a transducin- (G
t
)
dependent retinal degeneration pathway. The “equivalent-light hypothesis” proposed
that the cause of photoreceptor-cell death in these mutants is constitutive activation
of phototransduction cascade by the active form of rhodopsin or opsin which is
equivalent to continuous light exposure to photoreceptors even in the absence of
light (Lisman and Fain, 1995).

1.7.2. Visual G-protein Independent Retinal Degeneration: Toxic Complexes of
Rhodopsin/Arrestin
This mechanism was discovered in Drosophila, in which rhodopsin and its
regulatory protein arrestin can form stable and persistent complexes to cause retinal
degeneration through the clathrin-dependent endocytosis of these complexes into
cytoplasmic compartments (Alloway et al., 2000; Kiselev et al., 2000; Iakhine et al.,
2004). These include the norpA mutant encoding phospholipase C (PLC
β
) and
rdgC
306
mutant encoding rhodopsin phosphatase. Phosphorylation of arrestin by
calmodulin-dependent protein kinase II, and dephosphorylation of rhodopsin by
rdgC-encoded phosphatase are prerequisite for dissociation of rhodopsin/arrestin
complex. Any mutation disturbing these two processes can cause the accumulation
of rhodopsin/arrestin complexes in Drosophila. The lack of phosphatase in the rdgC
mutant results in the inability of R* dephosphorylation; the norpA mutant with
defective PLC
β
can lead to decreased Ca
2+
influx, which prevents arrestin
phosphorylation by a calcium-dependent protein kinase II (PKC II). Thus these two
21

mutations lead to accumulation of rhodopsin/arrestin complexes. When the
accumulated complexes are endocytosed from rhabdomere into cytoplasmic
compartments, they signal photoreceptor cell death (Alloway et al., 2000; Kiselev et
al., 2000). Retinal degeneration in these mutants cannot be rescued in G
q
knockout
background, which implicates a G-protein independent mechanism. It was further
supported in ninaE
pp100
rhodopsin mutant in Drosophila (Iakhine et al., 2004). Would
this mechanism be conserved in vertebrates? In physiological conditions, there are
R*-Arr transiently formed in both rhabdomere of Drosophila or rod outer segment in
vertebrates. Although endocytosis is not supported in the rod outer segment of
vertebrate, this process can occur in the equivalent site of Drosophila cytosol, the
inner segment of rods, where rhodopsin and all other proteins are synthesized and
diseases are initiated. We hypothesize that when the accumulation of
rhodopsin/arrestin complexes reaches a critical dose-threshold in the inner segment,
it initiates retinal degeneration in vertebrates. This will be further discussed in
Chapter 2.

1.7.3. Rhodopsin Misfolding
Some of ADRP-causing rhodopsin mutations lead to defective folding,
including mutations in the transmembrane, intradiscal or cytoplasmic domains, like
P23H. The in vitro study showed that the misfolded rhodopsin P23H can form
aggresomes in the cell and target to impair ubiquitin proteasome system to cause
retinal degeneration (Illing et al., 2002; Saliba et al., 2002).
22

1.7.4. Rhodopsin Mistrafficking
Rhodopsin is synthesized in the endoplasmic reticulum and Golgi apparatus
in the inner segment and transported in post-Golgi vesicles to rod outer segment
(Deretic, 1998; Deretic et al., 1998). Rhodopsin with C-terminal mutations can
usually fold correctly but can not traffic normally to the outer segment, possibly by a
defect in the targeting of the protein to rod outer segment (Illing et al., 2002; Saliba
et al., 2002), like S334ter, Q344ter. The mistrafficked rhodopsin that accumulates in
the inner segment is also suggested as a pathogenic mechanism to cause retinal
degeneration.

1.8. Thesis Outline
In this thesis, I and my colleagues concentrated on understanding the
pathological molecular mechanisms of retinal degeneration in a common blind
disease autosomal dominant retinitis pigmentosa (ADRP), and the mechanism
involved in rhodopsin phosphorylation, by employing a combination of genetic,
electrophysiological, biochemical and biophysical techniques.
In Chapter 2, the investigation was focused on a hypothesis that rhodopsin-
arrestin complexes cause retinal degeneration in the K296E rhodopsin mutant
transgenic mouse model. By manipulating K296E in different gene knockout
backgrounds, the relationship of the mutant rhodopsin with different protein partners
was examined. The morphology, immunocytochemistry, electroretinogram and other
biochemical assays implicated the rhodopsin/arrestin complex as underlying initiator
23

of retinal degeneration. It indicates that this mechanism is conserved from
invertebrates to vertebrates.
In Chapter 3, the special characteristics of K296E rhodopsin mutant
transgenic mice together with cone S-opsin transgenic mice were utilized to
investigate a hypothesized mechanism that underlies “high-gain phosphorylation”.
The cross-desensitization mechanism of rhodopsin, called transphosphorylation, was
confirmed to be present in vivo, in that non-activated rhodopsin can be
phosphorylated by rhodopsin kinase which was activated by nearby photoexcited
rhodopsin.
To have a better visualization and understanding of rhodopsin and mutant
rhodopsin turnover rate and possible mechanisms, Rhodopsin-Timer transgenic mice
were generated. In this construct, rhodopsin is tagged with a special fluorescent
protein which changes color with aging. This transgenic mouse presented some
indication of circadian rhythm of ROS renewal and some hint about misfolding and
mistrafficking for some rhodopsin mutants. This content is presented in Chapter 4.

24

Chapter 2*
Stable Rhodopsin/Arrestin Complex Leads to Retinal Degeneration
in a Transgenic Mouse Model of
Autosomal Dominant Retinitis Pigmentosa
Abstract
Over 100 rhodopsin mutation alleles have been associated with autosomal
dominant retinitis pigmentosa (ADRP). These mutations appear to cause
photoreceptor cell death through diverse molecular mechanisms. We show that
K296E, a rhodopsin mutation associated with ADRP, forms a stable complex with
arrestin that is toxic to mouse rod photoreceptors. This cell death pathway appears to
be conserved from flies to mammals. A genetics approach to eliminate arrestin
unmasked the constitutive activity of K296E and caused photoreceptor cell death
through a transducin-dependent mechanism that is similar to light damage.
Expressing K296E in the arrestin/transducin double knockout background prevented
transducin signaling and led to substantially improved retinal morphology, but did
not fully prevent cell death caused by K296E. The adverse effect of K296E in the
arrestin/transducin knockout background can be mimicked by constant exposure to
low light. Furthermore, we found that arrestin binding causes K296E to mislocalize
to the wrong cellular compartment. Accumulation of stable rhodopsin/arrestin
complex in the inner segment may be an important mechanism for triggering the cell
death pathway in the mammalian photoreceptor cell.
*This work has been accepted by Journal of Neuroscience and is in printing.
25

2.1. Introduction
Rhodopsin, a prototypical G-protein-coupled receptor (GPCR), is a light-
sensitive visual pigment in retinal rod photoreceptors. Photon absorption causes
isomerization of 11-cis retinal, the chromophore that is covalently attached to the
opsin apoprotein. This in turn causes conformational changes in the protein moiety
and subsequent activation of transducin, the visual G-protein. Like other GPCRs,
rhodopsin is deactivated sequentially by phosphorylation and arrestin binding.
Over 100 rhodopsin mutations have been linked to ~30% of patients
diagnosed with autosomal dominant retinitis pigmentosa (ADRP), a blinding
disorder that affects one in 3000 people (RetNet,
http://www.sph.uth.tmc.edu/RetNet/home.htm). Functional studies of some
rhodopsin mutants in ADRP have led to their classification into distinct groups that
implicate misfolding, mistrafficking and constitutive activity as underlying basis for
photoreceptor cell death (Sung et al., 1993; Kaushal and Khorana, 1994; DeCaluwe
and DeGrip, 1996; Deretic, 1998; Deretic et al., 1998; Mendes et al., 2005). Yet, a
functionally distinct group of mutations is suggested by the Drosophila visual system,
wherein light-dependent formation of stable rhodopsin/arrestin complex was
implicated in photoreceptor cell death (Alloway et al., 2000; Kiselev et al., 2000;
Iakhine et al., 2004). In Drosophila, preventing the formation of rhodopsin/arrestin
complex circumvents the degeneration phenotype. Whether this pathway is
conserved in the vertebrate retina is not known and cannot be assumed in view of
substantial differences that exist between animals in these two phyla with respect to
26

the light-induced modes of cell death, the arrangement of cellular compartments, and
the phototransduction signaling pathways. Nevertheless, some indirect evidence
suggests that rhodopsin/arrestin complex may be a pathogenic mechanism for certain
ADRP in human patients as well. For example, disruption of opsin transport by
conditional knockout of kinesin-II subunit KIF3A in mouse photoreceptor cells leads
to accumulation of both opsin and arrestin in the inner segment compartment of rod
cells prior to cell death (Marszalek et al., 2000). In addition, certain mutations that
affect the R135 residue of opsin lead to formation of opsin/arrestin complex and
disrupted receptor-mediated endocytic functions in a cell culture system (Chuang et
al., 2004). Another opsin mutant, K296E, was thought to cause ADRP by
constitutive stimulation of phototransduction, a notion supported by an in vitro
finding that recombinantly expressed K296E catalyzed GTP loading in transducin in
a light-independent manner (Robinson et al., 1992) However, in a transgenic mouse
model, K296E was phosphorylated and bound with arrestin in the rod photoreceptor
cells, and purified outer segments from these mice did not activate transducin in vitro
until the samples were stripped of arrestin and dephosphorylated (Li et al., 1995).
Thus, the basis for photoreceptor cell death mediated by K296E had heretofore
remained unknown.
Testing for a toxic effect of rhodopsin/arrestin complex in the vertebrate
visual system requires an experimental paradigm that allows us to manipulate and
control the formation of such complexes and to observe its effect on retinal
degeneration. Toward this end, we utilized transgenic mice expressing the K296E
27

opsin mutation. Formation of the rhodopsin/arrestin complex can be prevented by
breeding the K296E transgene into the arr1-/- background (Figure 2.1). However, in
the absence of arrestin, the constitutive activity of K296E may become unmasked,
thereby promoting photoreceptor cell death through another mechanism: persistent
stimulation of the phototransduction cascade (Fain and Lisman, 1993; Chen et al.,
1999b{Fain, 1999 #184; Fain and Lisman, 1999; Fain, 2006). Therefore, the
K296E
arr1-/-
mice were further crossed into the rod transducin α subunit knockout
(Tr-/-) background (Calvert et al., 2000) (Figure 1.1). In this study, we showed that
the retinal morphology of K296E
arr1-/-Tr-/-
mice improved significantly as compared
with age-matched K296E mice. Furthermore, we provide evidence that K296E is
hyperphosphorylated, and have discovered that its interaction with arrestin in the
inner segment causes its mislocalization to that cellular compartment. These results
provide direct evidence that rhodopsin/arrestin complex is toxic to vertebrate rod
photoreceptor cells. Moreover, they suggest that mislocalization of
rhodopsin/arrestin complex to the inner segment plays a role in initiating the
signaling pathway that leads to cell death.

2.2. Materials and Methods
2.2.1. Generation of Mouse Lines.
All experimental procedures were performed in accordance with regulations
established by the National Institutes of Health, as well as with the Society for
Neuroscience Policy on Animal Use in Neuroscience Research. K296E
28

Figure 2.1. Experimental scheme leading to rescue of K296E-induced retinal
degeneration.

29

transgenic mice that expressed human K296E mutant opsin were obtained from Dr.
Tiansen Li (Li et al., 1995). K296E transgenic mice were crossed with arr1-/-, Tr-/-

and arr1-/-Tr-/-

mice, respectively, to obtain K296E
arr1-/-
, K296E
Tr-/-
and K296E
arr1-/-
Tr-/-
mice. K296E-negative littermates in the respective genetic backgrounds were
used as age-matched controls. Unless clearly stated, all mice were born and raised in
darkness in order to avoid light-dependent retinal degeneration (Chen et al., 1999b;
Hao et al., 2002). K296E was also crossed with rhodopsin-/- mice to increase the
proportion of K296E protein in rod photoreceptor cells (Lem et al., 1999).

2.2.2. Retinal Morphometry.
Mice were sacrificed under infrared light. The superior pole of the cornea
was cauterized for orientation before enucleation. Eyecups were dissected,
embedded into epoxy resin and sectioned along the vertical meridian, as previously
described (Concepcion et al., 2002). Thickness of the outer nuclear layer (ONL) was
measured at 20 positions equally spaced along the retina (10 positions each in the
superior and inferior hemispheres). For each position, three measurements were
taken, and the average value of these three measurements was recorded.
Measurements were made using a camera lucida connected to a light microscope, a
WACOM graphics tablet (Vancouver, WA) and AxioVision LE Rel. 4.1. software
(Zeiss Co., Goettingen, Germany). Prior to each measurement session, the setup was
calibrated using a stage micrometer (Klarmann Rulings, Litchfield, NH).

30

2.2.3. Immunocytochemistry.
Dark adapted thirty-day-old K296E and K296E
arr1-/-
mice were treated with 0.5%
tropicamide and 2.5% phenylephrine hydrochloride to dilate the pupils prior to
exposure to diffuse white light (2000 lux intensity) for 15 min. Eyecups were then
fixed, infiltrated with sucrose, and 10 µm frozen sections were obtained as described
(Concepcion et al., 2002). Mouse monoclonal antibodies A11-82P and R2-12N
(gifts from Dr. P. A. Hargrave, University of Florida, Gainesville, FL) recognized
phosphorylated opsin and the amino-terminus of opsin, respectively. A rabbit
polyclonal antibody raised against the C10C10 epitope was used to visualize rod
arrestin (Mendez et al., 2003). Images were acquired on an Axioplan2 microscope
(Zeiss Co.). All images for each section were taken at the same detection gain.

2.2.4. Western Blot Analysis.
a) K296E phorphorylation assay. Thirty-day-old K296E transgenic mice and
their transgene-negative littermates were dark-adapted overnight. Retinas were
dissected under infrared light and exposed to 515-650 nm of light from a 100 W
quartz tungsten halogen lamp (Oriel Instruments, Stratford, CT). The number of
photons delivered was measured using a calibrated photodiode (United Detector
Technology Sensors, Inc., Hawthorne, CA), and light delivery was controlled by
neutral density filters (Oriel Instruments) and an electromagnetic shutter (Vincent
Associates, Rochester, NY). Retinas were exposed to light that caused 1%, 5%, 10%,
20% bleach, or kept in the dark. Rhodopsin phosphorylation was allowed to proceed
31

for an additional 10 min in the dark before the reaction was terminated by freezing in
liquid N
2
. Frozen retinas were then homogenized in 100 µl buffer (80 mM Tris, pH
8.0, 4 mM MgCl
2
) containing protease inhibitor cocktail at 1 tablet per 10 ml buffer
(Roche Diagnostics, Indianapolis, IN). DNase I (30 units, Roche Diagnostics) was
added and incubated at room temperature for 30 min. An equal amount of retinal
homogenate from each sample was loaded onto a 12% Bis-Tris SDS-PAGE gel
(Invitrogen Corp., Carlsbad, CA). Proteins were transferred onto nitrocellulose
membrane and incubated with A11-82P or R2-12N and visualized using enhanced
chemiluminescence (GE Healthcare, Piscataway, CA). b) Rhodopsin/arrestin
complex stability assay. The samples were prepared following the procedures
described previously (Li et al., 1995) and blotted with rabbit polyclonal arrestin
antibody raised against the C10C10 epitope.

2.2.5. GTP
γ
S Assay.
Retinas from arr1-/- or K296E
arr1-/-
mice were dissected under infrared light.
Ten retinas were pooled per tube and outer segments were isolated as described
(Tsang et al., 1998). Rhodopsin was quantified by Pierce’s BCA assay kit.
Transducin GTP
γ
S loading assays were performed as described previously (Robinson
et al., 1992).

32

2.2.6. Electroretinogram (ERG).
Mice were dark-adapted overnight (>12 h) and processed under infrared light.
The mice were anesthetized with an intraperitoneal injection of xylazine (10 µg/g
body weight) and ketamine (100 µg/g body weight) mixture. Pupils were dilated
with 0.5% tropicamide and 2.5% phenylephrine hydrochloride. A drop of
hydroxypropyl methylcellulose solution (Akorn Co., Buffalo Grove, IL) was placed
on the cornea to keep the eye moist and to establish electrical contact with the
corneal electrode. A steel needle reference electrode was placed subcutaneously
below the eye. The flash intensity and its spectral composition were controlled with
neutral density and narrow bandpass interference filters (500 nm ± 10 nm FWHM).
ERG signals were amplified by an AC/DC differential amplifier (A-M Systems, Inc.,
Carlsborg, WA), bandpass-filtered at 0.1-1000 Hz, sampled at 2000 Hz, and acquired
with a Digidata 1322A data acquisition board (Axon Instruments, Union City, CA)
using pClamp software (Axon Instruments). Sensitivity measurements were based
on normalized b-wave amplitudes. To smooth the b-wave traces, a Gaussian filter
with a bandwidth of 16.6 Hz at 3dB cutoff was applied as described (Lyubarsky et al.,
1999).

2.2.7. Sample Preparation for Liquid Chromatography Mass Spectrometry
(LC-MS) Analysis.
LC coupled to MS (LC-MS) and tandem mass spectrometry (LC-MS/MS)
has been an efficient technique for analyzing complex peptide mixtures to determine
33

composition and relative abundance. Sample preparation and mass spectrometry
analysis of rhodopsin phosphorylation was performed as described previously
(Kennedy et al., 2001; Shi et al., 2005b)). Synthetic carboxyl-terminal peptides
corresponding to mouse and human rhodopsin were used to determine their elution
profile and ionization efficiency. Phosphates were removed from rhodopsin by calf
intestine phosphatase treatment of retinal membranes from K296E
arr1-/-
and K296E
mice prior to LC-MS analysis.

2.3. Results
K296E transgenic mice were crossed into arr1-/-, Tr-/- and arr1-/-Tr-/-
backgrounds in order to test the hypothesis that persistent rhodopsin/arrestin
complex is toxic to vertebrate photoreceptor cells. Unless stated otherwise, all mice
were born and raised in darkness to eliminate the possible effect of light exposure on
retinal morphology. All mice were in a pigmented background. Dark-reared
K296E-negative control mice (arr1-/-, Tr-/- and arr1-/-Tr-/-) exhibited retinal
morphology that was indistinguishable from wildtype mice during the time course of
our study (Figure 2.2.A, E, I, M). Thus, the absence of arrestin, transducin, or both,
had no discernible impact on retinal morphology.

2.3.1. K296E-induced Retinal Degeneration Is Transducin-independent.
The ratio of K296E transcript to endogenous opsin mRNA had previously
been established as 0.25:1 in the line of transgenic mice used in this study (K296E-A)
34

Figure 2.2. Retinal morphology of K296E-induced retinal degeneration. Dark-reared K296E-negative
littermates in the arr1-/-, Tr-/- and arr1-/-Tr-/- genetic backgrounds displayed retinal morphology
similar to that of wildtype mice at 10 weeks of age (A, E, I, M). (B-D) Time course of retinal
degeneration in K296E mice. (F-H) Similar time course and morphological appearance were
observed in K296E
Tr-/-
mice. (J-L) While the rate of degeneration appeared similar, the outer segment
length was noticeably shorter when K296E was expressed in the arr1-/- background. (N-P) K296E-
induced retinal degeneration was noticeably slowed in the arr1-/-Tr-/- background. Scale bar = 25 µm.
35

(Li et al., 1995). This low expression level precludes overexpression of opsin as an
underlying cause of photoreceptor cell death (Olsson et al., 1992; Li et al., 1996).
Consistent with the previous report (Li et al., 1995), the K296E-A transgenic mice
showed a slow time course of progressive light-independent retinal degeneration
(Figure 2.2.B-D). Thinning of the ONL due to death of photoreceptor cells was
apparent at eight weeks of age. At ten weeks, the ONL was reduced by ~40% as
compared with the transgene-negative littermates. Slight disorganization of the outer
segment structure was also apparent, but the overall length of the outer segment was
not altered. The rate of degeneration increased when K296E was expressed in the
rhodopsin +/- background, which lends further support for the causative role of
K296E, rather than opsin overexpression, in promoting photoreceptor cell death
(Figure 2.3).
The time course of retinal degeneration and the appearance of retinal
morphology in K296E
Tr-/-
mice was indistinguishable from K296E
Tr+/+
(Figure 2.2.
F-H). These data confirm the conclusion reached by Li and colleagues that the
K296E mutation does not cause retinal degeneration through constitutive activation
of phototransduction (Li et al., 1995). Rather, the data is consistent with the current
hypothesis that formation of K296E/arrestin complex is the underlying basis for
photoreceptor cell death in this transgenic mouse model.

36

Figure 2.3. Retinal degeneration was more rapid when the proportion of K296E was
increased in the rhodopsin +/- background.

37

2.3.2. K296E Activates Transducin and Causes Retinal Degeneration in the
arr1-/- Background.
K296E transgenic mice were crossed into the arr1-/- background in order to
eliminate the formation of K296E/arrestin complex. As predicted, however, retinal
degeneration was also observed in K296E
arr1-/-
mice (Figure 2.2.J-L). Interestingly,
although the rate of degeneration was similar to that of K296E and K296E
Tr-/-
mice,
K296E
arr1-/-
retinas showed a different degeneration pattern; i.e., the outer segments
were noticeably shorter and more disorganized, and resembled the morphology of
light-damaged retinas of albino rats or pigmented arr1-/- mice exposed to low levels
of light (Li et al., 1995; Schremser and Williams, 1995a; Chen et al., 1999b),
suggesting that K296E activated transducin in the arrestin knockout background. To
test this notion, we sought to determine whether outer segment preparations from
K296E
arr1-/-
mice exhibited light-independent GTP γS loading of transducin in vitro.
Figure 2.4.A shows transducin activation by rod outer segment membranes isolated
from arr1-/- and K296E
arr1-/-
mice. As expected, both samples activated transducin in
a light-dependent manner due to the presence of endogenous rhodopsin. However,
no difference was seen between arr1-/- and K296E
arr1-/-
samples in the dark. Notably,
Li et al. observed light-independent GTP
γ
S loading of transducin by K296E only
after removal of arrestin and phosphatase treatment (Li et al., 1995; Schremser and
Williams, 1995a; Chen et al., 1999b). Thus, it is likely that phosphorylation greatly
reduced the constitutive activity of K296E (see also below).

38

Figure 2.4. In vitro and in vivo analyses of K296E catalytic activity in the arr1-/- background.
(A) GTP γS loading assay catalyzed by rod outer segment preparations from arr1-/- and
K296E
arr1-/-
retinas that contained 10 nM rhodopsin. Although catalytic activity of K296E
was expected to result in an increased rate of GTP γS loading in the dark-adapted samples, no
such activity was observed. (B) ERG recorded from five-week-old arr1-/- mice and their
K296E
arr1-/-
littermates. Five-week-old Tr-/- mice were used to illustrate the cone threshold.
Sensitivity was presented as normalized b-wave amplitude vs. light intensity.
39

We then utilized the electroretinogram (ERG) as an alternative method to
detect light-independent activity of K296E in the arr1-/- retina. If K296E activated
transducin in the arr1-/- background, then retinas from dark-reared K296E
arr1-/-
mice
should have behaved as if light-adapted and would have exhibited an elevated light
threshold. To determine whether this was the case, ERG responses were compared
between dark-adapted K296E
arr1-/-
mice and K296E-negative arr1-/- littermate
controls. Tr-/- mice were also included in the comparison to demarcate the cone
threshold. Figure 2.4.B shows that K296E
arr1-/-
mice displayed a ~100-fold decrease
in flash sensitivity when compared to arr1-/- mice. However, this lowered sensitivity
was still one log unit above the cone threshold, indicating that K296E
arr1-/-
rods were
not saturated. These results support the notion that, in the absence of arrestin,
K296E constitutively activated transducin, thereby acting as a source of “dark” light
to cause photoreceptor cell death in K296E
arr1-/-
mice.

2.3.3. K296E-induced Retinal Degeneration Is Slowed in the Absence of
Arrestin and Transducin.
K296E
arr1-/-
mice were further crossed into the Tr-/- background to prevent
retinal degeneration induced by persistent stimulation of phototransduction.
Remarkably, K296E
arr1-/-Tr-/-
mice exhibited substantially improved retinal
morphology when compared to age-matched K296E, K296E
arr1-/-
or K296E
Tr-/-
mice.
The number of photoreceptor cells, as reflected by the thickness of the ONL, was
similar to that of the transgene-negative controls, as was the length and the organized
40

structure of the outer segment (Figure 2.2.N-P). To better quantify the degree of
rescue, ONL thickness was measured at 20 positions, spaced apart equally, that
spanned the vertical meridian of the eye. As can be seen in Figure 2.5, the ONL
thickness of K296E
arr1-/-Tr-/-
retinas was similar to that of the transgene-negative
controls at the periphery of the retina, but the ONL was noticeably thinner at the
central region of the retina. Nevertheless, the arr1-/-Tr-/- background provided a
higher degree of rescue from K296E-induced retinal degeneration when compared
with K296E in the single knockout backgrounds (Figure 2.2 and Figure 2.5). These
results are consistent with the hypothesis that accumulation of stable
rhodopsin/arrestin complex is indeed toxic to mammalian photoreceptor cells.
However, preventing this complex formation was not sufficient to completely rescue
K296E-induced retinal degeneration.

2.3.4. Constitutive Activity of K296E Causes Retinal Degeneration in arr1-/-Tr
-/- Retinas.
We further explored the possibility that the constitutive activity of K296E is
the underlying cause of photoreceptor cell death observed in dark-reared K296E
arr1-/-
Tr-/-
mice. To test this hypothesis, arr1-/-Tr-/- mice were exposed to constant light for
one week to examine whether light-activated rhodopsin (R*) had an effect similar to
K296E on the retinal morphology of arr1-/-Tr-/- mice. As can be seen in Figure 2.6,
the central region of the light-exposed arr1-/-Tr-/- retina was severely damaged.
This pattern of light damage to the central retina closely mirrored that seen in
41

Figure 2.5. Quantification of K296E-induced retinal degeneration. For each group
(N = 4 to 8), ONL thickness was measured at 20 positions, equally spaced along the
vertical meridian of the retina. Three measurements were taken for each position,
and an average value was recorded. Each point represents mean ± SD obtained for
each group. Position 0 corresponds to optic nerve head. I, inferior; S, superior.

42

Figure 2.6. Pattern of retinal degeneration in dark-reared K296E
arr1-/-Tr-/-
mice is
recapitulated in light exposure of arr1-/-Tr-/- mice. Representative retinal sections
obtained from arr1-/-Tr-/- mice (pupils undilated) exposed to constant illumination
of 2000 lux for one week. A. Low magnification view of the whole retina. B. Higher
magnification of the boxed area shown in A. C. Morphometric measurements of the
outer nuclear layer thickness (N=3, error bars represent SD). S, superior (-10); I,
inferior (10). Scale bar = 50 µm.

43

K296E
arr1-/-Tr-/-
retinas, and was unlike the pattern seen in other rodent models of light
damage where sensitivity to light damage was exhibited by either the superior or the
inferior region of the retina (LaVail et al., 1987; Hao et al., 2002; Roca et al., 2004).
Thus, R*, as well as K296E, is able to signal through a transducin- and arrestin-
independent pathway to induce retinal degeneration.

2.3.5. K296E Is Colocalized with Arrestin in the Retina.
Immunocytochemistry was performed to verify that K296E colocalizes with
arrestin (Figure 2.7). Rhodopsin, in the active conformation, is a substrate for
rhodopsin kinase, which places multiple phosphate groups at a cluster of Ser and Thr
sites at rhodopsin’s carboxyl-terminus (Wilden and Kuhn, 1982; Palczewski et al.,
1988b; Palczewski and Benovic, 1991). Activated, phosphorylated rhodopsin is then
capped by arrestin binding (Wilden et al., 1986; Xu et al., 1997). Thus, arrestin is
expected to colocalize with R* in a light-dependent manner and K296E in a light-
independent manner, inasmuch as K296E is in an active conformation regardless of
light. A monoclonal antibody, A11-82P, binds strongly to multiply phosphorylated
rhodopsin, but not to unphosphorylated rhodopsin (Adamus et al., 1988). As shown
in Figure 2.7, A11-82P reactivity was observed only in the rod outer segment of
light-exposed wildtype retina where phosphorylated R* is localized. In contrast,
A11-82P reactivity was observed in both the dark-adapted and the light-adapted
retinas of K296E mice, indicating that K296E is constitutively phosphorylated. It is
also noteworthy that the A11-82P staining pattern in the wildtype retina is restricted
44

Figure 2.7. K296E was phosphorylated in a light-independent manner and
colocalized with arrestin in the photoreceptor cell layer in darkness and in light.
Wildtype and K296E mice were dark-adapted (A-C and G-I, respectively) or
exposed to light (D-F and J-L, respectively). Phosphor-opsin was visualized by
A11-82P, a mouse monoclonal antibody that preferentially binds multiple
phosphorylated species of rhodopsin (A, D, G and J). Retinal sections reacted with
the antibody against arrestin are shown in B, E, H and K. Merged images of A11-
82P and arrestin are shown in C, F, I and L. Scale bar = 25 µm.

45

to the outer segment where rhodopsin is localized, whereas, in the K296E retina, the
A11-82P reactivity is seen throughout the photoreceptor layer, indicating
mislocalization of phosphorylated K296E. Thus, K296E, being in an active
conformation, is a substrate for rhodopsin kinase in all compartments of the rod
photoreceptor cell, even in the absence of light.
Arrestin is known to exhibit a light-dependent distribution pattern in the
retina (Broekhuyse et al., 1985; Broekhuyse et al., 1987; Mangini and Pepperberg,
1988; Whelan and McGinnis, 1988; Peterson et al., 2003). In the dark, arrestin is
largely excluded from the outer segment compartment and resides primarily in the
inner segment and ONL (Figure 2.7.B). Upon light exposure, arrestin moves to the
outer segment (Figure 2.7.E). This light-dependent distribution of arrestin was not
seen in the K296E retina. Regardless of lighting conditions, arrestin appeared to be
distributed throughout the photoreceptor cell layer, mirroring the pattern of
phosphorylated K296E (Figure 2.7.H and K). The colocalization of arrestin with
phosphorylated K296E is consistent with the notion that they exist as a complex in
the rod photoreceptor cell.

2.3.6. K296E Is Retained in the Inner Segment and ONL through Its Interaction
with Arrestin.
We explored the cause of K296E mislocalization to the inner segment and the
ONL. It is known that the carboxyl-terminal domain of rhodopsin contains a
trafficking signal that is necessary and sufficient to direct polarized transport of
46

rhodopsin to the outer segment (Sung et al., 1994; Deretic, 1998; Deretic et al., 1998;
Tam et al., 2000; Concepcion et al., 2002). Immediately preceding this domain is a
cluster of Ser and Thr residues that, upon phosphorylation, participate in converting
arrestin from its latent, inactive state into a conformation that binds light-activated,
phosphorylated rhodopsin with high affinity (Hirsch et al., 1999; Vishnivetskiy et al.,
2000; Gurevich and Gurevich, 2004). Therefore, it is plausible that the binding of
arrestin may have masked the carboxyl-terminal domain of K296E that is normally
required for rhodopsin to traffic to the outer segment. Another possibility is that a
proportion of K296E is misfolded, inasmuch as misfolded rhodopsin is also retained
in the inner segment and becomes mislocalized (Olsson et al., 1992), and causes cell
death through protein aggregation and disruption of protein degradation (Kopito,
2000; Rajan et al., 2001; Illing et al., 2002). To distinguish between these two
possibilities, localization of rhodopsin and K296E was investigated in K296E
arr1-/-

retinas using an antibody against the amino-terminus of rhodopsin, R2-12N (Figure
2.8). In the wildtype retina, rhodopsin is localized predominantly to the membranous
outer segment. In the K296E retinas, however, rhodopsin immunoreactivity was
seen throughout the photoreceptor cell layer, a result similar to that obtained with the
anti-phosphorylated opsin antibody, A11-82P (Figure 2.8B). Remarkably, in the
absence of arrestin, K296E mutant opsin reactivity was shifted to the outer segment
(Figure 2.8D). These results suggest that K296E is correctly folded, but is retained
in the inner segment and ONL through its interaction with arrestin.

47

Figure 2.8. K296E was retained in the inner segment and ONL via its interaction
with arrestin. Rhodopsin was visualized using R2-12N, a mouse monoclonal
antibody that recognizes the amino-terminal 2-12 residues of rhodopsin (Adamus et
al., 1991). Rhodopsin localized to the outer segment in wildtype and arr1-/- retinas
(A, C). In the retina expressing K296E, rhodopsin reactivity was seen throughout
the retinal layers (B). In the K296E
arr1-/-
retina, all rhodopsin reactivity was seen in
the outer segment (D). OS, outer segment; IS, inner segment; ONL, outer nuclear
layer. Scale bar = 25 µm.

48

2.3.7. Hyperphosphorylation of K296E May Contribute to Stable Complex
Formation with Arrestin.
The results shown in Figure 2.4.B indicate that the rod responses from
K296E
arr1-/-
mice were not saturated. This result is somewhat surprising since the
level of K296E transcript was estimated to be 25% of total rhodopsin transcripts (Li
et al., 1995). This level of K296E protein expression would be equivalent to a
steady-state light exposure that bleached 25% of rhodopsin at any given time.
However, it is known that primate and mouse rods, saturation is reached upon ~400
R*·s
-1
to ~4000 R*·s
-1
, or ~0.001 – 0.01% bleach·s
-1
, respectively, under steady
background light (Baylor et al., 1984; Mendez et al., 2001; Makino et al., 2004),
which is a value substantially lower than 25% bleach·s
-1
. We quantified the level of
K296E protein using a mass spectrometry-based method (Kennedy et al., 2001; Shi
et al., 2005b) to directly assess the level of K296E protein. Shown in Figure 2.9 are
representative ion chromatograms of carboxyl-terminal peptides that correspond to
mouse rhodopsin and K296E released by endopeptidase Asp-N treatment of
K296E
RK-/-
retinal membranes. K296E
RK-/-
retina was used to ensure that neither
K296E nor rhodopsin would be phosphorylated, thus simplifying quantification.
Using this LC-MS method, we estimated the proportion of K296E to endogenous
mouse rhodopsin in the following genetic backgrounds (mean ± SD, N=3 for each
condition): K296E
RK-/-
, 1.04% ± 0.06%; K296E
arr1-/-
, 1.33% ± 0.03%; K296E, 4.5%
± 0.8%. Phosphates were removed by phosphatase treatment in the latter two
samples prior to LC-MS analysis. The discrepancy between the values obtained
49

Figure 2.9. Tandem liquid chromatography-mass spectrometric quantification of
K296E expression in mouse retinal homogenates. A. Ion chromatograms of eluted
synthetic peptides from reverse-phase HPLC corresponding to the terminal 19 C-
terminal residues of mouse rhodopsin and human opsin mutant K296E. When equal
amount of standard peptides were loaded, the relative detection efficiency of
K296E:endogenous rhodopsin was established to be 1.14:1. B. Representative ion
chromatogram of peptides released from Asp-N digestion of retinal membranes
obtained from one K296E
RK-/-
retina. After correcting for the ionization efficiency,
K296E expression level in this retina is estimated to be 1.08%.

50

from K296E, K296E
arr1-/-
and K296E
RK-/-
samples may reflect the shortened outer
segments observed when K296E is expressed in the arr1-/- or RK-/- genetic
backgrounds. These results indicate that the proportion of the K296E protein to
endogenous mouse rhodopsin was much lower than the value obtained by transcript
analysis.
A comparison between single photon responses obtained from RK-/- rods
(Chen et al., 1999a) and arr1-/- rods (Xu et al., 1997) indicates that phosphorylation
alone decreases the catalytic activity of R* by ~70%. Thus, an expression level of
1% K296E protein in the arr1-/- background is still expected to exceed the rod
saturation threshold of ~0.001% bleach·s
-1
even if phosphorylation decreased the
catalytic activity of K296E. However, it is known that heavily phosphorylated
rhodopsin exhibits greatly reduced catalytic activity in vitro (Arshavsky et al., 1985;
Wilden, 1995). For this reason, we used the A11-82P antibody, which preferentially
binds highly phosphorylated species of rhodopsin (Adamus et al., 1988), to compare
the extent of phosphorylation in K296E and in rhodopsin exposed to calibrated light
that produced 1%, 5%, 10% and 20% bleach. As can be seen in Figure 2.10.A, the
signal from retinas that expressed 4.5% K296E exceeded that of control retinas
exposed to 20% bleach, suggesting that K296E is phosphorylated to a greater extent
than is wildtype rhodopsin. Thus, hyperphosphorylation is likely the reason for our
inability to detect K296E-catalyzed GTP- γS loading of transducin in vitro (Figure
2.4.A) and its inability to saturate rod response in arr1-/- mice (Figure 2.4.B). To
compare the relative stability of K296E/arrestin complex to R*/arrestin complex,
51

Figure 2.10. K296E was highly phosphorylated and formed a stable complex with
arrestin. (A) Retinal homogenates from dark-adapted K296E retina and retinas
isolated from wildtype mice exposed to calibrated light that produced an estimated
0%, 1%, 5%, 10% and 20% bleach. Highly phosphorylated opsin was detected by
A11-82P antibody. Remarkably, K296E samples, with an estimated expression level
of 4.5 ± 0.8%, showed a much stronger signal than did wildtype retinas exposed to
20% bleach. (B) Membrane fraction from retinal homogenates obtained from dark-
adapted or light-exposed mice was subjected to hypotonic wash followed by urea
wash. The most stringent urea wash stripped most of the arrestin from the light-
exposed retina of the wildtype sample, but not from the K296E sample.

52

pellet fractions from retinal homogenates were subjected to sequential, increasingly
stringent washes (Figure 2.10.B). Similar amounts of arrestin were present in the
original pellet fractions of dark- or light-adapted retinas (Figure 2.10.B, unwashed).
After a hypotonic wash, the majority of arrestin was removed from the dark-adapted
wildtype sample but not from the K296E sample, while, in the light-adapted samples,
comparable amounts of arrestin were retained. Following the more stringent urea
wash, the remaining bound arrestin was removed from the light-treated wildtype
sample but not from the K296E sample. Thus, the K296E/arrestin complex appears
to be more stable than the R*/arrestin complex formed as a consequence of normal
light exposure. This increased stability may be attributed to the observed
hyperphosphorylation of K296E.

2.4. Discussion
The amelioration of K296E-induced retinal degeneration in the
arrestin/transducin double knockout provide direct evidence that persistent
rhodopsin/arrestin complex is a stimulus for photoreceptor cell death in the
vertebrate visual system. Thus, this retinal degeneration pathway appears to be
conserved from the invertebrate to the vertebrate visual system. In Drosophila,
endocytosis of rhodopsin/arrestin complex is a required step in this signaling
pathway. Further investigation is necessary to determine whether endocytosis is a
required step leading to cell death in the vertebrate retina.
53

It is important to note that certain naturally occurring mutations affecting
rhodopsin’s carboxyl-terminus, such as Q344ter and P347S, perturb rhodopsin
trafficking to the outer segment. The ensuing mislocalization is thought to be the
underlying basis for ADRP in these instances (Sung et al., 1994; Li et al., 1996;
Deretic, 1998; Deretic et al., 1998). In a previous study to investigate the effect of
rhodopsin mislocalization on retinal degeneration, we observed that mislocalization
of a trafficking defective mutant, S334ter, did not lead to retinal degeneration in a
transgenic mouse model when the protein was expressed at 10% of total rhodopsin
levels (Concepcion et al., 2002). Thus it appears unlikely that K296E, when
expressed at 4.5%, exerted its toxicity through mislocalization alone.
Inasmuch as arrestin binding to R* is a step essential to rhodopsin
deactivation, it can be expected that high levels of rhodopsin/arrestin complex are
formed in the rod outer segment during the course of bright light exposure. Unlike
the Drosophila photoreceptors, the outer segment compartment of vertebrate rods is
physically sequestered from the rest of the cell body via a thin connecting cilium and
appears to lack space for endocytic machinery; indeed, endocytosis has not been
observed in this compartment. How, then, does K296E/arrestin complex cause
photoreceptor cell death? We speculate that the cause is the inappropriate
accumulation of rhodopsin/arrestin complex in the inner segment compartments of
the rod cell where endocytosis is supported. Importantly, we show that K296E is
retained in the proximal compartments, where it appears to form a stable complex
with arrestin.
54

The 11-cis retinal chromophore is covalently attached to K296 of the opsin
protein through a protonated Schiff base linkage (Zhukovsky et al., 1991).
Mutations affecting K296 render the opsin protein incapable of binding retinal, and
disrupt the interaction between K296 and E113 that normally constrains the opsin
protein to an inactive conformation (Robinson et al., 1992). Hence, K296E exists in
an active conformation the moment it is synthesized in the inner segment, and is
phosphorylated and bound to arrestin in this compartment. The K296E/arrestin
complex appears to be stable, inasmuch as the ability of K296E to activate
transducin is silenced by arrestin binding. It appears that hyperphosphorylation of
K296E may be the underlying cause for this increased stability of K296E/arrestin
complex as well as its greatly decreased ability to activate transducin in the arrestin
knockout background. Further evidence in support of a stable complex derives from
our observation that transphosphorylation of non-activated rhodopsin was observed
only when K296 was expressed in the arrestin knockout background; when arrestin
was present, K296E lost its ability to activate rhodopsin kinase to phosphorylate
nearby, non-activated rhodopsin molecules (Shi et al., 2005b). In addition, ERG
measurements showed no difference in sensitivity between wildtype mice and
K296E mice (data not shown). These data support the notion that stable
rhodopsin/arrestin complex accumulated in the inner segment initiates a transducin-
independent signaling cascade that leads to photoreceptor cell death.
Formation of excessive rhodopsin/arrestin complex in the inner segment is
likely to occur as a result of other mutations. For example, rhodopsin R135L
55

mutants appear to be phosphorylated by rhodopsin kinase and bound to arrestin
in vitro (Shi et al., 1998). When the R135L mutant is introduced into murine retina
by in vivo electroporation, it appears to cause arrestin to accumulate around the cell
body, suggesting that R135L/arrestin complex may form within these cellular
compartments (Chuang et al., 2004). In addition, other naturally occurring mutations
that affect the trafficking domain on rhodopsin’s carboxyl-terminus will likely result
in accumulation of rhodopsin in the inner segment and ONL (Sung et al., 1994; Li et
al., 1996). Inasmuch as these mutations often do not affect phosphorylation or
arrestin binding, it is plausible that light activation of mislocalized mutant rhodopsin
leads to excessive rhodopsin/arrestin complex in these cellular compartments. By
the same token, defects in other components that affect normal rhodopsin transport to
the outer segment (such as the aforementioned KIF3A mutation) will have a similar
effect. Thus, excessive accumulation of rhodopsin/arrestin complex may define a
broader class of mutations that lead to blindness.
Drosophila expresses two visual arrestins where Arr2 is more abundant and
plays the major role in deactivating rhodopsin. Under normal light exposure, an
increase in cytoplasmic Ca
2+
concentration leads to Arr2 phosphorylation by
Ca
2+
/calmodulin-dependent protein kinase II (Kahn and Matsumoto, 1997).
Phosphorylation of Arr2 destabilizes its binding to rhodopsin, thereby preventing
accumulation of rhodopsin/Arr2 complex. In addition, Arr1 appears to play an
essential role in normal, light-induced rhodopsin endocytosis as well as in preventing
accumulation of toxic rhodopsin/Arr2 complex; when Arr1 is absent, retinal
56

degeneration ensues (Satoh and Ready, 2005). Thus, it appears that multiple
mechanisms regulate the accumulation of rhodopsin/Arr2 complex formation during
normal light exposure in the Drosophila photoreceptor cell. The mammalian rod
photoreceptor cells express two isoforms of rod arrestin encoded by one gene: a 48
kD full-length arrestin and a less abundant splice variant that is truncated at the
carboxyl-terminus (Palczewski et al., 1994; Smith et al., 1994; Smith, 1996). While
the splice variant resembles Drosophila Arr1, it does not appear to play a part in
mediating light-dependent rhodopsin endocytosis, inasmuch as expression of the
full-length protein alone in the arrestin knockout mice had little effect on retinal
morphology (Burns et al., 2006).
Retinal degeneration was not completely suppressed upon placement of
K296E in the arrestin/transducin double knockout background. The residual
degeneration appears to be caused by the activity of K296E, inasmuch as light
exposure recapitulated the same pattern of degeneration in arr1-/-Tr-/- retinas. Low
light induced retinal damage in pigmented mice caused by persistent
phototransduction should have been blocked in the transducin knockout background.
However, an earlier study showed that not all mice deficient in transducin are
resistant to this mechanism of light damage, suggesting a genetic modifier effect
(Hao et al., 2002). The nature of this signaling cascade leading to photoreceptor cell
death is not yet known. It is possible that rhodopsin signaling through other G-
proteins, Gi or Go, may be the underlying basis of this degeneration. Future studies
will be required to fully understand the complex effect of light on retinal physiology.

57

Acknowledgements
We wish to express our gratitude to Dr. Juan Korenbrot for his help in
assembling our electroretinogram (ERG) apparatus, and in the experimental design
and data analysis pertaining to ERG. We thank Dr. Tiansen Li for kindly providing
the K296E transgenic mice and Dr. Paul Hargrave for generous gifts of the
rhodopsin antibodies A11-82P and R2-12N. This work was supported by National
Institutes of Health (NIH) Grant EY12155 and the Arnold & Mabel Beckman
Macular Research Center.

58

Chapter 3*
Light Causes Phosphorylation of Non-Activated Visual Pigments
In Intact Mouse Rod Photoreceptor Cells

Abstract
Phosphorylation of G-protein coupled receptors (GPCRs) is a required step
in signal deactivation. Rhodopsin, a prototypical GPCR, exhibits high-gain
phosphorylation in vitro whereby hundred-fold molar excess of phosphates are
incorporated into the rhodopsin pool per molecule of activated rhodopsin. The
extent by which high-gain phosphorylation occurs in the intact mammalian
photoreceptor cell, and the molecular mechanism underlying this reaction in vivo,
are not known. Trans-phosphorylation is a mechanism proposed for high-gain
phosphorylation whereby rhodopsin kinase, upon phosphorylating the activated
receptor, continues to phosphorylate nearby non-activated rhodopsin. We used two
different transgenic mouse models to test whether trans-phosphorylation occurs in
the intact photoreceptor cell. The first transgenic model expressed a murine cone
pigment, S-opsin, together with the endogenous rhodopsin in the rod cell. We
showed that selective stimulation of rhodopsin also led to phosphorylation of S-
opsin. The second mouse model expressed the constitutively active human
rhodopsin mutant, K296E. K296E, in the arrestin -/- background, also led to

*This work was fulfilled with Dr. Guang Shi together, which has been published in Journal of
Biological Chemistry in 2005.
59

phosphorylation of endogenous mouse rhodopsin in the dark-adapted retina. Both
mouse models provide strong support of trans-phosphorylation as an underlying
mechanism of high-gain phosphorylation, and provide evidence that a substantial
fraction of non-activated visual pigments become phosphorylated through this
mechanism. Since activated, phosphorylated receptors exhibit decreased catalytic
activity, our results suggest that dephosphorylation would be an important step in
the full recovery of visual sensitivity during dark adaptation. These results may also
have implications for other GPCR signaling pathways.

3.1. Introduction
Visual pigments, such as rhodopsin and cone opsins, belong to a family of
G-protein-coupled receptors (GPCRs) that contain a cluster of Ser/Thr sites at their
carboxyl-termini. Visual pigments initiate G-protein signaling upon photon
absorption. As with other GPCRs, phosphorylation of the carboxyl-terminal Ser/Thr
sites, followed by arrestin binding, are required steps in signal deactivation (Burns
and Baylor, 2001). Rhodopsin phosphorylation is catalyzed by rhodopsin kinase
(GRK1, or RK), which is activated upon association with light-activated rhodopsin
(R*) in the MII conformation (Palczewski et al., 1988a; Palczewski et al., 1991).
In vitro and in vivo evidence shows that phosphorylated rhodopsin exhibits
diminished catalytic activity (Wilden et al., 1986; Chen et al., 1995b; Chen et al.,
1999a; Mendez et al., 2000), and that arrestin binding is required to fully terminate
R* signaling (Wilden et al., 1986; Xu et al., 1997).
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Since the discovery of light-activated rhodopsin phosphorylation, a number
of studies have reported that, in isolated rod outer segments, several hundred-fold
molar excess of phosphates are incorporated into the rhodopsin pool per mole of R*
(Aton, 1986; Binder et al., 1990; Chen et al., 1995a; Binder et al., 1996). Given that
each rhodopsin has been observed to incorporate only up to nine phosphates
(Wilden and Kuhn, 1982), the straightforward interpretation is that non-activated
rhodopsin molecules, which we designate here as R, are phosphorylated as well as
R*. This phenomenon has been termed high-gain phosphorylation (Binder et al.,
1990). Phosphorylation of non-activated rhodopsin has also been reported in
isolated frog retinas and in living frogs (Binder et al., 1996). In this system, ~1% of
R become phosphorylated following a 3% bleach (i.e., photon absorption by 3% of
rhodopsin), and, when living frogs are exposed to prolonged dim light, a higher
fraction of R (3%) becomes phosphorylated (Binder et al., 1996).
The mechanism by which R becomes phosphorylated in vivo is not known.
One possibility is that the phosphorylated MII decays to opsin and is reconstituted
with the 11-cis retinal to regenerate a visual pigment prior to dephosphorylation
(Biernbaum et al., 1991). Another possibility is trans-phosphorylation, which was
termed to describe a putative mechanism whereby rhodopsin kinase (RK), once
activated by associating with R*, phosphorylates a nearby R (Rim et al., 1997).
Evidence in support of this model includes the observation that R*-activated RK is
capable of phosphorylating an exogenously added peptide substrate (Palczewski et
al., 1988b; Palczewski et al., 1989; Brown et al., 1993; Dean and Akhtar, 1996). In
61

addition, structural studies on the carboxyl terminus of rhodopsin, using site-
directed spin labeling, revealed that it is disordered and highly mobile (Langen et al.,
1999). Therefore, given the high density of rhodopsin and its high diffusion rate on
disk membranes (Wey et al., 1981; Kaplan, 1984; Calvert et al., 2001), it is
plausible that carboxyl termini from neighboring R molecules would be accessible
to R*-activated RK.
A major challenge in attempting to demonstrate the presence of trans-
phosphorylation is the difficulty in unambiguously distinguishing between
phosphorylated R* and phosphorylated R in the same reaction mixture. As a direct
test of trans-phosphorylation, Rim and colleagues designed experimental protocols
based on a recombinantly expressed split receptor mutant of rhodopsin that was
assembled from two separately expressed fragments (Rim et al., 1997). This “split
rhodopsin” exhibits light-dependent phosphorylation. Importantly, it can be
distinguished from full-length rhodopsin in the same phosphorylation mixture by
virtue of its distinct electrophoretic mobility on a denaturing gel. Therefore, the
split rhodopsin can be exposed to light and mixed with the non-bleached full-length
rhodopsin, or vice versa, to test for the presence of phosphorylated R in the same
mixture that contains RK. However, despite numerous attempts using different
experimental configurations, this system was unable to provide evidence that non-
activated rhodopsin is trans-phosphorylated (Rim et al., 1997).
The experiments described above were performed using solubilized
receptors and receptors reconstituted into lipid vesicles. However, if the activated
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kinase needs to be physically associated with R*, or if it can diffuse only a short
distance prior to phosphorylating nearby non-activated R, then trans-
phosphorylation may proceed efficiently only on native disk membranes that
contain high concentrations of freely diffusible receptor molecules. To test this
hypothesis, we generated transgenic mice that expressed a shortwave-sensitive cone
pigment, S-opsin, in rod photoreceptors. This system allowed for preferential
activation of rhodopsin by long wavelength light when the two pigments are co-
expressed in the native rod disks, and phosphorylation of S-opsin would arise only
as a result of trans-phosphorylation. Furthermore, we used the rhodopsin K296E
transgenic mouse model, expressing human rhodopsin mutant K296E in mouse rod
photoreceptors, to provide additional evidence of the presence of trans-
phosphorylation.

3.2. Materials and Methods
3.2.1. S-opsin and K296E Transgenic Mouse Lines.
All mice were treated in accordance with the ARVO Statement for the Use of
Animals in Ophthalmic and Visual Research, as well as with USC IACUC
guidelines. S-opsin transgenic mice were mated with rhodopsin knockout mice
(Lem et al., 1999) to generate S-opsinrho+/- and S-opsinrho-/- mice. K296E
transgenic mice were bred with arrestin knockout mice to obtain K296Earr-/- mice
(Li et al., 1995; Xu et al., 1997). The S-opsin transgenic mice were born and raised
in a 12 hr/12 hr light/dark cycle. K296Earr-/- and arr-/- mice were born and raised
63

in constant darkness to prevent retinal degeneration resulting from constitutive
signaling (Xu et al., 1997).

3.2.2. Standard Peptides.
The non-phosphorylated and monophosphorylated peptides corresponding
to rhodopsin and S-opsin C-terminal sequences released by Asp-N cleavage were
synthesized by Biomer Technology (Concord, CA). The monophosphorylated S-
opsin C-terminal peptide contained 15N at the amino-N of residues V335 and V338.
The monophosophorylated rhodopsin C-terminal peptide had 15N at the amino-N
of A337 and A346 sites. The detection efficiency of each peptide species was
determined by integrating the area under each peak on the mass chromatogram,
normalized against peptide quantity measured by its OD205 nm absorbance from
the UV-vis chromatogram.

3.2.3. Light Stimulation and Sample Preparation for LC-MS.
The light source was a 100 W quartz tungsten halogen lamp connected to a
liquid light guide (Oriel Instruments, Stratford, CT). Light stimulation was
controlled by neutral density filters, interference filters (Oriel Instruments), and an
electromagnetic shutter (Vincent Associates, Rochester, NY). Light intensity was
measured using a calibrated photodiode (United Detector Technology Sensors, Inc.,
Hawthorne, CA) positioned at an equal distance to the retina. The current was
measured using a current amplifier (SR570 current preamplifier; Stanford Research
64

Systems, Sunnyvale, CA). The sample preparation was modified based on
published protocol (Kennedy et al., 2001). Dissected retinas were either kept in
darkness or exposed to calibrated light for 30 sec and incubated in darkness for a
period of time. The retinas were homogenized in 700 µl urea buffer (7 M urea, 5
mM EDTA, 20 mM Tris-HCl [pH 7.4]). The retinal membranes were digested with
50 µl of 10 µg/ml Asp-N protease (Roche, Inc.) in 10 mM ammonim bicarbonate
(pH 8.0) at room temperature overnight. The peptides were then acidified with
formic acid to pH < 2 and stored at -20°C.

3.2.4. LC-MS.
Twenty µl peptide samples were loaded onto a C18 capillary column
(Thermo Finnigan) in 0.08% HFBA at a flow rate of 5 µl/min. The peptides were
separated by a 5% - 30% acetonitrile gradient. The eluent was delivered to an LCQ
Deca XP Mass Spectrometer (Thermo Finnigan) to record the full MS and MS/MS
spectra. The fragmentation parameters used to break the parent ions by collision-
induced dissociation (CID) were: activation amplitude of 35%, activation Q of 0.25,
and activation time of 50 ms. Rhodopsin or S-opsin C-terminal peptides were
monitored in the mass detector with an isolation width of 1.5 centered on the
predicted m/z values of rhodopsin or S-opsin C-terminal peptides released by Asp-
N digestion. The most abundant ion of each peptide was monitored. Identities of ion
peaks on the mass chromatogram were confirmed by their MS/MS spectra. After

65

correcting for the detection efficiency of each peptide species, the area under each
peak was integrated to quantify amount of peptide.

3.2.5. Isoelectric Focusing.
The preparation of retinal sample and IEF gel electrophoresis to separate
phosphorylated rhodopsin species was performed as described previously (Adamus
et al., 1993). After separation on the IEF gel, the proteins were transferred to a
nitrocellulose membrane by capillary forces. Rhodopsin species were detected by
western blot using mAb 4D2, which recognizes the N-terminal domain of rhodopsin
(Laird and Molday, 1988). Procedures used in separating phosphorylated S-opsin
species were similar to those used for rhodopsin, except that the pH gradient of IEF
gel used to resolve phosphorylated S-opsins was pH 3-10, instead of pH 3-8, as was
the case for rhodopsin. After transferring the proteins to nitrocellulose membrane,
phosphorylated S-opsin species were detected by western blot using an antibody
raised against the N-terminus of S-opsin (Shi, 2004).

3.2.6. Mathematical Simulations of Trans-phosphorylation.
The simulations were performed on a 200 x 200 square array of rhodopsin.
These rhodopsin molecules were arranged as if on a grid, and is able to trans-
phosphorylate its neighbor to the north, south, east and west. The array assumed a
random distribution composed of 86% rhodopsin and 14% S-opsin (determined by
rhodopsin and S-opsin expression level in S-opsinrho+/- retinas). A small, 20 x 20
66

portion of this grid is illustrated in the top panel of Figure 3.7. We denoted the
subset of molecules corresponding to rhodopsins as R. Initially, a random
proportion, p, of R, was chosen to become light activated and phosphorylated.
Given the particular value of p used for this iteration, the exact proportion of
rhodopsin that would become phosphorylated was chosen by randomly sampling
from the entire population of R (Figure 3.7., middle). We denoted this set by P.
Each iteration had a different p value chosen at random such that it fell between 0
and 1. Subsequently, we chose a random subset of molecules and labeled them S,
corresponding to the proportion of S-opsin. These were chosen such that the final
proportion of S equaled 0.14 (Figure 3.7., top and middle). There was no overlap
between subsets P and S. Next, we simulated a process wherein each member of P
randomly selected just one of its neighboring molecules to become phosphorylated
(Figure 3.7., bottom). If the chosen molecule was type S, it is designated as S1P. If
the chosen molecule was type R, it became R1P. We recorded the total number of
molecules in S that underwent this latter conversion process and became S1P.
Simulations were also performed where each member of P was able to cause
phosphorylation of two or three neighbors.
We repeated this simulation process 2,500 times for each value of p in order
to obtain an accurate estimate of the average number of S molecules that were
converted to S1P status. If the mean proportion of such molecules over the course
of these 2,500 replicates came close to the target proportion of S1P’s, denoted by sp,
which is estimated experimentally, we recorded the value of p that generated this
67

data set. The entire procedure was repeated 5,000 times, which led to a set of
accepted p values, and yielded a report on the mean of these p’s. Subsequently, we
used this value to calculate the number of rhodopsin molecules that became R1P,
which includes members of both subset P and R1P resulted from neighbor selection
by P.

3.3. Results
We sought to investigate whether RK, once activated by R*, would
phosphorylate neighboring unbleached visual pigments in native disk membrane by
a trans-phosphorylation mechanism. To address this question, transgenic mice were
generated that expressed S-opsin (a mouse cone pigment) in rod photoreceptors
expressing endogenous rhodopsin. Importantly, S-opsin is a native substrate for RK
in murine cones (Weiss et al., 2001). Characterization of these mice showed that
ectopically expressed S-opsin localized exclusively to the rod outer segment (Shi,
2004), produced light responses (our unpublished results), and was capable of
undergoing light-stimulated phosphorylation (see below). Therefore, S-opsin
appears to be correctly folded and functional in transgenic rods.

3.3.1. Detection of Rhodopsin and S-opsin Phosphorylation.
Activation by 360-420 nm Light. To detect phosphorylation of S-opsin and
rhodopsin at their carboxyl termini, we adopted a mass spectrometry-based method
described by Kennedy et al. (Kennedy et al., 2001). Briefly, treatment of urea-
68

washed retinal membranes with endo-proteinase Asp-N released the carboxyl-
terminal peptide that contains all (6/6) or most (8/9) of the Ser/Thr phosphorylation
sites on rhodopsin and S-opsin, respectively, from the disk membranes. The
solubilized peptide mixture, including non-phosphorylated and phosphorylated
peptides, was separated by reverse-phase chromatography and detected by a
nanospray ionization ion trap mass spectrometer (Thermo Finnigan DECA XP) at
their specific mass-to-charge (m/z) ratios. Synthetic peptides derived from
rhodopsin and S-opsin carboxyl-terminal sequences were used to determine their
HPLC elution profiles and calculate the ionization efficiency of each peptide
(Figure 3.1.). Transgenic mice expressing S-opsin were crossed into rhodopsin +/-
or rhodopsin -/- genetic backgrounds (Lem et al., 1999) to obtain S-opsinrho+/- or
S-opsinrho-/- genotypes. To estimate the relative proportion of S-opsin to rhodopsin
in S-opsinrho+/- rod, retinal membranes were prepared from dark-adapted mice
wherein non-phosphorylated peptides predominated. The elution profiles of
peptides released from Asp-N cleavage of retinal membranes obtained from dark-
adapted mice were identified and confirmed by MS/MS. After correcting for their
differences in ionization efficiency (Figure 3.1.), we estimated the proportion of S-
opsin to be 14 ± 1 % (SD, N=3) of total pigment molecules (Figure 3.2.A).
We first tested whether S-opsin would undergo light-activated
phosphorylation in rod photoreceptors. Control experiments showed that peptides
corresponding to monophosphorylated rhodopsin and S-opsin of S-opsinrho+/-
retinas were readily detected after exposure to short wavelength light (360-420 nm)
69

Figure 3.1. Ion chromatograms of eluted peptides synthesized according to C-
terminal sequences of rhodopsin and S-opsin. The following peptides were
synthesized, separated by C18 reverse-phase chromatography, and detected by mass
spectrometry: R1P (rhodopsin C-terminal peptide monophosphorylated at site 343;
doubly charged; m/z = 974.0); R0P (non-phosphorylated rhodopsin C-terminal
peptide; doubly charged; m/z = 933.1); S1P (S-opsin C-terminal peptide
monophosphorylated at site 328; doubly charged; m/z = 1099.9); and S0P (non-
phosphorylated S-opsin C-terminal peptide; triply charged; m/z = 706.3). Of each
type of peptide, the most abundant ion with specific charge and m/z was monitored
in a narrow mass window of 1.5 D. The x-axis is time course of HPLC elution; the
y-axis is ion intensity. The area of integrated peak was normalized against a loaded
peptide amount measured by its absorption at 205 nm to derive the relative detection
efficiency of each peptide. R1P synthetic peptide contains
15
N isotopes at amino-N
of A337 and A346 sites, and S1P synthetic peptide contains
15
N isotopes at amino-N
of V335 and V338 sites. Therefore, their m/z values are 1 a.m.u./e greater than those
of wild-type peptides obtained from retinal samples (in Figure 3.2 and 3.3). R0P and
S0P synthetic peptides have the same m/z as wild-type peptides. The numbers
beside each shaded peak represent the number of ions detected prior to normalization
for ionization efficiency.

70

Figure 3.2. Phosphorylation of rhodopsin and S-opsin following exposure to 360-
420 nm Light. Ion chromatograms of R1P (monophosphorylated rhodopsin C-
terminal peptide; m/z = 973.0), R0P (non-phosphorylated rhodopsin C-terminal
peptide; m/z = 933.1), S1P (monophosphorylated S-opsin C-terminal peptide; m/z
= 1098.9), and S0P (non-phosphorylated S-opsin C-terminal peptide; m/z = 706.3),
from dark-adapted S-opsinrho+/- retinas (A), and 360-420 nm light exposed S-
opsinrho+/- (B), S-opsinrho-/- (C) and rho+/- (D) retinas. The rhodopsin and S-
opsin peptide sequences were indicated on top of R0P and S0P spectra,
respectively. The area of integrated peak was indicated. Both R1P and S1P ion
peaks contained a mixture of peptides singly phosphorylated at different sites.
Whereas different species of R1P eluted as one peak, S1P ions displayed two peaks,
arising from two groups of monophosphorylated peptides with slightly different
hydrophobicities. Each experiment was repeated at least three times.

71

that caused an estimated ~100% bleach of rhodopsin and S-opsin, followed by 10
min of dark incubation, (Figure 3.2.B), indicating that both S-opsin and rhodopsin
were phosphorylated following their activation by UV light. In the experiments
involving mass spectrometry, we chose to monitor only singly phosphorylated
species as readout for rhodopsin and S-opsin phosphorylation because they had
been shown to be a highly abundant species after light exposure (Kennedy et al.,
2001), and because of the relatively high sensitivity for their detection by the mass
spectrometer. In the absence of rhodopsin (S-opsinrho-/-), S-opsin also became
phosphorylated following stimulation by short wavelength light (Figure 3.2.C). As
predicted, rhodopsin by itself also became phosphorylated following exposure to
short wavelength light (Figure 3.2.D). These data indicate that the ectopically
expressed S-opsin is correctly folded and is a substrate for RK in native rod disk
membrane.

3.3.2. S-opsin Becomes Trans-phosphorylated following Activation of
Rhodopsin by Long (515-620 nm) Wavelength Light.
Murine S-opsin has maximum absorbance in the ultraviolet range (357 nm
(Jacobs et al., 1991; Lyubarsky et al., 1999)). At longer wavelengths, its absorbance
falls precipitously, and, at λ >= 515 nm, photon absorption by S-opsin is 104-fold
less efficient than rhodopsin (Jacobs et al., 1991; Lyubarsky et al., 1999). On the
other hand, at the peak absorption of S-opsin, rhodopsin can be readily activated,
albeit at ~25% of its maximum sensitivity at 500 nm (Lyubarsky et al., 1999). We
72

utilized the large separation of spectral sensitivity at long wavelengths to selectively
stimulate rhodopsin, and sought to determine whether S-opsin becomes
phosphorylated as a consequence of rhodopsin activation.
S-opsinrho+/- retinas were exposed to long wavelength (515-650 nm) light,
causing an estimated ~100% bleach of rhodopsin, and incubated in darkness for 10
min (Figure 3.3.). As expected, long wavelength light exposure led to efficient
phosphorylation of rhodopsin (Figure 3.3.A). When S-opsin was expressed in the
absence of endogenous rhodopsin, it did not become phosphorylated after exposure
to 515-620 nm light (Figure 3.3.B), verifying our expectation that S-opsin would
not be efficiently activated by long wavelength light under our experimental
settings. Interestingly, when retinas co-expressing S-opsin and rhodopsin were
exposed to long wavelength light, a substantial fraction of S-opsin’s carboxyl
terminus became phosphorylated (Figure 3.3.C), indicating that non-photolyzed
pigments can serve as substrate for activated RK.
The extent of S-opsin that becomes indirectly phosphorylated (trans-
phosphorylation) was examined at different times after bright light exposure that
was sufficient to bleach 100% of rhodopsin (Figure 3.4.A). Under this condition,
the concentration of RK became rate limiting such that rhodopsin phosphorylation
continued to increase as a function of time in dark incubation (Laitko and Hofmann,
1998; Kennedy et al., 2001). Figure 3.4.A illustrates the proportion of
phosphorylated peptides normalized to the amount of each pigment population.
Whereas the level of singly phosphorylated rhodopsin reached a steady state at 2
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Figure 3.3. Trans-phosphorylation of S-opsins following generation of R* by 515-
620 nm light. Ion chromatograms of R0P, R1P, S0P and S1P from S-opsinrho+/-
(A), S-opsinrho-/- (B), and rho+/- (C) retinas after 515-620 nm exposure. Each
experiment was repeated at least three times.

74

min, the level of S-opsin phosphorylation continued to increase thereafter, and, at
10 min, the proportion of monophosphorylated S-opsin reached the same relative
level as rhodopsin. The lag of S-opsin phosphorylation behind R* phosphorylation
suggests that R* is the preferred substrate for RK; as all available R* is
phosphorylated, RK becomes available to phosphorylate neighboring R.
If R* is the preferred substrate, then the relative amount of trans-
phosphorylation would likely be higher under low bleach conditions that generate
fewer R*. This notion was tested by comparing the levels of S-opsin
phosphorylation under different light intensities. As shown in Figure 3.4.B, the
level of S-opsin phosphorylation was minimal shortly following >100% bleach of
rhodopsin. This fraction increased two-fold following a longer period of dark
incubation as more R* deactivated. When the retina was exposed to 1.2% bleach,
the relative amount of trans-phosphorylation was substantially higher at the earlier
time point and remained stable over time (Figure 3.4.C). In fact, as much as 7%
rhodopsin was found to be mono-phosphorylated following 1.2% bleach, suggesting
a substantial gain to this reaction. Together, these data provide strong evidence that
trans-phosphorylation occurs in living photoreceptor cells under low light
conditions. These data are consistent with previous findings where the highest gain
of phosphorylation reported was for an exceedingly low bleach where one R* was
generated per disk. When multiple R*’s are generated per disk, the gain decreases
(Binder et al., 1990; Binder et al., 1996). One likely explanation for this observation,
as mentioned above and also stated by Binder et al., is that R* may be the preferred
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Figure 3.4. Rhodopsin phosphorylation and S-opsin trans-phosphorylation level as
a function of dark incubation time after exposure to different intensities of light.
Each data point represents the average of at least three independent experiments.
(A) Kinetics of rhodopsin and S-opsin phosphorylation following light exposure.
S-opsinrho+/- retinas were illuminated with 515-620 nm light for 30 sec, which
caused an estimated 100% bleach to rhodopsin. Retinas were then incubated in
darkness for 10 sec, 2 min, 5 min or 10 min. At each time point, phosphorylation
levels of rhodopsin and S-opsin were quantified as a normalized fraction of R1P to
the total amount of R1P and R0P, and S1P to the total amount of S0P and S1P,
respectively. (B) S-opsinrho+/- retinas were exposed to 515-620 nm light, which
bleached ~100%, and (C) ~1.2 % rhodopsin. Phosphorylation levels of rhodopsin
and S-opsin at 2 min and 5 min after bleach were quantified as in (A).

76

substrate for RK; at high bleach, R* would efficiently compete with R for
interactions with RK, which is at limiting concentration in the rod outer segment.
We used mathematical simulations to estimate the fraction of rhodopsin (R)
that becomes trans-phosphorylated. Simulations were based on the experimentally
derived value of S-opsin expression level and the amount of S-opsin that became
trans-phosphorylated following light stimulation that beached ~1.2% of rhodopsin.
Using mass spectrometry, we determined that the amount of trans-phosphorylated
S-opsin was ~7.5 % of total S-opsin after 1.2% bleach of rhodopsin (Figure 3.4.C).
Based on our simulations, we estimated that 4.9% of R would be trans-
phosphorylated, assuming that rhodopsins and S-opsins were equally good trans-
phosphorylation substrates. This value was in close agreement with the
experimentally assessed value (7.1 ± 2.9%) of the total pool of monophosphorylated
rhodopsin; this included directly phosphorylated R* and trans-phosphorylated R
(1.2% and 4.9%, respectively). It should be noted that these values underestimated
the total number of phosphorylated pigments because only monophosphorylated
species were monitored. Nevertheless, they represented an approximation of the
underlying process since monophosphorylated pigment appeared to be the most
abundant phosphorylated species under this light condition (see below).
We also simulated the amount of R* required to obtain the experimentally
derived value for phosphorylated S-opsin, if each R* was allowed to phosphorylate
one, two, or three neighbors. The simulation estimated the values to be 4.9%,
2.45% and 1.2% R*, respectively. A comparison of these values with our estimated
77

1.2% bleach, based on the calibrated light source, suggests that at least three R’s
become trans-phosphorylated for each R* generated.

3.3.3. Trans-phosphorylation Results in Multiply Phosphorylated S-opsin.
The sensitivity of mass spectrometric detection of peptides decreases as the
number of phosphate modifications increases. Although mass spectrometry offers
higher sensitivity and better quantification of low levels of singly phosphorylated
peptides, isoelectric focusing (IEF), followed by western blotting, provides an
alternative means by which to visualize multiply phosphorylated rhodopsins and S-
opsins. Murine rhodopsin contains six phosphorylatable sites on its carboxyl
terminus, and all six differentially phosphorylated species were clearly resolved by
IEF (Figure 3.5.A). Rhodopsin species incorporating up to four phosphates were
clearly detected ten seconds after 100% bleach, and species with five and six
phosphates appeared with increasing dark incubation time after the bleach. A visual
comparison of signals from these species indicates that they are present at
approximately equal molar ratios. These data also show that phosphorylation
reaches a steady state about 2 min after the 100% bleach, similar to the results
presented in Figure 3.4. Multiply phosphorylated rhodopsins incorporating up to
three phosphates were also observed after 1.2% bleach (Figure 3.5.A).
IEF followed by western blotting, using an amino-terminal antibody against
S-opsin, was employed to detect phosphorylated species of S-opsin (Figure 3.5.B).
Exposure of S-opsinrho-/- retina to 360-420 nm light led to S-opsin species that
78

Figure 3.5. Detection of phosphorylated rhodopsin and S-opsin species by
isoelectric focusing. (A) S-opsinrho+/- retinas were exposed to 515-620 nm light,
which caused the indicated rhodopsin bleaching levels; they were incubated in
darkness for the indicated period of time. Dk, dark-adapted control retina.
Rhodopsin species incorporating zero to six phosphates (numbers marked on right
side) were resolved on a pH 3-8 IEF gel and detected by western blot using mAb
4D2 antibody against the rhodopsin N-terminus. (B) Trans-phosphorylation results
in multiply phosphorylated S-opsin. S-opsin carrying zero to nine phosphates were
separated on pH 3-10 IEF gel and detected by western blot using an antibody
against the S-opsin N-terminus. The number of phosphates incorporated is
indicated on the right. Lane 1, S-opsinrho-/- retina exposed to 360-420 nm light,
causing ~100% bleach, followed by 10 min of dark incubation. Lanes 2-7, S-
opsinrho+/- retinas incubated in the dark (Dk), or exposed to 515-620 nm light for
the indicated bleach, followed by the indicated time in dark incubation. Lane 8,
rho+/- retina exposed to 515-620 nm light, causing ~100% bleach of rhodopsin.
Lane 9, S-opsinrho-/- retina exposed to intense 515-620 nm light and incubated in
the dark for 10 min. Lane 10, S-opsinrho-/-, RK-/- retina exposed to intense 360-
420 nm light, followed by 10 min in dark incubation. The band at the upper edge of
each lane is the trace of loading position; it overlaps with the band that represents
S-opsin containing nine phosphates (lane 1). * is a non-specific band unrelated to
S-opsin.

79

incorporated up to nine phosphates (lane 1). Selective stimulation of rhodopsin in
S-opsinrho+/- retinas by intense 515-620 nm light also led to multiply
phosphorylated S-opsin, particularly after a period in dark incubation (lanes 3-5).
Results from the trans-phosphorylated S-opsin indicate a different pattern of
phosphorylation when compared with direct stimulation (lane 1). After 1.2% bleach,
monophosphorylated species appeared to be most abundant (lanes 6, 7). As
expected, phosphorylation of S-opsin did not occur when S-opsinrho-/- retinas were
stimulated with 515-620 nm light (lane 9), or in the absence of RK (lane 10). Thus,
trans-phosphorylation gives rise to multiply phosphorylated, non-activated visual
pigment molecules.

3.3.4. Demonstration of Trans-phosphorylation in Transgenic Mice that
Express Human K296E Opsin.
K296E is a naturally occurring mutation in the rod opsin gene that leads to
autosomal dominant retinitis pigmentosa (Keen et al., 1991). In wild-type rhodopsin,
the 11-cis retinal is covalently attached to the lysine residue at position 296 of
rhodopsin (Bownds, 1967). Mutation at this position renders the opsin protein
incapable of binding 11-cis retinal, and also leads to constitutive activity of the
opsin protein in biochemical assays (Zhukovsky et al., 1991; Robinson et al., 1992).
When expressed in transgenic mice, however, K296E apparently is inactivated by
phosphorylation and arrestin binding and shows no evidence of constitutive activity
(Li et al., 1995).
80

Figure 3.6. Trans-phosphorylation of endogenous mouse rhodopsin in dark-adapted
K296E transgenic mouse retina. Dark-adapted retinas from K296E transgenic mice
and transgene negative controls were prepared under infrared light. Normalized
fraction of monophosphorylated C-terminal peptide released from endogenous
mouse rhodopsin was compared between K296E and control retinas in the arr +/+
(A) and arr -/- (B) backgrounds.

81

We utilized the rhodopsin K296E transgenic mouse that expressed human
K296E in rod photoreceptor cells as another model to demonstrate the presence of
trans-phosphorylation in vivo (Li et al., 1995). Since K296E is capable of activating
RK in the dark while the endogenous wild-type rhodopsin is unbleached, trans-
phosphorylation would result in phosphorylated mouse rhodopsin in the dark-
adapted K296E retinas. The carboxyl-terminal peptide of mouse rhodopsin has an
m/z value different from that of human rhodopsin; thus, phosphorylation of human
and mouse rhodopsin could be unequivocally determined by LC-MS detection of
the respective carboxyl-terminal phosphopeptides. As shown in Figure 3.6.A, no
differences in endogenous mouse rhodopsin phosphorylation were observed when
comparing dark-adapted retinas obtained from K296E transgenics and wild-type
mice (Figure 3.6.A). This result is consistent with the observation that K296E is
stably inactivated by arrestin binding and, therefore, is incapable of further
activating RK (Li et al., 1995). To unmask the constitutive activity of K296E, this
transgene was crossed into the arrestin -/- background. Indeed, in the absence of
arrestin, the constitutive activity of K296E led to readily detectable phosphorylation
of endogenous mouse rhodopsin (Figure 3.6.B). As expected, rhodopsin
phosphorylation in the dark was not affected by the absence of arrestin (Figure
3.6B). These results provide independent evidence of the presence of a trans-
phosphorylation mechanism in vivo.

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3.4. Discussion
Phosphorylation of non-activated rhodopsin has been observed in isolated
rod outer segment preparations, as well as in living frogs (Aton, 1986; Binder et al.,
1990; Chen et al., 1995a; Binder et al., 1996). However, the mechanism and the
extent by which this happens in intact photoreceptor cells of mammals have not
been addressed. In this study, we sought to investigate whether unbleached visual
pigments would become phosphorylated by RK through a trans-phosphorylation
mechanism. We utilized two transgenic mouse models: one that expressed a cone
shortwave pigment, and one that expressed a constitutively active human rhodopsin
mutant (K296E) in the rod photoreceptors. These mouse models enabled us to
specifically monitor the phosphorylation of unbleached visual pigment, as well as
R* molecules, in their native environment, and each model provided independent
evidence in strong support of trans-phosphorylation as a mechanism for high-gain
phosphorylation.
To detect the phosphorylation of visual pigments at their carboxyl termini, a
mass spectrometry method was employed. We chose to monitor
monophosphorylated peptides in order to report the presence of phosphorylation
events, since the levels of monophosphorylated species were more readily
detectable and quantifiable in our experiments as compared with multiply
phosphorylated peptides. Thus, the values we obtained from this method
represented an underestimation of the total number of phosphorylated pigment
molecules, especially under the condition of times after 100% bleach; in such
83

Figure 3.7. Simulation of trans-phosphorylation by light-activated rhodopsin. Top
panel: rhodopsin molecules are represented as white squares, and S-opsin molecules
are represented as grey squares. The distribution of rhodopsin and S-opsin is 86%
and 14%, respectively. Middle panel: light activated, phosphorylated rhodopsin is
represented as a black square. This diagram illustrates that ~1% of rhodopsin is
light activated. Bottom panel: Light activated rhodopsin causes trans-
phosphorylation of neighboring rhodopsin and S-opsin molecules, designated as
cross-hatched white squares and grey squares, respectively.

84

instances, the results from IEF showed that monophosphorylated species arising
from phosphorylated R and R* may represent only 1/6 of all of the phosphorylated
R*/R molecules. At lower bleach (1.2%), however, monophosphorylated species
appeared to be the more abundant species. Therefore, the estimates derived at this
light level may be more accurate. The IEF results also provide evidence that trans-
phosphorylation produces highly phosphorylated R that, upon photoisomerization,
will give rise to a diminished response.
Our results suggest that multiple non-activated rhodopsin molecules become
trans-phosphorylated for each R* generated. Given the high diffusion rate of
rhodopsin along the lipid bilayer (Wey et al., 1981; Kaplan, 1984; Calvert et al.,
2001), one can envision the R*/RK complex making contact with several non-
activated rhodopsin molecules and phosphorylating their flexible carboxyl termini.
Another possibility is that, once activated by associating with an R*, RK may be
able to diffuse some distance away and phosphorylate R. The dependency of the
gain of the phosphorylation reaction on the integrity of the rod outer segment
preparation, as well as the inability of Rim and colleagues to observe trans-
phosphorylation in solubilized proteins, are two lines of evidence against freely
diffusible, active RK (Binder et al., 1990; Rim et al., 1997). This does not, however,
rule out the possibility that active RK may be able to diffuse short distances alone.
Future experiments would be required to clearly distinguish between these
possibilities.

85

The highest level of trans-phosphorylation under our experimental
conditions, as indicated by S-opsin phosphorylation, was seen after 10 min of dark
incubation following exposure to intense light that caused 100% bleach of
rhodopsin. In this instance, up to 20% of the entire population of S-opsin became
phosphorylated. Although we did not explore the lighting condition that caused the
highest level of trans-phosphorylation, our results suggest that a substantial fraction
of non-activated visual pigments can be phosphorylated by this mechanism. Besides
trans-phosphorylation, another source of light-responsive, phosphorylated
rhodopsin may be from regeneration of phosphor-opsin that has not yet been
dephosphorylated. This is not unlikely, given the apparent slow rate of
dephosphorylation (Ohguro et al., 1995). Inasmuch as S-opsin was not activated in
our assay, our system did not report phosphorylated R from this additional pathway.
What may be the physiological consequence of photoactive, phosphorylated
visual pigment? It should be noted that, in a fully dark-adapted rod cell, suction
electrode recordings have shown that amplitude saturation was reached upon a flash
of light that excited ~80 rhodopsin molecules, as well as upon ~400 R*/sec under
steady background light (Baylor et al., 1984). Under the latter condition, up to 5%
R* can accumulate in 10 min given the lag of the phosphatase activity. It is
plausible, therefore, that, under certain steady-state lighting conditions, trans-
phosphorylation may have an impact on decreasing transduction gain and thereby
extending the range of rod response.

86

Another process by which trans-phosphorylation may have a profound
impact takes place when the retina switches from photopic (cone) vision to scotopic
(rod) vision. The switch between photopic and scotopic vision is a process crucial
to our ability to detect visual cues within the wide variation of environmental
illumination. During the switch from cones to rods, a period of dark adaptation is
required for the rod cell to recover maximal sensitivity after high bleach. Our data
indicate that the trans-phosphorylation reaction continues long after light exposure.
Since phosphorylated R* catalyzes transducin activation less efficiently, but is
deactivated more efficiently, photoisomerization of these phosphorylated R’s is
expected to give rise to a response with a slowed rising phase, a decreased
amplitude, and a shorter duration; and, hence, decreased sensitivity during the
period of dark adaptation. It is also important to consider that the presence of
phosphorylated R would be expected to affect the reproducibility of the single
photon response. It is well documented that the invariant shape and size of the
elementary light response are salient features of rods that underlie their ability to
encode our visual scene under dim light (Rieke and Baylor, 1998). In this situation,
dephosphorylation of R may be a limiting step in fully restoring visual sensitivity
during dark adaptation. In light of the recent discovery that other G-protein-coupled
receptors exist as dimers, or even as higher-structure oligomers (Angers et al.,
2002), trans-phosphorylation of non-activated receptors in close proximity to
activated receptors may also have an impact on other signaling pathways.
87

Acknowledgments
We thank Dr. Tiansen Li for providing the K296E transgenics, and Dr.
Robert Molday for providing the 4D2 rhodopsin antibody. We also thank Dr. James
Hurley for his helpful comments on the manuscript. This work was supported by
NIH Grant EY12155 (JC) and the Beckman Macular Research Center Grant (JC
and RL).

88

Chapter 4
Rhodopsin Turnover and Trafficking in
Rhodopsin-Timer (Timer) Transgenic Mice

4.1 Introduction
The detection of light by photoreceptors is the first stage in the process of
vision. The rod outer segment (ROS) consists of many hundreds of densely packed
membranous discs in vertebtrates photoreceptors (Young, 1971a), containing
necessary components to conduct phototransduction. The light pigment rhodopsin is
almost exclusively embedded in these discs of ROS. It constitutes more than 90% of
the total membrane protein with concentration of ~3mM (Liebman and Entine, 1968;
Papermaster and Dreyer, 1974; Harosi, 1975). Since rhodopsin amount is so
prominent in ROS, it is not only a phototransduction element but also a structural
protein in photoreceptors. Rhodopsin knockout leads to retinal degeneration resulting
from inadequate structural support (Lem et al., 1999). Proteins are synthesized in the
inner segment (IS) which is connected to the outer segment with an extremely
narrow cilium. The IS includes ellipsoid and myoid, characterized by densely packed
mitochondria or endoplasmic reticulum (ER), Golgi complexes and ribosomes,
respectively. Rhodopsins are synthesized in ER, modified in Golgi and then
transported by post-Golgi vesicles to the base of outer segment where they are
assembled into new discs, move towards the apical end of the cell and progressively

89

displace the older discs. The discs are normally phagocytosed by epithelial cells at
the apical end.
Tritium autoradiography was first utilized to study the turnover of rod outer
segment in the 1960s. The tritium-labeled amino acids were injected into a group of
mice at the same time, and the animals were sacrificed at different time/days to
enable observation of the whole turnover process. The results showed that labeled
outer segment moved an equal distance per day, suggesting that ROS undergoes
constant renewal and shedding (Young, 1967, 1971a, b). By the same method, it was
also revealed that rhodopsin does not undergo turnover once it is incorporated into
the membrane disc (Young, 1967; Hall et al., 1969; Young, 1971a). Newly-formed
discs displace the older discs at the base of ROS, and the oldest discs at the apical tip
of the outer segment are shed and phagocytosed by the retinal pigment epithelium. It
takes about 10 days in mice, rat and X. laevis for a single disc to migrate along the
rod outer segment before being shed at the apical tip; it takes from 9-13 days in
rhesus monkeys and other primates, depending on the length of ROS in the region
(Young, 1973). Although this method was effective, it was crude, as the investigators
had to track and compare different molecules in different mice to observe rhodopsin
turnover instead of the same molecule in the same ROS. Compared to the tritium-
labeled amino acids injection, fluorescence protein labeling is a more effective and
efficient method to track rhodopsin/ROS turnover speed.
Photoreceptor cells have an extremely high metabolism requirement,
consuming much more energy than other cell types in the retina. The average
90

number of discs stacked in a single rod outer segment is ~1100 in the parafovea, 920
in the perifovea and 790 in the periphery area of retina, which means on average
around 80-90 discs are renewed per rod per day, including all the disc proteins,
especially rhodopsin (Young, 1973). The average speed of 10 days for disc renewal
is extremely fast. Rhodopsin does not undergo turnover during the period that it
forms the structure protein in the disc membrane till phagocytosed by epithelium at
the apical tip of rod (Hall et al., 1969). So why does the photoreceptor cell spend so
much energy to renew and shed discs and rhodopsin? It was proposed that the fast
renewal process is a preventive maintenance for replacing “aging” rhodopsin “before
they wear out” (Young, 1967; Young and Droz, 1968; Young, 1971a; Anderson and
Fisher, 1976). However, there was no evidence that rhodopsin molecules can be
“light damaged” or otherwise “aged” within 10 days (Kaplan and Deffebach, 1978;
Baylor et al., 1979; Williams, 1984). At the time, it was thought that ROS/rhodopsin
turnover at a constant speed.
New insights into rhodopsin turnover came decades later: it was discovered
that photoreceptor shedding as well as its renewal are initiated by light in mice and
frogs (Basinger et al., 1976; LaVail, 1976a, b; Besharse et al., 1977; Bird et al.,
1988). It was further found that a diurnal rhythm regulated inner segment (IS) opsin
transport: opsin labeling was lowest at light onset, increasing by three- to four-fold
and remained until high 2 hours into the dark phase (Bird et al., 1988). Thus, the rod
outer segment is not renewed and shed at a constant rate, instead responding to
environmental lighting. Another observation was that ROS length and rhodopsin
91

synthesis and packing density in disc changed according to light and dark cycles
(Schremser and Williams, 1995a, b). It was proposed that fast ROS turnover is a
means to adapt to a new lighting environment. All these results suggest that
rhodopsin turnover is a highly dynamic process.
The fast turnover rate of rhodopsin provides a good opportunity to study
rhodopsin trafficking on a 10-day time scale. Numerous studies have suggested that
normal rhodopsin trafficking is very important for cell survival. Mutations like
Q344ter, P347S affect rhodopsin’s C-terminal region. Mutations that affect the last
five amino acids QVXPX-COOH, disrupt rhodopsin localization and lead to retinal
degeneration (Sung et al., 1994; Concepcion et al., 2002; Illing et al., 2002; Deretic,
2004; Tam and Moritz, 2006). However, less progress was made towards
understanding the turnover of mutant rhodopsin. Although there have been sporadic
reports about autophagy and endosome turnover (Reme et al., 1999; Chuang et al.,
2004), rhodopsin phagocytosis by RPE is considered to be the only means of
rhodopsin turnover in vertebrates photoreceptors. As mentioned previously,
fluorescent protein labelling is an alternative means to visualize rhodopsin turnover.
In this study, the new fluorescent protein Timer was applied to study rhodopsin
turnover. Timer has the ability to change color as a function of time (Terskikh et al.,
2000; Gagescu, 2001), maturing from green to orange to red. Such spectroscopic
changes make it possible to study the temporal and spatial behavior of rhodopsin, in
a single image, a significant advantage over the regular green fluorescent protein

92

(GFP). The high rate of ROS renewal and shedding makes a good subject for study
with Timer.
We made three mutant rhodopsin-timer constructs and generated two
wildtype rhodopsin-timer (Timer) transgenic mouse lines. By monitoring Timer
expression, we aim to get a better understanding of rhodopsin turnover/trafficking.
Three rhodopsin mutant-timer constructs were also made as Q344ter-Timer, P23H-
Timer and K296E-Timer. These represent three different categories of rhodopsin
mutants which cause autosomal dominant retinitis pigmentosa (ADRP) by
mistrafficking, misfolding, and constitutive active/toxic complex formation
respectively (see introduction page 20).

4.2. Meterials and Methods
4.2.1. Generation of non-aggregated Timer
The non-aggregated Timer construct was spliced together by using two
fragments of DNA. The first half was from Dsred2 in pDsred2-C1 (BD science
Clontech) digested by AgeI and StuI which includes the V105A mutation of Timer
and 3 additional mutations: R2A, K5E, K9T that are expected to decrease protein
aggregation; the second half was from pSCA1(ppANF-Timer) (generous gift from
Dr. Robert H. Chow), digested by StuI and NotI, that includes the second mutation
S197T. These 2 pieces of sequences were ligated together into pEGFP-N1 vector by
AgeI and NotI (Figure 4.1). This construct was transfected into the 293 cell line by
CaPO
4
-mediated transfection. Cells were observed by Total Internal Reflection
93

Fluorescence (TIRF) Microscopy. (The transfecion and TIRF was carried out in Dr.
Chow’s laboratory.)

4.2.2. Generation of Rhodopsin-Timer transgenic mice (Timer) and other
mutant rhodopsin constructs
The 1.1 kb mouse rhodopsin cDNA coding sequence was obtained by RT-
PCR with primers 5’SalI-Rho (5’ cgc GTCGAC atg aac ggc aca gag gg ), and primers
3’Rho-overlap primer (5’gtt ctc gga gga ggc cat ggc tgg agc cac ctg gct 3’). Timer
sequence was amplified by the 5’Timer1-overlap primer (5’ agc cag gtg gct cca gcc
ATG GCC TCC TCC GAG AAC 3’) and 3’Timer-1D4-ClaI primer (5’-
CCATCGATcta ggc tgg agc cac ctg gct ggt ctc cag gaa cag gtg gtg gc 3’) by adding
1D4 epitope, ClaI and a stop codon after Timer. Rhodopsin and Timer were then PCR
together with Timer attached to C-terminal of rhodopsin by primers 5’ SalI-Rho and
5’Timer1-1D4-ClaI primer and cloned into pBluescriptKSII+ vector using SalI and
ClaI sites. Before Rho-Timer insertion, the vector was inserted with 4.4 kb mouse
rhodopsin promoter at 5’ (KpnI and XhoI sites) and a 0.6 kb mpI fragment for
polyadenylation site at 3’ (BamH I site with direction selection) (Lem et al., 1991).
pRho-Timer was sequenced. The pRho-Timer plasmid was purified by CsCl
2
and
digested with Kpn I and NotI to yield the 6.6 kb insert fragment, which was then
purified by QIAquick gel extraction kit (Qiagen) and Elutip-D column. The 6.6 kb
fragment was microinjected into F1 hybrid zygotes from C57BI/6J and DBA/2J
strains following standard procedures.
94

95

Figure 4.1. Three-piece-ligation to generate non-aggregated form of Timer
construct. The non-aggregated Timer sequence was ligated by 2 pieces of DNA: the
first half was from Dsred2 digested by AgeI and StuI which includes V105A mutant
of Timer and 3 mutations R2A, K5E, K9T that decrease protein aggregation; the
second half was from pSCA1(ppANF-Timer) (generous gift from Dr. Robert H.
Chow) digested by StuI and NotI that includes the second mutation S197T. These 2
pieces of sequences were ligated together into pEGFP-N1 vector by AgeI and NotI.

pSCA1(ppANF-Timer)
NotI
pEGFP-N1
AgeI
pDsRed2-C1
AgeI
AgeI
V105A I161T S197A
R2A K5E K9T
StuI
V105A S197T
StuI NotI

Mutant rhodopsin Timer construct pQ344ter-Timer was made by the same overlap
PCR method with primers: 5’SalI-Rho, 3’Timer-Q344ter-ClaI (5’-CC ATC GAT cta
gct ggt ctc cag gaa cag gtg gtg gc 3’), and 5’Q344terTimer1-overlap primer (5’- aag
acg gag acc agc ATG GCC TCC TCC GAG AAC) and 3’Q344terRho- overlap
primer (5’ gtt ctc gga gga ggc cat gct ggt ctc cgt ctt 3’). pK296E-Timer and pP23H-
Timer were generated by site-directed mutagenesis with primers 5’K296Emut (5’ cc
agctttcttt gctgagagct cttccatc 3’), 3’K296Emut (5’ gat gga aga gct ctc agc aaa gaa agc
tgg 3’), and 5’P23Hmut (5’ gtg gtg cgg agc cac ttc gag cag ccg 3’) and 3’P23H (5’
cgg ctg ctc gaa gtg gct ccg cac cac 3’).

4.2.3 Genotype Analysis
Timer transgene positive mice in both lines were identified by transgene-
selective PCR amplification with primers RhoTimer-F (5’ ctc tgc cag ctt tct ttg ct 3’)
at the C-terminal of rhodopsin cDNA and RhoTimer-R (5’ CCT TGG TCA CCT
TCA GCT TC 3’) at Timer. Timer transgenic mice were bred into rhodopsin knock
out mice to obtain rhodopsin +/- or -/- background.

4.4.4. Retinal whole mount
All experimental procedures using mice were performed in accordance to
ARVO and USC IACUC guidelines. Mice were euthanized by CO
2
inhalation
followed by cervical dissociation. All mice were born and raised in 12h-light-12h-
dark cyclic light.
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The Timer transgenic, Timer
Rho+/-
, Timer
Rho-/-
retina were dissected under
light or infrared light and placed on a cryosection slide, with the photoreceptor layer
facing up. The retina was added with a drop of Vectorshild mounting media and
covered with cover slip. Then it was observed under Zeiss fluorescent microscope
(Zeiss Co.).

4.4.5. Vibrotome and confocol imaging
The procedure followed a modified protocol that was previously described
(Khodair et al., 2003). Briefly, Timer transgenic retina was dissected and embedded
in agarose and immerged in Ame’s solution (Sigma) and sectioned on vibratome
sectioning system. About 100um thick live-tissue sections were collected in Ame’s
solution and imaged by confocol microscopy. (The vibratome sectioning was
performed with help from Haruhisa Okawa in Dr. A. Sampath’s laboratory.)

4.4.6. EPON morphometry
Timer transgenics, Timer
Rho+/-
, Timer
Rho-/-
and their transgene negative
littermates were sacrificed, and the superior pole of the eye was marked by
cauterization before enucleation. Eyecups were then fixed overnight in 1/2
Karnovsky buffer (2% paraformaldehyde, 2.5% glutaraldehyde, and 0.1 M
cacodylate buffer, pH 7.2) and embedded into epon as described in Chapter 2. The
epon-embedded eyes were sectioned into 1 µm thickness and stained with
Richardson’s stain (0.5% methlene blue with 0.5% Borax, and 0.5% Azure II).
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4.4.7. Cryosection
Eyes were cauterized, enucleated to mark the superior pole, and fixed in 4%
paraformaldehyde buffer for 90 min at RT before wash with 0.1 M cacodylate (pH
7.2). Then eyecups were immersed in 30% sucrose buffer overnight followed by
embedding into O.C.T. (Tissue Tek). Sagital sections of retina were collected in 10
um thickness onto glass sildes, covered with Vectorshield mounting media and
coverslips, and observed under Zeiss fluorescence microscope.

4.4.8. Western blot
Timer transgenic, Timer
Rho+/-
, Timer
Rho-/-
retinas and their negative littermates
were dissected under infrared illumination and homogenized in buffer (80 mM Tris,
pH 8.0, 4 mM MgCl
2
) containing protease inhibitor cocktail at 1 tablet per 10 ml
buffer (Roche Diagnostics, Indianapolis, IN). DNase I (30 units, Roche Diagnostics)
was added and incubated at room temperature for 30 min. An equal amount of
retinal homogenate from each sample was loaded onto a 12% Bis-Tris SDS-PAGE
gel (Invitrogen Corp., Carlsbad, CA). Proteins were transferred onto nitrocellulose
membrane and incubated with rhodopsin N-terminal or C-terminal antibody, R2-12N
and 1D4 respectively, and visualized using enhanced chemiluminescence
(Amersham Pharmacia Biotech Inc., Piscataway, CA), or by infrared-labeled
secondary antibody on ODYSSEY Infrared Imaging system (LI-COR Biosciences).

98

4.4.9. Fluorospectrometry
Timer transgenic and their transgene negative littermates were dark-adapted
overnight and their retinas were removed and homogenized in buffer (80 mM Tris,
pH 8.0, 4 mM MgCl
2
) containing protease inhibitor cocktail at 1 tablet per 10 ml
buffer (Roche Diagnostics, Indianapolis, IN). Membrane and soluble fraction of
retina were separated by centrifugation at 50,000 rpm for 30 min in a Beckman TLA-
100 rotor. The pellet fractions were dissolved in solubilization buffer (20mM HEPES,
pH 7.5, 1mM MgCl
2
, 1mM CaCl
2
, 10mM NaCl, 0.1mM EDTA, 1% Dodecyle
maltoside) and shaken on a nutator for 3 hours at 4°C. After spinning for 15 min at
55,000rpm at 4°C, the supernatant part was collected. The collected soluble protein
and membrane protein fractions were measured on the fluorimeter (in Dr. Ralf
Langen’s laboratory), using 483 nm and 558 nm excitation, to measure 500-600 nm
emission respectively.

4.4.10. Cell culture and Immunocytochemistry (ICC)
Rhodopsin-Timer and WT rhodopsin sequence were cloned into pEGFP-N1
vector using BsrGI + NheI sites. Two constructs were transfected into COS cells
with Fugene6 kit (Roche). ICC was followed that WT rhodopsin were stained by
1D4 antibody and DAPI for nuclei staining. (This was done by Yukihiro Koike.)

99

4.3. Results
4.3.1. Generation of Timer transgenic mice
Timer (DsRed1-E5) fluorescence protein contains two amino acid
substitutions V105A and S197T which endow the fluorescence protein with a
distinct spectral property as well as increase of its fluorescence intensity: Timer
changes color from green to red as a function of time (Terskikh et al., 2000). Three
amino acid substitutions R2A, K5E and K9T were made to decrease protein
aggregation in DsRed2. By using three piece ligation, the non-aggregation portion of
DNA sequence from DsRed2 and the Timer-specific spectral mutations were pieced
together to make a new Timer sequence, and cloned into pEGFP-N1 plasmid where
EGFP was excised (Figure 4.1). This plasmid expression was tested in the 293 cells,
which showed changes color from green to red (data not shown).
By using RT-PCR from retinal RNA from C57Bl6 mice, rhodopsin cDNA
was amplified. It was further pieced together with Timer sequence at the end of
rhodopsin’s C-terminal by nested PCR, because it was reported C-terminal
attachment of fluorescence reporter protein has more advantages than C-terminal
insertion, e.g. higher efficency to activate transducin, to be phosphorylated by
rhodopsin kinase, better folding and normal traffic (Moritz et al., 2001; Jin et al.,
2003). After microinjecting the 6.6 kb expression construct into F1 hybrid zygotes
from C57BI/6J and DBA/2J strains, three founders were generated and confirmed by
PCR of DNA extracted from tail biopsies. Two founders produced progeny which
had fluorescent protein expression examined by wholemount and fluorescence
100

microscope. These two lines were mated with C57BL6 wildtype mice to obtain F1
and F2 transgenic progenies. The F2 were then sequentially mated with rho-/- to
generate Timer
Rho+/-
and Timer
Rho-/-
(Lem et al., 1999).

4.3.2. Rhodopsin-Timer fluorescence expression
The expression of Rhodopsin-Timer fusion protein was checked by
wholemount and vibratome sections of living tissue. Figure 4.2 showed the fusion
protein was able to traffic normally to ROS in both rho+/+ or rho+/- retinas.
Rhodopsin also changed color from green at the base of ROS to red at the apical tip
of ROS, matching aging of rhodopsin in Figure 4.3(A). However there was some
fluorescence retained in ONL shown in Figure 4.3.A-C, most likely in the membrane
of the outer nuclear layer, which might suggest that the C-terminus Timer tag might
cause some defect in trafficking or folding.
An interesting finding in Timer transgenic retina is its expression pattern in
ROS. Figure 4.2E is the magnified picture from the highlighted box in Figure 4.2D.
It clearly showed that Rho-Timer fluorescence appeared like threaded beads in ROS,
which suggests that the fusion protein was expressed/renewed in an intermittent
manner. It could even be counted as around 10 intervals. The mouse in Figure 4.3.D
was born and raised in the dark, which means that this expression pattern is
independent of light. It is well known that mice ROS turnover is around 10 days. Is
this related to the circadian rhythm? Or is it an effect of heterochromatin-associated
position-effect variegation (PEV)? We will discuss this later.
101

Figure 4.2. Timer expression in rhodopsin+/+ and +/- background seen in
wholemount of freshly dissected retinas. (A-C) Wholemount of Timer
Rho+/+
under
fluorescence microscope with 500 nm and 550 nm filter or both, respectively.
Rhodopsin-Timer fusion protein traffic normally to ROS. (D-E) In dark born and
raised Timer
Rho+/-
mice, Rhodopsin-Timer fusion protein showed similar expression
pattern as in rho+/+ background. (E) is the higher magnification of the highlighted
box from (D). It was shown that Rhodopsin-Timer fusion protein in ROS was in an
intermittent pattern, appearing like threaded beads. This existed in both rho+/+ and
+/- background in both light and dark raised animals, suggesting a light-independent
expression pattern.

102

Figure 4.3. Timer fusion proteins did not completely traffic to ROS, instead some of
them were retained in ONL. Confocol pictures of living Timer retina from vibratome
section. A.B.C. were difference panel pictures of the same Timer transgenic retina.
Magnification of confocol: 40X. Tissue was immerged in Ames’s solution. This
indicated Timer expression with the rhodopsin+/+ background was in both ROS and
ONL. It also showed that Timer fusion protein ages from base of ROS to apical tip as
the color changed from green to red.

103

4.3.3. Morphology of Timer transgenic mice was normal in rho+/+ and rho+/-
background but not in rho-/- background
Eyes from 4 weeks and 8 weeks old Timer, Timer
Rho+/-
, Timer
Rho-/-
and their
transgene negative littermates were enucleated and examined their morphology. The
4-week and 8-week-old Timer, Timer
Rho+/-
showed very similar morphology to their
transgene negative littermates WT. No retinal degeneration was seen in Figure 4.5
(A-E), with organized ROS structure and normal ONL thickness. This confirmed the
normal traffic of Rhodopsin-Timer to ROS, which helps maintain normal
morphology. It also indicates that Rhodopsin-Timer was not overexpressed.
Rhodopsin overexpression can induce photoreceptor cell death (Tan et al., 2001).
However Rhodopsin-Timer expression in rho-/- background was not able to
rescue retinal degeneration at 4 or 8 weeks. There was almost no ROS structure due
to a shortage of rhodopsin, the ROS major membrane protein (Figure 4.5.G-H). This
indicates that Rhodopsin-Timer expression is low, which fits well with our
experimental aim: observe rhodopsin turnover and trafficking without disturbing the
endogenous system by high expression of transgene.

4.3.4. Fluorescence in Timer transgenic mouse was largely from membrane
fraction
Because it was shown that part of Rho-Timer fusion proteins were retained in
the ONL, fluorimeter measurement for membrane and soluble fraction of retina was
applied. Surprisingly, the fluorescence was not detected in soluble fraction of retina
104

Figure 4.4. The morphology of Timer transgenic mice in different background rho+/+, +/-
and -/-. (A-C) Timer transgenic mice at 4weeks, 8 weeks (B-C) showed normal morphology
as WT (A). (D-E) Timer
Rho+/-
mice showed normal morphology as its negative control rho+/-
at 8 weeks. A-E suggested that Timer transgene expression did not significantly disturb
retinal morphology. It can be used as a good indicator of rhodopsin activity. (F-H) Timer
Rho-/-

showed neither ROS nor rescue of retinal degeneration from rho null. This suggested that
Rho-Timer was not able to target to ROS without presence of endogenous rhodopsin or that
its expression level is very low.

105

homogenate (Figure 4.5). This suggestes that Rhodopsin-Timer might be in the
membranous organelles instead of in cytosol in the ONL layer. In Figure 4.5, mouse
#3 didn’t show fluorescence in neither membrane nor soluble fraction, which means
that it was a false positive.

4.3.5. Rhodopsin-Timer fusion protein did not traffic to ROS in rho-/-
background
We further examined Rhodopsin-Timer fusion protein localization and
trafficking in rho-/- background (Figure 4.6), which was extremely different from its
localization in the presence of endogenous rhodopsin: the fluorescence was localized
in the perinulear region, and therewas no ROS localization (Figure 4.2). This shows
that Rhodopsin-Timer fusion protein does not target to ROS even with the presence
of intact traffic domain 1D4 at the C-terminal end. When Rho-Timer was expressed
in COS cells, it also showed mislocalization in the perinuclear area (Figure 4.6B),
different from what is seen for WT rhodopsin expression in COS cell (Figure 4.6C).

4.3.6. Rhodopsin-Timer was elusive in the Western blot
With hints from morphology of Timer
Rho-/-
that the expression level of
Rhodopsin-Timer might be low, retinas from Timer transgenic, Timer
Rho+/-
and

Timer
Rho-/-
and their transgene negative littermates were homogenized and loaded
into SDS-PAGE gel. Surprisingly, no Rhodopsin-Timer was found in any of these
samples when probed with two N-terminal and one C-terminal rhodopsin antibodies
106

Figure 4.5. Fluorescence in Timer transgenic mouse in rho+/- was from membrane
fraction. Fluoro-reading was taken at 480nm excitation and 490-600 nm emission for
green fluorescence. Mouse #3 in both membrane and soluble frantion were close to
zero which means it was a false positive transgenic moue, while mouse #1 and #2 all
showed 500 nm peak in membrane fraction after normalization by buffer background.

107

Figure 4.6. Rhodopsin-Timer showed mislocalization in rho-/- mice and COS cells.
(A) In transgenic mouse, Rhodopsin-Timer fusion protein did not traffic to ROS in
rho-/- background. The fluorescence appeared in the perinuclear area. The
Rhodopsin-Timer fluorescence expression level was many folds less than in rho+/+,
+/- background. (B) Rhodopsin-Timer was transfected and expressed in COS cells. It
also showed perinuclear localization. (C) is WT rhodopsin expressed in COS cells
without perinuclear localization.

108

4D2, R2-12N and 1D4 (data not shown). It was also reported in other Rhodopsin-
GFP transgenic frog that the transgene was not detected in Western (Moritz et al.,
2001). This might be due to the extremely low transgene expression.

4.4. Discussion
In this project, we tagged rhodopsin with fluorescence Timer, a protein that
changes color with aging, in order to monitor rhodopsin/ROS turnover rate. With this
tag, rhodopsin can be traced shortly after biogenesis to the time when it is
phagocytosed by retinal pigment epithelial cells. Two lines of Rhodopsin-Timer
transgenic mice were generated, examined and characterized, and these two lines
showed consistent properties in trafficking and expression. Three other rhodopsin
mutants from each class of mutations known to lead to ADRP were also fused with
Timer in plasmid. With the subretinal injection and electroporation technique that I
had successfully set up and practiced in the lab, these sequences can be injected and
expressed in young mice to mimic transgenic lines for future studies (Matsuda and
Cepko, 2004).
This study is still ongoing. From the current results, two interesting issues
came up, one is the protein trafficking with 1D4 ROS target signal, the other is the
rhodopsin turnover pattern.

109

4.4.1. Rhodopsin trafficking
Correct rhodopsin targeting and trafficking to the ROS is very important for
photoreceptors. For a number of rhodopsin mutants, mis-trafficking is thought to
underlie retinal degeneration (Sung et al., 1994; Li et al., 1996). Rhodopsin is first
synthesized in ER and modified in Golgi. Then it is vectorially transported long
distance from the post-Golgi network to ROS by post-Golgi vesicles (Deretic et al.,
1996). In this process, rhodopsin C-terminal binding to and recruitment of ADP-
ribosylation factor 4 (ARF4) to the trans-Golgi network are essential for post-Golgi
vesicle budding (Deretic et al., 2005). Several independent groups have indicated
that the last five amino acids of rhodopsin as the essential targeting sequence, and the
minimal essential sequence is VXPX-COOH (Deretic et al., 1996; Deretic et al.,
1998; Tam et al., 2000; Concepcion et al., 2002; Deretic, 2004; Deretic et al., 2004;
Shi et al., 2005a).
Ablation of this sorting signal causes a loss in the ability of rhodopsin to be
recognized by the sorting machinery at the trans-Golgi network. In our transgenic
mice with rho-/- background, the fusion protein Rhodopsin-Timer seems to lose its
capability to traffic to the ROS although it still bears the last eight C-terminal amino
acids. This observation is in huge contrast with the large percentage of Rhodopsin-
Timer in ROS when endogenous rhodopsin is co-expressed (Figure 4.2, Figure 4.3,
Figure 4.7). It is not surprising that a mislocalization mutant like S334ter can target
to ROS by co-transporting in the vectorial transport with presence of endogenous
rhodopsin C-terminal signal (Concepcion et al., 2002). However it is surprising that
110

Rhodopsin-Timer is incorrectly targeted, when it contains the C-terminal targeting
signal. The C-terminal signal is not sufficient to target Rhodopsin-Timer to ROS in
this case. It is possible that the Timer protein masks the 1D4 traffic domain that
normally recruits ARF4, so that Rhodopsin-Timer was not able to bud from Golgi.,
whereas Rhodopsin-Timer could be co-transported to ROS in the presence of large
quantity of endogenous rhodopsin in the rho+/+ or rho+/- background. This may
explain why the fluorescence signal assayed by fluorimetry was largely found in
membrane fraction in rho+/- background. The reason that the cytosolic fluorescence
signal was not detected is probably because of limited detection efficiency.
The C-terminus of rhodopsin has been suggested to participate in important
cellular processes distinct from phototransduction, in particular, it might be crucial
for macromolecular interaction (Deretic, 2006). A rhodopsin mutation Ter349Glu
containing an additional 51 amino acids in the C-terminus leads to rhodopsin
dysfunction and one of the most severe phenotype of ADRP (Bessant et al., 1999).
Similarly, in our transgenic mice, the large Timer protein was added, which might
impact significantly on rhodopsin function. Our data suggest that the intact
rhodopsin C-terminus is essential for its normal traffic and full function. It is also
noticeable that the expression level of the fluorescence was many folds lower in rho
-/- than in the rho+/+ or rho +/- background (Figure 4.2, 4.3., 4.6.). These results
further suggest that some functions of rhodopsin need the presence of fully intact C-
terminal sequence. The Ter349Glu rhodopsin mutant and possibly other mutants

111

might cause retinal degeneration because of interference with C-terminus function by
insertion of extra amino acids.
It is also possible that misfolding of Rhodopsin-Timer leads to retention in
the ONL. When Rho-Timer is expressed in COS cells, it showed a perinuclear
distribution, resembling inclusion bodies formed from aggregated misfolded proteins
(Figure 4.7B). It is possible that misfolded Rhodopsin-Timer may be degraded into
small pieces by autophagy or ubiquitin proteinase degradation. There was a report
that in the VPP transgenic mouse (P23H together with two other rhodopsin
mutations, V20G and P27L, in the same rhodopsin molecule. The latter two
mutations do not contribute to ADRP whereas P23H is the mis-folding rhodopsin
mutant.), VPP transported to ROS as well as to the perinuclear region, which
suggested that misfolded protein can also traffic to ROS in the presence of rhodopsin
(Wu et al., 1998). However in P23H transgenic mouse, it was reported that P23H
rhodopsin accumulated in outer nuclear layer and synaptic terminals (Roof et al.,
1994). These data present an inconsistent picture. Furthermore, misfolding was not
reported in any of previous rhodopsin-GFP transgenics or other protein-Timer
transgenics. Whether Rhodopsin-Timer is misfolded or not still needs further
investigation in the future.

4.4.2. Rhodopsin Turnover pattern in ROS
In 1960s and early 1970s, Young and colleagues developed the concept of
constant renewal and shedding of the ROS and ROS proteins, including rhodopsin
112

(Young, 1967; Hall et al., 1969; Young, 1969, 1971a; Bok and Young, 1972). It was
discovered in the late 1970s that ROS/rhodopsin renewal and shedding could be
stimulated and accelerated by environmental lighting (Basinger et al., 1976; LaVail,
1976a, b; Besharse et al., 1977). Some experiments showed that ROS renewal may
be a mechanism for adaptation to new envionmental lighting. For example, alteration
in ROS length, rhodopsin synthesis and packing density can lead to changes in
rhodopsin levels. This in turn can increase or decrease the response of photoreceptor
to light (Schremser and Williams, 1995a, b). These later discoveries demonstrated
that rhodopsin/ROS turnover does not occur at a constant rate as previously assumed;
instead it is a dynamic process that can respond to and change with environmental
cues.
In our Timer transgenic mice, although Rhodopsin-Timer fusion protein
could not traffic to the ROS by itself (Figure 4.7), it is largely located to ROS in the
presence of endogenous rhodopsin (Figure 4.2, 4.3). Thus it may still serve as a good
trace marker for the activity of the same batch of rhodopsins synthesized and
transported together. Rhodopsin-Timer showed a beaded pattern in the ROS, in both
rho+/+ and rho+/- background, in dark and light (Figure 4.2D, E). And the “bead”
number is around 10, which matches the reported rhodopsin turnover rate. Previous
studies have shown that ROS shedding follows a circadian rhythm that is
independent from suprachiasmatic nucleus (SCN), the major circadian rhythm
“clock” master for circadian behavior in mammals (Teirstein et al., 1980; Reme et al.,
1991; Terman et al., 1991; Terman et al., 1993). Furthermore, by monitoring
113

antibody-labeled rhodpsin, it was found that a diurnal rhythm of rhodopsin synthesis
occurs in the inner segment in cyclic light (Bird et al., 1988). Our results support the
notion that rhodopsin/ROS renews at a rhythmic and diurnal pace. From the dark
born and raised Timer transgenic retina, the “beaded” pattern was the same as those
in cyclic light reared mice, suggesting that there is a circadian rhythm of rhodopsin
renewal independent from light. This result is consistent with the previous finding
that ocular circadian rhythm is independent of the SCN. Our result is the direct
visualization including cells of fluorescence pattern, revealing that ROS turns over
with a diurnal/circadian rhythm.
An uneven rhodopsin expression pattern in ROS was also reported before in
Rhodopsin-GFP transgenic Xenopus laevis (Moritz et al., 2001). It also showed
fluorescence bands in ROS, however it did not follow diurnal pattern (much less than
10 bands) in fixed cryosection pictures. They speculated that the spaced fluorescence
bands in ROS originated from heterochromotin associated position-effect variegation
(PEV) as an artifact of transgenesis. Since both our two transgenic lines showed
exactly the same pattern, it is less probable that both of them have PEV at the same
time. It is possible that the retinal cryosections lost some of fluorescent signals and
resolution with fixative, as experienced from our experiment (Figure 4.7). The
fluorescence disappeared when fixed with formaldehyde and glutaldehyde in Figure
4.4.A. While the fluorescence was kept in fixative 4% formaldehyde shown in Figure
4.4.B-D. However it still lost some fluorescence as shown in 4.7B without ROS
signal.
114

Figure 4.7. Timer expression was not even in whole retina but in selected region in
cryosections. The fusion protein lost some signal in fixative like glutaldehyde, but
retained with paraformaldehyde. A-D indicated cryosection fixed at different fix
buffer and time.

115

More experiments are needed including Northern blotting to confirm RNA
level changes, Western blot using more efficient Timer antibody, in order to have
better a understanding of rhodopsin turnover.

116

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135

Abstract (if available)

Abstract Through the experiments described in this thesis, I strived to obtain a better understanding the function of rhodopsin in retinal degeneration and light adaptation. Over 100 rhodopsin mutation alleles have been associated with autosomal dominant retinitis pigmentosa (ADRP), a blindness disorder that affects one in 3000 people globally. These mutations appear to cause photoreceptor cell death through diverse molecular mechanisms. We show that Lys296Glu (K296E), a rhodopsin mutation associated with ADRP, forms a stable complex with arrestin that is toxic to mouse rod photoreceptors. This cell death pathway appears to be conserved from flies to mammals. Accumulation of stable rhodopsin/arrestin complexes in the inner segment may be an important mechanism for triggering cell death in the mammalian photoreceptor cells. Abnormal turnover of rhodopsin mutants could also underlie a mechanism leading to cell death. In order to investigate rhodopsin turnover rate, rhodopsin was tagged with a special fluorescent reporter Timer, which changes color with the function of time, to report rhodopsin turnover activity.

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Asset Metadata

Creator Chen, Jiayan (author)

Core Title New understanding of rhodopsin in retinal degeneration and high gain phosphorylation

School College of Letters, Arts and Sciences

Degree Doctor of Philosophy

Degree Program Neuroscience

Publication Date 11/22/2006

Defense Date 10/31/2006

Publisher University of Southern California (original), University of Southern California. Libraries (digital)

Tag OAI-PMH Harvest,photoreceptor,Retinal degeneration,rhodopsin

Language English

Advisor Chen, Jeannie (committee chair)

Creator Email jiayanch@usc.edu

Permanent Link (DOI) https://doi.org/10.25549/usctheses-m191

Unique identifier UC1164704

Identifier etd-Chen-20061122 (filename),usctheses-m40 (legacy collection record id),usctheses-c127-39461 (legacy record id),usctheses-m191 (legacy record id)

Legacy Identifier etd-Chen-20061122.pdf

Dmrecord 39461

Document Type Dissertation

Rights Chen, Jiayan

Type texts

Source University of Southern California (contributing entity), University of Southern California Dissertations and Theses (collection)

Repository Name Libraries, University of Southern California

Repository Location Los Angeles, California

Repository Email cisadmin@lib.usc.edu