University of Southern California Dissertations and Theses

Cone arrestin 4 contributes to vision, cone health, and desensitization of the dopamine receptor D4

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Cone arrestin 4 contributes to vision, cone health, and desensitization of the dopamine receptor D4

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Content 1

Title Page

Cone Arrestin 4 Contributes to Vision, Cone Health,
and Desensitization of the Dopamine Receptor D4

By Janise D. Deming

A Dissertation Presented to the
FACULTY OF THE USC GRADUATE SCHOOL
UNIVERSITY OF SOUTHERN CALIFORNIA
In Partial Fulfillment of the
Requirements for the Degree
DOCTOR OF PHILOSOPHY
(GENETIC, MOLECULAR AND CELLULAR BIOLOGY)
August, 2015

Copyright 2015 Janise D. Deming

2

Dedication

This work is dedicated to my loving family, including my husband Tyler. Their
encouragement and support has allowed me to become the person I am today, and I
am grateful every day for their presence in my life.
3

Acknowledgements

There are many people who contributed to the work in these pages and without
whom this dissertation would not have been possible.
First, I would like to thank my mentor Dr. Cheryl Craft for her unwavering support
and encouragement. She gave me the guidance I needed, as well as freedom,
materials, and space to pursue my project, and she never stopped believing that I could
complete this project. Thank you, Dr. Craft, for pushing me beyond my comfort zone, for
encouraging my participation in meetings and application for fellowships, and for setting
the tone for a positive lab environment where everyone feels welcome and included. It
has been a pleasure to spend the last four years in your lab.
Dr. Eun-Jin Grace Lee has been a second mentor to me for the last year, and her
input and advice has been invaluable.
Many people contributed to the work within these pages. I’d like to thank Bruce
Brown, Joseph Pak, Lawrence Rife, Yun-Sung Annette Eom, Dr. Jung-a Shin, Kayleen
Lim, and the lab teams of Dr. Machelle Pardue (Emory University) and Dr. Kathleen Van
Craenenbroeck (University of Ghent) for their significant experimental contributions to
this work. I would also like to thank Dr. Van Craenenbroeck and Dr. Pardue for their
critical reading and editing of manuscripts.
All of the members of the Mary D. Allen Laboratory for Vision Research have also
been wonderful friends and colleagues. In addition to those listed above, I’d like to thank
Erik Haw, Gerda Goette, Daphne Derose, Isabel Shen and Gloria Arciniega for keeping
me company, brightening my day, and making the lab a fun place to work. Special
thanks to Gloria for innumerable cups of tea and plates of food.
I would like to thank Ernesto Barron, Anthony Rodriguez, and Jose Gonzalez for
training and technical support using the confocal microscopes.
Thank you to Dr. Jeannie Chen and Dr. David Hinton for serving on my committee
and providing valuable insight and suggestions at each meeting.
I would like to especially thank Mrs. Dorie Miller for her generous financial support
throughout my time at USC. In addition, the Mary D. Allen Lab for Vision Research
would not exist without the generous donations of Mary D. Allen, and in loving memory I
thank her for her contributions.

4

Table of Contents

Introduction ............................................................................................ 11 Chapter 1:
1.1 G-protein Coupled Receptors and the Phototransduction Cascade .............. 11
1.2 Cone Arrestin 4 ............................................................................................ 14
1.2.1 Discovery of Cone Arrestin .................................................................... 14
1.2.3 Role in opsin shutoff .............................................................................. 15
1.2.4 Expression level and cellular localization .............................................. 19
1.2.5 Crystal structure and GPCR binding ..................................................... 20
1.2.6 Non-GPCR binding partners ................................................................. 21
1.2.7 Role in visual phenotypes ..................................................................... 23
1.3 Dopamine Receptor D4 ................................................................................. 25
1.3.1 The functions of dopamine and its receptors ......................................... 25
1.3.2 Function of D2-like receptors and DRD4 ............................................... 26
1.3.3 DRD4 polymorphisms and their relation to disease ............................... 28
1.3.4 The importance of DRD4 in photoreceptors and in vision ..................... 29
1.3.5 The shutoff of DRD4 .............................................................................. 30
Visual Cone Arrestin 4 Contributes to Visual Function and Cone Health .. Chapter 2:
............................................................................................................... 32
2.1 Introduction ................................................................................................... 32
2.2 Materials and Methods .................................................................................. 34
2.3 Results .......................................................................................................... 40
2.3.1 OKT studies reveal decreased visual acuity and contrast sensitivity in
Arr4
-/-
mice ...................................................................................................... 40
2.3.2 Physiological response of young Arr4
-/-
mice is abnormal compared to
WT .................................................................................................................. 41
2.3.3 Immunoblot analysis reveals a decrease in M- and S-opsin expression in
older Arr4
-/-
mouse retinas .............................................................................. 43
2.3.4 M-opsin expression is increased in the inferior retina of young Arr4
-/-

mice, but in older mice is similar to WT .......................................................... 44
2.3.5 M- and S-opsin cone numbers decrease with age in Arr4
-/-
mouse retinas
....................................................................................................................... 47
2.4 Discussion ..................................................................................................... 50

5

Characterization of Antibodies to Identify Cellular Expression of Chapter 3:
Dopamine Receptor D4 ...................................................................................... 56
3.1 Introduction ................................................................................................... 56
3.2 Materials and Methods .................................................................................. 58
3.3 Results .......................................................................................................... 61
3.3.1 Immunohistochemistry: Transfected Cells ............................................. 61
3.3.2 Immunoblots .......................................................................................... 61
3.3.3 Immunohistochemistry: mouse retinas .................................................. 63
3.3.4 Permanently transfected cell lines ......................................................... 64
3.4 Discussion ..................................................................................................... 65
Dopamine Receptor D4 Internalization Requires a Beta-Arrestin and a Chapter 4:
Visual Arrestin ..................................................................................................... 67
4.1 Introduction ................................................................................................... 67
4.2 Materials and Methods .................................................................................. 70
4.3 Results .......................................................................................................... 79
4.3.1 β-arrestins and visual arrestins are co-expressed in mouse retinas ...... 79
4.3.2 DRD4 is expressed in mouse cone photoreceptors .............................. 82
4.3.3 Co-IP of DRD4 with ARR4 .................................................................... 84
4.3.4 Internalization of DRD4 requires two different arrestins ........................ 85
4.3.5 Translocation of ARR4 to plasma membrane ........................................ 90
4.4 Discussion ..................................................................................................... 93
4.5 Conclusion .................................................................................................. 102
Conclusions .......................................................................................... 104 Chapter 5:
References ........................................................................................... 107 Chapter 6:

6

List of Tables
2.1 Mouse strains used in Arr4
-/-
studies
2.2 Primary antibodies used in Arr4
-/-
studies
3.1 Anti-DRD4 antibodies characterized
4.1 Mammalian expression plasmids used in DRD4 desensitization studies
4.2 Mouse strains used in DRD4 desensitization studies
4.3 Primary antibodies used in DRD4 desensitization studies
List of Figures
1.1 Summary of GPCR activation and desensitization
1.2 Cone recovery in visual arrestin knockout mouse models
1.3 Light-dependent translocation of ARR4 to cone outer segments
1.4 Crystal structure of salamander cone arrestin
1.5 Contrast sensitivity of zebrafish larvae lacking cone arrestin (Arr3a)
1.6 Drd4 mRNA expression in rat retina and pineal gland
1.7 Drd4 mRNA expression in mouse retina and daily cAMP accumulation in WT
and Drd4
-/-
mice
2.1 Contrast sensitivity and visual acuity in young and old WT and Arr4
-/-
mice
2.2 ERG single-flash recording from young and old WT and Arr4
-/-
mice
2.3 ERG flicker recording from young and old WT and Arr4
-/-
mice
2.4 Immunoblot of retinal M- and S-opsin expression in young and old WT and Arr4
-/-

mice
2.5 IHC of retinal M-opsin expression in young and old WT and Arr4
-/-
mice
2.6 IHC of retinal S-opsin expression in young and old WT and Arr4
-/-
mice
7

2.7 Number of M-cones and S-cones in retinal superior and inferior regions in young
and old WT and Arr4
-/-
mice; ONL thickness in young and old WT and Arr4
-/-
mice
3.1 IHC of DRD4 in HEK 293T comparing multiple commercial antibodies
3.2 Immunoblot of DRD4 in HEK 293T and mouse retinas comparing multiple
commercial antibodies
3.3 IHC of DRD4 in mouse retinas comparing multiple commercial antibodies
3.4 IHC of DRD4 in mouse retinas using the best candidate, SC N-20
3.5 IHC of HA-or FLAG-tagged DRD4 in permanently transfected HEK 293
4.1 Summary of internalization measurement methods
4.2 Immunoblot of β-arrestins in mouse retina; IHC of beta-arrestins in mouse retina
4.3 IHC of visual arrestins and beta-arrestins in mouse retina
4.4 IHC of DRD4 in HEK 293T and mouse photoreceptor inner segments
4.5 Co-IP immunoblot of ARR4 with HA-DRD4.4
4.6 IHC of HEK 293T with FLAG-DRD4.4 co-transfected with one arrestin
4.7 IHC of HEK 293T with FLAG-DRD4.4 co-transfected with two arrestins
4.8 Alternate methods of quantification: flow cytometry, in-cell western
4.9 IHC of FLAG-DRD4 and ARR4 in HEK 293T stimulated with DA

8

List of Abbreviations
A2AR Adenosine receptor 2A
AAV Adeno-associated virus
AC Adenylate cyclase
ADHD Attention deficit hyperactivity disorder
ANOVA Analysis of variance
ARR1 Arrestin 1
Arr3a Zebrafish cone arrestin 3a
Arr3b Zebrafish cone arrestin 3b
ARR4 Arrestin 4 (cone arrestin)
Arr-DKO Visual arrestin double knockout (Arr1
-/-
Arr4
-/-
)
B2AR Beta 2 adrenergic receptor
β-ARR1 β-arrestin 1 (arrestin 2)
β-ARR2 β-arrestin 2 (arrestin 3)
c/d Cycles/degree
cAMP Cyclic AMP (adenosine monophosphate)
CAR Cone arrestin (Arr4)
cDNA Coding DNA (deoxyribonucleic acid)
cd-s/m
2
Candela-seconds per square meter
CIS Cone inner segment
Co-IP Co-immunoprecipitation
COS Cone outer segment
CT Circadian time
Cx36 Connexin 36
DA Dopamine
DAPI 4',6-diamidino-2-phenylindole (nuclear stain)
deg/s Degrees/second
DMEM Dulbecco’s modified eagle’s medium
DR Dopamine receptor
DRD4 Dopamine receptor D4
9

EMEM Eagle’s minimal essential medium
ERG Electroretinography
FBS Fetal bovine serum
Fc Foot candle
GC1 Guanylate cyclase 1
GCL Ganglion cell layer
GDP Guanosine diphosphate
GPCR G-protein coupled receptor
GRK G-protein coupled receptor kinase
GTP Guanosine triphosphate
HEK Human embryonic kidney (cell line)
HRP Horseradish peroxidase
Hz Hertz
IB Immunoblot
IHC Immunohistochemistry
INL Inner nuclear layer
IPL Inner plexiform layer
IS Inner segment
Jnk3 c-Jun N-terminal kinase 3
kDa kiloDalton
L-cone Long-wavelength sensitive cone
L-DOPA L-3,4-dihydroxyphenylalanine (DA precursor)
LSM Laser scan microscope
M-cone Middle-wavelength sensitive cone
M-opsin Middle-wavelength sensitive opsin
mRNA Messenger RNA (ribonucleic acid)
MW Molecular weight
Nrl Neural retina leucine zipper
NSF N-ethylmaleimide Sensitive Factor
OCT Optimal cutting temperature medium
OKT Optokinetic tracking
10

ON Optic nerve
ONL Outer nuclear layer
OPL Outer plexiform layer
OS Outer segment
PAGE Poly-acrylamide gel electrophoresis
PBS Phosphate buffered saline
PCR Polymerase chain reaction
PFA Paraformaldehyde
P-Rh* Phosphorylated, light activated rhodopsin (Rh)
qRT-PCR Quantitative real-time PCR
RIPA Radio-immunoprecipitation assay (buffer)
rTHKO Retina-specific tyrosine hydroxylase knockout
SAG S-antigen (arrestin 1)
SDS Sodium dodecyl sulfate
SNP Single nucleotide polymorphism
S-cone Short wavelength-sensitive cone
S-opsin Short wavelength-sensitive opsin
TBS Tris buffered solution
TH Tyrosine hydroxylase
TUNEL Terminal deoxynucleotidyl transferase-mediated dUTP nick-end labeling
VNTR Variable number of tandem repeats
WT Wild-type

11

Introduction Chapter 1:
1.1 G-protein Coupled Receptors and the Phototransduction Cascade
The process of phototransduction is an intricate cascade of intracellular signaling
and cell-to-cell communication that begins in the retina, exits the eye through the optic
nerve, and ends in the visual processing centers of the brain. Light is first detected by
photoreceptor cells, of which there are two types: rods, which are primarily used in low-
light conditions; and cones, which are essential for high acuity vision in bright ambient
light. The photoreceptors send the light signal to downstream cells in the retina,
including bipolar cells and ganglion cells, before the signal travels to the brain.
Within the photoreceptors, the phototransduction process is a molecular signaling
cascade. First, a photon traveling into the retina hits a chromophore called opsin.
Opsins, including rhodopsin and the cone opsins, are members of a large protein family
called G-Protein Coupled Receptors (GPCRs). The GPCR superfamily consists of over
800 proteins in humans (Fredriksson et al., 2003), and they have wide biological
functions, including taste, olfaction, immune response, and sympathetic and
parasympathetic signaling (Pierce et al., 2002). They are seven transmembrane pass
proteins that transduce a signal from the outside of the cell to the inside by interacting
with an extracellular ligand and an intracellular heterotrimeric G-protein. GPCRs are
well known for their medical applications, and it is estimated that 40% of currently
available drugs target GPCRs (Tyndall and Sandilya, 2005; Lagerstrom and Schioth,
2008).

12

GPCRs work by a well-defined canonical mechanism (Figure 1.1) reviewed in
(Kroeze et al., 2003) in which ligand binding (for the opsins, the “ligand” is a photon)
induces a conformational change in the GPCR that allows it to interact with a
heterotrimeric G-protein. Before this interaction, the G-protein is bound to guanosine
diphosphate (GDP) and is anchored to the cell membrane. The ligand-bound GPCR
acts as a guanine nucleotide exchange factor for the G-protein, allowing GDP to
Figure 1.1 Summary of GPCR activation and desensitization. Before activation, the
GPCR and heterotrimeric G-protein (bound to GDP) do not interact. When an
agonist, or ligand, binds to the extracellular portion of the receptor, it induces a
conformational change in the receptor which allows it to act as a guanine exchange
factor for the G-protein. The G-protein swaps GDP for GTP, the α subunit
dissociates from the β/ γ dimer, and they each act downstream. The GPCR continues
to activate G-proteins until shutoff. The shutoff pathway begins with GRK
phosphorylation of the intracelllular portion of the GPCR, which can only take place
after anonist binding. After phosphorylation, arrestin binds to the phosphorylated
GPCR, preventing further G-protein activation. Often, arrestin binding leads to
internalization of the GPCR for recycling or degradation. Based on a figure from
Lefkowitz and Shenoy, 2005, Science.
13

dissociate from the G-protein and guanosine triphosphate (GTP) to bind in its place.
When GTP binds the heterotrimeric G-protein, the alpha subunit of the G-protein
dissociates from the beta and gamma subunits, travels into the cytoplasm, and
continues the downstream signaling pathway. Each GPCR can activate many G-
proteins of the same type, amplifying the signal until the GPCR is inactivated.
The inactivation pathway is important for termination of the GPCR signal. It begins
with phosphorylation of the intracellular portion of the GPCR by a G-protein coupled
receptor kinase (GRK). Once the GPCR has been phosphorylated, a protein called
arrestin binds the GPCR, preventing it from activating any additional G-proteins. Many
GPCRs are then internalized into the cell for degradation or to be recycled back to the
cell membrane (Ferguson, 2001; Shenoy and Lefkowitz, 2003).
Arrestins compose a family of four proteins that are essential for desensitization of
numerous GPCRs. In mammals, there are two visual arrestins, Arrestin 1 (ARR1, also
called S-Antigen or 48 kDa protein) (Wacker et al., 1977; Kuhn et al., 1984; Pfister et
al., 1985) and Arrestin 4 (ARR4, also called cone arrestin [CAR], X-arrestin, or ARR3 in
the NCBI gene nomenclature) (Murakami et al., 1993; Craft et al., 1994). Previous work
clearly demonstrated the functional roles of visual arrestins in mouse models in which
these two genes are individually or simultaneously ablated (Arr1
-/-
, Arr4
-/-
, or Arr1
-/-
Arr4
-/-
) (Chen et al., 1999; Nikonov et al., 2008; Brown et al., 2010; Huang et al., 2010).
The other two arrestins are called beta-arrestins ( β-ARR1 and β-ARR2), after the
beta-adrenergic receptors with which they were first shown to interact (Lohse et al.,
1990). They are expressed ubiquitously and interact with many different GPCRs,
shutting off their G-protein signal and sometimes activating downstream pathways of
14

their own (Lefkowitz and Shenoy, 2005). They are also responsible for mediating the
process of GPCR internalization, participating in the recruitment of components of the
cell’s endocytotic machinery (Lefkowitz and Shenoy, 2005; Gurevich and Gurevich,
2006).
1.2 Cone Arrestin 4
1

1.2.1 Discovery of Cone Arrestin

After the molecular identification of S-antigen, which was later renamed “rod”
Arrestin 1 (ARR1) (Shinohara et al., 1987), two ubiquitously expressing β-arrestins were
cloned using the original cDNA encoding ARR1 (Lohse et al., 1990; Benovic et al.,
1990). At this time, there were hints that other arrestins existed and were expressed in
red/green cone photoreceptors and pinealocytes, based on immunohistochemical and
immunoblot analysis with a panel of S-antigen monoclonal antibodies that recognized
ARR1 in all rods and only blue cones (Craft et al., 1990; Nir and Ransom, 1992). These
observations led to the eventual discovery of the fourth arrestin, which was
independently identified using two distinct cloning strategies. The first approach
employed a technique that identified genes on the X chromosome specifically
expressed in the retina using a retinal cDNA library and northern blot screen analysis.
Based on the sequence similarity to ARR1, this arrestin was named X-arrestin
(Murakami et al., 1993). Simultaneously, Craft, Whitmore and Wiechmann identified and
characterized the family of arrestins using a pineal gland cDNA expression library by
targeting an epitope-domain shared anchor of the three known arrestins in a novel
polymerase chain reaction (PCR) approach (Craft et al., 1994). Their cDNA encoded an
1
Large portions of this work (Section 1.2) were previously published in (Craft and Deming,
2014). Re-used with permission from Springer.
15

arrestin-like protein, which was localized with a hybridoma panel to the human
chromosome Xq13.1. Based on in situ hybridization studies, the cellular expression
pattern of the transcript demonstrated that it was highly enriched in pinealocytes and
cone photoreceptors, and was named “cone arrestin” (Arrestin 4, ARR4) (Craft et al.,
1994).
The development of specific antibodies for cone arrestin helped to clarify the cellular
expression pattern of ARR4 (Zhu et al., 2002). The “cone arrestin” was shown to be
specifically expressed in all cones; however, it has reduced expression in blue cones. In
contrast, “rod” ARR1 is highly expressed in all rod photoreceptors but is only expressed
in blue cones and not red/green cones in Macacas and human retinas (Zhang et al.,
2001; Craft et al., 2014).
1.2.3 Role in opsin shutoff
Because ARR4 has high amino acid sequence similarity to ARR1 and is highly
enriched in cone photoreceptors, researchers hypothesized that ARR4 acts in a similar
physiological manner to ARR1. Instead of interacting with rhodopsin, the hypothesis
was that ARR4 binds to light-activated, phosphorylated cone opsins, and subsequently
desensitizes them. This binding would prevent phosphatase 2A from dephosphorylating
the opsin complex and allowing it to be reactivated. However, until a decade after the
initial discovery of ARR4, there was insufficient evidence to support this hypothesis.
Because of the rod dominance of the mouse retina, it was difficult to isolate cone
photoreceptors and determine if ARR4 was involved in cone pigment shutoff, and how
that involvement occurred. In 2002, using retinas isolated from the neural retina leucine
zipper knockout mouse (Nrl
-/-
) developed by Swaroop and colleagues (Mears et al.,
16

2001), in which the rod progenitor cells develop into an enhanced S-cone phenotype,
Craft and her collaborators observed high expression of cone-specific genes. Using
immunoprecipitation, in vitro phosphorylation, and isoelectric focusing studies, they
verified ARR4 binding was specific to light-activated, G-protein receptor kinase 1-
(GRK1) phosphorylated S- and M-opsins in mice (Zhu et al., 2003). Backcrossing Nrl
-/-

with Grk1
-/-
mice to create double knockout mice, they revealed that when GRK1 is
absent, the cone pigments are not phosphorylated and ARR4 is unable to bind them in
a light-dependent manner. This was the first clear evidence that ARR4 acts in the way
that had been hypothesized since its discovery. Additional in vitro studies suggested
ARR4 participated in binding to light-activated phosphorylated cone opsins (Sutton et
al., 2005). However, it still did not show that ARR4 is required for the cone pigment
shutoff, but only that it binds to the cone opsins after they were light-activated and
subsequently phosphorylated by GRK1.
Craft and Pugh collaborated to clarify the contribution of ARR4 to cone pigment
shutoff utilizing the ARR4 knockout mouse (Nikonov et al., 2008). To their surprise, their
initial studies revealed no significant difference in the Arr4
-/-
cone pigment shutoff
response compared to the control in native murine cones. Previous studies had
demonstrated that in a transgenic mouse model where the cone arrestin was driven by
the rhodopsin promoter to be highly expressed in rods, ARR4 could only partially rescue
the light-induced rod degeneration and activated rhodopsin shutoff and recovery in Arr1
-
/-
retinas (Chan et al., 2007). ARR4 is expressed in cone photoreceptors and
pinealocytes, and ARR1 was discovered to be highly expressed in all mouse rods and
co-expressed with ARR4 in cones (Nikonov et al., 2008). They hypothesized that ARR1
17

may contribute to the cone pigment shutoff. To test this, they employed single (Arr1
-/-
,
Arr4
-/-
) and double (Arr1
-/-
Arr4
-/-
) knockout mice to determine if one or both visual
arrestins were necessary and sufficient for normal cone pigment shutoff. Using
electrophysiological patch recording from single cones of normal control mice, they
showed that after a bright light stimulus, there is essentially no response difference in
the cone recovery time between WT, Arr1
-/-
, and Arr4
-/-
. In contrast, Arr1
-/-
Arr4
-/-
double
knock-out (Arr-DKO) response had a significantly longer recovery time compared to the
single arrestin knockout genotypes (Figure 1.2) (Nikonov et al., 2008).
Further experiments probed the time course of phototransduction activated by S-
and M-cone opsins, respectively. Previously, it was observed that in a “dim-flash”
response of 360 nm and 510 nm light, the cone response has a linear relationship to
flash intensity and can independently be evaluated (Nikonov et al., 2006). Surprisingly,
the Arr-DKO cones exhibited a similar waveform response to the other genotypes until
they achieve 60% of their recovery to baseline. At this point, the recovery response of
the Arr-DKO cone diverged from the other groups, exhibiting a much slower tail phase
than the others, regardless of whether S- or M-opsin was activated by the flash
(Nikonov et al., 2008). Therefore, the normal inactivation of each isomerized S- or M-
opsin molecule requires at least one visual arrestin (ARR1 or ARR4) after a strong
bright light stimulus. This avoidance of saturation in steady illumination implies that the
phosphodiesterase activity generated by each photoisomerized cone opsin is
prolonged. Thus, the current state of ARR4 research indicates that ARR4 binds to and
desensitizes light-activated, phosphorylated cone pigments; however, ARR1 fulfills a
similar functional role if ARR4 is absent.
18

Figure 1.2 S-cone recovery is delayed in Arr-DKO mice, but not Arr1
-/-
or Arr4
-/-
.
A,D,G, and J represent ERG flash responses to 361 nm light (S-cone dominant) at
multiple intensities. B,E,H, and K magnifies the responses to the dimmest flashes
on a smaller time scale. C,F,I, and L plot the intensity of the flash compared to the
amplitude of response (top) and time to 40% recovery from saturation (bottom).
Re-used with permission from Elsevier; original figure appeared in Nikonov et. al.
2008, Neuron.
19

1.2.4 Expression level and cellular localization
The concentration of the visual arrestins in dark-adapted cones was measured and
compared to previous studies to reveal that the total quantity of ARR4 protein in each
cone is close to the quantity of cone opsin protein (Nikonov et al., 2008). In this study,
ARR1 expression in mouse cones was shown to be approximately 50-fold higher than
that of ARR4 (see Table 1, supplemental data) (Nikonov et al., 2008). By
immunohistochemical localization using specific antibodies unique for mouse ARR4
(LUMIj-mCAR), ARR4 is expressed in several cone photoreceptor cellular
compartments before and after light exposure (Zhu et al., 2002). Similar to ARR1
translocation studies in rod photoreceptors (Broekhuyse et al., 1985; Whelan and
McGinnis, 1988), ARR4 undergoes a light-dependent translocation from the cone
pedicles, cell bodies, and inner segments to the cone outer segments (Figure 1.3) (Zhu
et al., 2002). Similar light/dark ARR4 translocation studies have been performed using
bovine cone photoreceptors with the 7G6 monoclonal antibody, which also recognizes
cone arrestin (Zhang et al., 2003a). However, the translocation of ARR4 is not as robust
as that of ARR1; even after bright light exposure, a residual amount of ARR4 remains in
the cone pedicle, while ARR1 nearly completely translocates to the outer segments
(Zhu et al., 2002). In Grk1
-/-
mice, ARR4 translocation to outer segments is light-
dependent, even though the cone opsins remain unphosphorylated in this model (Zhang
et al., 2003b). This implies that the classical “on” pathway through the opsins to alpha-
transducin is not required for ARR4 translocation, and there is likely to be another light-
dependent pathway driving the translocation of ARR4. It has also been shown that light-
dependent ARR4 translocation does not take place in Guanylate Cyclase 1 knockout
20

(Gc1
-/-
) mice; however, ARR4 translocation can be restored when Gc1
-/-
mice are
treated with an adeno-associated virus (AAV) which replaces the Gc1 gene (AAV-GC1),
which rescues guanylate cyclase 1 cone function (Coleman and Semple-Rowland,
2005).
1.2.5 Crystal structure and GPCR binding
The generation of a crystal structure of the ARR4 protein allowed further
understanding of the functions of ARR4. In 2005, a crystal structure of the salamander
cone arrestin was solved (Figure 1.4) (Sutton et al., 2005). It was similar to the other
arrestin structures that were previously identified, having the canonical arrestin fold

Figure 1.3 Cone arrestin (red) is present throughout the cones in both light and
dark conditions. In light-adapted retinas, cone arrestin immunofluorescent signal is
higher in cone outer segments (COS) than cone inner segments (CIS), indicating a
light-dependent translocation of the arrestin. Re-used with permission from
Molecular Vision; original figure appeared in Zhu et. al. 2002, Molecular Vision.
21

consisting of two domains, each containing a β-strand “sandwich.” The β-strand
sandwich consists of two β- sheets joined by hydrophobic interactions. There was also a
single α-helix in the amino terminal (N)-domain. The investigators explored the binding
selectivity of ARR4 compared to ARR1 and β-ARR1. While ARR1 is highly selective for
light activated, phosphorylated rhodopsin (P-Rh*), and β-ARR1 is able to bind many
GPCRs, ARR4 has an intermediate binding selectivity. Its highest binding affinity was
for human green cone opsin (GCO), but it also was able to bind to the m2 muscarinic
cholinergic receptor, indicating that it may also be able to bind to other non-opsin
GPCRs. Thus, while the molecular structural details of ARR1 function have been well-
characterized, there is still much to discover regarding ARR4 and its function in cone
photoreceptors and pinealocytes.
While the β-arrestins have high amino acid sequence similarity to one another (76%
identical), the visual arrestins are less similar to each other (58% identical). ARR4
shares the same degree of similarity to β-ARR2 as to ARR1 (58%) (Craft and Whitmore,
1995). Perhaps this similarity to the β-arrestins is what confers ARR4 with its binding
capacity for GPCRs other than the opsins, while ARR1 maintains a very high preference
for P-Rh*.
1.2.6 Non-GPCR binding partners
In an in vitro proteomic study in cultured HEK 293 cells, β-arrestins were shown to
interact with both visual arrestins after stimulation with the beta-adrenergic agonist,
isoproterenol (Xiao et al., 2007). So far, no evidence exists that the heteromerization of
β-arrestins and visual arrestins has any functional significance, but they may work
22

synergistically and in conjunction with one another, leading to an intriguing, unexplored
area of inquiry.
As with the other arrestins, ARR4 has multiple confirmed non-receptor binding
partners, including c-Jun N-terminal kinase (JNK3) and E3 ubiquitin ligase MDM2.
ARR4 works together with these proteins to regulate their subcellular localization and
relocalize them from the nucleus to the cytoplasm (Song et al., 2007). Both of these
proteins can also bind the other arrestins to serve as scaffolds and recruitment modules

Figure 1.4 Crystal structure of salamander cone arrestin. A. The N (amino)-
domain is yellow and the C (carboxy)-domain is magenta. α-helix H1 is orange,
and the phosporylated-GPCR-specific binding region (including R166 and D287) is
blue. B. The structure of cone arrestin (red) is overlaid with that of visual arrestin 1
(green) and β-arrestin 1 (blue). Re-used with permission from Elsevier; original
figure appeared in Sutton et. al. 2005, Journal of Molecular Biology.
23

(Shenoy et al., 2001; Lefkowitz and Shenoy, 2005). Using a cell-based assay, Song and
collaborators identified individual N- and C-domains of cone and rod arrestins that
contain elements to bind JNK3 and to remove it from the nucleus. Unlike the N-domain
interaction of β-ARR2, MDM2 preferentially interacts with full-length ARR4 in the
“frozen” basal configuration, which mimics the conformation of free (GPCR unbound)
ARR4. Their ARR4 studies set the stage to analyze the precise identification of Jnk3
and MDM2 binding sites by site-directed mutagenesis (Song et al., 2007).
In yeast two hybrid screens of retinal cDNA libraries, other potential interactions
between ARR4 and novel candidates were identified, including RND2 (Zuniga et al.,
2002) and a cilia protein, ALS2CR4 (hypothetical protein FLJ33282) which was
discovered to be a transmembrane cilia protein and renamed TMEM 237 (Zuniga and
Craft, 2010). RND2 belongs to a family of small GTP-binding proteins that alter many
important cellular functions by affecting the actin cytoskeletal structure and stability
(Nishi et al., 1999). TMEM 237 is involved in the cilia transition zone and a gene defect
contributes to Joubert syndrome (Huang et al., 2011).
ARR4 is highly expressed in cones and pinealocytes, and it is reasonable to predict
that it is participating in other cellular pathways besides opsin pigment shutoff in these
cell types. The interactions with other proteins could be responsible for maintaining the
presence of ARR4 in the cone pedicle after light exposure (Zhu et al., 2002).
1.2.7 Role in visual phenotypes
Zebrafish studies have also provided evidence of the physiological role of ARR4 in
vision. Zebrafish have two genes orthologous to mouse Arr4, which are called Arr3a
and Arr3b. Unlike mouse cones, which express both visual arrestins, zebrafish cone
24

photoreceptors only express one visual arrestin per cone. M- and L-cones express
Arr3b, while S-cones express exclusively Arr3a. Zebrafish cones do not express the
ARR1 ortholog, ArrS (Renninger et al., 2011). In zebrafish larvae, which have a cone-
dominant retina, morpholino knockdown of Arr3b causes a delay in M- and L-cone
photoreceptor recovery. Because of technical limitations, S-cone photo-receptor
recovery could not be measured, but the group hypothesized that Arr3a is required for
S-cone recovery. Using optokinetic experiments, Arr3b was also shown to be necessary
for high temporal resolution in the L- and M-cones (Figure 1.5) (Renninger et al., 2011).
Mouse models utilizing the visual arrestin knockouts have shown a similar result.
Arr4
-/-
mice have a significant decrease in contrast sensitivity compared to Arr1
-/-
or
controls (Brown et al., 2012 53:ARVO E-Abstract 760/A637; Deming et al., 2015b).
Thus, although ARR1 can substitute for ARR4 in cone pigment shutoff, it may not be
able to substitute all of the functional roles that ARR4 has in cones. These other roles
are still under investigation, but the existence of an Arr4
-/-
mouse will allow further
characterization of the visual transduction pathways in which ARR4 is involved.

Figure 1.5 Contrast sensitivity in zebrafish larvae, measured through optokinetic
eye movements, is dependent on Arr3a, the cone arrestin specific for red and green
cones in zebrafish. Re-used with permission from John Wiley and Sons; original
figure appeared in Renninger et. al. 2008, European Journal of Neuroscience.
25

1.3 Dopamine Receptor D4
1.3.1 The functions of dopamine and its receptors
Dopamine (DA) is a catecholamine produced in neurons containing tyrosine
hydroxylase, the rate-limiting enzyme in its synthesis process. Dopamine has important
functions in many aspects of cellular signaling, including vision. Dopamine acts as a
ligand for five different GPCRs, named Dopamine Receptor (DR) D1 through D5
(O'Dowd, 1993; Gingrich and Caron, 1993; Seeman and Van Tol, 1994). The dopamine
receptors are classified into two groups that have opposite downstream effects in the
cells in which they are expressed. The D1-like subgroup consists of DRD1 and DRD5
and activates adenylate cyclase (AC), thereby increasing the concentration of cyclic
AMP (cAMP) in the cell. The D2-like subgroup, including DRD2, DRD3, and DRD4, acts
to inhibit AC, effectively decreasing cAMP concentration in the cell.
The production of dopamine is under circadian and light control, so the neurons that
produce DA rely on signals from the visual system to regulate their production of DA
(Iuvone et al., 1978). DA is produced in multiple areas of the brain and in a subset of
amacrine cells in the retina (Haeggendal and Malmfors, 1963). Humans who are
dopamine-deficient (as in Parkinson’s disease) display visual defects as well as the
more obvious motor symptoms of dopamine deficiency (Bodis-Wollner, 1990; Muller et
al., 1997; Bodis-Wollner, 2003; Laatu et al., 2004). They tend to have problems with
contrast sensitivity and color vision, and L-DOPA therapy can partially restore their color
vision (Buttner et al., 1994).

26

1.3.2 Function of D2-like receptors and DRD4
D2-like receptors are important for memory recording and cognition in the prefrontal
cortex and hippocampus (Tritsch and Sabatini, 2012). Stimulation of the D2-like
receptors is also related to neurite outgrowth, indicating an important role in
development and neuronal connectivity (Todd, 1992).
Antagonists of the D2-like receptors are used as anti-psychotics (Dean and Scarr,
2004; Newman-Tancredi and Kleven, 2011; Seeman, 2014). In particular, clozapine is a
powerful drug for schizophrenia (Rodova et al., 1973; Faltus et al., 1973; Gerlach et al.,
1974). DRD4 has a particularly high affinity for clozapine (Van Tol et al., 1991), but it is
still unclear whether DRD4 is the primary actor in the effects of clozapine or whether its
effects on DRD2 or DRD3 are responsible for its anti-psychotic properties (Sanyal and
Van Tol, 1997; Wong and Van Tol, 2003).
Agonists for the D2-like receptors are used as drugs for Parkinson’s disease, either
alone or in combination with L-DOPA (Lieberman et al., 1987; Lees, 1993; Jenner,
1995; Rascol, 1999; Montastruc et al., 1999; Gottwald and Aminoff, 2008). It is clear
that modulation of the D2-like receptors is under careful control in adults without these
neurological disorders. Too much or too little D2-like signaling can cause problems in
humans, so the on- and off-pathways controlling the downstream signaling of these
GPCRs are important areas of scientific study.
DRD4 expression is highest in the retina (Cohen et al., 1992; Klitten et al., 2008)
and pineal gland (Figure 1.6) (Matsumoto et al., 1995; Bai et al., 2008; Kim et al., 2010;
Gonzalez et al., 2012), but its expression is also present in significant amounts in the
prefrontal cortex, hippocampus, amygdala, hypothalamus (Meador-Woodruff et al.,
27

1994; Meador-Woodruff et al., 1995; Meador-Woodruff et al., 1996; Meador-Woodruff et
al., 1997; Ariano et al., 1997; Lidow et al., 1998). However, it is not restricted to the
nervous system; DRD4 expression has also been observed in the heart, kidney, and
lymphocytes (O'Malley et al., 1992; Bondy et al., 1996; Sun et al., 1998; Ricci et al.,
1998a; Ricci et al., 1998b; Amenta et al., 1999).
In recent years, the homo- and hetero-dimerization of GPCRs has been gaining
attention as having functional importance and as possible drug targets (reviewed in
(Ferre et al., 2010)). DRD4 has been reported to form homodimers (Van Craenenbroeck
et al., 2011), and this occurs in the endoplasmic reticulum, assisting in receptor folding
and targeting to the plasma membrane. It has also been shown to form heterodimers
with DRD2 (Borroto-Escuela et al., 2011).Although controversial, it has been shown in

Figure 1.6 Drd4 mRNA expression in rats is highest in the pineal gland and retina,
and that expression is circadian (A and B). Compared to Drd1 and Drd3 (C) or Drd1
and Drd5 (D), Drd4 is the most highly expressed of the three during night time in the
retina and pineal gland. Re-used with permission from Elsevier; original figure
appeared in Kim et. al. 2010, Molecular and Cellular Endocrinology.
28

vitro that DRD4 is able to heterodimerize with the adrenergic receptors α
1 β
and β
1
, and
this has functional significance in the pineal gland, where it is hypothesized that these
heteromers form in a circadian fashion and regulate melatonin synthesis (Gonzalez et
al., 2012).
1.3.3 DRD4 polymorphisms and their relation to disease
In humans, there are many polymorphisms of the DRD4 gene, in which between 2
and 11 imperfect repeats of a 16-amino acid sequence in the third intracellular loop of
the receptor are present (Van Tol et al., 1992; Lichter et al., 1993). These are referred
to as variable numbers of tandem repeats (VNTR), and many studies have been
conducted to determine the phenotypes associated with different VNTR genotypes. The
most common human alleles contain 2 or 4 repeats, named DRD4.2 and DRD4.4.
Longer VNTR alleles (7-11 repeats) have been associated with higher rates of novelty-
seeking (Ebstein et al., 1996; Benjamin et al., 1996; Pogue-Geile et al., 1998; Ray et al.,
2009) and an increased risk of attention deficit hyperactivity disorder (ADHD) (LaHoste
et al., 1996; Smalley et al., 1998; Holmes et al., 2000; Muglia et al., 2000; Sunohara et
al., 2000; Curran et al., 2001; Holmes et al., 2002; Tovo-Rodrigues et al., 2013), drug
and alcohol abuse (George et al., 1993; Kotler et al., 1997; Comings et al., 1999;
Hutchison et al., 2002; Shao et al., 2006; Chien et al., 2010), and overeating and
obesity (Poston et al., 1998; Levitan et al., 2004; Levitan et al., 2006; Ariza et al., 2012).
Although some studies have been unable to replicate these findings (Hinney et al.,
1999; Li et al., 2000; Mill et al., 2001; Todd et al., 2001; Luciano et al., 2004; Bakker et
al., 2005), it is generally accepted that the DRD4 gene locus has an association with at
least some of these traits, if not all of them.
29

Polymorphisms within the DRD4 gene promoter have also been associated with
disease and personality traits. Two single nucleotide polymorphisms (SNPs) at the -521
and -616 locations and a 120 base pair repeat in the promoter region of the DRD4 gene
have in some studies been shown to correlate with novelty seeking behavior (Ronai et
al., 2001; Schinka et al., 2002; Bookman et al., 2002), ADHD (Lowe et al., 2004;
Bellgrove et al., 2005; Yang et al., 2008; Thomson et al., 2013), schizophrenia (Lai et
al., 2010), and drug addiction (Ho et al., 2008; Lai et al., 2010). Other studies have
found no correlation, however (Mill et al., 2003; Kirley et al., 2004; David et al., 2008).
The varying results may be due to genetic differences between populations.
1.3.4 The importance of DRD4 in photoreceptors and in vision
Soon after its discovery, DRD4 was shown to play a role in AC shutoff in
photoreceptors (Cohen et al., 1992; Nir et al., 2002; Patel et al., 2003; Klitten et al.,
2008; Ivanova et al., 2008). Retinal DRD4 mRNA expression is circadian (Figure 1.6,
1.7) (Humphries et al., 2002; Bai et al., 2008; Kim et al., 2010), as is its regulation of
cAMP in the retina (Figure 1.7) (Jackson et al., 2011). This finding led to the
investigation of DRD4 as a modulator of connexin 36 (Cx36) phosphorylation, which
allows gap junctional coupling between photoreceptors (Lampe and Lau, 2000; Lampe
and Lau, 2004; Ouyang et al., 2005; O'Brien et al., 2012). It was soon discovered that
coupling between photoreceptors was indeed regulated by DRD4, in combination with
the adenosine A2 receptor (A2AR) (Li et al., 2009; Li et al., 2013). It has yet to be
discovered if the downstream signaling of DRD4 affects other cAMP-linked signaling
pathways in the retina, but electrophysiological studies (Jackson et al., 2012) and gene
30

expression studies (Hwang et al., 2013) indicate that DRD4 plays an important role in
electroretinography (ERG) responses and circadian expression of clock genes.
1.3.5 The shutoff of DRD4
As each of the dopamine receptors was discovered, researchers began to
determine their specific mechanisms of action, and how the receptors are desensitized
for effective regulation of their signaling pathways. It was quickly shown that DRD1 and
DRD2 undergo agonist-induced internalization (Kim et al., 2001; Mason et al., 2002;

Figure 1.7 A. Drd4 mRNA expression in mice is under circadian control. B. The
circadian expression pattern is diminished after two days of constant dark,
indicating that it is light-dependent. C. When Drd4 is not present in mouse retinas,
circadian regulation of cAMP accumulation is severely disrupted. Re-used with
permission from John Wiley and Sons; original figure appeared in Jackson et. al.
2011, European Journal of Neuroscience.
31

Namkung and Sibley, 2004; Macey et al., 2005) and soon after published that DRD3
and DRD5 also undergo a less robust internalization of the receptor.
In 1999, Watts and colleagues reported that DRD4 did not internalize (Watts et al.,
1999), and in a 2000 review of DRD4 research Van Tol cited unpublished observations
confirming this (Oak et al., 2000). In 2010, Spooren and colleagues published a detailed
study of DRD4 desensitization, verifying that the intracellular portion of the receptor is
phosphorylated, but that the phosphorylation is not altered by DA stimulation (Spooren
et al., 2010). They also showed through co-immunoprecipitation (co-IP) studies that
DRD4 binds β-ARR2, but that this binding is also independent of DA. In this study, β-
ARR recruitment to the plasma membrane was not observed with DA stimulation, but a
previous report did observe the translocation of β-ARR2 (Cho et al., 2006). This finding
raises more questions about the signaling and shutoff of DRD4; if it is constitutively
phosphorylated and bound to β-ARR2, how does it interact with its G-protein to inhibit
AC? Further, it is able to inhibit AC in spite of phosphorylation and apparent arrestin
binding, so how does the receptor become desensitized? The authors of the study
hypothesize that DRD4 signaling may have longer effects than the other dopamine
receptors, since it does not appear to be down-regulated through the same
mechanisms.

32

Visual Cone Arrestin 4 Contributes to Visual Function and Cone Chapter 2:
Health
2

2.1 Introduction
Arrestins compose a family of four proteins that are essential for desensitization of
numerous G-protein coupled receptors (GPCRs). In mammals, there are two visual
arrestins, Arrestin 1 (ARR1, also called S-Antigen or 48 kDa protein) (Wacker et al.,
1977; Kuhn et al., 1984; Pfister et al., 1985) and Arrestin 4 (ARR4, also called cone
arrestin [CAR], X-arrestin, or ARR3 in the NCBI gene nomenclature) (Murakami et al.,
1993; Craft et al., 1994). Previous work clearly demonstrated the functional roles of
visual arrestins in mouse models in which these two genes are individually or
simultaneously ablated (Arr1
-/-
, Arr4
-/-
, or Arr1
-/-
Arr4
-/-
) (Xu et al., 1997; Chen et al., 1999;
Nikonov et al., 2008; Huang et al., 2010; Brown et al., 2010).
After opsins have been light-activated, their shutoff begins with multiple
phosphorylations by G-protein coupled receptor kinase 1 (Grk1) for rhodopsin (Wilden
et al., 1986; Baylor and Burns, 1998), and depending on the species, either Grk1 or
Grk7 for cone opsins (Hisatomi et al., 1998; Weiss et al., 1998; Weiss et al., 2001; Chen
et al., 2001; Zhu et al., 2003). These phosphorylated opsins are subsequently bound by
either ARR1 or ARR4, which sterically inhibits the opsins from further activation of the
downstream G-protein, alpha ( α) transducin. ARR1 is primarily responsible for the
signal shutoff of rhodopsin in rods; in contrast, in murine cones both ARR1 and ARR4
are co-expressed and desensitize short (S-) or middle (M-) wavelength opsins (Nikonov
et al., 2008). Compared to ARR4, ARR1 has a 50 fold higher concentration in cone
photoreceptors and can substitute for ARR4 in the shutoff of S-opsin or M-opsin
2
This chapter was accepted for publication in Investigative Ophthalmology and Visual Science
and is in press (Deming et al., 2015b).
33

(Nikonov et al., 2008). When measuring the single cone photoreceptor physiological
response after a bright light stimulus, experiments demonstrate that mice without
expression of both visual arrestins have limited and delayed recovery, underscoring the
critical role of the visual arrestins in normal phototransduction shutoff.
Recent evidence suggests that ARR4 plays a unique role in other visual functions
for which ARR1 cannot substitute and vice versa (Brown et al., 2012 53:ARVO E-
Abstract 760/A637). Zebrafish retina studies of the ARR4 ortholog, Arr3a, have
demonstrated that it plays a vital role in maintaining the normal optokinetic response of
zebrafish larvae across temporal frequencies (Renninger et al., 2011).

In contrast,
ARR1, but not ARR4, was shown to modulate the mouse rod photoreceptor presynaptic
exocytotic function of N-ethylmaleimide Sensitive Factor (NSF) and to contribute to
normal light adaptation (Huang et al., 2010; Brown et al., 2010).
In this study, we further investigated Arr4
-/-
mice (Nikonov et al., 2008) to assess the
contribution of ARR4 to overall cone visual function. In previous published work, Arr4
-/-

mice had enhanced photopic ERG amplitudes and abnormal flicker response compared
to controls (WT) (Brown et al., 2010). We performed optokinetic tracking (OKT) studies
to determine if these abnormal photopic ERG amplitudes corresponded to downstream
functional changes in either visual acuity or contrast sensitivity. We also examined the
cone photoreceptor morphology of Arr4
-/-
mice as they age, to determine if the observed
abnormal retinal physiology and behavioral defects in younger mice contributed to
greater visual deficits or increased cone degeneration over time.

34

2.2 Materials and Methods
2.2.1 Mice
Arr4
-/-
mice were produced on a mixed C57Bl/6J-129SVJ strain (WT) background
(Nikonov et al., 2008) and were reared in a 12 hour:12 hour light:dark cycle and tested
at 2, 4, 7, and 9 months of age. The mice tested for OKT were 3-4 months old. The 2
and 4 month old mice were phenotypically identical, as were the 7 and 9 month old
mice, so they were separated into two groups: “younger” mice of 2 and 4 months old,
and “older” mice of 7+ months old. Mice of either sex were used for experimental
procedures. All animals were treated according to the guidelines established by the
Institute for Laboratory Animal Research (Guide for the Care and Use of Laboratory
Animals), conformed to the Association for Research in Vision and Ophthalmology
statement for the Use of Animals in Ophthalmic and Vision Research, and were
approved by the appropriate animal committees of the University of Southern California
and the Atlanta VA Medical Center.
Table 2.1 Mouse strains used in Chapter 2
Name Source Publication
WT (C57Bl/6J-129SVJ mix) C. Craft Nikonov 2008
Arr4
-/-
C. Craft Nikonov 2008

2.2.2 Optokinetic Tracking (OKT)
For OKT, mice were placed on a platform in the center of a virtual-reality chamber,
which is composed of four computer monitors (OptoMotry, Cerebral Mechanics,
Lethbridge, AB, Canada), as previously described (Aung et al., 2013). A vertical sine
wave grating rotated across the monitors at a speed of 12 deg/s. Mice were monitored
for reflexive head movements in the direction of the rotating gratings using a video
35

camera positioned above the animal. For visual acuity assessment, the grating started
at a 0.042 cycles/degree (c/d) spatial frequency with 100% contrast and increased in a
staircase paradigm until the maximum spatial frequency threshold was reached.
Contrast sensitivity curves were measured across five spatial frequencies (0.031, 0.064,
0.092, 0.103, and 0.192 c/d) (Prusky et al., 2004).
2.2.3 Electroretinography (ERG) analysis
Following 12 hours of dark adaptation, ERG studies were performed as previously
described in detail (Brown et al., 2010). Briefly, flash stimuli of 10 µs duration, from 0.2-
20 Hz were delivered through one arm of a bifurcated glass fiber optic to within 3 cm of
the corneal surface. The ultraviolet filter plate of the Xenon flash unit was removed to
permit transmittance of shorter wavelengths (<400 nm) and the delivery arm of the fiber
optic was affixed to the flash unit directly under, and 7 cm from, the Xenon flash bulb.
Continuous white light, 8 foot candles (fc) (200 cd-s/m
2
) was delivered through the other
arm of the fiber optic for 1 minute before the first flash response was recorded.
Additional responses were recorded every two minutes up to 15 minutes of light
adaptation. Following the light adaptation recordings, the white background light was
delivered for 1 additional minute before the single-flash recordings presented in Figure
2. Maximum flash intensity at the surface of the cornea was 2.01 log (cd-s/m
2
) and
calibrated with a photometer (UDT model S350 Laboratory Photometer with a Model
211 Illuminance Sensor Head, San Diego, CA). For some studies, reduction in intensity
(from 2.01 to -1.59 log cd-s/m
2
) was achieved by the use of both neutral density Wratten
filters and the 16X-1X flash intensity settings on the Grass visual stimulator. The optimal
flash intensity that resulted in the greatest difference between young WT and young
36

Arr4
-/-
was 2.01 log (cd-s/m
2
), so this was the intensity used for the studies presented
here.
2.2.4 Immunoblot (IB) analysis
Each eye was enucleated and the retina removed. After dissection, each retina was
flash frozen on dry ice and maintained at -80°C until use. Each frozen retina was
homogenized; 60 µg of protein per retina were resolved on replicate 10% SDS-PAGE,
transferred to nitrocellulose membranes (Li-Cor), incubated sequentially with antibodies
for anti β-actin (1:4,000) and either anti-S-opsin (1:5,000), or anti-M-opsin (1:5,000)
(Zhu et al., 2003). Appropriate secondary antibodies conjugated to a fluorophore (680
nm or 800 nm) allowed detection using the Li-Cor Odyssey infrared detection system.
Li-Cor Odyssey v. 3.1 was used to quantify the intensity of each band. The linear
relationship between the amount of immunoreactive protein on the membrane and the
fluorescence intensity detected by the Li-Cor Odyssey
TM
(Shutz-Geschwender et al.,
2004; Eaton et al., 2013) allows direct and quantitative comparisons between the ratio
of two proteins across multiple blots. Relative amounts of the opsins were calculated by
dividing the intensity of the M- or S-opsin band by the intensity of the β-actin band. The
average of the WT younger samples was set as 100% (Nikonov et al., 2008).
2.2.5 Immunohistochemistry (IHC)
Materials and methods were previously published for IHC (Zhu et al., 2003). Briefly,
the retinal sections were obtained from the eyes fixed in 4% paraformaldehyde in
phosphate buffered saline (PBS) for 1 hour on ice. Each lens was removed prior to
embedding in Optimal Cutting Temperature (OCT) medium (Sakura Finetechnical,
Torrance, CA) and snap frozen in liquid nitrogen. Frozen retinal sections were cut in a
37

cryostat at 10 µm thickness along the vertical meridian through the optic nerve and were
placed on SuperFrost Plus (VWR) glass slides (3 sections per slide).
Sections were rehydrated in PBS and blocked with blocking buffer (10%
ChemiBlocker (Millipore), 0.5% Triton X-100) for 30 minutes at room temperature, then
incubated at 4°C overnight with affinity purified rabbit polyclonal antibodies for anti-
mouse S-opsin or anti- mouse M-opsin peptide (dilution 1:1,000) (Zhu et al., 2003).
Sections were washed 3 times in PBS and incubated for an hour at room temperature in
Alexa Fluor 488-conjugated anti-rabbit secondary antibody (1:500) (Invitrogen,
Carlsbad, CA), then mounted with mounting medium with DAPI (Vectashield, Vector
Laboratories) and covered with a glass coverslip.
The same IHC procedures described above were used for whole-mount
immunological analysis, except for the following antibody incubation times: primary and
secondary antibodies were each sequentially incubated for 36 hours.
Slides were imaged using either a Leica DMR fluorescent microscope (Leica
Microsystems, Buffalo Grove, IL) using a 20x dry lens or a Carl Zeiss LSM-510 confocal
microscope with a 40x oil immersion lens equipped with a digital camera (SPOT
SP401–115, software version 3.5; Diagnostic Instruments, Inc., Sterling Heights, MI).
Intensity measurements: Images were recorded using the digital camera with
identical exposure times for all retinas studied. The average pixel intensity of the
photoreceptor layer was quantified in ImageJ (Reish et al., 2013) at equally spaced
regions across each retina section, from the ON to 2.1 mm away in either direction
(inferior and superior).

38

Table 2.2 Primary antibodies used in Chapter 2
Name Source Speci-
ficity
Species Appli-
cation
Dilution Publication
M-opsin
residues 3–16,
QRLTGEQTLD
HYED
C. Craft mouse
M-opsin
rabbit IHC
IB
1:1000
1:5000
Zhu 2003
S-opsin
residues 1–11,
MSGEDDFYLF
Q
C. Craft mouse
S-opsin
rabbit IHC
IB
1:1000
1:5000
Zhu 2003
β-actin (clone
AC-15)
Sigma
Aldrich
(#A1978)
mouse
β-actin
mouse IB 1:4000
Abbreviations: IHC: immunohistochemistry; IB: immunoblot; co-IP: co-immunoprecipitation

2.2.6 Cone cell counts
Confocal micrographs of the whole-mounted retinas (n=3 animals per group) were
taken at the focal level of the outer segments of M-opsin and S-opsin immunologically
stained cones, covering a 300 x 300 µm
2
area at both the superior and inferior regions
(1mm away from optic disc) of the retina. Each cone outer segment was then marked by
a visual dot, using Photoshop (Adobe), to facilitate accurate counting.
2.2.7 Outer nuclear layer (ONL) thickness measurement
DAPI staining was performed as described for IHC above. The DAPI fluorescence
for each entire retina section was captured using a Leica DMR fluorescent microscope
(20x objective) with a SPOT imaging program. Images were stitched together in
Photoshop (Adobe) to recreate the whole vertical section of the retina. Starting 0.3 mm
from the optic nerve, the number of layers of nuclei were counted (Brown et al., 2010)
every 0.6 mm on both the inferior and superior sides of the retina.
39

2.2.8 Statistical analysis
Statistical analysis was performed using Two-way ANOVA and Student’s t-tests
with GraphPad Prism 6 or Three-way ANOVA with SPSS statistical software. Huynh-
Feldt correction factors were used when the data failed the Machly’s test of sphericity.
Post-hoc tests were performed using Student’s t-tests with rough false discovery rate
correction factor (Benjamini and Hochberg, 1995) to decrease the chance of Type II
Errors.

40

2.3 Results
2.3.1 OKT studies reveal decreased visual acuity and contrast sensitivity in Arr4
-/-
mice
Based on our original hypothesis that visual acuity and/or contrast sensitivity would
be compromised in Arr4
-/-
mice, OKT was used to measure visual acuity and contrast
sensitivity in Arr4
-/-
mice compared to WT. Spatial frequency and contrast sensitivity
thresholds are commonly used as measurements of functional visual performance in
rodents. Because data from a previous study noted abnormally high photopic ERG
amplitudes in 1 month old Arr4
-/-
mice (Brown et al., 2010), OKT studies were performed
to determine the effect of the absence of Arr4 expression on visual function. Four month
old Arr4
-/-
mice have significantly reduced visual acuity thresholds compared to WT mice
(Figure 2.1A; Student’s t-test p<0.001). The average contrast sensitivity threshold for

Figure 2.1A: Visual acuity thresholds of 4 month old WT mice compared to Arr4
-/-
.
Arr4
-/-
mice have a significantly lower visual acuity threshold than WT (p<0.001).
1B: Contrast sensitivity thresholds (arbitrary units = a.u.) of 4 month old WT and
Arr4
-/-
mice at multiple frequencies (cycles/degree). At all mid-range frequencies,
Arr4
-/-
mice display lower contrast sensitivity thresholds compared to WT (*p<0.05
or **p<0.01). WT, wild type; Arr4, arrestin 4.
41

each genotype at multiple spatial frequencies (cycles/degree) is shown in Figure 2.1B.
Arr4
-/-
mice have significantly lower contrast sensitivity thresholds than WT mice at all
mid-range spatial frequencies (Two-way repeated ANOVA F(5,89) = 4.55, p<0.001).
2.3.2 Physiological response of young Arr4
-/-
mice is abnormal compared to WT
The functional vision deficits displayed in the OKT of 4 month old Arr4
-/-
mice
indicate a problem with visual signaling and/or downstream processing, although the
photopic ERG recording of 1 month old Arr4
-/-
mice was abnormally high. We
hypothesized that the abnormal signaling would lead to visual deficits in older Arr4
-/-

mice. In order to determine if age-related changes were occurring in the retina, photopic
ERGs were performed in age-matched Arr4
-/-
and WT mice at 2 months and 7+ months.
Representative waveforms to single flash stimuli from light-adapted mice are shown
in Figure 2.2A. In single flash tracings, the a-wave and b-wave amplitudes are shown
(Figure 2.2B and C). The younger Arr4
-/-
mice display larger photopic ERG amplitude
responses compared to WT controls, which is consistent with published results (Brown
et al., 2010).
Analyses of the a-wave amplitudes showed significant differences in amplitudes
between genotypes that were dependent on age (Two-way ANOVA interaction effect,
F(1,26)=5.03, p<0.05; Figure 2.2). We found a significant decrease in a-wave amplitude
across age in Arr4
-/-
mice (p<0.001),

but not WT mice. The a-wave amplitude in older
Arr4
-/-
mice was even significantly lower than older WT mice (p<0.05).
ERG b-wave amplitudes were significantly decreased in old versus young mice
(Two-way ANOVA main effect, F(1,25)=27.37, p<0.001), with WT mice decreasing by
28% (p<0.01) and Arr4
-/-
mice by 45% (p<0.001). In addition, younger Arr4
-/-
mice had
42

significantly higher b-wave amplitude than younger WT (p<0.05). Our results show that
both a-wave and b-wave amplitudes decrease with age in the Arr4
-/-
mouse.
Representative flicker waveforms are shown in Figure 2.3A. Younger Arr4
-/-
had
significantly higher amplitudes compared to older Arr4
-/-
at all frequencies (Three-way
ANOVA, frequency X age interaction F(2,14)=4.53, p<0.05). Thus, flicker ERG
responses decrease with age for Arr4
-/-
while WT responses remain unchanged.

Figure 2.2A: Representative photopic ERG tracings for 2 mo. Arr4
-/-
and WT mice.
IT
a
is the implicit time, or the time from the stimulus to the peak of the wave, for the
a-wave. IT
b
is the implicit time for the b-wave. 2B: A-wave amplitudes of younger (2
mo.) and older (7+ mo.) WT and Arr4
-/-
mice. Older Arr4
-/-
mice have significantly
lower amplitudes than younger Arr4
-/-
mice (***p<0.001) and older WT mice
(*p<0.05). 2C: B-wave amplitudes of younger and older WT and Arr4
-/-
mice.
Younger Arr4
-/-
mice have significantly higher b-wave amplitudes (*p<0.05), while
older Arr4
-/-
b-wave amplitudes are significantly decreased compared to younger
Arr4
-/-
amplitudes (***p<0.001). WT amplitudes also decrease with age (**p<0.01).

43

2.3.3 Immunoblot analysis reveals a decrease in M- and S-opsin expression in older
Arr4
-/-
mouse retinas
The decrease in photopic ERG signal in the older Arr4
-/-
mice led us to the
hypothesis that M-opsin or S-opsin expression level in the mice may correlate with the
observed differences in photopic ERG amplitudes. Immunoblot analysis of total retinal
protein was performed in order to determine if M-opsin or S-opsin expression correlates
with the observed differences in photopic ERG amplitudes (Figure 2.4). Immunoreactive
M-opsin protein expression decreases with age in Arr4
-/-
mice; however, no decrease
was observed in WT (Two-way ANOVA F(1,8)=15.60, p<0.01) (Figure 2.4B).
For immunoreactive S-opsin, no significant interaction is observed between age and
genotype, but each variable contributes independently to the variation between groups
(Two-way ANOVA main effects of age F(1,8)=37.19, p<0.001, and genotype
F(1,8)=43.29, p<0.001; Figure 2.4D). There is a significant difference between younger
Arr4
-/-
and older Arr4
-/-
mice (p<0.001), and older WT express more S-opsin than older
Arr4
-/-
mice (p<0.001).

Figure 2.3A: Representative photopic response ERG tracings for younger Arr4
-/-
and
WT mice with 10 Hz Flicker stimulus. 3B: Average b-wave amplitudes in response to
multiple flicker frequencies. At all frequencies, Arr4
-/-
mice at 2 months of age are
significantly higher than Arr4
-/-
mice at 7+ months. (**p<0.01 or *p<0.05). There are
no other significant comparisons.
44

2.3.4 M-opsin expression is increased in the inferior retina of young Arr4
-/-
mice, but in
older mice is similar to WT
The immunoblot studies determined differences in M-opsin and S-opsin expression
in the older Arr4
-/-
mice, but this analysis could not determine whether these changes in
protein expression occurred on the inferior region, superior region, or both regions of
the retina. In order to test our hypothesis that the change in cellular expression pattern
of M- and S-opsin is different in the superior and inferior regions of the retina, IHC
studies were done on retinal sections to compare immunological staining intensity in the
inferior vs. superior retina, as well as at defined distances from the optic nerve. M-opsin
intensities varied by location, age and genotype (Three-way ANOVA interaction,
F(6,76)=2.6, p<0.05). In general, M-opsin intensities were significantly lower in the

Figure 2.4A: Immunoblot analysis of total retinal protein homogenates from SDS-
PAGE for three WT and three Arr4
-/-
younger animals (2 mo.) and older animals (7+
mo.). Blot was probed for M-opsin and β-actin expression. 4B: Quantification of the
intensity of each M-opsin band (see methods). Arr4
-/-
younger is significantly higher
than Arr4
-/-
older (**p<0.01), and WT older is significantly higher than Arr4
-/-
older
(*p<0.05).4C: Immunoblot analysis of 60 µg of total retinal protein probed for S-opsin
and β-actin. 4D: Quantification of the intensity of each band. Arr4
-/-
older is
significantly lower than Arr4
-/-
younger (***p<0.001) and WT older (***p<0.001).
45

inferior retina compared to the superior retina across all genotypes and age (p<0.001).
In the inferior retina, younger Arr4
-/-
mice have significantly higher M-opsin intensity than
younger WT (p<0.05 or p<0.01) and older Arr4
-/-
mice (p<0.01 or p<0.001). In contrast,
M-opsin intensity in the WT mice does not change significantly with age.

Figure 2.5A: Representative images of IHC analysis of retinal sections for M-opsin
(green). DAPI nuclear stain is in blue. 5B: Quantification of fluorescent intensity of
M-opsin IHC staining. M-opsin intensities were higher in the superior retina for all
ages and genotypes (p<0.001). Additionally, younger Arr4
-/-
mice have significantly
higher M-opsin labeled fluorescent intensity than younger WT mice in the inferior
retina (*p<0.05 or **p<0.01). Younger Arr4
-/-
mice have significantly higher M-opsin
label fluorescent intensity than older Arr4
-/-
mice throughout the inferior retina as well
(**p<0.01 or ***p<0.001).
46

The overall S-opsin intensities showed an opposite pattern to the M-opsin
intensities, with significantly lower intensities in the superior retina (Three-way ANOVA,
main effect of location F(4, 59)=36.3, p<0.001; Figure 2.6). No differences in S-opsin
intensities for age or genotype were found.

Figure 2.6A: Representative images of IHC analysis of retinal sections labeled for S-
opsin (green). DAPI nuclear stain is blue. 6B: Quantification of IHC fluorescent
intensity of S-opsin reveals significantly higher S-opsin intensities in the inferior
retina across all ages and genotypes (p<0.001).
47

2.3.5 M- and S-opsin cone numbers decrease with age in Arr4
-/-
mouse retinas
Based on the immunoblot and IHC analysis results indicating that the older Arr4
-/-

mice express less M- and S- opsin, we hypothesized that the older Arr4
-/-
had
experienced cone dystrophy, resulting in a decrease in cone number. To clarify if the
differences in M- and S-opsin protein expression were due to differential expression in
each cone or a difference in total cone number, M- and S-opsin labeled cone
photoreceptor numbers were quantified directly. In mice, M- and S- cones have different
expression patterns in the inferior vs. superior retina, so these areas were counted
separately (Szel et al., 1992).
The number of immunoreactive M-opsin cones was significantly different in the
superior and inferior regions of the retina with interactions between age and genotype
(Three-way ANOVA F(1,8)=15.6, p<0.01; Figure 2.7A). There are significantly fewer M-
opsin cones in the inferior retina of older Arr4
-/-
compared to both younger Arr4
-/-

(p<0.001) and older WT (p<0.001). The pattern is similar in the superior retina, with
fewer M-opsin cones in older Arr4
-/-
compared to both younger Arr4
-/-
(p<0.01) and older
WT (p<0.01). The number of M-opsin cones in the inferior retina also decreases in WT
mice as they age (p<0.05).
S-opsin cone numbers were significantly lower in the superior retina compared to
the inferior retina, which depended on genotype or age (Three way ANOVA location X
genotype F(1,8)=9.8, p<0.01; location X age F(1,8)=17.4, p<0.01; genotype X age
F(1,8)=9.2, p<0.05). In the inferior retina, S-opsin cone numbers decrease in both the
WT (p<0.05) and Arr4
-/-
(p<0.01) mice across age. Additionally, older Arr4
-/-
retinas have
48

fewer S-opsin cones than older WT retinas in both the inferior (p<0.01) and superior
(p<0.01) retina.
We hypothesized that the photoreceptor loss was specific to cones and would not
affect the number of rods in the older Arr4
-/-
mice. To determine if the deterioration of

Figure 2.7A: Number of M-opsin IHC-stained cones in a 300x300 µm
2
area for young
and old WT and Arr4
-/-
mice. Inferior retina regions had fewer M-cones than superior
regions. In both the inferior and superior retina, older Arr4
-/-
mice have fewer M-opsin
cones than younger Arr4
-/-
(inferior ***p<0.001, superior ***p<0.01) and older WT
mice (Inferior ****p<0.001, superior ***p<0.01). In the superior retina, the number of
M-cones is decreased in older WT compared to WT younger (*p<0.05). 7B: Total
number of S-opsin IHC-stained cones in the same area as 7A. Superior regions of
the retinas had far fewer S-cones than the inferior regions (note y-axis split). In the
inferior retina, younger mice have significantly more S-opsin cones than older mice
for both WT (*p<0.05) and Arr4
-/-
(p<0.01). The older Arr4
-/-
also have significantly
fewer S-opsin cones than older WT in both the inferior (p<0.01) and superior
(p<0.01) regions of the retina. 7C: Outer nuclear layer (ONL) thickness, in number of
layers of nuclei, throughout the inferior and superior retina. There are no significant
differences between groups.
49

cones in older Arr4
-/-
corresponded with an overall degeneration of photoreceptors,
including rods, the thickness of the outer nuclear layer of the retina was measured for all
groups. We found no significant differences between genotype, age or location,
indicating that the photoreceptor outer nuclear layers remain the same in Arr4
-/-
and WT
mice across age. The results are consistent with a previous study of young Arr4
-/-
and
WT mice (Brown et al., 2010).

50

2.4 Discussion
2.4.1 WT young vs. Arr4
-/-
young
Visual ARR1 can substitute for ARR4 in both S-opsin and M-opsin mouse
phototransduction signal shutoff at the single cell level (Nikonov et al., 2008); however,
our results clearly demonstrate that the lack of ARR4 contributes to visual phenotype
abnormalities. For example, young Arr4
-/-
mice display enhanced photopic ERG b-wave
amplitudes (Figure 2.2C). We propose that this increase is due, at least in part, to a
surprising elevated M-opsin expression in the inferior retina (Figure 2.5). Single-cell
recordings of cone photoreceptors have confirmed that the expression level of the cone
opsins is positively correlated with cone functional signaling (Daniele et al., 2005;
Insinna et al., 2012). In addition, previous work in rodents has indicated that scotopic
ERG amplitudes are positively correlated with rhodopsin expression levels (Goto et al.,
1995; Gorbatyuk et al., 2007),

so it is feasible to predict that this elevated M-opsin
expression would also be correlated with higher photopic ERG amplitudes.
Although patients with enhanced S-cone ERG without widespread retinal
degeneration have been reported (Sieving, 1994; Kinori et al., 2011), supernormal cone
ERG amplitudes are not commonly observed. When rod phototransduction signaling is
compromised (Toda et al., 1999; Jaissle et al., 2001; Krebs et al., 2009), enhanced
cone ERG amplitudes are observed prior to degeneration, and a few molecules induce
an increase in photopic ERG amplitudes (Jurklies et al., 1996; Kapousta-Bruneau,
2000; Popova and Kupenova, 2011; Kim and Jung, 2012; Popova, 2014). Enhanced
photopic ERG amplitudes have also been observed in rats with streptozotocin-induced
diabetes mellitus (Olivier et al., 1990). Based on these other studies, Arr4
-/-
visual
51

deficits may provide additional links to understanding the cellular and molecular
mechanisms behind our observations.
In addition, Arr4
-/-
mice performed worse than WT mice in OKT behavioral
measurements of visual acuity and contrast sensitivity (Figure 2.1). A recent study in
zebrafish larvae observed that knockdown of expression of Arr3a, the ARR4 ortholog,
causes deficits in optokinetic responses to moving images across many temporal
resolutions (Renninger et al., 2011). These results indicate that cone arrestin is crucial
for contrast sensitivity, which in zebrafish is mediated by L- and M-opsin cones (Krauss
and Neumeyer, 2003; Orger and Baier, 2005), and this is consistent with our
observation that lack of ARR4 expression in mice leads to a decrease in visual acuity
and contrast sensitivity.
There are other rodent models that display defects in optokinetic responses, but
usually these functional deficits are accompanied by widespread photoreceptor
degeneration, increases in lens opacity, and/or decreases in ERG amplitudes (Lodha et
al., 2010; Wright et al., 2014; Lee et al., 2014; Aung et al., 2014). Notable exceptions to
this are the retinal amacrine cell-specific tyrosine hydroxylase targeted knockout mouse
(rTHKO) and knockout models of two GPCR dopamine receptors, Drd1 and Drd4,
respectively. Drd4
-/-
mice have a decrease in contrast sensitivity compared to WT, while
Drd1
-/-
mice have decreased spatial frequency thresholds compared to WT, and rTHKO
display decreases in both contrast sensitivity and visual acuity (Jackson et al., 2012). It
is clear from these data that dopamine and its receptors play an important role in the
optokinetic responses, although the molecular mechanism of this process is still under
investigation. Recent results indicate that ARR4 plays a role in the desensitization of
52

DRD4, which we hypothesize contributes to the decreased contrast sensitivity observed
in both Arr4
-/-
and Drd4
-/-
mice (Deming et al., 2013 54:ARVO E-Abstract 2452) (see
Chapter 4).
While the cellular and molecular mechanisms are still under exploration, the current
study demonstrates that expression of normal levels of ARR4 has other modulatory
roles in maintaining viable, metabolically healthy cone photoreceptors aside from cone
pigment shutoff. Because of the earlier appearance of these visual phenotypes by 2
months, we propose that these roles include developmental triggers and maintenance
of daily regulation of cone gene expression. This hypothesis is consistent with our
results, which indicate that each M-opsin expressing cone on the inferior side of the
young Arr4
-/-
retina produces more M-opsin protein than WT (Figure 2.5), while overall
M-opsin cone number is not significantly altered in younger Arr4
-/-
compared to WT
(Figure 2.7). The immunoblot analysis of M-opsin content did not show an increase in
M-opsin in the young Arr4
-/-
retina (Figure 2.4A), but we believe that because the
increase only occurred in the inferior region of the retina, the difference was not large
enough to detect using the total retina homogenate on the immunoblot.
2.4.2 WT old vs. Arr4
-/-
old
We also observe clear phenotypes in older Arr4
-/-
mice compared to WT. The total
immunoreactive retinal expression of M-opsin and S- opsin is decreased in older Arr4
-/-

mice, while WT remains unchanged (Figure 2.5 and 2.6). These data are consistent
with the M-opsin and S-opsin cone counts, which reveal an age-dependent
degeneration of cone numbers in older Arr4
-/-
mice (Figure 2.7A and B). In age-matched
WT mice, a small but not significant loss of cones occurs. Follow-up studies were
53

performed to examine hallmarks for cellular apoptosis using TUNEL staining in mouse
retina sections at 2, 5, 7, and 9 months, but no significant differences were noted
between Arr4
-/-
and WT (data not shown). This suggests that the observed cone
dystrophy occurs as a gradual degeneration over time. Outer nuclear layer (ONL)
thickness remained consistent with WT for both younger and older Arr4
-/-
(Figure 2.7C),
indicating that there is no significant rod degeneration. Thus, the degeneration observed
in the cones does not affect rod viability and should have no effect on rod structure or
function.
Since we observe a significant defect in photopic physiological signaling in the
younger Arr4
-/-
mice, the cone dystrophy observed in the older mice may be the result of
a slow but cumulative decrease in cone phototransduction shutoff. Alternatively, ARR4
may have other modulatory cellular and developmental partners and interact with GPCR
pathways in vivo that are unique and linked to ARR4’s functions. Although ARR4 does
translocate into the cone outer segments after bright light exposure, a significant
amount of ARR4 remains distributed throughout the synapse and the cell body of cones
(Zhu et al., 2002). The reason for this is still unclear, but we hypothesize that ARR4 is
essential for performing other critical functions in the cone photoreceptor synapse.
ARR4 can participate in non-GPCR pathways (Song et al., 2007), and ARR4 has also
been shown to interact with a photoreceptor ciliary transmembrane protein, Als2cr4
(TMEM237), which when mutated causes a Joubert syndrome related disorder (Zuniga
and Craft, 2010; Huang et al., 2011). This idea is also supported by the ability of ARR4
to bind non-opsin GPCRs, unlike the other visual arrestin, ARR1 (Sutton et al., 2005),
so ARR4 is likely to have further functions in cones that have not yet been discovered.
54

2.4.3 Arr4
-/-
younger vs. Arr4
-/-
older
In our experiments, the greatest phenotypic variations were observed between Arr4
-
/-
younger and Arr4
-/-
older groups. Even though the difference is not statistically
significant, in photopic ERG recordings and IHC M-opsin intensity measurements, the
data values of younger Arr4
-/-
are greater than those observed in the younger WT. For
example, unlike the photopic ERG b-wave pattern, the a-wave for younger Arr4
-/-
is
consistently higher than that of WT (Figure 2.2B). However, the older Arr4
-/-
amplitudes
are slightly reduced but not significantly different from the older WT. Because of this,
there is a significant difference between younger Arr4
-/-
and older Arr4
-/
, but not between
younger WT and older WT. Because of this, the older Arr4
-/-
mice phenotypically appear
more similar to their WT counterparts and may be misinterpreted to have corrected the
observed deregulated signaling in younger mice. In fact, if photopic ERGs were only
measured in older mice, you would reasonably conclude the ARR4 null mice were
physiologically “normal.”
In order to investigate this further, we evaluated the overall superior/inferior
distribution patterns and their intensity levels of M-opsin and S-opsin expression, in
addition to measuring their respective cone cell numbers. In the WT mice, the
distribution of M-opsin and S-opsin cones in the inferior and superior retina was
consistent with published results (Szel et al., 1992). We observed that the older Arr4
-/-

mice have lost M-opsin and S-opsin cones across their entire retinas. In rat models of
cone dystrophy, cone number is directly proportional to photopic ERG amplitude
(Sugawara et al., 2000). Based on the observed total cone number loss in the older
Arr4
-/-
mice, we would predict that the corresponding photopic ERG amplitudes would
55

be lower than older WT mice, but instead we were surprised to observe that the older
Arr4
-/-
ERG amplitudes are not significantly different from WT. Despite widespread cone
loss, the magnitude of photopic ERG response is similar. These data suggest that each
cone photoreceptor is sending an abnormally higher signal postsynaptically for both
young and older Arr4
-/-
mice, but the effect is masked by the widespread cone dystrophy
in the older animals.
Overall, this investigation leads us to conclude that ARR4 is a key component of
normal photopic visual signaling. Mice without ARR4 display widespread cone
dystrophy by 7 months of age, indicating that ARR4 is essential for long term cone
survival and high acuity vision over an animal’s lifetime. In parallel studies, we are
exploring alternative GPCR signaling pathways in cones and potential postsynaptic
communication to ON and OFF bipolar and inner retina relays that should reveal how
ARR4 is involved in maintaining high acuity vision. Even though no genetic defect has
yet been identified for the human ARR3 X-chromosomal linked cone arrestin, these
studies will contribute to a closer examination of patients with deficits in visual acuity
and contrast sensitivity or abnormal photopic ERG or flicker responses and may help in
understanding the etiology of other cone dystrophies.

56

Characterization of Antibodies to Identify Cellular Expression of Chapter 3:
Dopamine Receptor D4
3

3.1 Introduction
Dopamine plays an important but complex role in regulating vertebrate vision. It is
synthesized in a subpopulation of amacrine cells and diffuses throughout the retinal
layers to activate five types of dopaminergic G-protein-coupled receptors (GPCRs)
(Missale et al., 1998). In mice and zebrafish, the dopamine receptors D1 (DRD1) and
D4 (DRD4) contribute to various aspects of vision. In genetically engineered Drd1 and
Drd4 null mice, electroretinography and optokinetic tracking were used to examine
physiological and behavioral responses (Nir et al., 2002; Jackson et al., 2012). Other
studies revealed the importance of these GPCRs in the regulation of retinal clock genes
(Hwang et al., 2013), phosphorylation levels in photoreceptors (Pozdeyev et al., 2008),
and the opening and closing of gap junctions between retinal neurons (Li et al., 2009;
Hu et al., 2010; Li et al., 2013).
Tools to study DRD1 and DRD4 have been developed including GPCR-specific
agonists and antagonists and even a transgenic mouse expressing green fluorescent
protein (GFP)-tagged DRD4 (Gong et al., 2003), which are helpful but limited in
elucidating the function of each receptor. In addition, in situ hybridization studies have
clearly localized Drd4 mRNA in photoreceptors, the inner nuclear layer, and a
subpopulation of ganglion cells (Cohen et al., 1992; Klitten et al., 2008; Li et al., 2013);
however, the cellular localization and amount of endogenous DRD4 protein are still
unclear because of a lack of specific antibodies for IHC localization.
3
Large portions of this work (Chapter 3) have been accepted for publication in Advances in
Experimental Medicine and Biology, Retinal Degenerative Diseases: Mechanisms and
Experimental Therapy (Deming et al., 2015a), in press.
57

Although many antibodies for DRD4 are commercially available, only limited
published data are available on these reagents and none are reliable. It is notoriously
difficult to produce good antibodies against GPCRs, which are 7-transmembrane pass
cell surface receptors. Since there are five unique, but closely related, dopamine
receptors, it is even more challenging to find an antibody that is specific. Previous
characterizations of DRD4 antibodies have given confusing results, including varying
molecular weights (MW) of the DRD4 protein on denaturing acrylamide gels followed by
immunoblot (Gomez et al., 2002; Bavithra et al., 2012), and some with no molecular
weight listed (Chu et al., 2004; Li et al., 2007; Strell et al., 2009; Gonzalez et al., 2012).
Doubt has been cast on whether any of these DRD4 antibodies should be trusted at all
(Van Craenenbroeck et al., 2005; Bodei et al., 2009).
A reliable, specific antibody recognizing DRD4 to study the dopamine system, both
in the retina and in the brain, is essential for our project and would benefit others.
Furthermore, it can be a waste of time and resources to test multiple commercially
available antibodies only to discover that they are not specific to DRD4. In this study, we
characterized six anti-DRD4 antibodies using immunoblot analysis and IHC, both with
human DRD4 overexpressed in transfected HEK cells and with mouse retinas from
C57Bl/6J and Drd4
-/-
.

58

3.2 Materials and Methods
3.2.1 Mice
All animals were treated and protocols were approved by USC IACUC according to
the guidelines established by the Institute for Laboratory Animal Research. Breeders for
C57BL/6J and Drd4
-/-
(strain B6.129P2-Drd4
tm1Dkg
/J) mice were obtained from Jackson
Laboratory (Bar Harbor, MN) (Table 3.2). They were bred and their offspring were
reared in 12 hr light/12 hr dark cycling light conditions until sacrifice. Mice were
sacrificed in the dark at circadian time (CT) 0, before the lights were turned on. Eyes
were enucleated and eyecups were processed for IHC or retinas were stored at -80
o
C
for immunoblot analysis.
3.2.2 HEK293 Cell culture and transfection
HEK 293T/17 (HEK 293T) and HEK 293 cells were purchased from ATCC
(Manassas, VA) (Product number CRL-11268 and CRL-1573). They were maintained at
37
o
C, 5% CO
2
and used for experiments below 15 passages. For transient transfection,
HEK 293T were transfected with FuGENE 6 transfection reagent (Promega) for 48 hrs
before they were harvested for immunoblot or fixed and stained for IHC. For permanent
transfection, HEK 293 cells were transfected with FuGENE 6, and 72 hours later
selected with 1 mg/mL G418 antibiotic (corresponding to the neomycin resistance gene
present on the DRD4 expression plasmids). Cells were passaged and maintained with 1
mg/mL G418 in EMEM supplemented with 10% FBS.
3.2.3 DRD4 expression plasmids
Mammalian expression plasmids were generously provided by Dr. Van
Craenenbroeck. Plasmids used were pcDNA3-HA-DRD4.4, and pFLAG-DRD4.4 (Oak
59

et al., 2001; Van Craenenbroeck et al., 2005) (Table 3.1) They each express a common
variant of human DRD4, along with an HA- or FLAG-tag for labeling and verification.
3.2.4 Anti-DRD4 antibodies
Six anti-DRD4 antibodies were tested. Five were from Santa Cruz Biotechnologies
(2B9, D-16, H-50, N-20, R-20) and one was from Antibody Verify (AAS63631C) (Table
3.1). They each were raised against slightly different regions of rat or human DRD4
protein.
Table 3.1 Anti-DRD4 antibodies used in Chapter 3
Name Source Speci-
ficity
Species Appli-
cation
Dilution DRD4
amino
acids
DRD4
Region*
2B9 Santa Cruz
(#136169)
human
DRD4
mouse IHC
IB
1:100
1:200
176-185 TM4, EC3
D-16 Santa Cruz
(#31481)
human
DRD4
goat IHC
IB
1:100
1:200
Extracellular
domain
EC1, EC2,
EC3, EC4
H-50 Santa Cruz
(#25649)
human
DRD4
rabbit IHC
IB
1:100
1:200
141-190
Intracellular
EC2, TM3,
IC2, TM4,
EC3
N-20 Santa Cruz
(#31480)
human
DRD4
goat IHC
IB
1:100
1:200
N-terminus EC1
R-20 Santa Cruz
(#1439)
rat
DRD4
goat IHC
IB
1:100
1:200
335-385 IC3, TM6,
EC4, TM7
anti-
DRD4
Antibody
Verify(AAS6
3631C)
human
DRD4
rabbit IHC 1:100 122-182 EC2, TM3,
IC2, TM4,
EC3
Abbreviations: TM: Transmembrane region; EC: Extracellular region; IC: Intracellular region;
IHC: immunohistochemistry; IB: immunoblot; co-IP: co-immunoprecipitation

3.2.5 Immunoblot (IB) analysis
Retinas or frozen cell pellets were homogenized in 50 mM Tris, pH 7.6 plus
cOmplete protease inhibitor cocktail (Roche). They were sonicated to break apart DNA
and denatured with SDS sample loading buffer, then subjected to 10% SDS-
polyacrylamide gel electrophoresis (PAGE) and transferred to PVDF membranes
(Millipore). Anti-DRD4 (1:100 dilution) or anti-HA (1:1,000 dilution, Cell Signaling) (Table
60

3.3 and Table 3.1) primary antibodies were used in conjunction with HRP-conjugated
secondary antibodies (1:10,000 dilution, Bio-Rad or Santa Cruz). Hi-Blot
Chemiluminescence kit (Denville) was used for detection with film.
3.2.6 Immunohistochemistry (IHC)
Mouse retinas: Mouse eyes were enucleated and immediately fixed using published
methods (Zhu et al., 2002). They were fixed in 4% paraformaldehyde (PFA) for 10 min
(N-20) or 60 min (other anti-DRD4 antibodies). Eyes were cut into 20 µm sections (N-
20) or 10 µm sections (other anti-DRD4 antibodies). Retina sections were blocked with
normal donkey serum in PBS (N-20) or Chemiblocker (Millipore) (other DRD4
antibodies), and antibodies were diluted in PBS.
HEK 293: Cells were seeded onto glass slides in multi-well plates and given 24 hrs
to adhere. Cells were transfected (see above) and 48 hrs later rinsed with PBS and
fixed in 4% PFA. After fixation, cells were either blocked with Blotto (3% milk, 1 mM
CaCl
2
in TBS), or permeabilized with Blotto plus 0.01% Triton-X. Antibodies were diluted
in Blotto.
Retinas and transfected cells were labeled with anti-DRD4 primary antibodies (SC
antibodies at 1:50 dilution, Antibody Verify at 1:100) and anti-HA or anti-FLAG (at 1:500
dilution) for transfected cells (Table 3.3). After rinsing with PBS, cells were labeled with
fluorescent secondary antibodies (anti-goat, anti-rabbit, or anti-mouse AlexaFluor 488
and anti-rabbit or anti-mouse AlexaFluor 568, each at 1:500), mounted with Vectashield
mounting medium with DAPI (Vector Labs), and visualized using confocal fluorescent
microscopy (Zeiss LSM 510).
61

3.3 Results
3.3.1 Immunohistochemistry: Transfected Cells
Three SC antibodies, D-16, N-20, and H-50, labeled the DRD4-transfected cells
brightly and did not label non-transfected cells (Figure 3.1). The anti-DRD4 signal was
identical to the anti-FLAG signal and the cells were labeled both with and without
permeabilization, indicating an extracellular binding site (Figure 3.1). The other three
antibodies, R-20, 2B9, and AAS63631C, had little overlap with FLAG signal, indicating
that the anti-DRD4 signal was not specific to the tagged protein. Interestingly, there was
no signal in permeabilized non-transfected cells for all antibodies.
3.3.2 Immunoblots
To test specificity of each anti-DRD4 antibody on immunoblot analysis, C57BL/6J
and Drd4
-/-
total retinal proteins (collected at CT0) were electrophoresed simultaneously
on 10% SDS-PAGE, along with HEK 293T (not transfected) and HEK239T transfected
with HA-DRD4.4. The MW of human DRD4.4 is calculated to be 43.1 kiloDaltons (kDa),
while mouse DRD4 is 41.5 kDa. Out of six antibodies, only N-20 recognized human
DRD4, evidenced by two bands in the lane with transfected HEK 293T that were not in
the lane with untransfected HEK 293T. The molecular weight is higher than calculated
from the amino acid sequence (45 and 48 kDa), but this is normal and indicative of the
glycosylation of the receptor as it is processed in the endoplasmic reticulum and moves
to the plasma membrane (Lanau et al., 1997; Van Craenenbroeck et al., 2005) (Figure
3.2). The bands were verified to be HA-DRD4 by their overlap with the bands identified
by the anti-HA antibody (Figure3.2). In contrast, there were no distinct bands in the
62

Figure 3.1 IHC analysis of HEK 293T with or without FLAG-DRD4 transfection. Top
three rows demonstrate non-permeabilized cells to show that DRD4 is present on
the plasma membrane. SC N-20, D-16, and H-50 show bright staining of DRD4 (anti-
mouse, rabbit, or goat AlexaFluor 488, green, row 1) that clearly overlaps with anti-
FLAG ( anti-mouse or anti-rabbit AlexaFluor 568 red, row 2) staining (see merge,
row 3). Rows 4 through 6 demonstrate permeabilized cells to increase brightness
and verify specificity within cells. SC N-20, D-16, and H-50 continue to show strong
overlap with FLAG signal (red, row 7), while 2B9, AAS-63631C, and R-20 do not
overlap specifically. No detectable signal was observed in the non-transfected cells
(permeabilized), except with AAS63631C or SC R-20.
63

control retina lane that were not in the Drd4
-/-
samples. There were multiple non-specific
bands for all of the antibodies (Figure 3.2).
3.3.3 Immunohistochemistry: mouse retinas
All antibodies were tested on frozen sections of control and Drd4
-/-
mouse retinas
collected at circadian time (CT) 0, when DRD4 mRNA expression is highest. For all but
the SC N-20 antibody, there was no detectable difference between the signal intensity
of control versus knockout samples (Figure 3.3). The N-20 antibody showed a
significant signal in the ganglion cell layer that was not present in the Drd4
-/-
mouse
retina (Figure 3.4). There is also a small increase in signal in the inner segments of the

Fig. 3.2. Immunoblot analysis of all SC antibodies; first two lanes are Drd4
-/-
and C57
retina lysates; lanes three and four are HEK 293T (non-transfected) and HEK 293T
transfected with HA-DRD4.4. Anti-HA antibody specifically recognizes HA-DRD4.4
(top right, red arrow), and N-20 antibody also recognizes HA-DRD4.4, demonstrated
by the missing band in the non-transfected HEK 293T lane (right middle, red arrow).
No other antibodies recognize a band in the transfected lane over non-transfected;
and no antibodies specifically recognize Drd4 in C57 retina lysate over Drd4
-/-
(all
blots).
64

photoreceptor layer in control compared to Drd4
-/-
. No differences are observed in the
outer segment layer (OS), outer nuclear layer (ONL), outer plexiform layer (OPL), inner
nuclear layer (INL), or inner plexiform layer (IPL).
3.3.4 Permanently transfected cell lines
Four different cell lines were developed with permanent transfection of each DRD4
expression plasmid into HEK293. Expression was highest in the HEK293 plus pFLAG-
DRD4.4 line (Figure 3.5A). The HA-tagged DRD4 plasmids also expressed DRD4, but
at lower levels, as seen with IHC and immunoblot analysis using anti-HA antibodies
(Figure 3.5A and B). The specificity of the HA and FLAG signal to the appropriate cell
lines indicates that there was no contamination between cell lines.

Figure 3.3 IHC of control versus Drd4
-/-
retina sections using five different anti-DRD4
antibodies (green). Mice were sacrificed at CT0 (before lights on). All antibodies
displayed similar immunofluorescence intensity between WT and Drd4
-/-
over
multiple experiments.
65

3.4 Discussion
The results of this study demonstrate that of the antibodies tested, there is one clear
candidate for both mouse and human DRD4. SC N-20 stood out from the rest based on
its specific labeling of the control retina compared to Drd4
-/-
, and its specific immunoblot
labeling of human DRD4 in transfected cells. The three antibodies D-16, H-50, and N-20
all show immunological labeling of DRD4-transfected cells using IHC.
It is unclear why these antibodies do not reveal an obvious retinal difference
between control mice and Drd4
-/-
. This may be because, as a previous study suggested,
anti-DRD4 antibodies may recognize other dopamine receptors as well (Bodei et al.,
2009). If this is the case, DRD4 expression in the retina is not high enough, even at its
mRNA observed peak at CT0, to show significant signal above the other four dopamine

Fig. 3.4 IHC of control versus Drd4
-/-
retina sections using N-20 anti-DRD4 antibody
(green). Mice were sacrificed at CT0 (before lights on). The control retina compared
to Drd4
-/-
displays specific labeling in the ganglion cell layer (GCL) (arrows) and in
the photoreceptor inner segment layer (IS) (arrowheads).
66

receptors. Based on non-specific bands in immunoblot analysis, the anti-DRD4
antibodies may also recognize unrelated proteins of various sizes, additionally clouding
the signal of DRD3.
The differences between the in vitro and in vivo studies may also be due to
sequence differences. Most of the antibodies were raised against peptides based on the
sequence of human DRD4, not mouse. Since the transfected cells were overexpressing
human DRD4, small differences in the peptide sequence may make the antibodies
specific for human, and not mouse, DRD4. The only exception is R-20, raised against a
rat DRD4 peptide, which is similar to the mouse sequence with 94% identity between
mouse DRD4 and rat DRD4, versus 73% for mouse and human. This may explain why
R-20 did not recognize the human DRD4 in transfected cells.
Overall, no antibody was able to give clear and consistent results in all applications,
but SC N-20 is a clear favorite for future studies of DRD4 in both mouse retinas and
transfected cell lines.

Fig. 3.5 A, IHC of permanently transfected HEK 293 with multiple DRD4 constructs,
after multiple passages. HEK+HA-DRD4.2 and HEK+HA-DRD4.7 display weak
staining with anti-HA antibody, and the expression level of DRD4 appears low.
HEK293+FLAG-DRD4.4 is brightly stained with anti-FLAG antibody, and it has a
high expression level of DRD4. B, Immunoblot verifies the presence of HA-DRD4.2
and HA-DRD4.7 in their respective cell lines, with no contamination between cell
lines.
67

Dopamine Receptor D4 Internalization Requires a Beta-Arrestin and a Chapter 4:
Visual Arrestin
4

4.1 Introduction
Dopamine is a hormone and neurotransmitter that has important functions in many
aspects of neuronal signaling, including vision. Dopamine acts as a ligand for five
different G protein-coupled receptors (GPCRs), named Dopamine Receptor (DR) D1
through D5 (O'Dowd, 1993; Gingrich and Caron, 1993; Seeman and Van Tol, 1994).
The dopamine receptors are classified into two groups that have opposite downstream
effects in the cells in which they are expressed. The D1-like subgroup consists of DRD1
and DRD5, which activate adenylate cyclase (AC), thereby increasing the concentration
of cyclic AMP (cAMP) in the cell. The D2-like subgroup, including DRD2, DRD3, and
DRD4, acts to inhibit AC, effectively decreasing cAMP concentration in the cell.
One of the dopamine receptors with high retinal expression in rodents and humans
is DRD4 (Cohen et al., 1992; Matsumoto et al., 1995; Suzuki et al., 1995; Bai et al.,
2008; Klitten et al., 2008; Kim et al., 2010). Retinal and pineal gland DRD4 mRNA
expression oscillates daily in a circadian pattern, with the highest expression during
subjective night and lowest expression during subjective day (Humphries et al., 2002;
Fukuhara and Tosini, 2008; Bai et al., 2008; Klitten et al., 2008; Kim et al., 2010). Its
physiological action in photoreceptors is also circadian, although it counter-intuitively
contributes to daytime phenotypes instead of nighttime. DRD4 contributes to the
decrease in cAMP during the day (Jackson et al., 2011) and, in combination with the
adenosine 2A receptor (A2AR), decreases in gap junctional coupling during the day (Li
et al., 2009; Li et al., 2013). It is hypothesized that the action of DRD4 in the
4
This chapter was accepted for publication in Cellular Signalling and is in press (Deming et al.,
2015c)
68

photoreceptors is inhibited in the nighttime by G
s
-coupled GPCRs, such as A2AR, but
that this inhibition is overruled in the daytime by the increase in retinal DA, in spite of the
lower DRD4 expression during the day. There is no reported evidence of circadian
regulation of DRD4 expression in other cell types (Kim et al., 2010), so we hypothesize
that there is a unique mechanism for DRD4 desensitization and sequestration in the
photoreceptors and pinealocytes.
Because the dopamine receptors are GPCRs, they undergo the classical GPCR
activation and desensitization mechanisms. In this study, we focused on the
desensitization mechanism, in which ligand binding leads to G-protein coupled receptor
kinase (GRK) phosphorylation of serine and threonine residues of the intracellular loops
or carboxy-terminal tail of the receptor, followed by arrestin binding. There are two β-
arrestins, β-ARR1 and β-ARR2 (Lohse et al., 1990; Attramadal et al., 1992). The
binding of an arrestin often, but not always, leads to recruitment of endocytic machinery
and internalization of the receptor for degradation or recycling (reviewed in (Lefkowitz
and Shenoy, 2005)). The desensitization of the D2-like receptors has been studied, and
DRD2 and DRD3 are internalized after dopamine stimulation, as long as a β-arrestin is
present (Kim et al., 2001; Namkung and Sibley, 2004). The most recent research on
DRD4 showed that it is able to bind β-arrestin, but it is not internalized (Spooren et al.,
2010).
The previous work on DRD4 desensitization has investigated the role of the β-
arrestins because of their ubiquitous expression and interaction with many different
GPCRs (Spooren et al., 2010; Cho et al., 2010). However, there are two other arrestin
proteins expressed in mammalian tissues. These are the visual arrestins, ARR1 (also
69

called S-antigen or 48kDa protein) (Wacker et al., 1977; Kuhn et al., 1984; Pfister et al.,
1985; Shinohara et al., 1987) and ARR4 (also called cone arrestin, X-arrestin, or ARR3
in the NCBI gene nomenclature) (Murakami et al., 1993; Craft et al., 1994). The visual
arrestins have a distinct expression pattern. In rodents, ARR1 is highly expressed in the
photoreceptor rods and cones, and in pinealocytes (Craft et al., 1990). ARR4 is not
present in rods and is highly expressed in cone photoreceptors and pinealocytes. Two
studies have verified the presence of the β-arrestins in the retinal photoreceptors, as
well (Nicolas-Leveque et al., 1999; Cameron and Robinson, 2014). Although DRD4 is
highly expressed in photoreceptors (Cohen et al., 1992; Klitten et al., 2008; Li et al.,
2013), ARR1 and ARR4 were not included in the previous in vitro studies of DRD4
desensitization.
In this study, we sought to determine whether either of the two visual arrestins,
ARR1 or ARR4, may play a role in DRD4 desensitization in vitro.

70

4.2 Materials and Methods
4.2.1 HEK 293T cell culture and transfection
HEK 293T/17 (HEK 293T) cells were purchased from ATCC (Manassas, VA). They
were maintained in Dulbecco’s Modified Eagle’s Medium (DMEM) supplemented with
10% fetal bovine serum (FBS) at 37
o
C, 5% CO
2
and used for experiments below 15
passages. HEK 293T were transiently transfected with FuGENE 6 transfection reagent
(Promega) using a 3 µg DNA: 1 µL Fugene ratio for 48 hrs before they were harvested
for co-immunoprecipitation (co-IP) or fixed and stained for immunohistochemistry (IHC).
For experiments with two plasmids co-transfected, they were added in a 1:1
receptor:arrestin ratio. For experiments with three plasmids co-transfected, the amount
of each arrestin plasmid was decreased by half, so that the final ratio of
receptor:arrestin:arrestin was 2:1:1.
4.2.2 Mammalian expression plasmids
The DRD4-encoding plasmids used were pcDNA3-HA-DRD4.4, and pFLAG-
DRD4.4 (Oak et al., 2001; Van Craenenbroeck et al., 2005). They each express a
common variant of human DRD4, along with an human influenza hemagglutinin-derived
peptide (HA-) or FLAG-tag for labeling and verification. HA-tagged rat β-arrestin 1 and
β-arrestin 2 (pcDNA3-barr1-HA [#14693], pcDNA3-barr2-HA [#14692]), and FLAG-
tagged rat β-adrenergic receptor 2 (pcDNA3-FLAG-B2AR [#14697]) expression
plasmids (Tang et al., 1999; Luttrell et al., 1999) were obtained from AddGene
(Cambridge, MA). Human Arrestin 1 and Arrestin 4 cDNA plasmids were also used
(pCI-hSAG, p-noEGFP-hCAR [derived from pEGFP-hCAR]) (Li et al., 2002; Li et al.,
2003).
71

Table 4.1. Mammalian expression plasmids used in Chapter 4
Name Source Protein expressed Publication
pFLAG-DRD4.4 Van Craenenbroeck FLAG-DRD4.4 (human) Oak 2001
pcDNA3-HA-DRD4.2 Van Craenenbroeck HA-DRD4.2 (human) Oak 2001
pcDNA3-HA-DRD4.4 Van Craenenbroeck HA-DRD4.4 (human) Oak 2001
pcDNA3-HA-DRD4.7 Van Craenenbroeck HA-DRD4.7 (human) Oak 2001
pcDNA3-FLAG-B2AR Addgene (#14697) FLAG-B2AR (rat) Tang 1999
pcDNA3-barr1-HA Addgene (#14693) β-arr1-HA (rat) Luttrell 1999
pcDNA3-barr2-HA Addgene (#14692) β-arr2-HA (rat) Luttrell 1999
pcDNA3-barr1-FLAG Addgene (#14687) β-arr1-FLAG (rat) Luttrell 1999
pcDNA3-barr2-FLAG Addgene (#14685) β-arr2-FLAG (rat) Luttrell 1999
pCI-hSAG Craft/ A. Li ARR1 (human) Li 2002
pEGFP-hCAR Craft/ A. Li GFP-ARR4 (human) Li 2002
p-noEGFP-hCAR Craft/ J. Deming ARR4 (human) Derived from
pEGFP-hCAR

4.2.3 Mice
All animals were treated and protocols were approved by USC IACUC according to
the guidelines established by the Institute for Laboratory Animal Research. Breeders for
C57BL/6J and Drd4
-/-
(strain B6.129P2-Drd4
tm1Dkg
/J) (Rubinstein et al., 1997) mice were
obtained from Jackson Laboratory (Bar Harbor, ME). They were bred and their offspring
were reared in 12 hr light/12 hr dark cycling light conditions until sacrifice. Mice were
sacrificed in the dark at circadian time (CT) 0, before the lights were turned on. Other
mouse strains used for IHC and immunoblot analysis include Arr1
-/-
(Xu et al., 1997)
(generously provided by Jeannie Chen, University of Southern California), Arr4
-/-

(created and described in detail in (Nikonov et al., 2008) supplement), Arr-DKO
(Nikonov et al., 2008), β-arr1
-/-
(Conner et al., 1997) and β-arr2
-/-
(Bohn et al., 1999)
(eyes generously provided by Eugenia Gurevich, Vanderbilt University and Robert
Lefkowitz, Duke University). Eyes were enucleated and eyecups were processed for
IHC or retinas were dissected from eyes and stored at -80
o
C for immunoblot analysis.

72

Table 4.2 Mouse strains and genetically engineered knockout mice used in
Chapter 4
Name Source Publication
Arr4
-/-
C. Craft Nikonov 2008
Arr1
-/-
J. Chen Xu 1997
Arr1
-/-
Arr4
-/-
(Arr-DKO) C. Craft Nikonov 2008
C57Bl/6J Jackson Labs Poel 1958
Drd4
-/-
Jackson Labs Rubinstein 1997
β-arr1
-/-
(Arrb1
-/-
, Arr2
-/-
) R. Lefkowitz/ E. Gurevich Conner 1997
β-arr2
-/-
( βarr2-KO, Arr3
-/-
) R. Lefkowitz/ E. Gurevich Bohn 1999

4.2.4 Co-Immunoprecipitation (co-IP) and immunoblot analysis
For co-IP, transfected HEK 293T (see section 2.1) were incubated for 30 min in
DMEM with 10% FBS (regular medium), either with or without 10 µM DA. Cells were
placed on ice and harvested by aggressive pipetting with ice-cold phosphate buffered
saline solution (PBS), then centrifuged at 4
o
C and the supernatant was aspirated. Cell
pellets were frozen at -80
o
C until ready to use. Cell pellets were resuspended and lysed
in radioimmunoprecipitation assay (RIPA) buffer (Rondou et al., 2008) containing
cOmplete protease inhibitor cocktail (Promega) and the phosphatase inhibitor β-
glycerophosphate. The RIPA buffer with inhibitors was also used for all washes. 0.5 µL
rabbit polyclonal anti-hARR4 antibody Luminaire founder- human Cone Arrestin (LUMIf-
hCAR) (Zhang et al., 2003a) was added to each tube of cleared lysate and tubes were
rotated at 4
o
C for 3 hrs. 20 µL Protein A magnetic beads (Life Technologies) were then
added to each sample and incubated at 4
o
C overnight while rotating. The beads were
washed with ice-cold RIPA buffer three times before eluting bound proteins by
incubating at 37
o
C (n=2). The opposite co-IP, using anti-HA antibody, was also
performed (n=1).
73

Total cell lysate and co-IP eluate were subjected to 10% sodium dodecyl sulfate -
polyacrylamide gel electrophoresis (SDS-PAGE) and transferred to polyvinylidene
fluoride (PVDF) membranes (Millipore). Anti-HA (1:1000 dilution, Cell Signaling) and
LUMIf-hCAR (1:20,000 dilution) primary antibodies were used in conjunction with horse
radish peroxidase (HRP)-conjugated secondary antibodies (1:10,000 dilution, Bio-Rad).
Hi-Blot Chemiluminescence kit (Denville) was used for detection with film.
4.2.5 Immunohistochemistry
Mouse retinas: Mouse eyes were enucleated and immediately fixed using published
methods (Zhu et al., 2002). They were fixed in 4% paraformaldehyde (PFA) for 10 min
(for anti-DRD4 antibody N-20) or 60 min (for the anti-arrestin antibodies), washed with
PBS, and incubated in 30% sucrose overnight. Eyes were embedded in optimal cutting
temperature (OCT) medium, frozen, and cut into 20 µm sections (for N-20) or 10 µm
sections (for the anti-arrestin antibodies). Retina sections were blocked with normal
donkey serum in PBS (N-20) or Chemiblocker (Millipore) (anti-arrestin antibodies), and
antibodies were diluted in PBS. Retinas were incubated with an anti-DRD4 primary
antibody (N-20 anti-DRD4, Santa Cruz #31480 at 1:100 dilution), anti- β-ARR1 and β-
ARR2 (A2CT at 1:500 dilution) (Wei et al., 2003), anti-mARR4 (LUMIj-mCAR at 1:2,000
dilution) (Zhu et al., 2002), anti-SAG/ARR1 (monoclonal D9F2 at 1:20,000 dilution)
(Donoso et al., 1990), or anti- β-ARR2 (monoclonal H-9 Santa Cruz #13140 at 1:500
dilution). The corresponding secondary antibodies were anti-goat AlexaFluor 488, anti-
mouse AlexaFluor 488, anti-rabbit AlexaFluor 488, anti-rabbit AlexaFluor 568 or anti-
mouse AlexaFluor 568 (Life technologies). Slides were mounted with Vectashield hard-
74

set mounting medium with 4',6-diamidino-2-phenylindole (DAPI) (Vector Labs), and
visualized using confocal fluorescent microscopy (Zeiss LSM 510 or LSM 710).
HEK 293T: Cells were seeded onto glass slides in multi-well plates and given 24 hrs
to adhere. For N-20 anti-DRD4 labeling in HEK 293T, 48 hours after transfection
(Section 2.1), cells were rinsed with PBS and fixed using 4% PFA in PBS. Cells were
then blocked using Blotto (3% Milk, 1 mM CaCl
2
in tris-buffered saline solution (TBS)),
or permeabilized and blocked using Blotto-T (Blotto plus 0.01% Triton-X). FLAG-
DRD4.4 was labeled with M1 anti-FLAG antibody (Sigma) and N-20 anti-DRD4 antibody
(Santa Cruz Biotechnologies) together in Blotto. Secondary antibodies used were anti-
mouse AlexaFluor 568 and anti-goat AlexaFluor 488, respectively (Life Technologies).
For internalization experiments, 48 hours after transfection (Section 2.1), media was
removed and cells were incubated with anti-FLAG primary antibody (Sigma) at 4
o
C for 1
hr. DMEM with 10% FBS, with or without 10 µM DA (or 10 µM isoproterenol for B2AR),
was added to cells and they were incubated for the appropriate time (15 min to 60 min)
at 37
o
C, 5% CO
2
, then immediately fixed in 4% PFA in PBS. After fixation, cells were
permeabilized with Blotto-T.
Preliminary experiments indicated that 30 min of DA stimulation was optimal for
internalization of DRD4, so this time point was used for all internalization experiments.
For hARR4 translocation experiments, cells were also labeled with anti-hARR4 antibody
LUMIf-hCAR (Zhang et al., 2003b) and multiple DA stimulation times (15 min to 60 min)
were used to demonstrate and detect the cellular translocation of ARR4 over time. After
rinsing with PBS, cells were labeled with fluorescent secondary antibodies (anti-mouse
AlexaFluor 488 and, for hARR4 translocation, anti-rabbit AlexaFluor 568, each at 1:500
75

dilution), mounted with Vectashield hard-set mounting medium with DAPI (Vector
Labs), and visualized using confocal fluorescent microscopy (Zeiss LSM 510 or LSM
710).

Table 4.3 Primary antibodies used in Chapter 4
Name Source Specificity Species Application Dilution Publication
M1 anti-
FLAG
Sigma
Aldrich
(#F3040)
FLAG tag mouse IHC 1:500
anti-HA
(clone 6E2)
Cell
Signaling
(#2367)
HA-tag mouse IHC
IB
1:500
1:500

LUMIf-hCAR C. Craft human Arr4 rabbit IHC
IB
co-IP
1:1000
1:10,000
1:200
(Zhang et
al., 2003a)
LUMIj-mCAR C. Craft mouse Arr4 rabbit IHC 1:1000 (Zhu et al.,
2002)
D9F2 L. Donoso bovine S-
antigen/
Arr1
mouse IHC 1:10,000 (Donoso et
al., 1990)
A2CT R.
Lefkowitz
β-arr1 and
β-arr2
rabbit IHC
IB
1:500
1:10,000
(Wei et al.,
2003)
N-20 Santa
Cruz
(#31480)
human
DRD4
goat IHC
IB
1:100
1:200

β-arrestin 2 Santa
Cruz
(#13140)
human β-
ARR2
mouse IHC 1:500
Abbreviations: IHC: immunohistochemistry; IB: immunoblot; co-IP: co-immunoprecipitation

4.2.6 Other methods of measuring internalization
Flow cytometry: Transfected HEK 293T were transferred to 96-well V-bottom plates
and incubated 60 min with anti-FLAG antibody followed by 60 min with anti-mouse
AlexaFluor 488 secondary antibody, before incubating for 30 minutes at 37
o
C in DMEM
with 10% FBS, with or without 10 µM DA. The cells without DA served as an internal
negative control. After stimulation, cells were subjected to an acid wash (5 min, 50 mM
76

glycine, pH 2.0) to remove extracellular antibodies. Cells were rinsed in PBS and
analyzed for AlexaFluor 488 fluorescence using a FACSCalibur cytometer using BD
CellQuest software, and analysis was performed using Flowing Software version 2.5.0
(Perttu Terho, Turku, Finland) and GraphPad Prism (La Jolla, CA).
In-cell western: HEK 293T were grown and transfected in tissue-culture treated 96-
well flat bottom plates. Transfected cells were labeled with antibody, stimulated, and
subjected to acid wash as for flow cytometry, except that the secondary antibody was
anti-mouse IRDye 800 (Li-Cor). After acid wash, cells were rinsed in PBS and labeled
with Cell Tag 700 (Li-Cor) to label all cells. The plate was analyzed for fluorescence
intensity using Li-Cor Odyssey, and the ratio of 800 nm intensity to 700 nm intensity
was used for calculations.
4.2.7 Quantification and statistical analysis
Image analysis of co-IP results was performed using Image Studio Software (v.
3.1.4; LI-COR Biosciences, Lincoln, NE). Uniformly-sized rectangles were drawn around
each band, with background measurements of 3 pixels on all sides of each rectangle.
The mean intensity of the image within each rectangle was measured, and the mean
intensity of the background measurements was automatically subtracted from this
measurement by Image Studio. The background-subtracted intensities were compared
to the control, which was the sample that included DRD4, ARR4, but no DA stimulation
(set as 1). The ratio of each band intensity to the control (relative intensity) was plotted
on the charts in Figure 4.5.
Quantification of IHC internalization experiments was done by counting the total
number of cells in each image and the number of cells with clearly internalized
77

receptors. Counts from each of four experiments were averaged and the standard
deviation was calculated. The baseline was calculated by taking the average percent of
cells with internalized receptors of all of the unstimulated samples. Results were
reported relative to the baseline, and column statistics were performed using GraphPad
Prism (La Jolla, CA) to determine whether each stimulated sample was significantly
different from the baseline (p<0.05).
Quantification of colocalization experiments was performed using a similar method.
The total number of cells in each image expressing both DRD4 and ARR4 was counted,
and the number of cells displaying overlap of the two signals was also counted. The
percentage of cells displaying overlap was calculated for each image (n=3 to 9 for each
condition). The baseline was calculated by taking the average percent of cells with

Figure 4.1 Illustration describing the methods of measuring DRD4 internalization in
HEK 293T cells. A. Visual analysis using IHC. B. Flow cytometric analysis for
quantification. C. In-cell western assay for quantification.
78

DRD4 and ARR4 overlap in all of the unstimulated (no DA) samples. Results were
reported and column statistics were performed as for the internalization analysis above.
Flow cytometry: All experiments used the same cytometer settings and gating
parameters. The population of cells with strong AlexaFluor 488 fluorescence was
counted as “positive” for internalization. The final percentage of positive cells for each
group was calculated by taking the difference between the percentage of positive cells
in the stimulated group (with DA) and the unstimulated group (no DA). The average of 5
experiments was used.
In-cell western: Each experiment used a single 96-well plate, with each group
measured in quadruplicate. The ratio of 800 nm fluorescence intensity (internalized
receptors) to 700 nm (all cells) was calculated to determine the amount of internalized
receptors compared to the size of the population in the well. The average of the four
wells within each group was used, taking the difference between the 800nm:700nm
intensity ratio for the stimulated groups (with DA) and unstimulated groups (no DA) to
control for differences in transfection efficiency between groups.

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4.3 Results
4.3.1 β-arrestins and visual arrestins are co-expressed in mouse retinas
It is well-documented that visual arrestins, ARR1 and ARR4, are present at high
concentrations in mammalian photoreceptors (Pfister et al., 1985; Broekhuyse et al.,
1985; Whelan and McGinnis, 1988; Craft et al., 1990; Craft et al., 1994; Zhu et al.,
2002; Nikonov et al., 2008). The presence and expression level of β-arrestins in the
photoreceptors has been studied in three reports (Nicolas-Leveque et al., 1999;
Concepcion, 2007; Cameron and Robinson, 2014). First, we confirmed that the anti- β-
arrestin antibody we used, A2CT (Wei et al., 2003), recognizes both β-ARR1 and β-

Figure 4.2. Analysis of β-arrestins with immunoblot and immunohistochemistry (IHC)
analysis. A. Immunoblot analysis of mouse retinal homogenates resolved first on
SDS-PAGE from control (WT) and arrestin knockout mice to determine the specificity
of the rabbit polyclonal primary antibody A2CT, paired with an HRP-conjugated
secondary antibody. Arrows point out bands for β-ARR1 and β-ARR2, just above and
below 50 kDa. B. IHC of β-ARR1 and β-ARR2 using A2CT, followed with Alexa-Fluor
488 conjugated secondary antibody (green) in WT, β-arr1
-/-
, and β-arr2
-/-
mouse
retinas. Retina layers are labeled: OS, photoreceptor outer segments; IS,
photoreceptor inner segments; ONL, outer nuclear layer; OPL, outer plexiform layer.
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ARR2 using immunoblot analysis of mouse retinas (Figure 4.2A).
Immunohistochemical analysis of retinal sections was performed to verify that the β-
arrestin proteins can be detected in mouse photoreceptors. Consistent with published
results, our IHC demonstrates that β-arrestin 2 is present in the photoreceptor layer of
the mouse retina (Figure 4.2B), and also highly expressed in the outer plexiform layer,
inner nuclear layer, inner plexiform layer, and ganglion cell layer (data not shown).
Using retinas from β-arr1
-/-
and β-arr2
-/-
mice, we determined that β-arr2 is more highly
expressed than β-arr1 in the outer plexiform layer, outer nuclear layer, and inner and
outer segment layers. The retinas from the β-arr2
-/-
mouse have a greatly decreased
signal compared to WT (C57Bl/6J), while the retinas from the β-arr1
-/-
are similar to WT
in immunological staining intensity (Figure 4.2B). These results confirm that β-ARR2 is
present in the photoreceptor cell layer. There is a basal immunofluorescent signal in the
β-arr2
-/-
mouse retina, and this may be due to the presence of β-ARR1. However, it is
difficult to assess whether this signal is specific for β-ARR1 or a non-specific
background signal. Therefore, β-ARR1 may also be present in the same layers of the
retina as β-ARR2, although β-ARR2 is more highly expressed in the photoreceptor
layers.
Further IHC studies demonstrated that the visual arrestins (ARR1 and ARR4)
colocalize with β-ARR2 in the photoreceptor layer of WT mouse retinas (Figure 4.3).
Figure 4.3A-C demonstrates the cellular localization of ARR1 and ARR4 in a light-
adapted mouse retina. ARR1 is present in rods and cones (Figure 4.3A), while ARR4 is
present only in cones (Figure 4.3B). The immunoreactive signals overlap in the cone
photoreceptors (Figure 4.3C). The immunoreactive signal of β-ARR2 co-localizes with
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Figure 4.3. Immunohistochemistry analysis of WT mouse retina vertical sections, with
higher magnification focus on the photoreceptor outer segments (OS), inner segments
(IS), outer nuclear layer (ONL), and outer plexiform layer (OPL). A. Retina is
immunologically stained with mouse monoclonal anti-ARR1 primary antibody D9F2
(green). B. Retina is immunologically stained with rabbit polyclonal anti-ARR4 primary
antibody LUMIj-mCAR (red). C. Merge of A and B, which were prepared on the same
retinal section. Insets emphasize the overlap of the two signals. D. Retina is
immunologically stained with mouse monoclonal anti- β-ARR2 primary antibody H-9
(Santa Cruz) (green). E. Retina is immunologically stained with anti-ARR4 primary
antibody (same as B). F. Merge of D and E, which were prepared on the same retinal
section. G. Retina is immunologically stained with rabbit polyclonal anti- β-ARR1 and β-
ARR2 primary antibody A2CT (green). H. Retina is immunologically stained with anti-
ARR1 primary antibody (same as A) (red). I. Merge of G and H, which were prepared on
the same retinal section.
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ARR4 in mouse cones (Figure 4.3D-F), particularly in the pedicle of the cone and in the
cone inner and outer segments (Figure 4.3F insets). The immunoreactive signals of
both β-arrestins ( β-ARR1/ β-ARR2) overlap with the signal of ARR1 in mouse rod outer
segments and cone pedicles (Figure 4.3I insets).
4.3.2 DRD4 is expressed in mouse cone photoreceptors
We confirmed the immunoreactive expression of a DRD4 protein in the
photoreceptor regions where visual arrestins are highly expressed. Previously, DRD4
cellular localization has been studied through the use of in situ hybridization studies to
determine the location of DRD4 mRNA in rodent retinas (Klitten et al., 2008; Kim et al.,
2010; Li et al., 2013). This is likely because the antibodies used to identify DRD4 protein
expression have been ineffective and/or non-specific (Van Craenenbroeck et al., 2005;
Bodei et al., 2009). Recent evidence suggests that one anti-DRD4 antibody, N-20, is
specific for DRD4 in mouse retinas, because of its reduced signal in Drd4
-/-
retina
sections compared to wild-type (for mouse details, see section 2.3) (Deming et al.,
2015a) (in press). Figure 4.4A-F also demonstrates the specificity of N-20 for human
DRD4 in transfected HEK293T cells. The top row shows the specific overlap of the N-20
immunoreactive signal (green) with the FLAG tag signal (red) in HEK293T transfected
with FLAG-DRD4.4. N-20 does not demonstrate immunoreactive staining in non-
transfected HEK293T. Using the N-20 antibody, we confirmed the expression of DRD4
protein in mouse photoreceptor inner segments (Figure 4.4G). The DRD4 signal (green)
overlaps with the ARR4 signal (red) in mouse cone inner segments.
The specificity of the DRD4 antibody was determined by comparison of control
C57Bl/6J mouse retinas with Drd4
-/-
mouse retinas. The Santa Cruz Biotechnologies
83

antibody N-20 gave the most reliable results out of six tested. In transfected HEK293T,
N-20 clearly and specifically labels human DRD4 using IHC (Figure 4.4A-F) and
immunoblot analysis (Deming et al., 2015a) (in press). In immunoblot applications using
homogenized mouse retinas, no specific band for DRD4 was detectable using this
antibody. After multiple rounds of IHC protocol optimization, a specific and repeatable

Figure 4.4. Immunohistochemistry analysis of DRD4 in transfected HEK293T and
mouse photoreceptors. A-F. HEK293T transfected with FLAG-DRD4.4 (A-C) or non-
transfected HEK293T (D-F). A and D were immunologically labeled with anti-DRD4
goat polyclonal primary antibody N-20 (Santa Cruz), followed by an Alexa-Fluor 488
conjugated secondary antibody (green); B and E were labeled with M1 anti-FLAG
primary antibody followed by an Alexa-Fluor 568 conjugated secondary antibody
(red). C and F show a merge of A and B or D and E, respectively, in combination with
DAPI nuclear stain (blue). G. WT mouse retina section, with higher magnification
focus on the photoreceptor outer segments (OS), inner segments (IS), and outer
nuclear layer (ONL). DRD4 is labeled with N-20 (see A), followed by an Alexa-Fluor
488 conjugated secondary antibody (green). ARR4 is labeled with anti-ARR4 rabbit
polyclonal antibody LUMIj-mCAR, followed by an Alexa-Fluor 568 conjugated
secondary antibody (red).
84

signal was observed in retinas from wild-type mice compared to Drd4
-/-
. The differing
results between the two applications may be due to the different state of the DRD4
protein (tissue homogenized and denatured for immunoblot, versus fixed and frozen for
IHC), or different buffers and conditions needed for the two different detection methods.
4.3.3 Co-IP of DRD4 with ARR4
Co-IP studies were done to determine if DRD4 interacts directly with visual arrestins
when co-expressed in HEK293T and if this interaction is dependent on DA stimulation.
Co-IP using an antibody recognizing human DRD4, followed by immunoblot analysis,
showed that ARR4 was pulled down along with HA-DRD4.4, suggesting a protein-
protein interaction between ARR4 and DRD4 (Figure 4.5A) (n=1). This specific binding
interaction is only seen when both plasmids encoding DRD4 and ARR4 were co-
transfected into the HEK293T, and when the cells were stimulated with dopamine (DA)
for 30 min.
The interaction was verified by the reverse co-IP, using an anti-human ARR4
antibody followed by immunoblot analysis with an anti-HA antibody, and similar results
were obtained (Figure 4.5B) (n=2). Again, no detectable immunoreactive proteins from
the pull-down were observed without DA stimulation. The intensities of the bands in the
images were quantified using an image analysis software, and the results of each co-IP
are presented in Figure 4.5C and D. These results are in contrast to the pulldown of β-
ARR2 with DRD4, which occurs with or without DA stimulation (Spooren et al., 2010).
Co-IP with ARR1 was also tested with the appropriate controls and an ARR1-specific
antibody, but no detectable pulldown was observed with or without DA stimulation (data
not shown).
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4.3.4 Internalization of DRD4 requires two different arrestins
Classic GPCR internalization experiments were performed to determine under
which conditions, if any, DRD4 was internalized. The first set of experiments tested co-
expression of human DRD4 with one arrestin at a time, including both visual and β-
arrestins, and confirmed that DRD4 co-transfected with either β-ARR1 or β-ARR2 did

Figure 4.5. Immunoblot analysis of co-immunoprecipitation (co-IP) of HA-tagged
DRD4 with ARR4. A. HEK293T cells were transfected with HA-DRD4.4, ARR4, or
both (+/- lanes) and incubated for 30 min with either regular medium (See section
2.4) or regular medium plus 10 µM DA (+/- DA lanes). Co-IP pull-down was
performed using beads with anti-HA antibody, and both supernatant (unbound
protein) and eluate from beads were analyzed on SDS-PAGE, then analyzed using
immunoblot analysis (anti-ARR4, LUMIf-hCAR). Arrows emphasize the ARR4-
specific band in each blot at about 60 kDa. B. The reverse co-IP pulldown was
repeated using beads with anti-ARR4 antibody (rabbit polyclonal LUMIf-hCAR) and
immunoblot analysis (anti-HA) of supernatant and bead-bound protein. Arrows
emphasize the HA-DRD4.4 band in each blot at 50 kDa. C. Band intensities from the
co-IP blot (beads) in A were calculated, compared to the intensity of the band with
DRD4 and ARR4 but without DA (set as 1), and the ratios to this control were
charted. D. Band intensitites from the co-IP blot (beads) in B were calculated and
charted in the same way as C.
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not allow internalization (Figure 4.6A). This was consistent with published results
(Spooren et al., 2010). We additionally co-transfected DRD4 with each of the visual
arrestins, ARR1 and ARR4, and no receptor internalization was detectable when either
visual arrestin was co-expressed with DRD4 (Figure 4.6A).
In each experiment, a positive control ( β2 adrenergic receptor (B2AR) with β-ARR1,
stimulated by 10 µM isoproterenol) was also included to verify that the experimental
conditions allow internalization of a classically-sequestered GPCR (von Zastrow and
Kobilka, 1992). B2AR was clearly internalized when stimulated with its agonist,
isoproterenol (iso) (Figure 4.6C), indicating that the HEK293T cells we used contain the
essential GPCR internalization components, and that the stimulation conditions (30 min
at 37
o
C) allow internalization of at least one GPCR.
Next, two arrestins were co-transfected with DRD4 ( β-ARR1+ β-ARR2,
ARR1+ARR4, β-ARR1+ARR1, β-ARR1+ARR4, β-ARR2+ARR1, or β-ARR2+ARR4).
Preliminary experiments indicated that 30 min of DA stimulation (10 µM) was the
optimum time needed to observe internalization, so all further experiments were done at
this time point. After 30 min of DA stimulation, specific combinations of arrestins did
allow DRD4 internalization (Figure 4.7B). Internalization is observed with only three of
the six combinations: β-ARR1+ARR1, β-ARR1+ARR4, and β-ARR2+ARR4.
Visual results were confirmed by counting the number of cells with internalized
receptors compared to the total number of DRD4-expressing cells in the images (Figure
4.6B and 4.7C). The three combinations of arrestins allow internalization that is
significantly different from the baseline, which is set as the average values of all
unstimulated cells (no DA added to identical medium for the same incubation time). Two
87

of the combinations of arrestins allow a 6.7 fold greater number of cells demonstrating
DRD4 internalization compared to non-stimulated baseline ( β-ARR1+ARR4, p<0.01;
and β-ARR1+ARR1, p<0.05), while β-ARR2+ARR4 is 5.8 fold greater than baseline

Figure 4.6. Immunohistochemistry of DRD4 in vitro internalization experiment with
single arrestin co-expressed. A. HEK293T with FLAG-DRD4.4 (labeled with anti-
FLAG followed by anti-mouse AlexaFluor 488, green) co-transfected with one
arrestin in each sample: β-ARR1, β-ARR2, ARR1, or ARR4. The top row groups
received regular medium (DMEM with 10% FBS) for 30 min, while the bottom row
groups received regular medium plus 10 µM dopamine (DA) for 30 min. B.
Quantification of the results in A, presented as fold increase relative to baseline of
the average of all unstimulated control groups, set at 1 (dotted line). Values are
mean +/- standard deviation. C. HEK293T with FLAG-B2AR (labeled as in A) co-
transfected with β-ARR1, included as a positive control. The top panel was treated
with regular medium for 30 min, while the bottom panel received regular medium plus
10 µM isoproterenol (iso).
88

(p<0.01). The other combinations of arrestins are not significantly different from the
baseline.
Other methods were also used to quantify the internalization of DRD4 with
combinations of arrestins. Inspired by other GPCR internalization studies, we used flow
cytometry (Hislop and von Zastrow, 2011; Pampillo and Babwah, 2015) and in-cell

Figure 4.7. Immunohistochemistry of DRD4 in vitro internalization experiment with
two arrestins co-expressed. A and B. HEK293T with FLAG-DRD4.4 (labeled with
anti-FLAG followed by anti-mouse AlexaFluor 488, green) co-transfected with two
arrestins in each sample (labeled above each column). Top row groups were treated
with regular medium (DMEM+10% FBS) for 30 min, while bottom row groups were
treated with regular medium plus 10 µM dopamine (DA). Receptor internalization is
emphasized with white arrows. C. Quantification of the results in A and B, presented
as fold increase relative to baseline of the average of all unstimulated control groups,
set at 1 (dotted line). Values are mean +/- standard deviation.
89

western assays (Miller, 2004), and the results of these studies were similar to the
manual counting described above. These methods are summarized in Figure 4.1.
However, due to variation between experiments using these approaches, analysis of the
results did not reach statistical significance (Figure 4.8). The variation between
experiments could be due to differences in transfection efficiency, brightness of the
fluorophore at the time of sampling, or other experimental factors.
In our experiments, we co-transfected three cDNA plasmid constructs encoding
DRD4 and two arrestins at a time. Not all transfected cells will express all three
proteins, but with our experimental conditions and subsequent observations, the cells
with DRD4 expression usually (greater than 50%) also immunologically stained for the
appropriate arrestins; however, this observation was not quantified. In our quantification
of DRD4 internalization, in the positive groups about 30% of cells with DRD4 expression

Figure 4.8. Alternate methods for quantification of the internalization of DRD4 with
different arrestins present. A. Flow cytometry results, reported as the mean percent
of positive cells in the stimulated group minus the unstimulated group, ± standard
deviation. Arrows indicate the most positive groups that are consistent with IHC
results. B. In-cell western results, reported as the mean difference in intensity
between the stimulated group versus unstimulated, ± standard deviation. Arrows
indicate the most positive groups that are consistent with IHC results.
90

displayed DRD4 internalization, compared with about 5% in unstimulated groups.
Compared with the positive control, the β2 adrenergic receptor (B2AR), in which about
75% of cells display internalization of the receptor, this percentage is relatively low. The
lower percentage may be due to the decreased chance of co-transfecting three cDNA
plasmid constructs instead of two, in addition to the more subtle endocytosis observed
for DRD4 compared to B2AR.
4.3.5 Translocation of ARR4 to plasma membrane
An important feature of arrestin-mediated GPCR desensitization is translocation of
the arrestin from the cytoplasm to the plasma membrane. To study this, we looked at
the cellular immunological localization of ARR4 in transfected HEK293T cells with
different durations of DA stimulation (15 to 60 min), or without DA stimulation for the
same time periods. This allowed us to observe the cellular location of ARR4 over time,
including whether or not it translocates to DRD4 at the cell membrane and the length of
time it remains there. Without DA stimulation, ARR4 is diffusely immunologically stained
within the cytoplasm with little to no overlap with DRD4 at the plasma membrane
(Figure 4.9A,G, and M). Without β-ARR1 or β-ARR2, there is no significant increase in
the amount of DRD4 and ARR4 overlap (Figure 4.9A-F). When β-ARR1 or β-ARR2 is
present, after 20 min of DA stimulation ARR4 appears to be localized with DRD4 at the
plasma membrane (Figure 4.9I and O, large arrowheads), and this is prior to DRD4
internalization. At 30 min, ARR4 localizes with DRD4 at the plasma membrane (Figure
4.9J and P, large arrowheads) and on some, but not all, of the internalized DRD4
(Figure 4.9J and P, small arrowheads). After 45 or 60 min of DA stimulation, strong
91

Figure 4.9. Immunohistochemistry of FLAG-DRD4 (labeled with anti-FLAG followed
by anti-mouse AlexaFluor 488, green) and ARR4 (labeled with LUMIf-hCAR followed
by anti-rabbit AlexaFluor 568, red) in HEK293T cells. A-F. FLAG-DRD4 and ARR4
were co-transfected in all groups, and cells were incubated with regular medium (no
stimulation) or regular medium plus 10 µM dopamine (DA) for 15, 20, 30, 45, or 60
min. G-L. FLAG-DRD4, ARR4, and β-ARR1 were co-transfected in all groups, and
the same stimulation conditions as A-F were used. M-R. FLAG-DRD4, ARR4, and β-
ARR2 were co-transfected in all groups with the same stimulation conditions as A-F.
Stronger co-localization of ARR4 and DRD4 is emphasized with large arrowheads,
and internalization of DRD4 is emphasized with small arrowheads. S. Quantification
of the percent of cells with DRD4/ARR4 co-localization under each condition. The fill
color of the bars from white to dark gray indicate the length of incubation, with or
without DA. The shading within each time point emphasizes the groups that were
stimulated with DA. Each group was compared to the baseline (set at 1), which is the
average percent of cells displaying co-localization in all unstimulated groups. Column
statistics determined whether each group was statistically significant from the
baseline (*, p<0.05; **, p<0.01).
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internalization of DRD4 is still observed (Figure 4.9K,L,Q, and R, small arrowheads), but
only minimal co-localization of DRD4 and ARR4 is present (Figure 4.9K,L,Q, and R,
large arrowheads). The co-localization of DRD4 and ARR4 at these time points appears
similar to that of the cells stimulated for 15 min. Further statistical analysis (Figure 4.9S)
determined that the only conditions with DRD4/ARR4 co-localization above baseline
(calculated as the average percent of cells demonstrating co-localization of ARR4 and
DRD4 in the unstimulated populations) were DRD4+ β-ARR1+ARR4 and DRD4+ β-
ARR2+ARR4, with either 20 or 30 min of DA stimulation. All other populations were not
significantly higher than baseline, although at 15, 45, and 60 min the amount of co-
localization appears slightly higher when either β-ARR1 or β-arr2 is present (Figure
4.9S).

93

4.4 Discussion
In this study, we demonstrated that DRD4 and ARR4 interact in vitro in a dopamine-
dependent manner, and that DRD4 is internalized after dopamine stimulation, as long
as the correct combination of both a β-arrestin ( β-ARR1 or β-ARR2) and visual arrestin
(ARR1 or ARR4) are expressed in the same cell. We further showed that these in vitro
observations are potentially relevant and may be extended to systems in vivo, as DRD4
is expressed in mouse photoreceptor inner segments along with visual arrestins and β-
ARR2.
4.4.1 DRD4 expression in photoreceptors with visual arrestin
In the retina, visual arrestins are co-expressed with β-ARR2 (Figure 4.2B and 4.3)
and may contribute to desensitization of non-opsin GPCRs. Previously, DRD4 has been
shown to act through the G-protein cone transducin in mesencephalic cells (Yamaguchi
et al., 1997), indicating that the cone opsins (which are also GPCRs) and DRD4 have a
potential shared target of activation. We propose that they may also share components
in their desensitization pathway. The activated cone opsins can be bound and
desensitized by either ARR4 or ARR1 (Nikonov et al., 2008). In mouse retinas, we show
that DRD4 and ARR4 are both present in the inner segments of cones (Figure 4.4B),
which means that these two proteins may potentially interact and function together in
the mouse retina.
4.4.2 Co-immunoprecipitation of DRD4 with ARR4 in vitro
To test the potential interaction of DRD4 with ARR4 and the other three arrestins,
we set up in vitro experiments in which DRD4 was co-transfected with an arrestin in
HEK293T cells. We showed with co-IP studies that DRD4 and ARR4 do interact when
94

overexpressed in HEK293T cells, but only when the transfected cells are stimulated
with DA (Figure 4.5). Co-IP studies have previously shown that DRD4 interacts with β-
ARR1 and β-ARR2, but that this interaction is not dependent on DA stimulation
(Spooren et al., 2010). The same study also reported that the intracellular portion of
DRD4 is phosphorylated in HEK293T, and the phosphorylation is also not dependent on
DA stimulation but is instead observed constitutively (Spooren et al., 2010). DRD4
phosphorylation is likely performed by the GPCR kinase, GRK2, because GRK2 is
known to phosphorylate the other D2-like receptors (Kim et al., 2001), and GRK2 is
present in photoreceptors as well (de Almeida Gomes and Ventura, 2004).
The DA-independent phosphorylation and β-arrestin binding of DRD4 indicates that
these processes may not be a part of the DRD4 desensitization pathway, since DRD4
was not activated by DA. In contrast to these results, we show that the binding of ARR4
is dependent on DA stimulation, indicating that it is this interaction which is specific to
receptor desensitization. Because only the presence of both a β-arrestin and a visual
arrestin allows detectable DA-dependent internalization, the interaction of ARR4 or
ARR1 with DRD4 must be an essential step for the observed desensitization and
sequestration of the receptor.
Our co-IP experiments indicate that the overexpression of a β-arrestin is not
required for ARR4 binding to DRD4, although it is necessary for the observed
internalization of the DA-stimulated receptor. Based on the internalization results, we
repeated the co-IP with HEK293T that were co-transfected with plasmids encoding
DRD4, ARR4, and β-ARR1 to determine whether ARR4 would display increased
binding to DRD4 when β-ARR1 is also present. Any pulldown of DRD4 with ARR4 that
95

may have occurred was below detection level with our immunoblot analysis. This may
be because in preliminary immunoblot analysis, we observed that decreasing the
amount of plasmid DNA in the transfection mixture leads to a decrease in the total
amount of the corresponding protein in transfected cells (data not shown). Because of
this observation we hypothesize that the amount of each arrestin expressed in the
HEK293T is decreased when three plasmids are co-transfected instead of two (Section
2.1), so the amount of ARR4 may have been decreased below the detection limit in the
experiments including β-ARR1. Alternatively, β-ARR1 may compete with ARR4 for the
same DRD4 binding site(s), thereby decreasing the amount of ARR4 bound to DRD4 to
undetectable levels.
4.4.3 Internalization of DRD4
In further in vitro experiments, we demonstrated that the overexpression of each
individual arrestin with DRD4 does not allow sequestration of the receptor (Figure 4.6A
and B). This was in contrast to our positive control, B2AR, which demonstrated robust
internalization under identical conditions upon receptor activation by isoproterenol
(Figure 4.6C). However, we were interested in the mechanism of DRD4 desensitization
in photoreceptors and pinealocytes, where DRD4 expression is circadian. In these
highly specialized neurons, the two β-arrestins are expressed along with the two visual
arrestins. We hypothesized that a combination of arrestins would allow desensitization
and sequestration of DRD4, so we tested the co-expression of two arrestins at a time
with DRD4.
We were surprised by the initial and repeated results of these experiments, which
clearly demonstrated the requirement for at least two different arrestins to be co-
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expressed for DRD4 internalization (Figure 4.7). Interestingly, with β-ARR1 and β-ARR2
co-expressed, no internalization is observed. This indicates that the internalization we
observe may not occur in cells not expressing visual arrestins. Additionally,
combinations of either ARR1 and ARR4 or β-ARR2 and ARR1 do not display detectable
internalization. Because three out of six combinations of two arrestins with DRD4 do not
demonstrate internalization of DRD4, the internalization observed is not merely an
artifact of arrestin overexpression. Instead, the observed specific internalization is
dependent on the presence of a unique combination of arrestins and the stimulation of
DA.
In any population of transiently transfected cells, there will be a group of cells that
do not appear to express the transfected construct. Co-transfection of two or more
mammalian expression plasmids also complicates this problem, as it can be difficult to
determine which cells simultaneously express both proteins. Because of the inclusion of
the appropriate controls (unstimulated populations for each group, DRD4 with each
arrestin individually, and all six possible combinations of two arrestins), it is clear that
the internalization observed is due to the presence and expression of specific arrestins
and not differences in transfection efficiency between plasmids. In addition, the results
were replicated in multiple experiments (n=4) and with multiple expression constructs
for DRD4 (HA-DRD4.4, FLAG-DRD4.4) and β-arrestins (HA- βarr1, FLAG- βarr1, HA-
βarr2, and FLAG- βarr2). Interestingly, when we transfected the same constructs into
COS-7 cells (Gluzman, 1981; Pierce et al., 2000) and stimulated with DA, no
internalization was observed for any group (data not shown). Isoproterenol-stimulated
B2AR was still internalized robustly in COS-7 cells. This set of experiments in COS7
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cells suggests that HEK293T cells are unique, perhaps because of their human origin or
neuronal gene expression characteristics (reviewed in (Thomas and Smart, 2005)), that
allows the detectable internalization of DRD4, while COS-7 cells are missing at least
one critical component for DRD4 internalization.
From these experiments, it is unclear why DRD4 would require two arrestins. To our
knowledge, there is no other GPCR that requires the expression of two arrestins to
undergo internalization after desensitization, so DRD4 appears to be the first and
perhaps a unique GPCR that requires this mechanism for desensitization and/or
internalization. Since each arrestin alone is not sufficient to allow internalization of
DRD4, we hypothesize that each arrestin performs a specific and perhaps sequential
role in the desensitization and internalization process. Furthermore, we have shown that
the presence of two β-arrestins or two visual arrestins also does not allow
internalization. These cumulative observations lead us to propose a more defined
hypothesis. In each cell where internalization occurs, β-ARR1 or β-ARR2 performs a
similar role to one another, while visual ARR1 or ARR4 also perform a similar role that
is distinct from that of a β-arrestin.
While β-ARR1 and β-ARR2 can perform unique functions outside of GPCR
desensitization (reviewed in (Shenoy and Lefkowitz, 2003), they have in common an
ability to bind phosphorylated GPCRs and recruit endocytic machinery to begin the
process of clathrin-coated reuptake (reviewed in (Walther and Ferguson, 2013)). The
abiilty of visual arrestins to take part in clathrin-coated endocytosis has not been as
thoroughly investigated; however, there are no reports that ARR4 is able to interact with
proteins in the endocytic machinery. On the other hand, ARR1 is able to bind and
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modulate N-ethylmaleimide sensitive factor (NSF) (Huang et al., 2010) and Adaptor
protein 2 (AP2) (Orem et al., 2006; Moaven et al., 2013), proteins that are essential
components in the SNARE complex and the clathrin-coated endocytosis machinery,
repectively.
However, visual arrestins have in common their ability to bind GPCRs and to
desensitize light-activated, phosphorylated opsins in all photoreceptors (Kuhn et al.,
1984; Xu et al., 1997; Mendez et al., 2000; Nikonov et al., 2008). Based on these
classical studies, as well our cumulative data, we propose that the primary contribution
of visual arrestin involves a critical agonist-dependent binding to DRD4 and possibly a
comformational and affinity shift. Subsequently, β-arrestin, which binds to DRD4 with or
without DA stimulation, may then be able to recruit essential components of the
endocytic machinery once DRD4 is fully desensitized by visual arrestin, which it is
unable to do without visual arrestin binding. It is also possible that β-arrestin binds one
component of the internalization scaffold, such as clathrin, while visual arrestin binds
another component, such as AP2. Further studies are ongoing to determine the specific
role each arrestin contributes in the desensitization and internalization of DRD4.
4.4.4 Quantification of Internalization
There are many published methods of quantifying GPCR internalization in vitro.
Nearly all of these methods measure the decrease in the presence of receptors on the
cell membrane after internalization (reviewed in (Milligan, 2003)). In the case of DRD4
internalization, the amount of receptor that moves into the cell is small compared to the
total number of receptors that are expressed on the plasma membrane. This is clear
from the IHC images of DRD4 internalization, in which there is still a stronger
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immunofluorescent signal on the membrane compared to the signal of the internalized
receptors (Figure 4.7B).
The quantification results displayed in Figure 4.6B and 4.7C were obtained through
direct counting from IHC images obtained in four separate experiments. We counted the
total number of cells with DRD4 expressed and the number of cells with DRD4
internalized. The fraction of cells with DRD4 internalized was calculated for each group,
including the non-stimulated controls. The non-stimulated cells occasionally also appear
to display internalization (about 5% of the cells), so it was appropriate to use these as
controls to ensure that the internalization observed in the other groups was truly
agonist-dependent. The average of the non-stimulated control groups was set as a
baseline of 1, and all stimulated groups were compared to this baseline. The fold
difference was plotted in figure 4.6B and 4.7C.
4.4.5 Translocation of ARR4 to plasma membrane
A key feature of arrestin-mediated GPCR sequestration is migration of arrestin from
the cytoplasm to the plasma membrane in order to interact with the intracellular portions
(loops or C-terminus) of the GPCR (Oakley et al., 2002). One study reported that β-
ARR2 also translocates to the plasma membrane to co-localize with DRD4 in response
to DA stimulation (Cho et al., 2006), but a more recent report was unable to confirm the
observation (Spooren et al., 2010). With immunohistochemistry tools, we labeled DRD4
and ARR4 in co-transfected HEK293T cells to determine the subcellular localization of
ARR4 with or without the presence of β-ARR1 or β-ARR2, at multiple durations after DA
stimulation (Figure 4.9). The use of sequential DA stimulation timepoints allowed
observation of the subcellular location of ARR4 without stimulation (at 15, 20, 30, 45,
100

and 60 min), after stimulation but before DRD4 internalization (15 and 20 min), at the
peak of DRD4 internalization (30 min), and after DRD4 internalization has occurred (45
and 60 min). We noted that both DA and beta-arrestin were required for stronger co-
localization of DRD4 and ARR4 in HEK293T (Figure 4.9). This is in contrast to our initial
findings that over expression of beta-arrestin is not required for co-IP pulldown of ARR4
with DRD4 (Figure 4.5). The time course of ARR4 translocation we observed in vitro is
similar to the time course of visual arrestin translocation to the photoreceptor outer
segments, which occurs within 20 min of light exposure (Broekhuyse et al., 1985;
Whelan and McGinnis, 1988; Zhu et al., 2002; Zhang et al., 2003a).
The subcellular redistribution of ARR4 is similar to the translocation of β-arrestin
observed with other GPCRs, where β-arrestin translocation occurs rapidly and
continues through 30 min of stimulation (Oakley et al., 2002). Many GPCRs
demonstrate a much faster time course for β-arrestin translocation (Oakley et al., 2002),
which occurs within two min. of agonist stimulation, in contrast to 15-20 min. for ARR4
and DRD4. The time discrepancy may be explained by the relative rates of action of
these GPCRs compared to DRD4. Many GPCRs in neurons, muscle, and other tissue
require very fast activation and desensitization for sensitive response to signals.
DRD4, on the other hand, has been hypothesized to be a slower-acting GPCR,
based on its constitutive phosphorylation and β-arrestin binding, which may slow down
its signal in vivo (Spooren et al., 2010). In addition, our initial hypothesis is that DRD4
sequestration is important in regulating the diurnal physiology in photoreceptors and
pinealocytes, because of its unique circadian regulation in these specialized neurons.
This modulation would only be required once per day, when the lights turn on, to
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decrease DRD4’s presence on the photoreceptor membrane. The decrease in DRD4
mRNA expression and increase of dopamine synthesis has been observed within a brief
one hour window of light onset (Kim et al., 2010), so a relatively slow recruitment of
ARR1 or ARR4 and subequent desensitization would not impact the overall circadian
regulation of DRD4.

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4.5 Conclusion
These experiments reveal a surprising new potential mechanism for desensitization
and sequestration of GPCRs in cells where visual arrestins are highly expressed,
primarily in the photoreceptors and pinealocytes.
Our results support the hypothesis that DRD4 is desensitized and internalized by a
unique mechanism in the photoreceptors and pinealocytes that is dependent on a dual
expression of visual and beta-arrestin. DRD4, ARR1 and ARR4 are localized to inner
segments and synapses, and the ubiquitously expressed β-arrestins are also expressed
in photoreceptors and pinealocytes. The agonist-dependent internalization of DRD4 is
only observed in vitro when specific combinations of arrestins are present, and all of the
combinations that work contain both a β-arrestin and a visual arrestin.
Based on previous work, activation and expression of the DRD4 signal transduction
cascade in the mouse photoreceptors are critical for maintaining normal daytime high
acuity vision (Jackson et al., 2012). Likewise, without the expression of ARR4 in the
photoreceptors, a similar decrease in contrast sensitivity and normal visual acuity is
observed (Brown et al., 2012 53:ARVO E-Abstract 760/A637; Deming et al., 2015b)
(submitted for publication). We propose that the unique DRD4 desensitization process
demonstrated in this work plays a role in the fine tuning of DRD4 circadian expression
and downstream signaling, including gap junctional coupling and postsynaptic On-
and/or Off- bipolar signal.
In addition, the discovery that two arrestins are required for the internalization of one
GPCR indicates that this may also be the case for other GPCRs that were previously
thought to be resistant to internalization. Further investigation of these GPCRs,
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particularly those which are relevant to dopamine functions, may reveal that they can be
internalized when the proper arrestins are expressed.

104

Conclusions Chapter 5:
Cone arrestin 4 and the dopamine receptor D4 are essential for maintaining normal
visual function. Both proteins are relatively newly discovered, with discoveries in the
early 1990s, and both proteins are still likely to have many undiscovered functions in the
retina and elsewhere. Based on the electrophysiological and cone opsin expression
studies of the Arr4
-/-
mouse, it is clear that Arr4 is carrying out important functions in the
cones that contribute to cone function, as seen in the optokinetic and ERG studies, and
to protein expression, as seen for M-opsin in the young animals. Over time, Arr4 is even
more critical, as cones degenerate and ERG amplitudes decrease in older animals
lacking Arr4. Investigating these functions, we will continue to focus our work on this
important topic of research, so that the molecular mechanisms behind the observed
defective phenotypes can be determined. In future studies, we will investigate other
systems linked to similar defects, including the dopamine system (Popova and
Kupenova, 2011; Popova, 2014; Aung et al., 2014).
From the studies of DRD4 internalization, we revealed that one of the roles of Arr4
in cone health may be the regulation of DRD4 signaling. In in vitro experiments, DRD4
is able to internalize only when both a visual arrestin and a β-arrestin are present. In
other published studies, DRD4 signaling has been shown to be essential for connexin
phosphorylation in the photoreceptors, which controls gap junctional coupling between
rods and cones (Nir et al., 2002; Li et al., 2009; Li et al., 2013). This may provide a
partial explanation of how misregulation of DRD4 leads to problems with daytime and
nighttime vision (Jackson et al., 2012). DRD4 transcription in the retina and pineal gland
is under circadian control, so the removal and turnover of the receptors from the plasma
105

membrane at the appropriate time could be a critical mechanism for regulation of this
signaling pathway. The visual arrestins, including Arr1 and Arr4, may assist with this
removal, recycling and/or degradation, allowing DRD4 protein expression at the plasma
membrane to increase and decrease at the appropriate times during the day. Further
studies will investigate the functional significance of the DRD4 interaction with visual
arrestins by looking at connexin phosphorylation at multiple circadian times in Arr1
-/-
,
Arr4
-/-
, Arr1
-/-
Arr4
-/-
(Arr-DKO), β-arr1
-/-
, and β-arr2
-/-
mice. In addition, gene expression
studies investigating using quantitative real-time PCR (qRT-PCR) in Arr4
-/-
mice will
determine whether Arr4 expression impacts DRD4 mRNA expression, or the expression
of other clock genes that are modulated by DRD4 expression (Hwang et al., 2013).
Critical to the study of Drd4 in vivo are robust and specific antibodies that will
recognize Drd4 protein. Previous analysis of Drd4 expression has largely relied on in
situ hybridization to look at mRNA expression, because of a lack of specific antibodies.
We characterized six different commercial antibodies to determine their specificity for
Drd4, using the Drd4
-/-
mouse as a negative control. Three of six antibodies were
successful in vitro, specifically labeling human DRD4-transfected HEK 293T cells, and
overlapping with the HA- or FLAG- tag on the DRD4 constructs. However, in mouse
retinas there was no difference between control (WT, C57Bl/6J) and Drd4
-/-
for any
antibody except for N-20, which had specific immunological staining in the WT mouse.
This staining labeled the ganglion cell layer well and the photoreceptor inner segments
very dimly. This immno-staining was consistent with previous studies using in situ
hybridization (Cohen et al., 1992; Klitten et al., 2008; Li et al., 2013). Researchers
should be cautious when using commercial antibodies against DRD4, because the
106

results may vary between species (as seen with mDrd4 versus hDRD4 in cell lines) or
the antibodies may recognize similar proteins that are present in one tissue (e.g. mouse
retina) but not another (e.g. HEK 293T cells). The Santa Cruz N-20 antibody is still a
less than ideal antibody, because of the short fixation time required (10 min) and the
dimness of the signal. Future work will be needed to purify and characterize a more
specific and robust antibody that will work in multiple applications. A better antibody
may allow researchers to look at DRD4 cellular protein localization and whether it is
internalized as a part of its down regulation.
Overall this work demonstrated the importance of the DRD4 and ARR4 proteins, as
well as presented a case for their future study in the retina. They are both important for
maintaining high acuity vision, particularly in photoreceptor signal transduction and
downstream postsynaptic pathways, and establishing the link between the two proteins
opens up a new area of research.

107

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Abstract (if available)

Abstract In this dissertation, I investigated classical and alternative roles of Cone Arrestin 4 (ARR4). First, my collaborators and I described visual phenotypes of Arr4 null mice. Subsequently, we discovered a unique arrestin‐driven desensitization mechanism of dopamine receptor D4 (DRD4), a G‐protein coupled receptor (GPCR) involved in specific retinal physiological pathways. GPCRs are responsible for many biological processes, including but not limited to phototransduction, olfaction, sympathetic and parasympathetic signaling. Classical GPCR signaling occurs through a canonical activation pathway, followed by desensitization with the binding of an arrestin. ❧ Two closely related visual arrestins (ARR) play a critical role in shutoff of rod and cone phototransduction. When electrophysiological responses are measured for a single mouse cone photoreceptor, ARR1 expression can substitute for ARR4 in cone pigment desensitization

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Asset Metadata

Creator Deming, Janise D. (author)

Core Title Cone arrestin 4 contributes to vision, cone health, and desensitization of the dopamine receptor D4

School Keck School of Medicine

Degree Doctor of Philosophy

Degree Program Genetic, Molecular and Cellular Biology

Publication Date 07/02/2015

Defense Date 03/11/2015

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

Tag ARR4,arrestin,cone arrestin,dopamine receptor,dopamine receptor D4,DRD4,GPCR,G-protein coupled receptors,OAI-PMH Harvest,phototransduction,retina,vision,visual arrestin,β‐arrestin

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Language English

Contributor Electronically uploaded by the author (provenance)

Advisor Craft, Cheryl M. (committee chair), Chen, Jeannie (committee member), Hinton, David R. (committee member)

Creator Email janise.deming@gmail.com,jddeming@usc.edu

Permanent Link (DOI) https://doi.org/10.25549/usctheses-c3-584780

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