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Multimodal investigation of rod photoreceptor structure and function in healthy and diseased states

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Multimodal investigation of rod photoreceptor structure and function in healthy and diseased states

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Content MULTIMODAL INVESTIGATION OF ROD PHOTORECEPTOR STRUCTURE AND
FUNCTION IN HEALTHY AND DISEASED STATES.
by
Kasey Valerie Rose
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
(NEUROSCIENCE)
May 2024
Copyright 2024 Kasey Valerie Rose

ii
ACKNOWLEDGEMENTS
It indeed takes a village to nurture a Doctor of Philosophy, and I am incredibly fortunate to
be surrounded by a supportive network of mentors, colleagues, friends, and family who have been
my bedrock throughout this journey. In recognition of the profound impact these individuals have
had on my development as a scientist, I would like to extend my heartfelt thanks to all who
contributed to making this PhD thesis possible.
Mentors and Advisors
First and foremost, I am deeply grateful to my PhD advisor, Dr. Jeannie Chen, who, in
2013, recognized my latent potential as a researcher and took a chance by hiring me as a lab
technician. Her belief in my capabilities, expressed through her encouragement to apply to USC’s
Neuroscience Graduate Program, played a big role in shaping my academic journey. Dr. Chen’s
skilled management of my tendency to look for perfection during troubleshooting and data
gathering, along with her encouragement for independent research and thoughtful feedback on
my scientific hypotheses, fostered a nurturing environment that added to my scientific learning
and growth. My much-improved writing skills are a testament to her editorial advice. I will continue
to draw inspiration from her exemplary qualities.
I thank my dissertation committee members, Dr. Cheryl Craft and Dr. Aaron Nagiel, for
their support throughout my PhD journey. I genuinely value their constructive feedback on my
project goals, their thought-provoking ideas that opened up new research avenues, and their
valuable support offered through Fellowship nominations and post-PhD career advice. I also
extend my thanks to Dr. Robert Chow for imparting insights and historical context related to
calcium imaging, and to Dr. Rikard Frederiksen for his technical support, particularly in providing
expertise on infrared equipment and retinal slice preparation in darkness. I am indebted to Dr.
Andrey Andreev for investing countless hours in discussions on multiphoton microscopy, various

iii
image analysis techniques, and creative approaches for connecting IR monoculars to microscope
eye pieces. Our conversations transformed seemingly giant hurdles into manageable challenges
and were instrumental in guiding me through my PhD.
Finally, I express my gratitude to Dr. Gary Martin, my undergraduate mentor at Occidental
College, who kickstarted my career in laboratory research, and without whose influence I might
not have explored the exhilarating path of scientific research after college.
USC Community
I extend my appreciation to the Neuroscience Graduate Program’s faculty, administrative
staff, and fellow students for coordinating engaging retreats, symposia, and speaker series that
provided unique opportunities for discussions on new scientific topics. I would also like to thank
the 2017 cohort, the ZNI support teams, and the irreplaceable members of Dr. Chen’s lab. Their
camaraderie and support have been instrumental in both my professional and personal growth. I
will fondly remember our shared experiences, whether it was commiserating over failed
experiments and navigating the tribulations of the PhD process together, or simply enjoying good
conversations during lab lunches and game nights. Beyond the emotional support from this
community, I would like to sincerely acknowledge the mentors in Dr. Chen’s lab who patiently
guided me in mastering new techniques, enabling me to conduct the research that ultimately
resulted in this finalized manuscript. I want to express special thanks to individuals who have
played significant roles, including Dr. Tian Wang, Yun Yao, Helen He, Dr. Sowmya Lokappa,
Natalie Chen, A.J. Cooper, Dr. Hui Xu, Anakha Ajayan, Xinyan Liang, Dr. Zach Murdock, Dr. Shai
Porat, Dr. Judith Hirsh, Deanna Solorzano, Ramiro Montano, and Benilda Ramos-Cruz.
Friends and Family
I extend my heartfelt gratitude to my family members and the exceptional circle of friends
whose enduring love and encouragement have uplifted me throughout my PhD. The pursuit of a
doctoral degree can be both challenging and isolating at times, and our shared moments outside

iv
the lab, whether on the volleyball court, during hiking, or while enjoying each other’s company
over good food, played a significant role in easing some of the burden. It has been a privilege to
know each and every one of you, and here I thank my biggest cheerleaders:
My lifelong friendships: Samantha Calisto, Cheyenne Coxon, Emily DeMuro, Dr. Morgan
Ingemanson, Dr. Sai Upadhyayula, Bryan Jones, Dr. Miriam Winthrop, Dana Michels, Jon Sarris,
Luc Coulier, Alexandria Dunn, Dr. Nora Benavidez, Dr. Andrey Andreev, Dr. Kristina Shkirkova,
Howard Levy, Mike Sedgwick, Dr. Jillian Shaw, Marco and Liz Bärman, and Dr. Jose Gonzalez.
My parents: Kelly Rose and Sandra Jankowski, who have afforded me limitless
opportunities and support since my earliest of days, championing both my academic and
extracurricular pursuits unconditionally. Whether it was allowing me to undertake a chicken
articulation project for my 7th-grade science fair or cheering me on while I danced, skied, horseback rode, and played volleyball, their love granted me the freedom to explore diverse pursuits
and activities. It was within this nurturing home environment that I could freely experience both
failure and success without fear. This supportive backdrop has been instrumental in shaping the
person I am today and has empowered me to pursue my passions wholeheartedly. I will be forever
thankful to them.
My partner: Dr. Spyridon Michalakis. Words cannot fully capture the depth of my gratitude
for the countless ways he has made my PhD journey more manageable, fulfilling, and fun. He
ensured that I consistently carved out time for other passions beyond the lab, whether it was
playing beach volleyball every Sunday, embarking on food adventures in Japan and Greece, or
nurturing baby avocado plants. This intentional focus on meaningful experiences allowed me to
approach each day in the lab with a refreshed mindset, prepared to give my very best. Beyond
maintaining my sanity outside the lab, he always was available, regardless of the time of day or
his own fatigue, to serve as a sounding board for the multifaceted challenges presented by my
research projects. I am honestly so thankful for his unconditional love and the many ways he has
enriched my life.

v
TABLE OF CONTENTS
ACKNOWLEDGEMENTS............................................................................................................. ii
LIST OF FIGURES ......................................................................................................................vii
LIST OF TABLES......................................................................................................................... ix
LIST OF ABBREVIATIONS .......................................................................................................... x
ABSTRACT.................................................................................................................................. xi
CHAPTER 1 General Introduction ................................................................................................ 1
1.1 Anatomy of the mammalian retina. ..................................................................................... 1
1.2 Vertebrate photoreceptor cells............................................................................................ 2
1.3 Rod photoreceptor outer segment renewal......................................................................... 5
1.4 Rhodopsin........................................................................................................................... 8
1.5 Phototransduction signaling in rods .................................................................................. 10
1.6 Rod dark/light signaling..................................................................................................... 13
1.7 Role of calcium (Ca2+) in rod function ............................................................................... 18
1.8 Inherited retinal degenerations ......................................................................................... 21
CHAPTER 2 Light-regulation of Rhodopsin Distribution during Outer Segment Renewal in
Murine Rod Photoreceptors........................................................................................................ 24
2.1 Summary........................................................................................................................... 24
2.2 Introduction ....................................................................................................................... 25
2.3 Methods ............................................................................................................................ 27
2.4 Results .............................................................................................................................. 37
2.5 Discussion......................................................................................................................... 53
2.6 Supplemental Information................................................................................................. 60
CHAPTER 3 Charge Reversal of Rhodopsin’s Glu134Arg135 Impacts Transducin Localization in
Rods without Affecting Rhodopsin Phosphorylation ................................................................... 63
3.1 Summary........................................................................................................................... 63
3.2 Introduction ....................................................................................................................... 64
3.3 Methods ............................................................................................................................ 67
3.4 Results .............................................................................................................................. 74
3.5 Discussion......................................................................................................................... 83
3.6 Supplemental Information................................................................................................. 89

vi
CHAPTER 4 Imaging Compartmentalized Ca2+ Dynamics in Rod Photoreceptors.................... 91
4.1 Summary........................................................................................................................... 91
4.2 Introduction ....................................................................................................................... 92
4.3 Methods ............................................................................................................................ 95
4.4 Results ............................................................................................................................ 102
4.5 Discussion....................................................................................................................... 115
4.6 Supplemental Information............................................................................................... 121
CONCLUDING REMARKS....................................................................................................... 124
REFERENCES ......................................................................................................................... 128
APPENDIX A: MANUSCRIPT INFORMATION ........................................................................ 162

vii
LIST OF FIGURES
Figure 1.1 | Structure of mouse eye and retina.
Figure 1.2 | Morphology of vertebrate rod and cone photoreceptors.
Figure 1.3 | Diagram of rod outer segment (ROS) renewal.
Figure 1.4 | Structure of rhodopsin.
Figure 1.5 | The phototransduction cascade in a representative rod.
Figure 1.6 | Dark current in rod photoreceptors.
Figure 1.7 | Light-mediated changes in rod photoreceptors.
Figure 1.8 | Ca2+ signaling in different rod photoreceptor compartments.
Figure 2.1 | Expression of Rho-Timer in rod photoreceptors.
Figure 2.2 | Rho-Timer expression does not affect retinal structure or function.
Figure 2.3 | Diurnal changes of Rho-Timer transcript mirrors that of endogenous rhodopsin.
Figure 2.4 | Banding pattern of endogenous phosphorylated rhodopsin in the ROS.
Figure 2.5 | Light history affects Rho-Timer periodicity.
Figure 2.6 | Rho-Timer deficient bands corresponds to structural weak points in the ROS.
Figure 2.7 | Rho-Timer localization in disease models.
Figure S2.1 | Low detection of Rho-Timer with rhodopsin antibodies.
Figure S2.2 | Periodic banding pattern is maintained in darkness for Rho-TimerA mice.
Figure S2.3 | Dim, ambient light does not induce rod loss.
Figure S2.4 | ROS from Nckx1 knockout mice are more curved.
Figure 3.1 | Effect of charge reversal (Arg134Glu135, RE) within the conserved G-protein binding
motif of rhodopsin on retinal function in-vivo.
Figure 3.2 | Phosphorylation of light activated rhodopsin (R*) by rhodopsin kinase (GRK1) is
unchanged in REY retinae.

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Figure 3.3 | Impact of REY mutation on the translocation of transducin and arrestin between rod
compartments.
Figure 3.4 | Quantification of transducin and arrestin translocation in REY retinae.
Figure 3.5 | Diagram of the hypothetical mechanism for a stabilizing rhodopsin-transducin
complex in darkness.
Figure S3.1 | Peeling technique of lyophilized mouse retina for western blot.
Figure S3.2 | Confirmation of cone staining with arrestin-1 using peanut agglutinin (PNA).
Figure S3.3 | Rhodopsin bleaching during steady-state 20-lux light in-vivo.
Figure 4.1 | Preparation of retinal slices from Salsa6fiCre mice.
Figure 4.2 | Spontaneous Ca2+ transients in rod photoreceptors.
Figure 4.3 | Light-exposed Ca2+ signal in wildtype, transducin knockout, REY, and Nckx1 knockout
rods.
Figure 4.4 | Rod mitochondria buffer Ca2+ from between rod outer and inner segments.
Figure 4.5 | Light-insensitive REY rods exhibit light-evoked Ca2+ responses.
Figure 4.6 | Determination of absolute Ca2+ concentrations in rod compartments.
Figure S4.1 | Assessment of retinal preparation and Salsa6f in rods.
Figure S4.2 | Ca2+ measurement pipeline.
Figure S4.3 | Unique spatiotemporal profiles of Ca2+ flares.

ix
LIST OF TABLES
Table 2.1 | Table 2.1 Primer sequences used for quantitative real-time PCR.

x
LIST OF ABBREVIATIONS
RPE – Retinal pigmented epithelium
R(OS) – Rod (outer segment)
R(IS) – Rod (inner segment)
CB – Cell body
SN – Synaptic terminal
RP – Retinitis pigmentosa
CSNB – Congenital stationary night blindness
CNG channels – Cyclic nucleotide-gated channels
NCKX1 – Sodium/calcium-potassium exchanger 1
GPCR – G-protein coupled receptor
TM – Transmembrane
Rho-Timer – Rhodopsin-Timer
ARR1 – Arrestin
GNAT1 – Transducin
GRK1 – Rhodopsin kinase
WT-Rho – Rhodopsin
REY-Rho – Rhodopsin where ERY motif is switched to REY
VGCCs – Voltage-gated calcium channels
ER – Endoplasmic reticulum
WT – Wildtype

xi
ABSTRACT
The retina, a multilayered and light-sensitive tissue at the back of the eye, transforms light
into electrical impulses crucial for visual perception. Dysfunction in any of its cell types can result
in partial or total blindness. Particularly, the loss of rod photoreceptor cells, attributed to either
structural or functional impairment, significantly impacts dim light vision and can result in total
vision loss. Elucidating the structural and functional dynamics in both healthy and diseased states,
especially prior to cell death, is critical for understanding inherited retinal degeneration and
developing novel treatments.
The specialized light-sensitive rod photoreceptors are primed to capture photons due to a
unique structure known as the rod outer segment (OS), containing numerous stacked discs
studded with rhodopsin, the photopigment. The length of the OS is constantly maintained
(completely renewed every ~10 days), with daily incorporation of rhodopsin into new discs
essential for maintaining rod health and function. However, to date, the mechanism governing
rhodopsin incorporation into the new discs remains elusive. Chapter 2 centers on our investigation
into rhodopsin incorporation, where we use a novel fluorescent rhodopsin fusion protein
(Rhodopsin-Timer) to visualize rhodopsin’s arrangement in the OS under various conditions,
including classic cyclic light, artificial lighting environments, and diseased states. Our findings
suggest that rhodopsin incorporation into the OS is not entirely regulated by lighting cues and
works in conjunction with intrinsic cues from retinal clocks. We reveal that prolonged lighting
affects rhodopsin content in new and old discs, and regions with less rhodopsin (dim fluorescence
bands) reflect points of structural fragility. Additionally, in two autosomal recessive retinitis
pigmentosa mouse models, we observe significantly altered rhodopsin incorporation, suggesting
a potential role in disease pathogenesis.

xii
Rhodopsin, the prototypical G-protein coupled receptor (GPCR), undergoes a lighttriggered conformational change, which initiates the phototransduction cascade and the
movement of certain signaling proteins within different compartments of the polarized rod cell.
The highly conserved E(D)RY motif, present in all Class A GPCRs, stabilizes rhodopsin’s basal
and active conformational states, facilitating interactions with G-protein transducin, rhodopsin
kinase (GRK1), and arrestin. Despite indications from heterologous and modeling systems
suggesting the motif’s role in cognate protein interactions, its impact on transducin and arrestin
translocation and GRK1-mediated phosphorylation within intact rods in response to light remains
unclear. In Chapter 3, using a recently CRISPR-generated mouse model, RhoREY/REY (REY), we
explore the effects of the ER → RE charge reversal, finding no impact on rhodopsin
phosphorylation or arrestin movement/binding, indicating that this domain is not essential for their
interaction. However, REY-rhodopsin exhibits impaired transducin coupling, translocating away
from the OS only at very high light intensities. Furthermore, our results add additional evidence
for the existence of a stable, preassembled complex between rhodopsin’s ERY motif and
transducin in darkness.
Rod photoreceptor function and survival are intricately tied to Ca2+ homeostasis. Chronic
imbalances in intracellular Ca2+, either elevated or reduced, have been thought to cause
photoreceptor dysfunction in many retinal diseases, though the specific pathways leading to cell
death remain unclear. Limited direct evidence for these hypotheses stems from challenges in
measuring absolute intracellular Ca2+ concentrations simultaneously across different rod
compartments in healthy and diseased retinas. To address these gaps, Chapter 4 outlines the
development of a two-photon Ca2+ imaging protocol using a transgenic mouse line expressing a
genetically encoded ratiometric Ca2+ indicator exclusively in rods. This transgenic mouse line can
be crossbred with mouse models mimicking certain inherited retinal diseases. We show that the
protocol successfully enables 1) the preparation of live-retinal slices, 2) imaging of Ca2+ dynamics
through muti-photon microscopy, 3) “in-slice measurement” of absolute Ca2+ concentrations, and

xiii
4) investigation of functional properties of rods. Our approach allows for the study of rod Ca2+
signaling, offering insights into spatiotemporal Ca2+ dynamics in both healthy and diseased rods.

1
CHAPTER 1
General Introduction
1.1 Anatomy of the mammalian retina.
The mammalian retina, the eye’s innermost layer of tissue, plays a major role in capturing
and processing light for visual perception.
1 Acting similarly to a camera, the eye is designed to
focus light and project images from the external environment onto the retina at the back of the
eye (Figure 1.1). Light first passes through the refractive cornea and pupil (a natural aperture),
then is further focused by the lens onto the retina, a thin (~0.5 mm thick), multi-layered structure.
1
Here, photons must travel through the entire thickness of the retina before striking and activating
the light-sensitive cells, rods and cones. These photoreceptors, which are the most abundant cell
type in the retina, are responsible for converting light into electrical signals, a process known as
phototransduction.2,3 Subsequent neurons, including bipolar, horizontal, and amacrine cells,
further refine the neurosynaptic signals generated by the photoreceptors and encode additional
visual information.
4,5 This processed information concerning the photopic input from the
surrounding visual environment is finally conveyed to the brain’s visual centers via the axons of
ganglion cells for further visual processing and perception. Müller cells, the primary glial cell in
the retina, and the retinal pigmented epithelium (RPE), though not part of the retina, both play
crucial roles in supporting the function and health of the retina and the photoreceptor cells.
6,7
Together, these six primary neuronal cell types (Figure 1.1), comprising ~60 subtypes,8 work in
concert to manage the flow of visual information from light detection to signal modulation before
it reaches the brain.

2
Figure 1.1. Structure of mouse eye and retina. In the mouse eye, light first passes through the cornea
and is subsequently focused by a notably larger lens – characteristic of the murine species compared to
most non-nocturnal vertebrates – directly onto the retina at the back of the eye. The retina is an organized
and layered tissue containing six specific cell types: five neuronal (photoreceptor cells, bipolar cells,
horizontal cells, amacrine cells, and ganglion cells) and one glial (Müller cells). Located in the retina’s most
distal portion, mammalian photoreceptors (rods and cones) respond to visible light, which must traverse
through the entire retina to reach these cells. Neighboring the photoreceptors is the retinal pigmented
epithelium (RPE), essential for photoreceptor maintenance and survival. The retinal layers are delineated
as follows: PL, photoreceptor layer; ONL, outer nuclear layer; OPL, outer plexiform layer; INL, inner nuclear
layer; IPL, inner plexiform layer; GCL, ganglion cell layer. Graphic was created with BioRender.com.
1.2 Vertebrate photoreceptor cells
Rod and cone photoreceptors, which derive their names from the shapes of their outer
segments (OS), exhibit distinct morphological and functional characteristics. These highly
specialized and compartmentalized cells are divided into four main sub-regions: the outer
segment (OS), the inner segment (IS), the cell body (CB), and the synaptic terminal (SN) (Figure
1.2).
9,10 The OS, a modified sensory cilium, contains the light-sensitive photopigments and other
components crucial for phototransduction, arranged within tightly packed membranous discs. This
stacked disc configuration enhances the surface area for light absorption by the visual pigments.
11
Visible light Retina
Ganglion cell
Horizontal cell
Bipolar cell
Rod
Cone
Müller cell
Pigmented
epithelium
cell
Amacrine cell
Light
ONL
OPL
INL
IPL
GCL
PL
Cornea
Lens

3
In mice, the rod OS (ROS) is a cylindrical, rod-shaped sensory cilium, measuring ~25 µm in length
and 1.2 µm in diameter, filled with ~800-1000 densely arranged discs that occupy over two-thirds
of its volume.
10,12,13 Conversely, murine cone OS adopts a conical, tapered shape, being shorter
(~13 µm) and slightly wider (1.5 µm) compared to rods.14 A key distinguishing feature of vertebrate
cones is their “open discs,” which are contiguous invaginations of the OS plasma membrane,
unlike the “closed discs” found in rods11,15 (Figure 1.2). This contiguous disc structure in cone OS
allows unhindered protein diffusion between discs, whereas in the ROS, proteins are confined to
a singular disc, limiting diffusion.
Figure 1.2. Morphology of vertebrate rod and cone photoreceptors. Schematic diagram illustrates the
unique and compartmentalized structure of ciliary photoreceptors. Each cell includes an outer segment
(OS) and an inner segment (IS), interconnected by a connecting cilium (CC). An inset provides a crosssectional view of the CC, revealing axonemal microtubule doublets characteristic of non-motile cilia. In rods,
the OS contains densely stacked membrane-bound discs, whereas in cones, the discs remain contiguous
with the plasma membrane. The inner segment harbors essential organelles for energy production and
protein synthesis, including mitochondria (M) and endoplasmic reticulum (ER). The cell body (CB) houses
the nucleus, containing the cell’s genetic material. At the synaptic terminal (SN), vesicles containing
glutamate are released onto second-order retinal neurons. The inset highlights the photoreceptor synapse
OS
IS
CB
SN
M
ER
CC
Rod Cone

4
and the ribbon structure that anchors many releasable vesicles. This arrangement enables continuous and
graded synaptic transmission crucial for both dark and light conditions. Schematic diagram was adapted
from Dr. J. Chen and Dr. A. Sampath 201310 and was generated with BioRender.com.
A slender, non-motile connecting cilium (CC), characterized by a microtubule-based
axoneme lacking the central pair typically found in motile cilia, links the OS to the IS (Figure 1.2,
upper inset).16 This cilium is a highway between the OS and IS, which facilitates the massive
vertical flow of proteins and lipids essential for light detection (phototransduction) and the OS’s
structural integrity. The IS houses the cell’s metabolic machinery (e.g. mitochondria) alongside
the chief organelles for protein production, such as the Golgi apparatus and endoplasmic
reticulum (ER).9,10 The cell body contains the nucleus, serving as the cell’s control center and
repository of genetic material. At the synapse, specialized ribbon-type structures ensure the
constant flow of the excitatory neurotransmitter glutamate in darkness, dynamically adjusting its
release in response to light (Figure 1.2, lower inset).
17 These ribbons function akin to conveyor
belts, efficiently positioning glutamate-loaded vesicles near the active zone for swift and continual
release.10 The rod synapse (e.g., spherules) contains a single ribbon, while the cone synaptic
terminal (e.g., pedicles) contains around ten pre-synaptic ribbons.14,18
Rods and cones differ not just in structure but also in their functional capabilities,
particularly regarding light sensitivity and wavelength perception. Rods, capable of single photon
detection, are specialized to function in low-light conditions, enabling scotopic vision up to
luminance levels equivalent to starlight on a moonless night. Cones, on the other hand, are suited
for bright light conditions, supporting photopic vision at luminance levels equivalent to daylight.
Mesopic vision, which occurs under light levels such as moonlight, involves both rods and cones.
The unique spectral sensitivities of rods and cones are attributed to the specific photopigments
they express.19 Rods have a single type of visual pigment, rhodopsin, which primarily absorbs
light at ~500 nm.19,20 Cones, depending on the species, vary in the types of opsin they express.
In human retinas, cones express either one of three opsins – blue (430 nm), green (535 nm), and

5
red (560 nm) – each tuned to a specific wavelength, enabling a broad spectrum of color detection
(Figure 1.1).
21-23 On the other hand, murine cones contain two pigments: short-wavelength opsin
(S-opsin) and medium-wavelength opsin (M-opsin), with peak sensitivities at 360 nm and 510 nm,
respectively, indicating a blue-shifted sensitivity compared to humans.20,24,25 Moreover, most
vertebrates are rod dominant, with rods comprising as much as ~95-97% of photoreceptors.
14,26,27
Thus, any disruptions in the structure and function of rod photoreceptors can significantly impact
retinal health. The similarity between the rod-dominant visual sensitivity of mice and the human
peripheral retina28 makes mice an invaluable model for investigating rod structure and function in
physiological and pathophysiological states, which has proven to be immensely beneficial for the
research conducted in this thesis.
1.3 Rod photoreceptor outer segment renewal
Lifelong vision relies on the intricate interaction between the ROS and the RPE, a
monolayer of cells neighboring the rod photoreceptors. The ROS, exposed to high levels of light
daily, risks photo-oxidative damage that can lead to retinal degeneration if not properly
managed.29 The RPE plays a crucial role in mitigating this risk by removing damaged ROS tips
through phagocytosis, thereby preventing the harmful effects of continuous light exposure.7 In the
late 1960s, pioneering work conducted by Young and colleagues first demonstrated the dynamic
renewal process of the ROS (Figure 1.3).30,31 Using autoradiography to detect pulse-labeled H3
-
proteins (mainly rhodopsin) within the ROS, Young et al.30 observed a distinct band of radioactive
material migrating from the ROS base to the tip over time, where it was eventually phagocytosed
by the RPE (Figure 1.3).32,33 These experiments demonstrated the ROS’s ability to preserve its
length through a daily renewal process, wherein ~10% of its volume was continuously renewed
at the ROS base and simultaneously shed at its distal tip. In rodents, the entire length of the ROS
is completely renewed over a period of ~10-12 days30,34 (Figure 1.3). Given that the ROS lacks

6
the biosynthetic machinery for protein synthesis, its daily renewal hinges on the active production
and incorporation of ~2 million rhodopsin molecules,35 essential for capturing light and providing
structural stability. These molecules, equipped with precise targeting signals, journey from the rod
IS (RIS) to the base of the ROS. Concurrently, the ROS base forms new discs at a rate of about
75-100 per day through the expansion and pinching off of the ciliary plasma membrane to form
discrete, closed discs.36,37 These processes, involving protein incorporation, disc formation, and
ROS phagocytosis, are essential cellular mechanisms for the continuous ROS renewal and health
of the rod.
Figure 1.3. Diagram of rod outer segment (ROS) renewal. Illustration of the results from radioautographs
by Richard Young (1967)30 of murine retinas, showing the progression of a radioactive band (purple) in rods
following an injection of methionine3H. The experiment uncovered the dynamic process of ROS renewal,
showing that the discrete, stacked, and densely packed discs within the ROS are continuously added to
the base and shed at the tip over ~10 days in mammals. (1) Before the methionine3H injection, the RPE
ingests the tip of the ROS, ~80-90 discs a day. (2) Peritoneal injection of methionine3H allows for the
visualization of radioactively labeled proteins synthesized after the injection. (3) Thirty minutes postinjection, radioactivity becomes notably concentrated in the RIS, indicating the synthesis of new proteins
utilizing the radiolabeled amino acids. (4) One day after the injection, a distinct radioactive band becomes
RPE
ROS
RIS
CB
SN
Injection of radioactive
amino acids
Time
Day 1
1 3 4 5 6
2
Day 5 Day 10

7
visible at the base of the ROS. (5) The radioactive protein band progressively migrates distally toward the
tip of the ROS. (6) Labeled discs undergo phagocytosis by the neighboring RPE on day 10. ROS, rod outer
segment; RIS, rod inner segment; CB, cell body; SN, synaptic terminal. Diagrammatic summary is inspired
by Richard Young’s drawings and was recreated using BioRender.com.
Research in mammals, fish, and amphibians has revealed a universally conserved
process for ROS renewal,38-42 with temperature and light playing significant roles in modulating
ROS shedding rates. For instance, higher temperatures30,43 and increased light intensities30,34
lead to an increase in RPE phagocytosis, which in turn shortens the ROS. These findings reveal
that vertebrate ROS shedding and its subsequent phagocytosis by the RPE adheres to a daily
rhythm, peaking 1–2 hours after light onset.
33,44 Specifically, daily light exposure activates the
RPE’s phagocytic machinery and triggers the externalization of phosphatidylserine (the “eat me”
signal) at the ROS tip.45,46 This precisely coordinated process is essential for promptly eliminating
the older portion of the ROS (Figure 1.3). Remarkably, this daily rhythm of phagocytosis persists
in conditions of constant darkness or light,47,48 albeit with modified phagocytic rates that
accordingly influence the length of the ROS – slightly elongating in darkness and shortening in
light.49 Hence, RPE phagocytosis of ROS appears to be under circadian control,44,50 showing a
daily activity surge shortly after light onset, with patterns of ROS clearance continuing under
steady light conditions. The specific source of this circadian control, whether it originates in the
retina, the RPE, or a combination of both, remains unclear. Moreover, our understanding of the
mechanisms that regulate both the daily creation of discs and the integration of rhodopsin into
these nascent discs – actions that offset the removal of ROS tips – is currently the focus of
ongoing research. Disruptions to this ROS renewal process, such as mutations affecting protein
trafficking or ciliary machinery, often lead to photoreceptor degeneration.
51 The mechanisms
underlying ROS renewal, including how rhodopsin is integrated into new discs in both healthy and
degenerating rods, are topics of further exploration in Chapter 2.

8
1.4 Rhodopsin
Rhodopsin is a prototypical G-protein coupled receptor (GPCR), functioning as a signal
transducer that converts light into cellular responses. This light-sensitive pigment accounts for
over 90% of the total membrane proteins in ROS discs.52 Besides its primary function of photon
capture, rhodopsin also plays a vital role in maintaining the structure of the ROS. Studies involving
rhodopsin knockout mice53,54 or mutants with impaired trafficking signals, like Q344ter,55
demonstrate that deleterious mutations in rhodopsin can result in the development of faulty or
completely absent ROS. Structurally, rhodopsin consists of seven transmembrane (TM) helical
domains (H1-H7) (Figure 1.4A). It possesses a C-terminal “trafficking guide,”56 notably the 1D4
epitope, which directs the large-scale, unidirectional delivery of rhodopsin from the RIS to the
ROS base for incorporation into new discs. The N-terminal is oriented towards the intradiscal
space, while the flexible C-terminal resides in the cytosol. The C-terminus also contains
phosphorylation sites (Figure 1.4A, stars), several serine/threonine residues,57 which are
necessary for initiating rhodopsin’s deactivation.58
Rhodopsin is composed of two parts: the 11-cis retinal chromophore and the apoprotein
opsin. In darkness, 11-cis retinal is tightly packed within the interior of rhodopsin via a protonated
Schiff’s base linkage to the Lys296 residue on TM H7 (Figure 1.4B), which is further stabilized by
a salt bridge with its counterion Glu113.
59,60 In this configuration, the visual pigment is very stable.61-
63 Furthermore, this arrangement forces the bundle of helices to re-fold and is further stabilized
by another salt bridge between Glu247 (H6) and the ERY motif on H3, creating an “ionic lock” that
keeps rhodopsin in a stable, low-activity dark state64,65 (Figure 1.4B). The ERY motif, a critical and
highly conserved element of class A GPCRs, is indispensable for maintaining rhodopsin’s
structural stability in the dark and has functional consequences for its activation upon exposure
to light (Figure 1.4B).

9
Figure 1.4. Structure of rhodopsin. (A) Murine ROS typically contain around ~800-1000 discrete discs
densely packed with rhodopsin, the visual pigment (inset). Rhodopsin is composed of seven alpha helices
(H1-H7) spanning the disc membrane. The N-terminus is situated within the intradiscal space, while the Cterminus is exposed to the cytosolic space. Several phosphorylation sites (*) are located on the C-terminus.
On the cytosolic edge of transmembrane H3, a highly conserved motif (ERY) is located. (B) 3-D structure
of rhodopsin, with the light-sensitive visual chromophore, 11-cis retinal. 11-cis retinal is covalently bound
to rhodopsin via a protonated Schiff-base linkage to a lysine residue. The ERY motif (orange) plays a crucial
role in maintaining rhodopsin in an “ionic lock” state in darkness. (C) Upon exposure to light, the
photoisomerization of 11-cis retinal (left) to all-trans retinal (right) occurs, triggering a conformational
change in rhodopsin from its inactive state (R) to its activated state (R*). This transformation is fundamental
to the initiation of the visual phototransduction cascade. Graphic was created with BioRender.com.
Light absorption triggers the 11-cis retinal to isomerize to an all-trans conformation,
transforming rhodopsin into its activated state (R*) (Figure 1.4C). This activation disrupts the “ionic
lock” via a sequence of rotameric adjustments, leading to the formation of a stable binding cavity
on the cytoplasmic side of the disc membranes.66 This enables the ERY motif to interact with its
conjugate proteins involved in the phototransduction cascade,67,68 which is further explored in
Chapter 3. Following R* deactivation, the all-trans retinal is released, and biochemical reactions
facilitated by the adjacent RPE regenerate the 11-cis retinal chromophore.69 This allows opsin to
rebind with 11-cis retinal, restoring the light-sensitive rhodopsin and priming it to react to incoming
photons.
A B
C
R R*
Light
Light
11-cis retinal all-trans retinal
N-terminus
C-terminus
Intradiscal
Cytosolic
Disc
N-terminus
C-terminus
H7 H6 H5 H4 H3 H2 H1
Rhodopsin
* * * *
CHO O
H
Intradiscal
Cytosolic
E
R
Y
Intradiscal space
Cytosolic
11-cis retinal
ER
Y

10
1.5 Phototransduction signaling in rods
The phototransduction cascade is a complex biochemical process that transforms photons
into chemical signals. Rods’ capacity to mediate dim light vision primarily stems from two factors:
1) the remarkable sensitivity of rhodopsin, capable of detecting single photons, and 2) the
extensive signal amplification that a solitary rhodopsin molecule undergoes through various
stages of the phototransduction cascade. In darkness, the concentration of cGMP in the ROS is
high (~60 µM), with a small fraction freely circulating in the ROS cytoplasm.70 In this environment,
approximately 1% of cGMP-gated (CNG) channels remain open, facilitating a steady influx of Na+
and Ca2+ ions in the dark.
71-73 This setup renders the channels acutely sensitive to minor changes
in cGMP levels, enabling rods to produce a nuanced array of responses across a broad spectrum
of light intensities.
Activation of the phototransduction cascade begins when a photon is absorbed by
rhodopsin in the ROS disc membrane, inducing a conformational change to its activated state
(R*) (Figure 1.5A). The rapid lateral movement of proteins within the disc membrane74 allows R*
to quickly interact with and activate transducin, a heterotrimeric G-protein comprised of �, �, and
� subunits.75 In its inactive state, the �-subunit of transducin (T�) is bound to GDP. Activation of
transducin by R* triggers an exchange of GDP for GTP on the T� subunit, leading to the
dissociation of the T�-GTP from the rest of the complex (Figure 1.5A).76 This dissociation enables
R* to further activate additional transducin molecules. The quantity of T� units that become
activated hinges on the active duration of R*. In the case of mouse rods, R* has an estimated
lifespan of 80 ms, which leads to the activation of approximately 12 to 20 T�-GTP units for each
R* molecule.67,77 To enable swift activation in response to light, the existence of a pre-assembled,
transient complex between rhodopsin and transducin may play a crucial role in facilitating rapid
and efficient signaling,78,79 an aspect further explored in Chapter 3. Activated T�-GTP then
interacts with phosphodiesterase (PDE), lifting the inhibition of the catalytic PDE� and/or PDE�

11
subunits.
75,76 The activity of PDE leads to the hydrolysis of thousands of cGMP molecules into 5’-
GMP,80 consequently lowering cGMP levels and triggering the closure of CNG channels, which
diminishes the influx of Na+ and Ca2+ into the ROS (Figure 1.5A).10 This reduction in open CNG
channels hyperpolarizes the rod, leading to the closure of voltage-gated Ca2+ channels (VGCC)
in various rod compartments, which ultimately reduces glutamate release at the synapse.
Figure 1.5. The phototransduction cascade in a representative rod. (A) In darkness, CNG (cyclic
nucleotide-gated) channels remain open due to a high concentration of cGMP, permitting an influx of Na+
and Ca2+ ions. The process of converting light into an electrical signal occurs when photons activate the
visual pigment, rhodopsin. This activation leads rhodopsin to undergo a conformation change to an active
state (R*), which in turn activates many transducin molecules. This interaction prompts the exchange of
GDP for GTP on the � subunit (T�) of transducin, causing T� to detach from its �� subunits. The activated
T� can then bind and activate the enzyme phosphodiesterase (PDE), accelerating the breakdown of cGMP.
This reduction in cGMP levels results in the closure of CNG channels, stopping the flow of Na+ and Ca2+
into the rod outer segment and leading to membrane hyperpolarization. This change signals a reduction in
Rhodopsin
Cytosolic
Intradiscal
T! T! T! T! T! " " # #
Closed
Open Dark
Ca2+
Na+
PDE
Arrestin Recoverin
Closed
Closed
Ca2+ Ca2+
Cytosolic
Intradiscal
Cytosolic
Intradiscal
R*
R*
RK
PPP PPP
T! T! T! T! " #
!
# # "
!
# # " ! # # "
GAP complex GC
Ca2+
Na+
GCAPs
Transducin
PDE
Transducin
Activation Deactivation (R*) Deactivation (PDE/T
!)
A
B
C Ca2+
Na+
Open
Open

12
glutamate release at the synapse. (B) For the system to reset and detect new photons, R* must be
deactivated. This process begins with the addition of phosphate groups to R*’s C-terminal by rhodopsin
kinase (RK). Typically, RK’s activity is inhibited by Ca2+-bound recoverin in darkness. However, the
reduction of Ca2+ levels in light removes recoverin’s inhibition, allowing RK to phosphorylate R*.
Deactivation is fully completed once arrestin binds to the phosphorylated R*. (C) To reopen CNG channels,
both activated T� and PDE must be deactivated, and cGMP production needs to increase. The deactivation
of activated T�, leading to PDE returning to its inactive state, occurs through the hydrolysis of bound GTP
to GDP. This hydrolysis can naturally occur through T�’s intrinsic GTPase activity or be facilitated by the
GAP complex. Concurrently, the decrease in Ca2+ levels enables GCAPs (guanylyl cyclase-activating
proteins 1 and 2), now free of Ca2+, to activate guanylyl cyclase (GC), boosting cGMP synthesis. This
creates a feedback loop that reopens CNG channels, restoring the rod’s sensitivity to light. Modified from
Dr. J. Chen and Dr. A. Sampath 201310 and recreated using BioRender.com.
Reversing the activation process is essential for rods’ return to their basal dark-state. This
reversal hinges on the decrease of intracellular Ca2+ levels prompted by the closure of CNG
channels, which in turn leads to quenching of the phototransduction cascade through a series of
steps that aid in deactivating both active intermediates (R* and PDE) and replenishing cGMP.
Phosphorylation initiates the first step in deactivating R* (Figure 1.5B). In rod photoreceptors,
rhodopsin kinase (RK) sequentially phosphorylates a cluster of six to seven serine/threonine
residues located near the C-terminus of R*.
57,81,82 Recoverin, which interacts with RK in a Ca2+-
dependent manner, effectively inhibits RK activity in high Ca2+ conditions. However, when there
is a decrease in Ca2+ induced by light exposure, the recoverin-RK complex dissociates, thus
alleviating the inhibition of the kinase’s catalytic activity83,84 (Figure 1.5B). For timely inactivation
of R* and recovery of the photoresponse, a minimum of three phosphate groups are added to
rhodopsin’s C-terminus.
57,85 This multiple phosphorylation is necessary for high-affinity arrestin
binding,
86,87 which shields R*’s cytoplasmic tip from further T� interaction, and results in the
complete deactivation of rhodopsin (Figure 1.5B). Subsequently, rhodopsin transitions to opsin,
and all-trans retinal is regenerated to 11-cis retinal with the assistance of neighboring RPE cells.
The 11-cis retinal then reassociates with opsin, regenerating rhodopsin and priming it for another
photon absorption event.
Concurrently, the recovery phase also involves turning off both transducin and PDE
activity (Figure 1.5C). This process begins with the inactivation of T�-GTP through GTP hydrolysis

13
to GDP. While T� possesses the inherent ability to hydrolyze GTP, its rate of GTPase activity is
naturally slow.76 This rate is significantly increased by the presence of a GTPase activating protein
(GAP) complex, which regulates G-protein signaling.88 As a result, T�-GTP’s hydrolysis leads to
its separation from PDE�, allowing it to rebind to free T��. Following this dissociation, the PDE
complex is deactivated when PDE�� swiftly recombines with PDE�, halting the breakdown of
cGMP89 (Figure 1.5C). Simultaneously, a reduction in Ca2+ concentration within the ROS
significantly influences the activation of guanylate cyclase-activating proteins (GCAPs), which in
turn enhance cGMP production through guanylate cyclase (GC)90,91 (Figure 1.5C). As intracellular
cGMP levels return to their resting, dark-state concentrations, CNG channels reopen.76 This
reopening allows free Ca2+ to enter the cell, leading to the reinstatement of GCAPs’ inhibition of
GC, and consequently, the dark-state membrane potential is reestablished through the movement
of cations via the CNG channels.10 The intricate interplay between cGMP and Ca2+ signaling is
pivotal for the dynamic regulation of the phototransduction process.
1.6 Rod dark/light signaling
In dark-adapted rods, the movement of Na+ into the ROS through CNG channels and its
exit alongside K+ ions through Na+
/K+
-ATPase and K+ channels in the RIS, respectively, is known
as the “dark current”
92-94 (Figure 1.6). This phenomenon is sustained by a significant intracellular
presence of cGMP which ensures a subset of the CNG channels remain open, thus permitting a
consistent influx of Na+ and Ca2+.
76,95 The inward current primarily consists of Na+ ions (~
85%),95,96 with the remainder predominantly carried by Ca2+,
97,98 and a minimal contribution from
Mg2+.
99 To maintain the ionic equilibrium within rods, two essential extrusion mechanisms are
employed: the simultaneous efflux of Na+ and K+ from the RIS and the removal of Ca2+ through
the Na+
/Ca2+, K+ exchanger 1 (NCKX1) in the ROS. These mechanisms establish a depolarizing
membrane potential around -40 mV100,101 (Figure 1.6). When the rod is depolarized in darkness,

14
specialized L-type voltage-gated calcium channels (VGCCs), particularly the Cav1.4 channels
located at the synapse,102,103 are open, leading to Ca2+ influx. At the synapse, Ca2+ influx triggers
the fusion of synaptic vesicles with the plasma membrane, allowing the continuous release of the
neurotransmitter glutamate onto inner retinal cells in darkness10 (Figure 1.6).
Figure 1.6. Dark current in rod photoreceptors. Schematic overview of cationic currents in light and dark
conditions. In darkness, Na+ and Ca2+ ions flow into the ROS via cyclic nucleotide-gated (CNG) channels,
while Na+ is pumped out by Na+/K+ ATPase in the RIS. This process of inward/outward Na+ movement is
described as the dark current and keeps the rod depolarized around -40 mV. Concurrently, Ca2+ and K+
ions are extruded by the Na+/K+-Ca2+ exchanger 1 (NCKX1) in the ROS, and K+ efflux in the RIS is mediated
by K+ channels. Under these conditions, voltage-gated Ca2+ channels (Cav1.4) in the synapse (SN) allow
an inward flow of Ca2+, enabling continuous release of glutamate. In response to light, phototransduction
leads to the closure of the CNG channels and results in rod hyperpolarization (~60 mV), which decreases
the activity of Cav1.4 channels. This ultimately results in a reduction in glutamate release at the synapse.
ROS, rod outer segment; RIS, rod inner segment; CB, cell body. Diagram was inspired by Purves et al.104
and was created using BioRender.com.
The absorption of light by rhodopsin in the ROS triggers a signaling cascade, resulting in
the closure of CNG channels near the photon absorption site on the disc membrane105 (Figure
NCKX1
Na+/K+
ATPase
K+ channels
Cav1.4
CNG
channels
Dark Light
0 0
-40mV -60mV
- + - +
Ca2+
4 Na+
K+Ca2+
Na+
Ca2+
Na+
4 Na+
K+Ca2+
K+
2 K+
3 Na+
2 K+
3 Na+
X
X
ROS RIS SN
CB
Glutamate
dark
current
K+

15
1.6). This process alters the flow of cations across the ROS plasma membrane, hyperpolarizing
the ROS membrane potential (-70 mV).104 The suppression of the dark current, accompanied by
a wave of hyperpolarization propagating from the ROS to the synapse, induces the closure of
Cav1.4 channels. This closure transiently diminishes Ca2+ influx, leading to alterations in the rate
of glutamate release onto postsynaptic neurons106,107 (Figure 1.6). Concurrently, the ROS
experiences a decline in Ca2+ levels as a result of the closure of select CNG channels and the
efflux of Ca2+ facilitated by NCKX1. This reduction in Ca2+ concentrations triggers a vital calcium
feedback mechanism (described in the previous section), crucial for regulating the recovery phase
of the phototransduction cascade. This regulatory process enables the rod cell to transition back
to its dark state or adapt to brighter light intensities effectively.71,108,109
Illumination levels on Earth change by more than ten orders of magnitude throughout the
24 hour day.
110 To cope with this broad range of light intensities, rods use various strategies to
prevent oversaturation while adjusting their sensitivity and speed of photoresponse according to
ambient light levels. This process, termed light adaptation, significantly broadens the rod’s
operational range.109,110 For instance, the continual decrease of Ca2+ levels in response to light
leads to increased activation of GC by GCAPs (Figure 1.5C), which in turn boosts the production
of cGMP.109 The rise in cGMP concentration increases the amount of open CNG channels,
thereby restoring some of the circulating current and extending the rod’s operating range across
different illumination conditions. Nonetheless, under extremely bright conditions, such as direct
sunlight, the mechanisms employed by rods to extend their operating range are overwhelmed by
the massive activation of rhodopsin, leading to saturation.28 Typically, the transition between rods
and cones enables the visual system to detect a comprehensive spectrum of light intensities, from
single photons (rods) to sunlight conditions (cones).
In response to prolonged exposure to high levels of illumination, particularly within the
mesopic range, rods employ a sophisticated adaptation mechanism involving the substantial
translocation of two critical phototransduction proteins, transducin111 and arrestin112 (Figure

16
1.7).113,114 Under dark conditions, transducin exhibits a strong affinity for ROS disc membranes,
attributed to lipid modifications on its � and �-subunits – specifically, acylation115,116 and
farnesylation,117 respectively. Arrestin, conversely, displays a moderate affinity for microtubules
in darkness, a factor crucial for its localization to the RIS.112,118 Subsequently, in a light-dependent
manner, there is a reciprocal redistribution of transducin and arrestin within the rod cell.
Figure 1.7. Light-mediated changes in rod photoreceptors. Light-driven translocation of signaling
proteins, transducin and arrestin, in mammalian rods. In dark conditions, transducin resides in the disc
membranes of the ROS, anchored via lipid modifications on its � and �� subunits. Arrestin is predominantly
located in the RIS, drawn there by its affinity for microtubules. Upon light exposure, transducin is activated,
prompting the T� and �� subunits to disassociate and relocate to the RIS, a migration facilitated by their
reduced affinity for plasma membranes due to single lipid modifications. Simultaneously, light exposure
triggers arrestin to move towards the ROS, where it binds to its high-affinity partner, phosphorylated
rhodopsin. This reciprocal movement of proteins is a key component of rods’ light adaptation mechanisms,
preventing excessive signaling and damage in bright light conditions. ROS, rod outer segment; RIS, rod
inner segment; CB, cell body; SN, synaptic terminal. Figure was created using BioRender.com.
Continuous bright-light illumination prompts a significant proportion of transducin to leave
the ROS and redistribute throughout the entirety of the rod cell (Figure 1.7). After interacting with
Dark Light
Arrestin
Microtubules
Arrestin
T! " # P
"
#
P P
T!
Transducin Rhodopsin
Dissociated
ROS
RIS
CB
SN

17
R*, transducin dissociates into its separate subunits (�, ��), each bearing only a single lipid
modification. This alteration significantly reduces the subunits’ affinity for the disc membranes,119
thereby enabling their redistribution away from the ROS (Figure 1.7).120 This redistribution
becomes feasible only after surpassing a specific illumination threshold,111 wherein the activated
T�-GTP cannot be rapidly inactivated by the GAP complex,121 leading to its membrane
detachment followed by the relocation of T�� units to the RIS, aided by phosducin.122 This
translocation significantly decreases the quantity of transducin accessible for activation by R*,
leading to a reduction in signal amplification. Consequently, this adjustment decreases rod
sensitivity, enabling it to operate effectively at higher light intensities. Remarkably, recent studies
have demonstrated that this mechanism enables rods to retain functionality even under intense
light conditions,111,123 with evidence suggesting a recovery of up to 15% of their dark current due
to this adaptive translocation.124
On the other hand, arrestin migrates in the opposite direction, drawn towards the ROS by
the presence and accumulation of its high-affinity binding partner, phosphorylated rhodopsin (R*-
P), serving as the key driver of its localization within the ROS112,125 (Figure 1.7). Arrestin exhibits
an affinity for light-activated, phosphorylated rhodopsin (R*-P) that is approximately 10-20 times
higher than for both the non-activated, phosphorylated form (P-R) and the activated,
unphosphorylated counterpart (R*).126 This higher affinity explains arrestin’s relocation from the
RIS to the ROS in response to the large number of R*-P molecules under bright light conditions.114
With arrestin and rhodopsin present in approximately a 0.8:1 ratio,127,128 arrestin’s efficient
translocation under bright light is crucial for effectively silencing R*. The prevailing theory
suggests that the movement of arrestin to the ROS serves as a mechanism for enhanced
deactivation in response to increased light exposure.125,129 Normally, in darkness, arrestin levels
in the ROS are adequate to suppress single photon events. Yet, in bright light, the escalating
need to deactivate numerous R*-Ps quickly depletes the available arrestin in the ROS, prompting
its translocation to effectively silence the rod’s activity.114,123

18
The bidirectional movement of arrestin and transducin unfolds over several minutes, yet
the return to their original locations upon reverting to darkness takes upwards of an hour.111,125,130
This translocation process is driven by passive diffusion that does not require energy,112,129,131
with the movements of arrestin and transducin independent of each other.132 This mechanism not
only broadens the operational range of rods, but likely also serves a protective role, mitigating the
harmful consequences of extended light exposure.114,121,133
1.7 Role of calcium (Ca2+) in rod function
Ca2+ is highly compartmentalized in rod photoreceptors, and its homeostasis plays a
critical role in both rod survival and function.134,135 Contributing to the compartmentalized control
of Ca2+ in rods, specific channels and exchangers regulate Ca2+ flow within individual
compartments (Figure 1.8), with each channel/exchanger fulfilling a vital function within its
designated domain. Changes in intracellular Ca2+ concentrations initiate key processes within the
different sub-regions of the rod cell, including light adaptation and neurotransmitter release
(Figure 1.8).71,134,135 In darkness, the intracellular Ca2+ levels remain relatively elevated, ranging
between 300 nM and 2 µM,134 contingent upon the specific compartment. These Ca2+ levels are
high enough to regulate the phototransduction machinery in the ROS and glutamate release at
the synapse, yet remain below the threshold that would induce Ca2+-mediated cytotoxic
effects.136,137 Conversely, exposure to light precipitates a significant reduction in Ca2+ levels, which
can fall to a range of 30 nM to 700 nM,134 depending on the compartment. The dynamic fluctuation
of Ca2+ concentration within rods under varying light conditions highlights the importance of
controlling intracellular Ca2+ levels to preserve rod homeostasis and functionality (a topic further
explored in Chapter 4).

19
Figure 1.8. Ca2+ signaling in different rod photoreceptor compartments. Calcium ions serve as
essential second messengers in rod photoreceptors. The regulation of Ca2+ is compartmentalized, with
specialized channels/exchangers in each rod sub-region facilitating precise control. (1) The ROS plasma
membrane features cyclic nucleotide-gated (CNG) channels and Na+/K+-Ca2+ exchanger 1 (NCKX1), which
modulate Ca2+ entry/exit crucial for regulating the phototransduction feedback loop. In darkness, Ca2+ levels
are elevated, and light exposure reduces Ca2+ levels due to the closure of CNG channels. This subsequent
decrease in Ca2+, due to CNG channel closure and NCKX1 extrusion of Ca2+, releases inhibitory constraints
on GCAPs and guanylate cyclases. This leads to cGMP synthesis and opening of CNG channels and Ca2+
entry. (2) The RIS houses mitochondria and endoplasmic reticulum (ER), organelles essential for regulating
intracellular Ca2+ levels that control energy production and protein synthesis. Mitochondrial Ca2+
homeostasis is maintained by the mitochondrial Ca2+ uniporter (MCU) and Na+/Ca2+ exchanger (NCLX).
Similarly, the ER’s Ca2+ homeostasis is primarily governed by sarco/endoplasmic reticulum Ca2+ ATPases
(SERCA) and ryanodine receptors (RyR)/inositol-1,4,5-trisphosphate receptors (IP3R). Ca2+ efflux from the
RIS is facilitated by plasma membrane Ca2+-ATPases (PMCAs), while Ca2+ entry is mediated by voltagegated Ca2+ channels (VGCCs). Effective regulation of Ca2+ flux within organelles and across the plasma
membrane is essential for maintaining a critical balance of Ca2+ levels necessary for normal function. (3) At
the synapse (SN), Ca2+ entry through voltage-gated Ca2+ channels (Cav1.4) is essential for continual
release of glutamate-containing synaptic vesicles (SVs) from the synaptic ribbon. Light-induced
hyperpolarization leads to the closure of the Cav1.4 channels, leading to a graded decrease of glutamate
release to bipolar cells. ROS, rod outer segment; RIS, rod inner segment; SN, synaptic terminal; PDE6,
phosphodiesterase 6. Figure was created using BioRender.com.
1. NCKX1
ROS
2.
RIS
3.
SN
CNG
channel
Cav1.4
Ca2+
Ca2+
Ca2+
Na+
Glutamate
4 Na+
K+
Ca2+
Ca2+
Ca2+
VGCC
Synaptic ribbon
SVs
SERCA
Ca2+
Ca2+
RyR/IP3R
Ca2+
Mitochondria Endoplasmic
reticulum
MCU PMCA
Ca2+
Na+
NCLX
Ca2+/cGMP
[high]
Ca2+/cGMP
[low]
Dark
Light
Channels closed Channels open
PDE6
cGMP
GMP
Visual cascade
NCKX1
Ca2+ out
GTP
cGMP
GC/GCAPs
cations
in
CNG
Phototransduction
feedback
loop
Intracellular
Ca2+ stores
Signal
transmission

20
Calcium ions play an essential role in modulating sensory signaling cascade in the ROS,
particularly in signal termination and visual adaptation.135,138 As previously discussed, in dark
conditions, rods remain depolarized due to the influx of Na+
, Ca2+, and Mg2+ through CNG
channels (Figure 1.8.1). This influx is counterbalanced by the extrusion of Ca2+ by NCKX1,
maintaining Ca2+ levels in the ROS at approximately 300 nM in mice.139 Upon light exposure, the
closing of CNG channels and ongoing Ca2+ extrusion lowers cytoplasmic Ca2+ to approximately
30 nM.140,141 This reduction in Ca2+ triggers a Ca2+-dependent feedback loop involving Ca2+-
binding proteins GCAPs and recoverin (described in section 1.5:Phototransduction signaling in
rods), which is essential for swift termination of the photoresponse (Figure 1.8.1).94,95,135,142 Ca2+
therefore appears to function as a coordinator, orchestrating the activities of various signaling
pathways within the ROS.
The control of Ca2+ within the RIS involves detailed coordination between plasma
membrane channels/exchangers and internal cell organelles (Figure 1.8.2).143 In darkness,
constant depolarization activates voltage-gated Ca2+ channels (VGCC),
134,144 which, along with
the counteracting role of plasma membrane Ca2+-ATPases (PMCAs),145 leads to elevated Ca2+
levels within the vertebrate RIS (~600 nM).141 With a significant capacity for Ca2+ storage,146
mitochondria serve as critical modulators of Ca2+ in the RIS by employing specialized exchangers
and uniporters134,143 (Figure 1.8.2). This is vital for enhancing mitochondrial metabolism and ATP
production that powers various energy-dependent processes within the rod.147 Similarly, the ER
functions as an essential Ca2+ store,
143 which enhances the ER’s capability for protein synthesis
and proper protein folding. Through specific ATP-dependent channels and receptors, the ER
adjusts intracellular Ca2+ levels, maintaining a balance both in dim and bright light conditions
(Figure 1.8.2).143 Specifically, the release or depletion of Ca2+ in the ER significantly influences
RIS Ca2+ homeostasis in a light-sensitive manner. In darkness, the ER releases Ca2+ to support
photoreceptor signaling, and during extended periods of light exposure when Ca2+ levels drop, it

21
triggers the opening of additional Ca2+ channels in the RIS plasma membrane to ensure normal
cellular function.148
Rods are equipped with a distinctive synaptic terminal that facilitates the continuous
release of glutamate in the dark (Figure 1.8.3).
103,149 This is achieved by anchoring a substantial
number of synaptic vesicles to a specialized structure, known as a ribbon, located close to the
synaptic active zone.149,150 Positioned close to the ribbon, Cav1.4 channels ensure a steady flow
of Ca2+, elevating synaptic Ca2+ levels to about 2.2 µM in darkness.140 This robust Ca2+ influx
supports a rapid exocytosis rate, with approximately 400 vesicles released per second at each
rod synapse.
18,151,152 Light exposure, however, leads to photoreceptor hyperpolarization and
transient closure of Cav1.4 channels, slowing the exocytosis rate and thereby modulating
glutamate release, which in turn conveys light signals to the inner retina.103,134
1.8 Inherited retinal degenerations
Many retinal diseases, including retinitis pigmentosa (RP) and congenital stationary night
blindness (CSNB), lead to vision loss, and despite arising from diverse mutations in various
elements of the phototransduction cascade, these conditions uniformly converge on a singular
terminal event: photoreceptor cell death.
153-155 Furthermore, degeneration of rod photoreceptors
has been associated with disrupted Ca2+ homeostasis.134,156-158 Yet, definitive evidence clarifying
how Ca2+ dysregulation initiates photoreceptor cell death in RP and CSNB is still lacking. This
gap in knowledge is largely attributable to difficulties in simultaneously measuring intracellular
Ca2+ concentrations across all compartments of rod cells under normal and diseased states,135 a
subject further explored in Chapter 4. Such measurements are crucial for determining whether,
and specifically where within the rod cell, Ca2+ dysregulation plays a critical role in the apoptotic
mechanisms affecting rods. Nevertheless, elevated intracellular Ca2+ levels in certain genetic
retinal disorders have been implicated in activating calpains and caspases ,159 calcium-dependent
cysteine proteases. This Ca2+-related activation of cysteine proteases initiates a series of events

22
leading to DNA and protein fragmentation, along with the condensation of chromatin and the
cytoskeleton, ultimately resulting in the formation of apoptotic bodies.
Retinitis pigmentosa (RP), affecting ~15 million individuals globally, is among the most
prevalent retinal disorders.160 This group of inherited diseases is marked by progressive
degeneration of the retina, initially affecting rods before eventually leading to the deterioration of
cones. The initial symptom of RP is typically night blindness, owing to compromised rod function.
As the disease progresses, patients experience a narrowing visual field (“tunnel vision”), and
ultimately a loss of central vision, leading to total blindness. RP is genetically heterogeneous, with
over 50 genes identified to date.161 These genes are predominantly expressed in rod
photoreceptors, although a smaller proportion are specific to other retinal cell types, such as the
RPE.162 Autosomal recessive RP can result from mutations in genes that encode the subunits of
heterotetrameric CNG channels in the ROS, which consist of three CNGA1 subunits and one
CNGB1 subunit.163 Mutations in CNGB1 are responsible for around 4% of autosomal recessive
RP cases.164 CNGB1 knockout mice exhibit a phenotype mirroring the key pathologies observed
in RP patients, including a loss of rod photoreceptor function.
165 The absence of the CNGB1
subunit disrupts the formation of functional CNG channels in the ROS, leading to undeveloped
ROS and a gradual rod cell death over 4-6 months.165-167 The loss of CNGB1 in murine ROS is
additionally marked by a notable rise in cGMP levels,167 possibly attributed to reduced Ca2+
concentrations within the ROS, though this necessitates further validation. Moreover, protein
kinase G has been implicated in the cGMP-induced cell death pathway associated with CNG
channel loss-of-function mutations.167 Nevertheless, the potential involvement of other
mechanisms in ROS dysfunction in RP remains to be elucidated, a subject that Chapter 2 explores
further.
Congenital stationary night blindness (CSNB) represents a clinically and genetically
diverse collection of non-progressive retinal disorders, chiefly characterized by diminished night
vision and reduced visual acuity.168 Diagnosis of CSNB primarily hinges on the sustained integrity

23
of photopic vision, albeit with gradual rod photoreceptor degeneration. Various mutations
implicated in CSNB influence proteins integral to synaptic communication between rods and rod
bipolar cells, such as Cav1.4 channels,169 as well as those associated with phototransduction
processes, such as rhodopsin170 and transducin.171 Recent discoveries have also established a
linkage between CSNB and mutations in the ROS exchanger, NCKX1.
172 Experimental evidence
from a NCKX1-deficient mouse model has demonstrated that, consistent with the stationary
nature of CSNB, rods undergo gradual degeneration starting at 6 months of age, with advanced
cell death observed by 12 months.173 Remarkably, rods lacking NCKX1 retain a degree of
functionality, though they exhibit a delayed response recovery.173 Thus, these rods continue to
play a role in dim light vision by employing an alternative Ca2+ extrusion mechanism that does not
depend on NCKX1. It is probable that a compensatory mechanism, which may involve the
transference of Ca2+ into the adjoining RIS and its organelles, contributes to the regulation of Ca2+
concentration. This could help decelerate rod degeneration and safeguard cone-mediated vision.
However, the management of Ca2+ flux throughout the progression of the disorder remains
unclear. Consequently, NCKX1 knockout rods present a valuable model for investigating
abnormal Ca2+ homeostasis and dynamics within and across subcellular compartments, a topic
elaborated upon in Chapter 4.

24
CHAPTER 2
Light-regulation of Rhodopsin Distribution during Outer Segment
Renewal in Murine Rod Photoreceptors
Adapted from Current Biology (2024) manuscript. See Appendix A.
2.1 Summary
Vision under dim light relies on primary cilia elaborated by rod photoreceptors in the retina.
This specialized sensory structure, called the rod outer segment (ROS), comprises of hundreds
of stacked, membranous discs containing the light-sensitive protein rhodopsin, and incorporation
of new discs to the ROS is essential for maintaining rod’s health and function. ROS renewal
appears to be primarily regulated by extrinsic factors (light); however, results vary depending on
different model organisms. We generated two independent transgenic mouse lines where
rhodopsin’s fate is tracked by a fluorescently labeled rhodopsin fusion protein (Rho-Timer) and
show that rhodopsin incorporation into nascent ROS discs appears to be regulated by both
external lighting cues and autonomous retinal clocks. Live-cell imaging of ROS isolated from mice
exposed to six unique lighting conditions demonstrates that ROS formation occurs in a periodic
manner in cyclic light, constant darkness, and artificial light/dark cycles. This alternating
bright/weak banding of Rho-Timer along the length of the ROS relates to inhomogeneities in
rhodopsin density and potential points of structural weakness. In addition, we reveal that
prolonged dim ambient light exposure impacts not only the rhodopsin content of new discs, but
also that of older discs, suggesting a dynamic interchange of material between new and old discs.
Furthermore, we show that rhodopsin incorporation into the ROS is greatly altered in two

25
autosomal recessive retinitis pigmentosa mouse models, potentially contributing to the
pathogenesis. Our findings provide insights into how extrinsic (light) and intrinsic (retinal clocks,
genetic mutation) factors dynamically regulate mammalian ROS renewal.
2.2 Introduction
Primary cilia of vertebrate photoreceptor cells are specialized for photon capture and
house a unique composition of proteins and receptors responsible for initiating the biochemical
cascade (phototransduction) that constitutes the first steps in vision. Rod photoreceptors, which
are responsible for dim light vision, elaborate a long (~20-30 µm in rodents) and thin (~1.5 µm
diameter) ciliary structure called the rod outer segment (ROS) containing ~800 membranous
discs.12,174 The light-sensitive pigment rhodopsin constitutes more than 90% of the total
membrane protein in these discs,175 with a spatial density ~24,000 molecules/µm2
.
12,176 The
stacked disc layout within the ROS increases the probability of photon capture by rhodopsin. In
addition to phototransduction, rhodopsin is also an essential structural component, as evidenced
by rhodopsin knockout mice failing to form ROS and experiencing rod degeneration by 3
months.
53,54 The ROS undergoes daily synchronized renewal,
31 with new discs densely packed
with rhodopsin added at the base,
37,177 and ROS tips phagocytosed by neighboring retinal
pigmented epithelial (RPE) cells (Figure 2.1A).
32 Consequently, the length of mammalian ROS is
constantly maintained and completely renewed every 10-12 days.
30,178 Given the critical
relationship between rhodopsin incorporation into the ROS and its maintenance in
phototransduction and rod health, the coordinated process of ROS renewal has been an area of
active investigation.
The process of ROS renewal seen in many vertebrates appears to be 1) regulated by
light,
44,179 and/or 2) under circadian control.
47,180 The periodic disc synthesis and renewal of the
ROS are hypothesized to lead to inhomogeneities along its length. Indeed, when amphibians and
mammals are raised in cyclic light, the ROS exhibits a periodic banding pattern, which can be

26
visualized with Nomarski optics.
181,182 Interestingly, switching lighting conditions from cyclic to
constant light or darkness seems to correlate with the abolishment of visually distinct banding
patterns.
183 However, Nomarski optics reveals differences in refractive index, and the underlying
cause of this pattern remain unknown.
More recently, banding patterns in ROS have been observed in transgenic animals
expressing fluorescently tagged rhodopsin and arrestin.
184,185 In an albino rat model, Hsu et al.185
found that rhodopsin-GFP protein enters the ROS during the night in cyclic light-reared animals.
A similar observation of higher incorporation of rhodopsin-GFP during darkness was reported for
Xenopus ROS.
184 Based on this result, it was posited that cyclic light/dark cycles trigger a
mechanism that allows discs to have distinctive rhodopsin compositions, thus producing the
periodic banding pattern.
185 However, the effect of light on such banding patterns appears to vary
among model systems. For example, unlike in studies of Xenopus ROS, rhodopsin-GFP
incorporation into rat ROS was effectively halted at the onset of light.
Along with the ongoing lack of clarity surrounding ROS renewal in healthy retinas, how
rhodopsin is incorporated into the ROS during photoreceptor degeneration remains largely
unexplored. Disruption of the ROS’s structural integrity and the impaired localization of rhodopsin
are well-documented characteristics associated with several inherited forms of human blinding
diseases, including retinitis pigmentosa (RP) and congenital stationary night blindness (CSNB).
186
For example, cGMP-gated (CNG) channels and Na+
/Ca2+, K+ exchanger 1 (NCKX1) are localized
to the plasma membrane of ROS to tightly regulate the entry and extrusion of Ca2+ within the
ROS, and mutations in the genes encoding CNG channels and NCKX1 lead to diminished visual
function and progressive degeneration.
135,173 It is known that mutations in these genes lead to
disrupted ROS, misaligned discs, and rhodopsin mislocalization. Whether altered rhodopsin
incorporation into ROS may contribute to ROS disruption and retinal degeneration is not known.
To bridge these knowledge gaps surrounding endogenous rhodopsin localization during
ROS maintenance, we generated two independent transgenic mouse lines, Rhodopsin-Timer

27
(Rho-TimerA, Rho-TimerB), which express low levels of rhodopsin fused to Timer, a fluorescent
protein.187 This system allows us to visualize ROS renewal under different lighting regimens and
genetic backgrounds. We present the following findings: 1) in contrast to previous observations,
rhodopsin incorporation into the ROS is not halted by light exposure; 2) periodic banding occurs
in dark reared, light-naïve animals, pointing to intrinsic retinal clocks as regulators; 3) constant
light exposure affects nascent discs as well as older distal discs formed in darkness; 4)
inhomogeneity along the ROS reflects structural weak points; and 5) the periodic nature of
rhodopsin incorporation becomes greatly compromised in rods afflicted with Cngb1 and Nckx1
mutations prior to overt retinal degeneration.
2.3 Methods
Ethics statement
All experiments and procedures that involved live animals were reviewed and approved by the
University of Southern California Institutional Animal Care and Use Committee.
Animals
All mice were handled in strict accordance with NIH recommendations provided in the Guide for
the Care and Use of Laboratory Animals. All experiments were conducted using non-breeder
young adult mice (age 4-8 weeks), and each sex contributed to roughly half of the total animal
number in each experiment. Transgenic positive Rhodopsin-Timer (Rho-Timer) mice were either
in the C57BL/6 and Balb/c (JAX #001026) background, and wildtype littermates were used as
controls. Rho-Timer mice in the BALB/c background were bred either into the heterozygous or
knockout background of either the Cngb1 or Nckx1 genes. Cngb1 knockout mice (Cngb1-/-
)
167,188
and Nckx1 knockout mice (Nckx1-/-
)
173 were generated previously. All genotyping was performed

28
by Transnetyx, Inc. Genotyping information is available upon request. All mice were euthanized
by isoflurane inhalation, followed by cervical dislocation in the light immediately before imaging
unless otherwise stated.
Housing and light exposure conditions
Mice were housed in a 12h/12h light/dark cycle (6:00 am to 6:00 pm, except for those otherwise
mentioned), in transparent cages and had unrestricted access to pellet food and water. To
minimize light-induced photoreceptor damage, Rho-Timer mice in the albino BALB/c background
were housed on the bottom shelf of the vivarium cage rack (illuminance within cages during the
light cycle was ~6-lux). These mice lack pigmentation of the iris which facilitates more uniform
light reaching the retina. Mice housed in constant light, constant dark, or artificial light/dark
conditions were born and weaned in cyclic lighting and then were transferred to the respective
lighting conditions in temperature- and humidity-controlled rooms in the vivarium after four weeks
of age. Mice housed in constant light were exposed for 20 days to 100-lux light intensity. Mice
that were switched to 20 days of constant dark were placed in a light-tight darkroom. Mice reared
in complete darkness were born and housed in a light-tight room from birth until 4-8 weeks of age.
Intermittent cage changes for these mice were performed under infrared light with the use of
infrared goggles (BNVD Night Vision Binocular, Night Vision Depot, USA). During the artificial
light/dark cycling schedule, illumination of 100-lux followed by a period of total darkness was
controlled by a programmable timer.
Monomeric Timer construct
The first generations of Timer showed a tendency to form tetramers.
189 We introduced mutations
into the Timer construct known to minimize this tendency. The construct was generated by piecing
together two DNA fragments: Dsred2 in pDsred2-C1 (BD science Clontech) using AgeI and StuI
and pSCA1 (ppANF-Timer; a kind gift of Dr. R. Chow, University of Southern California) using

29
NotI and StuI. Dsred2 includes the V105A mutation of Timer, enhancing its fluorescence intensity,
alongside three additional amino acid substitutions (R2A, K5E, and K9T) which are expected to
minimize protein aggregation. pSCA1 includes the S197T mutation of Timer, which stabilizes the
green monomer.
187,189 These DNA fragments were ligated together into the pEGFP-N1 vector
using AgeI and NotI. To verify Timer expression, the construct was transfected into HEK-293T
cells using a Ca3(PO4)2-mediated transfection kit (InvitrogenTM), and the transfected cells were
visualized using Total Internal Reflection Fluorescence (TIRF) microscopy (courtesy of Dr. R.
Chow).
Generation of Rhodopsin-Timer (Rho-Timer) mouse
Mouse rhodopsin cDNA coding sequence and Timer sequence were obtained and amplified by
RT-PCR. PCR linked Timer to the C-terminus of rhodopsin with primers Sal I-Rho
(5’CGCGTCGACATGAACGGCACAGAGGG3’) and Timer1-1D4-Cla I
(5’CCATCGATCTAGGCTGGAGCCACCTGGCTGGTCTCCAGGAACAGG-TGGTGGC3’),
creating Rhodopsin-Timer (Rho-Timer). Rho-Timer was ligated into a pBluescriptKSII+ vector,
containing a 4 kb mouse rhodopsin promoter at 5’ (Kpn I and Xho I sites) and a 0.6 kb mp1
fragment for polyadenylation at 3’ (BamH I site with direction selection),
190 using Sal I and Cla I
sites. pRho-Timer was sequenced, and the plasmid was subsequently purified by CsCl2 and
digested with Kpn I and Not I to release the 6.6 kb insert fragment. The fragment was purified by
QIAquick gel extraction kit (Qiagen) and Elutip-D column (Whatman) and was microinjected into
F1 hybrid zygotes from C57BL/6J and DBA/2J strains following standard procedures. Rho-Timer
founder mice were identified by PCR screening using primers Timer-F
(5’GAGTTCATGCGCTTCAAGGT3’) and Timer-R (5’TGGTCTTCTTCTGCATCACG3’). Two
founders produced offspring that expressed Rho-Timer in rod photoreceptors and were
subsequently crossed with C57BL/6 mice. One founder line was kept in the C57BL/6 mice
background (Rho-TimerA) while the other was bred into the BALB/c background (Rho-TimerB).

30
Preparation and live-cell imaging of ex-vivo retinas from Rho-Timer positive mice
Eyes were harvested at 11:00h in their respective lighting conditions, the superior pole of each
eye was cauterized, the eyes were enucleated, and the retinas were isolated in Ames’-
bicarbonate buffer (1.9 g/L NaHCO3, 1 bottle/L Ames’ Medium (Sigma), pH 7.4, 280 mOsm)
equilibrated with 95% O2/5% CO2. Retinal whole-mounts were flattened and immobilized on a
piece of nitrocellulose membrane (0.2 mm, BIO-RAD) with the photoreceptor cells facing up.
191 A
small cut (1 mm) was made at the 0˚ (superior pole) for orientation. The retinal whole-mounts
were positioned on a glass slide and were inspected with a 20X/0.5 NA objective using a Zeiss
Axioplan2 microscope. Retinal vertical sections were acquired using a tissue slicer (Stoelting).
Briefly, isolated retinas were hemisected and embedded in 3% low melting agarose (Invitrogen).
Agarose-embedded retinas were incubated in cold Ames’-bicarbonate buffer and were
subsequently cut into 250 µm-thick slices. Live tissue sections were transferred to a glass-bottom
dish (NuncTM, Thermofisher Scientific), stabilized with a handmade U-shaped anchor, and imaged
for 20 min in carbogen-saturated Ames’-bicarbonate buffer using a confocal microscope (Leica
SP8 DIVE multiphoton system, 63X/1.3 NA oil objective, Chameleon Discovery laser (920 nm,
Coherent), and Leica 4Tune spectral hybrid detectors (green emission 505-535 nm and red
emission 565-590 nm)). Dissociated rods were obtained from the superior hemisphere of the
retina. The hemisected retina was incubated in a Delrin enclosure192 containing carbogensaturated Ames’-bicarbonate buffer plus 0.8% (vol/vol) rhodamine-labeled wheat germ agglutinin
(Vector Laboratories, Inc.), 0.1% (vol/vol) BSA, and 1mM MgCl2 for 30 min to label rod OS. Rods
were mechanically dissociated from the retina with a feather scalpel and successively flattened
with moist dialysis tubing (SnakeSkinâ
, PIERCE) and a 1.8 g metal cylinder. Rods were bathed in
carbogen-saturated Ames’-bicarbonate buffer and then imaged for 10 min. Care was taken to
image clusters of dissociated rods. Dissociated rods were visualized using a confocal microscope
(Leica SP8X with DMi8 microscope platform and HC PL APO CS2 63X/1.4 NA oil objective) and
hybrid detectors (GaAsP enhanced Avalanche Photodiodes). Illumination was provided by a

31
supercontinuum white light laser, and two-color imaging of Rho-Timer rods was carried out
sequentially (488 nm and 551 nm). To compensate for inverted dish reflection from 488 nm
excitation, LightGate (1.51-7.15 ns) was on throughout imaging. All procedures were performed
at room temperature unless otherwise noted.
Quantitative real-time PCR (qRT-PCR)
Animals were sacrificed at three-hour intervals over a 24h period: zeitgeber time (ZT) or circadian
time (CT) 0, 3, 6, 9, 12, 15, 18, 21, where ZT 12 and CT 12 equals to the time of light off at 18:00h
under cyclic light. The retinal extraction procedure was either carried out in dim red light or room
light (500-lux) depending on the lighting environment at the time of extraction. Isolated retinas
were immediately subjected to total mRNA extraction (BioRad, #7326820, Hercules, CA) before
quantification on a Nanodrop spectrophotometer (ND-1000, Thermoscientific). Total RNA was
reverse transcribed with a cDNA synthesis kit (NEB, #E6300, IPswitch, MA). The primer
sequences (designed previously 193 or by using Primer3) and reference sequence numbers for
the genes used in this study are listed in Table 2.1. qRT-PCR (~15 ng of cDNA, 1 mM of each
primer, SYBR Green Master Mix (Bio-Rad, #1725270, Hercules, CA)) was performed using the
Roche LightCycler 96 (Roche, Basel, Switzerland). Cycling parameters were denaturation at 95˚C
for 5 min, then 45 cycles of 94˚C for 60 s, 60˚C for 60 s, and 72˚C for 60 s. Upon completion, a
final extension was carried out at 72˚C for 8 min, which was followed by a product melting curve
(95˚C for 10 s, 65˚C for 60 s, 97˚C for 1 s). Reactions were run in triplicate, and the reference
gene �-actin was used as an internal control to normalize expression variability. The R package
“chipPCR”194 was used to analyze raw data, and normalized DCT values from each time point in
a specific lighting condition were then subtracted from its respective T = 0 (6:00h) DCT to
determine the relative changes in mRNA expression (2-DDCT method) across different lighting
conditions.
195

32
Table 2.1 Primer sequences used for quantitative real-time PCR. Primers were designed by Primer3
(Actb, Rho, Timer) on the specified RefSeq sequences and Arntl primers are as reported previously.193
Each primer forward/reverse pair yielded a single product. F = forward primer; R = reverse primer.
Electroretinogram (ERG)
2-month-old Rho-Timer mice and their nontransgenic littermates were dark-adapted overnight,
and ERG recordings were performed under dim red light. The mice were anesthetized by
intraperitoneal injection of a xylazine (10 μg/g bodyweight) and ketamine (100 μg/g bodyweight)
solution diluted in 1X PBS. 0.5% tropicamide and 2.5% phenylephrine hydrochloride solutions
were added to the right eye for pupil dilation. Hydroxylpropyl methycellulose solution was added
to both eyes to prevent eye dryness and to create an electrical contact between the corneal
electrode and the right eye. Insertion of a subcutaneous reference electrode was placed
proximate to the eye. Flashes from dim to bright light intensities (ranging from 0.1 mcd to 25 cd)
were delivered using The OcuScience HMsERG LAB System. The ERGs were acquired and
amplified by an AC/DC differential amplifier and subsequently filtered for analysis. Scotopic awave and b-wave amplitudes were obtained using OcuScience ERG Viewer software.
Retinal morphology
The superior pole of each eye was marked by cauterization. Enucleated eyes were fixed, the
cornea and lens were removed, and the eyecups were embedded in an epoxy resin as previously
described.
56 Briefly, eyes were placed into ½ Karnovsky buffer (0.1 M cacodylate buffer containing
2.5% glutaraldehyde and 2% formaldehyde, pH 7.2) for 5 min. The eyes were subsequently
Name RefSeq Size(bp) Sequence
actin, beta
(Actb)
NM_007393.5 146 F: GATTACTGCTCTGGCTCCTAGC
R: CGTTGTGGGAGGTGATGTCCAGCT
rhodopsin
(Rho)
NM_145383.2 89 F: TGTGGTCTTCACCTGGATCAT
R: GAACATTGCATGCCCTCAG
Timer Terskikh et al., 2000187 435 F: CTGTCCCCCCAGTTCCAGT
R: CGTTGTGGGAGGTGATGTCCAGCT
Bmal1
(Arntl)
NM_007489.4 127 F: TGGAACCCTAGGCCTTCATT
R: TTCGATCCAGTGTGGGAGAT

33
dissected, and the cornea and lens were removed. The eyecups were further fixed in ½ Karnovsky
buffer at 4˚C overnight. Fixed eyecups were rinsed in 0.1 M cacodylate buffer, fixed in 1% OsO4
for 1h, dehydrated in graded ethanol, and embedded in epoxy resin. The epon-embedded eyes
were sectioned at or near the vertical meridian into 1 µm sections using an ultramicrotome (Leica
Ultracut UCT, Leica Microsystems, Bannockburn, USA) for light microscopy. Sections were
stained, and images of the retinal morphology were acquired on a Zeiss Axioplan2 microscope
using a 63X/1.4 oil objective.
Western blot
Dark-adapted retinas were homogenized in 150 µl RIPA lysis buffer (50 mM Tris pH 8, 0.1% SDS,
150 mM NaCl, 0.5% NP40, 0.5% deoxycholate acid) containing 0.1 mM phenylmethanesulfonyl
fluoride, 0.02 mM Aprotinin, and 0.02 mM Leupeptin. DNase I (Roche) was added, and samples
were incubated at room temperature for 30 min. PierceTM BCA Protein Assay Kit (#23227)
determined total protein amount. An equal amount of protein from each sample was loaded onto
a 4-12% Bis-Tris SDS-PAGE gel (Invitrogen) and was then transferred to a nitrocellulose
membrane (Cytiva AmershamTM ProtranTM, pore size 0.2 µm, 45004011). Membranes were
incubated overnight at 4˚C with the following antibodies: mouse monoclonal anti-rhodopsin (1D4,
1:20,000, generously provided by Dr. R. Molday, University of British Columbia), mouse
monoclonal anti-rhodopsin (R2-12N, 1:10,000, a kind gift from Dr. P.A. Hargrave, University of
Florida), and rabbit polyclonal anti-Living ColorsⓇ DsRed antibody (DsRed, 1:1,000, #632496,
Takara Bio USA, Inc.). The membranes were incubated with peroxidase labeled horse anti-mouse
IgG antibody (H+L) (1:5,000, Vector, PI-2000-1) or fluorescently labeled secondary antibodies
(1:10,000, 926-31081, LI-COR) for 1h at room temperature. Blots were either treated with the
AmershamTM ECL Western blotting detection reagents (RPN2209) and imaged with the G:BOX

34
Chemi XX6 digital imaging system (Syngene) or were detected by the Odyssey CLx system (LICOR).
Immunocytochemistry
Retinal eyecups were prepared as described in the Retinal morphology section. For frozen
sections, the tissues were fixed in 4% formaldehyde in 1X PBS for 1h, rinsed with 1X PBS, and
placed into 30% sucrose in 0.1 M cacodylate buffer overnight at 4˚C. The eyecups were
embedded in Tissue-Tek O.C.T. Compound (Sakura), frozen in a dry ice/ethanol slurry, and
cryosectioned at 10 µm. The slices were blocked for 1h (5% donkey serum, 0.3% Triton X-100 in
1X PBS) and incubated overnight at 4˚C with the mouse monoclonal primary antibodies that
recognize either the N- or C-terminus of rhodopsin (4D2196 (1:250) and 1D4 (1:10,000),
respectively). The frozen retinal sections were incubated in Alexa Fluor 647-labeled secondary
antibody (1:200, Thermo Fisher Scientific). For retinal flat mounts, eyes were enucleated in room
light (500-lux) and directly fixed in 4% formaldehyde in 1X PBS. Once the cornea and lens were
removed, the remaining eyecups were fixed for an additional 2h. The RPE was then carefully
removed. Four relief cuts were placed at the edge of the retina, which was flattened and affixed
to a nitrocellulose membrane (photoreceptors pointing up). The whole-mounts were blocked
overnight at 4˚C (5% donkey serum, 0.3% Triton X-100 in 1X PBS) and incubated with either of
the following mouse monoclonal primary antibodies for 5 days at 4˚C: A11-82P (1:10, gift from
Dr. P.A. Hargrave, University of Florida) and G-8 (1:50, sc-8004, Santa Cruz Biotechnology). The
whole-mounts were then incubated with the Alexa Fluor 594-labeled secondary antibody (1:400;
Thermo Fisher Scientific). Wholemounts were embedded in 3% low melting agarose (Invitrogen)
and 250 µm-thick sections were generated using a tissue slicer (Stoelting). Retinal specimens
were viewed under a 40X/1.3 NA oil objective, and images were acquired on a Zeiss LSM800
confocal microscope (Carl Zeiss Meditec, Jena, Germany).

35
Rhodopsin concentration
The concentration of rhodopsin was measured by absorption spectroscopy (optical density vs
wavelength). Mice were dark-adapted overnight, and the retinas were isolated under infrared light.
Individual retinae were solubilized in 200 µL of 1X PBS containing 1% n-dodecyl-b-Dmaltopyranoside (850520P, Avanti Polar Lipids), 0.1 M PMSF and complete mini protease
inhibitor (Roche Applied Sciences), light protected with aluminum foil, and rocked overnight at
4˚C. Following solubilization, samples were centrifuged at 4,000 rpm for 3 min to remove
particulates. 100 µL of the supernatant from each sample was individually transferred to a quartz
cuvette to scan the UV/visible (270-700 nm) absorption spectra (DU640 Beckman Coulter).
Measurements of the absorption spectra were made before and after a 5 min bleach. Rhodopsin
concentrations were then calculated from Beer’s Law using (1) the difference in absorbance at
500 nm before and after full bleach by white light (5 min) and (2) its extinction coefficient (40,600
M-1
cm-1
). This value was normalized to the estimate of the total protein concentration at 280 nm,
and the rhodopsin concentration was subsequently normalized to average ROS length for each
condition. This analysis relied on the assumption that combining both rhodopsin and Timer as a
fusion protein does not alter their spectral properties. Based on the indistinguishable absorbance
spectra and previous findings regarding rhodopsin fusion proteins, this assumption was
considered reasonable.
184,197
Image analysis
Fluorescence images of Rho-Timer rod OS were analyzed using a custom ImageJ macro.
198 Line
profiles were manually drawn on each image, and the resulting profile data was saved. A custom
MATLAB code extracted the frequency information using fast Fourier transform (FFT) (MATLAB,
2023a). FFT spectra were interpolated to account for different image pixel size between samples,
normalized, and detrended to account for background. To find whether the banding pattern was
periodic, averaged FFT plots were transformed into periodograms and underwent the following

36
data analysis.
199 High-pass filtering (cut-off frequency 0.1 µm-1
) and normalization (so that each
trace intensity is between 0 to 1) were performed for each rod profile. A periodogram was then
extracted for each rod. Periodograms for each condition were averaged, and for each averaged
periodogram, a peak period (T) was found. For rod OS curvature analysis, a randomly selected
population of rods underwent image processing. ROS curvature from dissociated rods was
assessed using FIJI’s200 plug-in Kappa – Curvature analysis (version 2.0.0).201 For each
fluorescence image of Rho-Timer rods, a B-Spline curve was carefully drawn along the length of
the ROS starting from the base to the tip. The curvatures along the ROS length were extracted
and the average curvature/µm was plotted.
Statistical analysis
The number of mice and rods per animal (N) is given in the corresponding figure legends. Unless
otherwise mentioned, GraphPad Prism 10.0.0 for Mac and SPSS for Mac, version 29 (IBM Corp.,
Armonk, N.Y., USA) were used to perform statistical analysis and generate plots. Gaussian
distribution was evaluated using the Shapiro-Wilk test, and homogeneity of variance was
evaluated with Levene’s test. For normally distributed data, statistical analysis was performed
using an unpaired t-test (pairwise comparisons) or a one-way ANOVA followed by a post hoc
Tukey test (multiple comparisons). If samples deviated from normality, statistical analysis was
performed using the Kruskal-Wallis test with Dunn-Bonferroni’s correction for multiple comparison
testing. Effect size (Cliff’s delta, ‘effsize’ package202) was calculated in R (RDevelopment Core
Team, 2022). Gene expression rhythmicity was analyzed using Circacompare203 (‘remotes’
package204) and the cosinor method by fitting the 24h data to a cosine curve. Pairwise correlations
were computed using Pearson’s correlation coefficient between two groups. Fisher’s g-statistic
was calculated for each condition’s Fourier transform spectra to produce an associated p-value
(MATLAB). Statistical significance was set at � = 0.05 for all tests. Tests with p ≤ 0.05 were
deemed statistically significant.

37
2.4 Results
Generation of Rhodopsin-Timer as a reporter of rhodopsin distribution in mammalian ROS
We created two independent transgenic mouse lines (Rho-TimerA and Rho-TimerB)
expressing a fluorescent Rhodopsin-Timer fusion protein to investigate how lighting history and
two specific degenerative mutations affect rhodopsin incorporation into ROS in intact rods. To
ensure the fusion protein is properly delivered to the ROS, the last 8 amino acids of the carboxyterminal of rhodopsin (1D4 epitope) containing the trafficking motif56,205 were repeated after the
coding sequence for Timer197 (Figures 2.1B-C). The 4.4-kb 5’ region of the mouse rhodopsin
promoter
190 was used to drive transgene expression in rods (Figure 2.1C). As seen previously,
this promoter produced a pattern of expression in both founder lines where Rhodopsin-Timer
(Rho-Timer) fluorescence intensity was more evenly distributed in the dorsal-temporal
hemispheres and more sparse in the ventral-nasal hemispheres (Figure 2.1D).190 Orthogonal
sections through these retinas show Rho-Timer fluorescence in the ROS layer, exhibiting a
banding pattern along each ROS, as well as in the inner segment and outer nuclear layer (ONL)
of some cells in both lines (Figure 2.1E). The strikingly similar expression patterns in the ROS of
both independent lines (Figure 2.1F) suggest that transgene expression is driven by the rhodopsin
promoter rather than position effect variegation, a phenomenon arising from transgene integration
at the boundary of euchromatin and heterochromatin which can lead to stochastic temporal and
spatial expression.
206,207 The Rho-Timer fusion protein was initially generated to aid the study of
rhodopsin turnover, as Timer (DsRed-E5 mutant) was found to possess the unique ability of timedependent shift of fluorescence emission spectra from green to red in cultured cells.
187 However,
this property was inconsistent in rod cells expressing Rho-Timer (Figure 2.1F). Thus, we used
Timer as a generic fluorescent tag, specifically imaging the GFP-like chromophore state, for the
present study.

38
Figure 2.1. Expression of Rho-Timer in rod photoreceptors. (A) Schematic of the highly
compartmentalized rod and its interplay with neighboring retinal pigmented epithelium (RPE). ROS, rod
outer segment; RIS, rod inner segment; N, nucleus; SN, synaptic terminal. (B) Schematic of the ROS, discs,
and Rho-Timer. Rho-Timer (red) resides within ROS disc membranes. The cartoon model of Rho-Timer
contains the entire coding sequence of rhodopsin, which is fused to Timer (red) and is followed by the last
8 amino acids of the C-terminus of rhodopsin (ETSQVAPA). Timer lies on the cytoplasmic side of the disc
membrane. (C) Diagram depicting the Rho-Timer transgene construct. 1D4 epitope corresponds to the Cterminus of rhodopsin (ETSQVAPA). (D) Representative Rho-Timer retinal whole-mount (N = 4). Rhodopsin
promotor produces a dorsal (D) to ventral (V) expression gradient. Scale bar = 500 μm. T, temporal; N,
nasal. (E) Vertical slice from Rho-Timer retina. Periodic banding is seen as alternating Rho-Timer rich and
deficient fluorescent regions in the ROS. Scale bar = 10 μm. ONL, outer nuclear layer. (F) Representative
overlay (merged) images of fluorescent Rho-Timer in ROS from Rho-TimerA and Rho-TimerB mice housed
in cyclic light. Wheat germ agglutinin labeled ROS plasma membrane (red), and red and green rich bands
(yellow) are co-localized. TA: Rho-TimerA; TB: Rho-TimerB. Scale bar = 2 μm. Schematic figures were
created with BioRender.com.
To mitigate concerns surrounding over-expression of rhodopsin affecting rod function and
survival,
208,209 and thereby interfering with the normal fate of endogenous rhodopsin in the ROS,
we examined the expression of the transgene, total rhodopsin concentration, and the morphology
and function of rods from Rho-Timer transgenic mice. Quantitative real-time PCR (qRT-PCR) was
used to quantify Rho-Timer transcript levels relative to that of endogenous rhodopsin. Results
from qRT-PCR revealed that Rho-Timer transcripts were substantially lower (mean = 0.84%,
range = 0.6%-1.4%) than endogenous rhodopsin (Figure 2.2A). In addition, we performed direct
Timer 1D4
Mouse
Rhodopsin
cDNA
Rhodopsin Promoter
A B
C
D
D
V
T N
ROS
RIS
ONL
E
Cytoplasmic
RPE Intradiscal
ROS
RIS
N
SN
NH2
COOH
Cytoplasmic
Intradiscal
F Base Tip
TB
TA

39
measurements of total rhodopsin pigment concentrations using spectrophotometry. The results
revealed no identifiable differences in rhodopsin content between transgenic and wildtype retinas
(Figure 2B, p = 0.90). Western blots using antibodies against either the amino- (R2-12N)210 or
carboxyl- (1D4)211 terminus of rhodopsin detected abundant endogenous rhodopsin, but not the
higher molecular weight Rho-Timer fusion protein (Figure S2.1, A and B). These results indicate
that the protein expression level of Rho-Timer was below the detection limits of these wellestablished antibodies in western blots. We also performed immunocytochemistry using
antibodies targeting both rhodopsin’s N-terminus (4D2)196 and C-terminus (1D4) to look for
evidence of Rho-Timer mislocalization at the outer nuclear layer (Figure S2.1, C and D). The
results showed no consistent correlation between Rho-Timer signal and increased antibody
reactivity, suggesting minimal contribution of Rho-Timer to rhodopsin localization in this
compartment. Next we examined retinal morphology for evidence of degeneration and performed
electroretinograms to assess retinal function and found that retinal morphology was
indistinguishable between Rho-Timer and transgene-negative littermates (Figure 2.2C), and
electroretinograms of light responses were similar between these two groups of mice (Figures
2.2D and 2.2E). Together with the broad expression pattern of the transgene in the retina (Figure
2.1D), these results confirm that Rho-Timer was expressed at very low levels per rod cell and had
no detectable effects on retinal morphology or function.

40
Figure 2.2. Rho-Timer expression does not affect retinal structure or function. (A) Quantification RhoTimer and endogenous rhodopsin transcript levels by qRT-PCR over 24h (N = 24, 3 mice per time point
indicated by different shade of gray). Values are presented as the percentage of Rho-Timer/endogenous
rhodopsin (mean ± SD). (B) Rhodopsin concentration (nmol/retina) for WT and Rho-Timer retinae did not
differ significantly (t-test, p = 0.90, N = 9-12). (C) Rho-Timer retinal morphology, ROS structure, and ONL
thickness were similar to littermate WT retinae. Scale bar = 20 μm. WT, wildtype; Rho-Timer, RhodopsinTimer; RPE, retinal pigmented epithelium; ROS, rod outer segment; RIS, rod inner segment; ONL, outer
nuclear layer; INL, inner nuclear layer. (D) Scotopic electroretinogram (ERG) recordings from dark-adapted
2-month-old WT (black) and Rho-TimerB (red) littermates housed under cyclic light (N = 3 per group).
Responses were captured from a range of flash intensities from weak (0.001 and 0.01 cd*s/m2) to strong
(0.1, 3, and 25 cd*s/m2). (E) Intensity-response curves of a- and b-wave amplitude of the same mice shown
in (D). Rho-Timer mice show no differences in a- or b-wave amplitudes or response kinetics compared to
wildtype littermates (t-test, p > 0.05).
Rho-Timer transcript mirrors that of endogenous rhodopsin under different lighting
conditions
Rhodopsin mRNA levels exhibit a rhythmic diurnal expression pattern under normal cyclic
light.
212-214 To assess Rho-Timer’s suitability as an endogenous rhodopsin reporter, its expression
pattern should track that of endogenous rhodopsin. To evaluate this, rhodopsin and Rho-Timer
mRNA levels were quantified every 3h over a 24h period using qRT-PCR. Indeed, the expression
level of rhodopsin and Rho-Timer from mice raised in cyclic light exhibited substantial fluctuations
in expression over a 24h period (Figure 2.3A, top panels). BMAL1, a master clock gene that drives
rhythmic gene expression and is known to cycle in the retina under cyclic light,
215 was carried as
a positive control. Using CircaCompare,
203 a new statistical method designed to analyze and
Rho-Timer/Rhodopsin
mRNA expression (%)
Rhodopsin Concentration
(nmol/retina)
25
3
0.1
10-2
10-3
150ms
250µV
WildtypeRho-Timer
0.2
0.3
0.4
0.5
0.6
Rhodopsin Concentraion
(nmol/retina)
Wildtype
Rho-Timer
A D
B
C
WT RhoTimer
ns
0.5
1.0
1.5
Rho-Timer/Rhodopsin
mRNA expression (%)
WT
Rho-Timer
RPE
ROS
RIS
ONL
INL
RPE
ROS
RIS
ONL
INL
cd*s/m2
WT
Rho-Timer
E
a-wave
Amplitude (µV)
b-wave
Amplitude (µV)
10-4 10-3 10-2 10-1 100 101 102
0
125
250
375
500
a-wave
10-4 10-3 10-2 10-1 100 101 102
0
125
250
375
500
a-wave 2
10-4 10-3 10-2 10-1 100 101 102
0
300
600
900
b-wave
10-4 10-3 10-2 10-1 100 101 102
0
300
600
900
b-wave 2
Intensity (log cd*s/m2)
Intensity (log cd*s/m2)
20 µm
20 µm
-4 -3 -2 -1 0 1 2
-4 -3 -2 -1 0 1 2

41
compare rhythmic data, we found that BMAL1 was expressed in a periodic fashion in cyclic light
(p = 0.005), as expected. Rhodopsin (p = 0.01) and Rho-Timer (p = 0.03) transcript levels also
fluctuated within 24h, with closely matched expression patterns relative to each other, but out of
phase with BMAL1 (Figure 2.3A). Our results indicate that there is a peak in rhodopsin transcript
levels that occurs just after light onset with subsequent dwindling during the night (Figure 2.3A),
consistent with previous reports.
212
The diurnal variation in rhodopsin and Rho-Timer transcription could either be in response
to light, or reflect an internal circadian rhythm originating from retinal clocks.
44,216-218 Therefore,
we investigated whether rhodopsin’s cycling transcription occurs in the absence of light/dark
cycles. To test this, Rho-TimerB mice were born and reared in complete darkness in a light-tight
room or were cyclic light-reared and moved to constant dark or constant dim light (100-lux) for 20
days (20dD or 20dL, respectively). We determined that 20 days was sufficient time for the mice
to adjust to a new lighting environment since one week is required for mice to re-entrain to a new
lighting environment,219,220 and ~10 days is required for ROS renewal.30 qRT-PCR revealed that
the transcriptional oscillation of both rhodopsin and Rho-Timer was preserved in 20dD (p = 0.007;
p = 0.005), dark reared (p = 0.02; p = 0.004), and 20dL (p = 0.02; p = 0.03). Under these
continuous lighting environments, rhodopsin and Rho-Timer gene expression sustained robust
periodicity and synchronicity with a free-running period of ~24h (Figures 2.3B-3D). Of note, the
phase of rhodopsin and Rho-Timer rhythmic expression in cyclic light differed considerably from
that in 20dD and dark reared (Figures 2.3A-3C), such that the transcriptional maxima now
occurred during the subjective night. Expression of BMAL1 was slightly attenuated but continued
to cycle under 20dD (p = 0.04), with loss of its rhythmic expression pattern when mice were dark
reared (p = 0.24) or transferred into 20dL (p = 0.25). Interestingly, in 20dD, BMAL1 transcripts
appeared phase-shifted relative to that of rhodopsin and Rho-Timer when compared to the cyclic
light (Figures 2.3A and 2.3B). Statistical analysis confirms a lack of correlation between BMAL1
and rhodopsin/Rho-Timer (Pearson’s correlation, 20dD: p = 0.25). Together, these data show that

42
Rho-Timer transcription mirrors rhodopsin under different lighting conditions, and therefore serves
as a good reporter for endogenous rhodopsin gene expression. Further, the persistent rhythmicity
of Rho-Timer and rhodopsin transcripts found in 20dD, dark reared, and 20dL retinae suggest
that daily visual pigment transcription is in part regulated by a distinct clock-derived mechanism.
Figure 2.3. Diurnal changes of Rho-Timer transcript mirrors that of endogenous rhodopsin. Total
retinal RNA was extracted in 3h intervals over 24h and subjected to qRT-PCR to quantify expression
profiles of rhodopsin, Rho-Timer, and BMAL1 under different lighting conditions. Each transcript was
normalized to �-actin. The lighting conditions are: (A) cyclic light, (B) cyclic light-reared moved to 20 days
dark (20dD), (C) dark reared from birth, and (D) cyclic light-reared moved to 20 days constant light (20dL,
100-lux). Statistics calculated using CircaCompare203 confirms presence of periodicity (p ≤ 0.05), which are
given in the upper left corner of each panel (N = 3 for each time point, and qRT-PCR samples were run as
triplicates). ZT, Zeitgeber time; CT, Circadian time.
0 3 6 9 12 15 18 21
0.0
0.5
1.0
1.5
CT (hr)
Relative Expression (A.U.)
BMAL1 Light
0 3 6 9 12 15 18 21
0.0
1.0
2.0
3.0
4.0
CT (hr)
Relative Expression (A.U.)
Plot a function (timer) Timer Light
0 3 6 9 12 15 18 21
0.6
0.8
1.0
1.2
1.4
CT (hr)
Relative Expression (A.U.)
Plot a function (rho) Rhodopsin Light
0 3 6 9 12 15 18 21
0.0
0.5
1.0
1.5
2.0
2.5
CT (hr)
Relative Expression (A.U.)
BMAL 1 Dark rear
0 3 6 9 12 15 18 21
0.0
2.0
4.0
6.0
CT (hr)
Relative Expression (A.U.)
Timer Dark rear
0 3 6 9 12 15 18 21
0.0
1.0
2.0
3.0
CT (hr)
Relative Expression (A.U.)
Rhodopsin Dark rear Plot a function (rho)
0 3 6 9 12 15 18 21
0.0
0.5
1.0
1.5
2.0
2.5
BMAL1 Dark
CT (hr)
Relative Expression (A.U.)
0 3 6 9 12 15 18 21
0.0
1.0
2.0
3.0
CT (hr)
Relative Expression (A.U.)
Timer Dark 0 3 6 9 12 15 18 21
0.0
1.0
2.0
3.0
ZT (hr)
Relative Expression (A.U.)
BMAL 1 Cyclic
0 3 6 9 12 15 18 21
0.0
0.5
1.0
1.5
2.0
2.5
Rhodopsin Dark
CT (hr)
Relative Expression (A.U.)
0 3 6 9 12 15 18 21
0.0
0.5
1.0
1.5
2.0
Timer Cyclic
ZT (hr)
Relative Expression (A.U.)
0 3 6 9 12 15 18 21
0.0
0.5
1.0
1.5
2.0
ZT (hr)
Relative Expression (A.U.)
Rhodopsin Rhodopsin Cyclic Rho-Timer BMAL1
p=0.01 p=0.03 p=0.005
20 days Dark
(20dD)
20 days Light
(20dL)
Cyclic Light
A
B
D
Dark Reared
C
p=0.007 p=0.005 p=0.04
p=0.02 p=0.004 p=0.24
p=0.02 p=0.03 p=0.25

43
Diurnal incorporation of Rho-Timer and endogenous rhodopsin creates discrete banding
patterns in mammalian ROS
The diurnal fluctuation of rhodopsin transcript should lead to alternating rhodopsin
densities along the ROS if 1) protein translation is directly proportional to transcript levels and 2)
the rate of membrane disc formation is relatively constant. Such heterogeneous rhodopsin
densities along the ROS are often not observed using indirect immunofluorescence, likely due to
the high concentration of rhodopsin in this compartment (8.23 mM with respect to the ROS
cytoplasm).
12 However, Hsu et al.,185 using neuroaminidase treatment and high magnification
confocal images of immunofluorescence and immunoelectron microscopy, demonstrated that
endogenous rhodopsin display ~10 peak signals along the ROS in C57BL6/J mouse retinas. To
investigate this phenomenon further, we used the A11-82P antibody,
221 which binds
phosphorylated rhodopsin, as a surrogate to track endogenous rhodopsin in the ROS. A11-82P
immunolabeling of retinal sections from light exposed, cyclic light-reared Rho-Timer negative
littermates revealed pronounced fluorescent intensity fluctuations along the length of the ROS in
some cells (Figure 2.4A). The lack of banding pattern in all cells may be due to variable access
of the antibody to its epitope or variable light exposure to each rod. To validate that the banding
pattern of phosphorylated rhodopsin is not a reflection of varying concentration of rhodopsin
kinase (GRK1), which phosphorylates activated rhodopsin, we used an antibody against GRK1
(G-8). In sharp contrast with the discrete fluctuations observed with the A11-82P staining, the G8 staining appeared uniform within the entire ROS (Figure 2.4B). These results suggest that the
banding pattern of Rho-Timer fluorescence reflects the heterogeneous distribution of endogenous
rhodopsin along the ROS. Further, the results indicate that entry of rhodopsin into the ROS occurs
with a periodic rhythmicity in cyclic light and that this pattern could potentially be attributed, at
least in part, to rhodopsin’s oscillating transcription.

44
Figure 2.4. Banding pattern of endogenous phosphorylated rhodopsin in the ROS. Images of retinal
sections from 1-month-old wildtype (transgene-negative) littermates labeled with (A) anti-A11-82P
(recognizes phosphorylated rhodopsin) and (B) anti-G-8 (detects rhodopsin kinase) antibodies (gray). The
insets show a magnified image of ROS exhibiting a distinct banding pattern. Tissues were acquired from
mice acclimated to room light. Scale bar = 5 μm; N = 5. ROS, rod outer segment; RIS, rod inner segment.
To better understand the periodicity of endogenous rhodopsin’s entry into the ROS, we
next determined the spatial period of Rho-Timer fluorescence oscillations along the ROS in cyclic
light. We plotted the fluorescence intensity profile of individual ROS (Figure 2.5A1) and computed
the respective fast Fourier transform (Figure 2.5B1). The average number of fluorescent rich
bands observed from individual intensity profiles was equal to ~10 (Figures 2.5A1 and 2.5C) along
the length of an average ROS (22.8 µm, Figure 2.5D), consistent with previous reports that
mammalian ROS turnover takes ~10 days.
30 These data indicate that Rho-Timer enters the ROS
following a diurnal rhythm, where each bright/dim fluorescent band is generated per day. By
averaging the power spectra from 60 rods, we found a prominent peak at a spatial period of 2.44
µm (Figures 2.5B1 and 2.5E). This value represents the average spatial distribution of the periodic
variation, containing one dim and one bright fluorescent band that represent two distinct disc
ROS
RIS
A
B
ROS
RIS

45
compositions of Rho-Timer. To determine if the observed banding pattern is periodic, we
computed Fisher’s g-statistic which confirmed that Rho-Timer exhibits periodicity along the ROS’s
length (p < 0.01).
Figure 2.5. Light history affects Rho-Timer periodicity. Mice born and raised in cyclic light (except for
dark reared) were subjected to different lighting conditions. 1, cyclic light (N = 18); 2, 20 days in darkness
(20dD, N = 20); 3, dark reared in light-tight room (N = 18); 4, 20 days constant light (20dL, 100-lux, N = 20);
5, 5 days constant dark followed by 5 days constant light (5dD/5dL, 100-lux, N = 15); 6, 20 days of artificial
7h/7h light/dark schedule (7hL/7hD, 100-lux, N = 8). (A) Representative images of ROS (measured in µm)
and fluorescence intensity profiles from a single ROS (base to tip) from Rho-TimerB mice. In panel 6, the
white line captures one period (one dim and bright band), and the red line emphasizes the ‘doublet’ band
(two dim and bright bands) from 7hL/7hD exposed ROS. (B) Fast Fourier transform analysis of ROS
presented in terms of average peak banding period (inverted black triangle, B1, T = 2.44 µm; B2, T = 2.52
µm; B3, T = 2.71 µm; B5, T = 2.69 µm; B6, T = 1.2 µm (short period) and T = 2.32 µm (long period, ‘doublet’
band)). Fisher’s g-statistic confirms periodicity (p < 0.01, N = 50-80 rods). 20dL did not reach statistical
significance (B4, p = 0.15), and Fisher’s g test was unable to provide a p-value for 7hL/7hD (two peak
Distance_(inches)
Gray_Value
24.79µm
24.00µm
24.60µm
23.37µm
24.54µm
25.14µm
23.58µm
21.25µm
20.13µm
cyclic
20 d dark
Dark rear
19.31µm
20.08µm
18.01µm
Light
21.27µm
20.62µm
21.67µm
22.39µm
20.41µm
20.64µm
5D5L
7hr L/D
24.79µm
24.00µm
24.60µm
23.37µm
24.54µm
25.14µm
23.58µm
21.25µm
20.13µm
cyclic
20 d dark
Dark rear
19.31µm
20.08µm
18.01µm
Light
21.27µm
20.62µm
21.67µm
22.39µm
20.41µm
20.64µm
5D5L
7hr L/D
24.79µm
24.00µm
24.60µm
23.37µm
24.54µm
25.14µm
23.58µm
21.25µm
20.13µm
cyclic
20 d dark
Dark rear
19.31µm
20.08µm
18.01µm
Light
21.27µm
20.62µm
21.67µm
22.39µm
20.41µm
20.64µm
5D5L
7hr L/D
24.79µm
24.00µm
24.60µm
23.37µm
24.54µm
25.14µm
23.58µm
21.25µm
20.13µm
cyclic
20 d dark
Dark rear
19.31µm
20.08µm
18.01µm
Light
21.27µm
20.62µm
21.67µm
22.39µm
20.41µm
20.64µm
5D5L
7hr L/D
24.79µm
24.00µm
24.60µm
23.37µm
24.54µm
25.14µm
23.58µm
21.25µm
20.13µm
cyclic
20 d dark
Dark rear
19.31µm
20.08µm
18.01µm
Light
21.27µm
20.62µm
21.67µm
22.39µm
20.41µm
20.64µm
5D5L
7hr L/D
A
1
2
3
4
5
Base Tip
6
E
F
C
D
Distance_(inches)
Gray_Value
Gray_Value
Distance_(inches)
Gray_Value
Gray_Value
Distance_(inches)
Gray_Value
Gray_Value
Distance_(inches)
Gray_Value
Gray_Value
Distance_(inches)
Gray_Value
Gray_Value
Cyclic Light
20 days Dark (20dD)
Dark Reared
20 days Light (20dL)
F (a.u.)
5dD/5dL
F (a.u.)
7hrL/7hrD
B
1
2
3
4
5
6
0
100
0
Power, a.u.
p ≤ 0.01
Cyclic Light
1 1.5 2 2.5 3 3.5 4
Period, µm
10
5
0
p ≤ 0.01
20dD
p ≤ 0.01
Dark Rear
20dL
p ≤ 0.02
5dD/5dL
7hrL/7hrD
24.79µm
24.00µm
24.60µm
23.37µm
24.54µm
25.14µm
23.58µm
21.25µm
20.13µm
19.31µm
20.08µm
18.01µm
21.27µm
20.62µm
21.67µm
22.29µm
20.75µm
20.41µm
100
F (a.u.)
0
100
F (a.u.)
0
F (a.u.)
0
100
100
F (a.u.)
0
100
Power, a.u.
1 1.5 2 2.5 3 3.5 4
Period, µm
10
5
0
Power, a.u.
1 1.5 2 2.5 3 3.5 4
Period, µm
10
5
0
Power, a.u.
1 1.5 2 2.5 3 3.5 4
Period, µm
10
5
0
Power, a.u.
1 1.5 2 2.5 3 3.5 4
Period, µm
10
5
0
Power, a.u.
1 1.5 2 2.5 3 3.5 4
Period, µm
10
5
0
Timer B cyclic
Timer B cyclic
20 DD
20 DD
Dark Rear
Dark Rear
20 DL
20 DL
5D/5L
5D/5L
Timer B 7hrL/7hrD Timer B 7hrL/7hrD
2
8
14
20
Number of fluorescent rich bands
Lighting Conditions
* * *
Cyclic Light 20 dD Dark Rear 20 dL 5dD/d5L Timer B cyclic Timer B cyclic 20 DD
20 DD 20 DD (208)
20 DD (208) 20 DL 20 DL 20 DL/ 20 DD
20 DL/ 20 DD Dark Rear Dark Rear 5D/5L
5D/5L
8
6
4
2
14
12
10
Banding (diff Lighting conditions)
Timer B cyclic
Timer B cyclic
20 DD
20 DD
20 DL
20 DL
20 DL/ 20 DD
20 DD (208)
20 DD (208)
Timer B cyclic
Timer B cyclic 20 DD
20 DD 20 DD (208)
20 DD (208) 20 DL
20 DL 20 DL/ 20 DD 20 DL/ 20 DD Dark Rear
Dark Rear 5D/5L
5D/5L
8
6
4
2
14
12
10
Banding (diff Lighting conditions)
Timer B cyclic
Timer B cyclic
20 DD
20 DD
20 DL
20 DL
20 DL/ 20 DD
20 DD (208)
20 DD (208)
Timer B cyclic
Timer B cyclic 20 DD 20 DD 20 DD (208)
20 DD (208) 20 DL
20 DL 20 DL/ 20 DD
20 DL/ 20 DD Dark Rear
Dark Rear 5D/5L
5D/5L
8
6
4
2
14
12
10
Banding (diff Lighting conditions)
Timer B cyclic
Timer B cyclic
20 DD
20 DD
20 DL
20 DL
20 DL/ 20 DD
20 DD (208)
20 DD (208)
Timer B cyclic Timer B cyclic 20 DD 20 DD 20 DD (208)
20 DD (208) 20 DL
20 DL 20 DL/ 20 DD
20 DL/ 20 DD Dark Rear
Dark Rear 5D/5L
5D/5L
8
6
4
2
14
12
10
Banding (diff Lighting conditions)
Timer B cyclic
Timer B cyclic
20 DD
20 DD
20 DL
20 DL
20 DL/ 20 DD
20 DD (208)
20 DD (208)
Timer B cyclic
Timer B cyclic 20 DD 20 DD 20 DD (208) 20 DD (208) 20 DL
20 DL 20 DL/ 20 DD
20 DL/ 20 DD Dark Rear Dark Rear 5D/5L 5D/5L
8
6
4
2
14
12
10
Banding (diff Lighting conditions)
Timer B cyclic
Timer B cyclic
20 DD
20 DD
20 DL
20 DL
20 DL/ 20 DD
20 DD (208)
20 DD (208)
Timer B cyclic Timer B cyclic 20 DD 20 DD 20 DD (208)
20 DD (208) 20 DL
20 DL 20 DL/ 20 DD
20 DL/ 20 DD Dark Rear Dark Rear 5D/5L
5D/5L
8
6
4
2
14
12
10
Banding (diff Lighting conditions)
Timer B cyclic
Timer B cyclic
20 DD
20 DD
20 DL
20 DL
20 DL/ 20 DD
20 DD (208)
20 DD (208)
7hrL/7hrD
*
Cyclic
Light
20dD Dark
Rear
20dL 5dD/
5dL
7hrL/
7hrD
20
14
8
2
Number of fluorescent rich bands
* * * *
Timer B cyclic
Timer B cyclic
20 DD 20 DD
Dark Rear
Dark Rear
20 DL 20 DL
5D/5L
5D/5L
Timer B 7hrL/7hrD Timer B 7hrL/7hrD
10
15
20
25
30
Length (diff Lighting conditions)
Length of ROS (μm)
* *
Cyclic Light 20 dD Dark Rear 20 dL 5dD/5dL
Lighting Conditions Timer B cyclic Timer B cyclic 20 DD 20 DD 20 DD (208)
20 DD (208) 20 DL 20 DL 20 DL/ 20 DD 20 DL/ 20 DD Dark Rear
Dark Rear 5D/5L 5D/5L
8
6
4
2
14
12
10
Banding (diff Lighting conditions)
Timer B cyclic
Timer B cyclic
20 DD
20 DD
20 DL
20 DL
20 DL/ 20 DD
20 DD (208)
20 DD (208)
Timer B cyclic Timer B cyclic 20 DD
20 DD 20 DD (208)
20 DD (208) 20 DL
20 DL 20 DL/ 20 DD 20 DL/ 20 DD Dark Rear
Dark Rear 5D/5L
5D/5L
8
6
4
2
14
12
10
Banding (diff Lighting conditions)
Timer B cyclic
Timer B cyclic
20 DD
20 DD
20 DL
20 DL
20 DL/ 20 DD
20 DD (208)
20 DD (208)
Timer B cyclic Timer B cyclic 20 DD
20 DD 20 DD (208)
20 DD (208) 20 DL 20 DL 20 DL/ 20 DD 20 DL/ 20 DD Dark Rear Dark Rear 5D/5L 5D/5L
8
6
4
2
14
12
10
Banding (diff Lighting conditions)
Timer B cyclic
Timer B cyclic
20 DD
20 DD
20 DL
20 DL
20 DL/ 20 DD
20 DD (208)
20 DD (208)
Timer B cyclic
Timer B cyclic 20 DD 20 DD 20 DD (208) 20 DD (208) 20 DL 20 DL 20 DL/ 20 DD
20 DL/ 20 DD Dark Rear Dark Rear 5D/5L 5D/5L
8
6
4
2
14
12
10
Banding (diff Lighting conditions)
Timer B cyclic
Timer B cyclic
20 DD
20 DD
20 DL
20 DL
20 DL/ 20 DD
20 DD (208)
20 DD (208)
Timer B cyclic Timer B cyclic 20 DD 20 DD 20 DD (208) 20 DD (208) 20 DL 20 DL 20 DL/ 20 DD
20 DL/ 20 DD Dark Rear Dark Rear 5D/5L 5D/5L
8
6
4
2
14
12
10
Banding (diff Lighting conditions)
Timer B cyclic
Timer B cyclic
20 DD
20 DD
20 DL
20 DL
20 DL/ 20 DD
20 DD (208)
20 DD (208)
Timer B cyclic Timer B cyclic 20 DD
20 DD 20 DD (208) 20 DD (208) 20 DL 20 DL 20 DL/ 20 DD 20 DL/ 20 DD Dark Rear
Dark Rear 5D/5L
5D/5L
8
6
4
2
14
12
10
Banding (diff Lighting conditions)
Timer B cyclic
Timer B cyclic
20 DD
20 DD
20 DL
20 DL
20 DL/ 20 DD
20 DD (208)
20 DD (208)
Cyclic 7hrL/7hrD
Light
20dD Dark
Rear
20dL 5dD/
5dL
7hrL/
7hrD
30
25
20
Length of ROS (µm) 15
10
* *
Rhodopsin concentration
nmol/retina
Length of ROS
Cyclic Light 20 dD Dark Rear 20 dL 5dD/5dL
Lighting Conditions Cyclic Light (Timer) Norm
Cyclic Light (Timer) Norm
Dark Adapt (Timer) Norm
Dark Adapt (Timer) Norm
Light (Timer) Norm
Light (Timer) Norm
Dark Rear (Timer) Norm
Dark Rear (Timer) Norm
5D/5L (Timer) Norm
5D/5L (Timer) Norm
0.7
0.6
0.5
0.4
0.3
0.2
Cyclic Light (Timer) Norm
Cyclic Light (Timer) Norm
Dark Adapt (Timer) Norm
Dark Adapt (Timer) Norm
Light (Timer) Norm
Light (Timer) Norm
Dark Rear (Timer) Norm
Dark Rear (Timer) Norm
5D/5L (Timer) Norm
5D/5L (Timer) Norm
0.7
0.6
0.5
0.4
0.3
0.2
Cyclic Light (Timer) Norm
Cyclic Light (Timer) Norm
Dark Adapt (Timer) Norm
Dark Adapt (Timer) Norm
Light (Timer) Norm
Light (Timer) Norm
Dark Rear (Timer) Norm
Dark Rear (Timer) Norm
5D/5L (Timer) Norm
5D/5L (Timer) Norm
0.7
0.6
0.5
0.4
0.3
0.2
Cyclic Light (Timer) Norm
Cyclic Light (Timer) Norm
Dark Adapt (Timer) Norm
Dark Adapt (Timer) Norm
Light (Timer) Norm
Light (Timer) Norm
Dark Rear (Timer) Norm
Dark Rear (Timer) Norm
5D/5L (Timer) Norm
5D/5L (Timer) Norm
0.7
0.6
0.5
0.4
0.3
0.2
Cyclic Light (Timer) Norm
Cyclic Light (Timer) Norm
Dark Adapt (Timer) Norm
Dark Adapt (Timer) Norm
Light (Timer) Norm
Light (Timer) Norm
Dark Rear (Timer) Norm
Dark Rear (Timer) Norm
5D/5L (Timer) Norm
5D/5L (Timer) Norm
0.7
0.6
0.5
0.4
0.3
0.2
Cyclic Light (Timer) Norm
Cyclic Light (Timer) Norm
Dark Adapt (Timer) Norm
Dark Adapt (Timer) Norm
Light (Timer) Norm
Light (Timer) Norm
Dark Rear (Timer) Norm
Dark Rear (Timer) Norm
5D/5L (Timer) Norm
5D/5L (Timer) Norm
0.1
0.3
0.5
0.7
Copy of Finalized Rho Conc
Cyclic Light (Timer) Norm
Cyclic Light (Timer) Norm
Dark Adapt (Timer) Norm
Dark Adapt (Timer) Norm
Light (Timer) Norm
Light (Timer) Norm
Dark Rear (Timer) Norm
Dark Rear (Timer) Norm
5D/5L (Timer) Norm
5D/5L (Timer) Norm
0.1
0.3
0.5
0.7
Copy of Finalized Rho Conc
Cyclic
Light
20dD Dark
Rear
20dL 5dD/
5dL
0.7
0.5
0.3
0.1
Rhodopsin concentration
nmol/retina
Length of ROS
cyclic light
cyclic light
20dd
20dd
dark rear
dark rear
5D5L
5D5L
7hD/7HL low
7hD/7HL low
7hD/7HL High
7hD/7HL High
0
1
2
3
4
Peak Period (μm)
* *
*
Cyclic Light 20dD Dark Rear 5dD/5dL
Lighting Conditions
7hrL/7hrD
(upper)
7hrL/7hrD
(lower) Cyclic Light (Timer) Norm
Cyclic Light (Timer) Norm
NCKX1 Het
NCKX1 Het
CNBG1 Het
CNBG1 Het
4
3
2
1
Cyclic Light (Timer) Norm
Cyclic Light (Timer) Norm
NCKX1 Het
NCKX1 Het
CNBG1 Het
CNBG1 Het
4
3
2
1
Cyclic Light (Timer) Norm
Cyclic Light (Timer) Norm
NCKX1 Het
NCKX1 Het
CNBG1 Het
CNBG1 Het
4
3
2
1
Cyclic Light (Timer) Norm
Cyclic Light (Timer) Norm
NCKX1 Het
NCKX1 Het
CNBG1 Het
CNBG1 Het
4
3
2
1
Cyclic Light (Timer) Norm
Cyclic Light (Timer) Norm
NCKX1 Het
NCKX1 Het
CNBG1 Het
CNBG1 Het
4
3
2
1
Cyclic Light (Timer) Norm
Cyclic Light (Timer) Norm
NCKX1 Het
NCKX1 Het
CNBG1 Het
CNBG1 Het
4
3
2
1
cyclic light
cyclic light
20dd
20dd
dark rear
dark rear
5D5L
5D5L
7hD/7HL low
7hD/7HL low
7hD/7HL High
7hD/7HL High
0
1
2
3
4
Cyclic
Light
20dD Dark
Rear
5dD/
5dL
7hrL/
7hrD
7hrL/
7hrD
*
* * 4
Peak Period (µm) 3
2
1
0
short
long

46
banding periods) because it can only test one frequency. (C) Number of Rho-Timer rich bands are displayed
as box plots. The values (mean ± SD) are as follows: 10.5 ± 1.3 (cyclic light), 10.2 ± 0.9 (20dD), 8.5 ± 1.1
(dark reared), 1.2 ± 0.6 (20dL), 5.1 ± 1.3 (5dD/5dL), 15.2 ± 1.1 (7hL/7hD). The number of Rho-Timer rich
bands is significantly different in dark reared, 20dL, 5dD/5dL, and 7hL/7hD conditions from cyclic light
(Kruskal-Wallis, *p < 0.01). No difference was observed between cyclic light and 20dD (p = 0.37). N = 140-
330 rods. (D) Comparison of ROS length from mice exposed to the indicated lighting conditions. The mean
ROS length are as follows: cyclic light, 22.8 ± 2.1 µm; 20dD, 22.6 ± 2.1 µm; dark reared, 21.8 ± 1.6 µm;
20dL, 17.4 ± 1.7 µm; 5dD/5dL, 19.1 ± 2.2µm; 7hL/7hD, 22.0 ± 1.5 µm. ROS lengths were significantly
shorter in 20dL and 5dD/5dL compared to cyclic light (Kruskal-Wallis, *p < 0.01). Differences between
average ROS length from cyclic light vs dark reared and 7hL/7hD were negligible due to small effect size
(Cliff’s delta < 0.2). Gray bar is interquartile range from cyclic light. N = 130-350 rods. (E) Comparison of
peak period (µm) for individual rods from each condition (50-80 rods, mean ± SD). There is no significant
difference in peak period between cyclic light and 20dD (p = 0.10), whereas dark reared and 5dD/5dL
banding pattern exhibit a larger peak period compared to cyclic light (Kruskal-Wallis, *p < 0.01). 7hL/7hD
displayed two peak periods: one significantly smaller (*p < 0.01) and one indistinguishable from cyclic light
(p = 0.166). (F) Rhodopsin concentration was normalized to average ROS length for each condition and
did not differ significantly (t-test, p > 0.05, N = 7-9).
Light history and intrinsic clock regulate Rho-Timer’s entry into mammalian ROS
To establish whether the periodic banding pattern of Rho-Timer is driven solely by light
exposure184,185 or an intrinsic retinal circadian rhythm,
222,223 we examined Rho-Timer distribution
in the ROS from mice subjected to five different lighting regimens. Rho-TimerB mice were initially
reared in cyclic light for 4 weeks and then were moved to 20 days of constant dark (20dD), 20
days of constant dim ambient light (20dL), 5 days constant dark followed by 5 days constant light
(5dD/5dL), or 7h light/7h dark cycles (7hL/7hD, 20 days). Mice born and reared in total darkness
were also included to control for the effect of initial cyclic light history of previous groups.
The fluorescent banding pattern persisted when Rho-TimerB mice were switched from
cyclic light to 20dD, with a periodicity along the length of the ROS that was highly reproducible
(Figure 2.5A2) and visually identical to ROS data obtained from cyclic light (Figure 2.5A1). These
findings were also observed in Rho-TimerA mice (Figure S2.2). Averaging the power spectra from
80 rods showed a prominent peak at a spatial period of 2.52 µm (Figure 2.5B2), which was not
statistically different from cyclic light (Figure 2.5E, p = 0.10). Thus, the ~10 fluorescent rich bands
within the ROS after 20dD strongly suggest that the entry of Rho-Timer (and endogenous
rhodopsin by extension) into the ROS is not regulated solely by light, as others have previously

47
reported,
184,185 but also by a retinal circadian pacemaker that maintains the daily coordination of
rhythmic rhodopsin delivery to the ROS during prolonged darkness.
The persistent banding pattern seen in 20dD may have originated from entrainment during
the preceding cyclic light-rearing period. To explore this possibility, Rho-TimerB mice were born
and reared in absolute darkness, and imaging was performed 4-8 weeks after birth. Consistent
with both cyclic light and 20dD findings, we observed a distinct and well-defined periodic
fluorescent banding pattern (Figure 2.5A3). However, we found fewer bands (~8 bands, Figures
5A3 and 5C, p < 0.01) compared to those observed in the ROS isolated from cyclic light and 20dD
mice (~10 bands), and the bands had a longer spatial period of 2.71 µm (Figures 2.5B3 and 2.5E).
Assuming a similar rate of disc incorporation, our results indicate that the retinal clocks
responsible for the rhythmic incorporation of Rho-Timer (and rhodopsin) to the ROS of dark reared
mice exhibit a period exceeding 24h when not synchronized with the standard 24h light/dark cycle.
Considering the reported inhibitory effect of light on rhodopsin trafficking to the ROS185
and its influence on retinal clock function,
180,224,225 we sought to determine whether low ambient
light alone is capable of preventing Rho-Timer entry into the ROS. When Rho-TimerB mice were
switched from cyclic light to 20dL, the periodic banding disappeared (Figures 2.5A4 and 2.5C).
Surprisingly, individual intensity profiles showed moderately bright fluorescence with minimal
fluorescent variation along the ROS length, and Fisher’s g-statistic did not detect any periodic
pattern (p = 0.15, Figure 2.5B4). We exclude the possibility that this is a consequence of retinal
light damage as histological sections revealed a slight shortening of ROS but no thinning of the
photoreceptor layer (Figure S2.3).
The clear correlation between the number of fluorescent bands in cyclic light-reared mice
and the known rate of ROS turnover suggests that each bright/dim fluorescent band corresponds
to the segment of a ROS made per day. We next moved cyclic light-reared Rho-TimerB mice to 5
days dark (5dD) followed by 5 days light (5dL) to see how this change is recorded along the length
of the ROS. These animals exhibited banding along the distal region (towards the tip) of the ROS

48
that was produced during the 5 days of constant dark, while the ROS produced during the 5 days
of constant light did not exhibit a discrete banding pattern (Figure 2.5A5), resembling ROS formed
in 20dL (Figure 2.5A4). A representative fluorescence intensity profile from 5dD/5dL ROS clearly
shows a loss of banding at the ROS base, with periodic fluctuations in fluorescent intensities at
the tip (Figure 2.5A5). Analysis of the distal half of the ROS produced during the 5dD revealed a
banding pattern with a spatial period of 2.69 µm (Figures 2.5B5 and 2.5E), similar to that of 20dD
or dark reared mice. However, although banding periodicity was maintained to some degree, the
contrast between bright and dim bands at the distal half was less pronounced when compared to
bands formed under cyclic light, or 20 days darkness (Figure 2.5A), suggesting mixing of
rhodopsin between bright and dim discs. Our results suggest that 5 days of constant ambient light
not only affected the distribution of Rho-Timer in newer discs, but also had an impact on the
distribution of Rho-Timer in older discs made during the period of constant darkness.
To further evaluate the extent to which the timing of light exposure alone can impact the
system responsible for regulating rhodopsin incorporation into the ROS, we subjected mice to an
abnormally short 7h/7h light/dark (7hL/7hD) schedule. We found that isolated rods from RhoTimerB mice exposed to 7hL/7hD for 20 days also displayed prominent periodic banding along
the length of the ROS, with a significantly greater number of fluorescent bands (~15 bands,
Figures 2.5A6 and 2.5C, p < 0.01). Two distinct spatial periods are present along the ROS length
(Figure 2.5A6): a notably short spatial period of 2.2 µm (white line) and a longer spatial period of
2.32 µm (red line) that contains a ‘doublet’ band (Figures 2.5B6 and 2.5E). Given their ROS length
of 22 ± 2.5 µm (Figure 2.5D) and assuming a normal 10 day turnover time (240h), these findings
suggest that the formation of one bright and one dim band (T = 1.2 µm) likely occurs within the
subjective 14h day, consistent with the 7hL/7hD lighting schedule (240h/22 µm = 10.9h/µm; 1.2
µm x 10.9h = 13.1 ± 1h to produce one dim and one bright band). Taken together, our data
highlight the profound effect of light/dark cycles on rhodopsin incorporation into the ROS and the
potential of light to override retinal clocks.

49
Rhodopsin content along ROS length is maintained under different lighting conditions
Photostasis is a term used to describe the phenomenon that ROS length and rhodopsin
content change depending on the lighting environment.
226 We examined whether these changes
occurred in retinae from Rho-TimerB mice exposed to the five different lighting conditions. The
ROS from these mice were labeled by rhodamine-conjugated wheat germ agglutinin (Figure 2.1F,
red) and their length was manually measured by tracing the ROS from the base to the tip. The
measurements revealed no statistical difference between ROS length in cyclic light and 20dD,
(Figure 2.5E, p > 0.05), while measurements of ROS from 20dL, dark reared, 5dD/5dL, and
7hL/7hD revealed significant differences when compared to the cyclic light ROS length range (p
< 0.01, Kruskal-Wallis one-way ANOVA, Figure 2.5D). Upon closer analysis, Cliff’s delta statistic
revealed that the effect size estimates between cyclic light, dark reared, and 7hL/7hD were smallto-negligible in magnitude (Cliff’s delta < 0.2), indicating that the only significant variations in ROS
length were between cyclic light, 20dL, and 5dD/5dL. Interestingly, the average ROS length of
mice exposed to prolonged darkness (20dD, dark reared) was similar to that from mice raised in
cyclic light, in contrast to ROS lengthening expected from photostasis.
227 On the other hand, the
ROS shortened when mice were exposed to constant light for extended periods (20dL, 5dD/5dL),
a phenomenon consistent with photostasis.
49,228,229 Next, we measured rhodopsin content from
retinae collected from the different lighting conditions and then normalized this value to average
ROS length (Figure 2.5D). It appears that when normalized against ROS length, rhodopsin
content per unit length remains relatively similar across different lighting conditions (Figure 2.5F).
Strong and weak Rho-Timer fluorescent bands represent different points of fragility along
the ROS
In most cases, the ROS of intact rods from dissociated retinae are straight (Figures 2.1F
and 2.5A1), a characteristic suggestive of its stiffness or rigidity. However, a small subsect of rods
were bent, which may have been a consequence of the mechanical force exerted on them during

50
the retinal chopping process before confocal imaging. Notably, it has been shown, both
experimentally and through theoretical modeling, that mechanical stress can cause the ROS from
Xenopus rods to bend and break at points along which higher density of fluorescently tagged
rhodopsin is observed.
230 We therefore examined whether this is also the case for mammalian
rods or whether the breaking points occurred randomly. In contrast to Xenopus rods, images of
95 bent ROS (N = 30 mice; pooled from cyclic light and 20dD) revealed that the majority (93.7%)
of bending/breakage occurred at fluorescent-deficient regions (Figure 2.6). These results suggest
that the fluorescent pattern of Rho-Timer reflects structural inhomogeneities along the ROS
length, with bands exhibiting lower Rho-Timer density tending to be more flexible, bending or
breaking more easily, than Rho-Timer high-density bands.
Figure 2.6. Rho-Timer deficient bands corresponds to structural weak points in the ROS.
Representative live-cell images of bent ROS (measured in µm) from Rho-TimerB retinae and its respective
fluorescence intensity profile (N = 30 mice, 95 rods; pooled from cyclic light and 20 days dark). ROS bending
typically occurs within a low-density Rho-Timer fluorescent band. White arrows point to the bent region in
the ROS image and black arrows point to the corresponding region in the fluorescence profile.
Distance_(inches)
Gray_Value
Distance_(inches)
Gray_Value
Distance_(inches)
Gray_Value
Distance_(inches)
Gray_Value
Distance_(inches)
Gray_Value
0 Distance (µm) 24.89
0 22.84 Distance (µm)
0 Distance (µm) 27.53
0 20.86 Distance (µm)
0 27.59 Distance (µm)
Intensity, a.u. Intensity, a.u. Intensity, a.u. Intensity, a.u. Intensity, a.u.
150
5
24.89µm
22.84µm
27.53µm
20.86µm
27.59µm
150
5
150
5
150
5
150
5

51
Spatial organization of Rho-Timer reveals rhodopsin incorporation defects in two
autosomal recessive disease models
It has been well documented that CNG channel- and NCKX1-deficient rods exhibit
disrupted ROS morphology and altered rhodopsin concentrations at different stages of
degeneration.
166,173,231,232 Disrupted ROS is already evident at one month of age, with Cngb1-/-
rods appearing more affected (Figure 2.7A). At this time point, Nckx1-/- retinae173 exhibited
negligible rod loss while ~20% rod loss232 had transpired in Cngb1-/- retinae. Consistent with the
recessive nature of disease caused by mutations affecting these genes, the ROS from both
Cngb1+/- and Nckx1+/- rods appeared structurally intact (Figure 2.7A). To evaluate how rhodopsin
is incorporated into ROS in these disease models, we imaged ROS from cyclic light-reared 1-
month-old Cngb1+/-
, Cngb1-/-
, Nckx1+/-
, and Nckx1-/- mice co-expressing Rho-Timer. Nckx1+/- rods
exhibited a periodic pattern of ~10 alternating fluorescent rich and deficient bands that were
almost indistinguishable from wildtype Rho-Timer mice (p = 0.44, spatial period = 2.44 µm,
Figures 2.7B-7E). Interestingly, the spacing of Rho-Timer fluorescent rich bands in a large
proportion of Cngb1+/- rods was irregular (85 rods of 146 rods examined or 58%, Figure 2.7B),
and this uneven pattern led to a marginally wider spectrum of peak periods (Figure 2.7C), with an
average period of 2.32 µm (Figure 2.7D). Despite the irregular spacing, Cngb1+/- ROS exhibited
an average of ~10 bands along its length (Figure 2.7E). This subtle deviations in rhodopsin
incorporation did not have a noticeable impact on ROS morphology or length (Figures 2.7A, 2.7F,
and 2.7G). Lastly, the ROS isolated from Cngb1-/- and Nckx1-/- rods expressing Rho-Timer
revealed visibly irregular fluorescent banding patterns that lacked any consistent periodicity
(Figures 2.7B and 2.7C), with Fisher’s g-statistic failing to detect reliable periodic signals along
the length of the ROS (p > 0.05). Some regions within the ROS in both Cngb1-/- and Nckx1-/- rods
displayed a complete absence of banding alongside expanded areas featuring dim signals (Figure
2.7B), resulting in statistically significant lower number of fluorescent rich bands (Figure 2.7E).
Like wildtype Rho-Timer rods (Figure 2.6), the dim fluorescent regions of the ROS from Nckx1-/-

52
mice appeared to exhibit a greater tendency to bend (Figure S2.4A and Figure 2.7B). Fiji Kappa
curvature analysis revealed that Nckx1-/- ROS displayed higher total curvature along their length
compared to cyclic light-reared Rho-Timer mice (Figure S2.4B and S2.4C). Additionally, few
individual fluorescent rich bands displayed longitudinal rather than horizontal geometry in Cngb1-
/- rods expressing Rho-Timer (Figure 2.7B, asterisks), a phenotype reminiscent of the
longitudinally arranged discs discovered in Cngb1-/- ROS.
166,231,232
Figure 2.7. Rho-Timer localization in disease models. (A) Light micrograph of retinal sections from 1-
month-old Rho-Timer mice of the indicated genotype. ROS from Nckx1+/- and Cngb1+/- mice appears
Timer B cyclic
Timer B cyclic
NCKX1 Het
NCKX1 Het
NCKX1 KO
NCKX1 KO
CNGB1 Het
CNGB1 Het
CNGB1 KO
CNGB1 KO
0
5
10
15
20
Banding (TimerA/CNGB/NCKX1)
Number of fluorescent rich bands
* *
Cyclic
Light
NCKX1
Het
CNGB1
Het
NCKX1
KO Timer B cyclic Timer B cyclic 20 DD 20 DD 20 DD (208) 20 DD (208) 20 DL 20 DL 20 DL/ 20 DD 20 DL/ 20 DD Dark Rear Dark Rear 5D/5L
5D/5L
8
6
4
2
14
12
10
Banding (diff Lighting conditions)
Timer B cyclic
Timer B cyclic
20 DD
20 DD
20 DL
20 DL
20 DL/ 20 DD
20 DD (208)
20 DD (208)
Timer B cyclic Timer B cyclic 20 DD 20 DD 20 DD (208) 20 DD (208) 20 DL 20 DL 20 DL/ 20 DD 20 DL/ 20 DD Dark Rear Dark Rear 5D/5L 5D/5L
8
6
4
2
14
12
10
Banding (diff Lighting conditions)
Timer B cyclic
Timer B cyclic
20 DD
20 DD
20 DL
20 DL
20 DL/ 20 DD
20 DD (208)
20 DD (208)
Timer B cyclic
Timer B cyclic 20 DD 20 DD 20 DD (208) 20 DD (208) 20 DL
20 DL 20 DL/ 20 DD 20 DL/ 20 DD Dark Rear Dark Rear 5D/5L 5D/5L
8
6
4
2
14
12
10
Banding (diff Lighting conditions)
Timer B cyclic
Timer B cyclic
20 DD
20 DD
20 DL
20 DL
20 DL/ 20 DD
20 DD (208)
20 DD (208)
Timer B cyclic Timer B cyclic 20 DD 20 DD 20 DD (208) 20 DD (208) 20 DL
20 DL 20 DL/ 20 DD 20 DL/ 20 DD Dark Rear
Dark Rear 5D/5L
5D/5L
8
6
4
2
14
12
10
Banding (diff Lighting conditions)
Timer B cyclic
Timer B cyclic
20 DD
20 DD
20 DL
20 DL
20 DL/ 20 DD
20 DD (208)
20 DD (208)
Timer B cyclic Timer B cyclic 20 DD 20 DD 20 DD (208) 20 DD (208) 20 DL 20 DL 20 DL/ 20 DD 20 DL/ 20 DD Dark Rear
Dark Rear 5D/5L 5D/5L
8
6
4
2
14
12
10
Banding (diff Lighting conditions)
Timer B cyclic
Timer B cyclic
20 DD
20 DD
20 DL
20 DL
20 DL/ 20 DD
20 DD (208)
20 DD (208)
CNGB1
KO Nckx1
+/-
Cngb1
+/-
Nckx1
-/-
Cngb1
-/-
Cyclic
Light
* *
Number of fluorescent rich bands15
10
5
0
Timer B cyclic
Timer B cyclic
NCKX1 Het
NCKX1 Het
NCKX1 KO
NCKX1 KO
CNGB1 Het
CNGB1 Het
CNGB1 KO
CNGB1 KO
0
5
10
15
Banding (TimerA/CNGB/NCKX1)
Timer B cyclic
Timer B cyclic
NCKX1 Het
NCKX1 Het
NCKX1 KO
NCKX1 KO
CNGB1 Het
CNGB1 Het
CNGB1 KO
CNGB1 KO
0
5
10
15
20
25
30
35
Length (TimerA/CNGB/NCKX1)
Length of ROS (μm)
* *
Cyclic
Light
NCKX1
Het
CNGB1
Het
NCKX1
KO Timer B cyclic Timer B cyclic 20 DD 20 DD 20 DD (208) 20 DD (208) 20 DL 20 DL 20 DL/ 20 DD 20 DL/ 20 DD Dark Rear
Dark Rear 5D/5L 5D/5L
8
6
4
2
14
12
10
Banding (diff Lighting conditions)
Timer B cyclic
Timer B cyclic
20 DD
20 DD
20 DL
20 DL
20 DL/ 20 DD
20 DD (208)
20 DD (208)
Timer B cyclic Timer B cyclic 20 DD 20 DD 20 DD (208) 20 DD (208) 20 DL 20 DL 20 DL/ 20 DD 20 DL/ 20 DD Dark Rear Dark Rear 5D/5L 5D/5L
8
6
4
2
14
12
10
Banding (diff Lighting conditions)
Timer B cyclic
Timer B cyclic
20 DD
20 DD
20 DL
20 DL
20 DL/ 20 DD
20 DD (208)
20 DD (208)
Timer B cyclic Timer B cyclic 20 DD
20 DD 20 DD (208) 20 DD (208) 20 DL 20 DL 20 DL/ 20 DD 20 DL/ 20 DD Dark Rear
Dark Rear 5D/5L 5D/5L
8
6
4
2
14
12
10
Banding (diff Lighting conditions)
Timer B cyclic
Timer B cyclic
20 DD
20 DD
20 DL
20 DL
20 DL/ 20 DD
20 DD (208)
20 DD (208)
Timer B cyclic Timer B cyclic 20 DD 20 DD 20 DD (208) 20 DD (208) 20 DL 20 DL 20 DL/ 20 DD 20 DL/ 20 DD Dark Rear Dark Rear 5D/5L 5D/5L
8
6
4
2
14
12
10
Banding (diff Lighting conditions)
Timer B cyclic
Timer B cyclic
20 DD
20 DD
20 DL
20 DL
20 DL/ 20 DD
20 DD (208)
20 DD (208)
Timer B cyclic Timer B cyclic 20 DD 20 DD 20 DD (208) 20 DD (208) 20 DL 20 DL 20 DL/ 20 DD 20 DL/ 20 DD Dark Rear
Dark Rear 5D/5L 5D/5L
8
6
4
2
14
12
10
Banding (diff Lighting conditions)
Timer B cyclic
Timer B cyclic
20 DD
20 DD
20 DL
20 DL
20 DL/ 20 DD
20 DD (208)
20 DD (208)
CNGB1
KO
A
22.40µm
25.31µm
22.36µm
27.14µm
25.83µm
23.35µm
23.78µm
23.54µm
22.94µm
22.80µm
21.97µm
25.62µm
11.76µm
10.40µm
10.29µm
12.33µm
12.07µm
Nckx1 +/-
B Base Tip
C Nckx1 +/-
Nckx1 -/-
Cngb1 +/-
Cngb1 -/-
p ≤ 0.01 p ≤ 0.01
RPE
ROS
+/- -/-
Nckx1 Cngb1
Distance_(inches)
Gray_Value
Distance_(inches)
Gray_Value
Base Tip
22.40µm
25.31µm
22.36µm
27.14µm
25.83µm
23.35µm
23.78µm
23.54µm
22.94µm
22.80µm
21.97µm
25.62µm
11.76µm
10.40µm
10.29µm
12.33µm
12.07µm
Distance_(inches)
Gray_Value
Nckx1 -/-
F (a.u.)
0
100
21.97µm
22.40µm
25.31µm
22.36µm
22.80µm
23.54µm
22.94µm
23.78µm
27.14µm
25.83µm
23.351µm
25.62µm
F (a.u.)
0
100
F (a.u.)
0
100Cngb1 +/-
D
E
F
G
Cyclic
Light
Nckx1
+/-
Cngb1
+/-
Nckx1
-/-
Cngb1
-/-
*
*
Cngb1 -/-
10.40µm
10.29µm
11.76µm
10.35µm
12.07µm
F (a.u.)
0
100
Distance_(microns)
Gray_Value
Power, a.u.
1 1.5 2 2.5 3 3.5 4
Period, µm
10
5
0
Power, a.u.
1 1.5 2 2.5 3 3.5 4
Period, µm
10
5
0
Power, a.u.
1 1.5 2 2.5 3 3.5 4
Period, µm
10
5
0
Power, a.u.
1 1.5 2 2.5 3 3.5 4
Period, µm
10
5
0
Cyclic Ligtht TimerB Norm
Cyclic Ligtht TimerB Norm
NCKX1 HET (Timer) Norm
NCKX1 HET (Timer) Norm
NCKX1 KO (Timer) Norm
NCKX1 KO (Timer) Norm
CNGB1 HET (Timer) Norm
CNGB1 HET (Timer) Norm
CNGB1 KO (Timer) Norm
CNGB1 KO (Timer) Norm
0.0
0.2
0.4
0.6
0.8
Rho Conc (norm rod death data points)
Cyclic Ligtht TimerB Norm
Cyclic Ligtht TimerB Norm
NCKX1 HET (Timer) Norm
NCKX1 HET (Timer) Norm
NCKX1 KO (Timer) Norm
NCKX1 KO (Timer) Norm
CNGB1 HET (Timer) Norm
CNGB1 HET (Timer) Norm
CNGB1 KO (Timer) Norm
CNGB1 KO (Timer) Norm
0.0
0.2
0.4
0.6
0.8
Rhodopsin concentration
nmol/retina
Length of ROS
Timer B cyclic Timer B cyclic 20 DD
20 DD 20 DD (208)
20 DD (208) 20 DL 20 DL 20 DL/ 20 DD
20 DL/ 20 DD Dark Rear Dark Rear 5D/5L
5D/5L
8
6
4
2
14
12
10
Banding (diff Lighting conditions)
Timer B cyclic
Timer B cyclic
20 DD
20 DD
20 DL
20 DL
20 DL/ 20 DD
20 DD (208)
20 DD (208)
Timer B cyclic
Timer B cyclic 20 DD
20 DD 20 DD (208) 20 DD (208) 20 DL
20 DL 20 DL/ 20 DD 20 DL/ 20 DD Dark Rear
Dark Rear 5D/5L 5D/5L
8
6
4
2
14
12
10
Banding (diff Lighting conditions)
Timer B cyclic
Timer B cyclic
20 DD
20 DD
20 DL
20 DL
20 DL/ 20 DD
20 DD (208)
20 DD (208)
Timer B cyclic
Timer B cyclic 20 DD
20 DD 20 DD (208) 20 DD (208) 20 DL 20 DL 20 DL/ 20 DD
20 DL/ 20 DD Dark Rear
Dark Rear 5D/5L
5D/5L
8
6
4
2
14
12
10
Banding (diff Lighting conditions)
Timer B cyclic
Timer B cyclic
20 DD
20 DD
20 DL
20 DL
20 DL/ 20 DD
20 DD (208)
20 DD (208)
Timer B cyclic
Timer B cyclic 20 DD
20 DD 20 DD (208)
20 DD (208) 20 DL
20 DL 20 DL/ 20 DD 20 DL/ 20 DD Dark Rear
Dark Rear 5D/5L
5D/5L
8
6
4
2
14
12
10
Banding (diff Lighting conditions)
Timer B cyclic
Timer B cyclic
20 DD
20 DD
20 DL
20 DL
20 DL/ 20 DD
20 DD (208)
20 DD (208)
Timer B cyclic Timer B cyclic 20 DD
20 DD 20 DD (208) 20 DD (208) 20 DL
20 DL 20 DL/ 20 DD
20 DL/ 20 DD Dark Rear Dark Rear 5D/5L
5D/5L
8
6
4
2
14
12
10
Banding (diff Lighting conditions)
Timer B cyclic
Timer B cyclic
20 DD
20 DD
20 DL
20 DL
20 DL/ 20 DD
20 DD (208)
20 DD (208)
Cyclic
Light
Nckx1
+/-
Cngb1
+/-
Nckx1
-/-
Cngb1
-/-
Rhodopsin concentration
nmol/retina
Length of ROS
0.8
0.6
0.4
0.2
0.0
*
Length of ROS (µm)
35
30
25
20
15
10
5
0
* *
*
ns
Cyclic Light Nckx1 +/- Cngb1 +/-
4
Peak Period (µm) 3
2
1
Cyclic Light (Timer) Norm
Cyclic Light (Timer) Norm
NCKX1 Het
NCKX1 Het
CNBG1 Het
CNBG1 Het
1
2
3
4
Cyclic Light (Timer) Norm
Cyclic Light (Timer) Norm
Dark Adapt (Timer) Norm
Dark Adapt (Timer) Norm
Light (Timer) Norm
Light (Timer) Norm
Dark Rear (Timer) Norm
Dark Rear (Timer) Norm
5D/5L (Timer) Norm
5D/5L (Timer) Norm
0.7
0.6
0.5
0.4
0.3
0.2
Cyclic Light (Timer) Norm
Cyclic Light (Timer) Norm
Dark Adapt (Timer) Norm
Dark Adapt (Timer) Norm
Light (Timer) Norm
Light (Timer) Norm
Dark Rear (Timer) Norm
Dark Rear (Timer) Norm
5D/5L (Timer) Norm
5D/5L (Timer) Norm
0.7
0.6
0.5
0.4
0.3
0.2
Cyclic Light (Timer) Norm
Cyclic Light (Timer) Norm
Dark Adapt (Timer) Norm
Dark Adapt (Timer) Norm
Light (Timer) Norm
Light (Timer) Norm
Dark Rear (Timer) Norm
Dark Rear (Timer) Norm
5D/5L (Timer) Norm
5D/5L (Timer) Norm
0.7
0.6
0.5
0.4
0.3
0.2

53
organized, while the ROS structure from Nckx1-/- and Cngb1-/- mice was more disrupted. N = 3 mice. Scale
bar = 20 μm. RPE, retinal pigmented epithelium; ROS, rod outer segment. (B) Representative images of
ROS from Nckx1+/-
, Nckx1-/-
, Cngb1+/-
, Cngb1-/- mice expressing Rho-Timer (N = 8-9 mice) with their
respective fluorescence intensity profiles (measured in µm). Wheat germ agglutinin (red) labeled ROS from
Cngb1-/- to demarcate the length and shape. Asterisks highlight vertically oriented Rho-Timer rich discs
(green). (C) Fast Fourier transform analysis of ROS presented in terms of average peak banding period
(inverted black triangle, Nckx1+/-
, T = 2.44 µm; Cngb1+/-
, T = 2.32 µm). Fisher’s g-statistic confirms
periodicity (p ≤ 0.01, N = 50-80 rods from N = 8-9 mice). Nckx1-/- (p = 0.06) and Cngb1-/- (p = 0.16) did not
reach statistical significance. (D) Comparison of peak period (µm) for individual ROS (50-80 rods, mean ±
SD). There is no significant difference in peak period among genotypes that display periodic banding
(Kruskal-Wallis, p > 0.05). (E) Number of Rho-Timer rich bands (mean ± SD) are as follows: 9.8 ± 0.98
(Nckx1+/-
), 5.1 ± 3.2 (NCKX1-/-
), 9.8 ± 1.1 (Cngb1+/-
), 3.1 ± 1.0 (Cngb1-/-
). The number of Rho-Timer rich
bands is significantly different in Nckx1-/- and Cngb1-/- (Kruskal-Wallis, *p < 0.01). N = 8-9 mice and 190-
250 rods. (F) Rhodopsin concentration was normalized to average ROS length (N = 5-7 mice). T-test
revealed a significant difference in the rhodopsin concentration for Nckx1-/- retinae (*p ≤ 0.05). There is no
statistical difference in rhodopsin concentration in Cngb1+/-
, Cngb1-/- (correcting for 20% rod loss), and
Nckx1+/- retinae. (G) Comparison of ROS length by genotype. The mean ROS length are as follows: cyclic
light, 22.8 ± 2.1 µm; Nckx1+/-
, 23.3 ± 2.3 µm; Nckx1-/-
, 20.9 ± 3.2 µm; Cngb1+/-
, 22.6 ± 1.9 µm; Cngb1-/-
, 10.3
± 2.0 µm. ROS lengths from knockout retinas were significantly shorter (Kruskal-Wallis, *p < 0.01, Cliff’s
delta; Nckx1-/-
, 0.4; Cngb1-/-
, 0.9). Gray bar is interquartile range from cyclic light. N = 8-9 mice, 190-250
rods.
Considering the presence of the irregularly spaced banding pattern observed in ROS
isolated from Cngb1+/-
, Cngb1-/-
, and Nckx1-/- mice, we assessed whether rhodopsin content or
ROS length were altered in these mutant mice. The absence of CNG channels and NCKX1 led
to shortened ROS when compared to control mice (p ≤ 0.01, Cliff’s delta; Nckx1-/-
, 0.4; Cngb1-/-
,
0.9, Figure 2.7G). For Nckx1-/- rods, the rhodopsin content, when normalized against ROS length,
revealed a reduction (p = 0.02, Figure 2.7F). This result correlates well with the imaging data that
show large segments of weak Rho-Timer fluorescence in these ROS (Figure 2.7B) which may
reflect regions of low rhodopsin density. On the other hand, Cngb1+/- and Nckx1+/- retinae showed
no noticeable reductions (p > 0.05, Figure 2.7F). For Cngb1-/- retinas, the normalized rhodopsin
content per shortened ROS was not significantly different from control mice after correcting for
20% rod loss at this age (p = 0.51).166,232
2.5 Discussion
The process of ROS maintenance has been a topic of interest due to the importance of
this specialized structure for phototransduction and rod survival. To enable tracking of rhodopsin’s

54
lifetime within the ROS, we generated two independent and stable transgenic mouse lines.
Focusing primarily on Rho-TimerB mice, we found that our fluorescently tagged rhodopsin fusion
protein 1) contributes to ~0.8% of endogenous rhodopsin transcript, 2) traffics properly to the
ROS, 3) does not alter endogenous rhodopsin concentration, retinal structure, or rod function,
and 4) mirrors the transcription pattern of native rhodopsin. These findings establish Rho-Timer
as an accurate reporter of endogenous rhodopsin that allowed us to visualize rhodopsin
incorporation into the ROS under various experimental lighting conditions and diseased states.
Because the ROS is renewed every ~10 days, Rho-Timer fluorescence pattern serves as a visual
tool to chronical the history of rhodopsin incorporation during this timeframe. This, in turn, allowed
us to gain insights into the interplay between external cues (light) and internal clock that regulate
the circadian rhythm of the organism as it adapts to the environment.
Discrete periodic banding patterns within the OS have been observed in Xenopus and
rodents using Nomarski optics. The direct underlying cause of this phenomenon, whether it is due
to protein or lipid inhomogeneity, is not known and awaits future investigations. In support of
rhodopsin contributing to heterogeneity in ROS, Hsu et. al.,185 showed ~10 fluorescent bands
along the ROS of wildtype mice using an optimized protocol for antibody binding to native
rhodopsin. Another line of evidence, using fluorescently labeled arrestin that bind light-activated
phosphorylated rhodopsin as a readout, showed a similar banding pattern in rat185 and Xenopus184
rods, with the number of bands corresponding to the number of days for ROS formation. Our
immunofluorescent labeling of endogenous light-activated phosphorylated rhodopsin with the
A11-82P antibody (Figure 2.4A) is consistent with these findings. This rhythmicity of endogenous
rhodopsin incorporation into the ROS may be the result of shifting balance between rhodopsin
synthesis and new lipid disc formation. The periodicity of rhodopsin synthesis is reflected at the
transcript level and may be maintained downstream through translation233 and transport steps as
well. Consequently, rhodopsin’s heterogeneity along the ROS would be maintained if the rate of
lipid disc formation is relatively constant throughout the day. Indeed, transmission electron

55
microscopy tomograms of the base of mouse ROS reveal relatively constant number of new open
discs throughout the day.36 In contrast to these results obtained in mice, direct
microspectrophotometric measurement of rhodopsin pigment concentration along the length of
the ROS from transgenic Xenopus did not detect variations in rhodopsin density.184 The reason
for this discrepancy is not clear, but may be related to the different mechanisms of rhodopsin
transport from the inner segment to the cilium base between amphibians and mammals,16,234,235
and hence rhodopsin incorporation into the ROS. Together, these data support the notion that
native rhodopsin incorporation into ROS discs in the mouse retina varies depending on the time
of day, and that the rhythmic banding of rhodopsin fusion proteins reflects endogenous rhodopsin
heterogeneity.
Previous reports using tagged rhodopsin constructs in rats185 and Xenopus184 show a strict
regulation of transgene incorporation into the ROS by light. For example, Hsu et al.185 showed
that Rho-GFP is retained in the cell body when animals were exposed to light and only enters the
ROS upon transitioning into darkness. When kept in darkness for 4 days, the exogenous
rhodopsin showed a homogeneous distribution along the ROS formed during that period. In the
Xenopus model, the Rho-GFP reliably records periods in which animals were kept in the light by
dim fluorescence and periods in darkness by bright fluorescence along the ROS.184 Contrary to
these reports, our results show that light does not act as a gate to constrain rhodopsin
incorporation into the ROS. For instance, when cyclic light-reared transgenic mice were
transitioned into darkness (20dD), fluctuations in Rho-Timer fluorescence intensities were
maintained and were indistinguishable from those observed under cyclic light (~10 bands, Figure
2.5A2). This periodicity was also reflected at the transcript level (Figure 2.3B). The reason for the
disparity between ours and the Xenopus study may be related to different mechanisms of
rhodopsin transport as mentioned above. As for the rat study, a possible explanation may lie in
differences in the intensity of light exposure. While both animal models were albinos, Rho-TimerB
mice were exposed to 100-lux, whereas the rat model (Sprague Dawley) was raised in 200-lux.

56
Notably, 270-lux is sufficient to cause severe damage and rod cell death in these albino rats.
236
Differences in the transgene construct and methodology of transgene incorporation
(electroporation vs. pronuclei injection) may also contribute.
Remarkably, the periodic banding pattern and rhythmic gene expression persisted in the
ROS from mice reared in total darkness (~8 bands, Figures 2.3C and 2.5A3). Earlier studies have
demonstrated that robust daily rhythms in the retina underlie essential physiologic processes,
including OS phagocytosis44,179,222 and visual pigment gene expression.
213,214 These rhythms have
also been shown to persist under constant darkness,
180,213,237,238 suggesting they are regulated by
autonomous, self-sustaining clocks located within the retina.
217 Our data are consistent with these
findings, where Rho-Timer expression and delivery to the ROS exhibit continuous rhythmicity in
darkness, confirming that these processes are regulated by a circadian clock. Notably, Rho-Timer
banding in the ROS isolated from dark reared, light-naïve mice displayed ~8 bright fluorescent
bands with a broader spatial period (T = 2.71 µm). This result indicates that the circadian clock
responsible for the periodic delivery of rhodopsin to the ROS has a period of ~28h. This finding
agrees with a recent discovery involving C57BL/6J mice which revealed that layer-specific clocks
demonstrate robust interdependence and can produce a longer, self-sustaining oscillatory period
of ~26-29h.239 We also observed a striking similarity between number of bands and peak spatial
period in cyclic light and cyclic light-reared mice transitioning into 20dD (Figure 2.5), indicating
that the periodicity in darkness can be entrained by light. Together, our results indicate that
rhodopsin delivery is controlled by external light cues and endogenous circadian clocks that can
be entrained to the ~24h day.
217,218
The interplay between external light cues and molecular clocks appears to be remarkably
plastic, as can be seen in Rho-Timer mice exposed to the abnormally short 7hL/7hD lighting
schedule. Their rods had notably higher number of Rho-Timer rich bands (~15) along the ROS
length (Figure 2.5A6), where one dim and rich fluorescent band reflected the subjective 14h day.
This result is consistent with recent studies in C57BL/6J mice and Syrian hamsters subjected to

57
abnormal 24h light:dark:light:dark (LDLD) cycles, which effectively ‘bifurcate’ behavioral rhythms
into two rest and activity intervals per 24h.240,241 Together, these results reveal unexpected
plasticity in the circadian system beyond the range of conventional entrainment.
We used BMAL1 transcript as a positive control in our qRT-PCR experiments because
BMAL1/Clock are highly conserved component of the molecular circadian clock and forms part of
the core feedback loop that interact with cellular metabolism. Indeed, its expression pattern was
periodic under cyclic light and 20dD, consistent with previous observation in mammals.
215,238,242
However, BMAL1 transcript was not rhythmic after 20dL and under dark rearing whereas both
endogenous rhodopsin and Rho-Timer showed strong rhythmicity under all lighting conditions.
This uncoupling suggests that BMAL1 is not responsible for rhythmic transcription of rhodopsin
or Rho-Timer. Our observation aligns with previous research indicating that BMAL1 expression is
absent in mouse rods215,243,244 and is primarily restricted to other retinal cell types.
217 Therefore,
the expression pattern seen in our BMAL1 qRT-PCR data might be influenced by cell-specific
rhythmic expression patterns that are not synchronized with each other.
243 Future studies are
necessary to further elucidate the regulation of rhodopsin transcription and ROS renewal
dynamics, either by an individual or a network of autonomous retinal clocks.
239
Photostasis is a widely accepted long-term adaptation phenomenon that describes ROS
plasticity under different lighting conditions to optimize photon catch. In these studies, albino rats
placed in constant light or constant dark led to ROS that are shortened or lengthened,
respectively.226 We observed a shortening of ROS in mice exposed to 20dL but no lengthening of
ROS in the dark conditions (20dD, dark reared). The lack of ROS lengthening in darkness was
also noted by Cunea et al.245 using pigmented and albino mice with sample sizes allowing for a
statistically rigorous study. These results put in question photostasis as a comprehensive
mechanism for long-term light adaptation.
Early pioneering work by Richard Young using pulsed radiolabeled amino acid in rats,
mice and frog demonstrated the principle of ROS renewal through addition of newly formed discs

58
at the base and older discs displaced distally. During this movement the concentration of
radioactivity in the band itself was stable, giving rise to the interpretation that the labeled protein
is displaced distally as a stable unit.30 Our current findings demonstrate that individual discs within
ROS are not entirely static, isolated membranous units: we found that rearing mice in 5dD/5dL
not only disturbed the periodic rhodopsin incorporation into newly formed discs, but it also affected
the banding pattern at the distal length of the ROS previously formed in darkness. This result
suggests that light exposure caused an exchange of rhodopsin between the older discs formed
in darkness. This light-induced mixing of rhodopsin may also explain the absence of bands in the
ROS formed during constant light despite rhythmic transcript expression during the 24h period
(Figure 2.3D). Our data is in agreement with previous findings using salamander rods incubated
with fluorescent polar and membrane tracers, present mostly in open discs at the ROS tip.
246
Once loaded onto these select discs, the fluorescent tracers can diffuse longitudinally along the
ROS, supporting an exchange of soluble and membrane material between discs.
246 These
observations challenge the previous notion that ROS discs are static entities once they are
formed.
The ROS’s unique structure makes it prone to injury when mechanically disturbed. Our
data provides evidence that the ROS tends to break at sites of low rhodopsin concentration, as
indicated by the dim Rho-Timer band. Therefore, the inhomogeneity of the fluorescent banding
appears to reflect subtle, yet tangible structural inhomogeneities associated with rhodopsin along
the ROS. We speculate that the dim fluorescent bands exhibit a decrease in resistance to bend
because of the lower density of rhodopsin, as it has been demonstrated that membrane stiffness
is modified by integral membrane proteins.247 In contrast, in Xenopus ROS expressing a
fluorescent rhodopsin reporter transgene, the ROS preferentially bent or fractured at bands of
high fluorescence.
230 Differences in ROS properties between mouse and Xenopus, such as
rhodopsin packing density and lipid bilayer composition, or differences in orientation and type of
mechanical stress, could account for the variations in ROS breakage. Nevertheless, it is evident

59
that there are regions along the length of the ROS that are more flexible, and mutations that affect
rhodopsin packing may further destabilize the ROS and promote bending and breakage.
Rhodopsin is a key structural element in the ROS. Given the profound impact mutations
in CNG channels and NCKX1 have on rod structure and function,
173,231 resulting in rhodopsin
mislocalization, disordered discs, and disrupted ROS, we investigated whether defect in
rhodopsin incorporation into the ROS is an early event in the disease. Using young Cngb1-/- and
Nckx1-/- mice at an age prior to overt rod loss, we observed extensive regions where Rho-Timer
fluorescence was minimal along the ROS, implying a substantial disruption in the mechanism
governing rhodopsin delivery into new discs. In Cngb1 heterozygotes, which to date has no
reported phenotypic or functional defects, irregular Rho-Timer banding pattern was also
observed. Such altered rhodopsin incorporation may have subtle effects on the ROS architecture
and disc composition. Indeed, we found that dissociated Nckx1-/- rods, characterized by irregular
banding patterns and larger regions of dim fluorescence, exhibited greater curvature (Figure
S2.4). The reason behind the altered packing of rhodopsin into nascent discs, and whether this
has functional consequences in the diseased state, are beyond the scope of the current study. It
is tempting to speculate that the rhythmic banding of rhodopsin in the normal ROS may offer a
balance of flexibility/rigidity to better withstand mechanical stress and is therefore physiologically
advantageous.

60
2.6 Supplemental Information
Figure S2.1. Low detection of Rho-Timer with rhodopsin antibodies. (A and B) Western blots of retinal
lysates from dark-adapted one-month-old Rho-TimerB mice and wildtype littermates (N = 3). Blots were
probed with antibodies to rhodopsin’s C- and N-terminus (1D4 and R2-12N, respectively) and DsRed
(green), from which Timer was derived (highlighted in white box). Rhodopsin appeared as monomer, dimer
and trimer, demonstrating its propensity to oligomerize, particularly when it is in the opsin form. The
apparent molecular weight for Rho-Timer (~170 kDa, as seen using DsRed antibody, arrows) on the
western is larger than the predicted molecular weight (66 kDa), indicating that the protein is aggregated.
Aggregation may have arisen from rhodopsin’s and/or Timer’s tendency to oligomerize. Aggregated RhoTimer was not detected using mouse monoclonal antibodies 1D4 (A and B, black box) and R2-12N (B). In
panel (B), the membrane was cut to separate monomer from the oligomerized rhodopsin because the
monomer signal was much higher than the oligomerized forms. This separation allowed higher exposure
time for the detection of Rho-Timer signal at ~170 kDa. Nevertheless, no signal was detected. Two protein
amounts (~6 µg, ~3 µg) were loaded for each genotype. M, molecular weight marker (kDa); WT, wildtype.
(C and D) Localization of Rho-Timer (green) and rhodopsin (magenta) was examined in one-month-old
Rho-TimerA (C) and Rho-TimerB (D) retinae (N = 3). Cryosections were probed with established antibodies
against rhodopsin’s C-terminus (1D4) or N-terminus (4D2). Intrinsic Rho-Timer fluorescence is seen in a
subset of rod inner segments (RIS) and plasma membrane within the outer nuclear layer (ONL). Rhodopsin
(1D4 and 4D2, magenta) immunoreactivity is also present in the ONL and RIS. Merged images show where
Rho-Timer and rhodopsin labeling are co-localized. The lack of consistent correlation between Rho-Timer
signal and signals from either rhodopsin antibodies (1D4 and 4D2) suggests that the presence of RhoA B
WTRhoTimerWTRhoTimer
WTRhoTimerWTRhoTimer
Rho
Monomer
Rho
Dimer
Rho
Trimer
1D4 R2-12N
93
53
41
170
M
125
70
30
22
18
14
235
6 µg 3 µg 6 µg 3 µg
Rho
Monomer
93
53
41
170
M
125
70
30
22
235
Rho
WT
-
Timer
RhoTimer
WT
DsRed and 1D4
Rho
WT
-
Timer
RhoTimer
WT
1D4
Rho
Dimer
Rho
Trimer
DsRed
6 µg 3 µg 6 µg 3 µg
C D
RIS
ONL
RIS
ONL
RIS
ONL
RIS
ONL

61
Timer does not significantly contribute to the total amount of rhodopsin within rods expressing Rho-Timer.
Nuclei are stained with DAPI (blue). Scale bar = 5 µm.
Figure S2.2. Periodic banding pattern is maintained in darkness for Rho-TimerA mice. Live-cell
images of ROS isolated from Rho-TimerA mice housed in 20 days darkness (20dD, N = 3). ROS (measured
in µm, base to tip) display stereotyped banding pattern seen in cyclic light and 20dD Rho-TimerB retinae.
ROS, rod outer segment; RIS, rod inner segment; N, nucleus; SN, synaptic terminal. Rod diagram was
created with BioRender.com.
Figure S2.3. Dim, ambient light does not induce rod loss. Light micrographs of representative retinal
sections collected from cyclic light-reared Rho-TimerB mice exposed to 20 days constant light (20dL, 100-
lux, N = 3) or 20 days constant darkness (20dD, N = 3). The ROS from Rho-TimerB mice in 20dL were
visibly shorter and disorganized compared to 20dD. However, the number of nuclei rows in the ONL was
comparable between mice housed in 20dL and those kept in 20dD, and no apoptotic nuclei were observed.
Base Tip
22.57µm
23.15µm
21.87µm
23.13µm
21.89µm
20.08µm
SN N RIS ROS
RPE
ROS
RIS
ONL
INL
GC
20dL 20dD
20 µm 20 µm

62
Scale bar = 20 µm. RPE, retinal pigmented epithelium; ROS, rod outer segment; RIS, rod inner segment;
ONL, outer nuclear layer; INL, inner nuclear layer; GC, ganglion cell layer.
Figure S2.4. ROS from Nckx1 knockout mice are more curved. (A) Live-cell images of ROS (measured
in µm) from cyclic light-reared Rho-Timer positive Nckx1-/- mice (N = 8). The representative ROS images
reveal that the ROS appears prone to bending at dim fluorescent regions (white arrows). Each fluorescence
profile highlights the corresponding bent regions (black arrows). (B) Dissociated ROS from wildtype RhoTimerB mice have exhibit distinct curvature types (straight, slight curve, and curved). Analysis using FIJI’s
Kappa (�) plug-in allowed quantification of curvature along the length of each ROS, yielding average
curvature/µm values for the different curvature sub-populations (straight ≤ 0.01; 0.01 > slight curve < 0.05;
curved ≥ 0.05). Scale bar = 2 µm. (C) Comparison of ROS curvature found in Rho-Timer positive wildtype
(N = 100 rods) and Nckx1-/- (N = 73 rods) retinae. The bar graph illustrates the distribution of ROS curvature
types (straight, slight curve, and curved) as percentages of the total ROS population observed in Rho-Timer
positive wildtype (light gray) and Nckx1-/- (dark gray) mice. Eighty-five percent of the dissociated ROS from
wildtype mice were straight or slightly curved, while 15% were curved. Conversely, ROS from Nckx1-/- mice
exhibited a tendency to bend, where notably 46% of the measured ROS were curved, and only 44%
appeared straight or slightly curved. The scatter plot of average curvature (µm-1) from both Rho-Timer
wildtype (circles) and Nckx1-/- (squares) rods is superimposed onto the bar graph. This provides visual
representation of the actual ROS curvature values that belong to each curvature sub-population found in
wildtype and Nckx1-/- rods. Taken together, these results indicate that Nckx1-/- rods are notably less linear.
The right y-axis is log scaled, allowing easy visualization of low values (<0.001) and higher values (<0.1).
Distance_(inches)
Gray_Value
Distance_(inches)
Gray_Value
Distance_(inches)
Gray_Value
0 Distance (µm) 24.42 Distance_(inches)
Gray_Value
0 Distance (µm) 24.01
0 Distance (µm) 23.85
Distance_(inches)
Gray_Value
0 Distance (µm) 26.15
0 Distance (µm) 26.34
Distance_(inches)
Gray_Value
0 Distance (µm) 21.73
Intensity, a.u.
80
0
Intensity, a.u.
80
0
Intensity, a.u. 80
0
Intensity, a.u.
80
0
Intensity, a.u.
80
0
Intensity, a.u.
80
0
24.42µm
26.15µm
21.73µm
24.01µm
26.34µm
23.85µm
A B
C
Straight Base Tip Curvature (µm-1)
0.0028
Slight Curve
Base Tip Curvature (µm-1)
0.0111
Curved
Base Tip Curvature (µm-1)
0.213
0.407
Straight
Slight Curve
Curved
0
10
20
30
40
50
Percentage (%)
Rho-Timer
Nckx1-/-
0.001
0.01
0.1
1
Rho-Timer
Nckx1-/-
Average curvature (µm-1)

63
CHAPTER 3
Charge Reversal of Rhodopsin’s Glu134Arg135 Impacts Transducin
Localization in Rods without Affecting Rhodopsin Phosphorylation
3.1 Summary
The highly conserved G-protein binding motif, Glu134/Asp134-Arg135-Try136 (E/DRY), is a
characteristic feature present all known class A G-protein-coupled receptors (GPCRs).
Rhodopsin, a prototypical GPCR, is required for visual perception, and its 3-D structure and
molecular interactions with conjugate proteins have been intensively studied. The charge reversal
from ERY to Arg134
-Glu135
-Try136 (REY) has been demonstrated, both by heterologous expression
and in-vivo studies, to uncouple light-activated rhodopsin (R*) from transducin activation without
impacting rhodopsin’s light absorption properties. However, in vivo, it remains unclear how this
mutation affects R*’s coupling to key phototransduction deactivation proteins, such as rhodopsin
kinase (GRK1) and arrestin. Using a knock-in mouse line expressing REY-rhodopsin (REY-Rho),
we monitored how this mutation affects R* signaling, R* phosphorylation, and light-activated
protein translocation. We verify that REY-Rho significantly diminishes rod sensitivity to light within
a live animal and observe a concurrent decrease in transducin translocation. We also find that
the charge reversal ERY → REY does not impact the phosphorylation of R* by GRK1 and the
light-dependent subcellular movement of arrestin. Furthermore, our results provide additional
support for the existence of a stable, preassembled complex between rhodopsin and transducin
in darkness.

64
3.2 Introduction
G protein-coupled receptors (GPCRs) are a biologically important and functionally diverse
family of membrane proteins. Their primary function involves converting external stimuli, such as
hormones, ions, and photons, into intracellular signaling events,248,249 thereby influencing various
physiological processes, ranging from immune system modulation to sensory transduction.
250
Defined by their characteristic seven transmembrane (TM) helical architecture,251 all activated
GPCRs exhibit a shared mode of signaling, recruiting heterotrimeric G-proteins (�, �, � subunits),
arrestins (�-arrestins), and GPCR kinases (GRKs) in a ligand-specific manner to transduce or
terminate the signal.252-254 The categorization of GPCRs into six distinct classes (A-F) relies on
identifying highly conserved amino acid motifs and residues within each specific class, which
frequently contribute to the receptor’s function.255,256
Rhodopsin, a prototypical class A GPCR and the first structurally determined, highresolution GPCR,
257 has served as a model system for uncovering the functional mechanisms
underlying GPCR signaling.
258 Responsible for dim light vision in the vertebrate eye, rhodopsin
consists of the apoprotein opsin linked to a chromophore (11-cis-retinal) and is densely packed
in the outer segment (OS) discs of rod photoreceptors to optimize photon capture.2,259,260 Upon
photon absorption, photoactivated rhodopsin (R*) initiates a signaling cascade, involving GDP-toGTP exchange on its cognate G-protein, transducin, resulting in the dissociation of individual T�-
GTP and T�� subunits. Simultaneously, rapid quenching of R*’s activity involves sequential
phosphorylation by G-protein kinase-1 (GRK1),
57,58,261 followed by the binding of visual arrestin-1
(ARR1),
262,263 resulting in complete deactivation of R*.264 These tightly controlled mechanisms
ensure both the rapid activation and deactivation of the signaling cascade in response to photon
capture by rhodopsin, essential for single-photon detection and responsivity in dim light.124,265-267
Given the critical role of these timely interactions for rod function, extensive research has
been carried out to uncover binding interfaces and key amino acids influencing rhodopsin

65
signaling and its downstream effectors. An exceptionally conserved stretch of residues, the ERY
motif (Glu134-Arg135-Try136),268 located at the cytoplasmic end of transmembrane domain 3 (TM3),
has received considerable attention. Recent advances in computational and structural biology
have illuminated the crucial role of the ERY motif in governing vertebrate rhodopsin
conformation269-271 and its coupling/recognition of cognate proteins.68,272-275 This insight is
reinforced by earlier findings from numerous biochemical studies, specifically targeting the
charged Glu134-Arg135 (ER) pair. Mutations to the ERY domain exerted notable effects, influencing
the coupling of R* to transducin,
276-279 R* phosphorylation by GRK1,
278,280,281 and possibly, arrestin
binding.
278 Additionally, specific mutations promote constitutive activation of rhodopsin,269,282,283
by disruption of the salt bridge, or ionic lock, between the charged side chain Glu247 on TM6 and
the charged ER pair, thereby destabilizing the dark-state rhodopsin. These findings highlight the
highly dynamic nature of interactions involving the ERY motif for rhodopsin function in the
phototransduction cascade. Despite extensive exploration of the 3-D structure and molecular
interactions with conjugate proteins, our understanding of how transducin, GRK1, and arrestin
interact with the ERY motif in-vivo and within the proper cellular micro-environment remains
incomplete.
A well-explored rhodopsin mutation involves the charge reversal of ERY to REY (Arg134-
Glu135-Try136), functionally uncoupling R* from transducin activation while preserving REYrhodopsin (REY-Rho) absorption properties.276,277,284 To access the consequences of ERY →
REY in-vivo, a knock-in mouse line (denote RhoREY/REY, or REY) was recently generated.67 As
anticipated, REY rods were highly insensitive to light, ~16,000-fold less sensitive than wildtype
counterparts,67,285 validating previous observations of reduced coupling between REY-Rho* and
transducin. In stark contrast to rat retinas ectopically expressing an ERY mutant (R135L)280,286,
which exhibited hyperphosphorylation and rapid cell death, REY retinas did not degenerate and
maintained normal light absorption properties and expression of essential signaling proteins.67
Notably, time-averaged photoresponses from REY rods displayed slower deactivation kinetics,

66
likely a consequence of REY-Rho*’s temporally-distributed activation of transducin.
67 However,
whether this phenomenon can be attributed to a delay, or impairment, in REY-Rho*
phosphorylation and arrestin binding remains unclear.
Rod photoreceptors, beyond their function in photon-to-signal conversion, exhibit a
phenomenon involving light-induced movement of key signaling molecules within the polarized
rod cell.114,129 In darkness, transducin is sequestered to the OS disc membranes due to its high
affinity for membranes, driven by lipid modifications on T� and T� subunits.287,288 Conversely,
ARR1 resides predominately in the inner segment (IS), anchored by its affinity for
microtubules112,118,289 and self-oligomerization status.
290 Upon light exposure, activated transducin
separates into T�-GTP and T�� subunits, significantly reducing their membrane affinities,
291
which ultimately results in their translocation to the IS and synaptic terminal (SN).
111 ARR1, on
the other hand, moves to the OS, driven entirely by its substantially higher affinity for R* and
phosphorylated rhodopsin (R*-P).112,125 This light-dependent redistribution of the signaling
proteins only occurs once the light intensity exceeds a critical threshold,121,125 serving as a
potential mechanism for rod survival292 and adaptation to brighter lighting conditions.
111,123,124
Despite the pivotal role played by this bidirectional energy-independent movement of transducin
and ARR1,
112,120 establishing a direct link between mutations affecting the highly conserved ER
charged pair and protein translocation remains challenging, primarily due to the limited availability
of mouse lines.
Using the RhoREY/REY mouse line, we address this research gap by examining the
interaction of rhodopsin’s ER tandem pair with cognate proteins in rod photoreceptors across
varying lighting conditions. Leveraging the non-degenerative nature of REY retinas, we probe the
effects of the ERY → REY mutation on R* signaling, R* phosphorylation, and light-activated
protein translocation. Our findings confirm that the REY mutation significantly reduces rod
sensitivity to light, resulting in marked impairment of T�-GTP translocation under low light (20-
lux) and bright light (5,000-lux) conditions. Of note, this mutation has no discernible effect on R*

67
phosphorylation by GRK1 or the light-dependent subcellular movement of arrestin, a stark
departure from findings in heterologous REY-Rho expression studies. Unexpectedly, the ERY →
REY mutation impacts the localization of transducin in darkness, with a significant amount
situated in the IS. This observation hints at the existence of a stabilizing dark-state
rhodopsin/transducin complex that ensures rapid cascade activation and aids in sequestering
transducin to the OS discs in the dark.
3.3 Methods
Ethics statement
All animal experiments were approved by the University of Southern California Institutional Animal
Care and Use Committee.
Mouse breeding and husbandry
Animals were handled in accordance with the recommendations provided in the Guide for the
Care and Use of Laboratory Animals. Non-breeder adult male and female mice aged 2-3 months
were used in this study. Mice were housed under cyclic light (12h/12h light/dark) in transparent
cages with adlib food and water and were dark-adapted overnight before experiments. Lightinsensitive RhoREY/REY (REY) mice were obtained from the laboratory of Dr. King-Wai Yau, John
Hopkins University School of Medicine. This mouse line was generated using the CRISPR/Cas
system in the Gcaps+/- background and was backcrossed to C57BL/6 mice.67 Mouse genotypes
were determined by Transnetyx, Inc., and information is available upon request. All mice were
euthanized by isoflurane inhalation followed by cervical dislocation.

68
Light exposure
Mice were subjected to two different illumination levels, 20-lux (low light) or 5,000-lux (bright light),
for specified durations. Before exposure to light, mice were anesthetized by intraperitoneal
injection of a xylazine (10 μg/g bodyweight) and ketamine (100 μg/g bodyweight) cocktail diluted
in 1X PBS. Pupillary dilation was achieved in both eyes by adding both 0.5% tropicamide and
2.5% phenylephrine hydrochloride solutions. To maintain corneal hydration, a hydroxylpropyl
methycellulose solution was applied to both eyes. The dim, 20-lux illumination was achieved
through LED light bars (Commercial Electric, 500 lumens, color temperature 2700K, #54195201)
combined with neutral density filters (B&H #210 ND 0.6 and #209 ND 0.3 filters). The 5,000-lux
lightbox enclosure utilized fluorescent lighting (B&H #POLB1118L2, Lithonia lighting #676883).
Body temperature was maintained during light exposure by heating pads.
Estimation of luminance
Facilitating comparisons with electroretinogram (ERG) lighting luminance values involved
transforming photopic lux measurements (captured by a Sekonic Speedmaster L-858D-U Light
Meter) into approximate scotopic cd/m2 values. The conversion followed the precedent set by
Lyubarsky et al.,293 wherein 1 photopic lux (at 555 nm) equates to 0.64 scotopic lux (at 507 nm)
for incandescent light (color temperature 2700 K). The conversion was considered reasonably
accurate for our set-up, given that the LED light bar shares the same color temperature (2700 K)
as the fluorescent lighting. Utilizing the equivalence 1 lux = 1 cd・sr/m2
, the expression for scotopic
lux becomes 1 lux = 0.64 scotopic cd・sr /m2
. To convert from cd・sr/m2 to cd/m2
, we employ the
formula: 1 sr = 2π (1-cos(θ/2)), where θ is the solid angle subtended by the light source (sr,
steradian). Using θ = π for the geometry of the light bar, we arrive at 1 photopic lux/sr ~ 0.1
scotopic cd/m2
. Note the dependence of these conversions on the differing geometries of the
Ganzfeld stimulus (θ=120°) and the light bar (θ=180°).

69
Electroretinogram (ERG)
3-month-old RhoREY/REY and C57BL/6 mice were dark-adapted overnight, and all procedures
conducted on the recording day were performed under dim red light. Animals were anesthetized
by intraperitoneal injection of a xylazine (10 μg/g bodyweight) and ketamine (100 μg/g
bodyweight) solution diluted in 1X PBS. Tropicamide (0.5%) and phenylephrine hydrochloride
(2.5%) solutions were added to dilate the pupil of the right eye. To keep the eyes hydrated,
hydroxylpropyl methycellulose solution was added to both eyes. A reference electrode was placed
subcutaneously near the right eye, and the OcuScience HMsERG LAB System delivered a
sequence of flashes from ranging from 0.1 mcd to 25 cd. The resulting ERGs were captured and
amplified by an AC/DC differential amplifier and filtered for analysis. OcuScience ERG Viewer
software facilitated the extraction of scotopic a-wave and b-wave amplitudes.
Isoelectric focusing (IEF)
Mice were dark-adapted overnight before assessing rhodopsin phosphorylation levels under two
conditions: (1) dark-adapted or (2) low light exposure (20-lux, 5 min). The retinae were isolated
either under infrared light or immediately after low light exposure, frozen in liquid nitrogen, doublewrapped in aluminum foil, and stored at -80˚C until further processing. IEF was performed
following established protocols.192,294 Briefly, frozen retinae were homogenized with a Polytron in
Buffer A (25 mM HEPES pH 7.5, 100 mM EDTA, 50 mM NaF, 5 mM Adenosine, 0.1 M PMSF and
complete mini protease inhibitor (Roche Applied Sciences)) and were centrifuged for 15 min
(19,000 x g). The resulting pellet was washed in 10 mM HEPES (pH 7.5) and resuspended in
Buffer B (10 mM HEPES pH 7.5, 0.1 mM EDTA, 50 mM NaF, 5 mM Adenosine, 1 mM MgCl2, 2%
(v/v) BSA, 0.1 M PMSF and complete mini protease inhibitor), supplemented with an excess of
11-cis-retinal (~1,200 pmol). The sample was light-protected and rocked overnight at 4˚C. The
following day the samples were pelleted by centrifugation, washed, and solubilized for 3h at 4˚C
in Buffer C (20 mM HEPES pH 7.5, 0.1 mM EDTA, 50 mM NaF, 5 mM Adenosine, 1 mM MgCl2,

70
10 mM NaCl, 1% (w/v) n-dodecyl-�-D-maltoside (DDM), 1 mM dithiothreitol (DTT), 0.1 M PMSF
and complete mini protease inhibitor). After centrifugation, glycerol was added to the supernatant
to achieve a 50% final concentration. Samples were loaded onto a pre-focused acrylamide gel
(40% (v/v) acrylamide, 1% (w/v) DDM, 13.33% (v/v) glycerol, 3.8% (v/v) Pharmalyte pH 2.5-5 and
2.5% (v/v) Pharmalyte 5-8 (GE Healthcare)) positioned on a Multiphore II Electrophoresis Unit
(Amersham Pharmacia). The gel was focused for 2 h at 23 W constant power and blotted for 45
min onto a nitrocellulose membrane (Cytiva AmershamTM ProtranTM, pore size 0.2 µm, 45004011).
The membrane was incubated overnight at 4˚C with the mouse monoclonal antibody against
rhodopsin’s N-terminus (R2-12N, 1:5,000, a kind gift from Dr. P.A. Hargrave, University of Florida)
followed by an incubation with a fluorescently labeled secondary antibody (1:10,000, 926-31081,
LI-COR). The bands were detected by the Odyssey CLx system (LI-COR), and the intensity of
the bands was quantified using ImageJ198 following established methods.192,294
Immunocytochemistry
Eyes were harvested either after dark adaptation or immediately following exposure to a specific
lighting condition (20-lux or 5,000-lux) for 30 min. Before enucleation, the superior pole of the
cornea was cauterized for orientation, and the eyes were fixed in 4% formaldehyde in 1X PBS for
5 min. The eyes were dissected into eye cups (cornea, lens, and vitreous removed) and then
further fixed for an additional 1h. The eye cups underwent a series of rinses in 1X PBS followed
by 0.1 M cacodylate buffer (pH 7.2). The samples were cryoprotected in 30% sucrose in 0.1M
cacodylate buffer (pH 7.2) overnight at 4˚C, embedded in Tissue-TeckÒ O.C.T. compound, and
flash-frozen in a dry ice/ethanol slurry. Retinal sections of 10 µm thickness were obtained using
a cryostat (CM 3050 S, Leica Microsystems). The slides were evaluated with the aim of capturing
sections representing a specific region of the retina (proximate to the optic nerve). Slides meeting
this criterion then underwent a 1h blocking step (10% donkey serum, 0.3% Triton X-100 in 1X
PBS). Using a hydrophobic barrier pen, each slide was divided into two regions, enabling the

71
independent treatment of neighboring sections with rabbit polyclonal antibodies against rod
arrestin (C10C10,295,296 1:600) and transducin (GNAT1, 1:2,000, Invitrogen, #PA5,28336). The
sections were rinsed and incubated with anti-rabbit AlexaFluorÒ 488 secondary antibody (1:400,
Jackson Immuno Research Laboratories). All sections were double-stained using the biotinylated
antibody against rhodopsin (1D4,297 1:300), rinsed, and incubated with rhodamine-labeled Avidin
D (1:200, Vector Laboratories) and DAPI (1:5,000). Retinal sections were inspected using a
20X/0.8NA air objective and were acquired with a Zeiss LSM800 confocal microscope (Carl Zeiss
Meditec, Jena, Germany).
Western blot
Mice were kept in darkness or were exposed to 15 min of either 20-lux or 5,000-lux illumination.
After euthanasia, eyes were immediately harvested, and retinal samples were prepared quickly
(under 5 min per mouse) in a darkroom under infrared light using a dissecting microscope fitted
with infrared converters (TNV/MOD3 Bravo monoculars, Tactical Night Vision Company, USA).
Samples were prepared following the lyophilized peeling technique outlined by Rose et al. (Figure
S3.1).298,299 Briefly, dissected retinas were hemisected in cold Ringer’s buffer (130 mM NaCl,
3.6 mM KCl, 2.4 mM MgCl2, 1.2 mM CaCl2, 10 mM HEPES, 0.02 mM EDTA, pH 7.4), trimmed
into two flat rectangles, and carefully placed on 5 x 2.5 mm filter paper pieces (WhatmanÒ Grade
1 filter paper, GE Health, USA). The retinal ganglion cell layer was positioned to face the filter
paper, ensuring direct contact between them. The samples were dried, light protected, frozen in
liquid nitrogen, and lyophilized for 30 min (FreeZone 4.5L Benchtop Freeze Dry System,
Labconco, MO, USA). The freeze-dried samples were peeled with ScotchTM tape to obtain distinct
fractions: +ROS, +RIS, and -ROS/RIS-depleted tissue (-OIS). The resultant fractions were
homogenized in RIPA lysis buffer (50 mM Tris pH 8, 0.1% SDS, 150 mM NaCl, 0.5% NP40, 0.5%
deoxycholate acid) supplemented with 0.1 mM phenylmethanesulfonyl fluoride, 0.02 mM

72
Aprotinin, and 0.02 mM Leupeptin. After a 30 min incubation at room temperature with DNase I
(Roche), the total protein content of each sample was determined using the PierceTM BCA Protein
Assay Kit (#23227). Equal amounts of protein from each sample were electrophoresed on a 4-
12% Bis-Tris SDS-PAGE gel (Invitrogen). The separated proteins were then transferred onto a
nitrocellulose membrane (Cytiva AmershamTM ProtranTM, pore size 0.2 µm, 45004011) and
incubated overnight at 4˚C with the desired primary antibodies: rabbit anti-arrestin (C10C10,295,296
1:5,000), rabbit anti-transducin (GNAT1, 1:3,000, Invitrogen, #PA5,28336), and rabbit antiG�5L/S (CT215,300 1:2,000). Fluorescent-labeled secondary antibodies (1:10,000, 926-31081, LICOR) and the Odyssey CLx system (LI-COR) were used to visualize the signals. Density analysis
of the individual bands was performed using Fiji200 and a modified gel sum normalization method
adapted from Degasperi et al.301 was used. For each independent experiment, the fluorescent
signals for the proteins of interest in each compartment were normalized against the combined
signals from all retinal compartments.
Rhodopsin concentration
Procedures were performed under dim red light. Mice were euthanized in either complete
darkness or after the right eye was exposed to 20-lux for 15 min. The retinae were then extracted
through a slit in the cornea302 and solubilized in 1X PBS containing 1% n-dodecyl-b-Dmaltopyranoside (850520P, Avanti Polar Lipids), 0.1 M PMSF and complete mini protease
inhibitor (Roche Applied Sciences). The samples were light-protected and stored at 4˚C overnight.
Samples were centrifuged at 4,000 rpm for 3 min, and the dark absorption spectra of the
supernatant were determined between 270-700 nm using a UV/visible spectrophotometer
(DU640 Beckman Coulter). Rhodopsin was completely bleached with a white LED light for 5 min,
after which another scan was performed to capture the post-bleach spectra. With the estimated
REY-mutant molar extinction coefficient (40,4000 ± 1,400 M-1
cm-1
)
67 closely resembling that of
wildtype rhodopsin (40,600 M-1
cm-1
), the molar extinction coefficient for wildtype rhodopsin was

73
used in the calculation of rhodopsin concentration. This determination was derived from the
difference spectra at 500 nm (subtracting the post-bleach from the pre-bleach) using Beer’s Law.
The spectra were zeroed at 700 nm and normalized to the estimate of the total protein
concentration at 280 nm.
Image analysis
Fluorescence images of fixed retinal slices were analyzed using Fiji200, and quantification of the
relative amount of arrestin and transducin in the ROS and inner compartments followed an
established method.120 Briefly, a portion of an image was selected using the rectangle tool to
include the entire length of a population of photoreceptor cells (spanning from ROS to ST). This
selection represents the total fluorescence (Ft) for a specific antibody within these rods. A second
rectangle was introduced within the same area to exclude the ROS, capturing the fluorescence
only in the RIS, CB, and ST (Fi). The integrated fluorescent intensity signals from both areas (Ft
and Fi) were measured, background fluorescence was subtracted, and fluorescent intensity
values were normalized to account for differences between independent experiments. To address
intensity variations across experiments, Fi/Ft ratios were determined. Additionally, an equation for
area normalization was formulated to accommodate differences in rectangle sizes arising from
varying rod lengths. This equation is as follows: %&
!"#
#
!"$
$
' ()
!"Ā
Ā * + 1-
"!
, where f is the
fluorescence ratio Fi/Ft, A denotes the ratio of the RIS/CB/ST area to the total area
(ROS/RIS/CB/ST), and Ā represents the average of the area ratios A across different samples.
Three distinct rectangular areas were selected within each image using Fiji200, and the mean Fi/Ft
ratios ± SD were calculated following normalization.

74
Statistical analysis
Initial studies revealed no gender-specific differences, therefore the data from both genders were
pooled for further analysis. Unless otherwise noted, the data are presented as mean ± standard
deviation, with the number of mice (N) indicated in the figure legends. GraphPad Prism 10.0.0 for
Mac and SPSS for Mac, version 29 (IBM Corp., Armonk, N.Y., USA) were used to perform
statistical analysis and generate plots. Normality and homogeneity of variance were assessed
using the Shapiro-Wilk and the Levene’s test, respectively. When data was normally distributed,
either an unpaired Welch’s t-test (pairwise comparisons), one-way ANOVA followed by a post hoc
Tukey test (multiple comparisons), or a two-way ANOVA with “lighting condition” and “genotype”
as between-group factors followed by a Šídák post hoc test (multiple comparisons) were
employed. Non-parametric statistical comparisons were made using Kruskal-Wallis one-way
ANOVA followed by a Dunn’s Multiple Comparison Test. Statistical significance was set at � =
0.05 for all tests, and p ≤ 0.05 was considered statistically significant.
3.4 Results
Mutant REY-rhodopsin are phosphorylated normally by rhodopsin kinase (GRK1)
Previous research concluded that rods from RhoREY/REY (REY) mice exhibit no discernable
structural or downstream functional consequences, except for a significant reduction in coupling
between light-activated REY-rhodopsin (REY-Rho*) and transducin.67 This disrupted coupling
renders these rods highly insensitive to light, as demonstrated by the absence of detectable
responses in suction electrode recordings, even with flash strengths up to ~10,000 photons•µm2
,
67 a level of illumination that typically saturates wildtype (WT) rods.303,304 To verify the attenuated
REY-Rho* signaling and determine the optimal white-light illumination for activating REY-Rho, we
performed in-vivo electroretinograms (ERGs) under scotopic conditions (Figure 3.1). ERG
recordings from age-matched 2-month-old WT mice exhibited the typical biphasic wave pattern

75
in response to increasing light stimuli, whereas both scotopic a- and b-waves from REY animals
were drastically diminished (Figure 3.1A). Normalized intensity-response functions (Figure 3.1B)
further demonstrate the significant reduction in REY a- and b-wave amplitudes for all illuminations
(p ≤ 0.05). These findings align with earlier observations indicating a reduced coupling between
REY-Rho* and transducin, thereby contributing to the overall light insensitivity.67,285 Based on the
response-intensity profiles, we found that white-light illumination between scotopic 1-3 cd*s/m2
(~20-lux in our experimental set-up, see Methods) determines the threshold at which REY-Rho is
activated.
Figure 3.1. Effect of charge reversal (Arg134Glu135, RE) within the conserved G-protein binding motif
of rhodopsin on retinal function in-vivo. (A) Scotopic ERG responses from 2-month-old wildtype (WT,
black) and REY (red) mice (N = 3-4). ERG traces depict a series of white light flash intensities ranging from
weakest (10-300mcd*s/m2) to strongest (1-25cd*s/m2). (B) Intensity-response data for the maximal a- and
b-wave amplitudes of the same mice shown in (A). Amplitudes of rod cell-evoked scotopic a- and b-waves
were reduced in REY mice (red) compared to age-matched WT mice (black).
We next sought to examine the extent to which the charge reversal of ERY → REY
impacted rhodopsin phosphorylation (R*-P) in-vivo that electrophysiological recordings couldn’t
conclusively determine.
67 Age-matched WT and REY mice were subjected to a 5 min exposure
of 20-lux light, and then rhodopsin phosphorylation levels were assessed by isoelectric focusing
-3 -2 -1 0 1 2
0
100
200
300
400
WT
REY
-3 -2 -1 0 1 2
0
100
200
300
400
500
600
700
WT
REY
New_b-wave_points
25cd*
WT
WT
WT
REY
REY
REY
REY
WT
10cd*
WT
WT
WT
REY
REY
REY
REY
WT
3cd*
WT
WT
WT
REY
REY
REY
REY
WT
1cd*
WT
WT
WT
REY
REY
REY
REY
WT
300mcd*
WT
WT
WT
REY
REY
REY
REY
100mcd*
WT
WT
WT
REY
REY
REY
REY
30mcd*
WT
WT
WT
REY
REY
REY
REY
WT
10mcd*
WT
WT
WT
REY
REY
REY
REY
30mcd*
WT
WT
WT
REY
REY
REY
REY
WT
25cd*
WT
WT
WT
REY
REY
REY
REY
WT
1cd*
WT
WT
WT
REY
REY
REY
REY
WT
300
100
30
10
mcd*s/m2
WT
REY
cd*s/m2
25
10
3
1
A B
a-wave
Amplitude (µV)
b-wave
Amplitude (µV)
100ms
100µV
100ms
200µV
Intensity (log cd*s/m2)
Intensity (log cd*s/m2)

76
(IEF), a technique that clearly resolves the differently phosphorylated species of rhodopsin (0-6)
based on their isoelectric points. Dark- and light-exposed WT rhodopsin (WT-Rho) and REY-Rho
were separated according to their unique isoelectric points and, unexpectedly, exhibited
comparable levels of phosphorylation (Figure 3.2A). Subsequent analysis of the distribution of the
phosphorylated species for both WT-Rho and REY-Rho revealed no significant differences
(Figure 3.2B, p > 0.05), suggesting comparable GRK1 activity. Collectively, our results reinforce
the critical role of the ERY motif in R* interaction with transducin in-vivo; however, the ERY motif
does not appear to be involved in the binding and phosphorylation of R* by GRK1 in intact rods.
Figure 3.2. Phosphorylation of light activated rhodopsin (R*) by rhodopsin kinase (GRK1) is
unchanged in REY retinae. (A) The status of phosphorylation was examined in 3-month-old WT (N = 8)
A
B
20-lux dark 20-lux dark
WT REY
0P
1P
2P
3P
4P
5P
6P
*
*
*
0P 1P 2P 3P 4P 5P 6P
0.0
0.1
0.2
0.3
0.4
0.5
Fraction of P-rhodopsin
WT
REY
ns REY
WT
0P 1P 2P 3P 4P 5P 6P
0.0
0.1
0.2
0.3
0.4
0.5
Fraction of P-rhodopsin
WT
REY
0P 1P 2P 3P 4P 5P 6P
0.0
0.1
0.2
0.3
0.4
0.5
Fraction of P-rhodopsin
WT
REY

77
and REY (N = 7) retinae using IEF. Retinas were either isolated in darkness or immediately after 5 mins of
20-lux light exposure. The representative IEF gel illustrates the segregation of different rhodopsin species
based on the total number of attached phosphate groups (mouse rhodopsin has 6 putative phosphorylation
sites at its C-terminus), which is specified on the left side of the gel. The charge reversal of ERY→REY
altered the isoelectric point (pI) of REY rhodopsin, impacting the separation of phosphorylated rhodopsin
species. Asterisks demarcate bands arising from rhodopsin’s interaction with dodecyl-maltoside impurities.
(B) Band intensities from IEF gels were quantified to determine the fraction of phosphorylated rhodopsin in
retinal samples from WT (white circles, N = 8) and REY (red circles, N = 7) mice exposed to darkness or
20-lux light. Densities from the phosphorylated species (1-6P) were summed and divided by all rhodopsin
species (0-6P). This ratio established the fraction of phosphorylated rhodopsin in that retinal sample. The
distribution of P-rhodopsin (phosphorylated rhodopsin) for WT was statistically indistinguishable from REY
(Welch’s t-test, p > 0.05).
REY mutation impairs transducin localization without affecting arrestin translocation
Considerable research efforts have been directed to unraveling the mechanisms, timing,
and lighting thresholds that govern the reciprocal light-induced movement of transducin and
arrestin (ARR1) between inner and outer segments of rod cells.111,114,130,132 Given the importance
of ARR1 binding to R*-P for recovery kinetics,
108 alongside the presumed involvement of the ERY
motif in the interaction of rhodopsin with arrestin (ARR1)195,305,306 and transducin,67,111 we next
focused on examining the impact of the REY charge reversal on the translocation of both ARR1
and the �-subunit of transducin (T�-GTP, GNAT1). Consistent with the light-insensitive nature of
REY retinas, GNAT1’s immunoreactivity in retinal slices following 30 mins of sustained exposure
to 20-lux and 5,000-lux, was largely confined to the rod outer segments (ROS), with weak labeling
observed in the rod inner segments (RIS, Figure 3.3A). Faint signals were also detected in the
cell bodies (CBs) and synaptic terminals (SNs) of retinal slices exposed to 5,000-lux, though most
of the staining was detected in the ROS (Figure 3.3A). In age-matched WT mouse retinae, GNAT1
immunoreactivity was observed in the RIS, CB, and ST, with noticeably stronger fluorescent
signal in these compartments under 5,000-lux compared to 20-lux illumination conditions (Figure
3.3A), consistent with previous reports.130,307 Surprisingly, following overnight dark adaptation,
GNAT1 immunolabeling consistently exhibited a moderate signal in the RIS of REY retinas,
whereas the signal observed in WT retinae was predominantly concentrated in the ROS (Figure
3.3A). On the other hand, the immunostaining of retinal cross-sections from dark-adapted and

78
light-exposed (20-lux and 5,0000-lux) REY and WT mice showed a similar ARR1 fluorescence
pattern (Figure 3.3A), where the signal transitions from the cytoplasmic space of the RIS, CB, and
ST (in darkness) to being detected in the ROS (5,000-lux). In the 20-lux light-exposed retinas
stained with antibodies targeting ARR1, we confirmed, through peanut agglutinin (PNA) staining,
that the bright signal originating from cone-like structures was from genuine cones308 (Figure
S3.2). This staining pattern may be attributed to the movement of ARR1 from the inner
compartments of rods towards the ROS, thereby accentuating the prevalence of ARR1
expression in cones.
Next, we evaluated the redistribution of GNAT1 and ARR1 under different lighting
conditions (dark, 20-lux, and 5,000-lux), by quantifying their relative immunofluorescence
intensities. This involved calculating the ratio of fluorescence intensities Fi/Ft, with Fi denoting the
signal in the RIS/CB/ST, and Ft denoting the signal in all rod compartments (Figure 3.3B). Our
results confirmed a significantly elevated GNAT1 signal in the inner compartments (RIS/CB/ST)
of REY retinae in darkness compared to WT (Figure 3.3C, p ≤ 0.001). Additionally, we found a
notable reduction in GNAT1 translocation to the inner compartments in both the 20-lux and 5,000-
lux compared to WT (p ≤ 0.05). Conversely, ARR1 relative fluorescence was indistinguishable
between REY and WT retinae across all conditions (p > 0.1); however, a robust translocation from
the inner compartments to the ROS was not observed, possibly due to obstructed access of the
antibody to its epitope. Together, these data show that the localization and light-induced
translocation of GNAT is compromised in the light-insensitive REY rods, while ARR1 remains
unaffected.

79
Figure 3.3. Impact of REY mutation on the translocation of transducin and arrestin between rod
compartments. (A) Representative confocal images of retinal cross-sections from 3-month-old WT (N = 4)
and REY (N = 4) mice isolated either in darkness or following 30 mins of light exposure at intensities of 20-
lux and 5,000-lux. Cryosections were probed with antibodies against GNAT1 (transducin, gray), ARR1
(arrestin, yellow), and rhodopsin (red). ERY→REY charge reversal does not impact retinal morphology and
localization of rhodopsin to the ROS in REY rods. Scale bar = 10 µm. (B) Schematic depicting the selection
process for regions of interest (ROIs) within rod photoreceptor cells, delineating either the inner
compartments (RIS, CB, ST; Fi, red box) or the entire rod (ROS, RIS, CB, ST; Ft, blue box). The ratio of
inner compartment fluorescence (Fi) to total fluorescence (Ft) was used to determine the relative distribution
(Fi/Ft) of transducin (gray) and arrestin (yellow) in a population of WT and REY rods before and after light
exposure. Rod diagram was created with BioRender.com. (C) Graphs summarizing relative distribution
(Fi/Ft) of transducin (GNAT1) and arrestin (ARR1) within a population of rods from WT (open circles, N = 4)
and REY (red circles, N = 4) mice exposed to either darkness (dark gray), 30 mins 20-lux (light gray), or 30
mins 5,000-lux (yellow). GNAT1 labeling is elevated in the inner compartments in dark-adapted REY rods
compared to WT (two-way ANOVA with Šídák post hoc test, *p ≤ 0.001). After 30 mins 20-lux or 5,000-lux
ARR1
5,000-lux 20-lux Dark
WT REY
WT
WT dark
REY dark
WT 20-lux 30min
REY 20-lux 30min
WT lightbox 30 min
REY Lightbox 30min
0
10
20
30
40
50
Relative flourescence in IS/CB/ST
as a percent of total (Fi/Ft)
WT dark
REY dark
WT 20-lux 30min
REY 20-lux 30min
WT lightbox 30 min
REY Lightbox 30min
WT dark
REY dark
WT 40 lux 30min
REY 40 lux 30min
WT Lightbox 30min
REY Lightbox 30min
0
20
40
60
80
100
Relative flourescence in IS/CB/ST
as a percent of total (Fi/Ft)
WT dark
REY dark
WT 40 lux 30min
REY 40 lux 30min
WT Lightbox 30min
REY Lightbox 30min
A
B
ns
*
*
*
GNAT1
Percentage of total relative
fluorescence (Fi
/Ft
)
Percentage of total relative
fluorescence (F/
i Ft
)
C
Ft
Fi
WT REY
GNAT1 ARR1
ROS
RIS
CB
SN
ROS
RIS
CB
SN
ROS
RIS
CB
SN
ROS
RIS
CB
SN WT dark
REY dark
WT 40 lux 30min
REY 40 lux 30min
WT Lightbox 30min
REY Lightbox 30min
0
20
40
60
80
100
Relative flourescence in IS/CB/ST
as a percent of total (Fi/Ft)
WT dark
REY dark
WT 40 lux 30min
REY 40 lux 30min
WT Lightbox 30min
REY Lightbox
30min
REY
20-lux 5,000-lux WT dark
REY dark
WT 20-lux 30min
REY 20-lux 30min
WT lightbox 30 min
REY Lightbox 30min
0
50
40
30
20
10
Relative flourescence in IS/CB/ST
as a percent of total (Fi/Ft)
WT dark
REY dark
WT 20-lux 30min
REY 20-lux 30min
WT lightbox 30 min
REY Lightbox 30min
Dark WT dark
REY dark
WT 20-lux 30min
REY 20-lux 30min
WT lightbox 30 min
REY Lightbox 30min
0
50
40
30
20
10
Relative flourescence in IS/CB/ST
as a percent of total (Fi/Ft)
WT dark
REY dark
WT 20-lux 30min
REY 20-lux 30min
WT lightbox 30 min
REY Lightbox 30min
WT dark
REY dark
WT 20-lux 30min
REY 20-lux 30min
WT lightbox 30 min
REY Lightbox 30min
0
50
40
30
20
10
Relative flourescence in IS/CB/ST
as a percent of total (Fi/Ft)
WT dark
REY dark
WT 20-lux 30min
REY 20-lux 30min
WT lightbox 30 min
REY Lightbox 30min
20-lux 5,000-lux WT dark
REY dark
WT 20-lux 30min
REY 20-lux 30min
WT lightbox 30 min
REY Lightbox 30min
0
50
40
30
20
10
Relative flourescence in IS/CB/ST
as a percent of total (Fi/Ft)
WT dark
REY dark
WT 20-lux 30min
REY 20-lux 30min
WT lightbox 30 min
REY Lightbox 30min
WT dark
REY dark
WT 20-lux 30min
REY 20-lux 30min
WT lightbox 30 min
REY Lightbox 30min
0
50
40
30
20
10
Relative flourescence in IS/CB/ST
as a percent of total (Fi/Ft)
WT dark
REY dark
WT 20-lux 30min
REY 20-lux 30min
WT lightbox 30 min
REY Lightbox 30min
WT dark
REY dark
WT 20-lux 30min
REY 20-lux 30min
WT lightbox 30 min
REY Lightbox 30min
0
50
40
30
20
10
Relative flourescence in IS/CB/ST
as a percent of total (Fi/Ft)
WT dark
REY dark
WT 20-lux 30min
REY 20-lux 30min
WT lightbox 30 min
REY Lightbox 30min
Dark

80
light exposure, GNAT1 fluorescence in the inner compartments (Fi) of REY rods is significantly lower than
WT rods (two-way ANOVA with Šídák post hoc test, *p ≤ 0.05). No significant differences in ARR1
localization in the three lighting conditions were observed between WT and REY rods (two-way ANOVA
with Šídák post hoc test, p > 0.1). ROS, rod outer segment; RIS, rod inner segment; CB, cell body; SN,
synaptic terminal.
The quantification of immunofluorescence signals for understanding protein distribution
poses inherent challenges, including susceptibility to photobleaching, staining variability, and the
potential impact of epitope masking artifacts.309 To better resolve ARR1 and GNAT1 movement
within the individual rod compartments, lyophilized retinas from WT and REY mice underwent a
sequential peeling process, yielding +ROS, +RIS, and -OIS (without outer and inner segment)
enriched fractions.298,299 These isolated fractions, retrieved either in darkness, or after 20-lux or
5,000-lux exposure (15 min), were subjected to western blotting, with a representative blot shown
in Figure 3.4A. A whole retinal sample served as a control, and G�5L/G�5S served as a quality
control for ROS purity.
294,310 G�5L, a part of the transducin GAP complex,
88,311 is predominantly
found in the ROS, whereas its shorter isoform, G�5S, is primarily expressed in other retinal
cells.
300 In agreement with our retinal staining results (Figure 3.3A), ARR1 and GNAT1 revealed
similar findings in regard to protein subcellular distributions across all REY and WT samples
(Figure 3.4A). Notably, ARR1 exhibited a clearer shift from the inner compartments (+RIS and -
OIS) to the ROS in response to both 20-lux and 5,000-lux light exposure, suggesting our
immunofluorescence data may not have fully captured the extent of ARR1’s translocation.

81
Figure 3.4. Quantification of transducin and arrestin translocation in REY retinae. (A) Representative
western blots of isolated retinal layers from dark- and light-exposed (5,000-lux and 20-lux, 15 mins) retinas
from WT and REY mice (N = 4-6 per condition). G�5L/G�5S served as a loading control. Molecular weight
(kDa) is provided on the left side of the first blot. Whole retinal homogenate (labeled ‘Retina’) is included
on each blot. GNAT1 (transducin); C10C10 (arrestin); G�5L/G�5S (G protein subunit G�5 long and short
isoforms). +ROS, rod outer segment only; +RIS, rod inner segment only; -OIS, without outer/inner
segments. (B-D) Signals derived from GNAT1 (top) and ARR1 (bottom) in each rod compartment were
quantified, normalized, and plotted. Isolated +ROS, +RIS, and -OIS fractions were acquired from WT (open
circles) and REY (red circles) mice (N = 4-6 per condition) exposed to either (B) darkness (dark gray bar),
(C) 20-lux exposure (light gray bar), or (D) 5,000-lux exposure (light yellow bar). Dark-to-light-exposed
peeled WT samples (20-lux and 5,000-lux) exhibited typical translocation patterns for both GNAT1
(movement away from the ROS) and ARR1 (movement towards the ROS). REY retinas displayed impaired
GNAT translocation and localization. Significant differences were observed in the relative amount of GNAT1
in the RIS of dark-adapted REY rods compared to dark-adapted WT controls (B, top panel, Welch’s t-test,
*p = 0.03). Translocation of GNAT1 away from the ROS in response to 20-lux (Welch’s t-test, *p = 0.04)
and 5,000-lux (Welch’s t-test, *p = 0.017) was significantly reduced in REY samples compared to WT (C
and D, top panels). A significant difference was noted in the amount of GNAT1 translocation to the cell
body/synaptic terminal (-OIS) in the 20-lux lighting condition (C, top panel, Welch’s t-test, *p = 0.003). REY
ARR1 dark-to-light translocation for each lighting condition did not differ from the WT samples (B-D, bottom
panels, Welch’s t-tests, p > 0.05). Rod diagram was created with BioRender.com.
ROS RIS OIS
0.0
0.1
0.2
0.3
0.4
0.5
0.6
0.7 WT Dark
REY Dark
A
Retina
+ROS
+RIS
-OIS
+ROS
+RIS
-OIS
+ROS
+RIS
-OIS
WT WT REY
GNAT1
ARR1
G!5L
G!5S
Dark 5,000-lux 5,000-lux
REY WT REY
kDa Dark 20-lux 20-lux
48
40
44
39
* *
* *
WT dark
REY dark
WT 40 lux 30min
REY 40 lux 30min
WT Lightbox 30min
REY Lightbox 30min
0
20
40
60
80
100
Relative flourescence in IS/CB/ST
as a percent of total (Fi/Ft)
WT dark
REY dark
WT 40 lux 30min
REY 40 lux 30min
WT Lightbox 30min
REY Lightbox
30min
REY
WT
GNAT1 relative density
(% of total)
Retina
+ROS
+RIS
-OIS
+ROS
+RIS
-OIS
+ROS
+RIS
-OIS
B
+ + -
ROS RIS OIS
0.0
0.2
0.4
0.6
0.8
1.0 WT Dark
REY Dark
ROS RIS OIS
0.0
0.2
0.4
0.6
0.8
1.0 WT 20-lux
REY 20-lux
ROS RIS OIS
0.0
0.2
0.4
0.6
0.8
1.0
WT 5,000 lux
REY 5,000 lux
C
GNAT1 relative density
(% of total)
D
GNAT1 relative density
(% of total)
ARR1 relative density
(% of total)
ROS RIS OIS
0.0
0.1
0.2
0.3
0.4
0.5
0.6
0.7 WT 40lux
REY 100mcd
ARR1 relative density
(% of total)
ROS RIS OIS
0.0
0.1
0.2
0.3
0.4
0.5
0.6
0.7
WT 5,000 lux
REY 5,000 lux
ARR1 relative density
(% of total)
Dark 20-lux 5,000-lux
+ + -
+ + -
+ + - + + -
+ + -

82
We next measured the relative amount of GNAT1 and ARR1 present in the +ROS, -RIS,
and -OIS enriched fractions by densitometric analysis. In the dark-adapted state, the +ROS
fraction contained ~80% of the total GNAT1 present in rods for both WT and REY retinae. This
observation aligns with the expected high concentration of transducin in the ROS in
darkness.111,312-314 We also confirmed a substantial increase (~50%) in GNAT1 content within the
+RIS fraction from dark-adapted REY compared to its WT counterparts (Figure 3.4B, p = 0.03).
After a 15 min continuous exposure to dim, 20-lux light, which did not result in measurable
rhodopsin bleaching (Figure S3.3), we observed a clear relocation of GNAT1 from the +ROS
fraction to the inner compartments (+RIS and -OIS) in WT retinae (Figure 3.4C). This resulted in
significant differences in GNAT1 localization for both +ROS (p = 0.04) and -OIS (p = 0.003)
fractions (Figure 3.4C) between WT and REY samples. The absence of GNAT1 translocation in
REY mice under 20-lux light exposure (p > 0.1) is likely the contributing factor to this difference.
Under the 5,000-lux condition, GNAT1 translocation is even more pronounced in WT retinae, with
~55% of the GNAT1 in the ROS relocating to the inner compartments (Figure 3.4D). This
substantial reduction in GNAT1 within +ROS fraction greatly differs from REY (p = 0.017), despite
REY samples displaying some signs of movement from the +ROS to the +RIS fraction (Figure
3.4D). In addition, we confirmed no significant differences in ARR1 localization in darkness and
ARR1 translocation in response to light between WT and REY mice (Figures 3.4C and 3.4D).
Unexpectedly, the translocation of ARR1 to inner compartments in response to 5,000-lux was not
as robust (Figure 3.4D), and no discernible differences were observed between the translocation
profiles of REY and WT in both 20-lux and 5,000-lux light exposures (p > 0.2). From these data,
we reaffirm that the light-induced translocation of ARR1 remains unaffected in REY mice, and
that the critical threshold for GNAT1 translocation is shifted to higher intensities, likely attributed
to the low efficiency of REY-Rho* and transducin coupling. Thus, our in-vivo assessment indicates
that the ERY motif does not play a role in coordinating the interaction of ARR1 with R*-P. Instead,
our findings suggest that mutations in the ERY motif could potentially disrupt pre-formed

83
complexes between rhodopsin and transducin in ROS discs, leading to the localization of a
modest portion of GNAT1 in the RIS in darkness.
3.5 Discussion
The ERY motif, a highly conserved tripeptide sequence in class A GPCRs, functions as a
key element within rhodopsin, regulating its conformational states that mediate context-specific
interactions with its cognate proteins.271 Although extensive research has revealed the motif’s
involvement in transducin binding and activation,276,284 our understanding of the functional role of
the ERY domain within the native rod cell micro-environment has been limited until recently. The
RhoREY/REY mouse line provides additional evidence highlighting the disruptive impact of mutations
in this domain on the normal coupling between activated rhodopsin (R*) and transducin.67 Notably,
this mutation in rod cells translates into robust light-insensitivity. Here, we further explore the
functional consequences of the REY-Rho mutation in vivo, specifically examining its influence on
GRK1 interactions, arrestin affinity, and its potential impact on light-induced protein translocation
between subcellular compartments in rods. We show that 1) REY-Rho undergoes normal
phosphorylation by GRK1, 2) there is no change in arrestin (ARR1) binding affinity and
translocation, 3) light-induced translocation of transducin (GNAT1) from the ROS to inner
compartments is only observable under intense illumination (5,000-lux plus), and 4) remarkably,
a significant amount of transducin (~20% of total) is found in the RIS of REY rods in darkness.
The interaction between GRK1 and R* primarily involves rhodopsin’s second and third
cytoplasmic loops,315,316 as evidenced by a 50% reduction in GRK1 activation following cleavage
of the third loop.316 Previous reports using site-directed mutagenesis at the cytoplasmic end of
TM3 demonstrated that replacing the charged amino acids in the ERY motif with neutral
counterparts, or reversing the ER (-/+) charged pair to RE (+/-), led to either
hyperphosphorylation278,280,281,317, or reduced phosphorylation,281 respectively. In a recent
molecular dynamics simulation featuring a naturally occurring mutated rhodopsin (R135L), it was

84
observed that the hydrogen bond between Arg135 and rhodopsin’s C-terminus, particularly at
important phosphorylation sites Thr342 and Ser343, was disturbed. This disruption allowed the Cterminus tail to be more exposed, providing easier access to residues that are substrates for
GRK1.318 This structural finding is consistent with the in-vitro data, where neutral modifications at
the Arg135 residue result in hyperphosphorylation. Drawing on the aforementioned
structure/function findings, it is tempting to speculate that the ER → RE charge reversal might
influence REY-Rho’s structure, thereby leading to the observed reduced interaction with GRK1 in
the heterologous expression study.281 Lending indirect support to this hypothesis, in-vivo
investigations using REY mice also revealed slower recovery kinetics in intact rods.67 Contrary to
these reports, we found that REY-Rho* can bind and activate GRK1 normally in-vivo, resulting in
no identifiable differences in phosphorylation states between REY-Rho* and WT-Rho* (Figure
3.2). Thus, our findings suggest that the ER → RE charge reversal in rhodopsin’s ERY motif does
not appear to impact the orientation of residues crucial for recognizing and modulating GRK1
binding. This contradicts the notion that the ERY motif may be important in rhodopsin kinase
binding and activation, as proposed by previous modeling studies.68,275,318 Conversely, our data
supports the conclusions presented by Yue et al., who propose that the slow response kinetics
from REY rods is likely attributed to the temporally-distributed activation of transducin by REYRho*, rather than issues with GRK1 phosphorylation.67 Since IEF is not highly sensitive, subtle
changes possibly eluded detection, necessitating further investigations for confirmation. The
reason for the ER → RE mutation causing decreased phosphorylation rates in biochemical
studies compared to in-vivo results is unclear, but it might be related to the misfolding of
exogenous REY-Rho, limiting access to GRK1. Indeed, Yue et al. indicated that
microspectrophotometry measurements suggested slightly lower expression of REY-Rho than
WT-Rho, suggesting some instability in REY-Rho. However, our study did not detect such
differences (Figure S3). Together, our IEF results challenge previous assumptions that the ERY
motif is directly involved in the interaction of R* and GRK1 in-vivo.

85
Previous biochemical and modeling data suggests that ERY motif may be the
promiscuous binding site for both transducin activation276,281 and arrestin binding,273 via interaction
with both arrestin’s finger loop68,319 and transducin �-subunit’s (T�) C-terminal tail.320,321 While
prior studies manipulated/modeled these proteins outside the native cellular environment, we
investigated the promiscuity of this domain within rod photoreceptors by utilizing light exposure to
visualize protein translocation. Consistent with the light-insensitive nature of REY rods, our results
indicated that our dim light condition (20-lux) could not trigger detectable transducin translocation.
The 20-lux light intensity (equivalent to ~1-3 cd/m2
, i.e. mesopic conditions322) is typically bright
enough to nearly saturate wildtype rods.
111 In these brighter conditions, the number of activated
transducin molecules exceeds the rate of GTP hydrolysis,121 and the reduced membrane affinity
of acyl group116,121 on T� enables its detachment from ROS discs, leading to its redistribution
within the rod. This phenomenon was seen in our dim-light exposed WT rods but not in REY rods,
likely attributable to limited activation of a sizeable portion of transducin. As anticipated, the
threshold for transducin translocation in REY mice was shifted to a much higher light intensity,
where a 30 min exposure to 5,000-lux resulted in only small but detectable movement of
transducin to the inner compartments (Figures 3.3 and 3.4). This reduced translocation of
transducin observed in bright conditions is consistent with previous findings indicating that REY
rods operate under illuminations levels commonly associated with photopic vision.67 Moreover,
given that rhodopsin activation67 and phosphorylation (Figure 3.2) appear unaffected in REY mice,
it was unsurprising that the arrestin translocation to these high affinity binding partners112,125
remained unaffected. Consequently, our data further substantiate the idea that light-driven
translocation of arrestin does not depend on transducin signaling.132 Although the ER → RE
modification has the potential to impact the electrostatic interaction between the ERY motif and
arrestin’s negatively charged finger loop,323 our results revealed no discernible differences in the
localization of light-activated arrestin, suggesting that the ERY motif does not play a crucial role
in enabling arrestin binding. Thus, structural data of rhodopsin-arrestin interactions319,323 may not

86
necessarily translate into measurable effects on their coupling in intact photoreceptors.
Interestingly, arrestin translocation to the ROS in 20-lux and 5,000-lux was not as robust as
anticipated, a departure from observations at similar intensities.
113,130,132 We suspect that the
anesthesia during light exposure may have contributed to this discrepancy. Our translocation data
reinforces the significant role of the ERY domain in facilitating the interaction between lightactivated rhodopsin and transducin while simultaneously revealing its non-essential role in
arrestin recognition/binding.
Activation of the phototransduction cascade requires rapid interaction between rhodopsin
and transducin, commonly thought to occur by a ‘collisional coupling’ mechanism because of the
relatively fast lateral diffusion of rhodopsin within the ROS disc membrane.324,325 Recent studies
revealed an ordered arrangement of rhodopsin in track-like structures in disc membranes,79,326
potentially limiting its lateral mobility. Another proposed model for the fast activation kinetics
involves a dynamic pre-coupling of rhodopsin and transducin in darkness, supported by insilico327-329 and in-vitro studies.78,79 In our investigation, we unexpectedly observed a higher
concentration of transducin in the RIS of REY rods in darkness compared to WT( Figures 3.3 and
3.4), suggesting an impact of the REY mutation on the localization of transducin. We propose that
the ER → RE charge reversal impacted the formation of pre-assembled complexes between
rhodopsin and transducin in darkness, subsequently affecting the sequestration of transducin to
the ROS discs (Figure 3.5). Apart from the lipid tails conventionally acknowledged for retaining
heterotrimeric transducin in the ROS disc membranes,330,331 these transient complexes with
rhodopsin in the dark may additionally play a role in stabilizing transducin within the membrane.
Supporting this concept, if ~25% of transducin in ROS indeed exists in a pre-coupled state in
darkness,78 our observation of ~10% more total transducin in the RIS aligns well with this idea,
especially when accounting for the membrane affinity of transducin. Other factor(s) impacting
transducin’s dissociation from the ROS discs remain to be investigated.

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Figure 3.5. Diagram of the hypothetical mechanism for a stabilizing rhodopsin-transducin complex
in darkness. The interaction between rhodopsin (blue) and transducin (pink) is crucial for initiating vision
in rod photoreceptor cells. A dynamic, pre-formed complex between rhodopsin and transducin in darkness
would increase the speed of activation of the phototransduction cascade. This interaction may also help
sequester a portion of transducin in the ROS disc membrane, together with the lipid modifications on the
transducin subunits (�, �). Our results indicate that rhodopsin’s ERY motif is involved in promoting the
coupling of native rhodopsin and transducin in the dark prior to light stimuli. ROS, rod outer segment; RIS,
rod inner segment; CB, cell body; SN, synaptic terminal. This graphic was created with BioRender.com.
In support of our hypothesis concerning the disruption of this pre-coupling, comparative
structural analysis demonstrates that Arg135 in the ERY motif not only participates in the “ionic
lock,” maintaining rhodopsin in its basal inactive state, but also transiently interacts with T�’s Cterminus in darkness.328,329,332 However, it remains unclear whether simulation data of the ERY
motif interaction with transducin is directly comparable to in-vivo structural and functional
outcomes. Nevertheless, our results indicate that ER → RE charge reversal impacted transducin
localization in darkness, making it plausible that it also impacted the formation of rhodopsintransducin pre-complexes. Furthermore, while mislocalization of REY-Rho could potentially
explain elevated levels of transducin to the RIS in darkness, rhodopsin staining did not reveal any
signal differences between WT and REY retinas67 (Figure 3.3A), affirming this is unlikely a

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consequence of improper trafficking of both REY-Rho and transducin to the ROS. Building upon
the concept that transducin translocation reduces the number of molecules R* interacts with in
bright conditions, we posit that the movement of a portion of transducin to the RIS in darkness
might also contribute to the light insensitivity of REY rods when exposed to light. Previous
investigations into pre-assembled rhodopsin-transducin complexes in darkness utilized
solubilized rhodopsin and transducin.78,333 However, the transient nature of these potential
interactions, coupled with the impact of detergents on quaternary structures,334 makes it difficult
to firmly establish the existence of these complexes in-vivo. Overall, our results obtained with the
REY mouse line lend additional support to the existence of a stabilizing, pre-assembled complex
between rhodopsin’s ERY motif and transducin in darkness, suggesting that this dynamic precomplex could be an essential step in the photoresponse in-vivo.

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3.6 Supplemental Information
Figure S3.1. Peeling technique of lyophilized mouse retina for western blot. Diagram of the main steps
for the tape peeling procedure to separate the rod outer segment (+ROS), rod inner segment (+RIS), and
the remaining retinal layers (-OIS) from lyophilized retinas. Peeling workflow was generated with
BioRender.com and was adapted from Rose et al. (2017 and 2021).298,299
Figure S3.2. Confirmation of cone staining with arrestin-1 using peanut agglutinin (PNA).
Representative frozen retinal section from a WT mouse, after a 15 min 20-lux light exposure, incubated
with ARR1 (yellow, left panel) and PNA (red, middle panel) antibodies. The right panel represents the
superimposed image of ARR1 and PNA labeling. Cones are co-labeled with ARR1 and PNA, and the inset
shows a magnified image. Nuclei are labeled with DAPI (blue). ROS, rod outer segment; RIS, rod inner
segment; CB, cell body; SN, synaptic terminal. Scale bar = 10 µm.
Place tape on lyophilized retina
Tape
Filter paper
Retina
Peel tape away
+ROS
+RIS
-OIS
Separate ROS and RIS with tape
Tape
Tape
+ROS
+RIS
ROS
RIS
CB
SN
ARR1 PNA Overlay

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Figure S3.3. Rhodopsin bleaching during steady-state 20-lux light in-vivo. Quantification of total
amount of rhodopsin per retina present in WT (N = 6, white circles) and REY (N = 6, red circles) retinas
after exposure to darkness (dark gray) or 20-lux light (15 mins, light gray). 20-lux does not generate a
measurable loss of pigment (one-way ANOVA, p = 0.21).
WT (dark)
REY (dark)
WT (20-lux)
REY (20-lux)
0.0
0.1
0.2
0.3
0.4
0.5
0.6
0.7
Rhodopsin Concentraion
(nmol/retina)
WT (dark)
REY (dark)
WT (20-lux)
REY (20-lux)
WT (dark)
REY (dark)
WT (20-lux)
REY (20-lux)
0.00
0.25
0.50
0.75
Rhodopsin Concentraion
(nmol/retina)
WT (dark)
REY (dark)
WT (20-lux)
REY (20-lux) ns
0.5
0.7
WT REY WT REY WT (dark)
REY (dark)
WT (20-lux)
REY (20-lux)
0.00
0.25
0.50
0.75
Rhodopsin Concentraion
(nmol/retina)
WT (dark)
REY (dark)
WT (20-lux)
REY (20-lux)
Dark
20-lux
Rhodopsin Concentration
(nmol/retina)
0.6
0.4
0.3
0.2
0.1
0

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CHAPTER 4
Imaging Compartmentalized Ca2+ Dynamics in Rod Photoreceptors
4.1 Summary
Calcium (Ca2+) homeostasis is essential for sensory transduction and survival of the lightsensitive and compartmentalized rod photoreceptor cell. Chronic dysregulation of intracellular
Ca2+, a key second messenger, is implicated in photoreceptor dysfunction observed in many
inherited retinal diseases, such as congenital stationary night blindness (CSNB), and is
conjectured to play a role in initiating cell death. While models of vertebrate photoreceptor
degeneration have been extensively explored, direct evidence linking changes in Ca2+
homeostasis to cell death is hampered by the lack of physiological and pathophysiological
measurements of intracellular Ca2+ concentrations in all rod compartments. Moreover, the extent
to which the propagation of Ca2+ ions between compartments contributes to the maintenance of
homeostatic Ca2+ levels remains unclear. To address these issues, we developed a two-photon
Ca2+ imaging protocol that enables the investigation of compartmentalized Ca2+ dynamics in
transgenic murine rods that selectively express a genetically encoded ratiometric Ca2+ indicator,
Salsa6f. This approach permits the monitoring of various Ca2+-related properties, including Ca2+
flares, mitochondrial Ca2+ compartmentalization, and light-evoked Ca2+ changes across rod
compartments. Additionally, our in-situ (‘in-slice’) calibration protocol ensures accurate
determination of absolute Ca2+ concentrations within each compartment’s intracellular milieu. The
versatility of this rod-specific transgenic mouse line, which can be bred into various genetic
backgrounds, combined with our Ca2+ imaging pipeline, offers a promising approach for
unraveling Ca2+ signaling intricacies in both healthy and diseased states.

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4.2 Introduction
Vision is mediated by rod and cone photoreceptors, specialized sensory neurons in the
retina responsible for converting photons into electrical signals. Rods, essential for dim light
vision, exhibit a polarized structure consisting of distinct regions – the rod outer segment (ROS),
rod inner segment (RIS), cell body (CB), and synaptic terminal (SN). In darkness, rods are
tonically depolarized,335 characterized by elevated levels of free intracellular calcium (Ca2+) within
each compartment.134 Exposure to light activates the phototransduction cascade, which lowers
Ca2+ levels in the ROS and initiates a hyperpolarizing wave that propagates to the SN. This
ultimately results in a transient decrease in synaptic Ca2+ levels.18,134,141 Calcium influx and
clearance in the ROS are tightly regulated by cyclic nucleotide-gated (CNG) channels and the
Na+
/K+
, Ca2+ exchanger 1 (NCKX1).135,336,337 In non-ROS regions, Ca2+ primarily enters through
L-type voltage-gated Ca2+ channels (VGCC) and is extruded by plasma membrane Ca2+-ATPases
(PMCA).141,145,338 The quasi-independent regulation of compartment-specific Ca2+ flux contributes
to the unique physiological functions of each compartment. For instance, changes in Ca2+ levels
in the ROS modulate phototransduction response recovery339,340 and light adaptation,
71,134 while
at the synapse, Ca2+ influx sustains the tonic release of glutamate-containing vesicles from the
specialized ribbon complex,17,341 thereby mediating graded light responses to downstream retinal
cells. Additionally, Ca2+ buffering and sequestration by the mitochondria and endoplasmic
reticulum (ER) in the RIS influence energy metabolism342,343 and protein synthesis.344,345 Thus, the
maintenance of Ca2+ homeostasis is crucial for ensuring normal rod function and is regulated
through complex and dynamic interactions between plasma membrane channels/exchangers and
internal organelles.
Historically, investigations into Ca2+ dynamics in healthy rod cells have primarily relied on
electrical recordings134,346,347 or fluorescent Ca2+ imaging,141,145,348 often conducted in reptilian or
amphibian models due to their larger size (~8-11 µm diameter) compared to rodents (~1.2 µm).
While electrophysiological techniques offer insights into Ca2+ currents, Ca2+ indicators enable

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cytosolic Ca2+ measurements. Seminal work by Krizaj and Copenhagen145 demonstrated
compartmentalization of Ca2+ in isolated salamander rods using the ratiometric Ca2+ dye Fura2AM, as evidenced by minimal to no changes observed in the RIS/SN upon LiCl-induced Ca2+
elevations in the ROS.145 Subsequent studies using Ca2+ dyes further investigated dark-state and
light-stimulated Ca2+ levels in individual compartments in various species, including Gecko
(ROS142, SN349), salamander (ROS,350 RIS,141 SN141,351), frog (ROS352,353) and mouse (ROS,139
CB,148 SN140). However, both ratiometric and single-wavelength synthetic dyes have several
limitations, including lack of cell specificity during loading, unsuitability for long-term studies due
to dye leaks, and interference with physiological conditions, potentially resulting in the production
of toxic byproducts (formaldehyde),
354,355 or the inhibition of plasma membrane ion
transporters.356,357 Consequently, researchers have turned to genetically encoded Ca2+ indicators
(GECIs) for improved visualization of Ca2+ dynamics in photoreceptors.343,358,359 While GECIs offer
advantages, such as cell-type specific targeting and long-term measurement capabilities,
nonetheless, they present a drawback: fluorescence intensity varies with expression levels,
potentially differing between cells, rendering them unsuitable for quantitative measurements.
Despite considerable research on compartmentalized Ca2+ dynamics in rods, accurate
physiological measurements of intracellular Ca2+ and simultaneous visualization of Ca2+ dynamics
across all compartments in intact mammalian rods remain unexplored.
Excessive Ca2+ and abnormal Ca2+ homeostasis, frequently arising from mutations in
genes encoding key phototransduction or structural proteins, is believed to detrimentally impact
rod function and longevity,135,157,158,360 with degeneration likely mediated by Ca2+-activated
calpains or caspase proteases.144,361,362 In support of this hypothesis, elevated cytosolic Ca2+
concentrations have been directly measured using Ca2+ indicator dyes in rd1 rods.157 These rods
exhibit inherited retinal degeneration stemming from a mutation in the �-subunit of PDE-6
(phosphodiesterase-6),
363,364 resulting in heightened CNG channel opening and elevated Ca2+
levels. Moreover, several studies have indicated that apoptotic cell death in these rods may be

94
facilitated by Ca2+-activated calpain365,366 and caspases.361,367 However, the use of Ca2+ blockers
aimed at delaying rod degeneration in response to Ca2+ overload in inherited retinal disease
models, such as the rd1 mouse model, has yielded inconsistent results.136,368-371 Conversely, low
Ca2+ levels have been implicated as a likely initiator of cell death in arrestin knockout158,372 and
RPE65 knockout rods,373 which suffer from continuous activation of the phototransduction
cascade (known as ‘equivalent light’).
374,375 Despite extensive mouse modeling of inherited retinal
diseases and advances in Ca2+ imaging techniques, the absence of physiological and
pathophysiological measurements of Ca2+ levels in all rod compartments impedes our ability to
map the exact sequence of Ca2+-related events leading to apoptosis. Particularly, since the
organelles that execute cell death reside in the RIS, the manner in which perturbations in Ca2+ in
the ROS and synapse are transmitted to the RIS to initiate cell death remains unclear.135 Direct
and accurate measurements of Ca2+ concentration in healthy rods, along with assessment of its
alteration due to various mutations, would significantly enhance our understanding of the
connection between Ca2+ dynamics in individual compartments and the process of cell death.
To explore spatiotemporal Ca2+ dynamics and assess intracellular Ca2+ concentrations
across all compartments of murine rods in both healthy and degenerative mutants, we utilized the
Salsa6f reporter mouse.376 This transgenic mouse model expresses a genetically encoded
ratiometric Ca2+ indicator (GERCI), consisting of a Ca2+-insensitive red fluorescent protein
(tdTomato) fused with the Ca2+-sensitive GCaMP6f protein.377 We selectively induced Salsa6f
expression in rod photoreceptors by using the iCre75 mouse line.378 Here, we outline a protocol
and strategies for imaging compartmentalized cytosolic Ca2+ levels in retinal slices from
transgenic mice expressing Salsa6fiCre. This protocol covers the preparation of ex-vivo retinal
slices, guidelines for 2-photon imaging, and approaches to evaluating Ca2+ compartmentalization.
This includes 1) experiments validating spontaneous spike-like activity at rod terminals and the
role of mitochondria as Ca2+ buffers between the ROS and RIS, 2) crossbreeding examples of
relevant control mouse models, 3) details on light-evoked Ca2+ dynamics, and 4) an in-situ Ca2+

95
calibration protocol for obtaining absolute Ca2+ values in each compartment. Additionally,
introducing the SalsaiCre mouse into various degenerative backgrounds can provide insights into
the role of Ca2+ pathophysiology in rod dysfunction. This guide aims to extend the requisite toolset
for investigating rod physiology and Ca2+ dynamics within and between compartments of healthy
and dysfunctional mammalian rods. Valuable insights gained from this approach could ultimately
guide the development of therapies aimed at restoring Ca2+ homeostasis and
preventing/minimizing vision loss.
4.3 Methods
Ethics statement
All animal experiments were approved by the University of Southern California Institutional Animal
Care and Use Committee.
Animals
Animals were handled in accordance with the recommendations provided in the Guide for the
Care and Use of Laboratory Animals. Non-breeder adult male and female mice aged 1-3 months
were used in this study. Details of rhodopsin-iCre (iCre75 or iCre),378 Salsa6f,376 Gnat1-/-379,
RhoREY/REY (REY),67 and Nckx1-/-173 mice have been previously described. Rod-specific,
ratiometric calcium reporter strains (Salsa6fiCre, Gnat1-/-
/Salsa6fiCre, REY/Salsa6fiCre, and Nckx1-/-
/Salsa6fiCre) were initially created by crossing iCre75378 mice and LSL-Salsa6f (JAX# 031968)
mice. iCre75 mice selectively express Cre recombinase in rod photoreceptors.378 LSL-Salsa6f
mice carry the Gt(ROSA)26SorpCAG-FRT-LSL-Salsa6f-WPRE-bGHpA-AttB/P allele with a floxed-STOP cassette
upstream of Salsa6f (TdTomato-V5-GCaMP6f) to prevent widespread expression of the
ratiometric calcium indicator.376 This crossbreeding resulted in mice exclusively expressing the

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ratiometric calcium indicator (Salsa6f) in rods. The Salsa6fiCre mouse line was further mated with
Gnat1-/-
,
379 REY67 (a gift from Dr. King-Wai Yao), and Nckx1-/-173 mice (a gift from Dr. Vladimir J.
Kefalov). All mice were either generated or backcrossed on the C57BL/6 background (JAX#
000664). Salsa6fiCre mice were maintained as hemizygous. Genotypes were determined by
Transnetyx, Inc., and information is available upon request. Mice were housed under cyclic light
(12h light/12h dark) in transparent cages and had unrestricted access to food and water. All mice
were euthanized by isoflurane inhalation, followed by cervical dislocation.
Slice preparation
Eyes were harvested from euthanized mice, and all procedures were performed at room
temperature (RT). The preparation of retinal slices for Ca2+ imaging was performed following an
adapted procedure.380,381 Briefly, the eyes were enucleated, and the retinal dissection proceeded
in a textured 35 x 10 mm petri-dish filled with carbogenated (95% O2/5% CO2) Ames’ Bicarbonate
buffer (1.9 g/L NaHCO3, 1 bottle/L Ames’ Medium (Sigma), pH 7.2, 280 mOsm)). The cornea,
lens, and vitreous body were removed, and the resulting eye cup was further processed by
carefully removing the retinal pigmented epithelium (RPE) away from the retina. The isolated
retina was cut into two small rectangular pieces (Figure 4.1A) using a feather scalpel (No. 10),
and each piece was flattened by cutting the curved edges of the retina. The hemisected retinas
were subsequently embedded in 3% low melting agarose (Invitrogen) at temperatures no higher
than 32°C and vertically sliced (photoreceptor layer facing up) using a Stoelting tissue slicer
(Stoelting, #51424), generating 250 µm-thick sections (Figure 4.1B). Slices were transferred to a
custom-fabricated Delrin chamber192 using a paint brush and were incubated in 95% O2/5% CO2-
saturated Ames’-bicarbonate buffer for 1-2h at RT before imaging. Photoreceptors are viable for
~12 h when stored in oxygenated Ames’ Bicarbonate buffer.

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Two-photon Ca2+ imaging
The live-tissue retinal sections were maintained in carbogen-saturated Ames’-bicarbonate buffer
and secured to a dish with a bottom inset cover glass (NuncTM, Thermofisher Scientific) using a
handmade U-shaped (“harp”)382 anchor with nylon strings attached perpendicularly to each arm
of the U (Figure 4.1C). Each tissue section was imaged for a maximum duration of 10 mins.
Images were acquired using a multiphoton confocal microscope (Lecia SP8 DIVE imaging
system, 63X/1.3 NA glycerin objective) equipped with Hybrid Detectors (HyD)(4Tune, Leica,
emission at 480-540 nm for GCaMP6f and 590-640 nm for tdTomato). Illumination was provided
by a tunable, ultrafast Chameleon Discovery laser at 960 nm (Coherent, CA). While 920 nm
wavelength is optimal for GCaMP6f excitation, 960 nm wavelength was chosen to mitigate the
potential of indirect photoreceptor activation by the fluorescence emitted from GCaMP6f activity
(emission 500-550 nm). In particular, photons emitted by GCaMP6f could inadvertently excite
rhodopsin382 (peak absorbance at 500 nm), especially when imaging under dark-adapted
conditions. The 960 nm wavelength strikes a balance, ensuring sufficient excitation for both
GCaMP6f and tdTomato while minimizing emitted photons that could inadvertently excite rods,
thereby aiding in the capture of the ratiometric Ca2+ signals in rods expressing Salsa6f. To further
minimize tissue exposure resulting from GCaMP6f emission, rapid scan times (8,000 Hz) using
the SP8 resonant scanner were used to capture rapid light-induced changes (~200 ms).
Fluorescence signals were easily detected up to 150 µm in depth with 960 nm, and rod cells were
imaged at least 20 µm into the retinal slice to avoid capturing potentially damaged cells. To
minimize phototoxicity risks, a combination of strategies was employed, including high scan
speeds (8K Hz) and low laser output power (20%, 1.3 W). Continuous monitoring of laser power
at the imaging plane (≤ 300 µW), with the assistance of a power meter (StarBright, OPHIR), further
contributed to this strategy. All parameters, including laser power and gain, remained consistent
across experiments. Time-lapse images of x = 97 µm and y = 97 µm were acquired at 167 ms
intervals for 5 min (pixel dwell time = 72 ns), and single snapshot images of x = 148 µm and y =

98
x = 148 µm were acquired in 240 ms (pixel dwell time = 216 ns) using Leica LASX software (Leica
Microsystems).
Image analysis of Ca2+ data
All live-cell Ca2+ imaging data were analyzed using Leica LASX software and Fiji200. Raw images
were processed, and look-up tables (LUT) were applied to aid in the visualization of fluorescence
in both single snapshot and time-lapse images. Specifically, a pseudocolored “Fire” LUT was
assigned to GCaMP6f images, while a gray LUT was applied to tdTomato images. Mean
fluorescence intensity (gray value) measurements for distinct rod compartments (outer segment
(OS), inner segment (IS), cell body (CB), synaptic terminal (SN)) were acquired by manually
delineating regions of interest (ROIs) within each sub-cellular compartment. Special attention was
given to tracing signals from a select number of cells (1-5), depending on the ease of isolating
individual cellular compartments (OS and SN tended to be more challenging). Before calculating
fluorescence intensity ratios for Salsa6f, an ROI positioned outside the retinal slice served to
establish baseline fluorescence in both channels for background subtraction (Figure S4.1A, white
box). Subsequently, the normalized (background-subtracted) GCaMP6f and tdTomato
fluorescence signals were obtained, and the GCaMP6f/tdTomato ratio for each ROI was
determined. For time-lapse imaging, normalized fluorescence signals for both GCaMP6f and
tdTomato were calculated using the formula ∆F/F0 = (F(t)-F0)/F0, with F0 an averaged fluorescence
intensity value derived from the initial 15 baseline images, and F(t) the mean fluorescence
intensity value at time t. Towards the end of continuous 5-min time-lapse imaging, tdTomato
experienced minor photobleaching.383 To correct for the exponential decay of tdTomato
fluorescence during time-lapse imaging, a FIJI plug-in “Bleach Correction” (version 2.1.0)384 was
applied. The resulting GCaMP6f/tdTomato ratios were then plotted for each rod compartment of
interest. In instances where noticeable XY-drift occurred during the 5 min live-cell imaging, the

99
time-series stacks underwent processing in FIJI using the StackReg385 plug-in with the ‘Rigid
Body’ transformation, and the GCaMP6f and tdTomato signals were subsequently analyzed.
“In-slice” Ca2+ calibration
Eleven Ca2+ calibration solutions were prepared using a calcium calibration buffer kit (Molecular
Probes, C3008MP) and the reciprocal dilution approach, which involved the mixing of known
quantities of EGTA and CaEGTA. To control Ca2+ concentration in intact rods, 250 µm-thick live
retinal slices underwent a 45 min incubation at RT in a 1:1 solution containing a specified free
Ca2+ concentration (0 nM, 17 nM, 38 nM, 65 nM, 100 nM, 150 nM, 225 nM, 351 nM, 602 nM, 1.35
µM, or 39 µM) and Ca2+-free Ames’ buffer (8.8 g/L Ames’ Media w/o CaCl (US Biological), pH 7.2,
560 mOsm adjusted with D-glucose), before imaging. The 1:1 ratio of Ca2+-free Ames’ buffer to
Ca2+ calibration solutions served to maintain an osmolarity of 280 mOsm in retinal slices, ensuring
physiological conditions without altering free Ca2+ values. This adjustment is made possible by
the robust chelating properties of EDTA, which remains unaffected by added solution. To interrupt
intrinsic Ca2+ regulation, each calibration solution was supplemented with specific agents: 10 µM
ionomycin (Ca2+ ionophore, Sigma-Aldrich), 5 µM thapsigargin (an inhibitor of Ca2+ uptake into
ER, Sigma-Aldrich), 10 µM m-Cl-CCCP (which collapses the mitochondrial membrane to block
Ca2+ uptake, Sigma-Aldrich), and 10 µM nigericin (K+ ionophore that stimulates Ca2+ release from
mitochondria, Sigma-Aldrich). Each retinal slice was exclusively exposed to and imaged in one of
the eleven Ca2+ calibration solutions, with the calibration progressing sequentially from the lowest
(0 nM) to the highest (39 µM) free Ca2+ concentrations. The “in-slice” calibration procedure for
different rod compartments followed established protocols.386,387 Briefly, images from eleven
different free Ca2+-buffered solutions (0 nM – 39 µM) were processed in FIJI, and the normalized
fluorescence ratios for each ROI were acquired (see Image analysis of Ca2+ data). Retinal slices
with normal rod morphology were included in the analysis. The normalized GCaMP6f/tdTomato
ratios specific to each rod compartment from three separate trials were plotted against the eleven

100
free Ca2+ concentrations and fitted with the Hill equation, Y= Rmin + [(Rmax - Rmin)/(1+ 10(logKd-logX)*n)],
where Rmin is the ratio at zero free Ca2+ (0 nM), Rmax is the ratio at saturating Ca2+ (39 µM), X
represents the GCaMP6f/tdTomato ratio, n is the Hill coefficient, and Kd is the apparent
dissociation constant. This fitting approach helped determine both Kd and n values for the ROS,
RIS, CB, and SN, providing key parameters for calculating an estimate of absolute Ca2+
concentrations in each rod compartment from normalized Salsa6f fluorescence ratios.
Modulation of Ca2+ exogenously
Retinal slices were anchored using “harp” weights within a specialized holder (Fig. 4.1C), where
a glass coverslip (25mm circle no. 1, VWR) was securely affixed to the lower part of the holder
using vacuum grease. The assembly was inserted into a custom perfusion chamber/platform
designed to provide both inflow and suction, enabling laminar flow across the retinal sample. For
KCl depolarization experiments, a cold Ringer’s solution (7.2 g/L NaCl, 0.17 g/L CaCl2, 0.37 g/L
KCl, pH 7.2, ~280 mOsm) was introduced into the perfusion chamber for at least 1 min before
switching to a cold high KCl solution (100 mM KCl in Ringer’s, 280 mOsm), which was perfused
at a constant rate of 1 ml/min using a Masterflex® L/S drive system (Cole-Parmer, 7520-60). To
alter mitochondrial Ca2+ uptake, Ru360 (Sigma-Aldrich, 557440) was prepared following
established protocols.388 Fresh Ru360 solutions were prepared for each experiment, starting with
an initial dilution in cold MilliQ water (5 mM) before further diluting it in Ames’ Bicarbonate buffer
to achieve a concentration of 10 µM. Live retinal slices were then incubated in 10 µM Ru360,
bubbled with 95% O2/5% CO2, within a 3 ml glass dish inside a custom-fabricated Delrin
enclosure for a minimum of 1h on ice. Following incubation, the slices were transferred to the
perfusion setup and were promptly subjected to depolarization using a cool 100 mM KCl solution
with a flow rate of 1 ml/min. Time-lapse images were processed according to the methods outlined
in the Image analysis of Ca2+ section.

101
Light-stimulation
Near-infrared lasers were used to stimulate rod photoreceptors389,390 without inducing tissue
damage (< 10 mW at sample).391 Leica’s fluorescence recovery after photobleaching (FRAP)
module was utilized to capture laser-induced rod cell stimulation. Initially, ROIs were manually
drawn using the Leica software to include the entire ROS in individual retinal slices, marking the
area for 960 nm exposure. The laser power was set at 50% for the “bleach” frames (960 nm, 0.65
mW), resulting in laser illumination that approximately matched low-photopic conditions.
Subsequently, the parameters for pre-flash (10), flash (10), and post-flash (600) frames were
defined, with frames acquired every 167 ms (6 fps). The tissue depth for imaging ranged from 20
µm to 60 µm, and the imaging sequence parameters for FRAP remained constant between
experiments. The subsequent retinal time-series images were analyzed (see Image analysis of
Ca2+ data). The 10 flash frames that specifically target the ROS were excluded from data
processing due to saturation of the ROI.
Statistical analysis
The number of mice (n) is specified in the corresponding figure legends, and data are presented
as a mean ± standard deviation. GraphPad Prism 10.1.1 for Mac was used to generate plots and
perform statistical analysis. Normality was assessed using the Shapiro-Wilk test. When data was
normally distributed, statistical analysis was performed with a t-test (pairwise comparisons). In the
case of non-normal distribution, the Kruskal-Wallis one-way ANOVA followed by Dunn’s multiple
comparison test was performed. Statistical significance was set at � = 0.05, and results with p ≤
0.05 were considered statistically significant.

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4.4 Results
Expression of Salsa6f in rods allows visualization of spontaneous Ca2+ flares.
We developed a transgenic mouse line expressing a non-FRETing genetically encoded
ratiometric Ca2+ indicator (GERCI) in rods by crossbreeding commercially available mouse
strains, Salsa6f376 and rhodopsin-iCre.
378 Salsa6f, consisting of the Ca2+-sensitive GCaMP6f
indicator fused to a Ca2+-insensitive red fluorescent protein, tdTomato,376 is targeted to the
ROSA26 locus for Cre-dependent expression. Co-expression with rhodopsin-iCre targets the
Ca2+ indicator exclusively to rod photoreceptors. The presence of the Ca2+-insensitive tdTomato
serves multiple functions: its uniform distribution throughout the rod cell enables visualization of
rod morphology and permits ratiometric imaging. Transgenic mice can carry either one or two
allelic copies of Salsa6f, offering control over the brightness of the Ca2+ indicator in rods.376 For
this current study, we used mice heterozygous for Salsa6f.
Characterization of Salsa6f expression in rods involved the generation of vertical retinal
sections. Precise positioning of the flattened retina in agarose for vertical sectioning (Figures 4.1A
and 4.1B) was crucial for imaging consistent signals from Salsa6f in all rod compartments. During
2-photon imaging, live retinal slices were stabilized using custom ‘harp’ weights, and careful
examination under brightfield illumination ensured all rod subcellular compartments were clearly
visible (Figure 4.1C). Flipping the slice provided an additional opportunity to locate the correct
focal plane encompassing the entire photoreceptor layer. In light-exposed retinal slices, we
observed widespread Salsa6f expression in all rod compartments (Figure 4.1C), with rods
displaying normal morphology. Although the tdTomato signal appeared uniform throughout the
entire rod cell, it was dimmer in the rod outer segment (ROS), likely due to the limited cytoplasmic
space (~31%)12 for the Ca2+ biosensor. Consistent with expectations for light-adapted retinas,
wherein the rod membrane is hyperpolarized and Ca2+ levels are low,134 we consistently observed

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a reduction in GCaMP6f signal throughout the entire rod cell layer (Figure 4.1C). Elevated
GCaMP6f fluorescence observed in distinct sub-cellular compartments under light-exposed
conditions may serve as an indicator of tissue damage incurred during the tissue slicing process.
In support of this, vibratome sectioning, known to mechanically disrupt the delicate ROS during
sectioning, resulted in heightened GCaMP6f signals in the ROS compared to vertical slices
generated with the Stoelting chopper (Figures 4.1C and S4.1A). We also confirmed that
homozygosity increased the brightness of Salsa6f about two-fold376 in live retinal slices (Figure
S4.1B and S4.1C), and Salsa6f expression remained stable until at least 8 months of age (data
not shown).376 The ubiquitous expression of Salsa6f in all rod sub-cellular compartments enables
simultaneous monitoring of numerous rods and distinct compartments, allowing for the collection
of multiple Ca2+ responses in a single experiment. By combining vertical slices prepared from
Salsa6fiCre mouse retina with 2-photon imaging, we can visualize the Ca2+ concentration in all rod
compartments.
Figure 4.1. Preparation of retinal slices from Salsa6fiCre mice. (A) Isolated retinas are bisected and
flattened for agarose embedding. The dotted rectangles denote the target areas for subsequent sectioning.
(B) A rectangular piece of flattened retina from (A) is embedded in 3% low-melt agarose. Vertical 250 µmthick sections are generated, with a focus on harvesting slices from the central region of the retina. (C)
Schematic representation of the imaging chamber, a retinal slice with Salsa6f (GCaMP6f-tdTomato)-
expressing rods, and a specialized “harp” weight. The rectangular inset shows a live-cell image (merged)
B
A
Retina
Region for
sectioning
INL
D
V
Agarose block
Sections
250 µm Retina
C
ROS
RIS
CB
SN
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Salsa6f slice
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captured from light-adapted rods selectively expressing the ratiometric Ca2+ indicator, Salsa6f (GCaMP6ftdTomato), using iCre75. GCaMP6f and tdTomato fluorescence signals are featured in green and red,
respectively. ROS, rod outer segment; RIS, rod inner segment; CB, cell body; SN, synaptic terminal. Scale
bar = 5 µm. Schematics were generated with BioRender.com.
Time-lapse imaging of rod cells is important for monitoring and evaluating Ca2+ dynamics
under physiological and pharmacological conditions. Therefore, we monitored Salsa6f signal and
rod cell morphology over a 3.5 min time period to ensure that our 2-photon imaging parameters
did not induce cell death over extended periods. During these imaging sessions, we observed
spontaneous calcium flares, i.e., sudden bursts of increased GCaMP6f fluorescence, in intact
Salsa6fiCre retinal slices (Figure 4.2A). These flares primarily occurred in distinct compartments of
individual rod cells, notably in the rod inner segment (RIS), CB, and SN (Figure 4.2A, colored
insets), with no Ca2+ transients observed in the ROS. For example, in the representative
Salsa6fiCre retinal slice shown in Figure 4.2A, a bright fluorescence increase in GCaMP6f was
observed in the SN at 00:33 s but was not seen thereafter, while another GCaMP6f signal
increase in the RIS reoccurred four times throughout the imaging period. Furthermore, these
flares predominantly originated in the CB and SN (Figure 4.2A, arrows), exhibiting random and
varied spatiotemporal patterns. However, due to the uniform expression of Salsa6f in all rod cells,
distinguishing whether flashes propagated within an individual cell proved challenging. As noted
by others,352,358,392,393 these Ca2+ transients in light-adapted Salsa6fiCre rods appear to be
independent of phototransduction, since, following each flare, we detected a subsequent Ca2+
decrease, as expected in a light-exposed rod (Figure 4.2A). Although abnormal Ca2+ signals are
typically associated with cell death,
394 the occurrence of these Ca2+ transients does not appear to
be correlated with rod cell death, as we did not observe a sustained increase or hyperactivity of
the GCaMP6f signal typically associated with thermal damage.395 Moreover, no laser-induced
morphological changes were observed (Figure 4.2A). To further verify that the 2-photon laser did
not induce cell death, we measured the laser power reaching the sample at the imaging plane
(~300 µW), which was significantly below the power required to cause damage (10 mW).
391

105
Figure 4.2. Spontaneous Ca2+ transients in rod photoreceptors. (A) Montage capturing spontaneous
Ca2+ flares over 3.5 min (210 s) in a retinal slice from a light-adapted wildtype Salsa6fiCre retina (n = 6). The
GCaMP6f fluorescence signal is presented in pseudo-color (LUT Fire, color bar 0-4095), enhancing
visualization of the Ca2+ transient flashes. Boxes spotlight specific rod subcellular compartments
experiencing Ca2+ transients (purple: inner segment (IS), red: cell body (CB), green: synaptic terminal (SN)),
which are graphed in (B). White arrows indicate additional cells with observed Ca2+ flashes. Scale bar = 10
µm. (B) Graphs illustrating the time course of spontaneous Ca2+ transients in the IS, CB, and SN from three
representative cell compartments (boxed in A). GCaMP6f/tdTomato ratio against time is plotted for each
individual compartment.
Next, we quantified our time-lapse image series using FIJI200 to ascertain the
spatiotemporal profiles and amplitudes of these spontaneous Ca2+ flares. Regions of interest
(ROIs) were manually drawn around each targeted compartment to isolate both GCaMP6f and
tdTomato signals (Figure S4.2A), from which three parameters were derived as described below.
First, mean intensity values were extracted for each individual compartment identified (RIS, CB,
SN). Subsequently, to normalize the GCaMP6f and tdTomato signals, background fluorescence
was subtracted from each channel (Figure S4.2B-D, top panels). Relative fluorescent signals
were then calculated using the ratios of normalized GCaMP6f/tdTomato, and the resulting signal
profiles were plotted (Figures 4.2B and S4.2B-D, bottom panels). We observed substantial
heterogeneity in the spatiotemporal profiles and amplitudes of the Ca2+ flares within the rod
population and across the distinct sub-compartments (Figures 4.2B, S4.2B-D, and S4.3). For
0 70 140 210
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106
instance, Ca2+ flashes in the RIS and CB often displayed moderate brightness (Figures S4.3A
and S4.3B), while the GCaMP6f signal in the synapse tended to be brighter (Figures 4.2B and
S4.2B-D). Additionally, the frequency of Ca2+ flares varied, occurring either once or multiple times
throughout the imaging period (Figure 4.2B). Our analysis revealed these flashes could persist
for durations ranging from 2 s up to 70 s (n = 10). By employing this quantitative methodology to
assess Ca2+ changes in rods expressing a ratiometric Ca2+ indicator, Ca2+ signals can be
systematically compared and evaluated across diverse physiological and pathophysiological
contexts.
Light-adapted Ca2+ levels in diverse mouse models
The phototransduction cascade involves several elements that are uniquely expressed in
rods. Through genetic manipulation of these components, researchers can begin to elucidate the
intricacies of Ca2+ dynamics in both healthy and diseased rods. With this in mind, we directed our
attention to studying Salsa6fiCre mice introduced into different genetic backgrounds. To establish
controls for examining dark-state Ca2+ levels, we utilized two transgenic mouse models: Gnat1-/-
,
379 lacking functional photoreception due to the absence of � transducin, and RhoREY/REY (REY),
67
which harbor a mutation in rhodopsin’s ERY motif that greatly diminishes its coupling to
transducin, necessitating photopic stimulus for function. Gnat1-/-
/Salsa6fiCre rods permit the
determination of basal dark Ca2+ levels but hinder the investigation of light-induced Ca2+ changes.
REY/Salsa6fiCre rods, on the other hand, enable the examination of both dark-state and lightstimulated Ca2+ changes under bright photopic light. Notably, Gnat1-/- and REY rods exhibit a
normal dark current,
67,379 and therefore, are expected to reflect basal dark-state Ca2+ levels under
dim room light conditions. As anticipated, two-photon imaging of light-exposed Gnat1-/-
/Salsa6fiCre
and REY/Salsa6fiCre rods revealed elevated GCaMP6f fluorescence in all compartments (Figure
4.3A). Subsequently, we directed our attention towards measuring fluorescence signals
specifically from the ROS and SN of Gnat1-/-
/Salsa6fiCre and REY/Salsa6fiCre rods. This decision

107
was motivated by the well-documented observation that these two compartments exhibit
heightened levels of Ca2+ in darkness.134 To this end, we manually delineated ROIs within the
intended compartments (Figure 4.3B), selecting small clusters of 1-5 cells, and compared the
differences in GCaMP6f/tdTomato ratios between light-sensitive and light-insensitive mice (n =
100-140 rods). As expected, Ca2+ levels were significantly higher in both the ROS and SN from
light-insensitive Gnat1-/-
/Salsa6fiCre and REY/Salsa6fiCre rods compared to wildtype Salsa6fiCre (p
= 0.0001). Surprisingly, the GCaMP6f fluorescence in the ROS and SN compartments from
REY/Salsa6fiCre rods were even brighter than the signals from Gnat1-/-
/Salsa6fiCre (p = 0.001).
Figure 4.3. Light-exposed Ca2+ signal in wildtype, transducin knockout, REY, and Nckx1 knockout
rods. (A) Vertical slices through light-exposed wildtype (WT), transducin knockout (Gnat1-/-
), REY, and
Nckx1-/- retinas expressing Salsa6f. Pseudo-color images (top row, LUT Fire, color bar 0-4095) depict
steady-state, light-adapted Ca2+ levels captured from GCaMP6f fluorescence. Bottom panel images show
the Ca2+ insensitive fluorophore tdTomato (gray). In light-exposed REY, TR-/-
, and Nckx1-/- rods the
WT Gnat1-/- REY Nckx1-/-
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108
GCaMP6f signal is noticeably stronger compared to WT rods, indicating elevated intracellular Ca2+ levels.
Scale bar = 5 µm. (B) Schematic drawing of a retinal slice with Salsa6f (GCaMP6f-tdTomato)-expressing
rods, indicating selected regions of interest (ROIs) at the rod outer segment (ROS) and synaptic terminals
(SN) of individual rods. ROIs in these specific cellular compartments were chosen for extracting the mean
fluorescence intensity of GCaMP6f and tdTomato. ONL, outer nuclear layer; INL, inner nuclear layer; GC,
ganglion cells. Diagram was generated with BioRender.com. (C-D) Quantification of the light-adapted Ca2+
signals (GCaMP6f/tdTomato ratios) isolated either from the ROS (C) or ST (D) in transgenic WT (white
circles, n = 6, 105 rods), Gnat1-/- (gray circles, n = 4, 117 rods), REY (red circles, n = 6, 144 rods), and
Nckx1-/- (dark red, n = 2, 57 rods) retinae expressing Salsa6f. A significant difference between genotypes
was detected within either the ‘ROS’ or ‘SN’ group using Kruskal-Wallis one-way ANOVA followed by
Dunn’s multiple comparison test (*p ≤ 0.002). No significant differences were observed between lightexposed Ca2+ signals in the SN between WT and Nckx1-/- rods (p = 0.55). RIS, rod inner segment; CB, cell
body.
Encouraged by the versatility of the Salsa6fiCre mouse line, we proceeded to crossbreed
Salsa6fiCre with Nckx1-/- mice,173 a model for congenital stationary night blindness (CSNB), to
investigate Ca2+ levels in degenerating retinas. In the Nckx1-/- mouse model, the absence of the
Na+
/K+
-Ca2+ exchanger 1 (NCKX1), the sole ROS ion exchanger responsible for Ca2+ clearance,
is presumed to result in elevated intracellular Ca2+ levels. However, the slow degeneration of rods
in this model suggests the involvement of alternative Ca2+ extrusion mechanisms.
173 Our Ca2+
imaging data show that compared to wildtype Salsa6fiCre rods, rods isolated from light-adapted 2-
month-old Nckx1-/-
/Salsa6fiCre mice have elevated fluorescence signals in the ROS, RIS, and CB
(Figure 4.3A, right panel). To further investigate this observation, we used our established ROI
approach (Figure 4.3B) to analyze fluorescence intensity in the ROS and SN of Nckx1-/-
/Salsa6fiCre
retinas. Our analysis demonstrates that loss of NCKX1 led to a significant increase in Ca2+ levels
in the ROS (p = 0.0002) but not in the SN (p = 0.55) compared to light-exposed wildtype Salsa6fiCre
retinas (n = 50-100 rods). Together, these findings highlight the utility of Salsa6fiCre in various
genetic backgrounds for elucidating the impact of both benign and deleterious mutations on
compartmentalized Ca2+ homeostasis.
Mitochondria act as a Ca2+ barrier in healthy rods
Recent studies using Ru360, a mitochondrial Ca2+ uptake inhibitor, have provided
evidence supporting the role of mitochondria as buffers between inner and outer segments of

109
cone photoreceptors,
359,396 thereby implying their role in Ca2+ uptake for maintaining Ca2+
compartmentalization. However, mammalian cones possess ~2-10 times more mitochondria in
their inner segment ellipsoid region than rods due to their heightened metabolic demands.397 The
question of whether rod mitochondria serve a comparable role as a Ca2+ sink between the ROS
and RIS remains unexplored. To examine Ca2+ compartmentalization in rods, we adopted a
perfusion strategy outlined by Giarmarco et al.359 in conjunction with our 2-photon imaging setup.
Initially, hyperpolarized (light-exposed) rods expressing Salsa6f were depolarized by perfusing a
Ringer’s solution containing 100 mM KCl over a ~1.5 min-period across the sample (Figure 4.4A).
The application of KCl activates L-type voltage-gated Ca2+ channels (VGCCs) located in the RIS,
CB, and SN, permitting the influx of Ca2+ ions into the cell. High KCl perfusion resulted in a
substantial increase in the GCaMP6f signal in Salsa6f expressing rods, beginning in the SN
(green box, Figure 4.4A) and propagating towards the RIS (purple box, Figure 4.4A). While the
elevated GCaMP6f signal persisted in the SN throughout the imaging period, transient elevations
in Ca2+ were observed in the other compartments (CB and IS). Notably, no change in fluorescence
intensity was observed in the ROS (Figure 4.4A), supporting the hypothesis that mitochondria
might play a role in Ca2+ compartmentalization.
To further explore the role of mitochondria in regulating Ca2+ dynamics between the RIS
and ROS, Salsa6fiCre retinal slices were pretreated with 10 µM Ru360,359 a selective inhibitor of
mitochondrial Ca2+ uniporter (MCU), for 1h before time-lapse imaging. Initial observations
revealed a slight elevation in GCaMP6f fluorescence in the ROS before KCl perfusion, suggesting
a deviation from the normal Ca2+ flux between the RIS and ROS (Figure 4.4A, left panel).
Following depolarization of the Ru360-treated retinal slices with 100 mM KCl, we captured a
transient increase in GCaMP6f fluorescence in the ROS around 00:33 s (Figure 4.4B). This finding
contrasts directly with the lack of response observed in the ROS in untreated retinal slices.
Additionally, the depolarizing wave (SN → RIS) propagated more rapidly in the Ru360 treated
samples, taking ~30 s to reach the RIS, compared to ~50 s in untreated samples (Figures 4.4A

110
and 4.4B). This finding is further supported by the almost synchronous rise in GCaMP6f
fluorescence observed in the SN and ROS. We also noted that the GCaMP6f signal in the RIS
did not return to its baseline level, unlike in wildtype Salsa6fiCre rods. Collectively, these
observations suggest that mitochondria are responsible for efficient Ca2+ uptake, with any
disruption to mitochondrial function potentially perturbing Ca2+ homeostasis within each
compartment.
Figure. 4.4. Rod mitochondria buffer Ca2+ from between rod outer and inner segments. Montages
from time-series imaging of light-adapted Salsa6f retinal slices during the application of a 100 mM KCl
solution to either (A) a wildtype retina (n = 8), (B) a wildtype retina pre-incubated for 1h with Ru360 (n = 6),
or (C) a Nckx1 knockout retina (n = 3). High KCl solution reached the retinal slice around ~30 s into the
experiment, triggering a wave of depolarization that propagated from the synaptic terminal (SN) to the other
rod compartments. When retinas experience mitochondrial dysfunction (B-C), Ca2+ flux between the inner
segments (IS) and outer segments (OS) is observed. Four time-stamped images from each condition (AC) were chosen to showcase Ca2+ influx over the period of KCl depolarization (100 s), and four regions of
interest (box/circle) in each rod compartment were chosen for ROI analysis. The temporal profiles of these
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111
Ca2+ signals (GCaMP6f/tdTomato ratio) are presented on the far right of each panel, with line profiles and
boxes color coordinated to emphasize the regions of interest (gray: outer segment (OS), purple: inner
segment (IS), red: cell body (CB), green: synaptic terminal (SN). GCaMP6f is presented in a pseudo-color
(Fire, LUT, scale 0-4095). Scale bar = 10 µm.
The transient increase in Ca2+ levels detected in the ROS of Ru360-treated Salsa6f slices
suggests the involvement of the ROS exchanger, NCKX1, in extruding the Ca2+ flux from the RIS
when mitochondria are dysfunctional. To explore the potential consequences of altered cytosolic
Ca2+ levels resulting from the absence of NCKX1 in the ROS, we utilized the Nckx1-/-
/Salsa6fiCre
mouse line. In experiments similar to those described previously, 100 mM KCl was applied to
light-exposed Nckx1-/-
/Salsa6fiCre retinal slices. Consistent with observations from Ru360 treated
slices, the GCaMP6f signal was observed propagating from the SN to the ROS (Figure 4.4C).
Moreover, in the absence of the sole Ca2+ extrusion component in the ROS, we observed a
sustained increase in the GCaMP6f signal in the ROS and in the RIS (Figure 4.4C). This result
implies impaired mitochondrial function in Nckx1-/- rods, which permitted Ca2+ flux between the
RIS and ROS compartments. These results demonstrate that Salsa6f expression in rods can
serve as a valuable tool when investigating pharmacologically-evoked changes in Ca2+ in healthy
and diseased rods.
Light-evoked Ca2+ dynamics in REY rods
Previous investigations of dark-to-light Ca2+ concentrations and dynamics in mammalian
rods focused primarily on individual compartments,139,140 leading to an incomplete assessment
across all regions simultaneously. Given the technical challenges associated with imaging the
highly sensitive rods, particularly due to the extreme photopigment concentration in the ROS,382
we opted to record light-evoked Ca2+ responses from REY rods expressing Salsa6f. REY rods
are highly insensitive and operate largely in bright photopic conditions,
67 offering greater flexibility
with the lighting environment during sample preparation. To mitigate sensor flooding during light
stimulation, we used 2-photon stimulation at 960 nm, which was capable of initiating the

112
phototransduction cascade390,398 without overwhelming the HyD sensors. Leveraging Leica’s
microscope’s fluorescence recovery after photobleaching (FRAP) module, we delivered timed
flashes to an ROI in REY/Salsa6fiCre retinal slices. Specifically, we traced an ROI encompassing
the entire ROS layer from REY/Salsa6fiCre retinal slices (Figure 4.5A, yellow ROI) and then
presented a high-powered (~0.65 mW) pulse using the 960 nm laser, lasting for 2.5 s. We then
continued to collect Salsa6f signals for an additional minute to monitor Ca2+ changes. Following
the flash, we detected a transient decrease in GCaMP6f fluorescence in both the SN and ROS
(Figures 4.5A and 4.5B). However, no discernible changes in the GCaMP6f signal were detected
in the RIS or CB (Figure 4.5A). Additionally, we found that the light pulse paradigm could be
repeated multiple times successively, confirming the viability of the slices (data not shown).
Figure. 4.5. Light-insensitive REY rods exhibit light-evoked Ca2+ responses. (A) Montage of a
representative retinal slice from REY mice expressing Salsa6f. REY mice, characterized by their
Pre-flash Recovery
60
80
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120
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Recovery
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113
insensitivity to light under photopic conditions, still exhibit responses to light, rendering them valuable for
investigating Ca2+ dynamics in response to bright light stimuli. The yellow-highlighted region indicates
where the light stimulation is directed (960 nm, 2-photon, 2.5 s). Two regions of interest (ROIs) in the outer
segment (OS) and synaptic terminal (SN) are highlighted by a gray rectangle and green circle, respectively.
The far-left panel displays the GCaMP6f signal pre-flash, the middle panel shows the post-flash signal, and
the far-right panel illustrates the fluorescence recovery in the OS and SN following the flash. The OS and
SN signals diminish after the light pulse, subsequently increasing to near pre-flash fluorescence levels after
~1 min. GCaMP6f signal is pseudo-colored (Fire, LUT, scale 0-4095). Scale bar = 5 µm. Inner segment, IS;
CB, cell body. (B) Light-induced changes in OS (gray) and SN (green) fluorescence intensity recorded in
the corresponding ROI shown in (A). The GCaMP6f/tdTomato ratio sharply decreases after the 2.5 s light
stimulation and gradually recovers. There is a more pronounced recovery in the SN, potentially attributed
to the beaching of GCaMP in the OS after light stimulation (960 nm, 2-photon). (C) Percent change in the
OS and SN fluorescence pre-flash and after a 1.5-min recovery period (n = 4, 11 retinal sections). Following
light exposure, OS fluorescence recovers ~80% of its pre-flash fluorescence. In contrast, the SN recovers
100% of its fluorescence. Data presented as mean ± SD.
To quantify light-evoked Ca2+ responses in the ROS and SN, we analyzed the response
rise of the Ca2+ profiles after light-onset and found that the half-life (t ½), the time it takes to reach
half-maximal fluorescence intensity from the plateau value, was statistically shorter in the ST (t ½
= 10.13 ± 4.2 s) compared to the ROS (t ½ = 18.68 ± 7.2 s, p = 0.004, n = 11 rod clusters). We
also noted that the post-flash fluorescence intensity from GCaMP6f in the ROS did not return to
its pre-flash fluorescence value, in contrast to the SN fluorescence, which fully recovered (Figures
4.5B and 4.5C). This discrepancy suggests that the concentrated 960 nm pulse may have partially
bleached the GCaMP fluorescent protein.
399 Despite this observation, no aberrant Ca2+ changes
were detected in the RIS or CB, and the rod’s morphology remained stable throughout the imaging
session, indicating no damage was inflicted on the rods themselves. Our findings demonstrate
that REY rods serve as a valuable mouse model for investigating dark-to-light Ca2+ dynamics and
that 2-photon light stimulation of rhodopsin in the ROS can induce hyperpolarization at the
synapse.
In-situ Ca2+ calibration of Salsa6f reveals unique milieu within individual rod compartments
To accurately assess intracellular Ca2+ concentrations in all compartments of live, intact
rods, we performed in-situ calibrations by incubating Salsa6fiCre retinal slices for 45 mins in eleven
Ca2+-buffered solutions containing specific compounds that disrupt the rod cell’s intrinsic ability to

114
regulate Ca2+.
386,400 Representative images from 4 out of the 11 different free-Ca2+ buffered
solutions are shown in Figure 4.6A, revealing a stable GCaMP6f signal in all rod compartments
that steadily increases in response to elevations in free Ca2+ levels. Surprisingly, each
compartment exhibited distinct responses to specific external Ca2+ solutions. For instance, in
retinal slices incubated in 371 nM Ca2+-buffered solution, the RIS appeared substantially brighter
compared to other compartments, with the SN region being the dimmest (Figure 4.6A). To
account for this heterogeneity between compartments, we created four separate calibration
curves for Salsa6f, each corresponding to a specific rod compartment (Figures 4.6B-4.6E). We
plotted the normalized fluorescence ratios against the eleven free Ca2+-buffered solutions and
then fitted the data extracted from each compartment with the Hill equation to estimate the
apparent dissociation constant (Kd) and Hill coefficient (n). The Kd values are 318.8 nM (ROS),
614.6 nM (RIS), 647.2 nM (CB), and 467.7 nM (SN), and the Hill coefficient values are 1.98 (ROS),
1.40 (RIS), 1.15 (CB), and 1.55 (SN). Based on these results, we conclude that Salsa6f in rods is
sensitive enough to detect changes in intracellular Ca2+ from ~100 nM – 3 µM, a range consistent
with physiologic Ca2+ concentrations previously measured in dark-adapted rods.139,140,148 This
observation suggests that each distinct compartment in rod photoreceptors likely possesses
unique Ca2+ buffering capacities owing to their intracellular environment. Furthermore, it highlights
the importance of calibrating the Ca2+ indicator within the chosen cell or sub-compartment to
guarantee precise cytosolic Ca2+ measurements in both physiological and pathophysiological
states.

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Figure 4.6. Determination of absolute Ca2+ concentrations in rod compartments. (A) Representative
2-photon images of in-situ calibrations performed on retinal slices from transgenic Salsa6f mice under four
distinct external Ca2+ conditions (0 nM, 150 nM, 371 nM, and 1.35 µM). Slices underwent a 45-min
incubation in the specified external free Ca2+ concentrations, in the presence of 10 µM ionomycin, 5 µM
thapsigargin, 10 µM CCCP, and 10 µM nigericin, before imaging. As the free Ca2+ concentration increases,
the GCaMP6f fluorescence signal intensifies while the tdTomato signal remains constant. Top panels,
GCaMP6f (pseudo-color images, LUT Fire, color bar 0-4095). Bottom panels, tdTomato (gray). Scale bar
= 10 µm. OS, outer segment; IS, inner segment; CB, cell body; SN, synaptic terminal. (B-E) Normalized
GCaMP6f and tdTomato intensities in each rod compartment (OS, IS, CB, ST) were used to calculate the
steady-state Salsa6f ratios (GCaMP6f/tdTomato) across 10 free Ca2+ solutions (17 nM, 38 nM, 65 nM, 100
nM, 150 nM, 225 nM, 351 nM, 602 nM, 1.35 µM, and 39 µM). These ratios were then plotted against free
Ca2+ concentrations and fitted with a four parameter Hill equation to obtain Kd and Hill coefficients for each
rod compartment. These determined variables will aid in the conversion of GCaMP6f/tdTomato ratios into
approximate cytoplasmic free Ca2+ concentrations specific to each individual rod compartment. The
goodness of fit (R2) is displayed on each graph, and Kd and Hill coefficients are presented in the bottom
right corner. Data are representative of three independent experiments (n = 3, 45 rod cells per solution).
4.5 Discussion
Research on Ca2+ dynamics in vertebrate rod photoreceptors has traditionally utilized
fluorescent Ca2+ dyes141,145,348,352 and genetically encoded Ca2+ indicators (GECIs)358,359,396 to
monitor individual compartments. However, a comprehensive understanding of Ca2+
homeostasis in the polarized rod requires simultaneous examination of Ca2+ dynamics in all
101 102 103 104 105
0.0
0.2
0.4
0.6
0.8
1.0
1.2
1.4
Ca2+ external (nM)
GCaMP6f/tdTomato (F/F0)
Cell Body
Kd= 647.2nM
Hill (n) = 1.148
R2=0.95
101 102 103 104 105
0.0
0.5
1.0
1.5
Ca2+ external (nM)
GCaMP6f/tdTomato (F/F0)
Inner Segment
Kd= 614.6nM
Hill(n) = 1.4
R2=0.95
101 102 103 104 105
0.0
0.2
0.4
0.6
0.8
1.0
Ca2+ external (nM)
GCaMP6f/tdTomato (F/F0)
Synapse
Kd = 467.7nM
Hill (n) = 1.567
R2=0.96
A O
S
I
S
C
B
S
T
tdTomato
LUT = Fire
0 nM Ca2+ 150 nM Ca2+
OS
IS
CB
SN
GCaMP6f
371 nM Ca2+ 1.35 µM Ca2+
OS
IS
CB
SN OS
IS
CB
SN
OS
IS
CB
SN
D
B
101 102 103 104 105
0.0
0.7
1.4
Ca2+ external (nM)
GCaMP6f/tdTomato (F/F0)
Outer Segment
Kd = 318.8nM
Hill (n) = 1.976
R2=0.94
C E
0
4095
101 102 103 104 105
0.0
0.5
1.0
1.5
Ca2+ external (nM)
GCaMP6f/tdTomato (F/F0)
Inner Segment
Ratio
101 102 103 104 105
0.0
0.5
1.0
1.5
Ca2+ external (nM)
GCaMP6f/tdTomato (F/F0)
Inner Segment
Ratio
101 102 103 104 105
0.0
0.5
1.0
1.5
Ca2+ external (nM)
GCaMP6f/tdTomato (F/F0)
Inner Segment
Ratio
101 102 103 104 105
0.0
0.5
1.0
1.5
Ca2+ external (nM)
GCaMP6f/tdTomato (F/F0)
Inner Segment
Ratio

116
compartments. Here, we introduce a transgenic mouse line, Salsa6fiCre, one of the first
mammalian models to express a ratiometric Ca2+ indicator in all rod compartments. This mouse
line enables comprehensive and accurate monitoring of cytoplasmic Ca2+ changes throughout
the entirety of the rod photoreceptor. Our study presents a versatile protocol for investigating
Ca2+ signaling in different regions of rods, adaptable to various physiological and
pathophysiological conditions. We demonstrate the utility of Salsa6fiCre in diverse genetic
backgrounds, including light-insensitive (Gnat1-/-379 and RhoREY/REY (REY)67) and congenital
stationary night blindness (Nckx1-/-
)
173 models. We also emphasize the importance of in-situ
calibration of the Ca2+ sensor to ensure accurate intracellular Ca2+ measurements. Lastly, our
work confirms several Ca2+-related phenomena in rods, such as spontaneous Ca2+ transients in
light-adapted healthy rods and the role of mitochondria in regulating Ca2+ flux between the RIS
and ROS in both healthy and diseased rods. Insights gleaned from using this protocol may
elucidate how disruptions in Ca2+ signaling in the ROS trigger cell death. These findings could
guide the development of therapeutic strategies aimed at improving visual acuity by restoring
Ca2+ homeostasis in photoreceptors.
Rod photoreceptors, unlike typical neuronal cells, are sensitive to light. This light sensitivity
poses significant challenges for experiments aimed at exploring rod physiology in darkness, as
conventional optical imaging systems, whether using white light or laser illumination, trigger
photoresponses. Two-photon microscopy, utilizing infrared lasers beyond the visible spectrum,
offers a promising solution by reducing the likelihood of exciting the light-sensitive tissue during
imaging. In the present study, we utilized the 960 nm wavelength for 2-photon imaging of
Salsa6fiCre rods, which greatly exceeds rhodopsin’s peak sensitivity at ~497 nm. Despite this,
previous research has demonstrated that high-energy illumination could inadvertently activate
rods through direct 2-photon excitation,
382,390,398 likely due to the high concentration of rhodopsin
in the ROS (8.2 mM).12 This effect may be further amplified by the indirect excitation of rhodopsin
from photons emitted by GCaMP6f fluorescence during imaging of Salsa6fiCre retinal slices.
382 As

117
a result, studying physiological Ca2+ dynamics in Salsa6fiCre rods beyond the mesopic regime
using 2-photon microscopy presents significant challenges. Nevertheless, adopting strategies
that reduce 2-photon laser intensity and increase scan speed and frame rate during time-lapse
acquisition could reduce direct and indirect photoreceptor stimulation. In addition, using infrared
light and infrared eyepieces can enable localization of the focal plane in dark-adapted retinal
slices before 2-photon imaging. Furthermore, the use of near-infrared genetically encoded Ca2+
indicators401 can reduce indirect excitation of rhodopsin. Despite concerted efforts to manage
rhodopsin activation, complete avoidance remains difficult with current methods. A different
approach to addressing the challenges associated with imaging the retina involves utilizing lightinsensitive mouse models, such as the Gnat1-/- and REY strains used in this study. These models
serve as valuable controls, aiding in the establishment of baseline dark-adapted Ca2+ levels in
rods. Determining dark-state compartmentalized Ca2+ concentrations in these light-insensitive
rods provides a benchmark against which the impact of 2-photon imaging can be evaluated. By
comparing these baseline Ca2+ levels with the dynamic dark-to-light Ca2+ changes observed in
wildtype Salsa6fiCre, researchers can gain additional insights into physiological Ca2+ levels in all
compartments.
Recent investigations have revealed the occurrence of spontaneous Ca2+ transients or
“sparks” in healthy, light-adapted zebrafish cones,
358,359 mouse cones,392,393 and isolated rod outer
segments from frogs.
352 Despite the undetermined origin of these Ca2+ flashes, their consistent
appearance across different model organisms piques significant physiological interest. In our
study, we also detected similar spontaneous Ca2+ flares in wildtype Salsa6fiCre rods (Figure 4.2),
primarily in synaptic terminals (SN) and cell bodies (CB), with sporadic occurrences in the rod
inner segment (RIS) and an absence in the rod outer segment (ROS). These events might stem
from transient Ca2+ influx through L-type voltage-gated Ca2+ channels (VGCCs), known to operate
to some extent even at hyperpolarized membrane potentials (e.g. -60 to -70 mV).
402-404
Alternatively, the transients may arise from internal Ca2+ mobilization processes, specifically

118
through ryanodine receptor (RyR)-mediated Ca2+-induced Ca2+ release (CICR),
351,405 or via storeoperated Ca2+ entry (SOCE) through Ca2+ channels or transient receptor potential (TRP)-like
channels.
143,148 RyRs, present in photoreceptor endoplasmic reticulum (ER), contribute to
maintaining Ca2+ homeostasis for normal function in dark and low light conditions405,406 by
amplifying VGCC-driven Ca2+ influx through CICR. This process might explain the spontaneous
Ca2+ “sparks” observed, arising either from RyR’s constitutive activity at hyperpolarized
potentials143 or from CICR following VGCC’s transient activation.407 Conversely, in saturating light
conditions, the depletion of ER stores induces SOCE via Ca2+ channels and/or TRP-like channels
to prevent extended intracellular Ca2+ decreases.
148 The spontaneous Ca2+ flares we observed in
Salsa6fiCre synapses and cell bodies may be mediated by SOCE, given the essential role of Ca2+
for gene expression408,409 and modulating tonic synaptic signaling under saturating
conditions.
107,143,410 Although the specific contributors to spontaneous Ca2+ “sparks” remain
elusive, utilization of ryanodine receptor blockers, L-type VGCC antagonists, or other relevant
pharmacological agents could aid in elucidating the origins of these flashes. Furthermore,
crossbreeding Salsa6fiCre mice with other mouse models of inherited retinal degeneration holds
promise in uncovering the significance of these flares in both physiological and pathophysiological
contexts.
The precise regulation of Ca2+ ions is essential for the functionality and survival of
photoreceptors and is distinctly compartmentalized.134,145,348 For instance, Ca2+ fluctuations
originating in either the outer or inner segments do not easily propagate through the connecting
cilium that bridges these two compartments.
145,352,359 The high density of mitochondria in the
ellipsoid region of cone inner segments has emerged as a critical factor in establishing functional
and spatially restricted Ca2+ domains.
359,396 The specific localization of mitochondria appears to
help limit cytosolic Ca2+ diffusion between outer and inner segments by acting as a Ca2+
reservoir.
359,396 Consistent with these findings, our study further confirms the role of mitochondria
as barriers impeding Ca2+ flux between the RIS and ROS in rods. Inhibition of mitochondrial Ca2+

119
uptake with Ru360, a mitochondrial Ca2+ uniporter (MCU) inhibitor, disrupts the formation of
distinct cellular Ca2+ domains upon KCl stimulation, as evidenced by the increase in GCaMP6f
fluorescence intensity observed in functional murine ROS (Figure 4.4B). Despite the higher
mitochondrial density397 and energy requirements411,412 in vertebrate cones compared to rods, the
role of mitochondria in sequestering Ca2+ ions appears to be integral in establishing
compartmentalized Ca2+ gradients in both types of photoreceptors. This recognition of
mitochondria as pivotal regulators of compartmentalized Ca2+ concentrations adds a new
dimension to our understanding of photoreceptor physiology. Future research investigating
mitochondria’s involvement in handling Ca2+ flux between the ROS and RIS in healthy and
diseased rods holds promise for formulating potential therapeutic interventions.
Paradoxically, both excessive Ca2+ influx136 and abnormally low Ca2+ levels158 have been
implicated in photoreceptor cell death. However, the precise mechanism by which aberrant Ca2+
concentrations within the ROS are transduced to the RIS to induce cell death remains elusive.
Under normal physiological conditions, Ca2+ ions are cleared from the ROS via the Na+
/Ca2+-K+
exchanger 1 (NCXK1).134 Surprisingly, rods lacking NCKX1 retain partial functionality, albeit with
delayed photoresponse recovery, and degenerate very slowly,173 indicating the presence of
alternative mechanisms for Ca2+ extrusion from the ROS. We hypothesize that mitochondria may
serve as compensatory agents in buffering elevated Ca2+ levels when ROS mechanisms are
insufficient, potentially contributing to the slow response recovery seen in Nckx1-/- rods. This extra
Ca2+ burden may lead to mitochondrial dysfunction,
157,413 subsequently disrupting Ca2+
homeostasis in the ER and/or RIS during the disease progression, ultimately activating calpains
and caspases, the executioners of cell death.154,157,414 Indeed, in light-adapted rods, the elevation
of ROS Ca2+ levels following KCl perfusion in 2-month-old Nckx1-/-
/Salsa6fiCre rods indicates
disrupted mitochondrial buffering and Ca2+ compartmentalization early in the disease (Figures
4.3A and 4.4C). In a somewhat analogous case, Nckx2-/- mice, lacking NCKX2 extrusion in cone
outer segments, also exhibit delayed response recovery but without cone degeneration.415 It is

120
plausible that the superior Ca2+-buffering capacity of cone mitochondria,146,416 compared to that of
rods, accounts for this discrepancy. This suggests that enhanced Ca2+ buffering might shield
cones from vulnerabilities to aberrant Ca2+ changes, which could otherwise be harmful to rods.
Interestingly, in double knockout Nckx1-/-
/Mcu-/- mouse rods, Ca2+ sequestration by MCU is not
involved in photoresponse recovery,396 indicating the existence of alternative pathways for Ca2+
entry into mitochondria. Taken together, Nckx1-/- rods appear to experience mitochondrial
dysfunction during disease progression, warranting further investigation into Ca2+ dynamics and
its involvement in cell death pathways.
The crossbreeding of Salsa6fiCre mice with mouse models of inherited retinal degeneration,
particularly rd1 or arrestin (Arr1) knockout mice, offers compelling opportunities for investigating
the mechanisms underlying rod degeneration. In rd1 mice, degeneration is attributed to Ca2+
overload, stemming from a loss-of-function mutation in the beta subunit of phosphodiesterase-6
(Pde6b), resulting in cGMP accumulation.363,364 Conversely, in Arr1-/- rods, characterized by
uncontrolled phototransduction (“equivalent light”),374,375 a Ca2+ deficiency is implicated in cell
death.188,372 Recent multi-omics analysis of rd1 mouse retinae revealed early occurrences of
mitochondrial stress and metabolic disruptions,383 suggesting a link between elevated Ca2+ in the
ROS and mitochondrial dysfunction in the RIS as precursors to cell death. However, the precise
involvement of low Ca2+ levels in the ROS in either instigating mitochondrial or ER dysfunction or
initiating cell death remains undetermined. Visualizing Ca2+ dynamics and compartmentalization
with Salsa6fiCre will aid in clarifying these hypotheses. This protocol serves not only to deepen our
understanding of rod physiology, but also to establish a foundation for unraveling the intricate
interplay of Ca2+ signaling pathways that contribute to rod degeneration.

121
4.6 Supplemental Information
Figure. S4.1. Assessment of retinal preparation and Salsa6f in rods. (A) Representative vibratome
images of a Salsa6fiCre retinal slice. Pseudo-color image (left panel, LUT Fire, color bar 0-4095) shows the
light-adapted Ca2+ levels from GCaMP6f fluorescence. The middle and left panels display the Ca2+-
insensitive fluorophore tdTomato (gray) and the merged GCaMP6f-tdTomato signals, respectively. White
rectangles highlight rod OS with elevated GCaMP6f signals, indicating tissue damage during the sectioning
process. To minimize vibrational damage, a vertical slicing method was used. Scale bar = 5 µm. (B)
Representative images of retinal slices from Salsa6f heterozygous (Salsa6f+) or homozygous (Salsa6f+/+)
mice. Light-exposed rods from homozygous Salsa6fiCre mice exhibit elevated fluorescence in all
compartments compared to heterozygous mice. n = 6. Scale bar = 5 µm. (C) Normalized tdTomato
fluorescence from light-exposed homozygous or heterozygous Salsa6fiCre retinas (n = 4, 64 rods). Salsa6f+/+
appears ~ 2-fold brighter than Salsa6f+. The tdTomato mean fluorescence value was extracted from the
RIS compartment and normalized to the max pixel intensity value for a 12-bit image (4094). OS, outer
segment; IS, inner segment; CB, cell body; SN, synaptic terminal.
A
B
OS
IS
CB
SN
GCaMP6f tdTomato GCaMP6f-tdTomato
0
4095
GCaMP6f tdTomato GCaMP6f-tdTomato
OS
IS
CB
SN
0
4095
Salsa6f+ Salsa6f+/+
OS
IS
CB
0 SN
4095
C
Salsa6f+/+
Salsa6f+
0
20
40
60
80
100
tdTomato Fluorescence (A.U.)
Transform of Data 2
Salsa6f+/+
Salsa6f+

122
Figure. S4.2. Ca2+ measurement pipeline. (A) Images of a light-adapted retinal slice expressing the Ca2+
indicator Salsa6f immediately before time-lapse imaging. GCaMP6f is presented in a pseudo-color (Fire,
LUT, scale 0-4095), and tdTomato is displayed in gray. The merged image is shown in the far-right panel.
Three distinct regions of interest (ROIs) are delineated around rod synaptic terminals (SN), and their
fluorescence profiles are recorded over a 5-min period (300 s) for each channel. A white rectangle indicates
the area where background fluorescence measurements are acquired for each channel. OS, outer
segment; IS, inner segment; CB, cell body. Scale bar = 5 µm. (B-D) Upper graphs present raw, prenormalized ROI data from GCaMP6f and tdTomato time-series recordings captured over a 5-min period.
Background fluorescence measurements, corresponding to the white rectangle in (A), are shown for
GCaMP6f (purple) and tdTomato (gray). The fluorescence intensity profiles for tdTomato (bright red) and
GCaMP6f (bright green) are also presented for each ROI. The lower graphs illustrate the normalized
GCaMP6f/tdTomato ratios for each ROI (1-3). This image analysis pipeline enables accurate examination
of Ca2+ dynamics within specific subcellular compartments.
0 100 200 300
0
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800
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Time (s)
GCaMP6f/tdTomato (F/F0)
Green Background
Red background
ST red
ST Green
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Time (s)
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Fluorescence Intensity A.U.
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Green Background
Red background
ST red
ST Green
0 100 200 300
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Fluorescence Intensity A.U.
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ST red
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Time (s)
Fluorescence Intensity A.U.
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Red background
ST red
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A
GCaMP6f tdTomato GCaMP6f-tdTomato
OS
IS
CB
SN
0 100 200 300
0.0
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1.2
Time (s)
GCaMP6f/tdTomato (F/F0)
ST 2
0 100 200 300
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1.0
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Time (s)
GCaMP6f/tdTomato (F/F0)
ROI 1 ROI 2 ROI 3
B C D
0
4095
–Background
–SN
–SN
–Background
–Background
–SN
–SN
–Background
–Background
–SN
–SN
–Background
0 100 200 300
0.0
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1.2
Time (s)
GCaMP6f/tdTomato (F/F0)
ST 2
Ratio
0 100 200 300
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123
Figure. S4.3. Unique spatiotemporal profiles of Ca2+ flares. Representative graphs depict spontaneous
Ca2+ flares from (A) inner segments and (B) cell bodies of rod photoreceptors expressing Salsa6f over a 3-
min period (180 s). These Ca2+ flashes exhibit diverse patterns and durations yet maintain relatively
consistent amplitudes within each specific compartment. n = 3.
0 30 60 90 120 150 180
0.0
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0.4
0.6
CB 1
Time (s)
Ratio
GCaMP6f/tdTomato
0 30 60 90 120 150 180
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Time (s)
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GCaMP6f/tdTomato
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Time (s)
Ratio
GCaMP6f/tdTomato
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0.0
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GCaMP6f/tdTomato
CB 1
Inner Segment Cell Body
0 30 60 90 120 150 180
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CB 1
Time (s)
Ratio
GCaMP6f/tdTomato
0 30 60 90 120 150 180
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A B

124
CONCLUDING REMARKS
This dissertation endeavors to advance our understanding of the intricate mechanisms
that contribute to rod photoreceptor structure and function, as well as the consequences of
inherited genetic mutations on these critical biological elements. By utilizing a variety of
methodologies, including live cell imaging, western blots, immunocytochemistry, and isoelectric
focusing (IEF), along with fluorescent-reporter and retinal degeneration mouse lines, we were
able to investigate three fundamental areas: 1) the mechanisms driving incorporation of rhodopsin
into rod outer segments, 2) the role of rhodopsin’s conserved ERY motif in mediating proteinprotein interactions involved in the phototransduction cascade, and 3) the compartmentalization
of Ca2+ dynamics and homeostasis. Through a series of in-depth studies employing multimodal
approaches, this body of work sheds light on the complex systems that sustain photoreceptor
morphology and physiology and the impact that disruptions can have on rod’s morphofunctional
equilibrium. Our findings not only deepen our comprehension of photoreceptor cell biology but
may also help identify promising avenues for developing innovative therapeutic approaches to
address retinal degenerative diseases.
The use of our novel Rhodopsin-Timer fusion protein to study rhodopsin incorporation has
provided significant insights into the architecture of the ROS across different lighting conditions
and genetic backgrounds. Our research demonstrates that the incorporation of rhodopsin, a
protein critical for light sensitivity and ROS structural integrity, is modulated by both external
lighting cues and internal circadian clocks. Moreover, we identify regions along the ROS with
reduced rhodopsin content as potential points of vulnerability in its structure, suggesting that
irregular rhodopsin distribution in the ROS may contribute to the etiology of retinal pathogenesis.
A promising future research direction is the elucidation of the circadian elements facilitating
rhodopsin incorporation into the ROS. Given that BALB/c and C57BL/6 mouse strains used in our

125
study lack melatonin but express dopamine, investigating the potential regulatory roles of
melatonin and dopamine, both neuromodulators of retinal clocks, in rhodopsin incorporation,
could offer valuable insights into the clock-related mechanisms governing ROS morphogenesis.
Additionally, crossbreeding Rhodopsin-Timer with clock-deficient mice, mice harboring
membrane trafficking or ciliogenesis disorders, or mice carrying mutations in other
phototransduction proteins, may yield deeper insights into the cellular mechanisms underlying
rhodopsin delivery to the ROS and its implications for disease pathogenesis. Advancing this
research, the application of light-sheet microscopy to image live, ex-vivo eyes from RhodopsinTimerB mice emerges as a promising technique for real-time observation of ROS renewal and
rhodopsin incorporation.
In examining the highly conserved ERY motif in rhodopsin using the RhoREY/REY (REY)
mouse line, the findings presented in this dissertation call into question established beliefs
regarding the motif’s role in promiscuous coupling with essential phototransduction proteins. Our
research demonstrates that, contrary to outcomes from heterologous expression studies and
theoretical models, in-vivo expression of REY-rhodopsin confirms its exclusive interaction with
transducin, rather than with rhodopsin kinase (GRK1) and arrestin. Notably, our findings offer
empirical evidence supporting the hypothesis of a stabilizing, pre-assembled complex involving
rhodopsin’s ERY motif and transducin in darkness, a phenomenon previously elusive to direct
detection through NMR and chromatography techniques due to its transient nature. Future
research avenues should explore the potential nuances in arrestin translocation and GRK1
interaction. For instance, exposing REY mice to prolonged photopic illumination might uncover
previously undetected anomalies in arrestin translocation, an effect not observed within the 30-
min timeframe of our experimental conditions. Furthermore, a thorough assessment of GRK1
activity, using isolated ROS membranes from REY retinas supplemented with purified murine
GRK1, could shed light on any subtle disruptions to GRK1 functionality not identified by IEF.
Finally, the documented increase in intracellular Ca2+ levels in light-adapted REY rods, compared

126
to transducin knockout rods as detailed in Chapter 4, warrants further exploration, despite the
current uncertainty regarding the precise pathways involved.
The creation of our Salsa6fiCre transgenic mouse line has opened up new avenues for
exploring Ca2+ dynamics across all compartments, adding to our understanding of how Ca2+
imbalances contribute to degenerative retinal diseases. Our findings reaffirm recognized Ca2+
signaling behaviors observed in cone photoreceptors, including spontaneous Ca2+ flares and
mitochondrial Ca2+ management, highlighting the utility of Salsa6fiCre for investigating Ca2+
dynamics. Integral to our Ca2+ imaging methodology is the in-situ Ca2+ calibration of Salsa6f,
which ensures precise measurement of Ca2+ concentrations within the distinct microenvironments of the polarized rod. Future studies, using 2-photon microscopy with specific
imaging parameters tailored to reduce physiological interference, will be capable of visualizing
Ca2+ changes in dark-adapted rods in response to light exposure. Furthermore, forthcoming
studies will illuminate whether, and in what ways, variations in Ca2+ levels (either high or low), due
to mutations affecting the phototransduction in the ROS, contribute to cellular dysfunction. Of
special interest is our ongoing investigation into Ca2+ dynamics that underpin the gradual
degeneration observed in Nckx1-/- rods. By mapping the spatiotemporal Ca2+ dynamics and
concentrations that precipitate cell death, alongside monitoring caspase and calpain activity
through in-situ enzymatic assays, future work can help clarify how abnormal Ca2+ levels induce
photoreceptor cell death.
This dissertation marks the culmination of my journey thus far in the field of visual
neuroscience, a path that has been both challenging and deeply rewarding. Under Dr. Chen’s
guidance, I’ve had the privilege of spearheading the development of novel techniques, including
the lyophilized peeling process and the Ca2+ imaging pipeline. The terzetto of research endeavors
presented here did more than just refine my technical and critical thinking skills; it instilled a
patience learned through countless hours of troubleshooting, honed through challenges posed by
the notoriously tricky IEF technique. My exploration into the complex landscape of retinal research

127
has fostered a profound appreciation for the intricacies of the rod photoreceptor cell, guiding my
ambition to uncover fundamental processes that could illuminate therapeutic opportunities. As I
present this work, it is my hope that it will add valuable insights to the scientific community and
contribute to shaping future research directions.

128
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APPENDIX A: MANUSCRIPT INFORMATION
Manuscript adapted for Chapter 2:
Light regulation of rhodopsin distribution during outer segment renewal in murine rod
photoreceptors.
Current Biology (2024). Manuscript accepted February 2024.
https://doi.org/10.1016/j.cub.2024.02.070
Kasey Rose1
, Natalie Chen1
, Andrey Andreev2
, Jiayan Chen1
, Vladimir J Kefalov3
, and
Jeannie Chen1,4,*
1
Zilkha Neurogenetic Institute, Department of Physiology and Neuroscience, Keck School of
Medicine of University of Southern California, Los Angeles, CA, 90033, USA.
2
Division of Biology and Biological Engineering, California Institute of Technology, Pasadena,
CA, 91125, USA.
3
Department of Ophthalmology, Gavin Herbert Eye Institute, University of California, Irvine,
CA, 92697, USA.
Author contributions: Conceptualization, K.R., Jeannie C.; Methodology, K.R., Jeannie C., and
V.J.K.; Formal Analysis, K.R., N.C., A.A., and Jeannie C.; Investigation, K.R., N.C., Jiayan C,
A.A.; Writing — Original Draft, K.R. and Jeannie C.; Writing — Review & Editing, K.R., A.A.,
N.C., Jeannie C., and V.J.K.; Funding Acquisition, Jeannie C.; Resources, Jeannie C. and
V.J.K.
Acknowledgements: This work was supported by National Institutes of Health Grants
EY12155, EY027193, EY027387 (Jeannie C.) and Vision Core Grant P30EY029220 to Roski
Eye Institute, USC. We thank Dr. Seth Ruffins from USC’s Optical Imaging Facility for
technical help; Ying Liu from USC Translational Research In Vivo Core for assistance with
ERG; Drs. Sowmya Lokappa, Hui Xu, and Spyridon Michalakis for proofreading and helpful
comments on the manuscript; Dr. Shai Porat for providing coding support (CircaCompare);
Dr. Hui Xu for technical contributions to this work (epon block preparation); and Dr. Janos
Peti-Peterdi and Dr. Georgina Gyarmati for imaging support. We are also grateful to our
summer research scholar Rahul Kumar for providing technical support in the early stages of
the project.

Abstract (if available)

Abstract The rod photoreceptor, a highly polarized light-sensitive cell in the retina, features a unique structure, the rod outer segment (OS), that houses rhodopsin, the photopigment. Maintenance of proper OS length and rhodopsin incorporation into new discs is essential for rod health and function. Light exposure triggers a conformational change in rhodopsin, initiating the phototransduction cascade and movement of signaling proteins (e.g. arrestin, transducin) within distinct cellular compartments. Mutations, particularly in conserved residues like the E(D)RY motif within rhodopsin, can compromise its stability and interaction with key phototransduction proteins. Furthermore, calcium (Ca2+) ions, vital intracellular messengers, modulate phototransduction cascade gain, with Ca2+ homeostasis critical for rod cell survival. Dysfunction in any of these systems can impair dim light vision and lead to blindness. Understanding the mechanisms regulating rod structure and function is key to addressing inherited retinal degeneration and developing novel treatments. Here, we investigate mechanisms governing rhodopsin incorporation into the OS using a novel fluorescent rhodopsin fusion protein, revealing insights about rhodopsin incorporation dynamics under various lighting regimens and genetic backgrounds. We also explore how a charge reversal mutation (ER to RE) in rhodopsin’s ERY motif affects its interaction with cognate proteins, suggesting a potential stabilizing dark-state rhodopsin/transducin complex for rapid phototransduction activation. Additionally, we present a protocol enabling spatiotemporal measurement of Ca2+ dynamics in all rod compartments simultaneously, offering a novel framework with which to understand rod physiology and pathophysiology. Our multimodal approach sheds new light on physiologically important structural and functional mechanisms, setting the stage for advanced research.

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Asset Metadata

Creator Rose, Kasey (author)

Core Title Multimodal investigation of rod photoreceptor structure and function in healthy and diseased states

School College of Letters, Arts and Sciences

Degree Doctor of Philosophy

Degree Program Neuroscience

Degree Conferral Date 2024-05

Publication Date 05/17/2024

Defense Date 04/26/2024

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

Tag arrestin,calcium,calcium homeostasis,calcium imaging,circadian clock,CNG channels,E(D)RY motif,genetically encoded calcium indicator,G-protein coupled receptor,NCKX,night blindness,OAI-PMH Harvest,photoreceptor,phototransduction,pre-coupling,primary cilia,protein translocation,retina,Retinal degeneration,retinitis pigmentosa,rhodopsin,rod,rod outer segment,transducin,two-photon microscopy,vision

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Language English

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