LIGHT-INDEPENDENT PATHOLOGY OF RHODOPSIN MISLOCALIZATION
by
PHILIP EDWARD ROPELEWSKI
Submitted in fulfillment of the requirements for the degree of Doctor of Philosophy
Department of Pharmacology
CASE WESTERN RESERVE UNIVERSITY
May 2020
CASE WESTERN RESERVE UNIVERSITY SCHOOL OF GRADUATE STUDIES
We hereby approve the thesis/dissertation of Philip Edward Ropelewski candidate for the degree of Doctor of Philosophy.
Committee Chair Monica Montano
Committee Member Marvin Nieman
Committee Member Alan Tartakoff
Committee Member Yoshikazu Imanishi
Committee Member Johannes von Lintig
Date of Defense April 1st, 2020.
*We also certify that written approval has been obtained for any proprietary material contained therein.
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Table of Contents
List of Tables…………………………………………………………...……...….....viii
List of Figures………………………………………………………..……...…..……ix
Acknowledgements…………………………………………………….……...... xi
List of Abbreviations………………………………………………….……..……...xiii
Abstract…………………………………………………………….....…………….…xv
Chapter 1: Introduction……………………………………………………………….1
1.1 Efficient rhodopsin trafficking is required for vision……………….1
1.2 Mechanism and Pathway of rhodopsin biosynthesis and
transport………………………………………………………….……..….2
1.3 Rhodopsin mutations are the major causes of inherited
blindness…………………………………………………………….……..7
1.4 Electrochemical interactions between IS and OS are essential for
vision………………………………...……………………………………...8
1.5 Currently available approaches to study rhodopsin
transport…………………………………………………………………....9
1.6 Aims of dissertation………………………………………………….….14
Chapter 2: Mislocalized rhodopsin is degraded via the lysosomal pathway
2.1 Introduction……………………………………………………………....15
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2.2 Materials and Methods………………………………………………….16
2.2.1 Animals…………………………………………………………..16
2.2.2 Reagents…………………………………………………………17
2.2.3 Molecular cloning………………………………………………..17
2.2.4 Transgenesis of Xenopus laevis………………………………17
2.2.5 Photoconversion of Dendra2 in tadpole retina…………….…18
2.2.6 Treatment of tadpoles by lysosome inhibitor……………...... 19
2.2.7 Preparation of retina explant for confocal imaging…………..19
2.2.8 Confocal microscopy, Image analysis, quantification, and
statistical analysis………………………………………………...……20
2.3 Results and Discussion………………………………………………...21
2.3.1 Class I mutant rhodopsin is degraded in lysosomes………..21
2.3.2 Mislocalized rhodopsin is trafficked from the plasma
membrane to intracellular lysosomes………………………………..26
2.4. Conclusions……………………………………..……………………….28
Chapter 3: NKA is co-degraded with class I mutant rhodopsin and becomes downregulated on the IS PM of rod photoreceptors…………………………..30
3.1 Introduction……………………………………………………………….30
3.2 Materials and methods………………………………………………….31
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3.2.1 Animals…………………………………………………………..31
3.2.2 Reagents…………………………………………………………32
3.2.3 Transgenesis of Xenopus laevis.…………………….………..32
3.2.4 Immunohistochemistry………………………………………….33
3.2.5 Reverse transcription and quantitative PCR…………………34
3.2.6 Treatment of tadpoles by lysosome inhibitor…………………35
3.2.7 SDS-PAGE and immunoblotting………………………………35
3.2.8 Preparation of retina explant for confocal imaging…….…….36
3.2.9 Confocal microscopy, Image analysis, quantification, and
statistical analysis……………………………..………………………37
3.3 Results and discussion………………………………………………...38
3.3.1 The Na+ /K+-ATPase is downregulated in cells expressing
class I mutant rhodopsin…………….…………………..……………38
3.3.2 NKAα downregulation is correlated to the expression level of
RhoQ344ter-Dend2 in individual rods……………..…….……………..41
3.3.3 NKAα is co-trafficked with RhoQ344ter-Dend2 to lysosomes in
vivo...... 43
3.4 Conclusions………………………………………………………………45
Chapter 4: Dysfunction of NKA results in photoreceptor degeneration…..44
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4.1 Introduction………………………………………………………………46
4.2 Materials and methods…………………………………………………47
4.2.1 Animals…………………………………………………………..47
4.2.2 Reagents………………………………………………………...47
4.2.3 Transgenesis of Xenopus laevis………………………………47
4.2.4 Immunohistochemistry…………….……………………………48
4.2.5 SDS-PAGE and immunoblotting………………………………50
4.2.6 Treatment of tadpoles by NKA inhibitor………………………51
4.2.7 Confocal microscopy, Image analysis, quantification, and
statistical analysis………………………………..……………………51
4.3 Results and discussion…………………….…………………………..52
4.3.1 Rod photoreceptors expressing RhoQ344ter-Dend2 have
shorter, disorganized OSs…………………………...………………52
4.3.2 Pharmacological inhibition of NKAα with digoxin mimics the
degenerative phenotype observed in RhoQ344ter-Dend2…………..56
4.4 Conclusions……………………………………..………………………61
Chapter 5: Alternate routes of mislocalized rhodopsin removal…....……..65
5.1 Introduction………………………………………………………..…….65
5.2 Materials and methods………………..…………………….…………67
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5.2.1 Animals………………………………………………...... 67
5.2.2 Reagents…………………………………………………………68
5.2.3 Molecular cloning………………………………………………..68
5.2.4 Transgenesis of Xenopus laevis……………..………………..69
5.2.5 Immunohistochemistry………………………………………….69
5.2.6 Preparation of retina explant for confocal imaging…….…….70
5.2.7 Confocal microscopy, Image analysis, quantification, and
statistical analysis ……………………...…………..…………………70
5.3 Results and discussion……………………….………………………..71
5.3.1 IS-mislocalized rhodopsin is secreted and uptaken by RPE
cells...... 71
5.3.2 RPE cells do not contribute to the degradation of NKA under
normal physiological conditions…………………………...…………74
5.3.3 NKAα is not co-secreted along with class I mutant rhodopsin
as microvesicles……………………………………...………………..76
5.3.4 NKAα is potentially degraded within rod photoreceptors…...79
5.4 Conclusions ……………………………………………………..……….81
Chapter 6: Conclusions and Future Directions…………………..…………….84
Bibliography…………………………………………………………..………………91
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List of Tables
Table 1. Summary of effects caused by mislocalized rhodopsin or drug treatment…………………………………………………………..……………………55
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List of Figures
Figure 1. Biogenesis of rhodopsin in the endoplasmic reticulum and formation of
rhodopsin transport carriers in the Golgi………………………………………..…….3
Figure 2. Vectorial trafficking of rhodopsin from the Golgi to the distal end of the connecting cilium………………………………………………………………………..5
Figure 3. Application of Dendra2 for imaging of rhodopsin trafficking in live
Xenopus laevis rod photoreceptor cells…………………………………………...…11
Figure 4. Monitoring removal of mislocalized mutant rhodopsin (Q344ter) in live
Xenopus laevis rod photoreceptor cells……………………..………………………13
Figure 5. Class I mutant rhodopsin accumulates in lysosomes after inhibition of
lysosome-mediated degradation………….………………………………………….22
Figure 6. Mislocalized rhodopsin is trafficked from the plasma membrane to
intracellular lysosomes………………………………..………………………………25
Figure 7. Na+ /K+-ATPase (NKAα) is downregulated in rods expressing
RhoQ344ter-Dend2……………………………………….………………….…………..40
Figure 8. Degree of NKAα downregulation is dependent on the amount of class I
mutant rhodopsin…………………………………………………..………….……….42
Figure 9. RhoQ344ter-Dend2 induces internalization of NKAα………….…………..44
Figure 10. Rods expressing RhoQ344ter-Dend2 exhibit disorganized rod OSs.….53
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Figure 11. Inhibition of NKA results in disorganized rod OSs of Xenopus laevis...... 58
Figure 12. NKA inhibitors structurally distinct from digoxin induce disorganization of rod OSs……………………………………………….……………………………..60
Figure 13. Class I mutant rhodopsin is released in microvesicles before being engulfed by RPE cells………………….…………..………..………………………..72
Figure 14. NKAα is not released in extracellular vesicles under normal physiological conditions………………..……………………………………………..75
Figure 15. IS fragments containing NKAα are phagocytosed by RPE at late stage of photoreceptor degeneration……………………………………………………….77
Figure 16. NKAα proteins are degraded intracellularly in rod photoreceptors…..80
Figure 17. Destiny of mislocalized rhodopsin and Na+/K+-ATPase………………87
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Acknowledgements
I would first like to acknowledge my mentor, Dr. Yoshikazu Imanishi, for
giving me the opportunity to join his lab first as a research assistant and then as
a graduate student. I cannot thank him enough for instructing me in all sorts of
laboratory techniques, scientific writing, time management, and much more. I
consider myself extremely lucky to have to have been able to learn under him
during this adventure. I would next like to thank the members of my thesis
committee including Dr. Monica Montano, Dr. Marvin Nieman, and Dr. Alan
Tartakoff. Thank you for all of your time spent in committee meetings, your feedback, and your support. I would also like to thank Dr. John Mieyal for his support and guidance through this program.
I would like to thank all current and former members of the Imanishi lab:
Kerrie Lodowski, for teaching me retina dissection techniques and confocal microscopy, and for setting the foundation for my thesis project; Ina Nemet, for always offering her help and encouragement in my experiments, her advice, and for the kindness she showed; Guilian Tian, for all of the knowledge and secret techniques she passed to me about molecular biology, cell culture, and more;
Sanae Sakami, for her assistance with experiments, unparalleled selflessness and patience; Richard Lee, for his constant support, troubleshooting, and friendship. I would also like to thank more recent members of the Imanishi lab including Sultana Jahan, Mohammed Rashid, and Hemavathy Harikrishnan, as well as all of the student helpers, including Eric Hlosek, who have aided our lab.
Lastly, I would like to acknowledge and thank the National Institutes of Health for
xi supporting this project, including grant numbers R01EY028884, R21EY027292, and T32EY007157.
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List of abbreviations
4OH 4-hydroxytamoxifen
AAV adenosine-associated virus
BA1 bafilomycin A1
BDNF brain-derived neurotrophic factor
CNTF ciliary neurotrophic factor
EEA1 early endosome antigen 1
ER endoplasmic reticulum
ERG electroretinogram
DPF days post-fertilization
GAP GTPase-activating protein
GDNF glial-derived neurotropic factor
GPCR G protein-coupled receptor
OS outer segment
IACUC Institutional Animal Care and Use Committee
IMCD inner medullary collecting duct
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IP immunoprecipitation
IS inner segment
LAMP1 lysosome-associated membrane protein 1
NKA sodium-potassium ATPase
NS not significant
NT nontransgenic
PM plasma membrane
PR photoreceptor
Q1 retina expressing RhoQ344ter-Dend2
RP retinitis pigmentosa
RPE retinal pigment epithelium
RTC rhodopsin transport carrier
SD standard deviation
TTX 3,4,5,6-tetrahydroxylxanthone
WT wild type
XOP Xenopus opsin promoter
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Light-Independent Pathology of Rhodopsin Mislocalization
Abstract
By
PHILIP EDWARD ROPELEWSKI
Rhodopsin mislocalization is frequently observed in retinitis pigmentosa
(RP) patients. For example, class I mutant rhodopsin is deficient in the VXPX trafficking signal, mislocalizes to the plasma membrane (PM) of rod photoreceptor inner segments (ISs), and causes autosomal dominant RP.
Mislocalized rhodopsin causes photoreceptor degeneration in a manner independent of light-activation. In this thesis project, I took advantage of Xenopus laevis models of both sexes expressing wild-type human rhodopsin or its class I
Q344ter mutant fused to Dendra2 fluorescent protein to characterize a novel light-independent mechanism of photoreceptor degeneration caused by mislocalized rhodopsin. I found that rhodopsin mislocalized to the PM is actively internalized and transported to lysosomes where it is degraded. This degradation process results in the downregulation of a crucial component of the photoreceptor IS PM: the sodium-potassium ATPase α-subunit (NKAα). The downregulation of NKAα is not because of decreased NKAα mRNA, but due to cotransport of mislocalized rhodopsin and NKAα to lysosomes or
xv autophagolysosomes. In a separate set of experiments, I found that class I mutant rhodopsin, which causes NKAα downregulation, also causes shortening and loss of rod outer segments (OSs); the symptoms frequently observed in the early stages of human RP. Likewise, pharmacological inhibition of NKAα led to shortening and loss of rod OSs. These combined studies suggest that mislocalized rhodopsin leads to photoreceptor dysfunction through disruption of the PM protein homeostasis and compromised NKAα function. In these disorders, rhodopsin-laden microvesicles are secreted into the extracellular milieu by afflicted photoreceptor cells. Through fluorescent labeling techniques, I demonstrated that RPE cells are capable of engulfing these vesicles which contain rhodopsin, but not NKA. Under normal physiological conditions, NKA is not shed as microvesicles to extracellular space, but is degraded intracellularly.
Therefore, there is a unique sorting mechanism which packages IS PM- mislocalized rhodopsin, but not NKA, into vesicles for secretion. These studies provide novel insights into protein homeostasis in the photoreceptor IS PM.
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Chapter 1: Introduction
The material in this Chapter was adapted from Nemet, I, Ropelewski P, and
Imanishi Y, 2015: Rhodopsin trafficking and mistrafficking: signals, molecular components, and mechanisms. Progress in molecular biology and translational science 132:39-71, and also from Nemet I, Ropelewski P, and Imanishi Y, 2015:
Applications of phototransformable fluorescent proteins for tracking dynamics of cellular components. Photochemistry and Photobiology Sci. 14(10); 1787-1806.
1.1 Efficient rhodopsin trafficking is required for vision
Photoreceptor cells are polarized neurons which are responsible for the visualization of light in a process known as phototransduction (Frederick et al.,
2019). To accomplish this feat, photoreceptors of vertebrate species utilize photosensitive outer segments (OSs), which is a modified ciliary compartment housing the phototransduction machinery (Baehr et al., 2019). Rhodopsin, the visual pigment for rod photoreceptor cells, is a seven transmembrane G protein- coupled receptor (GPCR) and comprises ~90% of the membrane proteins of the
OS (Papermaster and Dreyer, 1974; Nathans, 1992; Kevany and Palczewski,
2010). For efficient photon capture, rhodopsin concentration in the OSs can reach as high as 4 mM (Nickell et al., 2007), and to accomplish such high concentration, new rhodopsin molecules must be constantly synthesized at the
1 high rate of more than 1 million molecules per day. Because the OS itself does not contain ER or ribosomes for protein synthesis, rhodopsin and all other OS components must first be synthesized in the inner segment (IS) before being transported to the OS via the connecting cilium (Wang and Deretic, 2014; Nemet et al., 2015). To facilitate rhodopsin trafficking to the OS, its ciliary targeting motif
VXPX exists on the C-terminus (Mazelova et al., 2009). This motif, which is conserved for other ciliary localized GPCRs (Wang and Deretic, 2014), allows rhodopsin to be efficiently transported to the connecting cilia and incorporated into growing evaginations of plasmalemma, which mature into disk membranes
(Nemet et al., 2015).
1.2 Mechanism and Pathway of rhodopsin biosynthesis and transport
Rhodopsin is first synthesized in the endoplasmic reticulum (ER) before entering the Golgi apparatus (Figure 1). In the lumen of the ER, rhodopsin is glycosylated at asparagines 2 and 15 on its N-terminus which is critical for the protein stability (Tam and Moritz, 2009) and plays a role in its trafficking and incorporation into the OS (Murray et al., 2015). Rhodopsin enters the Golgi apparatus through COPII-mediated vesicle budding (Barlowe et al., 1994;
Schekman and Orci, 1996). Initial studies of post-Golgi rhodopsin transport used a cell-free system, which reconstitutes rhodopsin transport carrier (RTC) budding in vitro (Deretic and Papermaster, 1991). This model was effective in identifying the interaction between the small GTPase Arf4 and the C-terminus of rhodopsin
(Deretic et al., 2005). When Arf4 was rendered deficient of GTP hydrolysis in
2
Figure 1. Biogenesis of rhodopsin in the endoplasmic reticulum and formation of rhodopsin transport carriers in the Golgi. After folding in the ER, rhodopsin is transported to the Golgi where rhodopsin interacts with Arf4, FIP3, ASAP1, Rab8 and Rabbin8. Then, rhodopsin transport carriers bud off from the trans-Golgi network. IFT20 and Rabaptin5 probably bind to the carrier, but the timing of the binding is unclear.
3
Xenopus rod photoreceptors, rhodopsin trafficking was disrupted
(Mazelova et al., 2009). Using mouse inner medullary collecting duct (IMCD) cell model, it was demonstrated that ASAP1, an Arf4 GTPase-activating protein
(GAP), helps rhodopsin enter and accumulate in the primary cilia, and knockdown of ASAP1 prohibits entry to the cilia and causes accumulation of rhodopsin at its base (Wang et al., 2012). Using co-IP assays, ASAP1 was revealed as a scaffold protein for the Rab11a-Rabin8-Rab8 complex which departs from the trans-Golgi network and directs rhodopsin to the cilia (Wang et al., 2012). After departure from the Golgi, rhodopsin-laden vesicles are thought to be transported to the cilia via microtubules (Figure 2). As evidence, Tctex-1, a dynein light chain, was shown to interact with rhodopsin’s C-terminus via a yeast two-hybrid assay (Sung et al., 1999). Mutations in the C-terminus of rhodopsin reduced binding of Tctex-1 (Sung et al., 1999). Additionally, defects in the Dynein gene lead to disorganized and shortened OS in vertebrate models (Krock et al.,
2009; Insinna et al., 2010). Additionally, KIFC1, a kinesin motor, has been shown to interact with ASAP1 and may be involved in the exit of ciliary receptors (Lee et al., 2018).
While the process of entering the cilia is not completely understood yet, studies on zebrafish models suggest Rab8 indirectly associates with intraflagellar transport protein 20 (IFT20) through a protein called elipsa (Omori et al., 2008).
IFT20, in addition to its role in rhodopsin transport to the cilia, aids in rhodopsin release from the Golgi body; defects in the IFT20 gene cause rhodopsin accumulation in the Golgi apparatus in mouse model (Keady et al., 2011). It is
4
Figure 2. Vectorial trafficking of rhodopsin from the Golgi to the distal end of the connecting cilium. Dynein and microtubules may mediate the vectorial trafficking of rhodopsin transport carriers from the Golgi to the basal body. The BBSome, IFT particles, and kinesin may mediate the trafficking of rhodopsin within the transition zone to the region of new disk synthesis. Two possible mechanisms for rhodopsin delivery are proposed: the first is the fusion of rhodopsin transport carriers with the inner segment periciliary plasma membrane followed by trafficking of rhodopsin-containing rafts to the nascent disks (left panel); the second is the trafficking of rhodopsin transport carriers directly to the nascent disks (right panel).
5
important to note that this model of rhodopsin transport is still under
debate. For example, although those studies suggest the essential roles of
various monomeric GTPases, genetic studies of mice suggested that Arf4, Rab8,
and Rab11 are dispensable for OS-localization of rhodopsin (Ying et al., 2016;
Pearring et al., 2017).
Once reaching the base of the connecting cilium, rhodopsin must pass
through the transition zone, also called connecting cilium, in order to reach the
OS. Real-time visualization of rhodopsin within this region of the connecting
cilium has not been achieved, though there is evidence that rhodopsin is
transported via the ciliary plasma membrane (Chadha et al., 2019). The IFT
machinery forms complexes called the IFT-A and IFT-B particles (Rosenbaum
and Witman, 2002) which localize to the ciliary region of photoreceptor cells
(Luby-Phelps et al., 2008). Defects in a particle A component induce rhodopsin mislocalization to the photoreceptor plasma membrane (Crouse et al., 2014), while defects in particle B components are known to cause aberrant trafficking of rhodopsin and OS defects (Pazour et al., 2002; Tsujikawa and Malicki, 2004).
Therefore, the IFT machinery is thought to play an essential role of trafficking rhodopsin within the connecting cilia. In this region, microtubules are present with their plus-ends directed towards the OS. A motor protein associated with plus-
end directed trafficking on microtubles, kinesin KIF3A, is known to interact with
the IFT machinery (Scholey, 2013) and therefore may be involved in rhodopsin
trafficking in this region. In the KIF3A knockout mouse model, various degrees of rhodopsin mislocalization accompany photoreceptor degeneration (Lopes et al.,
6
2010), although OS targeting of rhodopsin was not totally inhibited (Marszalek et al., 2000)}(Jimeno et al., 2006)}(Avasthi et al., 2009). Morpholino knockdown of another microtubule motor protein, KIF17, resulted in impaired rhodopsin targeting to OS resulting in OS defects in zebrafish model (Insinna et al., 2008).
Controversially, a KIF17 mutant zebrafish line demonstrated normal OS and rhodopsin localization, complicating the interpretation regarding the roles of
KIF17 in rhodopsin trafficking (Zhao et al., 2012). In addition to microtuble motor proteins, an actin-based motor called myosin VIIa is also evidenced to participate in rhodopsin trafficking within the cilia. Defects in myosin VIIa resulted in accumulation of rhodopsin in the connecting cilium, but did not result in overt OS defects or photoreceptor cell death, indicating that its function is supplementary and not indispensable for OS-localization of rhodopsin. Miscoordination of these trafficking components can lead to rhodopsin mislocalization which is the subject of my thesis project.
1.3 Rhodopsin mutations are the major causes of inherited blindness
Rhodopsin mutations are the major causes of inherited blinding disorders collective called as rhodopsin retinitis pigmentosa (Mendes et al., 2005; Wright et al., 2010); they comprise approximately 25% of autosomal dominant forms of retinitis pigmentosa (RP); Over 100 distinct point mutations of the rhodopsin gene have been identified to be RP-causative (Malanson and Lem, 2009).
Among them, Mutations which disrupt the cilia-targeting (VXPX) motif result are categorized as class I mutations and result in rhodopsin mislocalization and
7 some of the most severe forms of progressive blindness. In retinitis pigmentosa caused by rhodopsin mislocalization, initial symptoms include shortening or loss of OSs and compromised vision. Those symptoms are followed by death of rod and cone photoreceptors (Fliegauf et al., 2007). While rhodopsin mislocalization is prevalent in class I rhodopsin mutants (Milam et al., 1996), rhodopsin mislocalization is more broadly observed in the RP patients associated with non- rhodopsin mutations (Li et al., 1995; Milam et al., 1998). Increasing evidence suggests rhodopsin mislocalization is the cause of photoreceptor dysfunction and degeneration, however, it is mechanistically unclear how rhodopsin mislocalization causes photoreceptor degeneration. Investigating the pathology of rhodopsin mislocalization is the major subject of this thesis project.
1.4 Electrochemical interactions between IS and OS are essential for vision
Photoreceptor OS is separated from IS by connecting cilium (Steinberg et al., 1980; Obata and Usukura, 1992; Pugh, 2015). Signals generated by photoreceptor OSs are conveyed to the ISs through electrochemical interaction.
Rhodopsin molecules existing in the OS disks are able to absorb photons, which induce a conformational change of the molecule resulting in the activatation of its cognate G-protein, transducin (Lee et al., 2010). When transducin becomes activated, a series of biochemical reactions lead to reduced [cGMP] and closure of the cGMP-gated channels in the OS plasma membrane (Nemet et al., 2014), thus barring entry of cations such as Na+ and Ca2+ into the photoreceptor, and causing the cell to become hyperpolarized. For this process, Na+/K+-ATPase,
8
which resides on the IS PM plays critical roles because and the ATPase pumps
Na+ out of the cell against its gradient in place of K+ (Schneider and Kraig, 1990).
It is currently unclear if this OS-IS interaction is disrupted in retinitis pigmentosa.
Investigating this interaction under pathological conditions is a primary goal of this thesis project.
1.5 Currently available approaches to study rhodopsin transport
More than five decades ago, Dr. Richard Young and his colleagues studied the process of renewing biological structures utilizing in vivo radiolabeling and autoradiography. These early studies allowed studying, for example, the renewal of OS proteins over an extended period of time (Young, 1967). In this technique, radiolabeled amino acids are randomly assimilated into newly synthesized proteins. This approach provided essential information about the renewal of OS proteins, but specific OS components such as rhodopsin could not be discriminated from the pool of various radiolabeled proteins. Since the beginning of this century, investigators started to harness genetic labeling methods to specifically visualize rhodopsin in photoreceptor cells. For example, mouse and frog models expressing rhodopsin fused to green fluorescent protein
(GFP) were generated (Moritz et al., 2001; Chan et al., 2004; Wensel et al.,
2005). Those models allowed to investigate the static localization and mislocalization of rhodopsin and its mutants. This technique, however, lacked the
advantage of the autoradiography technique which provided distribution pattern
of recently-synthesized proteins.
9
More recent studies thus improved the genetic labeling methods to
visualize outer segment dynamics. In order to study OS growth rates, one
innovative model utilized a heat-shock promotor to induce the expression of a red fluorescence protein fused to a transmembrane region to facilitate its incorporation into the OS disk membranes (Willoughby and Jensen, 2012). Such
a system is useful as it allows tracking of the movement of a subset of disk
membranes within the OS, similar to autoradiography setting. Therefore, the
turnover rate of the OS can be calculated. This system does not allow, however,
studying the protein synthesis or transport under native conditions, as heat shock
would alter the expression of chaperone proteins or may cause additional stress
responses. To ameliorate these issues, another model was developed in which a
rhodopsin reporter molecule was induced by heat-independent mechanism. This
technique instead utilized an inducible promoter which is activated only after
treatment with 4-hydroxytamoxifen (4OH) (Hsu et al., 2015); such Cre-Lox
systems are useful for inducing expression of a specific protein which can be
tracked from that point onward (Wang et al., 2005). Such utility is advantageous
for determining protein expression under different conditions such as dark/light.
This tool was used to show that light regulates the transport of rhodopsin
molecules by sequestering them in the Golgi apparatus under light conditions,
and are released under dark conditions. The technique, however, lacks the
necessary spatial resolution to study rhodopsin transport, because of the lapse in
gene editing after treatment with 4OH. The exact time of protein synthesis is
difficult to determine because of the time required for 4OH to distribute to the
10
Figure 3. Application of Dendra2 for imaging of rhodopsin trafficking in live Xenopus laevis rod photoreceptor cells. (A) Representation of photoconversion (PC) of Dendra2. Dendra2, which is fused to human rhodopsin (Rho-Dend2-1D4), is green prior to PC, and is irreversibly changed to red after PC. New Rho-Dend2-1D4 (green) is synthetized in the inner segment and trafficked to the outer segment where it gets incorporated into disks. (B, C) Live Xenopus laevis rod photoreceptor cells expressing Rho-Dend2-1D4. Newly synthetized Rho-Dend2-1D4 can be observed in the inner segment shortly after PC (B, left panel, arrow); Rod outer segments are renewed over 2 - 6 days (B, middle and right panel). (C) Vesicles containing newly synthetized Rho-Dend2-1D4can be seen moving from the inner segment towards the outer segment. Images are maximum projections of optical slices. Scale bars = 10 µm.
11 ocular tissue as well as the stochastic nature of cre-recombination which may or may not occur.
In the past decade, our laboratory has developed a novel technique which allows studies of rhodopsin transport at high-spatiotemporal resolution, surpassing currently available methods. This technique took advantage of the
Dendra2 fluorescent protein, which is a green-to-red photoconvertible fluorescent protein. This property allows all existing rhodopsin fusion proteins to be irreversibly converted to exhibit red fluorescence while newly synthesized proteins will fluoresce in green (Figure 3). Therefore, like the inducible system, temporal information such as age of the proteins can be attributed to understand the destiny of those specific population of proteins. We can follow the destiny of those fusion proteins either for days (up to 6 d, Figure 3B). Because Dendra2 proteins are photoconverted instantaneously, the technique allows tracking of rhodopsin transport which occurs in the time regime of seconds (Figure 3C). Our group previously used this approach to demonstrate that class I mutant rhodopsin is continuously removed, and then renewed to the IS PM (Figure 4)
(Lodowski et al., 2013). Knowledge that mislocalized rhodopsin was being removed from the IS PM allowed us to investigate the mechanism behind this removal and consequences of this process.
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Figure 4. Monitoring removal of mislocalized mutant rhodopsin (Q344ter) in live Xenopus laevis rod photoreceptor cells.
(A) RhoQ344ter-Dendra2 mislocalizes to the inner segment plasma membrane including calyceal processes (arrows). Over time mislocalized rhodopsin is renewed at the plasma membrane (arrows). At 2 days post-photoconversion (2 d post-PC), the plasma membrane is yellow because contains both old (red) and new (green) RhoQ344ter-Dendra2. At 6 days post-photoconversion (6 d post-PC), old RhoQ344ter-Dendra2 was removed and new RhoQ344ter-Dendra2 was added, resulting in green fluorescent signal on the plasma membrane. Images are maximum projections of confocal sections. Scale bar = 10 µm. (B) Schematic representations of hypothetical mechanisms involved in removal of mislocalized rhodopsin, and how a green-to-red PC fluorescent protein can be used unravel those mechanisms. MVB stands for multivesicular body.
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1.6 Aims of dissertation
The current literature suggests that there is a light-independent
mechanism of photoreceptor degeneration that is caused by rhodopsin
mislocalization. The purpose of this dissertation is to investigate the mechanism
of photoreceptor degeneration induced by rhodopsin mislocalization which
occurs in the absence of light, as well as to determine the methods rod cells
employ to remove mislocalized rhodopsin. In Chapter 2, the lysosomal pathway is explored as an avenue for mislocalized rhodopsin degradation. In Chapter 3, the plasma membrane protein homeostasis of rods expressing class I mutant rhodopsin is examined. In Chapter 4, the relationship between photoreceptor cell health, including IS-OS interactions, and rhodopsin mislocalization is explored. In
Chapter 5, alternate mechanisms rod photoreceptors employ to remove
mislocalized rhodopsin, specifically vesicular secretion, will be explored. Lastly,
Chapter 6 will offer conclusions and suggest new directions for future studies
into the pathology caused by rhodopsin mislocalization.
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Chapter 2: Mislocalized rhodopsin is degraded via the lysosomal pathway
The material in this Chapter was adapted from Ropelewski P, Imanishi Y, 2019:
Disrupted plasma membrane protein homeostasis in a Xenopus laevis model of retinitis pigmentosa. The Journal of Neuroscience 39(28):3025-18.
2.1 Introduction
Mislocalized rhodopsin causes photoreceptor degeneration in both light- dependent and -independent manners (Alfinito and Townes-Anderson, 2002;
Tam et al., 2006; Concepcion and Chen, 2010). Light activation of rhodopsin leads to ectopic activation of G protein-mediated cascade and subsequent photoreceptor degeneration (Alfinito and Townes Anderson, 2002). While light exposure may hasten the onset of photoreceptor degeneration, the underlying mechanism of light-independent photoreceptor cell death remains unsolved.
Light independent effects include morphological changes in the plasma membrane (PM) which leads to neurite sprouting, suggesting the direct effect of rhodopsin mislocalization is on the PM (Milam et al., 1996; Tam et al., 2006).
Therefore, I sought to understand how rhodopsin mislocalization induces PM toxicity in a light-independent manner.
15
Rod photoreceptors have an intrinsic mechanism to remove IS-
mislocalized rhodopsin. For example, in photo-damaged rat retinas, rod photoreceptors initially exhibit IS-mislocalized rhodopsin which disappears following the recovery (Edward et al., 1993). More recently, our laboratory found that class I mutant rhodopsin mistrafficked to the IS PM is continuously eliminated (Lodowski et al., 2013). Rhodopsin is normally eliminated through phagocytosis of rod OSs by the retinal pigment epithelium (RPE) (Young, 1967).
However, rhodopsin mislocalized on the IS PM has no direct access to the RPE, and therefore the mechanism(s) of mislocalized rhodopsin elimination were unclear. Therefore, I explored the intracellular mechanisms rod photoreceptors employ to remove mislocalized rhodopsin. Intracellular class I mutant rhodopsin partially co-localized with markers of early endosome (EEA1) and lysosome
(LAMP1) (Lodowski et al., 2013). These observations led us to question if mislocalized rhodopsin is continuously transported to IS lysosomes for
degradation.
2.2 Materials and Methods
2.2.1 Animals
All animal procedures were approved by the Institutional Animal Care and
Use Committee (IACUC) at Case Western Reserve University. Adult female and
male frogs were purchased from Nasco and housed at 16° C under a 12-h
light/12-h dark cycle. All tadpoles used for experiments were housed at 16° C in
16
24-h darkness. Tadpoles were fed spirulina (Nuts.com) until metamorphosis was
complete, and then were fed pelleted frog brittle (Nasco). Both male and female
tadpoles were used for all experiments.
2.2.2 Reagents
Unless otherwise specified, all reagents were purchased from either
Fischer Scientific or Sigma-Aldrich.
2.2.3 Molecular cloning
DNA expression vectors were generated by standard methods combining
PCR, DNA recombination, and site-directed mutagenesis. The plasmid vectors
containing XOP-Rho-Dend2-1D4 and RhoQ344ter-Dend2 were previously
generated (Lodowski et al., 2013). XOP-Rho-Dend2-1D4 is designed to express human rhodopsin followed by Dendra2 fluorescent protein and the last 8 amino acids of rhodopsin which serve as the epitope for the monoclonal antibody 1D4
(Xie et al., 2011). The addition of the 1D4 epitope does not inhibit the function or localization of rhodopsin, and results in proper targeting of the fusion protein to
the OSs (Xie et al., 2011; Lodowski et al., 2013). RhoQ344ter-Dend2 is designed to express class I mutant (Q344ter) of human rhodopsin missing the last five amino acids (QVAPA) followed by Dendra2 fluorescent protein. All the vectors contained polyadenylation signals following the coding and non-coding regions.
2.2.4 Transgenesis of Xenopus laevis
Transgenic Xenopus laevis were produced using the intracytoplasmic sperm injection (ICSI) method following the previously published procedure
17
(Sparrow et al., 2000; Smith et al., 2006; Lodowski et al., 2013). Expression
vectors for Rho-Dend2-1D4 and RhoQ344ter-Dend2 were purified from bacteria cultures using EndoFree Plasmid Midi Prep Kit (QIAGEN), and then used to isolate DNA fragments containing the XOP promoter, coding/noncoding regions, and polyadenylation signal for transgenesis. Fully developed tadpoles were
screened for the presence of Dendra2 fluorescence in their eyes at days post-
fertilization (7 DPF). Tadpoles were immersed in 6% methylcellulose to prevent
movement while probed for green fluorescence in their eyes using a Leica
MZ16F stereoscope (Leica Microsystems). Tadpoles were classified into one of
the three following categories based on the expression level of the transgene:
low, medium, or high as assessed by intensity of green fluorescence in their
eyes. In order to prevent photobleaching, light-dependent photoreceptor
degeneration, and unintended photoconversion of fluorescent protein Dendra2,
tadpoles were reared in 24-h darkness.
2.2.5 Photoconversion of Dendra2 in tadpole retina
Dendra2 fluorescent protein in Xenopus tadpole eye was photoconverted
as described previously (Lodowski et al., 2013; Lodowski and Imanishi, 2015).
Xenopus tadpoles were anesthetized in 0.026% tricaine methanesulfonate and
then immobilized in 6% methylcellulose covered with a protective light filter to
prevent tissue damage. Light from a 405 nm laser placed 10 cm above the
tadpole was directed at both eyes for a total of 15 - 20 minutes with 30 second
breaks between each minute. The successful and nearly complete
photoconversion was confirmed via inspection of green and red fluorescence by
18
Leica MZ16F stereoscope. After photoconversion, tadpoles were subjected to small molecule treatment as described below, or sacrificed and imaged immediately.
2.2.6 Treatment of tadpoles by lysosome inhibitor
Animals were treated in 0.1x Mark’s Modified Ringer (MMR) solution (10x
MMR composition: 1 M NaCl, 20 mM KCl, 20 mM CaCl2, 10 mM MgCl2, 50 mM
HEPES, pH 7.5) (Lodowski et al., 2013) containing 0.1% DMSO and bafilomycin
A1 (lysosome inhibitor, 100 nM). Bafilomycin A1 was initially diluted in DMSO,
and then applied to 5 ml of 0.1x MMR buffer containing a single tadpole.
Bafilomycin A1 treatments lasted for 24 h. As controls, animals were placed in
0.1x MMR containing 0.1% DMSO.
2.2.7 Preparation of retina explant for confocal imaging
After euthanasia and decapitation, whole eyes were extracted from
tadpole heads using surgical tools. Retinas were immediately placed into a glass
bottom dish (P35G-1.5-14-C, MatTek Corporation) containing equilibrated
modified Wolf medium (D-glucose, 700 mg/L, 30 mM NaHCO3, 55% MEM, 31%
sodium-free BBS, 10% FBS) (Lodowski et al., 2013). Equilibration of Wolf
medium was achieved by incubating the dish containing the medium in a gas
chamber supplied with 5% CO2 and 95% OS (Airgas) (Nemet et al., 2014). The
glass bottom dishes were coated with Cell-Tak Cell and Tissue Adhesive
(Corning). RPE were carefully removed from the neural retinas which were then
placed onto the dish, and then a circular glass coverslip (12 mm diameter and
19
0.13 to 0.17 mm thickness) was used to seal and flatten the retinas before
imaging. Retinas were kept alive in Wolf medium with constant flow of 5% CO2
and 95% O2 in a humid imaging chamber (Tokai Hit). For labeling of lysosomes,
retinas were incubated in equilibrated modified Wolf medium containing 1:100 dilution of LysoTracker Red DND-99 (Thermo Fisher Scientific) for 30 min prior mounting on MatTek dish and imaging.
2.2.8 Confocal microscopy, Image analysis, quantification, and statistical analysis
All images were acquired using a Leica TCS SP2 laser scanning confocal/multiphoton microscope system equipped with four lasers for excitation:
488 nm argon ion, 543 nm HeNe, 633 nm HeNe, and tunable Chameleon XR
Ti:Sapphire laser (Leica Microsystems) as described previously (Lodowski et al.,
2013). A HCX PL APO CS 40.0 X 1.25 oil UV objective lens was used for imaging. Diameters of vesicular structures were measured using ImageJ’s line function tool (Schneider et al., 2012). For the quantitative studies involving drug treatments, the same laser power and imaging condition were used for all the samples. The cytoplasmic space was highlighted manually and its mean fluorescence intensity measured using ImageJ’s freehand selection tool
(Schneider et al., 2012). The background noise was determined from blank space where no cells were observed, and subtracted from the measurements. All the quantitative data were obtained from 4 independent animals (or structures thereof) are represented as mean ± SD. In comparing two populations, *p <
0.001 by Student’s t-test was considered statistically significant.
20
2.3 Results and Discussion
2.3.1 Class I mutant rhodopsin is degraded in lysosomes
We previously demonstrated that class I mutant rhodopsin is actively
eliminated from rod photoreceptor IS PM (Lodowski et al., 2013). To characterize the removal process of class I mutant rhodopsin and compare it to that of wild
type rhodopsin in living cells, Q344ter— a class I mutant human rhodopsin with
the five terminal amino acids truncated— and wild type human rhodopsin were
fused to Dendra2 fluorescent protein and individually expressed in Xenopus
laevis rod photoreceptors. As reported previously for endogenous rhodopsin, wild
type rhodopsin fused to Dendra2 specifically localized to rod OSs (Figure 5A,
Rho-Dend2-1D4, Control) (Lodowski et al., 2013). As reported for the localization
of untagged class I mutant rhodopsin in transgenic animal models (Sung et al.,
1993; Sung et al., 1994; Tam et al., 2006; Concepcion and Chen, 2010), the
majority of class I mutant rhodopsin fused to Dendra2 mislocalized to the IS PM
(Figure 5A, RhoQ344ter-Dend2, Control) (Lodowski et al., 2013). Wild type
rhodopsin correctly localizes to the rod OS and is removed via phagocytosis by
the neighboring RPE cells (Young, 1967; Besharse et al., 1977). However,
because IS PM mislocalized rhodopsin has no direct access to the RPE, we
asked whether RhoQ344ter-Dend2 is degraded within lysosomes of rod
photoreceptor cells. Lysosome-mediated degradation was inhibited by
bafilomycin A1 (BA1, 100 nM) in Xenopus laevis rods either expressing Rho-
Dend2-1D4 or RhoQ344ter-Dend2 (Figure 5A).
21
Figure 5. Class I mutant rhodopsin accumulates in lysosomes after inhibition of lysosome-mediated degradation. (A) Confocal imaging of live Xenopus laevis rod photoreceptors expressing either Rho- Dend2-1D4 (left panels) or RhoQ344ter-Dend2 (right panels). Outer segment (OS) and inner segment (IS) structures are labeled in a representative rod cell of each image. Prior to the imaging, the animals were treated with DMSO (Control) or 100 nM bafilomycin A1 (BA1) for 24 h. In control retinas, neither Rho-Dend2-1D4 nor RhoQ344ter- Dend2 accumulated significantly in vesicular structures of the ISs. After BA1 treatment, RhoQ344ter-Dend2 accumulated in vesicular structures (arrowheads) in the ISs, but Rho- Dend2-1D4 did not accumulate in vesicular structures. (B) In retinas expressing either Rho-Dend2-1D4 or RhoQ344ter-Dend2, we assessed the percentage of rod photoreceptor cells which exhibit vesicles > 1 μm in diameter. BA1 treatment of photoreceptors expressing Rho-Dend2-1D4 increased this percentage from 0.9 ± 1.0% (untreated) to 5.1 ± 6.2% (NS, p = 0.23). BA1 treatment of photoreceptors expressing RhoQ344ter-Dend2 increased this percentage from 8.2 ± 2.7% to 59.4 ± 10.0% (p < 0.001), indicating that RhoQ344ter-Dend2, but not Rho-Dend2-1D4, accumulates upon BA1 treatment. (C) Confocal imaging of live Xenopus laevis rod photoreceptors expressing RhoQ344ter- Dend2 and labeled with LysoTracker Red dye. LysoTracker Red signal co-localized with RhoQ344ter-Dend2 in untreated cells (Control, arrowheads), and the amount of RhoQ344ter- Dend2 co-localized with LysoTracker Red increased after pre-treatment with BA1 (BA1, arrowheads). Scale bars = 10 μm.
22
After 24 h of treatment, RhoQ344ter-Dend2 frequently accumulated in
vesicular structures within the rod ISs (Figure 5A, RhoQ344ter-Dend2, BA1,
arrowheads) with a size distribution (diameter) of 2.07 ± 0.63 μm (n = 126
vesicles from 7 animals). In the absence of BA1 treatment, RhoQ344ter-Dend2
rarely accumulated in vesicular structures with diameters larger than 1 μm
(Figure 5A, RhoQ344ter-Dend2, Control). In animals expressing RhoQ344ter-Dend2 and treated with BA1, 59.4 ± 10.0% of rod photoreceptors exhibited vesicular structures with diameter larger than 1 μm (Figure 5B, based on 64 cells each
from n = 4 animals), whereas untreated animals exhibited these structures in only
8.2 ± 2.7% of rod photoreceptors (p < 0.001) (Figure 5B, based on 61 cells each
from n = 4 animals). These observations suggest that RhoQ344ter-Dend2 is prone to lysosome-mediated degradation, and that inhibition of this degradation via BA1 treatment resulted in the intravesicular accumulation of RhoQ344ter-Dend2. As a control, animals expressing Rho-Dend2-1D4 did not significantly accumulate in intracellular structures (> 1 μm) after BA1 treatment (Figure 5A, Rho-Dend2-1D4,
BA1). Untreated rods expressing Rho-Dend2-1D4 exhibited intracellular structures (> 1 μm) at a low frequency of 0.9 ± 1.0% (Figure 5B, based on 57
cells each from n = 4 animals). After BA1 treatment, the number of rods exhibiting these structures increased to 5.1 ± 6.2% (Figure 5B, based on 64 cells each from n = 4 animals), but not in a statistically significant manner (NS, p =
0.23). These observations indicate that Rho-Dend2-1D4 is marginally degraded by lysosomes within rod photoreceptors, and confirm previous findings that the
23
majority of wild type rhodopsin is degraded by RPE cells (Young, 1967; Besharse
et al., 1977).
To identify these intracellular organelles containing RhoQ344ter-Dend2,
LysoTracker Red (Pierzynska-Mach et al., 2014) was employed to specifically label lysosomes in photoreceptor cells. In rod ISs of untreated animals incubated with LysoTracker Red dye, LysoTracker-positive lysosomes frequently contained
RhoQ344ter-Dend2 (Figure 5C, Control, arrowheads). The majority of LysoTracker- positive lysosomes contained RhoQ344ter-Dend2 (68.0 ± 16.2% based on 32
lysosome-positive cells each from n = 4 animals). BA1 prevents acidification of
lysosomes/autophagolysosomes through inhibition of the V-ATPase of the
lysosome, which is responsible for pumping H+ ions into the compartment
(Yamamoto et al., 1998). Because LysoTracker Red only labels low-pH
organelles, it did not reproducibly label lysosomes of BA1-treated cells. Because
BA1 is a reversible inhibitor of the V-ATPase (Yamamoto et al., 1998), we
improved labeling of lysosomes by lifting BA1 treatment for 1 h prior to labeling
with LysoTracker Red (Figure 5C, BA1, arrowheads). Under these conditions,
accumulated RhoQ344ter-Dend2 co-localized with LysoTracker Red in 84.3 ±
5.3% of all lysosome-labeled cells observed (based on 31 lysosome-positive cells each from n = 4 animals). Therefore, the majority of these intracellular organelles containing RhoQ344ter-Dend2 are lysosomes or autophagolysosomes.
24
Figure 6. Mislocalized rhodopsin is trafficked from the plasma membrane to intracellular lysosomes. (A) Experimental design to determine the origin of proteins accumulating in the ISs of RhoQ344ter-Dend2 cells. Prior to photoconversion, the majority of RhoQ344ter-Dend2 is localized to the IS PM. After photoconversion and BA1 treatment (BA1, 24 h), red protein should accumulate if RhoQ344ter-Dend2 is being trafficked from the PM to the IS lysosomes (model i.). If RhoQ344ter-Dend2 is not internalized, then red RhoQ344ter-Dend2 will not accumulate significantly in the IS lysosomes (model ii.) (B) Immediately following photoconversion, rods expressing RhoQ344ter-Dend2 were imaged (0 h). Photoconversion was efficient as confirmed by the loss of green fluorescence (0 h, Green). The majority of red RhoQ344ter-Dend2 was observed on the IS PM (0 h, Red, PM, arrowhead). After 24 h of BA1 treatment, RhoQ344ter-Dend2 accumulated intracellularly in the ISs of rods (BA1, arrowheads). A nominal amount of green RhoQ344ter-Dend2 was observed after 24 h of retina culture (BA1 and Control, Green). This observation indicates that new RhoQ344ter-Dend2 was synthesized in these retinas. (C) Average red fluorescence intensities (a.u./area) were measured for intracellular regions of ISs in individual rods. The histograms indicate frequency of cells (y-axis) for each fluorescence intensity range (x-axis). Gray arrowheads indicate the average fluorescence intensity of each condition (0 h, BA1, or Control). Red fluorescence intensity values in the ISs were 36.7 ± 13.9 a.u. for time = 0 h condition, 29.4 ± 16.8 a.u. for 24 h DMSO-treated Control group, and 60.2 ± 16.8 a.u. for the BA1-treated group. Animals were aged 9 DPF upon photoconversion (0 h) and were sacrificed and imaged at 10 DPF (BA1 and Control), with a total of 250 cells for each condition measured from n = 5 animals. Scale bar = 10 μm.
25
2.3.2 Mislocalized rhodopsin is trafficked from the plasma membrane to
intracellular lysosomes
Two alternative explanations are plausible regarding the source of RhoQ344ter-
Dend2 in lysosomes: one possibility is that RhoQ344ter-Dend2 is continuously removed from the PM and trafficked to lysosomes, and another possibility is that
RhoQ344ter-Dend2 is sent to lysosomes directly after its synthesis in the Golgi
apparatus. In order to distinguish between these two possibilities, we took
advantage of the photoconversion property of Dendra2. Dendra2 is a
photoconvertible fluorescent protein engineered to irreversibly shift its emission
peak from shorter (507 nm, green) to longer (573 nm, red) wavelength (Nemet et al., 2015a). This photoconversion technique allowed us to study the destiny of photoconverted red RhoQ344ter-Dend2 by discriminating it from newly synthesized
green RhoQ344ter-Dend2 coming from the Golgi apparatus. If RhoQ344ter-Dend2 is
actively transferred from the PM to lysosomes, then we would expect a statistically significant increase of red proteins in the IS after 24 h of BA1 treatment (Figure 6A, model i). If, however, PM-localized RhoQ344ter-Dend2 is not
being transferred to the IS lysosomes (Figure 6A, model ii), then we would not
expect a significant increase of intracellular red proteins after BA1 treatment.
Photoconversion of rod photoreceptors expressing RhoQ344ter-Dend2 allowed us to label existing fusion proteins in red (Figure 6B, 0 h, Red). Most red RhoQ344ter-
Dend2 in these rods was observed on the IS PM (Figure 6B, 0 h, Red, PM,
arrowhead). Photoconversion of RhoQ344ter-Dend2 was highly efficient; significant
26
green fluorescence was not observed after photoconversion (Figure 6B, 0 h,
Green). After 24 h of BA1 treatment following the photoconversion, both green
and red RhoQ344ter-Dend2 accumulated in intracellular vesicular structures (Figure
6B, BA1, arrowheads). The majority of Dendra2 present in these structures exhibited red fluorescence (Figure 6B, BA1, Red arrowheads), thus supporting
our model in which PM-localized RhoQ344ter-Dend2 was internalized (Figure 6A,
model i.). As controls, a set of animals expressing RhoQ344ter-Dend2 were
photoconverted and then placed in 0.1x MMR buffer containing 0.1% DMSO (the
vehicle we used for BA1) for 24 h. As expected, RhoQ344ter-Dend2 did not
significantly accumulate in the ISs of untreated rods (Figure 6B, Control).
Because RhoQ344ter-Dend2 did not accumulate under these conditions, the accumulation of RhoQ344ter-Dend2 observed in the BA1-treated group must have
been due to lysosome inhibition and not due to other experimental conditions.
After 24 h, newly synthesized green RhoQ344ter-Dend2 was observed on the PM of both treated and untreated rods (Figure 6B, BA1 and Control, Green). Green protein was also observed in the intracellular structures of BA1-treated rods
(Figure 6B, BA1, Green, arrowheads). Therefore, under our experimental conditions, rod photoreceptors were healthy and capable of synthesizing and trafficking new RhoQ344ter-Dend2.
We confirmed through quantitative analysis that red RhoQ344ter-Dend2 accumulated after inhibition of lysosomes. Red fluorescence within the IS cytoplasm of individual rods from each experimental group was measured, and rods were categorized based on intracellular fluorescence intensities within their
27
ISs (Figure 6C). Overall, there were variable amounts of red fluorescence in the
ISs for all the conditions tested; this was likely because our transgenic system results in variable expression of the transgene in individual rods (Lodowski et al.,
2013). Despite the variance, BA1-treated rod cells (Figure 6C, BA1) more frequently exhibited high amounts of red fluorescence in their ISs (Figure 6C, compare BA1 to 0 h and Control). On average, BA1-treated rods exhibited significantly (p < 0.001) more red fluorescence in their ISs than in control rods;
BA1-treated rods had 2.03 ± 0.89 times more red fluorescence than control rods and 1.62 ± 0.71 times more red fluorescence than 0 h rods (based on n = 250 cells from 5 animals for each condition). The increased amount of red Dendra2 observed in BA1-treated rods is largely contributed by vesicular accumulated proteins (Figure 6B, BA1, Red, arrowheads), whereas red protein observed in untreated rods were contributed by proteins existing diffusively in the IS (e.g. endoplasmic reticulum). Red fluorescence in the IS decreased after 24 h in untreated rods, suggesting that red protein existing at 0 h had exited structures such as ER after 24 h. In summary, these results statistically support model i
(Figure 6A) in which PM-localized RhoQ344ter-Dend2 is actively removed from the
PM and translocated to IS lysosomes for degradation.
2.4. Conclusions
By taking advantage of class I mutant rhodopsin which is deficient in ciliary targeting, we proved a novel mechanism of photoreceptor toxicity caused by mislocalized rhodopsin in retinal ciliopathies. In this Chapter, I provided
28
evidence that mislocalized rhodopsin is continuously internalized, transported to
lysosomes, and degraded. This lysosome-mediated degradation likely results in disruption of the protein homeostasis of the rod IS PM where class I mutant rhodopsin is locally added and removed at rapid rates. Class I mutant rhodopsin causes photoreceptor degeneration in a manner independent of rhodopsin activation by light (Tam et al., 2006; Concepcion and Chen, 2010). In this study, all the animals were reared in 24-h darkness until subjected to live microscopy experiments, and therefore served as a model for light-independent disruption of photoreceptor structure and function.
These studies demonstrate the strengths of pharmacological approaches which allowed lysosome inhibition as photoreceptor degeneration and dysfunction manifest in Xenopus model of RP. By combining live confocal
microscopy technique and photoconversion property of Dendra2 fluorescent protein, we were able to mechanistically interrogate how class I mutant rhodopsin
molecules are degraded from the plasma membrane. This novel approach is
potentially useful for analyzing the pathological basis of RP caused by other
rhodopsin mutation, such as Class II rhodopsin mutation which is actively
degraded, but the degradation process is not fully elucidated.
29
Chapter 3: NKA is co-degraded with class I mutant rhodopsin and becomes
downregulated on the IS PM of rod photoreceptors
The material in this Chapter was adapted from Ropelewski P, Imanishi Y, 2019.
Disrupted plasma membrane protein homeostasis in a Xenopus laevis model of
retinitis pigmentosa. The Journal of Neuroscience 39(28):3025-18.
3.1 Introduction
As mentioned in Chapter 1, a massive amount of rhodopsin molecules
(106 - 108 molecules) are synthesized in individual rod cells every day. In
autosomal retinitis pigmentosa due to class I rhodopsin mutation, at least half the
amount (0.5 x 106 - 108 ) of rhodopsin molecules are delivered to the IS PM,
which demonstrates smaller membrane area than collective disk membranes in
the OS. Within disk membranes, rhodopsin molecules form crystal-like structures
(Fotiadis et al., 2003) because of their high concentrations. Thus the high concentration of rhodopsin in the IS PM will potentially affect the homeostasis of
endogenous IS PM proteins. Moreover, as described in Chapter 2, constant
misrouting and removal of the mislocalized rhodopsin to/from the IS PM would
have negative impact on IS PM proteins. Studies focusing on the effect of
30 rhodopsin mistrafficking to the IS PM membrane homeostasis has been scarce in the past, and is the focus of this Chapter 3.
Photoreceptor inner segments (ISs), where rhodopsin mislocalizes, play critical roles in maintaining the dark current. In the OSs, photon absorption by rhodopsin triggers a series of biochemical events which result in hydrolysis of cGMP, closure of cGMP-gated channels, and reduced inward flow of Na+ and
Ca2+. In the IS, Na+ /K+-ATPase pumps out Na+ in exchange for K+ that is then conducted through K+ channels. Phototransduction-mediated changes in the net flow of Na+ and K+ constitute the basis for signal transmission from OS to IS, eventually leading to reduced glutamate release from the synaptic terminus (Yau and Baylor, 1989). Such a massive amount of rhodopsin incorrectly delivered to the IS PM would likely affect the local concentration of other transmembrane proteins, and may also disturb the cell’s ion homeostasis (Demontis et al., 1995).
We studied the homeostasis of Na+ /K+-ATPase in rod photoreceptors expressing class I mutant rhodopsin.
3.2 Materials and Methods
3.2.1 Animals
All animal procedures were approved by the Institutional Animal Care and
Use Committee (IACUC) at Case Western Reserve University. Adult female and male frogs were purchased from Nasco and housed at 16° C under a 12-h
31
light/12-h dark cycle. All tadpoles used for experiments were housed at 16° C in
24-h darkness. Tadpoles were fed spirulina (Nuts.com) until metamorphosis was
complete, and then were fed pelleted frog brittle (Nasco). Both male and female
tadpoles were used for all experiments.
3.2.2 Reagents
Unless otherwise specified, all reagents were purchased from either
Fischer Scientific or Sigma-Aldrich.
3.2.3 Transgenesis of Xenopus laevis
Transgenic Xenopus laevis were produced using the intracytoplasmic
sperm injection (ICSI) method following the previously published procedure
(Sparrow et al., 2000; Smith et al., 2006; Lodowski et al., 2013). Expression
vectors for NKAα-PSmOrange, Rho-Dend2-1D4, and RhoQ344ter-Dend2 were
purified from bacteria cultures using EndoFree Plasmid Midi Prep Kit (QIAGEN), and then used to isolate DNA fragments containing the XOP promoter,
coding/noncoding regions, and polyadenylation signal for transgenesis. Fully
developed tadpoles were screened for the presence of Dendra2 fluorescence in
their eyes at days post-fertilization (7 DPF). Tadpoles were immersed in 6%
methylcellulose to prevent movement while probed for green fluorescence in their
eyes using a Leica MZ16F stereoscope (Leica Microsystems). Tadpoles were
classified into one of the three following categories based on the expression level
of the transgene: low, medium, or high as assessed by intensity of green
fluorescence in their eyes. In order to prevent photobleaching, light-dependent
32
photoreceptor degeneration, and unintended photoconversion of fluorescent protein Dendra2, tadpoles were reared in 24-h darkness. A male animal expressing high-level of RhoQ344ter-Dend2 (F0) was 10 raised to maturity (> 1 y
old) and sacrificed to collect sperm, which were then used to create 2nd
generation (F1) animals. This sperm was also used for generating tadpoles which
co-expressed PSmOrange-NKAα and RhoQ344ter-Dend2. For this purpose, the sperms were subjected to ICSI method described above with linearized DNA containing XOP-PSmOrange-NKAα.
3.2.4 Immunohistochemistry
Xenopus laevis eyes were fixed as previously described (Lodowski et al.,
2013). Tadpoles were first euthanized in a 0.26% tricaine and 0.52% sodium bicarbonate solution (in dH2O) and decapitated. Their heads were fixed in 4%
paraformaldehyde (Electron Microscopy Sciences) in 0.1 M phosphate buffer
(PB, NaHPO4 and Na2HPO4 dissolved in MilliQ H2O, pH 7.4) for 6 h at RT. Fixed
heads were incubated in 5% sucrose in phosphate buffer for 30 min, and then
subsequently washed in 10%, 15%, and 20% sucrose solutions in 0.1 M PB. The
heads were then incubated overnight at 4 ºC in a solution containing 20%
sucrose and optimum cutting temperature compound (O.C.T. compound, Sakura
Finetek) in a 2:1 ratio. The following day, heads were aligned in Tissue-Tek
cryomolds (Sakura Finetek) and frozen in isopentane cooled by liquid N2. Frozen
tissue sections (12 μm) were cut using a Leica CM1850 cryostat (Leica
Microsystems) and collected on Superfrost Plus microscopy slides. Sections
were dried and stored at -80 ºC until use. Slides were removed from -80 ºC
33
storage conditions, warmed using a 37 ºC incubator (CCC 0.5d, Boekel
Scientific), and then sections were encircled using an ImmEdge pen (Vector
Laboratories). Sections were rehydrated and blocked in a 1.5% normal goat
serum diluted in phosphate-buffered saline solution (PBS, mM NaCl, 2.7 mM
KCl, 8 mM Na2HPO4, 2 mM KH2PO4) for 30 min at RT. Blocking solution was
then removed and primary antibody (1:100) diluted in PBS containing 0.1% Triton
X-100 (PBST) was added; sections were incubated at RT for approximately 14 h
and then washed 3 times with PBST for 10 min each. After the washes, sections
were incubated in PBST containing anti-mouse Cy3 (1:100) and Hoechst dye
(1:1000) for 1 h at RT followed by an additional 3 washes with PBST for 10 min
each. After these washes, one drop of Vectashield mounting medium (Vector
Laboratories) was added to a square coverslip and placed over the section. Nail
polish (Electron Microscopy Sciences) was used to seal the coverslip, and the
sections were dried at RT for > 30 min before imaging. Antibodies used for
immunofluorescence imaging were as follows: mouse anti-Na+ /K+-ATPase
(Lebovitz et al., 1989) (Hybridoma Bank, AB_2166869), mouse anti-
peripherin/rds (anti-xrds 2D4, a kind gift from Dr. Robert Molday, Univ. of British
Columbia), mouse XAP-2 (clone 5B9, Hybridoma Bank), Hoechst (Thermo Fisher
33342), and anti-mouse-Cy3 conjugate (Jackson, 115-165-166).
3.2.5 Reverse transcription and quantitative PCR
Total RNA was isolated from Xenopus retinas using TRIzol (Invitrogen)
according to the manufacturer’s instructions. Approximately 5 µg of RNA was
subjected to reverse transcription using iScript Reverse Transcription Supermix
34
for RT-qPCR (Biorad). The resulting cDNA was used as a template for realtime
PCR analyses. Following oligonucleotide primers were designed for amplifying
NKA and -tubulin transcripts: NKA-F (5-GTC AGA CAG TTA CCG CGT AGC
CAC C-3); NKA-R (5-CTT TAT CCA CTC AGG GGT TGT GGG AGG-3); - tubulin-F (5-ACA CGG CAT TGA TCC TAC AG-3); -tubulin-R (5-AGC TCC TTC
GGT GTA ATG AC-3). NKA and -tubulin transcripts were amplified and quantified using iQ SYBR Green Supermix (Biorad) and a StepOnePlus
RealTime PCR System (Applied Biosystems). StepOne software (v2.3) was used for analysis
3.2.6 Treatment of tadpoles by lysosome inhibitor
Animals were treated in 0.1x Mark’s Modified Ringer (MMR) solution (10x
MMR composition: 1 M NaCl, 20 mM KCl, 20 mM CaCl2, 10 mM MgCl2, 50 mM
HEPES, pH 7.5) (Lodowski et al., 2013) containing 0.1% DMSO and bafilomycin
A1 (lysosome inhibitor, 100 nM). Bafilomycin A1 was initially diluted in DMSO,
and then applied to 5 ml of 0.1x MMR buffer containing a single tadpole.
Bafilomycin A1 treatments lasted for 24 h. As controls, animals were placed in
0.1x MMR containing 0.1% DMSO.
3.2.7 SDS-PAGE and immunoblotting
Tadpoles were euthanized in water containing 0.26% tricaine and 0.52%
sodium bicarbonate before decapitation and eyeball removal. After removal of
the lens and RPE cell layer, both isolated retinas from a single tadpole were
homogenized via sonication in 100 μL of SDS buffer (0.1% SDS v/w in PBS)
35
containing cOmplete Mini Protease Inhibitor Cocktail (Roche) and 0.1% β- mercaptoethanol. Proteins in these samples were resolved by SDS- polyacrylamide gel electrophoresis using a standard method employing a Mini-
PROTEAN Tetra Cell (Biorad) powered by a Powerpac HC (Biorad). Transfers
were performed by a semi-dry transfer cell (Trans-Blot SD Semi-Dry Transfer
Cell, Biorad) following the manufacturer’s recommended protocol. Membranes
were blocked in a 5% (w/v) dried milk (Nestle Carnation) in PBS for 30 min prior
to antibody incubation. Primary antibodies were added to PBS containing 0.1%
tween for 16 h at 4°C. Membranes were washed in PBS containing 0.1% tween 3
times for 10 min each, and then incubated in secondary antibody solution for 1 h
at RT. Antibodies and dilutions used were as follows; anti-β tubulin (Chu and
Klymkowsky, 1989) (Hybridoma Bank AB_2315513, RRID AB_2315513, 1:500),
anti-Na+/K+ ATPase α (Santa Cruz M7-PB-E9, 1:500), and anti-rhodopsin (B6-30,
2 μg/ml; a kind gift from Dr. David Salom, Case Western Reserve Univ., RRID:
AB_2572379, 1:3000). Secondary antibody used for immunoblotting was anti-
mouse IgG-HRP (SC-2005, Santa Cruz, 1:1000). After secondary antibody
incubation, membranes were washed 3 times for 10 min each before being
exposed to HyGLO Chemiluminescent HRP Antibody Detection Reagent Quick
Spray (Denville Scientific). After HRP exposure, membranes were immediately
imaged by a Tanon 5200 developer (Tanon Science and Technology Co.).
3.2.8 Preparation of retina explant for confocal imaging
Tadpoles were euthanized, decapitated, and their eyes were removed as
described previously (Lodowski et al., 2013). The RPE layers were separated
36
from the retinas, which were immediately placed into a glass-bottom dish (P35G-
1.5-14-C, MatTek Corporation) containing modified Wolf medium (D-glucose, 700 mg/L, 30 mM NaHCO3, 55% MEM, 31% sodium-free BBS, 10% FBS) (Lodowski et al., 2013). Equilibration of Wolf medium was achieved by incubating the dish containing the medium in an incubation chamber (Tokai Hit) supplied with 5%
CO2 and 95% O2 (Airgas) (Nemet et al., 2014). The glass-bottom dishes were
coated with Cell-Tak Cell and Tissue Adhesive (Corning). A circular glass
coverslip (12 mm diameter and 0.13 to 0.17 mm thickness) was used to seal and
flatten the retinas before imaging.
3.2.9 Confocal microscopy, Image analysis, quantification, and statistical
analysis
All images were acquired using a Leica TCS SP2 laser scanning
confocal/multiphoton microscope system equipped with four lasers for excitation:
488 nm argon ion, 543 nm HeNe, 633 nm HeNe, and tunable Chameleon XR
Ti:Sapphire laser (Leica Microsystems) as described previously (Lodowski et al.,
2013). A HCX PL APO CS 40.0 X 1.25 oil UV objective lens was used for
imaging. For the quantitative studies involving drug treatments, the same laser
power and imaging condition were used for all the samples. The cytoplasmic
space was highlighted manually and its mean fluorescence intensity measured
using ImageJ’s freehand selection tool (Schneider et al., 2012). The background
noise was determined from blank space where no cells were observed, and
subtracted from the measurements. All the quantitative data were obtained from
4 independent animals (or structures thereof) are represented as mean ± SD. In
37
comparing two populations, *p < 0.001 by Student’s t-test was considered
statistically significant.
3.3 Results and Discussion
3.3.1 The Na+ /K+-ATPase is downregulated in cells expressing class I
mutant rhodopsin
In healthy rod photoreceptors, rhodopsin does not accumulate on the IS
PM. In rods afflicted by ciliopathy, the daily accumulation, removal, and
degradation of PM-mislocalized rhodopsin would likely have negative effects on
the PM protein homeostasis. An essential component of the photoreceptor IS PM
is the Na+ /K+-ATPase which plays important roles in maintaining the Na+ and K+
gradients required for the photoresponses of cone and rod cells (Wetzel et al.,
1999). In our experiments, RhoQ344ter-Dend2 and Rho-Dend2-1D4 are expressed
under the regulation of the Xenopus rhodopsin promoter which drives expression
specifically in rods (Mani et al., 2001). By immunofluorescence microscopy,
robust and consistent expression of Na+ /K+-ATPase alpha subunit (NKAα) was observed in cone cells (Figure 7A, white asterisks). In contrast, the expression
levels of NKAα were attenuated in the rod cells expressing RhoQ344ter-Dend2
(Figure 7A, RhoQ344ter-Dend2, arrowheads). The amount of NKAα on the IS PM of
rods expressing Rho-Dend2-1D4 appeared to be higher than that of rods
expressing RhoQ344ter-Dend2 (Figure 7A, Rho-Dend2-1D4 and RhoQ344ter-Dend2,
arrowheads), indicating the Q344ter class I mutation is the cause of NKAα
38
downregulation. NKAα downregulation was independently confirmed via
immunoblot analysis, which revealed an approximate 50% decrease in NKAα in
retinas expressing RhoQ344ter-Dend2 (Q1) compared to non-transgenic (NT) retinas (p < 0.01) (Figures 7B and 7C, IB). This modest decrease, compared to the decrease observed in individual rods, was consistent with the presence of
NKAα in secondary retinal neurons which were not affected by RhoQ344ter-Dend2.
Based on quantitative PCR analysis, we found that downregulation of NKAα protein is not due to reduced mRNA levels. The delta Ct value compares relative expression of a pair of genes (Mahanty et al., 2017). When we quantified NKAα transcript amounts relative to β-tubulin’s, the delta Ct values werenot significantly different between non-transgenic (NT) retinas and retinas expressing RhoQ344ter-
Dend2 (Q1) (Figure 7C, qPCR). Because of the lack of significant change in
gene expression and the significant difference in the protein level, we surmised
that NKAα is downregulated by one or more posttranslational processes, and
likely by lysosome-mediated degradation.
39
Figure 7. Na+ /K+-ATPase (NKAα) is downregulated in rods expressing RhoQ344ter-Dend2. (A) Immunofluorescence labeling of NKAα (red) in retinas expressing either Rho- Dend2-1D4 (Left column) or RhoQ344ter-Dend2 (right column). NKAα signal on the PM of cells expressing wild type rhodopsin was relatively uniform (Rho-Dend2- 1D4, NKAα, arrowheads) while NKAα signal was noticeably reduced or absent from cells expressing class I mutant rhodopsin (RhoQ344ter-Dend2, NKAα, arrowheads). NKAα expression was robust in cone cells (white asterisks). (B) Representative immunoblot comparing the expression levels of NKAα and β-tubulin in non-transgenic retinas (NT) or retinas expressing RhoQ344ter-Dend2 (Q1). Molecular weight (MW) is indicated in kDa. (C). Left side of graph (IB, delimited by dotted line): In nontransgenic retinas (NT) or retinas expressing high amount of RhoQ344ter-Dend2 (Q1), the protein amounts of NKAα were quantified and normalized to the amounts of β-tubulin protein. NT value was standardized to 1.00. (Exact values were 1.00 ± .0.19 for NT and 0.50 ± 0.12 for RhoQ344ter-Dend2, based on n = 6 animals, p < 0.01). Right side of graph (qPCR): the amount of NKAα transcript relative to β-tubulin transcript was assessed by quantitative PCR. Delta Ct values of non-transgenic and high-expression RhoQ344ter-Dend2 retinas were compared (4.60 ± 0.27 for RhoQ344ter-Dend2 retinas and 4.42 ± 0.78 for non- transgenic retinas based on n = 5 animals, p = 0.63). Scale bar = 10 μm
40
3.3.2 NKAα downregulation is correlated to the expression level of
RhoQ344ter-Dend2 in individual rods.
In F0 transgenic retinas expressing either RhoQ344ter-Dend2 or Rho-Dend2-
1D4, individual rods express varying amounts of the protein product likely due to position-effect variegation (Moritz et al., 2001). Individual rods were categorized based on the expression levels (Low and High) of RhoQ344ter-Dend2. We found that the higher the expression level of RhoQ344ter-Dend2, the less NKAα was present on the IS PM (Figure 8A, Low Q and High Q, arrowheads). As controls, nontransgenic rods (NT) and rods expressing Rho-Dend2-1D4 (WT) did not exhibit downregulations of NKAα (Figure 8A, NT and WT, arrowheads).
Quantification of NKAα in the apical region of the IS PM (Figure 8A, NKAα, arrowheads) revealed that while rods with low expression (Low Q) of RhoQ344ter-
Dend2 had reduced NKAα presence on the IS PM compared to non-transgenic and Rho-Dend2-1D4 transgenic rods, rods with high expression (High Q) of
RhoQ344ter-Dend2 had the lowest amounts of NKAα compared to all groups tested
(p < 0.001) (Figure 8B, compare High Q to Low Q, NT, and WT). The inverse relationship between the expression levels of RhoQ344ter-Dend2 and NKAα is consistent with a model in which these two proteins are co-degraded: higher expression of RhoQ344ter-Dend2 would result in more active lysosomal degradation, which would coincidentally lower the NKAα levels.
41 zz
Figure 8. Degree of NKAα downregulation is dependent on the amount of class I mutant rhodopsin. (A) Direct comparison of NKAα immunofluorescence in a rod cell not expressing RhoQ344ter-Dend2 (non-transgenic, NT), a rod cell expressing low amount of RhoQ344ter-Dend2 (Low Q), a rod cell expressing high amount of RhoQ344ter-Dend2 (High Q), and a rod cell expressing Rho-Dend2-1D4 (WT). Arrowheads indicate the apical region of the IS PM. Scale bar = 10 μm (B) Quantification of NKAα immunofluorescence observed in (A). Compared to NT and WT rods, NKAα was downregulated in rods expressing class I mutant rhodopsin in a manner depending on the expression level of RhoQ344ter-Dend2. Rods with low expression of RhoQ344ter- Dend2 (Low Q) had higher quantities of NKAα on the PM than rods with high expression of RhoQ344ter-Dend2 (High Q). By ANOVA Tukey post-hoc test, all the pairs (a vs. b, a vs. c, and b vs. c) were significantly different (p < 0.001, based on 52 cells from n = 4 animals for each condition) except for between WT and NT rods (a, p = 0.58), which indicated that the presence of the wild type rhodopsin transgene did not affect the expression of NKAα.
42
3.3.3 NKAα is co-trafficked with RhoQ344ter-Dend2 to lysosomes in vivo
Co-degradation of RhoQ344ter-Dend2 and NKAα would require these two
proteins to be in the same lysosomal compartments. To visualize the possible
colocalization in vivo, we fused NKAα to fluorescent protein PSmOrange (Subach
et al., 2011) which emits orange fluorescence distinguishable from green
Dendra2 fluorescence. PSmOrange-NKAα and RhoQ344ter-Dend2 were coexpressed in Xenopus tadpoles which were then treated with BA1 in order to inhibit lysosome-mediated degradation (Figure 9A). After 24 h of BA1 treatment,
PSmOrange-NKAα and RhoQ344ter-Dend2 accumulated together in intracellular vesicles of rod ISs (Figure 9A, BA1, arrowheads). Based on the fluorescence intensities, the amount of PSmOrange-NKAα in the ISs was approximately 2.15 times higher (p < 0.001) in BA1-treated rods than in untreated rods (Figure 9B,
compare NKAα+Q344ter with (+) and without (-) BA1). To test if lysosomal accumulation of PSmOrange-NKAα was dependent on RhoQ344ter-Dend2, we
repeated the experiment using animals expressing PSmOrange-NKAα, but not
RhoQ344ter-Dend2 (Figure 9C). After BA1 treatment, PSmOrange-NKAα did not
significantly accumulate (NS, p = 0.35) in the ISs over untreated rods (Figure 9B,
compare NKAα with (+) and without (-) BA1). These results indicate that
lysosomal localization of PSmOrange-NKAα was induced by RhoQ344ter-Dend2.
43
Figure 9. RhoQ344ter-Dend2 induces internalization of NKAα.
(A) RhoQ344ter-Dend2 (green) was expressed together with NKAα fused to PSmOrange fluorescent protein (red) in rod photoreceptors. These transgenic tadpoles aged 9 DPF were treated for 24 h with either DMSO (Control) or 100 nM BA1 (BA1). BA1-treated rods exhibited vesicles containing both RhoQ344ter-Dend2 and PSmOrange-NKAα in the ISs. (B) Comparative quantification of intracellular fluorescence from PSmOrange-NKAα in either the presence (+) or absence (-) of 100 nM BA1 and either with (+Q344ter) or without coexpression of RhoQ344ter-Dend2. Nearly equal amounts of PSmOrange-NKAα were observed among untreated and BA1-treated cells expressing only PSmOrange-NKAα (NS, p = 0.35 based on 80 untreated cells and 140 BA1-treated cells from n = 4 animals each). Conversely, BA1-treated cells expressing both PSmOrange-NKAα and RhoQ344ter-Dend2 had significantly increased amounts of PSmOrange-NKAα in the ISs compared to untreated cells expressing both PSmOrange-NKAα and RhoQ344ter-Dend2 (p < 0.001, based on 160 untreated cells and 173 BA1-treated cells from n = 4 animals each). This difference suggests that the presence of RhoQ344ter-Dend2 was required to bring PSmOrange-NKAα to the IS lysosomes for degradation. (C) Cells expressing only the PSmOrange-NKAα fusion protein without RhoQ344ter-Dend2 did not significantly accumulate PSmOrange-NKAα upon treatment with BA1. Scale bars = 10 μm.
44
3.4 Conclusions
In this Chapter, I found that mislocalized rhodopsin induces co- degradation of NKAα, and this degradation led to severe downregulation of
NKAα on the IS PM. Downregulation of NKAα was not due to overexpression of transgene in our Xenopus model, but due to specific effect of class I mutant
RhoQ344ter-Dend2 mislocalized to the IS PM. In our experiments, the transgene is
not overexpressed, and is only observed at levels between 1 – 5% of
endogenous rhodopsin (Lodowski et al., 2013). Moreover, transgenic expression
of wild type Rho-Dend2-1D4 did not alter the amount of NKAα. The effect we
observed in our class I model may also be occurring in other inherited blinding
disorders which exhibit rhodopsin mislocalization. Rhodopsin mislocalization is
commonly observed in a subset of Joubert Syndrome (Lessieur et al., 2017),
Bardet-Biedl syndrome (Nishimura et al., 2004; Simons et al., 2011), as well as
many other forms of retinitis pigmentosa in which rhodopsin is not the defective
gene (Hagstrom et al., 2001; Gao et al., 2002; Adamian et al., 2006). Therefore,
further studies are warranted to determine if NKAα downregulation is occurring in
these diseases.
45
Chapter 4: Dysfunction of NKA results in photoreceptor degeneration
The material in this Chapter was adapted from Ropelewski P, Imanishi Y, 2019:
Disrupted plasma membrane protein homeostasis in a Xenopus laevis model of
retinitis pigmentosa. The Journal of Neuroscience 39(28):3025-18.
4.1 Introduction
In RP, the first observable signs of photoreceptor degeneration are the
shortening and loss of the photoreceptor OSs (Milam 1998), compromising the
vision of afflicted patients. The mechanism leading to the compromised OS
structures remain uncharacterized.
Studies in Chapter 3 indicates that NKA concentration is low in the IS PM of a RP model. Therefore, we asked if the attenuated function of NKA was a contributing factor in causing OS defects. Animal models deficient in NKA demonstrate rapid rod photoreceptor degeneration (Luan et al., 2014; Landfried et al., 2017) . In these models, however, NKA deficiencies occur during the retina
development, and cannot emulate the progressive rod degeneration which
occurs after complete maturation of rod photoreceptors. Instead of genetic
manipulation, pharmacological methods are well-established to inhibit NKA
functions in developed animals and humans (Demontis et al., 1995). For
example, a NKA inhibitor, digoxin, is utilized for treating human patients with
46 atrial fibrillation, but the extended treatment results in vision defects of both animals and human (Landfried et al., 2017). Thus we tested the hypothesis that compromised NKA function is the root cause of vision defects observed in RP patients, and such compromised NKA function results in shortening or loss of OS structures. Toward the goal of testing this hypothesis, I combined genetic and pharmacological models together with state of the art microscopy techniques.
These analyses answered the question of the effect of NKA inhibition on rod photoreceptor cell health and maintenance.
4.2 Materials and Methods
4.2.1 Animals
All animal procedures were approved by the Institutional Animal Care and
Use Committee (IACUC) at Case Western Reserve University. Adult female and male frogs were purchased from Nasco and housed at 16° C under a 12-h light/12-h dark cycle. All tadpoles used for experiments were housed at 16° C in
24-h darkness. Tadpoles were fed spirulina (Nuts.com) until metamorphosis was complete, and then were fed pelleted frog brittle (Nasco). Both male and female tadpoles were used for all experiments.
4.2.2 Reagents
Unless otherwise specified, all reagents were purchased from either
Fischer Scientific or Sigma-Aldrich.
4.2.3 Transgenesis of Xenopus laevis
47
Transgenic Xenopus laevis were produced using the intracytoplasmic
sperm injection (ICSI) method following the previously published procedure
(Sparrow et al., 2000; Smith et al., 2006; Lodowski et al., 2013). Expression
vectors for Rho-Dend2-1D4 and RhoQ344ter-Dend2 were purified from bacteria cultures using EndoFree Plasmid Midi Prep Kit (QIAGEN), and then used to isolate DNA fragments containing the XOP promoter, coding/noncoding regions, and polyadenylation signal for transgenesis. Fully developed tadpoles were
screened for the presence of Dendra2 fluorescence in their eyes at days post-
fertilization (7 DPF). Tadpoles were immersed in 6% methylcellulose to prevent
movement while probed for green fluorescence in their eyes using a Leica
MZ16F stereoscope (Leica Microsystems). Tadpoles were classified into one of
the three following categories based on the expression level of the transgene:
low, medium, or high as assessed by intensity of green fluorescence in their
eyes. In order to prevent photobleaching, light-dependent photoreceptor
degeneration, and unintended photoconversion of fluorescent protein Dendra2,
tadpoles were reared in 24-h darkness.
4.2.4 Immunohistochemistry
Xenopus laevis eyes were fixed as previously described (Lodowski et al.,
2013). Tadpoles were first euthanized in a 0.26% tricaine and 0.52% sodium
bicarbonate solution (in dH2O) and decapitated. Their heads were fixed in 4%
paraformaldehyde (Electron Microscopy Sciences) in 0.1 M phosphate buffer
(PB, NaHPO4 and Na2HPO4 dissolved in MilliQ H2O, pH 7.4) for 6 h at RT. Fixed
heads were incubated in 5% sucrose in phosphate buffer for 30 min, and then
48
subsequently washed in 10%, 15%, and 20% sucrose solutions in 0.1 M PB. The
heads were then incubated overnight at 4 ºC in a solution containing 20%
sucrose and optimum cutting temperature compound (O.C.T. compound, Sakura
Finetek) in a 2:1 ratio. The following day, heads were aligned in Tissue-Tek
cryomolds (Sakura Finetek) and frozen in isopentane cooled by liquid N2. Frozen tissue sections (12 μm) were cut using a Leica CM1850 cryostat (Leica
Microsystems) and collected on Superfrost Plus microscopy slides. Sections
were dried and stored at -80 ºC until use. Slides were removed from -80 ºC
storage conditions, warmed using a 37 ºC incubator (CCC 0.5d, Boekel
Scientific), and then sections were encircled using an ImmEdge pen (Vector
Laboratories). Sections were rehydrated and blocked in a 1.5% normal goat serum diluted in phosphate-buffered saline solution (PBS, mM NaCl, 2.7 mM
KCl, 8 mM Na2HPO4, 2 mM KH2PO4) for 30 min at RT. Blocking solution was then removed and primary antibody (1:100) diluted in PBS containing 0.1% Triton
X-100 (PBST) was added; sections were incubated at RT for approximately 14 h and then washed 3 times with PBST for 10 min each. After the washes, sections were incubated in PBST containing anti-mouse Cy3 (1:100) and Hoechst dye
(1:1000) for 1 h at RT followed by an additional 3 washes with PBST for 10 min
each. After these washes, one drop of Vectashield mounting medium (Vector
Laboratories) was added to a square coverslip and placed over the section. Nail
polish (Electron Microscopy Sciences) was used to seal the coverslip, and the
sections were dried at RT for > 30 min before imaging. Antibodies used for
immunofluorescence imaging were as follows: mouse anti-Na+ /K+-ATPase
49
(Lebovitz et al., 1989) (Hybridoma Bank, AB_2166869), mouse anti- peripherin/rds (anti-xrds 2D4, a kind gift from Dr. Robert Molday, Univ. of British
Columbia), mouse XAP-2 (clone 5B9, Hybridoma Bank), Hoechst (Thermo Fisher
33342), and anti-mouse-Cy3 conjugate (Jackson, 115-165-166).
4.2.5 SDS-PAGE and immunoblotting
Tadpoles were euthanized in water containing 0.26% tricaine and 0.52% sodium bicarbonate before decapitation and eyeball removal. After removal of the lens and RPE cell layer, both isolated retinas from a single tadpole were homogenized via sonication in 100 μL of SDS buffer (0.1% SDS v/w in PBS) containing cOmplete Mini Protease Inhibitor Cocktail (Roche) and 0.1% β- mercaptoethanol. Proteins in these samples were resolved by SDS- polyacrylamide gel electrophoresis using a standard method employing a Mini-
PROTEAN Tetra Cell (Biorad) powered by a Powerpac HC (Biorad). Transfers were performed by a semi-dry transfer cell (Trans-Blot SD Semi-Dry Transfer
Cell, Biorad) following the manufacturer’s recommended protocol. Membranes were blocked in a 5% (w/v) dried milk (Nestle Carnation) in PBS for 30 min prior to antibody incubation. Primary antibodies were added to PBS containing 0.1% tween for 16 h at 4°C. Membranes were washed in PBS containing 0.1% tween 3 times for 10 min each, and then incubated in secondary antibody solution for 1 h at RT. Antibodies and dilutions used were as follows; anti-β tubulin (Chu and
Klymkowsky, 1989) (Hybridoma Bank AB_2315513, RRID AB_2315513, 1:500), anti-Na+ /K+ 225 ATPase α (Santa Cruz M7-PB-E9, 1:500), and anti-rhodopsin
(B6-30, 2 μg/ml; a kind gift from Dr. David Salom, Case Western Reserve Univ.,
50
RRID: AB_2572379, 1:3000). Secondary antibody used for immunoblotting was anti-mouse IgG-HRP (SC-2005, Santa Cruz, 1:1000). After secondary antibody incubation, membranes were washed 3 times for 10 min each before being exposed to HyGLO Chemiluminescent HRP Antibody Detection Reagent Quick
Spray (Denville Scientific). After HRP exposure, membranes were immediately imaged by a Tanon 5200 developer (Tanon Science and Technology Co.).
4.2.6 Treatment of tadpoles by NKA inhibitor
Animals were treated in 0.1x Mark’s Modified Ringer (MMR) solution (10x
MMR composition: 1 M NaCl, 20 mM KCl, 20 mM CaCl2, 10 mM MgCl2, 50 mM
HEPES, pH 7.5) (Lodowski et al., 2013) containing 0.1% DMSO and either digoxin (NKA inhibitor, 3.5 μM), ouabain (NKA inhibitor, 1 μM), wedelolactone
(NKA inhibitor, 3.5 μM), istaroxime (NKA inhibitor, 3 μM), or 3,4,5,6-
Tetrahydroxylxanthone hydrate (NKA inhibitor, 10 μM). All inhibitors were initially diluted in DMSO, and then applied to 5 ml of 0.1x MMR buffer containing a single tadpole. NKA inhibitor treatments lasted for 3-7 days. As controls, animals were placed in 0.1x MMR containing 0.1% DMSO. For all treatments that lasted longer than 24 h, the 0.1x MMR solutions containing the drug in 0.1% DMSO were replaced daily in experimental groups, whereas 0.1x MMR solutions containing
0.1% DMSO were replaced daily in control groups.
4.2.7 Confocal microscopy, Image analysis, quantification, and statistical analysis
51
All images were acquired using a Leica TCS SP2 laser scanning
confocal/multiphoton microscope system equipped with four lasers for excitation:
488 nm argon ion, 543 nm HeNe, 633 nm HeNe, and tunable Chameleon XR
Ti:Sapphire laser (Leica Microsystems) as described previously (Lodowski et al.,
2013). A HCX PL APO CS 40.0 X 1.25 oil UV objective lens was used for
imaging. Lengths of outer segments were measured using ImageJ’s line function tool (Schneider et al., 2012). All the quantitative data were obtained from 4 independent animals (or structures thereof) are represented as mean ± SD. In comparing two populations, *p < 0.001 by Student’s t-test was considered statistically significant.
4.3 Results and Discussion
4.3.1 Rod photoreceptors expressing RhoQ344ter-Dend2 have shorter,
disorganized OSs
We labeled RhoQ344ter-Dend2 transgenic retinas (21 DPF) with
antiperipherin/rds (P/rds), a marker for rod and cone OS disk membranes
(Arikawa 1992; Tian 2014). Rod cells expressing RhoQ344ter-Dend2 had
noticeably disorganized OSs compared with those expressing Rho-Dend2-1D4
52
Figure 10. Rods expressing RhoQ344ter-Dend2 exhibit disorganized rod OSs. Immunofluorescence labeling of outer segments (OSs) employing (A) anti-P/rds (red) and (B) XAP-2 (red) antibodies. Retina sections (21 DPF) expressing either Rho-Dend2- 1D4 (green) or RhoQ344ter-Dend2 (green) were labeled. (C) Rods with lower NKAα expression demonstrate shorter OSs. NKAα expression levels in RhoQ344ter-Dend2- positive rods (21 DPF) were normalized to average NKAα levels (100 %) in non- transgenic rods (21 DPF). Rods with three different ranges of NKA expression levels (80 – 100 %, 40 – 60 %, and 0 – 20 %) were analyzed. (D) Representative immunoblot comparing 21 DPF retina samples of either non-transgenic (NT) or RhoQ344ter-Dend2- expressing (Q1) animals. Primary antibodies against β-tubulin and P/rds were used in combination with anti-mouse HRP for chemiluminescence detection of the signals. (E) Representative blot comparing 21 DPF retina samples of either non-transgenic (NT) or RhoQ344ter-Dend2 (Q1) retinas. Primary antibody against rhodopsin (B6-30) was used in combination with anti-mouse HRP for chemiluminescence detection of the signals originating from endogenous Xenopus rhodopsin. Molecular weight (MW) marker is in kDa for both (D) and (E). (F) Quantification of immunopositive signals in (D) and (E). NT average value was standardized to 1.00 for both P/rds (normalized to β-tubulin) and total endogenous rhodopsin, for readily comparison with groups expressing RhoQ344ter-Dend2. For immunoblot against P/rds, the relative values were 1.00 ± 0.23 for NT and 0.52 ± 0.42 for RhoQ344ter-Dend2 (based on n = 6 animals, p = 0.025). For immunoblot against total endogenous rhodopsin, the relative values were 1.00 ± 0.21 for NT and 0.14 ± 0.37 for RhoQ344ter-Dend2 (based on n = 6 animals, p < 0.001). Scale bars = 10μm
53
(Figure 10A). RhoQ344ter-Dend2 retinas exhibited fewer and shorter P/rdspositive
OSs compared with retinas expressing Rho-Dend2-1D4 (Figure 10A, P/rds). In
addition to anti-P/rds, which labels both cone and rod OSs, we also used a XAP-
2 antibody that specifically labels Xenopus rod OSs (Figure 10B; Viczian 2009;
Choi 2011). Consistent with the results obtained for P/rds-positive OSs,
RhoQ344ter-Dend2 transgenic retinas contained ~30% fewer (p < 0.001; based on
retinas from n = 6 animals) XAP-2-positive OSs (per 180 µm retina length; Table
1) than Rho-Dend2-1D4 transgenic retinas (Figure 10B, XAP-2). Additionally,
RhoQ344ter-Dend2 rods were ~44% shorter than Rho-Dend2-1D4 rods (p < 0.001;
based on 65 OSs from n = 6 animals for each condition; Table 1). To clarify the
relationship between NKA downregulation and OS defects, we measured the
NKA levels and OS lengths in the same individual rods expressing RhoQ344ter-
Dend2. Rods expressing RhoQ344ter-Dend2 had variable degrees of NKA downregulation (Figure 10C). For this analysis, we categorized individual rods
(192 measurements from n = 6 individual animals) into three groups depending on the severity of the downregulation. Rods with NKAα levels similar (80 –100%) to the nontransgenic values had OSs that were 16.0 -- 4.0 µm long, and thus comparable to OS lengths of nontransgenic rod photoreceptors (Table 1, untreated). Rods with NKAα downregulation demonstrated significantly shorter rod OSs (p < 0.001); rods that contained 40 –60% of the normal NKAα level had
OSs that were 11.3 ± 4.2 µm long, whereas rods that expressed 20% of the
average nontransgenic NKAα level had OSs that were 6.1 ± 3.5 µm long
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(Figure 10C). This analysis indicated that NKAα downregulation coincides with
the shortening of rod OSs.
4.3.2 Pharmacological inhibition of NKAα with digoxin mimics the degenerative phenotype observed in RhoQ344ter-Dend2
As additional tests to gauge OS loss, retinas from 21 DPF transgenic
tadpoles were subjected to immunoblotting analyses using antibodies against
P/rds and rhodopsin (Figure 10 D, E). The amount of P/rds, normalized to the
expression levels of -tubulin, was nearly 50% lower (p = 0.025) in Q1 retinas
compared with NT retinas (Figure 10 D, F; representative blot based on n = 6
animals). The amount of endogenous Xenopus rhodopsin was ~85% lower (p <
0.001) in Q1 retinas than in NT retinas (Figure 10 E, F; n = 6 animals). While
there were significant differences in OS structure and number between
RhoQ344ter-Dend2 and Rho-Dend2-1D4 retinas, rod nuclei were only marginally
lost in RhoQ344ter-Dend2 retinas at this stage: the average number of nuclei was
only 10% lower in retinas expressing RhoQ344ter-Dend2 than in retinas expressing
Rho-Dend2-1D4 (Table 1; p = 0.017 based on n = 6 animals). This minor
decrease suggests that OS shortening and loss both precede rod cell death.
These results indicate that rhodopsin mislocalization, as well as NKA
downregulation, contribute to OS shortening and loss, which are the beginning
signs of retinal disease.
To examine whether compromised NKA activity can be causative to the
shortening or loss of rod OSs, we treated nontransgenic animals (starting age 14
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DPF) with digoxin, which specifically inhibits NKA. The integrity of the OS
structures was assessed by P/rds (Figure 11A) and XAP-2 (Figure 11B)
immunofluorescence. After 3 d, retinas from digoxin-treated animals exhibited
shorter and fewer OS structures than untreated animals. On average, the rod
OSs of 3 d treated retinas were ~20.0% shorter (p < 0.001; based on analyzing
96 OSs from n = 4 untreated animals and 120 OSs from n = 4 treated animals)
and 30% fewer (p < 0.03; based on retinas from n = 4 animals each for treated
and untreated) than those of untreated retinas. After 7 d of treatment, this effect
was exacerbated. On average, the rod OSs of 7 d digoxin-treated retinas were
~29% shorter (p < 0.001; based on analyzing 250 OSs from n = 5 untreated
animals and 170 OSs from n = 5 treated animals; Table 1) and ~34% fewer (p <
0.001; based on retinas from n = 5 animals each for treated and untreated; Table
1) than those of untreated retinas. The magnitude of OS shortening and loss
observed in 7 d digoxin-treated animals, which were 21 DPF at the time of
analysis, was similar to that seen in RhoQ344ter-Dend2 animals, which were also analyzed at 21 DPF (Table 1). Although OS structures were affected by the digoxin treatment, the number of nuclei per retina area (180 µm) remained nearly identical between treated and untreated animals for both 3 (p = 0.96; based on n
= 4 animals) and 7 (p = 0.56; based on n = 5 animals) day treatments (Table 1).
This result is similar to the pathogenic process of RP beginning with OS
shortening and loss before the onset of rod cell death (Milam et al., 1998).
Immunoblotting revealed a 30% decrease in the total endogenous rhodopsin
levels in digoxin-treated (7 d) retinas (Figure 11C), which is consistent with the
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Figure 11. Inhibition of NKA results in disorganized rod OSs of Xenopus laevis. Xenopus tadpoles (non-transgenic) were treated with digoxin, which is a specific inhibitor of NKA. Control animals were treated with 0.1% DMSO and were housed in the same conditions. (A) Immunofluorescence labeling of photoreceptor OSs (OS layer labeled as “OS”) employing anti P/rds after 3 or 7 days of either 3.5 μM digoxin treatment (Digoxin) or no treatment (Control). Digoxin-treated retinas had fewer and shorter OS structures compared to control retinas; this effect was exacerbated with longer treatment time (compare 3 and 7 days). (B) Immunofluorescence labeling of rod OSs (OS layer labeled as “OS”) employing XAP-2 antibody after 3 or 7 days of digoxin treatment. Treated retinas (Digoxin) had shorter and fewer OS structures than untreated retinas (Control). (C) Representative immunoblot comparing endogenous rhodopsin levels between digoxin-treated (Digoxin) and untreated (Untreated) retinas. Mean rhodopsin signal in untreated retinas was standardized to 1.00 (a.u.). The relative values were 1.00 ± 0.31 (n = 6 animals) for untreated retinas and 0.69 ± 0.14 (n = 6 animals) for digoxin-treated retinas. Molecular weight (MW) marker is in kDa. Scale bars = 10 μm
58
occurrence of OS loss and shortening. These results suggest that
compromised NKA function, whether via pharmacological inhibition or co-
degradation with RhoQ344ter-Dend2, leads to compromised rod OSs and reduced
amounts of rod OS membrane proteins as represented by P/rds and rhodopsin.
To rule out possible nonspecific effects of digoxin, we tested four
additional NKA inhibitors that are structurally distinct from digoxin (Fig. 12; Table
1) and hence unlikely to share off-targets. Treatments with these four NKA inhibitors, for up to 7 d, resulted in time-dependent shortening and loss of rod
OSs as observed for digoxin-treated retina (Figure 12A–D, compare 3 and 7 d).
As with the studies of digoxin, photoreceptor outer segments were visualized by
P/rds and XAP-2 immunofluorescence. Ouabain is a steroidal glycoside more
potent and selective than digoxin (Gable et al., 2017), whereas istaroxime is a
steroidal but non-glycoside molecule (De Munari et al., 2003). After 7 d of treatment with ouabain, rod OSs were 17–23.5% (p < 0.001) shorter and 12–
22.5% fewer (p < 0.05) on average compared with age-matched untreated
retinas (Table 1). We also tested the effects of NKA inhibitors that are unrelated
to steroidal or glycoside NKA inhibitors, and thus do not share the mechanism of
action with digoxin. Wedelolactone and 3,4,5,6-tetrahydroxyxanthone hydrate
(TTX) are coumestan and xanthone-based structures, respectively (Wang and
O’Doherty, 2012), and do not share a structural backbone with digoxin. In
inhibiting NKA, these classes of molecules demonstrate different mechanisms of
action and binding to NKA (Poˆc¸as et al., 2003; Zhang et al., 2010; Wang and
O’Doherty, 2012). After wedelolactone or TTX treatment, the rod OSs were 8 –
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Figure 12. NKA inhibitors structurally distinct from digoxin induce disorganization of rod OSs. Photoreceptor OSs (OS layer labeled as “OS”) were labeled by anti-P/rds antibody (red) which binds to both cone and rod OSs, or by XAP-2 antibody which specifically binds to rod OSs (red). (A) Animals were treated with 1.0 μM ouabain for 3 or 7 days. (B) Animals were treated with 3.5 μM wedelolactone for 3 or 7 days. (C) Animals were treated with 3.0 μM istaroxime for 3 or 7 days. (D) Animals were treated with 10.0 μM 3,4,5,6-tetrahydroxylxanthone (TTX) for 3 or 7 days. Rod OSs were shorter and fewer after treatment with all the NKA inhibitors. Scale bars = 10 μm
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13.7% shorter (p = 0.043 for wedelolactone and p < 0.001 for TTX) and 23.3–
25.7% fewer in number (p < 0.001) than untreated retinas. Various degrees of
OS shortening and loss were observed among these inhibitors, potentially because of their different pharmacokinetics and other pharmacological properties. Nevertheless, these treatments invariably did not result in the statistically significant changes in the number of nuclei (Table 1; p > 0.05 for all untreated vs treated combinations). These experiments generally support the notion that NKA inhibition emulates the symptoms of early stage RP: OS shortening and loss without the significant loss of rod photoreceptor cells.
4.4 Conclusions
Separate lines of evidence indicate that compromised NKAα function, either due to pharmacological inhibition or downregulation by mislocalized rhodopsin, triggers shortening and loss of rod OSs. This finding is significant because rhodopsin mislocalization, OS shortening, and loss of OS structures are the most commonly observed events in retinitis pigmentosa patients (Li et al.,
1995; Milam et al., 1998), but their cause-effect relationships has been unclear until this study. The finding that NKAα downregulation causes photoreceptor dysfunction underscores the important role of NKAα in photoreceptor cell health and function. NKAα is expressed in high density on the IS PM of rod photoreceptors (Schneider et al., 1991; Wetzel et al., 1999). NKAα is an essential component of rod photoreceptors, and is responsible for generating the dark current and maintaining ion homeostasis within these cells (Demontis et al.,
61
1995; Roy et al., 2013). Shortening of OSs is possibly a homeostatic mechanism
rods employ to maintain the current per OS plasma membrane area, when the
corresponding current in the IS is reduced due to NKA downregulation. In the
retina of Drosophila, RNAi-mediated knockdown of NKAα resulted in loss of
photoresponse and rapid degeneration of photoreceptors (Luan et al., 2014).
Similarly, in the retina of mice, inhibition of NKAα by digoxin caused
photoreceptor-specific cell death (Landfried et al., 2017). In our model, NKAα
became downregulated due to constant co-degradation with mislocalized
rhodopsin. This downregulation of NKAα resulted in shortening and loss of rod
OSs; these defects were recapitulated by pharmacological inhibition of NKAα
pumping function. The degree of NKAα downregulation was 83.4 ± 11.2 % in
rods expressing a high-level of RhoQ344ter-Dend2. This degree of downregulation was similar to the extent of NKA inhibition (~ 80%) (Gable et al., 2017) by the
inhibitor concentrations (3.5 µM for digoxin and 1 µM for ouabain) introduced in
this study. Thus, those studies strongly suggest that NKA downregulation
induced by rhodopsin mislocalization is sufficient to cause rod OS shortening and
loss. NKAα inhibitors with distinct physicochemical and pharmacological
properties invariably caused rod OS loss and shortening without appreciable loss
of rod photoreceptors, emulating the conditions of early stage RP patients.
In addition to maintaining ion homeostasis, an additional function of NKA proteins has been proposed: as a binding partner for retinoschisin (Molday et al.,
2007), which is a photoreceptor adhesion molecule. Such adhesion is likely critical for photoreceptor-Muller glia and photoreceptor-bipolar cell interactions
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(Molday et al., 2007; Friedrich et al., 2011). Neurite sprouting accompanies
rhodopsin mislocalization in retinitis pigmentosa (Li et al., 1995; Tam et al.,
2006), which is suggestive of defective cell-cell interaction and adhesion.
Therefore, unlike pharmacological inhibition of NKAα, physical removal of NKAα induced by rhodopsin mislocalization may result in additional adverse effects to the PM due to the structural roles of Na+/K+-ATPase proteins.
In retinitis pigmentosa patients, shortening and loss of OSs are observed prior to photoreceptor cell death and compromise their vision (Milam et al., 1996; Oh et al., 2014). Class I mutant rhodopsin, mislocalized to the IS PM, causes shortening and loss of rod OSs at the early stages of pathogenesis. Our
observation that total endogenous Xenopus rhodopsin is reduced by
approximately 85% in RhoQ344ter-Dend2 retinas is consistent with previous finding that retinas expressing class I mutant rhodopsin had low amount of endogenous rhodopsin (Tam et al., 2006; Lodowski et al., 2013). Similar to RP patients, our
Xenopus model did not demonstrate significant death of rod photoreceptors.
Sustained downregulation or inhibition of NKA, however, would lead to rod photoreceptor cell death as observed for NKAβ2 knockout mouse (Molthagen et
al., 1996), digoxin-treated mouse (Landfried et al., 2017), and NKA-deficient
Drosophila (Luan et al., 2014) models. In the early stages of pathogenesis, as
tested in our study, the occurrence of OS loss/shortening is well-separated from the onset of cell death events. The number of nuclei per same retinal area stayed fairly consistent between retinas expressing Rho-Dend2-1D or RhoQ344ter-Dend2,
and also between untreated retinas or retinas treated with digoxin, indicating that
63 compromised NKAα function initially results in OS loss/shortening. This result is supported by our previous study in which only a nominal number of TUNEL- positive nuclei were observed in retinas expressing class I mutant rhodopsin
(Lodowski et al., 2013). Therefore, our Xenopus model will be useful to understand the early pathogenesis process of human retinitis pigmentosa.
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Chapter 5: Alternate routes of mislocalized rhodopsin removal
The material in this Chapter was adapted from a manuscript that is currently under consideration from eNeuro.
5.1 Introduction
Photoreceptor and RPE cells evolved a unique symbiotic relationship to
maintain the structure and function of the photoreceptive outer segments. Each
day, approximately 5 – 10 % of the OS is shed and engulfed by RPE cells, which
digest the components—the majority of which is rhodopsin—in phagosomes
located within the cytoplasm (Young, 1967; Kevany and Palczewski, 2010). This
relationship is disrupted in retinal ciliopathies in which the majority of rhodopsin
molecules are no longer destined to the OSs and instead mislocalize to the inner
segment plasma membrane (IS PM) (Sung et al., 1994; Li et al., 1996; Nishimura
et al., 2004; Deretic et al., 2005; Adams et al., 2007; Concepcion and Chen,
2010; Hollingsworth and Gross, 2013; Nemet et al., 2015). In various animal
models exhibiting rhodopsin mislocalization, rod photoreceptors expel rhodopsin-
laden vesicles which accumulate in the interphotoreceptor space (Li et al., 1996;
Hagstrom et al., 1999; Concepcion and Chen, 2010; Lodowski et al., 2013). The
interphotoreceptor space is in constant contact with RPE microvilli, which are
optimally positioned for phagocytic activities (Strauss, 2005). Increasing evidence
suggests various neurons shed vesicles as means of communication and
removing unwanted materials under neurodegenerative conditions (Nagarajah,
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2016; Fowler, 2019). More recently, RPE cells have been reported to take up
extracellular vesicles in an in vitro cell culture model (Nicholson et al., 2019).
Thus, as the first step in understanding the in vivo function of these
photoreceptor-derived vesicles, we asked if RPE cells are capable of engulfing
them in a manner analogous to OS phagocytosis. Such studies will shed light on
the symbiotic relationship between RPE and photoreceptors under disease
states.
Unlike the degradation of OS membrane proteins which has been
relatively well-characterized (Strauss, 2005; Kevany and Palczewski, 2010), little
is known about the degradation of IS PM proteins which lack access to the RPE
cells. Thus, we have initiated an effort in understanding the renewal of IS PM
proteins, especially focusing on class I mutant rhodopsin. In addition to vesicle-
mediated removal described above, mislocalized class I mutant rhodopsin is
degraded intracellularly: once reaching the IS PM, mislocalized rhodopsin
becomes internalized and subsequently degraded by lysosomes. IS PM
component Na+/K+-ATPase (NKA) plays a critical role in maintaining both the
dark current of photoreceptor cells (Yau and Baylor, 1989) and interactions
between bipolar and photoreceptor cells (Molday et al., 2007; Friedrich et al.,
2011). The lysosome-mediated removal of class I mutant rhodopsin induces co-
internalization and co-degradation of native NKA, compromising the structure
and function of rod photoreceptors. Another IS PM protein, HCN1 channel, plays a role in the normal electrophysiological response of photoreceptor cells, and its deficiency worsens the symptoms of retinitis pigmentosa (Schon et al., 2016).
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HCN1 contains a di-arginine ER retention signal which negatively regulate PM
transport (Pan et al., 2015a), suggestive of ER-associated degradation prior to
exiting the ER and reaching the PM. This mechanism appears to be important for
regulating the expression level of HCN1 at the level of IS PM (Pan et al., 2015b).
Despite improved understanding of HCN1 degradation during biosynthesis or
NKA degradation under pathological states, it is currently unknown if and how
rod photoreceptors coordinate intracellular and intercellular mechanisms for
degradation of endogenous IS PM proteins under normal physiological
conditions.
We investigated the fate of microvesicles shed by rod photoreceptor cells
expressing class I mutant rhodopsin. Toward the goal, we employed genetic
labeling technique which allowed clarifying the origin and destination of these
secreted vesicles. Then we asked if there is a sorting mechanism for packaging
specific IS PM protein to the microvesicles; we accomplished this by comparing
IS PM-mislocalized rhodopsin and a major IS PM resident protein, NKA. To
comprehensively understand the mode of renewing NKA, we compared the
extracellular and intracellular mechanisms of degrading NKA. Those studies will
provide novel insights into the protein homeostasis of rod photoreceptor cells
both in pathological and normal physiological conditions.
5.2 Materials and Methods
5.2.1 Animals
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All animal procedures were approved by the Institutional Animal Care and
Use Committee (IACUC) at Case Western Reserve University. Adult female and
male frogs involved in transgenesis were purchased from Nasco and housed at
16° C under a 12-h light/12-h dark cycle. All tadpoles used for experiments were
housed at 16° C in 24-h darkness. Tadpoles were fed spirulina (Nuts.com). Both
male and female tadpoles were used for all experiments.
5.2.2 Reagents
Unless otherwise specified, all reagents were purchased from either
Fischer Scientific or Sigma-Aldrich.
5.2.3 Molecular cloning
DNA expression vectors were generated by standard methods combining
PCR, DNA recombination, and site-directed mutagenesis. The expression vector
containing the Xenopus opsin promotor (XOP) followed by GFP-NKAα3 (Laird et
al., 2015) was modified. The vector contained an AgeI site upstream of GFP-
NKAα. Site-directed mutagenesis was used to create an additional AgeI site after
the region encoding GFP; GFP coding region was then removed using AgeI
enzyme and replaced with cDNA encoding Dendra2 fluorescent protein. The
resulting DNA fragment was introduced downstream of XOP and upstream of
NKAα using In-Fusion cloning kit (Takara). The plasmid vectors containing XOP-
Rho-Dend2-1D4 and XOP-RhoQ344ter-Dend2 were previously generated
(Lodowski et al., 2013). All the vectors contained polyadenylation signals following the coding and non-coding regions.
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5.2.4 Transgenesis of Xenopus laevis
Transgenic Xenopus laevis were produced using the intracytoplasmic
sperm injection (ICSI) method following the previously published procedure and
screened for the presence of Dendra2 fluorescence (Sparrow et al., 2000; Smith
et al., 2006; Lodowski et al., 2013). In order to prevent photobleaching, light-
dependent photoreceptor degeneration, and unintended photoconversion of
fluorescent protein Dendra2, tadpoles were reared in 24-h darkness.
5.2.5 Immunohistochemistry
Xenopus laevis retinal sections (12 µm) were generated and stored at -80
C as described previously (Lodowski et al., 2013). Slides were removed from -80
ºC storage conditions and warmed using a 37 ºC incubator (CCC 0.5d, Boekel
Scientific). After thawing the sections, all the following procedures were
conducted at room temperature. Individual sections were encircled by an
ImmEdge pen (Vector Laboratories). Sections were then rehydrated and blocked
in a 1.5% normal goat serum diluted in phosphate-buffered saline solution (PBS,
137 mM NaCl, 2.7 mM KCl, 8 mM Na2HPO4, 2 mM KH2PO4) for 30 min. Blocking
solution was then removed and primary antibody mouse anti-NKAα (Santa Cruz
Biotechnology, M7-PB-E9,1:100) diluted in PBS containing 0.1% Triton X-100
(PBST) was added; sections were incubated for approximately 14 h and then washed 3 times with PBST for 10 min each. After the washes, sections were incubated in PBST containing phalloidin conjugate (Alex Fluor 633 phalloidin
A22284), Hoechst 33342 dye (1:500), and goat anti-mouse IgG conjugated either with Alexa-488 (Jackson ImmunoResearch Laboratories, 115-545-166, 1:100) or
69
with Cy3 (Jackson ImmunoResearch Laboratories, Inc, 115-165-166,1:100) for 1
h followed by 3 additional 10 min washes with PBST. After these washes,
individual sections were mounted with a coverslip and one drop of
VECTASHIELD mounting medium (Vector Laboratories). Nail polish (Electron
Microscopy Sciences) was used to seal the coverslip, and the sections were
dried for > 30 min before imaging.
5.2.6 Preparation of retina explant for confocal imaging
Tadpoles were euthanized, decapitated, and their eyes were removed as
described previously (Lodowski et al., 2013). The RPE layers were separated
from the retinas, which were immediately placed into a glass-bottom dish (P35G-
1.5-14-C, MatTek Corporation) containing modified Wolf medium (D-glucose, 700 mg/L, 30 mM NaHCO3, 55% MEM, 31% sodium-free BBS, 10% FBS) (Lodowski et al., 2013). Equilibration of Wolf medium was achieved by incubating the dish containing the medium in an incubation chamber (Tokai Hit) supplied with 5%
CO2 and 95% O2 (Airgas) (Nemet et al., 2014). The glass-bottom dishes were
coated with Cell-Tak Cell and Tissue Adhesive (Corning). A circular glass
coverslip (12 mm diameter and 0.13 to 0.17 mm thickness) was used to seal and
flatten the retinas before imaging.
5.2.7 Confocal microscopy, Image analysis, quantification, and statistical
analysis
All images were acquired using a Leica TCS SP2 laser scanning
confocal/multiphoton microscope system equipped with four lasers for excitation:
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488 nm argon ion, 543 nm HeNe, 633 nm HeNe, and tunable Chameleon XR
Ti:Sapphire laser (Leica Microsystems) as described previously (Lodowski et al.,
2013). A HCX PL APO CS 40.0 X 1.25 oil UV objective lens was used for imaging. Diameters of vesicular structures were measured using ImageJ’s line function tool (Schneider et al., 2012). For the quantitative studies involving drug treatments, the same laser power and imaging condition were used for all the samples. The cytoplasmic space was highlighted manually and its mean fluorescence intensity measured using ImageJ’s freehand selection tool
(Schneider et al., 2012). The background noise was determined from blank space where no cells were observed, and subtracted from the measurements. All the quantitative data were obtained from 4 independent animals (or structures thereof) are represented as mean ± SD. In comparing two populations, *p <
0.001 by Student’s t-test was considered statistically significant.
5.3 Results and Discussion
5.3.1 IS-mislocalized rhodopsin is secreted and uptaken by RPE cells.
Based on the past studies (Li et al., 1996; Concepcion and Chen, 2010;
Lodowski et al., 2013), class I mutant rhodopsin molecules are secreted as vesicles that accumulate in the interphotoreceptor space. We reproduced these findings in a Xenopus laevis model that expresses class I mutant (Q344ter) rhodopsin fused to Dendra2 in rod photoreceptor cells. Rhodopsin-laden vesicles were released into the extracellular space of retina explant culture (Figure 13A,
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Figure 13. Class I mutant rhodopsin is released in microvesicles before being engulfed by RPE cells (A) Live retina explant was imaged at 9 DPF. Vesicles (arrowheads) were frequently observed in the extracellular space surrounding rods expressing class I mutant rhodopsin, RhoQ344ter-Dend2. (B) Cross sections of retinas expressing RhoQ344ter-Dend2 (Dend2, green) 444 labeled with phalloidin-633 (Actin, red) and Hoechst 33342 dye (Nuclei, blue). Microvesicles containing RhoQ344ter-Dend2 were in close contact with the actin filaments of the RPE microvilli (arrowheads), and within RPE cytoplasmic space (indicated by double headed arrows). Microvesicles were observed both at 9 DPF (top panels) and 21 DPF (bottom panels). Large OS fragments (asterisks) also existed within the RPE cell layer (double-headed arrows). (C) Cross sections of retinas expressing Rho-Dend2-1D4 (Dend2, green) were labeled with phalloidin-633 (Actin, red) and Hoechst 33342 dye (Nuclei, blue). OS fragments (asterisks) containing Rho-Dend2-1D4 were visible in cytoplasmic space (double-headed arrows) of the RPE cells. Retinas were imaged either at 9 DPF (top panels) or at 21 DPF (bottom panels). (D) Size distribution of green fluorescent structures/vesicles found in the RPE of animals expressing RhoQ344ter-Dend2 (Q344ter, based on 180 structures from n = 4 animals) or Rho-Dend2-1D4 (Wild type, based on 180 structures from n = 4 animals) at 21 DPF. Scale bars = 10 μm and Zoom =1 μm.
72
arrowheads). As apical extensions of RPE cells are involved in phagocytic
activities in the interphotoreceptor space, we asked whether RPE cells are
capable of engulfing and clearing these vesicles. Apical microvilli and basal
plasma membrane of RPE cells are densely coordinated with F-actins (Bonilha et al., 1999) which were labeled with fluorescently-conjugated phalloidin (Figure
13B and C, Actin). Cytoplasmic portions of the RPE cells were barely labeled and readily recognizable from apical and basolateral borders (Figure 13B and C, double-headed arrows). In retinas expressing RhoQ344ter-Dend2, microvesicles
associated with the RPE microvilli (Figure 13B, Zoom, arrowheads), indicating
these vesicles were in the process of being engulfed. We obtained two lines of
evidence that a significant fraction of microvesicles released by RhoQ344ter-
Dend2-positive rods are engulfed by RPE cells. First, approximately 40% of all
vesicular structures in the cytoplasm of the RPE (Figure 13D, Q344ter) had
diameters within the 95% confidence interval (0.34 – 0.38 µm) of the size
distribution demonstrated by microvesicles (Figure 13A). Second, because
RhoQ344ter-Dend2 is synthesized in rods under the regulation of the Xenopus
rhodopsin promoter, these RhoQ344ter-Dend2-containing microvesicles within the
RPE must have originated from rod cells. Thus, under pathological conditions,
RPE cells are capable of engulfing rhodopsin-laden microvesicles that were
secreted into the interphotoreceptor space.
Under normal conditions, wild type rhodopsin is known to be shed in OS
membrane fragments which are phagocytosed by RPE cells. Consistently, wild
type rhodopsin fused to Dendra2 (Rho-Dend2-1D4) was observed in
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phagosomes within RPE cells (Figure 13C, Merge/Zoom, asterisks). Structures
containing Rho-Dend2-1D4 were significantly larger (2.89 ± 1.31 µm, n = 180
structures) than the extracellular vesicles released by rods expressing RhoQ344ter-
Dend2 in retina explant culture (Figure 13A, arrowheads, 0.36 ± 0.14 µm, n =
180 vesicles, p < 0.001). While the majority of RhoQ344ter-Dend2 mislocalizes to the IS PM of rod cells, a small fraction still reaches the OS, likely due to co- trafficking with endogenous rhodopsin (Concepcion and Chen, 2010). Thus, larger OS fragments exhibiting Dendra2 fluorescence were also observed in the
RPE cytoplasm of animals expressing RhoQ344ter-Dend2 (Figure 13B, Merge,
asterisks). Unlike the RPE cells of animals expressing RhoQ344ter-Dend2, in which
40% of vesicular structures are at the size of microvesicles, nearly all the structures in the RPE layer of animals expressing Rho-Dend2-1D4 animals were
larger than microvesicles (Figure 13D, compare Q344ter to Wild Type),
confirming that wild type rhodopsin is shed and phagocytosed as large OS
fragments but not as microvesicles.
5.3.2 RPE cells do not contribute to the degradation of NKA under normal physiological conditions
The above experiments suggested that IS PM mislocalized rhodopsin is packaged in microvesicles that are engulfed by RPE cells. We asked if a similar mechanism removes IS PM resident proteins. NKA is highly expressed on the photoreceptor IS PM (Kwok et al., 2008), and is essential for maintaining the dark current (Schneider and Kraig, 1990). Thus, we focused on the degradation of the alpha 3 subunit of NKA (NKAα) which is enriched in photoreceptor neurons
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Figure 14. NKAα is not released in extracellular vesicles under normal physiological conditions
(A) Cross sections of retinas expressing Dend2-NKAα were labeled with phalloidin-633 (Actin, Red) and Hoechst 33342 dye (Nuclei, blue). Retinas were 9 DPF (top panels) or 21 DPF (bottom panels). For each panel, Dend2-NKAα fluorescence was either imaged directly (Dend2, green) or NKAα (both endogenous and Dend2-NKAα) was visualized by immunofluorescence (+NKAα ab, green). (B) Live retina explant was imaged at 9 DPF. NKAα-containing extracellular vesicles were not observed in the vicinity of cells expressing Dend2-NKAα. Scale bars = 10 μm.
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(Wetzel et al., 1999). To emulate the above experiments visualizing class I mutant rhodopsin, the NKA alpha 3 subunit was fused to Dendra2 (Dend2-NKAα) and expressed in rod photoreceptors under the regulation of Xenopus rhodopsin promoter. To visualize the borders of RPE cells, retinas expressing Dend2-NKAα were stained with fluorescently-conjugated phalloidin (Figure 14A, Actin, red). In these retinas, Dend2-NKAα was not observed in the RPE cells (Figure 14A,
Dend2, RPE), indicating that the photoreceptors were not secreting Dend2-
NKAα. Consistently, Dend2-NKAα was not released to the extracellular space as vesicles in live retina explant culture (Figure 14B). To exclude the possibility that vesicles were present but not visible due to our limited detection sensitivities of
Dendra2, we enhanced the fluorescence signal by labeling these samples with anti-NKAα (Figure 14A, +NKAα ab). Regardless of rigorous labeling techniques, no vesicles were observed in the RPE layer. We obtained similar results with wild type (Nontransgenic) animals which did not express Dend2-NKAα. In these animals, endogenous NKAα, labeled by immunofluorescence, was not observed in microvesicles extracellularly or within RPE cells (Figure 15A, Nontransgenic), confirming RPE cells do not contribute to the degradation of endogenous NKAα under normal physiological conditions.
5.3.3 NKAα is not co-secreted along with class I mutant rhodopsin as microvesicles
In rods expressing class I mutant rhodopsin (RhoQ344ter-Dend2), NKAα is downregulated because of its co-degradation with RhoQ344ter-Dend2 by lysosomes. As our current study demonstrates engulfment of vesicles containing
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Figure 15. IS fragments containing NKAα are phagocytosed by RPE at late stage of photoreceptor degeneration (A) Cross sections of retinas that express either no transgenic protein (Nontransgenic) or class I mutant rhodopsin fused to Dendra2 (RhoQ344ter-Dend2). Retinas were 9 DPF (top two rows) or 21 DPF (bottom two rows). Sections were labeled with anti-NKAα antibody (NKAα, cyan), phalloidin-633 (Actin, red), and Hoechst 33342 dye (Nuclei, blue). Photoreceptor-derived NKAα was not shed in microvesicles either at 9 DPF or 21 DPF, whereas large vesicular structures (arrowheads) containing NKAα were occasionally seen in contact with RPE microvilli or inside RPE layer at 21 DPF. (B) Histogram indicating the size distribution of structures containing NKAα within the RPE layer. Retinas expressing RhoQ344ter-Dend2 were analyzed at 21 DPF. (C) A cross section of 21 DPF retina expressing RhoQ344ter-Dend2 and labeled with anti-NKAα (NKAα, cyan), phalloidin-633 (Actin, red) and Hoechst 33342 dye (Nuclei, blue). In this view, a large inner segment fragment containing NKAα (arrowhead) is being engulfed by an RPE cell. Scale bars = 10 μm. 77
RhoQ344ter-Dend2 by RPE cells, we asked whether NKAα was also co-degraded
by this route. As described in Chapter 3, rods expressing RhoQ344ter-Dend2,
actively co-degrade NKAα proteins around 9 days DPF. At this early stage (9
DPF), RhoQ344ter-Dend2 is actively released as vesicles from rods (Figure 13A)
and engulfed by RPE cells (Figure 13B). We employed anti-NKAα antibody to
label endogenous NKAα in retinas expressing RhoQ344ter-Dend2 at 9 and 21 DPF
(Figure 15A, RhoQ344ter-Dend2, NKAα, cyan). NKAα was not observed in
RhoQ344ter-Dend2-positive vesicles inside the RPE cells at 9 DPF (Figure 15A,
RhoQ344ter-Dend2, RPE). Thus, NKAα is not secreted together with RhoQ344ter-
Dend2 as microvesicles. In later stage (21 DPF) animals expressing RhoQ344ter-
Dend2 (Figure 15A, 21 DPF, RhoQ344ter-Dend2), photoreceptor degeneration advances (Lodowski et al., 2013). Likely because of rod degeneration, we observed fewer vesicles (*p < 0.001) in the retinas of older (21 DPF) animals
(16.7 ± 3.3 vesicles per 100 µm retina) than in younger (9 DPF) animals (33.9 ±
5.6 vesicles per 100 µm retina length). Coincidentally to the degenerative events,
NKAα proteins originating from IS PM were occasionally observed in structures in contact with RPE microvilli or within the cytoplasm of the RPE (Figure 15A, 21
DPF, arrowheads). These structures frequently contained RhoQ344ter-Dend2, indicating they originated from rods experiencing rhodopsin mislocalization. The size of these structures (1.33 ± 0.94 µm, based on 138 structures, Figure 15B)
was significantly (*p < 0.001) larger than that of the extracellular microvesicles
containing RhoQ344ter-Dend2 (0.36 ± 0.14 µm) in retina explant culture (Figure
13A). Occasionally, large NKAα-containing fragments comparable in size to a rod
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IS were seen in the process of being engulfed by RPE cells (Figure 15C,
arrowhead). Thus, the NKAα-containing structures observed at 21 DPF are likely cellular debris from ISs which degenerated as a result of RhoQ344ter-Dend2
expression. In retinas not expressing RhoQ344ter-Dend2, we did not observe
photoreceptor-derived NKAα in RPE cells either at 9 DPF or 21 DPF (Figure 14A,
9 DPF and 21 DPF). In summary, RPE cells are capable of engulfing cellular
debris containing IS PM protein NKAα as a result of photoreceptor degeneration,
but NKAα itself is not co-secreted together with class I mutant rhodopsin as
microvesicles.
5.3.4 NKAα is potentially degraded within rod photoreceptors
IS PM protein NKAα is not degraded via vesicle release and absorption by the
RPE cells under normal physiological conditions. Therefore, we explored if
intracellular mechanisms contribute to the degradation of NKAα. Two potential
contributors are lysosomes and proteasomes. To test the in vivo degradation of
NKAα, we treated animals expressing Dend2-NKAα with a lysosome inhibitor
bafilomycin A1 (BA1) and/or a proteasome inhibitor bortezomib (Bort) for various
lengths of time (Figure 16A). As controls, animals were treated only with the
vehicle (DMSO) for 24 hr (Figure 16A, Control). BA1 (100 nM) did not
significantly increase the amount of intracellular Dend2-NKAα (Figure 16B, p >
0.5 for 24 – 72 h BA1-treated vs Control), suggesting lysosomes do not
contribute to the NKAα degradation. Bort (100 nM) increased the amount of
Dend2-NKAα in the IS cytoplasm significantly after 24 (37.0% increase) and 72
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Figure 16. NKAα proteins are degraded intracellularly in rod photoreceptors (A) Representative images of rod cells from animals expressing Dend2-NKAα and treated with either the vehicle (DMSO, Control), lysosome inhibitor (100 nM Bafilomycin A1, BA1), proteasome inhibitor (100 nM Bortezomib, Bort), a combination of both the inhibitors (Combo), or Marizomib (Mariz) for 24 – 72 h. Scale bar = 5 μm. (B) Quantification of the cytoplasmic Dend2-NKAα fluorescence in ISs for each condition in (A). Mean fluorescence intensities are indicated on y-axis (arbitrary unit, a.u.). Asterisks indicate p < 0.001 by Student’s t-test. At least 92 rod cells (n = 92) from 4 independent animals were subjected to quantification for each condition.
80
(40.6% increase) h of treatment (Figure 16B, p-values: <0.001 for 24 h, <0.001 for 72 h). Similar accumulations were observed when animals received combined treatment of BA1 and Bort for 24 (58.7% increase) or 72 (61.6% increase) h
(Figure 16B, p-values: <0.001 for 24 h and 72 h). To independently confirm that the inhibition of the proteasome led to the intracellular accumulations, we treated animals expressing Dend2-NKAα with marizomib (250 nM, Mariz), which is a potent inhibitor of all three proteasomal activities of the 20S subunit (Ma and
Diao, 2015). The amount of Dend2-NKAα within the IS cytoplasm increased significantly after 24 (46.4%) and 72 (49.7%) h of treatment (Figure 16B, p- values: <0.001 for 24 h, <0.001 for 72 h). Considering that proteasome inhibition resulted in the accumulation of otherwise degraded Dend2-NKAα proteins, the proteasome is likely involved in the degradation of NKAα proteins. Further studies are necessary to corroborate this finding because of the small effect of
Bort on protein accumulation and the toxic nature of these molecules which precluded long-term longitudinal studies.
5.4 Conclusions
In this Chapter, we demonstrated that microvesicles secreted by rod photoreceptors are engulfed by and incorporated into RPE cells. Those microvesicles are distinct from shed rod OS fragments phagocytosed by RPE cells. Under pathological conditions as observed for rods expressing class I mutant rhodopsin, rhodopsin-laden microvesicles are actively shed into the interphotoreceptor space (Li et al., 1996; Concepcion and Chen, 2010; Lodowski
81 et al., 2013). The sizes of the vesicles (95% confidence interval of 0.34 – 0.38
µm) in our Xenopus laevis model were comparable to the sizes of vesicles (0.1 –
0.2 µm) in murine model of rhodopsin class I mutation (Concepcion and Chen,
2010), indicating that shedding of rhodopsin-laden vesicles is a conserved mechanism to expel mislocalized rhodopsin from rod photoreceptors. The past studies indicate that mouse models of ciliopathies caused by Tulp1 and IFT88 mutations manifest rhodopsin mislocalization and accumulation of rhodopsin- laden vesicles as observed in this study (Hagstrom et al., 1999; Pazour et al.,
2002). Those proteins are considered essential for cilia-associated signaling and intraflagellar transport (Hagstrom et al., 2001; Nemet et al., 2015). In the Tulp1- deficient murine model, the number of rhodopsin-laden vesicles peaks at early stages (17-21 days old) when photoreceptors are still maturing (Hagstrom et al.,
2001). Consistently, there were more vesicles in the early stage (9 DPF) than the later stage (21 DPF) in our Xenopus model. Anatomically, the interphotoreceptor space where these vesicles are shed is accessible by RPE apical membrane extensions that are optimal for phagocytic activities. To clarify the source of the microvesicles, we fluorescently labeled the cargo of the microvesicles with
RhoQ344ter-Dend2. Because RhoQ344ter-Dend2 was specifically expressed in rods under the regulation of rhodopsin promoter, we concluded that fluorescent vesicles observed in RPE originated from rod photoreceptors. Engulfment of these vesicles is potentially neuroprotective as it both removes rhodopsin molecules which cannot productively engage in phototransduction (Alfinito and
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Townes-Anderson, 2002), prevents over-accumulation of vesicular structures in the interphotoreceptor space, and potentially allows biomolecules to be recycled.
Those microvesicles did not contain NKAα, which is the major component
of the photoreceptor IS PM. Thus removal of class I mutant rhodopsin through
this route will not result in the downregulation of NKAα. To compare this behavior
of class I mutant with another IS PM protein under the same condition, we
expressed Dend2-NKAα in rods under the Xenopus rhodopsin promoter. Despite utilizing the same expression system, we did not observe Dend2-NKAα secreted in microvesicles when we expressed Dend2-NKAα. Therefore, microvesicle secretion does not appear to be a normal route of degradation for IS PM-resident protein. Packaging of mislocalized rhodopsin into microvesicles is a unique cellular response to the expression of class I mutant rhodopsin, and there are specific sorting mechanisms which differentiate class I mutant rhodopsin and
NKA.
To study the behavior of endogenous NKAα, we employed anti-NKAα antibody which specifically labeled NKAα in the photoreceptor IS PM layer, but did not label the NKAα proteins which is localized to the apical membrane of RPE cells (Hu and Bok, 2001). Under conditions of rhodopsin mislocalization, we did not observe NKAα in the RPE layer of tadpoles at 9 DPF, when photoreceptor degeneration is not prominent and microvesicles are actively released. In later stages (21 DPF) when photoreceptor degeneration becomes apparent, we started observing structures containing NKAα around the microvilli and within the
RPE cells. We also captured a large NKA-containing IS fragment in the process
83
being engulfed by RPE cells. In line with this notion, IS components are
phagocytosed by the RPE cells during photoreceptor degeneration caused by
class II mutant rhodopsin (Sakami et al., 2019). Thus, these experiments confirm
that endogenous NKAα proteins are not secreted as microvesicles under
physiological or pathophysiological conditions. Considering microvesicles are not
removing NKAα from rods, we explored intracellular degradation pathways within
rods. In Chapter 3, I demonstrated that PSmOrange-NKAα fusion protein was not
degraded by lysosomes in rod photoreceptors in the absence of rhodopsin
mislocalization. Using pharmacological approaches in this Chapter, we found that
NKAα is likely degraded by proteasomes, but not by lysosomes, within rods.
Collectively, these studies demonstrate that NKAα and class I mutant rhodopsin are eliminated from rods by distinct degradation pathways.
Chapter 6: Conclusions and Future Directions
The major goals of my thesis projects have been to understand the homeostasis of rhodopsin molecules and to study the mechanisms underlying dysfunction and degeneration of rod photoreceptor cells in an animal model of inherited retinal ciliopathies caused by rhodopsin mislocalization. In Chapters 2-
4, I demonstrated that mislocalized rhodopsin is actively degraded by lysosomes.
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It disrupts the PM homeostasis of rod photoreceptors through activating lysosome-mediated degradation of IS PM membrane protein, NKA. This led to the downregulation of NKA on the IS PM and photoreceptor dysfunction/degeneration. In addition to this intracellular removal mechanism, I described in Chapter 5 that mislocalized rhodopsin molecules are packaged into microvesicles which then are released and engulfed by RPE. This project revealed a novel pathogenic mechanism applicable to various forms of blinding disorders caused by rhodopsin mislocalization while providing insights into photoreceptor-RPE cell interactions in inherited blinding disorders.
In addition to the process described in Chapters 2 – 4, lysosome-mediated degradation is associated with various photoreceptor degenerative disorders
(Chen et al., 2013; Bogea et al., 2015; Yao et al., 2018), but the context and consequences of the degradation differ among these diseases. For example, in a light-dependent photoreceptor degeneration model, activation of lysosomes and autophagy is neuroprotective (Chen et al., 2013), but in a mouse model of a class II rhodopsin gene mutation (Sakami et al., 2011; Sakami et al., 2014), activation exacerbates the photoreceptor degeneration (Yao et al., 2018). In the case of retinal diseases caused by mislocalized rhodopsin, our study suggests lysosome-mediated degradation is both beneficial and detrimental (Figure 17).
On one hand, lysosomes are utilized to remove and eliminate toxic rhodopsin species mislocalized on the PM, but on the other hand, lysosomes cause collateral damage by co-eliminating NKAα. Other than NKAα, other IS membrane proteins, which are physiologically relevant and essential, might also be
85
degraded together with mislocalized rhodopsin and therefore downregulated on
the IS PM (Figure 17). Thus, in relation to rhodopsin mislocalization, additional
global consequences of rhodopsin degradation to the photoreceptor proteome
should be characterized in the future to fully comprehend the mechanisms of
retinal ciliopathies.
Chapter 5 described a novel process of microvesicle secretion promoted
by rhodopsin mislocalization. Recent studies indicate that microvesicles are
means of intercellular communication (Lee and Kim, 2017; Vidal-Gil et al., 2019).
Contents of microvesicles, such as microRNA and signaling proteins, mediate phenotypic changes of the recipient cells (Schorey and Harding, 2016; Roballo et al., 2019). RPE cells undergo significant morphological (Jones et al., 2016) and metabolic changes (Wang et al., 2016) in diseased retinas. In this study, we tracked microvesicles by fluorescently labeling them. The future scope of this project will be to elucidate the functions of these vesicles in relation to neuronal degeneration or protection. Our improved labeling technique would be useful for
identifying and purifying those photoreceptor-derived vesicles, which then would
allow analysis of their components and further our understanding of intercellular
communication in the pathological retina.
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Figure 17. Destiny of mislocalized rhodopsin and Na+/K+-ATPase. Class I mutant rhodopsin (green) is mislocalized to the inner segment (IS) plasma membrane (PM) of rod photoreceptors. Mislocalized rhodopsin is internalized to the IS, bringing native IS PM protein Na+/K+-ATPase with it to intracellular lysosomes for degradation. Simultaneously, a distinct mechanism releases class I mutant rhodopsin as vesicles which are taken up by neighboring RPE cells.
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Because of highly heterogeneous nature of retinitis pigmentosa, investigators attempted to develop a generally-applicable strategy for preventing photoreceptor deneneration. Those are collectively called neuroprotection which was also applied to animal models harboring class I rhodopsin mutation
(Athanasiou et al., 2018). In these approaches retinas are supplemented with neurotrophic factors whose major advantage is that they act on neuronal survival instead of targeting a disease causative gene products (Athanasiou et al., 2018).
For example, overexpression of brain-derived neurotrophic factor (BDNF) resulted in a delay in the onset of photoreceptor cell death and an improved electroretinogram (ERG) recordings in class I mutant rhodopsin mice (Okoye et al., 2003). In class I model of RP, rods degenerate first and cones becomes dormant by losing their OS structures. Supplementation with human ciliary neurotrophic factor (CNTF) has been demonstrated to regenerate the OSs of cones, improving the visual function of S344ter rat model (Li et al., 2010). In a similar set of studies, glial-derived neurotropic factor (GDNF) was overexpressed in the retinas of class I mutant rhodopsin rats, which resulted in sustained functional rescue (Dalkara et al., 2011). Therefore, overexpression of neurotrophic factors demonstrated partial success in preserving vision of animals with class I mutant rhodopsin. Nevertheless, in these approaches, the root causes of the diseases, mutant rhodopsin and NKA downregulation, is not directly targeted, and the mechanisms of preventing photoreceptor degeneration is still not fully elucidated.
88
To circumvent the issues of non-specific neuroprotection approaches,
other approaches were proposed in the past to specifically target or reduce the
amounts of mutant rhodopsin genes. One avenue of targeting class I mutant
rhodopsin is to use gentamycin in combination with aminoglycosides which allow
read-through of stop codon resulting in the addition of the ciliary targeting signal to the translated protein (Guerin et al., 2008; Gregory-Evans et al., 2012;
Hollingsworth and Gross, 2012). Such targeting of aberrant rhodopsin mRNA has
been shown to improve the morphology of the afflicted retina (Gregory-Evans et
al., 2012), however the efficacy of read-through therapy is not sufficient to completely prevent the generation of class I mutant rhodopsin. Another approach used an AAV to introduce an shRNA that targets both mutant and wild type rhodopsin mRNAs transcribed from endogenous genome. This approach results in downregulation of all rhodopsin molecules including mutant rhodopsin which is toxic to the cells. The same AAV carries a human rhodopsin gene which is rendered resistant to the shRNA, allowing sustained expression of functional rhodopsin molecules (Cideciyan et al., 2018). This is a promising therapeutic approach which is proven successful in treating canine model of inherited retinal degeneration. The approach, however, has not been tested on models of class I rhodopsin mutants and further studies are necessary to define the efficacy on human subjects.
My study elucidated the mechanism of photoreceptor degeneration caused by rhodopsin mislocalization. Such knowledge would be harnessed to develop novel therapeutic strategies. The lysosomal pathway promotes NKAα
89 degradation during rhodopsin mislocalization due to co-trafficking of IS PM- mislocalized rhodopsin with NKAα to cytoplasmic lysosomes, serving as a common mechanism leading neuronal dysfunction and degeneration. Likewise, accelerated degradation of NKAα by lysosomes leads to its downregulation and exacerbates disease conditions such as damage of lung tissue or oxidative stress of kidney proximal tubule cells (Thevenod and Friedmann, 1999; Helenius et al., 2010). In these diseases in which rhodopsin mislocalization is prevalent, enhancing NKA levels could be a viable strategy. One possible way to increase
NKA level is lysosome inhibition by small molecules. However, this approach’s shortcoming is it would also prevent degradation of class I mutant rhodopsin, which is neurodegenerative. Another possible way to enhancing NKA levels is to deliver NKA genes via AAV; transgenic NKA expression specifically to rods would restore its presence to the IS PM. Unlike lysosome inhibition, this AAV approach would not result in preventing degradation of the mutant proteins. Such a therapeutic study would also further confirm that NKA downregulation is the cause of retinal degeneration in the long term.
90
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