Functions to Regulate Moesin Phosphorylation During

CLIC FUNCTIONS TO REGULATE MOESIN

PHOSPHORYLATION DURING DROSOPHILA

RHABDOMERE FORMATION

A Thesis

Presented to

The College of Arts and Sciences

Ohio University

In Partial Fulfillment

of the Requirements for Graduation with Honors

from the College of Arts and Sciences

with the degree of

Bachelor of Science in Molecular and Cellular Biology

Kara J. Finley

May 2015

ABSTRACT

ERM family proteins have a critical role in the formation of actin-rich structures through the process of linking the cytoskeleton to the plasma membrane. In

Drosophila melanogaster, Moesin, the sole ERM protein, is responsible for the development of the stacks of thousands of actin-based microvilli that form the rhabdomeres of photoreceptors in the compound eye. When phosphorylated, active

Moe can bind both F-actin and the phospholipid PIP2; dephosphorylation inactivates

Moe and leads to the dissociation of F-actin. The phosphoregulation of Moe is accomplished through the activities of various proteins. Moe binds to the phospholipid

PIP2, which is produced through the action of Sktl. Slik kinase then phosphorylates

Moe, thereby activating it and promoting binding of the actin cytoskeleton. This Moe- actin complex is next connected to a membrane-bound protein via the protein Sip1

(Drosophila homolog of EBP50). Dephosphorylation of Moe can then occur. The chloride intracellular channel protein Clic is implicated to have a role in this pathway.

Our study focuses on determining the function and location of Clic in this cycle. We overexpressed Slik, both individually and in combination with Sip1 or Sktl, with a

GMR driver and compared these in the wild type and a Clic deficient background by means of external morphology and internal eye morphology using a transmission electron microscope. Excess Slik kinase or Sktl, and the combination of both increases the phosphorylation of Moe, leading to larger rhabdomere formations; loss of Clic enhances these effects. Clic deficiency appears to result in a slight rescue of the phenotype of offspring with excess Slik kinase and Sip1, leading to a more regular

rhabdomere organization. Our results suggest that Clic functions at multiple points in

Moe phosphoregulation.

ACKNOWLEDGMENTS

First and foremost, I would like to thank Dr. Tanda of the Ohio University

Department of Biological Sciences for being a great instructor and mentor. It was through his mentoring and patience that I was able to pursue an honors thesis. I am grateful for everything he has taught me in the lab, as well as for his advice about my plans for the future. My time at Ohio University would not have been the same without him.

Furthermore, I would like to thank Dr. Berryman of the Ohio University

Department of Biomedical Sciences for his help and advice, as well as for his previous research on this topic. I would also like to thank Dr. Robert Hikida of the Ohio

University Department of Biomedical Sciences for his contributions to this project, including embedding, staining, and sectioning of tissue samples, along with his instruction on how to use the Ultramicrotome. His expertise in the use of transmission electron microscopy contributed to the informative electron micrographs used in my thesis.

This work was supported by the Provost’s Undergraduate Research Fund

(PURF) at Ohio University. Additionally, I would like to thank Dr. John Kopchick for his donations to form the John J. Kopchick Molecular and Cellular

Biology/Translational Biomedical Sciences Undergraduate Student Support Fund, from which I received funding for this project.

TABLE OF CONTENTS

Introduction...... 7

I. Drosophila melanogaster photoreceptor differentiation...... 8

II. Rhabdomere formation...... 10

III. ERM proteins...... 13

IV. Moesin regulation...... 14

V. Chloride intracellular channels...... 15

VI. Chloride intracellular channels and Moesin...... 17

VII. Rho signaling...... 18

VIII. Rho, ERM proteins, and Moe...... 18

Experimental Design...... 21

I. Use of Drosophila melanogaster as a model organism...... 21

II. Research questions and hypotheses...... 23

Materials and Methods...... 24

I. Fly strains and genetics...... 24

II. Maintenance of fly cultures...... 26

III. External eye imaging...... 27

IV. Fixation, staining, and embedding...... 27

V. Sectioning...... 28

VI. Western blot...... 29

VII. Immunofluorescence...... 31

Results...... 34

I. Clic is involved in rhabdomere morphogenesis...... 34

II. Clic functions antagonistically to Moe phosphorylation...... 37

III. Clic functions synergistically to Moe phosphorylation...... 43

Discussion...... 47

Future Directions...... 52

Literature Cited...... 55

INDEX OF FIGURES

Figure 1. Schematic drawing of the Drosophila adult ommatidium...... 12

Figure 2. EM picture of normal ommatidium...... 12

Figure 3. Current model for Moesin activation...... 15

Figure 4. Stereocilia in jitterbug mice...... 16

Figure 5. Western blot of larval salivary glands...... 17

Figure 6. Model of ERM, Clic, and Rho complex...... 19

Figure 7. RNAi gene knockdown schematic...... 26

Figure 8. EM of WT vs. Clic109 ommatidia...... 34

Figure 9. EM of GMR control ommatidium...... 35

Figure 10. EM of MoeRNAi WT vs. MoeRNAi Clic109 ommatidia...... 36

Figure 11. Co-localization assay in pupal retinas...... 37

Figure 12. LM of UAS-Sktl WT vs. UAS-Sktl Clic109 eye tissue sections...... 38

Figure 13. EM of UAS-Slik WT vs. UAS-Slik Clic109 ommatidia...... 39

Figure 14. EM of UAS-Sktl + UAS-Slik in WT vs. Clic109ommatidia...... 40

Figure 15. EM of UAS-Slik + UAS-MoeWT.myc in WT vs. Clic109 ommatidia...... 41

Figure 16. Western blot of adult head tissue...... 43

Figure 17. EM of UAS-Slik + UAS-Sip1 in WT vs. Clic109 ommatidia...... 45

Figure 18. LM of UAS-Sip1 in WT vs. Clic109 adult eye exterior...... 45

Figure 19. EM of UAS-Pp1-87B (x2) in WT vs. Clic109 ommatidia...... 46

Figure 20. Model of Clic activity in Moe phosphorylation...... 51

Figure 21. Mosaic analysis with a repressible marker schematic...... 53

INTRODUCTION

Hearing loss affects up to 360 million people worldwide (Atkinson et al.,

2014). When exposed to loud noise, damage occurs to the stereocilia in the ears, which can lead to cell death of sensory cells. Often, this occurs over a long period of exposure to noise and is typically found in the elderly; however, hearing loss is becoming more prevalent in the younger generations due to damaging sounds, such as loud music, construction noise, and gunfire. According to the National Institute on

Deafness and Other Communication Disorders, of those affected by hearing loss in

America, about 26 million cases are caused by exposure to harmful noises. Common treatments include cochlear implants and hearing aids; however, hearing aid production currently only covers 10% of the global need (Disorders, 2010). Since there is a drastic gap between supply and demand, alternative methods for treating deafness must be studied.

An understanding of the genetic and protein pathways responsible for the development and functioning of the stereocilia, which are similar structures to microvilli, is crucial to developing new methods of treatment. The proteins responsible for proper formation of stereocilia in humans have been studied recently in mice

(Salles et al., 2014). These proteins include the ERM proteins, or Ezrin, Radixin, and

Moesin. ERM proteins function to anchor the actin cytoskeleton to the membrane, producing the microvilli associated with sensory cells (McClatchey, 2014). Studying these protein pathways and their interactions in mice can be difficult; however, using other organisms to screen for these interactions can be easily accomplished.

Fortunately, Drosophila melanogaster, the common fruit fly, has only one

ERM protein: Moesin (Moe), and numerous genetic tools in which to study it. One important function of Moe in Drosophila is to form the microvilli stacks that make up the photosensitive structures, or rhabdomeres, in the photoreceptors of the compound eye (Polesello and Payre, 2004). Therefore, rhabdomere formation, size, and shape can be used to study the regulation and function of Moe. The compound eye of Drosophila acts as a more simplistic model with which to study ERM protein involvement in the formation of microvilli structures related to human and mouse stereocilia.

I. Drosophila melanogaster photoreceptor differentiation

The compound eye of Drosophila melanogaster is composed of 750 cone- shaped units termed ommatidia (Hsiao et al., 2012). Each ommatidium contains a trapezoidal pattern of eight photoreceptive cells, four cone cells, three mechanosensory bristles, two primary pigment cells, six secondary pigment cells, and three tertiary pigment cells (Pichaud, 2014). The photoreceptive cells are comprised of light-sensitive membrane stacks dubbed rhabdomeres (R1-R8) (Wolff and Ready,

1993).

Differentiation of the photoreceptor cells begins during the imaginal disc stage of development with the expression of the Atonal (Ato) transcription factor, which initiates a cluster of three presumptive photoreceptors to form (Dokucu et al., 1996).

One of these cells will differentiate into the presumptive R8 cell and the other two will form the presumptive R2 and R5. The formation of R2 and R5, rather than the formation of three R8 cells, is accomplished through the inhibition activities of both

rough (ro), a DNA binding protein, and the transcription factor senseless (sens), a zinc finger transcription factor that can activate ato (Kumar, 2012). In the presumptive R2 and R5 cells, sens expression is absent, leaving ro expression active, which in turn suppresses sens and leads to adoption of the R2/5 fate (Frankfort et al., 2001). In the presumptive R8 cell, sens is actively expressed and inhibits ro activity, allowing for ato activation and adoption of the R8 fate (Kumar, 2012).

For R3/4 specification, the ligand Spitz is first secreted by the R8 cell, leading to EGF receptor signaling in the R2/5 cells (Freeman, 1994). EGFR signaling from the

R2/5 cells to the R3/4 cells leads to activation of the transcription factor Pointed and phosphorylation by MAP kinase and subsequent degradation of the transcription factor

Yan, which is responsible for inhibiting photoreceptor formation in the neighboring undifferentiated cells (Kumar, 2012). R3 and R4 are determined through different pathways, namely the Wingless and Notch pathways. According to Domingos et al., these cells must adopt the correct fate to produce proper ommatidial chirality and planar polarity in the retina. To ensure this, frizzled is activated in R3, which in turn activates the expression of Delta. The Delta ligand then activates the Notch receptor in

R4, leading to differential photoreceptor adoptions (2004). R1 and R6 are produced through expression of the Bar locus, which encodes two transcription factors, during the second mitotic wave of the imaginal disc (Kumar, 2012).

R7 is the last photoreceptor to be specified. The R7 fate is induced by Delta production in R1/6, which activates the Notch pathway in the presumptive R7 cell, leading to expression of the membrane-associated receptor tyrosine kinase Sevenless

(Kumar, 2012). According to Reinke and Zipursky, Sevenless physically interacts with the integral protein Bride of Sevenless found on the R8 cell. This contact instructs the presumptive R7 cell to assume the R7 fate (1988). Interestingly, the R3/4 cells contain

Sevenless as well, but the R7 fate is prevented through the actions of Ro and Seven-up

(Svp), which may be activated through EGF receptor signaling (Kumar, 2012).

Additionally, the spalt (sal) genes are activated in R3 and R4, leading to downstream expression of seven-up and the maintenance of the R3/R4 identity (Domingos et al.,

2004).

II. Rhabdomere formation

During the pupal stage of development, a 90° shift in the apical pole orients the apical base of the future photoreceptive cells to align and face the center of the ommatidium

(Knust, 2007). The apical membrane is then compartmentalized into the central and subapical membranes: the central membrane will form the compact rhabdomere structure, while the subapical membrane forms the stalk membrane that will connect the rhabdomere to the zonula adherens (Knust, 2007). Apical-basal polarity proteins, such as the Crumbs complex and the Par complex, accumulate in the apical domain

(Hwa and Clandinin, 2012). The apical plasma membrane begins to form irregular folding structures around 35% pupal development (Ready, 2002). Increased membrane traffic to the apical membrane expands the membrane, which is then folded by the cytoskeleton into the microvilli-rich stacks (Karagiosis and Ready, 2004). The expanded membrane extends to the retinal floor between 35 and 50% pupal development (Ready, 2002). During formation, the rhabdomeres begin to separate

from one another, forming a central lumen, the inter-rhabdomeral space, into which the eight rhabdomeres project (Knust, 2007). The stalk of the rhabdomere begins to stiffen due the formation of a cytocortical membrane scaffold formed by proteins, such as Stardust, Crumbs, and beta-Heavy Spectrin (Ready, 2002). At 55% pupal development, the microvilli undergo elongation and packing, forming tightly associated bundles. Individual microvilli may be bound together through the action of

Chaoptin, the mutant of which is known to result in separated microvilli (Ready,

2002). The mature photoreceptor’s rhabdomere contains approximately 60,000 microvilli, making up approximately 90% of the plasma membrane of the cell (Wolff and Ready, 1993). The microvilli are oriented in differing directions, known as rhabdomere twisting, in order to enhance the absorption of both polarized and unpolarized light (Baumann and Lutz, 2006). After terminal differentiation, the rhabdomeres R1-R6 comprise the peripheral regions of the ommatidium, while R7 and

R8 are located in the center (Dickson and Hafen, 1993). A schematic representation of one ommatidium is represented (Fig. 1). R7 and R8 have smaller cross sections than

R1-R6 and are stacked vertically one on top of the other, so only seven rhabdomeres are visible in any horizontal plane (Fig. 2) (Kumar, 2012).

Fig. 1. Longitudinal and various horizontal cross- sections of the adult ommatidium. The locations of R1-R8 are indicated. As shown, rhabdomeres R1-R6 extend the entire longitudinal length of the ommatidium, while R7 and R8 are stacked upon one another. From Wolff and Ready (1993).

Fig. 2. Normal ommatidium with seven rhabdomeres. The traditional trapezoidal shape can be seen. R1-R7 are marked, as well as the stalk (S), the inter-rhabdomeral space (IRS), and the adherens junctions (AJ). From Karagiosis and Ready (2004).

III. ERM proteins

Rhabdomeres are formed from interactions between ERM proteins and the actin cytoskeleton. The ERM family is highly conserved amongst species and is composed of Ezrin, Radixin, and Moesin (Polesello and Payre, 2004). ERM proteins are members of the band 4.1 superfamily, meaning they share a common string of approximately 300 amino acids composing the FERM (Four point one Ezrin, Radixin,

Moesin) domain (Fievet et al., 2007). ERM proteins contain two distinct regions: the amino terminal and carboxy terminal regions. The N-terminal region (N-ERMAD), which is comprised of the FERM domain, can interact with various integral membrane proteins. These integral proteins include CD43, CD44, and ICAM-1, -2, and -3

(Yonemura et al., 2002). The FERM domain also binds the PDZ domain of EBP50, the orthologue of which is Sip1 in Drosophila (McClatchey, 2014). The C-terminal region (C-ERMAD) binds to F-actin of the cytoskeleton. Normally, the C-ERMAD interacts with the FERM domain, preventing cytoskeletal interactions (Roubinet et al.,

2011). In order for the C-ERMAD to interact with F-actin, the ERM protein must be activated to the open conformation, breaking the intramolecular attraction between the

C-ERMAD and the N-ERMAD (Roch et al., 2010). Activation of ERM proteins requires FERM domain binding of phosphatidyl-inositol (4,5)-bisphosphate (PIP2) along with phosphorylation of a threonine residue located near the C terminus

(Polesello and Payre, 2004). First, the FERM domain is recruited to regions of the membrane rich in PIP2, where binding occurs. Once this binding occurs, the regulatory threonine residue, Thr558 in Moe, is more available for phosphorylation, typically by

kinases (Fehon et al., 2010). Once activated, the ERM protein can bind to the cytoskeleton, anchoring microfilaments to the membrane. Moe is the sole ERM protein in Drosophila (Roubinet et al., 2011), making Drosophila a useful model organism for studying ERM protein interactions without fear of compensatory interactions. During rhabdomere formation, phosphorylated (active) Moe accumulates at the base of the stalk and binds to the rhabdomere terminal web (RTW) to secure the cytoskeleton (Ready, 2002).

IV. Moesin regulation

The activation of Moe requires two steps: binding of Moe to PIP2, and subsequent phosphorylation (Fig. 3). Inactivated, dephosphorylated Moe binds to the membrane-bound phospholipid PIP2. PIP2 is synthesized by type I phosphatidylinositol kinases (PIP5KI) , also known as Skittles (Sktl) in Drosophila

(Roch et al., 2010). Threonine residue phosphorylation, and thereby activation, occurs through the action of the Sterile 20 family kinase Slik (Hughes et al., 2010). Once

Moe is activated, interactions with various proteins transfer the phosphorylated Moe from PIP2 to an as of yet unknown integral protein. Evidence has shown that Sip1 physically interacts with both Slik and Moe, suggesting the formation of a complex during the phosphorylation of Moe (Hughes et al., 2010). Scaffolding proteins may bind to the activated complex and cross-link Moe to additional integral membrane proteins (Jiang et al., 2014).

Dephosphorylation occurs through the action of Pp1-87B phosphosphatase

(Kunda et al., 2012). It is currently unclear whether Moe must be bound to specific

proteins for dephosphorylation or if dephosphorylation can occur in any complex once activation takes place.

Fig. 3. One current model for Moesin activation. Moe binds to PIP2, thereby causing a conformation change that increases the chances of phosphorylation and activation by Slik kinase. This model involves a Sip1-Slik kinase complex. From Hughes et al. (2010).

V. Chloride intracellular channels

Chloride intracellular channels (CLICs) were first identified by their ion channel activity, but alternative activities for CLICs have been suggested (Littler et al.,

2008). CLIC proteins can be found as both soluble globular proteins and insoluble integral membrane proteins (Littler et al., 2010). The structure of the protein contains a ~240 residue CLIC module belonging to the glutathione S-transferase (GST) fold superfamily (Littler et al., 2008). A conserved cysteine residue found on the Clic protein structurally resembles an enzymatic active site (Jiang et al., 2014).

Numerous studies have found CLIC functions as a poorly selective ion channel, possibly requiring a high pH for activation. In addition, CLIC has been implicated to interact with the actin cytoskeleton and ERM proteins (Littler et al.,

2010). Previous studies of CLIC5 found that CLIC is required for the proper formation and maintenance of stereocilia bundles involved in hearing in mice. Using CLIC5- deficient jitterbug (jbg) mice, Salles et al. demonstrated that the localization of

Taparin, protein tyrosine phosphatase receptor Q (PTPRQ), and Radixin to the base of the stereocilia is affected by loss of CLIC5, suggesting the formation of a structural complex between these proteins. In addition, jbg/jbg mice demonstrate degenerate stereocilia with fusing (Fig. 4) between individual bundles, leading to loss of hearing and balance. The interaction between CLIC5 and the actin cytoskeleton is further supported by the loss of Taparin, PTPRQ, or Radixin, which leads to stereocilial defects similar to jbg mice, as well as deafness (2014).

While there are six vertebrate CLIC proteins, Drosophila only contains one,

Clic (Jiang et al., 2014). With Moesin as the only ERM protein and Clic as the only

Clic protein, Drosophila melanogaster is an ideal model organism to study the possible activity of Clic to interact with ERM proteins.

Fig. 4. Formation of stereocilia in heterozygous and homozygous jitterbug mice. jbg/+ (E) demonstrate normal stereocilia development while jbg/jbg (F) show fused stereocilia. From Salles et al. (2014).

VI. Chloride intracellular channels (CLICs) and Moesin

In addition to CLIC studies in mice, unpublished data produced by Dr. Soichi

Tanda, Dr. Mark Berryman, and Regan Price of the Ohio University Department of

Biological Sciences suggests that Moe and Clic interact in Drosophila. The first experiment was accomplished using extracts from 3rd instar larval salivary glands. A western blot was performed to detect phosphorylated Moe levels in each of two groups: the wild-type control group and a loss-of-function Clic109 group. The results

(Fig. 5) demonstrate that levels of phosphorylated Moe are much higher in the Clic109 background than the wild-type background. This suggests Clic functions to decrease the phosphorylation of Moe in the salivary glands, either through inhibition of phosphorylation or increase in phosphatase activity.

Fig. 5. 3rd instar larva salivary gland extracts labelled for Clic and phosphorylated Moesin. The Clic109 mutant shows drastically increased levels of phosphorylated Moesin compared to the wild- type. The absence of Clic in the Clic109 mutant demonstrates that the Clic109 allele is an effective protein null allele.

The second experiment conducted involved co-immunoprecipitation of Moe and Clic, again in 3rd instar larval salivary gland extracts. A Clic-GFP fusion construct was produced and expressed in the salivary glands before extraction. The Clic-GFP proteins were precipitated using rabbit anti-GFP antibodies. Moe was pulled down during this experiment, demonstrating that Clic and Moe physically interact. The

nature of this interaction is as of yet unclear and could be due to direct contact or indirect contact through the binding of other proteins to both Clic and Moesin.

VII. Rho Signaling

Rho proteins are small GTPases involved in many cellular processes, including the cell cycle, membrane transport, cytoskeletal dynamics, and cell polarity

(Buchsbaum, 2007). Active, GTP-bound Rho enhances signals, often from cell surface receptors, to downstream effectors (Schwartz, 2004). Activation of Rho occurs through guanine-nucleotide-exchange factors (GEFs), while GTPase-activating proteins (GAPs) deactivate Rho’s GTPase ability by enhancing GTP hydrolysis to the inactive GDP form (Buchsbaum, 2007).

Many Rho-family GEFs contain PH (pleckstrin homology) and DH (Dbl homology) domains involved in various functions (Buchsbaum, 2007). These functions include membrane targeting, control of exchange activity, and regulation of phospholipid and protein interactions. The PH domain is also involved in protein translocation, leading to localized Rho activation (Buchsbaum, 2007).

VIII. Rho, ERM proteins, and Moesin

Previous studies demonstrate that Rho may be involved in ERM protein activation. Rho-kinase has been shown to phosphorylate and activate ERM proteins in certain types of cells (Yonemura et al., 2002). Specifically, the phosphorylation of

Moe by Rho/Rho-kinase has been found to have a critical role in forming microvilli- like structures in monkey kidney cells (Oshiro, 1998). In addition, PIP2 production is affected by Rho signaling (Hirao et al., 1996). Phosphatidylinositol-4-phosphate 5-

kinase, responsible for PIP2 formation, is activated by Rho and Rho-kinases

(Schwartz, 2004). As stated, an increase in PIP2 production increases binding of Moe, allowing for subsequent Moe phosphorylation and activation.

Recent studies demonstrate CLIC4 is also involved in the RhoA signaling pathway. RhoA activation affects the cytosolic location of CLIC4 by promoting translocation of CLIC4 to plasma membranes containing RhoA-activated G-protein coupled receptors (Ponsioen et al., 2009). In addition, RhoA accumulates in regions of the membrane containing high concentrations of ERM proteins (Hirao et al., 1996).

This data, coupled together, indicates ERM proteins attract RhoA, which in turn enhances CLIC4 translocation to the membrane location. This theory is demonstrated by Jiang et al. (2014), in which Moe, Clic, and Rho complex to form the actin cytoskeleton anchor (Fig. 6).

Fig 6. Model of ERM, Clic, and Rho complex. Speculative model in which the ERM protein binds PIP2, EBP50, Rho, and Clic, as well as F- actin. From Jiang et al. (2014).

According to Speck et al., Moesin is an additional player in Rho signaling. In

Drosophila, Moe acts to regulate cell-signaling events by working antagonistically to

Rho. Moe mutants display apical protrusions where normal microvilli-like structures should be found in epithelial cells. Reduction of Rho results in more normal microvilli structures with fewer protrusions, suggesting Moe functions to oppose Rho. These

results indicate negative regulation of Rho by ERM proteins. Rho signaling involves

ERM proteins as downstream effectors, meaning a negative feedback loop may exist between ERM proteins and Rho. This feedback loop is reminiscent of the relationship between GTPase Rac and the ERM-related tumor suppressor Merlin (2003).

Considering Moe and Clic may be involved in the Rho signaling pathway, additional measures must be taken to distinguish Rho effects on Moe phosphorylation levels. This control was accomplished in two ways: UAS elements and Moe tagged with myc. Using the UAS/GAL4 system abolishes any feedback mechanisms involved in transcriptional regulation. While this does not exclude inhibitory protein interactions, the UAS genes will not be affected by inhibition at a genetic level due to the unnatural promotor construct. For western blot purposes, Moe.myc constructs were used to distinguish between Rho-affected Moe levels and Rho-independent Moe. This additional level of control ensures the resulting Moe phosphorylation levels are affected only by Moe phosphorylation pathway proteins rather than extraneous influences.

EXPERIMENTAL DESIGN

I. Use of Drosophila melanogaster as a model organism

In the last couple of decades, Drosophila melanogaster has become known as an appropriate and easy-manipulated model organism. Not only can Drosophila be used to determine mechanistic properties, such as protein-protein interactions and signaling pathways, but it can also be used to study many human diseases. It has been determined that approximately 75% of all human disease genes have conserved or similar sequences found in the Drosophila genome, corresponding to about 700 disease genes that can be efficiently studied in the organism, including certain developmental, cardiovascular, and metabolic diseases (Bier, 2005). In addition, cancer formation and cancer drug screening can be studied using flies. Most important to this experiment, the genes required for the auditory and visual systems are relatively conserved (Bier, 2005).

In addition to homologous genes, Drosophila can be easily genetically manipulated during experiments. The GAL4/UAS system and RNA interference

(RNAi) are established in flies and used for a variety of studies. Genetic manipulations in mice, for example, are much more complicated and take a larger amount of time and money to accomplish. In flies, the relatively short life cycle and easy breeding makes changes to the genome easy, fast, and affordable (Bier, 2005). The

Bloomington Drosophila Stock Center at Indiana University provides a wide variety of fly lines to choose from in order to form experimental crosses.

Drosophila also contain fewer redundant genes in their genome than mammalian systems. The usefulness of Drosophila can be appreciated in this regard when conducting protein interaction assays, as well as knockout studies. The human

CLIC5 was first established to interact with the actin cytoskeleton, including the ERM protein Ezrin, in human placental microvilli by Berryman and Bretscher (2000).

Subsequent studies determined that the CLIC5 mouse homologue co-localizes with

Radixin in the base of stereocilia (Salles et al., 2014). In order to fully understand the interactions between CLICs and the ERM proteins, our study uses Drosophila as a model organism. In our experiment, the focus is on the proteins Clic and Moesin.

Moesin is the only ERM protein in Drosophila, meaning there are no extraneous effects due to Ezrin and Radixin interference. Additionally, Clic is the only CLIC protein, unlike the six CLIC genes found in mammalian systems (Littler et al., 2008), making the loss-of-function Clic109 fly line completely devoid of any Clic function and preventing cooperative effects of similar proteins.

Since previous studies of CLIC-ERM interactions have focused on actin-rich structures (microvilli and stereocilia), the Drosophila photoreceptor is an excellent structure to focus on. The photosensitive organelle of the photoreceptor is the rhabdomere, composed of 60,000 microvilli in part formed through Moesin-F-actin binding (Wolff and Ready, 1993). Using knowledge of rhabdomere formation and structure, the interactions between Drosophila Clic and the ERM protein Moesin can be identified.

II. Research questions and hypotheses

The intent of this research is to understand the interactions between Clic and

Moesin in Drosophila rhabdomere formation. To accomplish this, rhabdomere formation, size, and shape in varying genetic backgrounds, including those in a Clic109 mutant, was analyzed. Transmission electron microscope images of the cross-sections of ommatidia provided qualitative data for analysis of each cross. Conclusions are based on the fundamental idea that increases in phosphorylated Moe levels cause an increase in rhabdomere width due to excessive microvilli formation; on the other hand, a decrease in phosphorylated Moe levels leads to a decrease in rhabdomere width.

We hypothesize that Clic directly interacts with at least one of the proteins involved in the phosphorylation or dephosphorylation of Moe, including Moe itself.

Two possible roles for Clic exist: inhibition of phosphorylation or enhancement of phosphorylation. If Clic functions to inhibit phosphorylation, either through inhibition of Slik kinase or Sktl, or through an as of yet unknown influence, the rhabdomeres are expected to increase in width in the Clic109 mutants. Alternatively, if Clic functions to augment phosphorylation, such as through the inhibition of phosphatase, the rhabdomeres in Clic109 mutants are anticipated to be smaller than in the wild-type. We predict that Clic functions to inhibit Slik kinase, thereby inhibiting the phosphorylation of Moe. If Clic does act to inhibit Slik kinase, we expect to see enlarged, widened rhabdomeres in the Clic109 mutant crosses.

MATERIALS AND METHODS

I. Fly strains and genetics

Fly strains were obtained from the Bloomington Drosophila Stock Center at

Indiana University, as well as provided by Richard Fehon and Sarah Hughes. The

GAL4-UAS system was used to increase or decrease expression levels of proteins that are known to affect Moesin function in the rhabdomeres (Duffy, 2002). The GAL4 activator was driven using eyeless or GMR (glass multiple reporter) tissue-specific promoters, both of which are known to be specific to the eye tissues (Li et al., 2012).

GMR-GAL4 has been demonstrated to cause irregular ommatidial arrays in heterozygotes at 29°C and homozygotes at 25°C (Kramer and Staveley, 2003). Due to this, a GMR control at 25°C was produced and analyzed using external eye morphology and transmission electron microscope imaging of the ommatidial structures (Fig. 7). UAS elements and GAL4 drivers are not endogenous in

Drosophila, so no additional genes are affected by their use in theory. Transgenes for the protein of interest were combined with the upstream activator sequences (UAS) and can only be activated when the GAL4 driver protein binds to this sequence

(Duffy, 2002). In this way, gene expression is induced only in fly strains where the

GAL4 driver and UAS responder are present and only in the eye tissues of the strains used in these studies. Flies contained balancer chromosomes in addition to the

GAL4/UAS elements. Balancer chromosomes are necessary to prevent recombination of inserted elements, and thereby inactivation of experimental constructs. Balancer chromosomes typically contain dominant markers to ensure each subsequent fly

generation has a balancer chromosome present (Greenspan, 2004). The flies were selected for using dominant mutant markers: Curly (Cy), Stubble (Sb), Tubby (Tb), and Bar (B). Flies were crossed using standard procedures (Greenspan, 2004).

To induce gene expression, a transgene is produced through the attachment of a UAS element to the wild-type gene, under which transcription can then be driven by crossing with a GAL4-containing fly strain. This system can also be used to decrease expression levels through the use of RNA interference (RNAi). Interference is achieved through connecting a UAS site with a specifically designed DNA sequence that corresponds to the gene to be decreased. When activated, the UAS-controlled

DNA produces a double stranded RNA (dsRNA) molecule that is then targeted by the protein Dicer (Sifuentes-Romero et al., 2011). Dicer processes the dsRNA to form small interfering RNAs (siRNAs) approximately 20-26 nucleotides long (Kasai and

Kanazawa, 2012). The siRNA is separated into two single stranded RNA (ssRNA) molecules. One strand, termed the guide strand, is integrated into the RNAi-induced silencing complex (RISC) (Sifuentes-Romero et al., 2011). The ssRNA then undergoes complementary base pairing with the mRNA of the target gene, causing the mRNA to be degraded by RISC (Fig. 7). This decreases the available mRNA copies of the gene, decreasing the overall gene expression (Sifuentes-Romero et al., 2011).

Dicer can be overexpressed in these systems in order to facilitate efficient gene knockdown.

Fig. 7. RNAi gene knockdown. I: siRNA formation. (a) dsRNA is targeted by Dicer, producing (b) an siRNA molecule. (c) the siRNA molecule is separated into single stranded RNA, the guide strand is (d) incorporated into RISC, and complementary mRNA is degraded. From Sifuentes-Romero et al. (2011).

II. Maintenance of fly cultures

Fly stocks were maintained at approximately 20°C. Experiments were conducted at 19°C, 22°C, 25°C, or 28°C, depending upon the genotype. Fly food was prepared and cooked using a mixture of: 2280g Jazz mix, 120g soy powder, 360g yeast, and 12L of water. Crosses were cultured at 19°C or 22°C until about day 5, during the middle third instar larval stage at which rhabdomere formation begins, and kept at 19°C, 22°C, or moved to 25°C for enhancement of the GAL4-UAS system.

Some crosses were maintained at 19°C and 22°C due to decreased viability, caused by the genetic manipulations (Greenspan, 2004). After selection, adult flies were maintained at 19°C for about 5 days before fixation.

III. External eye imaging

Once selected for, appropriate genotypes were frozen at -20°C and imaged using a Nikon DXM-1200 camera with Nikon Act-1 software programing on a Nikon

SMZ1000 microscope. These images were then sized using Photoshop to produce 4cm x 5cm pictures. These pictures were examined for changes in external eye morphology through comparison with the wild type or control strains.

IV. Fixation, staining, and embedding

The antennae and proboscis were removed from anesthetized Drosophila and then the head was removed. Heads were fixed in a 0.1M cacodylate buffer at pH 7.4 containing 2.5% gluteraldehyde and 2.5% paraformaldehyde. All subsequent solutions were made in a 0.1M cacodylate buffer. The initial fixation was done for at least 4 hours to overnight at room temperature. The heads were then fixed for at least 6 hours in new fixative solution containing 1% tannic acid. Then the heads were rinsed for 10 minutes in cacodylate buffer and post-fixed in 2% osmium tetroxide for 15 minutes to

2 hours. Following a short rinse in cacodylate buffer, the heads were treated in the dark with 2% aqueous uranyl acetate at room temperature for 2-4 hours. The use of osmium tetroxide and uranyl acetate adds heavy metals to specific structures, producing electron density contrast for transmission electron microscopy (TEM). Last, the heads were rinsed in cacodylate buffer, dehydrated in an ethanol series, and submerged in propylene oxide for 15-30 minutes to remove remaining ethanol.

The heads were next embedded in Epon/Araldite or Spurr’s low viscosity resin. The components of the Epon/Araldite resin mixture include: Epon 812 (13.1g),

DDSA (27.7g), Araldite 502 (10g), and DMP-30 (1.5ml) or BDMA (1.75ml). The tissues were immersed in 50% propylene oxide, along with 50% resin, overnight, then in 100% resin for 2 hours at room temperature. The resin was polymerized for 48 hours following immersion at 60°C. Spurr’s low viscosity resin mixture contained

DER 736 (14ml), ERL 4221 (18ml), DMAE (0.6ml), and NSA (48ml). Following kit directions, the heads were placed through a graded series of propylene oxide and

Spurr’s resin for 5 days. The resin was then polymerized at 70°C for 8-12 hours. Both types of resin mixtures underwent thorough mixing and at least a 1 hour degassing process before use.

V. Sectioning

Embedded heads were trimmed so that one compound eye was located at the surface of the resin block. The surface of the resin was then cut until the lens of the eye was exposed. Eye tissues were cut into approximately 0.6µm thick sections using glass knives on a Reichert Ultracut E Ultramicrotome. Sections were mounted on slides and stained with 0.7% toluidine blue in 0.7% borax, then covered with coverslips using Cytoseal 60 low viscosity mounting medium (Richard-Allan

Scientific). Eye tissue sections were imaged using a Nikon Labphot-2 microscope on both 40x and 100x magnification. The images were taken using a Nikon DXM-1200 camera with Nikon Act-1 software programing. Using Photoshop software, the images were transformed into black and white and the contrast was manually or automatically enhanced for optimal rhabdomere detection.

For TEM use, tissues were cut into ultrathin sections 60-80nm thick with a

Diatom diamond knife using a Reichert Ultracut E Ultramicrotome. The tissue sections were fixed on either naked or formvar-coated grids, then stained with 2% uranyl acetate and lead citrate and inspected using a JOEL 1010 transmission electron microscope functioning at 80kV. Electron micrographs were imaged at 1,000X-

80,000X using a Gatan digital camera fixed inside the microscope.

VI. Western blot

Western blot analysis was carried out on whole adult Drosophila heads. First, flies were anesthetized using CO2 and heads were removed using forceps (Mishra and

Knust, 2013). The heads were then quickly centrifuged to the bottom of the tube and crushed to release the proteins. Double strength Laemmli sample buffer was produced containing: 160mM HCl-Tris at pH 6.8, 4% SDS, 20% glycerol, 0.012% bromophenol blue, 2% β-mercaptoethanol, 10mM EGTA, and water to make the total volume 100mL (Laemmli, 1970). The sample buffer was added to the mixture and the mixture was then boiled for either 3 or 5 minutes. A 12% mini-maxi lower gel was produced using the following reagents: 54.2% acrylamide A, 6.03% water, 0.377M

Tris at pH 8.7, 0.15% SDS, 0.05% TEMED, and 0.05% ammonium persulfate. The upper gel was mixed to 14.8% acrylamide A, 69.1% water, 0.13M Tris at pH 6.8,

0.16% SDS, 0.12% TEMED, and 0.12% ammonium persulfate. The extracted protein mixtures were loaded into the gel and the gel was run at 50 mAmp constant current.

Once the gel was finished running, a Millipore Immobilon PVDF blot membrane and six pieces of Whatman 3mm CHR Chromatography paper were cut to

the dimensions of the gel. The membrane was wet briefly in reagent-grade methanol, and then rinsed with deionized water. The gel was removed from the running apparatus, nicked in the lower left corner, and placed in a dish with deionized water.

During the sandwich-making process, the gel was poked using a syringe needle in the locations of the pre-stained molecular weight markers to retain the location of the markers after transfer. The sandwich was built in the following order from the anode carbon electrode: two pieces of Whatman paper saturated in anode buffer I, one piece of Whatman paper in anode buffer II, blotting membrane, gel, and three pieces of

Whatman paper saturated in cathode buffer. Anode buffer I (pH 10.4) contained 0.3M

Tris and 20% MeOH; anode buffer II (pH 10.4) contained 0.025M Tris and 20%

MeOH; and cathode buffer (pH 9.4) contained 0.025M Tris, 40mM amino caproic acid, and 20% MeOH. Bubbles were rolled out from the sandwich and the transfer apparatus (Integrated Separation Systems) was closed. The transfer apparatus was run at 1mA/cm2 for 1 hour (Kyhse-Anderson, 1984).

Once transfer was complete, the membrane was blocked in 5% BSA in TBST for 30 minutes. The membrane was incubated on a shaker in the primary antibody diluted in 1% BSA, TBS, and 0.1% Tween20 at pH 7.4 at 4°C overnight. The membrane was next washed with TBS and 0.01% Tween20 (TBST) three times and then three times for 5 minutes at room temperature. The secondary peroxidase- conjugated protein A was diluted in 1% TBS and TBST and the membrane was immersed and incubated for 2-3 hours at room temperature. Last, the membrane was rinsed with TBST three times quickly and washed three times at 5 minutes each.

After washing, enhanced chemiluminescence reagents (Millipore) were used to image the western blot. An aliquot of 0.7mL of reagent A and reagent B were pipetted directly onto the blot surface and mixed. This mixture was run over the surface of the membrane multiple times to ensure a reaction. The membrane was then imaged using a BioRad ChemiDoc XRS+ Imaging System.

VII. Immunofluorescence

To analyze the co-localization of Clic and phosphorylated Moesin, immunofluorescence staining was performed on retinas at 50% pupal development.

The fly strains overexpressed a Clic-GPF construct to be used for antibody detection.

White pupae were collected and incubated at 25°C for 50 hours prior to dissection

(Bainbridge and Bownes, 1981). Dissection instructions were adapted from Walther and Pichaud (2007). Dissection proceeded first by removal of the puparium surrounding the head. The pupae were then transferred to PBS solution and dissection scissors were used to cut the pupae in half, releasing the anterior portion, including the head region. Using forceps, the head region was cleared of residual tissues so that the cuticle and brain/eye tissues remained. The cuticle was then secured using the forceps while a syringe was used to shoot PBS into the cuticle, releasing the brain and eye tissues from the cuticle. The retinas, connected by the optic lobes and brain, were then transferred to fresh PBS.

Immunofluorescence staining was performed using methods adapted from

Hsiao et al. (2012). The retinas were fixed in ice-cold TCA for 30 minutes. The retinas were then transferred to G-PBS (30mM glycine-PBS). The retinas were permeabilized

in 0.5% Triton X-100/G-PBS for 30 minutes at room temperature. G-PBS was next used to wash the retinas three times for 5 minutes each. Blocking took place at 4°C overnight in G-PBS with 4% BSA. The retinas were incubated in a 10 % solution (in

4% BSA/G-PBS) of rabbit anti-P-ERM (Cell Signaling) and a 10% solution of mouse anti-GFP mAb for 2 hours at room temperature. The retinas were then rinsed three times in G-PBS and washed three times for 5 minutes. Next, the eye tissues were incubated in a 0.33% solution (in G-PBS) of donkey anti-mouse-Alexa 546 and a

0.33% solution of donkey anti-rabbit-Alexa 488 for 1 hour at room temperature. The retinas were again rinsed three times in G-PBS and incubated three times for 5 minutes. Three final rinses were performed in PBS. The final dissection then took place, releasing the retinas from the remaining tissues.

Following immunofluorescence staining, the retinas were mounted onto slides as follows. First, the tissues were placed concave down onto SuperFrost slides using a small amount of PBS. A coverslip bridge was produced to prevent squishing of the retinas by placing two 22x22 coverslips on both the right and left regions of the slide.

Excess PBS was removed using Kim wipes. Next, 60µL of Prolong Gold mountant was dropped over the retinas. A final 22x22 coverslip was placed on the mountant such that the ends of the coverslip extended onto the coverslip bridge. Finally, the medium was allowed to cure in the dark for 1 day.

The retinas were imaged on a Zeiss LSM 510 confocal microscope using Zen

2007 software. An Argon laser and HeNe laser were used to excite the green and red dyes, respectively. Each laser was set at 1 Airy unit and the lowest possible laser

power to ensure 100% pixel saturation. Other settings were based on what produced the best image of fluorescent structures. When possible, the same settings were used for both the experimental and control retina images. The images were processed by producing a maximum intensity projection. Differences between the experimental and control images were then analyzed through comparison.

RESULTS

I. Clic is involved in rhabdomere morphogenesis

In order to determine what proteins Clic might interact with, ectopic gene expression was induced in both normal and Clic-deficient backgrounds. The wild-type w1118 and Clic109 mutant adults show very similar morphology (Fig. 8A, B). The most notable difference is that the Clic109 rhabdomeres appear more rectangular than in the wild-type. In addition, to ensure the GMR driver did not drastically interfere with rhabomere formation and structure, a GMR wild-type cross at 25°C was produced to provide a control. The addition of a GMR driver appears to form more rectangular rhabdomeres but overall is unaffected (Fig. 9A). Additionally, since the majority of the crosses were raised and maintained at temperatures lower than 25°C, the effect of the GMR driver on rhabdomere morphology is expected to be minimal.

A B

Fig. 8. Electron micrographs of wild-type vs. Clic109 ommatidia. A: Cross-section of wild-type adult ommatidium. R1-R7 arrange in a trapezoidal pattern. B: Cross- section of an adult Clic109 mutant ommatium. R1-R7 remain in regular formation, but rhabdomeres appear rectangular and boxy instead of the typical rounded shape. EM pictures were provided by Dr. Hikida of OUHCOM.

A B

Fig. 9. Electron micrograph and external morphology of a GMR control raised and maintained at 25°C. A: The effect of the GMR driver on rhabdomere morphology is trivial. Rhabdomeres appear slightly box-shaped and generally larger than in the wild-type control (Fig. 8A). Provided by Dr. Hikida of OUHCOM. Scale bar: 3µm. B: External morphology of the GMR control compound eye. Scale bar: 1mm.

To demonstrate that Moe is indispensable for rhabdomere formation, Moe

RNAi was first performed in both the wild type and Clic109 background. RNAi was driven using the GMR-GAL4 driver. The resulting MoeRNAi genotype was expected to depict smaller, irregular rhabdomeres or loss of rhabdomere formation altogether.

Since RNAi is a gene knockdown, rather than a gene knockout, some Moe function is expected to remain. In the wild-type background, the RNAi completely eliminates any resemblance of rhabdomere structures (Fig. 10A). Since Moe is involved in linking the actin cytoskeleton to the plasma membrane, this result is expected when Moe is knocked down. In the Clic109 ommatidium, around four rhabdomere structures can be distinguished, but each of these remain smaller than normal and irregularly shaped

(Fig. 10B). These results indicate Clic negatively impacts rhabdomere formation, possibly through the inhibition of Moe phosphorylation.

A B

Fig. 10. Electron micrograph of MoeRNAi in wild-type vs. Clic109. A: Normal rhabdomere structure is completely abolished. B: At least four distinct rhabdomere structures (one shown with an arrow) can be seen; an increase in rhabdomere size compared to wild-type is apparent. Scale bars: 3µm. EM pictures provided by Dr. Hikida of OUHCOM.

To determine whether Clic and Moe occupy the same intracellular location, a co-localization assay was performed. Using antibody staining and confocal microscopy, Clic and phosphorylated Moe were determined to co-localize at the rhabdomere structures at 50% pupal development (Fig. 11). This co-localization suggests an interaction between the two proteins, strengthening the argument for

Clic’s involvement in the Moe phosphorylation pathway.

Fig. 11. Co-localization assay for Clic and phosphorylated Moe (p- Moe). Performed in 50% pupal development retinas. A UAS-Clic- GFP element was driven by the GMR GAL4 driver and stained using anti-GFP antibodies (green). Phosphorylated Moe was stained using anti-pMoe antibodies (red). The co-localization of p-Moe and Clic occurs at the forming rhabdomere structures (see arrow) and the cell cortex.

II. Clic functions antagonistically to Moe phosphorylation

The co-localization assay, loss-of-function Clic109 mutant, and apparent antagonistic effect on Moe phosphorylation levels suggests that the Clic protein is heavily involved in the phosphorylation of Moe in regards to rhabdomere morphogenesis. Which specific proteins are directly impacted by Clic’s function remains to be understood. In order to determine this missing piece, the gene expression of some of the notable genes involved in the Moe phosphorylation pathway was altered using the UAS/GAL4 system in both wild-type and Clic109 mutants.

Since the protein Sktl has a known function in Drosophila, the interaction between Clic and Sktl was first examined. Ectopic expression of Sktl leads to an increase in PIP2 formation and should therefore lead to an increase in Moe phosphorylation (Roch et al., 2010). Sectioning and EM results portray very irregular rhabdomere formation, including larger, broken rhabdomeres with a slight boxy shape

(Fig. 12A). The traditional trapezoidal pattern of formation is completely abolished

and rhabdomeres appear to be growing out of cellular regions other than the apical domain. These results suggest phosphorylation levels of Moe are greatly increased and regulation of Moe location in the cell is hindered. In the Clic109 mutant, the ommatidial structure appears to be greatly malformed (Fig. 12B). Many ommatidium appear to contain no rhabdomeres or fewer rhabdomeres than normal; the rhabdomeres that are present are greatly disfigured in shape. These rampant deformities make it difficult to determine the effect that the loss of Clic has on rhabdomere development.

Electron microscope images of the UAS-Sktl, Clic109 crosses are currently in production in order to produce more distinguishable data from which to draw a solid conclusion.

A B

Fig. 12. Light microscope images of UAS-Sktl wild-type vs. Clic109 ommatidia. A: Wild-type rhabdomeres appear irregular (arrow); some ommatidia are missing rhabdomeres. B: Clic-deficient rhabdomeres (arrow) are difficult to distinguish and many appear to be missing. Scale bars: 10µm.

The next experiment involved the interaction of Slik kinase. Slik is a sterile 20 family kinase responsible for phosphorylation, and therefore activation, of Moe

(Hughes et al., 2010). As expected, the UAS-Slik flies exhibited expanded, larger

rhabdomeres, confirming excessive phosphorylation of Moe (Fig. 13A). In Clic- deficient flies, the rhabdomeres appeared more enlarged; they were much wider, split in random places, and the resulting ommatidia appeared to have lost the original size and pattern (Fig. 13B). These results suggest Clic works in an opposing fashion to

Slik. In conjunction with the results obtained from the Sktl experiments, Clic likely works in opposition to the activation and phosphorylation of Moe.

A B

Fig. 13. Electron micrograph of UAS-Slik wt vs. Clic109 ommatidia. A: Wild-type rhabdomeres appear wider than normal and splitting can be seen; eight rhabdomeres are visible in the cross-section. B: Clic109 mutant demonstrates much broader rhabdomeres than the wild-type and excessive splitting of the rhabdomere structure. Scale bar: 3µm. EM pictures provided by Dr. Hikida of OUHCOM.

In continuing with this pattern, the effects of both Slik and Sktl on phosphorylation levels in the rhabdomere were studied. The rhabdomeres in the combination of both Slik and Sktl (Fig. 14A) appear more irregular than those with just Slik alone (Fig. 13A). In ommatidium from Slik and Sktl flies, the normal trapezoidal pattern of rhabdomere formation is abolished and rhabdomere size ranges from slightly widened to much smaller than usual (Fig. 14A). In many places, the

rhabdomeres appear to be growing into one another and cell borders cannot be easily seen. Rhabdomeres also seem to be produced from abnormal locations in the cells, suggesting not only excessive phosphorylation levels but also decreased ability to regulate the intracellular location of Moe. In Clic-deficient flies, the rhabdomeres retain these irregular formations, and appear much wider in size (Fig. 14B). The borders between rhabdomeres are difficult to distinguish and splitting of the structures is very common.

A B

Fig. 14. Electron micrograph of UAS-Slik and UAS-Sktl wt vs. Clic109 ommatidia. A: Wild-type rhabdomeres appear in an irregular pattern; some rhabdomeres are smaller than normal while others are much broader. B: Clic-deficient rhabdomeres are very irregular in shape and pattern (arrows); individual rhabdomere structures are indistinguishable and excessive splitting of the widened rhabdomeres can be seen. Scale bar: 3µm. EM pictures provided by Dr. Hikida of OUHCOM.

To add another level of control to the experimental design, we took advantage of the UAS-MoeWT.myc contruct, which increases wild-type Moesin levels when under a GAL4 driver. This exogenous Moe allowed us to eliminate any possible signaling feedback mechanisms that might inhibit or enhance the Moe promoter activity to affect the levels of Moe; endogenous Moe levels were unaffected. The

addition of both Moe and Slik kinase are expected to increase Moe phosphorylating levels, leading to larger rhabdomeres. The combination of both UAS-Slik and UAS-

MoeWT.myc constructs was compared in both wild-type and Clic-deficient backgrounds. In wild-type flies, the rhabdomeres appear very expanded and connected together. Finger-like projections can be seen protruding from the rhabdomere body on the surface facing the lumen (Fig. 15A). In Clic-deficient flies, the rhabdomeres appear generally wider and very irregularly shaped (Fig. 15B). The rhabdomeres contain more noticeable projections as well. The projections and widening of the rhabdomeres suggests an increase in phosphorylated Moe in the Clic109 mutant. This implies Clic acts to decrease the phosphorylation levels of Moe, since an increase in

Moe phosphorylation is seen when Clic activity is abolished.

A B

Fig. 15. Electron micrograph of UAS-Slik and UAS-MoeWT.myc in wt vs. Clic109 mutant. A: Rhabdomeres form box-like, widened rhabdomeres (arrow) in the wild- type background. At least eight rhabdomeres can be discerned. B: Rhabdomeres appear to be growing into one another in the Clic109 mutant; deciphering the total number of rhabdomeres in one ommatidium is difficult. Excessive splitting of the rhabdomeres is common, resulting in “fingerlike” projections of microvilli (arrow). Scale bar: 3µm. EM pictures provided by Dr. Hikida of OUHCOM.

In addition to the qualitative analysis of the external and internal eye morphology, western blot techniques were used to quantitatively study the Moe phosphorylation levels in the eye tissues (Fig. 16). Using mitochondrial ATP synthetase as a loading control, p-ERM levels were compared between various UAS constructs. Quantification of the difference in p-ERM levels was provided using the ratio of phosphorylated Moe to total Moe for each UAS construct. The western blot results demonstrate that Slik kinase alone increases phosphorylated Moe levels the most, up to 3.8 times more than the control, while Sktl and Sip1 have a minor effect, both coming in at approximately 1.5 times the control. Slik kinase and Sip1 together, however, show a 6.7 fold, significant increase in Moe phosphorylation levels, suggesting a synergistic effect on Moe phosphorylation between Slik kinase and Sip1.

Thus, the quantitative data matches the qualitative data assumption that expanded rhabdomeres are associated with increased levels of phosphorylated Moe.

None

Sktl

Slik

Sip1

Slik + Sip1

P-Moe.myc

Total Moe.myc

Mitochondrial ATP synthetase

Fig. 16. Western blot of phosphorylated Moe.myc. Phosphorylated Moe is demonstrated by the first arrow. Unmarked signals specify endogenous Moe levels (both phosphorylated and total). Numbers at the top of each lane indicate the relative strength of the signal as compared to the wild-type with no UAS elements (None). Note that Slik kinase increases the phosphorylated Moe by a factor of 3.8, while the combination of Slik kinase and Sip1 produces a 6.7 fold increase. Produced with the assistance of Dr. Berryman of OUHCOM.

III. Clic functions synergistically to Moe phosphorylation

The protein Sip1, an orthologue of the human EBP50, is suggested to function with Slik kinase and Moe (Hughes et al., 2010). The conventional view, in contrast to

Sarah Hughes’ view (Hughes et al., 2010), places Sip1 in a membrane-stabilizing complex with Moe, PIP2, and the actin cytoskeleton (Jiang et al., 2014). In order to test both possible cooperative effects, crosses containing both UAS-Sip1 and UAS-

Slik transgenes were produced and compared with UAS-Slik only flies. External features appear very similar and normal in both constructs; however the internal morphology is very different. With the addition of Slik kinase only, the rhabdomeres appear slightly elongated with irregular boxy shapes, as seen in previous experiments

(Fig. 13A). While electron microscope images for UAS-Sip1 alone are currently

lacking, the external morphology appears very normal, suggesting the ommatidial and rhabdomeral structures are probably unaffected (Fig. 18A, B). With both Slik and

Sip1, however, the rhabdomeres appear much larger and more misshapen (Fig. 17A).

Many rhabdomeres appear to be split and jagged, as well as growing from abnormal cellular locations. This suggests a synergistic effect between Slik kinase and Sip1. In contrast, Clic-deficient flies show a marked decrease in rhabdomere size (Fig. 17B).

Rhabdomeres still appear to be jagged on the apical surface and splitting is common, yet the organization of the ommatidium is much more normal than in the wild-type background. Unlike previous results, the loss of Clic actually appears to decrease the phosphorylated Moe levels in the UAS-Slik and UAS-Sip1 rhabdomeres. These results suggest the conventional view of Sip1 function is more applicable to this study.

If Sip1 functions to enhance Slik kinase activity, the loss of Clic would affect both the

Slik alone and Sip1+Slik crosses in a similar fashion. The UAS-Slik in Clic109 appears to show increased phosphorylation levels (Fig. 13B). In contrast, the combined UAS-

Slik and UAS-Sip1 in Clic109 appear to show decreased phosphorylation levels (Fig.

17B). This data suggests Sip1 functions in the conventional view and opens up the possibility that a direct interaction between Sip1 and Clic is taking place.

A B

Fig. 17. Electron micrograph of combined UAS-Slik and UAS-Sip1 in wt vs. Clic109 ommatidia. A: Wild-type rhabdomeres are much wider than normal (arrow) and appear to be growing out of various abnormal locations in the ommatidium. B: Clic109 mutants depict smaller rhabdomeres than the wild-type but remain broader than normal rhabdomeres; splitting of the structure can be seen (arrow). Scale bar: 3µm. EM pictures provided by Dr. Hikida of OUHCOM.

A B

Fig. 18. Light microscope images of UAS-Sip1 in wt vs. Clic109 flies. A: Wild-type flies show normal compound eye development. B: Clic109 flies have normal compound eye development. Scale bars: 1mm.

The proteins Slik kinase, Sktl, and Sip1 have all been implicated to be involved with the phosphorylation of Moe. In contrast, the phosphatase Pp1-87B functions to dephosphorylate Moe (Roubinet et al., 2011). To fully understand the actions and

interactions of Clic in Moe phosphorylation, Pp1-87B needed to be studied. To do this, two copies of the UAS-Pp1-87B construct were introduced into both wild-type and Clic109 mutant crosses. In wild-type ommatidia, some rhabdomeres appeared relatively normal sized, while others were either missing or drastically reduced in size

(Fig. 19A). In contrast, the Clic109 mutants displayed radically diminished rhabdomeres (Fig. 19B). Many rhabdomeres also appear to be missing. These results indicate that Clic may function to inhibit phosphatase activity, thereby contributing to the retention of phosphorylated Moe.

A B

Fig. 19. Electron micrograph of UAS-Pp1-87B (2 copies) in wt vs. Clic109 ommatidia. A: Wild-type ommatidium contains seven rhabdomere structures. Four rhabdomeres appear to be normal size and shape, while the other three are much smaller than is typical (arrow). B: Two distinct rhabdomere structures can be seen in the Clic-deficient background. All rhabdomeres are profoundly diminished in size (arrow). Scale bar: 3µm. EM pictures provided by Dr. Hikida of OUHCOM.

DISCUSSION

The formation of human stereocilia in the auditory system and Drosophila rhabdomeres in the visual system share many conserved pathways. With this knowledge, scientists are able to study the pathways involved in the formation of actin-rich microvilli structures Drosophila. In this study, Drosophila is an optimal model organism due to the presence of only one orthologue for redundant genes in humans. These orthologues include the ERM protein Moe, the chloride intracellular protein Clic, and Sip1.

My study involved elucidating the influences that Clic has on the Moe phosphorylation pathway. Using the loss-of-function Clic109 mutant, Clic’s involvement in Moe phosphorylation was demonstrated. Results suggest Clic functions in several ways to both enhance and inhibit Moe phosphorylation. Clic109 mutants with UAS-Slik and the combinations of both UAS-Slik and UAS-Sktl, and

UAS-Slik and UAS-MoeWT.myc demonstrate an increase in phosphorylated Moe levels when compared to those in the wild-type background. These observations suggest Clic may function to decrease or inhibit Moe phosphorylation. This inhibition can be accomplished in two ways: inhibition of Sktl or inhibition of Slik. Clic109 mutants with two copies of UAS-Pp1-87B and a combination of both UAS-Sip1 and

UAS-Slik depict decreases in Moe phosphorylation levels when compared to those in the wild-type, suggesting Clic enhances Moe phosphorylation or inhibits Moe dephosphorylation. Clic therefore has a possible role in inhibition of Pp1-87B

phosphatase or the stabilization of the scaffolding complex containing Sip1, Moe, and the actin cytoskeleton.

Hassan et al (1998) showed that skittles encodes a PIP5 kinase, which is responsible for producing PIP2. Since PIP2 is required for the phosphorylation and activation of Moe, it makes logical sense that Moe phosphorylation increases when more PIP2 is present. While the western blot (Fig. 15) demonstrates that levels of phosphorylated Moe do in fact increase in UAS-Sktl flies, the increase is marginal and light microscope pictures show disfigured rhabdomeres rather than expanded rhabdomeres. Taking these results into consideration, the relationship between increased gene expression levels of Sktl and increased production of PIP2 needs to be studied. Increased levels of PIP2 in the membrane possibly causes membrane instability and therefore breakdown of forming rhabdomeres, contributing to the fewer rhabdomere structures seen in both the UAS-Sktl wild-type and Clic109 backgrounds.

This possibility can be tested histochemically using a PIP2 marker to determine whether an increase in Sktl actually leads to an increase in PIP2 in the apical membranes of photoreceptors. In addition, Rho kinase can directly induce PIP5K to produce more PIP2 (Hirao et al., 1996). Since activated Moe may form a negative feedback loop with Rho (Speck et al., 2003), this increase in PIP5K activity may be decreasing Rho activity and thereby decreasing PIP2 production enough to produce a lower effect on Moe phosphorylation levels than expected.

Rho signaling not only affects PIP2 production, but also Clic localization

(Ponsioen et al., 2009) and possibly Moe phosphorylation (Yonemura et al., 2002).

Since the results suggest two opposing roles for Clic, namely, inhibition and enhancement of phosphorylated Moe, a possible explanation for these results involves the activity of Rho. RhoGDI complexed with inactive Rho binds to activated ERM proteins, which in turn cause the release of Rho so Rho can bind and be activated by

GTP (Jiang et al., 2014). Rho influences the production of PIP2, which is required for

ERM activation, and in turn ERM proteins activate Rho. This forms the Rho-ERM positive feedback loop. In contrast, the negative feedback between Rho and Moe is illustrated by the work of Speck et al. (2003). Speck et al. observed the formation of large protrusions forming where microvilli should form in Moe-/- mutants; rescue of this phenotype was seen in Moe-/- Rho-/+ double mutants. The Moe-/- mutants had increased Rho levels, and the Moe-/- Rho-/+ knockout restored the Moe-Rho balance and formed more normal structures (2003). The increased levels of Rho when Moe is knocked out suggests Moe is required to inhibit Rho function and a balance between the proteins must be maintained in order to protect the integrity of involved cellular structures. In order to study Rho signaling, a possible future direction involves western blots to detect levels of Rho in each experimental cross, as well as in the Clic109 background. If Rho levels are drastically affected, the effects of Rho must be taken into account to fully understand Moe phosphoregulation.

There are now several new possibilities for Clic’s function in Moe phosphorylation (Fig. 20). Interestingly, in the first half of the model, during the actual phosphorylation of Moe, Clic appears to function as an inhibitor of phosphorylation.

In the second half of the model, the anchorage of Moe to the membrane and to F-actin

and the dephosphorylation of Moe, Clic appears to prevent Moe from dephosphorylation. This intermediate step, the transition between the PIP2-Moe-actin complex and the scaffold complex involving Sip1, seems to be the changeover point for the function of Clic as well. This suggests that Clic may function in an as-of-yet unidentified way in the intermediate step. This function may include the transition between membrane complexes or the recruitment of stabilizing factors involved in the scaffold complex. In order to determine which proteins Clic directly interacts with, co- immunoprecipitation will be performed using a UAS-Clic-GFP construct pulled down by anti-GFP antibodies. The proteins pulled down during this assay will provide an understanding of which proteins in the pathway Clic can be directly affecting. For instance, if Clic directly interacts with Sip1, Moe, and the actin cytoskeleton, Clic is likely involved in the formation of the scaffold complex. If Clic only interacts with

Moe and Slik, then Clic likely functions to either enhance or inhibit Slik kinase activity. With this information, a more accurate model can be developed.

Fig. 20. Model of possible Clic activity in Moe phosphorylation. Blue diamonds indicate Clic has a negative effect on Moe phosphorylation levels, including inhibition of Sktl or Slik kinase. Orange diamonds indicate Clic has a positive effect on Moe phosphorylation levels, including stabilization of the scaffold complex or inhibition of phosphatase.

FUTURE DIRECTIONS

To determine the precise activities of Clic in the Moe phosphorylation pathway, a variety of assays must be utilized in the future. As previously stated, co- immunoprecipitation can be used to detect what specific proteins are directly interacting with Clic. In addition, western blots can be used to determine whether Rho signaling is affected by either the introduced UAS transgenes or the loss of Clic function in Clic109 mutants. The process of producing a western blot assay comparing

Moe phosphorylation levels between the Clic109 crosses and the wild-type crosses is currently underway. The western blot will provide quantitative data on the relative

Moe phosphorylation level difference between the wild-type and Clic109 crosses. This data will also be used to back up the qualitative data procured from the electron micrographs.

Additionally, rhabdomere formation in UAS-Slik or UAS-Sktl wild-type crosses can be studied using mosaic analysis with a repressible cell marker (MARCM)

(Wu and Luo, 2006). In MARCM, one organism is both experimental and a control; in this case, one ommatidium will contain both experimental and control photoreceptors, allowing for comparison of rhabdomere development without the added complexity of two different individuals. To use MARCM analysis, first one chromosome must be altered such that one arm of the chromosome contains the UAS transgene to be studied

(in this case Sktl or Slik kinase), while the other arm contains a GAL80 repressor.

Both of these elements are closely associated with an FLP-recombination target

(FRT). The cross will also contain a GAL4 driver, along with a flippase (FLP)

activated by an ey-GAL4 driver, and a reporter gene, usually a UAS-GFP transgene.

The GAL80 repressor inhibits GAL4 from activating any UAS elements. In the experiment, GMR-GAL4 will be used, which is active during eye morphogenesis.

When FLP is activated, through an ey-GAL4 driver, it functions to recombine the arms of the chromosome on which the FRTs are found (Fig. 21).

Fig. 21. Mosaic analysis with a repressible marker. Heat shock-activated flippase causes recombination at FRTs, forming two distinct daughters. One daughter lacks GAL80, so UAS transgenes, including GFP, are activated by GAL4. One daughter contains two copies of the GAL80 transgene, producing no GFP. From Lee and Luo (2001).

The result, after mitosis, is one cell contains both GAL80 transgenes (on separate arms of the chromosome) and one homozygous mutant cell containing no GAL80 transgenes and two copies of the UAS transgene. The homozygous mutant cell will also have active GFP to distinguish it from homozygous wild-type cells (Wu and Luo,

2006). In addition to UAS-Slik and UAS-Sktl transgenes, this process can also be used in Clic109 mutants using UAS-Clic transgenes. In this way, Clic-deficient mutant photoreceptors and wild-type photoreceptors can be studied in the same retina. This

data would provide additional evidence for Clic’s involvement with various proteins of the Moe phosphorylation pathway.

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