Science, Faculty of
Zoology, Department of
DSpace
UBCV
Roth, Sophie Faith
Master of Science - MSc
University of British Columbia
In the nervous system, glial cells play an important role in ensheathing and protecting nerves from damage, however, the employed mechanisms to properly ensheath nerves are still unknown. In Drosophila melanogaster, the perineurial glia (PG) are a conserved class of glia that surround peripheral nerves and ensure proper ensheathment through interactions with the extracellular matrix (ECM). Glia-ECM interactions occur through focal adhesions (FAs) comprised of integrins and intracellular proteins including talin and vinculin. Loss of FAs leads to the loss of the perineurial glial sheath and disruption of nervous system function. Integrins also associate with the transmembrane protein Basigin (aka Neuroplastin) in the PG. Loss of Basigin leads to constriction across nerves causing deformation of the glial membrane, cytoskeleton and the overlying ECM. The loss of Basigin phenotype can be rescued by a loss of integrin, leading to the hypothesis that Basigin knockdown in the PG leads to increased integrin activation. We performed FLIM-FRET (Fluorescence Lifetime Imaging Microscopy - Förster Resonance Energy Transfer) using paired donor-acceptor sensors inserted in talin that vary in sensitivity to force to quantify changes in integrin activation. Compared to controls, we observed a significant decrease in the FLIM-FRET efficiency in Basigin knockdown animals. This result implies that the sensor is being stretched due to the lengthening of talin, likely because of increased integrin activation. However, the excitation lasers for both the donor and acceptor were turned on when collecting FLIM-FRET images, which led to a false positive FRET signal with donor-only controls and prompted the reinterpretation of results. We also observed a significant increase in the integrated density of vinculin in the PG when Basigin is reduced. Since vinculin binds and stabilizes talin in an extended state, an increase in vinculin recruitment suggests an increase in talin lengthening and thus an increase in integrin activation. Overall, these data provide further evidence that the phenotypes observed with the loss of Basigin are likely caused by an increase in integrin activation. These findings support the proposed role for Basigin as a negative regulator of integrin activation allowing for the modulation of glia-ECM adhesion and overall nerve morphology.
UNDERSTANDING THE ROLE OF BASIGIN IN REGULATING INTEGRIN ACTIVATION AT GLIAL FOCAL ADHESION COMPLEXES IN THE PERIPHERAL NERVES OF DROSOPHILA MELANOGASTER by Sophie Faith Roth B.Sc., Wilfrid Laurier University, 2021 A THESIS SUBMITTED IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF MASTER OF SCIENCE in THE FACULTY OF GRADUATE AND POSTDOCTORAL STUDIES (Zoology) THE UNIVERSITY OF BRITISH COLUMBIA (Vancouver) March 2024 © Sophie Faith Roth, 2024 ii The following individuals certify that they have read, and recommend to the Faculty of Graduate and Postdoctoral Studies for acceptance, the thesis entitled: Understanding the role of Basigin in regulating integrin activation at glial focal adhesion complexes in the peripheral nerves of Drosophila melanogaster submitted by Sophie Faith Roth in partial fulfilment of the requirements for the degree of Master of Science in Zoology Examining Committee: Dr. Vanessa Auld, Professor, Zoology, UBC Supervisor Dr. Calvin Roskelley, Professor, Cellular & Physiological Sciences, UBC Supervisory Committee Member Dr. Ninan Abraham, Professor, Zoology, UBC Additional examiner Additional Supervisory Committee Members: Dr. Kenji Sugioka, Assistant Professor, Zoology, UBC Supervisory Committee Member iii Abstract In the nervous system, glial cells play an important role in ensheathing and protecting nerves from damage, however, the employed mechanisms to properly ensheath nerves are still unknown. In Drosophila melanogaster, the perineurial glia (PG) are a conserved class of glia that surround peripheral nerves and ensure proper ensheathment through interactions with the extracellular matrix (ECM). Glia-ECM interactions occur through focal adhesions (FAs) comprised of integrins and intracellular proteins including talin and vinculin. Loss of FAs leads to the loss of the perineurial glial sheath and disruption of nervous system function. Integrins also associate with the transmembrane protein Basigin (aka Neuroplastin) in the PG. Loss of Basigin leads to constriction across nerves causing deformation of the glial membrane, cytoskeleton and the overlying ECM. The loss of Basigin phenotype can be rescued by a loss of integrin, leading to the hypothesis that Basigin knockdown in the PG leads to increased integrin activation. We performed FLIM-FRET (Fluorescence Lifetime Imaging Microscopy - Förster Resonance Energy Transfer) using paired donor-acceptor sensors inserted in talin that vary in sensitivity to force to quantify changes in integrin activation. Compared to controls, we observed a significant decrease in the FLIM-FRET efficiency in Basigin knockdown animals. This result implies that the sensor is being stretched due to the lengthening of talin, likely because of increased integrin activation. However, the excitation lasers for both the donor and acceptor were turned on when collecting FLIM-FRET images, which led to a false positive FRET signal with donor-only controls and prompted the reinterpretation of results. We also observed a significant increase in the integrated density of vinculin in the PG when Basigin is reduced. Since vinculin binds and stabilizes talin in an extended state, an increase in vinculin recruitment suggests an increase in talin lengthening and thus an increase in integrin activation. Overall, these data provide further evidence that the phenotypes observed with the loss of Basigin are likely caused iv by an increase in integrin activation. These findings support the proposed role for Basigin as a negative regulator of integrin activation allowing for the modulation of glia-ECM adhesion and overall nerve morphology. v Lay summary Two cell types can be found in the nervous system of animals, neurons, which generate electrical signals between the brain and muscles, and glia, which play important roles in protecting and maintaining the health of the nervous system. Glial cells wrap nerves which contain the neurons that generate the electrical signals. As well, the relationship between the glia and their surroundings is important to ensure that the morphology of the nervous system is maintained despite movement or growth. We have collected further evidence to support the importance of a protein called Basigin in the glia of fruit flies, the loss of which is detrimental to the animal. Our data supports that Basigin negatively regulates the activation of a complex of proteins that mediate the interaction between the glia and their surrounding environment. Basigin could play a similar role in the nervous system of vertebrates as a mediator of glial interactions. vi Preface The work in this thesis will be incorporated into a manuscript in preparation as: Title: Basigin negatively regulates integrin activation to modulate perineurial glia-ECM adhesion in Drosophila. Authors: Sophie F Roth and Vanessa J Auld. For this thesis, I performed all experiments and contributed to experimental design, data analysis, writing and editing. Vanessa J Auld contributed to experimental design, data interpretation and analysis, writing and editing. vii Table of contents Abstract ......................................................................................................................................... iii Lay summary ................................................................................................................................. v Preface ........................................................................................................................................... vi Table of contents .......................................................................................................................... vii List of figures ................................................................................................................................ ix List of abbreviations ..................................................................................................................... x Acknowledgments ....................................................................................................................... xii Introduction ................................................................................................................................... 1 Drosophila peripheral glial morphology ..................................................................................... 2 Focal adhesions ........................................................................................................................... 4 Integrin activation ................................................................................................................. 6 Talin-protein interactions at FAs ........................................................................................... 8 Integrin-extracellular matrix interactions at focal adhesions ............................................ 10 Integrin-Basigin interactions ..................................................................................................... 12 Thesis questions ........................................................................................................................ 13 Materials and methods ............................................................................................................... 20 Fly strains and genetics ............................................................................................................. 20 Immunolabeling and image processing ..................................................................................... 20 FLIM-FRET ............................................................................................................................... 21 Statistical analyses .................................................................................................................... 23 Results .......................................................................................................................................... 26 Talin and β-integrin are present in perineurial glial focal adhesions ........................................ 26 Basigin co-localizes to glial focal adhesions and its knockdown causes the ruffle phenotype. 27 FRET sensors in talin are expressed in peripheral nerves ......................................................... 28 Knockdown of Basigin in the perineurial glia leads to a decrease in intramolecular FRET .... 29 The knockdown of Basigin in the PG causes a decrease in intermolecular FLIM-FRET ........ 32 Investigating the higher-than-expected donor alone FRET efficiencies ................................... 34 The knockdown of Basigin leads to an increase in vinculin ..................................................... 36 Discussion..................................................................................................................................... 55 Loss of Basigin in the perineurial glia causes an increase in integrin activation ...................... 55 Intermolecular FLIM-FRET measurements ........................................................................ 60 viii Loss of Basigin promotes vinculin recruitment and ECM binding at glial FAs ....................... 62 References .................................................................................................................................... 68 ix List of figures Figure 1: Vertebrate versus invertebrate peripheral nerve morphology ........................................ 15 Figure 2: Focal adhesion complex ................................................................................................ 16 Figure 3: Focal adhesion complex activation ............................................................................... 17 Figure 4: Talin protein structure .................................................................................................... 18 Figure 5: Talin dimer orientation .................................................................................................. 19 Figure 6: FRET sensors and single fluorophore controls inserted into talin ................................ 24 Figure 7: Lifetime of heterozygous donor .................................................................................... 25 Figure 8: Talin is needed for proper formation of focal adhesions in the perineurial glia ............ 39 Figure 9: Basigin co-localizes with focal adhesions in peripheral nerves and knockdown causes a ruffle phenotype ............................................................................................................................ 41 Figure 10: FRET sensors inserted endogenously into talin are expressed in peripheral nerves ... 42 Figure 11: Basigin RNAi decreases intramolecular FLIM-FRET with F40 sensor in peripheral nerves ............................................................................................................................................ 44 Figure 12: Basigin RNAi differentially affects the two FRET sensors ........................................ 46 Figure 13: Basigin RNAi may decrease intermolecular FLIM-FRET in peripheral nerves ......... 48 Figure 14: FLIM-FRET experiments should be performed by only exciting the donor .............. 50 Figure 15: Basigin RNAi causes increased recruitment of vinculin at focal adhesions in peripheral nerves ........................................................................................................................... 52 Figure 16: Basigin RNAi leads to increased activation of focal adhesion complexes ................. 54 x List of abbreviations ABS Actin binding site Bsg Basigin CD8 Cluster of differentiation 8 CD147 Cluster of differentiation 147 CNS Central nervous system ECM Extracellular matrix EMMPRIN Extracellular MMP inducer EYFP Enhanced yellow fluorescent protein FA Focal adhesion FAK Focal adhesion kinase FLIM Fluorescence lifetime imaging microscopy FRET Förster resonance energy transfer GFP Green fluorescent protein IBS Integrin binding site ILK Integrin linked kinase mAB Monoclonal antibody mCh mCherry MCT Monocarboxylate transporter MIDAS Metal ion-dependent adhesion site MMP Matrix metalloproteinase MS Multiple Sclerosis Disorder Mys myospheroid NMJ Neuromuscular junction NMSC Non-myelinating Schwann cell PBS Phosphate buffer solution PG Perineurial glia PIP2 Phosphatidylinositol 4,5-biphosphate PLA Proximity ligation assay xi PMCA Plasma membrane Ca2+-ATPase PNS Peripheral nervous system RFP Red fluorescent protein RIAM Rap1-interacting-adaptor-molecule RNAi RNA interference ROI Region of interest SPG Subperineurial glia VASP Vasodilator-stimulated phosphoprotein VNC Ventral nerve cord WG Wrapping glia YPet Yellow fluorescent protein for energy transfer αPS Alpha position specific βPS Beta position specific xii Acknowledgments I would like to start by thanking my supervisor Vanessa for your support and words of encouragement throughout my degree, as well as for being a kind and compassionate mentor. Thank you also to my committee members Cal and Kenji for their time and ideas over the past two years. I also need to thank the LSI imaging facility, specifically Guang Gao, for all of your help. Many thanks are also required for all of my lab mates, whose friendship got me through the hardest of times. Thank you especially to Zazil, for sharing not only a home with me but also so many laughs, and for sharing your loving heart with everyone you meet. Thank you, Katie and Sravya, for being so supportive from day one and always being willing to lend an ear or give advice. Your mentorship and friendship mean so much to me. Thank you also to Nat, not only for my beautiful pine tree tattoo, but for being one of the loveliest humans I have ever met. I’m very grateful to know your kindness and humor, and of course, am grateful for your willingness to always lend me books. Thanks to the other wonderful humans I’m lucky to call friends: Celena, Megan, Colin and Carling. Thank you to my parents, Gwen and Don, for embracing my desire to move across the country so fully, and always coming for a visit when needed. Thank you for the many Sunday evening facetimes, and for always being my biggest cheerleaders. Finally, a huge thank you to my partner Max. Your unconditional love and friendship mean the world to me, and I’m so grateful for our time spent together in BC. I can’t wait to see more of the world by your side. 1 Introduction The nervous system is a complex network of neuronal cells and glial cells that work in concert with the other major systems of the body to coordinate many processes including movement, digestion, and fight or flight responses. While research has mainly focused on the function of neurons, the contributions of glia to the nervous system are becoming increasingly appreciated. Glia are found throughout the central nervous system (CNS) and peripheral nervous system (PNS), and they perform a large variety of functions. Glia play a vital role in maintaining the blood-brain and -nerve barriers, surveying the nervous system for immune-related threats, providing nutrient and ion homeostasis support, and guiding axons during development (Lago-Baldaia et al., 2020). Another key function of glia is to wrap and insulate axons through myelination or glial sheath formation. In vertebrate peripheral nerves, the axons at the core of each nerve are ensheathed by either myelinating or non-myelinating Schwann cells. The ensheathing glia are surrounded by a layer of perineurial glial cells to create individual fascicles. Myelinating Schwann cells myelinate large caliber axons to generate saltatory conduction and allow for the electrical impulses to travel at a faster rate (Muppirala et al., 2020). The majority of studies have focused on the roles of myelinating Schwann cells, as hypomyelination is associated with a range of diseases including Multiple Sclerosis Disorder (MS) and Alzheimer’s, while hypermyelination is associated with a range of movement disorders including Tourette syndrome and epilepsy (Knowles et al., 2022). Conversely, non-myelinating Schwann cells (NMSCs) form Remak bundles by wrapping multiple small diameter axons such as autonomic axons and sensory c-fibers, which are responsible for relaying pain information to the brain (Campana, 2007; Kidd et al., 2013; Harty and Monk, 2017; Wei et al., 2019). The disruption of NMSCs lead to disrupted nociceptive signaling, and increased density of NMSCs are associated with hypersensitivity to physical and thermal stimuli (Furusho et al., 2009; Harty and Monk, 2017). 2 NMSC function is implicated in a range of diseases (Ioghen et al., 2020) and yet our knowledge of the functions of NMSCs is lacking in comparison to myelinating Schwann cells. In vertebrates, both myelinated and non-myelinated axons are embedded in a matrix of collagen surrounded by an outer layer of perineurial glia (Figure 1). The perineurium is composed of long flat perineurial glial cells that possess a double basal lamina. Perineurial glial cells are connected via tight junctions to allow for the protection of the nerve fibers from inflammation and even assists in nerve repair following injury (Parmantier et al., 1999; Lewis and Kucenas, 2014). The perineurium is likely CNS-derived, and plays a role in guiding motor axons as well as ensuring proper Schwann cell differentiation during development (Kucenas, 2015). However, research targeting the specific functions and roles of the perineurial glia with regards to protecting the Schwann cells within each fascicle is still ongoing. Revealing all of the roles that the perineurium plays in maintaining nerve health and morphology is a continuing effort (Kucenas, 2015). Drosophila peripheral glial morphology Fruit flies (Drosophila melanogaster) are a model through which we can study how glial cells wrap nerves during development as well as how the glial sheath is maintained (Yildirim et al., 2018). Drosophila lack myelin but possess non-myelinating glia that wrap and protect nerves throughout development similar to vertebrate ensheathing glia in NMSCs (Matzat et al., 2015; Harty and Monk, 2017). Drosophila peripheral nerves in the 3rd instar larval stages possess three layers of glia that form the glial sheath around axons which are encapsulated by a neural lamella formed by extracellular matrix (ECM) (Freeman and Doherty, 2006). The outermost glial layer is known as the perineurial glia (PG), then the subperineurial glia (SPG) and finally the innermost wrapping glial layer (WG) (Freeman and Doherty, 2006) (Figure 1). The WG make direct 3 contact with axons and play a role in mediating the transduction rate and precision of neuronal signaling (Kottmeier et al., 2020). The SPG contains the septate junctions necessary for establishing the blood brain barrier and blood nerve barrier (Stork et al., 2008). The outermost PG is conserved with the perineurium in vertebrate peripheral nerves. In the peripheral nervous system (PNS), the PG make contact with the ECM which is deposited by hemocytes (Evans et al., 2010; Xie and Auld, 2011). The interaction between the ECM and the perineurial glia is vital to maintaining the glial sheath (Meyer et al., 2014; Xie and Auld, 2011). Similar to vertebrates, the functions of the PG in Drosophila are still being uncovered, although so far it is known that the interaction between the PG and the ECM is vital to the functioning of the nervous system and health of the animal (Xie & Auld, 2011; Meyer et al., 2014; Yildirim et al., 2018). The PG arise from neuroblasts in the CNS late in embryogenesis and migrate to the PNS between the ECM and SPG, where they continue dividing during larval and early pupal stages (Stork et al., 2008; Yildirim et al., 2018). When matrix metalloproteinases (MMPs) are either overexpressed or mutated in the PG during larval stages, the ventral nerve cord (VNC) appears elongated (Meyer et al., 2014), demonstrating that a delicate balance of ECM deposition is required for proper nervous system development in which the PG play a likely role. There are two conserved complexes that are expressed by the PG to facilitate ECM connection: the dystrophin-glycoprotein complex and the integrin adhesion complex known as focal adhesion complexes. In Drosophila, disruption of the focal adhesions that facilitate the interaction between the glia and ECM lead to failure of the glia to wrap nerves and death of the animal (Xie and Auld, 2011). As well, loss of integrins in the glia leads to an elongated VNC, similar to when MMPs are overexpressed (Meyer et al., 2014). Integrin is the key focal adhesion transmembrane protein with a single alpha subunit and a beta subunit that binds the cytosolic 4 adaptor protein talin intracellularly. Both integrins and talin show a strong degree of conservation between vertebrates and Drosophila. In vertebrate myelinating Schwann cells, integrin is involved in many stages of development including Schwann cell differentiation and axonal sorting (Denda and Reichardt, 2007; Barros et al., 2011; Muppirala et al., 2020). However, the role of integrins and talin in NMSCs or perineurial glia in vertebrates is not known. The anatomical similarities shared between vertebrates and invertebrates in the perineurial glia and ECM allows for the peripheral nerves of Drosophila to be studied as a model to understand the role of PG-ECM interactions in the perineurial glial cells of vertebrates (Rodrigues et al., 2011). This thesis will focus on how the morphology of the PG and ECM is disrupted in the Drosophila peripheral nerve when focal adhesions are dysregulated. Focal adhesions Focal adhesions (FAs) are large protein complexes that bridge the intracellular actin cytoskeleton to the ECM to allow for the transduction of force across the membrane, leading to potential changes in signalling. They can range from two to five microns in size (Kanchanawong & Calderwood, 2023). FAs are found in a wide range of tissues and are vital to facilitating tissue adhesion and cell migration. FA structure can be conceptualized in three layers: the integrin signalling layer, the force transduction layer, and the actin regulatory layer (Legerstee and Houtsmuller, 2021; Kanchanawong and Calderwood, 2023) (Figure 2). The integrin signalling layer includes the integrin heterodimer with one alpha and one beta subunit, as well as a range of intracellular proteins that can include: integrin linked kinase (ILK), paxillin, kindlin and focal adhesion kinase (FAK). Integrin is a transmembrane protein that binds ligand in the ECM including collagen, perlecan and laminins, while the cytosolic leg of beta subunit binds the head domain of talin intracellularly. Next, the force transduction layer includes the cytosolic adaptor protein talin and vinculin, which both possess actin binding domains. When activated, talin 5 unfolds and exposes vinculin binding sites along its rod domain, which allows for vinculin binding and increased recruitment of actin (Gingras et al., 2005; del Rio et al., 2009). Lastly, the actin regulatory layer includes proteins zyxin and VASP (vasodilator-stimulated phosphoprotein) which both play a role in regulating actin recruitment to FAs (Legerstee and Houtsmuller, 2021). Together, the three layers work together to facilitate communication between the ECM and the cytoskeleton of cells. In Drosophila, analysis of integrin function is simplified because only five alpha and two beta subunits are present (Brown et al., 2000). The alpha subunits include αPS1 to αPS5 (with αPS1, 2, 3 most conserved with clear vertebrate homologues), and the beta subunits βPS and βυ. βPS is considered the main beta subunit since it has the widest expression pattern and is an essential gene, while animals deficient in βυ are still viable (Narasimha and Brown, 2013). Two alpha subunits have been identified in the peripheral nerve, αPS2 (inflated) is found primarily in the PG, while αPS3 (scab) is found in the SPG and WG (Xie and Auld, 2011). αPS1 (multiple edematous wings) is not present. The βPS subunit (myospheroid or mys) is found throughout all three glial layers. Loss of mys in the PG leads to failure of the glia to wrap the nerve whereby the PG can be found only along the length of one side of the nerve rather than surrounding and encapsulating the nerve (Xie and Auld, 2011). Similarly, αPS2-RNAi leads to PG wrapping defects but they are less severe compared to βPS-RNAi, but when both αPS2 and αPS3 were knocked down in the PG, the severity of the wrapping defects increased (Xie and Auld, 2011). This result implied a potential redundancy of αPS2 and αPS3 in the PG. Of the intracellular components of the FA, there is a single talin homologue, rhea, which is highly homologous to vertebrate talin isoforms 1 and 2 (Brown, 2002). The loss of mys and talin in the optic stalk glia leads to defects in perineurial glial morphology and axon migration in the CNS (Xie et al., 2014). 6 Talin can also be found throughout the peripheral nerve, and knockdown of talin in the PG results in a similar phenotype as mys RNAi where the glia fail to wrap the nerve, revealing the close association between these two proteins in the PG (Xie and Auld, 2011). Overall, FAs mediate the connection in the peripheral nerve between the PG and ECM, and disruption to FA components lead to glial wrapping defects. However, the dynamic modulation of glia-ECM interactions through FA activation is still being elucidated today. Integrin activation Integrin activation is an important first step in FA formation and increased integrin activation is associated with the strengthening of cell adhesion (Shattil et al., 2010). Therefore, integrin activation can be thought of as synonymous with overall FA activation. Integrins perform bi-directional signalling to the ECM and cytoskeleton for mechanotransduction, as well as for cell differentiation and development (Hu and Luo, 2012). In its inactive form, the extracellular domains of alpha and beta integrin are bent down towards the membrane (Xiong et al., 2001) (Figure 3). In this bent conformation, the binding of ECM ligand is not favourable, although integrin has been shown to bind ligand in solution (Luo et al., 2007). Binding of divalent cations such as Mg2+ or Mn2+ to the MIDAS (metal ion-dependent adhesion site) of alpha integrins increases its affinity for ligand binding by attracting the negatively charged residues of ligand, whereas binding of Ca2+ has an inhibitory effect (Luo et al., 2007). An inhibitory salt bridge is also present between the alpha and beta subunits within the transmembrane domain to stabilize the closed conformation (Kadry and Calderwood, 2020). Integrin can be activated by outside-in signaling whereby the alpha subunit first binds ligand in the ECM to initiate FA formation. Conversely, integrin can be activated via inside-out activation. Inside-out activation involves the initial binding of talin or kindlin, an integrin co-7 activator, intracellularly to induce the extension of the alpha and beta subunits extracellularly, as well as promote disruption of the salt bridge (Sun et al., 2019; Haydari et al., 2020). When activated, integrin may possess an intermediate affinity state or a high affinity state (Luo et al., 2007; Kadry and Calderwood, 2020) (Figure 3). The difference between the two conformations is observed in the orientation of the alpha and beta headpieces. In the high affinity state, an open headpiece conformation is observed whereby the angle between the head and hybrid domains of beta integrin becomes larger compared to in the intermediate state (Takagi et al., 2002; Takagi et al., 2003; Xiao et al., 2004). This conformational change promotes ligand binding in alpha integrin and stabilizes the high affinity state. Furthermore, the open headpiece conformation leads to a separation in the alpha and beta leg subunits by approximately 70 Å that stabilizes integrin extension (Xiao et al., 2004; Luo et al., 2007). Talin binding also plays a role in integrin activation by inducing conformational changes including a change in the tilt angle of integrin in the membrane, leading to destabilization of the alpha and beta subunit interaction and promoting integrin activation (Kim et al., 2012; Ginsberg, 2014). Interestingly, talin mutants in Drosophila have shown little to no effect on the ability of integrins to bind ligand-mimetic antibodies, suggesting that integrin is primarily activated via outside-in activation in this organism (Brown et al., 2002; Helsten et al., 2008; Ellis et al., 2014). Rather, the head domain of talin plays an important role in reinforcing adhesions via the promotion of integrin clustering (Brown et al., 2002; Ellis et al., 2014). Integrin clustering can occur either through the binding of ECM ligand or through the dimerization of talin molecules (Legerstee and Houtsmuller, 2021). As FAs grow and mature, integrin shows advanced clustering as more proteins are recruited (Goult et al., 2021). To assist in this process, inactive talin is recruited by activated talin and the newly formed dimer binds a neighboring integrin molecule 8 via paxillin, an FA scaffolding protein, allowing for FAs to mature (Lu et al., 2022). Talin, as a mediator between integrin and the actin cytoskeleton, binds many proteins to modulate the interaction on either side of the membrane. Talin-protein interactions at FAs Talin is a large cytosolic adaptor protein that is vital to FA function because of its link between ECM-bound integrins and the internal actin cytoskeleton. In Drosophila, animals deficient in the single talin isoform fail to form connections between integrin and the actin cytoskeleton (Brown et al., 2002). Talin is composed of approximately 2500 amino acids formed into head and rod domains connected by a linker (Klapholz and Brown, 2017; Goult et al., 2021) (Figure 4). Talin possesses an N-terminal FERM domain containing integrin-binding site 1 (IBS1) where it binds the cytosolic leg of βPS. The rod domain is formed by 13 α-helical bundles containing a variety of binding sites including those for vinculin and actin (Klapholz and Brown, 2017). When talin is inactive, the rod domain inhibits the binding of βPS (Goksoy et al., 2008; Ellis et al., 2013). Phosphatidylinositol 4,5-biphosphate (PIP2) promotes activation of talin by unfolding the protein, thus exposing the β-integrin binding site (Goksoy et al., 2008), while RIAM (Rap1-interacting-adaptor-molecule), is important for bringing talin to the membrane (Ellis et al., 2013; Chang et al., 2014; Hu et al., 2016). The application of force on talin allows for unfolding of the helical bundles which exposes otherwise hidden vinculin binding sites (del Rio et al., 2009). Once bound, vinculin stabilizes the unfolded conformation of talin (Yao et al., 2014). While talin can possess up to 11 vinculin binding sites (Gingras et al., 2005), vinculin only possesses a single talin-binding D1 domain, as well as other protein binding sites. Similar to talin, vinculin is formed by head and rod domains, and performs autoinhibition via folding. Through the rod domain, vinculin can also bind actin to provide additional mechanical support while the head domain is involved in regulating integrin clustering (Humphries et al., 2007). 9 Interestingly, hyperactive vinculin, but not talin, in Drosophila leads to animal death (Tanentzapf et al., 2006; Maartens et al., 2016). Hyperactive vinculin lethality was likely mediated through the ability of vinculin to recruit closed talin in the cytosol to form nascent aggregates that lacked integrin and actin recruitment (Maartens et al., 2016). Overall, vinculin plays an important role at FAs by stabilizing the unfolded conformation of talin and recruiting additional actin proteins. Furthermore, a common method of studying how proteins such as talin and vinculin experience force is through the use of FRET (Förster resonance energy transfer) sensors composed of donor and acceptor fluorophores linked through a flexible peptide (Austen et al., 2013). Many functions of the various binding domains on talin have been investigated using endogenous FRET sensors where the stretching of talin under force leads to a decrease in energy transfer from the donor to the acceptor and thus a decrease in the overall FRET efficiency. Examples include the use of FRET sensors in Drosophila to investigate the amount of force talin experiences at muscle attachment sites during development (Lemke et al., 2019), although most of this work has been completed in cell culture with mouse talin1 (Austen et al., 2015; Kumar et al., 2016; Ringer et al., 2017; Driscoll et al., 2020). For example, the use of FRET sensors in talin has allowed for the functions of the various actin binding sites (ABS) along the rod domain to be investigated. ABS2 is found in the middle of the rod domain and is important for recognizing ECM stiffness (Kumar et al., 2016). In contrast, ABS3 is found at the C-terminus and is responsible for differentially regulating the tension at central versus peripheral adhesions, where peripheral adhesions are often under more tension (Kumar et al., 2016). ABS3 is also the least mechanically stable domain of the talin rod, and so it plays a key role in the initial mechanosensing of forces (Yao et al., 2016). ABS3 encompasses the dimerization domain of talin which is composed of a five-helix bundle including a THATCH domain that enhances actin 10 binding when the proteins are dimerized (Gingras et al., 2008). The formation of antiparallel talin homodimers is vital to inducing ligand binding in integrins and forming large FA complexes (Lu et al., 2022). Using molecular dynamic simulations, the talin dimer has been shown to switch between an orientation in which the two talin proteins are forced together, leading to a collapsed talin dimer, or the FAs may be forced apart, leading to a more energetically favorable conformation of an elongated talin dimer (Golji and Mofrad, 2014). Similarly, in Drosophila, talin dimers have been shown to organize differently in various tissues, for example, the talin proteins may orient parallel (i.e. elongated dimer) or perpendicular (i.e. collapsed dimer) to the membrane depending on if integrin is bound at both integrin-binding site 1 (IBS1) and IBS2, or just IBS1 (Klapholz et al., 2015) (Figure 5). The orientation of the talin dimer to the membrane can also be affected by mechanical stimulation (Golji and Mofrad, 2014; Hu et al., 2016; Owen et al., 2022). Overall, talin experiences forces at its N-terminus from integrin-binding, as well as at its C-terminus, from dimerization and actin-binding. The forces acting on FA proteins are very dynamic, and often a change in conformation of one FA protein leads to consequent changes in associated proteins as well. These conformational changes are also influenced by the binding of integrin to ligand in the ECM. Integrin-extracellular matrix interactions at focal adhesions At FAs, the head domain of alpha integrins bind ligand in the ECM. The ECM layer is composed of proteoglycans like perlecan and transmembrane proteins like syndecan, as well as fibrous proteins including collagens, fibronectins and laminins (Frantz et al., 2010). During Drosophila morphogenesis, the ECM in the nervous system and epithelia is remodelled. During embryogenesis, germ band retraction allows for the remodelling of the epithelium (Lynch et al., 2014). Integrins and their interaction with the ECM proteins, laminins, have been found to play a key role in this process (Schöck and Perrimon, 2003). The end of embryogenesis is marked by 11 dorsal closure, whereby the hole produced by germ band retraction is sealed over. Both αPS3 and βPS mutants show defects in dorsal closure (Reed et al., 2004; Narasimha and Brown, 2004). Integrin-ECM interactions are also vital for nervous system remodeling. During larval stages, the PG secrete proteases such as matrix metalloproteinases (MMPs) to remodel the matrix (Meyer et al., 2014; Skeath et al., 2017). The overexpression of MMPs leads to the loss of integrin-ECM interactions and causes ventral nerve cord (VNC) lengthening in the CNS and glial wrapping defects in the PNS (Xie and Auld, 2011; Meyer et al., 2014). In Drosophila, extensive research has been done on the role of ECM receptors such as integrins in the roles of remodelling of the neuromuscular junction (NMJ), and the growth and guidance of axons (Broadie et al., 2011; Myers et al., 2011), however, little is known regarding the role that integrins play in the glia in terms of remodelling of the nervous system. So far, it has been shown that the stiffness of the ECM can regulate the migration of retinal glial cells in the CNS of Drosophila and that loss of integrins leads to ECM softening as well as faulty glial migration (Kim et al., 2014; Tavares et al., 2015). ECM stiffness and deposition are tightly regulated in order to mediate forces at FAs to maintain overall cell and membrane morphology. For example, tension across talin is reduced on stiffer substrates compared to softer substrates as shown with FRET sensors (Kumar et al., 2016; Ringer et al., 2017). Interestingly, incorporation of perlecan has been shown to decrease ECM stiffness, whereas collagen-IV and integrins promote stiffness (Pastor-Pareja and Xu, 2011; Ge et al., 2021). Furthermore, integrin signaling was shown to be sufficient in regulating lox expression, a gene that encodes Lysyl oxidase, an enzyme that regulates ECM stiffness (Kim et al., 2014). We know that the glial layers and ECM present in larval peripheral nerves are remodelled during pupation (Yildirim et al., 2018), however we do not know the role that 12 integrins play in mediating glia-ECM interaction during development nor how integrin-ECM interactions are regulated in glia. Integrin-Basigin interactions Basigin (Bsg) is a transmembrane glycoprotein that has previously been shown to associate with integrins in the perineurial glia of Drosophila (Hunter et al., 2020). Vertebrate Bsg is also known as CD147 or EMMPRIN (extracellular MMP inducer), and is part of a superfamily with Embigin and Neuroplastin; although the single Drosophila Bsg family member, Basigin, shows a stronger homology to Neuroplastin. Drosophila Bsg and vertebrate Neuroplastin both possess 2-3 extracellular Ig domains with varying numbers of glycosylation sites as well as a conserved transmembrane domain that includes a fully conserved glutamate residue (Zhu et al., 2017). Vertebrate Bsg has been shown to localize to the CNS in mice and be involved in neuronal interactions (Fan et al., 1998) whereas Neuroplastin is primarily expressed in the CNS and non-neural tissues (Langnaese et al., 1997). Similarly, Bsg is expressed by neurons in Drosophila and is involved in dendrite morphogenesis (Shrestha et al., 2021). Bsg is also expressed in the visual system of Drosophila and is important for glial migration around photoreceptor axons (Curtin et al., 2007), similar to integrins (Xie et al., 2014). Therefore, Bsg plays a number of roles in the nervous system in both vertebrates and invertebrates. Bsg has been shown to be multifunctional, and possesses many binding partners across cell types, including monocarboxylate transporters (MCTs) (Kirk et al., 2000), and as the name EMMPRIN suggests, Bsg also plays a role in inducing MMPs to degrade ECM (Muramatsu, 2016). As well, Bsg often acts as a chaperone by bringing proteins to the membrane, including PMCA (plasma membrane Ca2+-ATPase) (Schmidt et al., 2017), and MCTs in retinal Muller glial cells, which are vital to glycolysis for energy production (Philip et al., 2003). Interestingly, Bsg 13 has been found to be overexpressed on the cell surface of some types of tumors, such as glioma and melanoma (Tian et al., 2013; Zeng et al., 2014), and the deleterious phenotypes are sometimes mediated through an interaction with integrin (Muramatsu and Miyauchi, 2003; Muramatsu, 2016). For example, Bsg was shown to activate β1 integrin, likely through interaction with the MIDAS in hepatoma cells (Li et al., 2012; Wu et al., 2015). Bsg has been shown to interact with integrin in photoreceptor cells of cultured insect cells where loss of Bsg leads to disruption of the cytoskeleton (Curtin et al., 2005). In Drosophila, Bsg has been shown to interact with integrins in various tissues including during embryogenesis (Reed et al., 2004), at larval neuromuscular junctions (Beumer et al., 1999; Besse et al., 2007), and at the perineurial glial membrane (Hunter et al., 2020). Thesis questions Previous work in the lab has showed that Bsg and β-integrin are associated with one another in the perineurial glia, demonstrated by a positive proximity ligation assay signifying a distance of 40nm or smaller, and knockdown of Basigin leads to a ruffle phenotype where glial and ECM morphology are grossly affected (Petley-Ragan, 2016; Hunter et al., 2020). When Bsg is knocked down in the PG, ventral nerve cord shortening is observed, as well as ruffles in the glial membrane of peripheral nerves, indicating morphological defects in both the CNS and PNS (Hunter et al., 2020). Interestingly, these phenotypes are supressed with a reduction in myospheroid (β-integrin) expression and are partially supressed with a reduction in rhea (talin) expression (Hunter et al., 2020). This leads to the hypothesis that Bsg potentially interacts with integrins at the PG membrane and may be involved in negatively regulating the activation of integrins to mediate glia-ECM interaction. Therefore, the defects seen with Bsg knockdown are likely a result of integrin dysregulation, potentially leading to too much cell adhesion between the glia and ECM. However, the nature of the interaction between Basigin and β-integrin has yet 14 to be fully elucidated. This thesis will investigate how Basigin interacts with β-integrin in the perineurial glia of Drosophila. 15 Figure 1: Vertebrate versus invertebrate peripheral nerve morphology. (A) Cross-section through a vertebrate peripheral nerve (left) and a peripheral nerve of Drosophila melanogaster (right). In vertebrates, axons are ensheathed by myelinating Schwann cells or grouped into non-myelinating Remak bundles where they are directly wrapped by glial cells. Schwann cells and Remak bundles are set in a dense collagen-rich endoneurium and wrapped by the perineurium, which contains tight junctions. The nerve is ensheathed by extracellular matrix (ECM) in the epineurium. In Drosophila, axons are wrapped directly by non-myelinating glia, which are then ensheathed by the subperineurial glia containing a septate junction. The perineurial glia make contact with the ECM in the neural lamella. (B) Longitudinal section of Drosophila peripheral nerve showing glial layers. Adapted from Rodrigues et al., 2011. Made in BioRender. 16 Figure 2: Focal adhesion complex. The integrin signalling layer includes transmembrane receptor integrin, composed of alpha and beta subunits. Alpha integrins bind ligand in the extracellular matrix (ECM), while beta integrin binds kindlin (integrin co-activator) and integrin-linked kinase. The force transduction layer includes cytosolic adaptor proteins talin and vinculin which connect integrin to the actin cytoskeleton. The actin regulatory layer includes proteins such as α-actinin, VASP (vasodilator-stimulated phosphoprotein) and zyxin. Made in Biorender. 17 Figure 3: Focal adhesion complex activation. In its inactive state, integrin alpha and beta heads are bent towards membrane and do not bind intracellular proteins. In the intermediate affinity state, integrins bind ligand in the extracellular matrix and talin intracellularly. In the fully active form, integrin has a high affinity for ligand and the headpiece conformation of beta integrin has changed to promote ligand binding. This change in headpiece conformation of beta integrin allows for the separation of the alpha and beta legs to promote integrin extension. Integrin clustering may also occur through dimerization of talin, leading to an increased recruitment of vinculin and actin. Made in Biorender. 18 Figure 4: Talin protein structure. Talin is composed of a head FERM domain (F0-F3) joined by a linker to the rod domain made up of α-helices (R1-R13). Rap1 interacts with F0 to promote talin activation and promote RIAM association (Rap1-interacting-adaptor-molecule). F3 binds FAK (focal adhesion kinase), RIAM and PIPKIγ which are involved in activating talin. Talin possesses two integrin binding sites (IBS), one at F3 (IBS1) and a second at R11 (IBS2). Talin possess 11 vinculin binding sites spanning R2-R4, as well some binding sites further down the rod. There are three actin binding sites (ABS), ABS1 spans F2-F3, ABS2 spans R4-R8 and ABS3 spans R13 to the dimerization domain. ABS3 is the most important site for binding actin due to its ability to stabilize strong forces. Talin can form antiparallel homodimers through a THATCH domain at the C-terminus which also promotes binding of actin to ABS3. Adapted from Klapholz and Brown, 2017. Made in Biorender. 19 Figure 5: Talin dimer orientation. Talin dimers have been shown to organize differently in various tissues. When the talin head domain only binds integrin binding site 1 (IBS1), this leads to a collapsed talin dimer (dimer is more perpendicular to the membrane; left). Alternatively, if talin binds both IBS1 and IBS2, this leads to an elongated dimer (dimer is parallel to the membrane; right). The orientation of the dimer can be influenced by mechanical stimulation from actin forces at the C-terminus. Made in Biorender. 20 Materials and methods Fly strains and genetics The following Drosophila strains were used: talin-RNAi (GD12050) (Dietzl et al., 2007) and basigin-RNAi (GD989, v2798 (RNAi 1) and GD15718, v43306 (RNAi 2)) were from the Vienna Stock Centre (VDRC). The following lines were from the Bloomington Drosophila Stock Center: UAS-Dicer2 (Dietzl et al., 2007), UAS-LacZ (Brand and Perrimon, 1993), UAS-mCD8::RFP (Yang et al., 2008), UAS-mCD8::mCherry (RRID:BDSC_27391) and Viking::GFP (Morin et al., 2001). 46F-GAL4 (Xie and Auld, 2011) was from the Kyoto Stock Centre (DGRC). Bsg::mCherry was previously generated (Hunter et al., 2020). The following fly lines were gifted by Dr. Frank Schnorrer (Max Planck Institute of Biochemistry): talin-1-F40, talin-C-F40, talin-1-HPst, talin-C-HPst, talin-1-YPet, talin-C-YPet, talin-1-mCherry, talin-C-mCherry (Lemke et al., 2019) (Figure 6). All RNAi experiments were performed at 25℃. Only Bsg-RNAi experiments used Dicer2 in the background. Immunolabeling and image processing Wandering third instar larvae were dissected and fixed as previously described (Clayworth et al., 2023). Larvae were fixed in 4% paraformaldehyde in phosphate buffer solution (PBS) for 15 minutes (10 minutes for βPS antibody). The following antibodies were used for immunofluorescence: mouse anti-βPS (CF.6G11, DSHB) at 1:10 (Brower et al., 1984), mouse anti-futsch at 1:2000 (22C10, DSHB) and rabbit anti-Drosophila Vinculin at 1:500 (gifted by Dr. Michael Krahn, University of Münster) (Heiden et al., 2021). Secondary antibodies used were goat-anti-mouse or goat-anti-rabbit conjugated to Alexa647 at 1:300 (Invitrogen, Toronto, Canada). Fluorescent images were taken using a Deltavision microscope (Applied Precision, 21 Mississauga, Canada) using a 60X oil immersion objective (NA 1.4) with a step size of 0.2 µm. Z-stacks were deconvolved with Softworx image processing software using a point spread function measured with a 0.2 µm fluorescent bead (Invitrogen, Toronto, Canada) in Vectashield (Vector Laboratories, Burlington, Canada). Stacks were further processed using Fiji (Schindelin et al., 2012) and Adobe Photoshop (Adobe Systems). For integrated density calculations, the mean fluorescence intensity was measured in Fiji and multiplied by the area of the region of interest. FLIM-FRET Wandering third instar larvae were placed in a breathable chamber inside a glass coplin jar filled with cotton soaked in diethyl ether (ThermoFisher Scientific) for 5-7 minutes for anaesthetization (Kakanj et al., 2020). Once asleep, larvae were arranged on agar pad (3% agar in water; select agar - Invitrogen) and secured with a coverslip (Skinner et al., 2013). Larvae were imaged on a Leica TCS Sp8 laser scanning confocal microscope with a 25X long-working distance water immersion objective (NA 0.95). Images were acquired with a scanning velocity of 600Hz, a spatial resolution of 512 x 512 pixels and a zoom factor of 2.0X, resulting in an image field coverage of 232.12µm x 232.12µm2. For intramolecular (Figures 11 and 12) and intermolecular (Figure 13) FRET experiments, FLIM images were collected with both the donor (YPet; 517 excitation) and acceptor (mCherry; 587 excitation) excitation lasers on. As well, determination of the unquenched donor lifetime (Figure 7) was measured with both donor and acceptor excitation lasers on. FLIM and FLIM-FRET experiments in Figure 14 were measured using only the donor excitation laser. In all FLIM-FRET experiments, regions of interest were selected spanning the width of multiple peripheral nerves at various distances from the ventral 22 nerve cord. All ruffled areas observed with Basigin-RNAi were also assayed. FLIM (fluorescence lifetime imaging) was calculated using Leica LASX software and ROIs were averaged per animal. The donor lifetime (YPet – yellow fluorescence protein for energy transfer) was determined to be approximately 2.9 ns and was used in FRET calculations as a mono-exponential donor (Figure 7) (Ringer et al., 2017). YPet is derived from EYFP, whose lifetime is also approximately 2.9 ns (Bajar et al., 2016). For Förster resonance energy transfer (FRET) measurements, larvae expressing the endogenous sensors (donor – YPet; acceptor – mCherry) (Lemke et al., 2019) were imaged as described above and the apparent FLIM-FRET efficiency was calculated using LASX software (see calculation below). 𝑬𝑬 = 𝟏𝟏 − 𝝉𝝉𝒒𝒒𝒒𝒒𝒒𝒒𝒒𝒒𝒒𝒒𝒒𝒒𝒒𝒒𝒒𝒒𝝉𝝉𝒒𝒒𝒒𝒒𝒒𝒒𝒒𝒒𝒒𝒒𝒒𝒒𝒒𝒒𝒒𝒒𝒒𝒒𝒒𝒒 E = FRET efficiency, 𝜏𝜏quenched = lifetime of donor in presence of acceptor, 𝜏𝜏unquenched = lifetime of donor only (2.9 ns) Two sensors of varying sensitivity to force were used – F40 sensitive to 1-6pN and HPst sensitive to 9-11 pN (Grashoff et al., 2010; Austen et al., 2015; Lemke et al., 2019). Sensors varied by the strength of linker peptide between the donor and acceptor. The linker in the F40 sensor was composed of a Flagelliform peptide (Grashoff et al., 2010), whereas the linker in the HPst sensor was a stable Villin headpiece (Austen et al., 2015). Sensors were either inserted in between the head and rod domain of talin (~amino acid 456), referred to as the N-terminus sensor, or at the C-terminus of talin (Figure 6). 23 Statistical analyses All statistical tests were performed using GraphPad Prism 9 (GraphPad Software, La Jolla, CA). Normality was tested for and a One-way ANOVA with Tukey’s post-hoc test or a Kruskall-Wallis test with Dunn’s multiple comparisons were performed. For two-group comparisons, an unpaired t-test was performed. All graphs show the mean and standard deviation. 24 Figure 6: FRET sensors and single fluorophore controls inserted into talin. C – sensor/single fluorophore inserted at C-terminus, 1 – sensor/single fluorophore inserted at N-terminus. F40 sensor linker between donor (D – YPet) and acceptor (A – mCherry) is composed of flagelliform protein and is sensitive to 1-6pN of force, whereas HPst linker is a stable villin headpiece sensitive to 9-11pN of force (Ringer et al., 2017; Lemke et al., 2019). Made in BioRender, adapted from Lemke et al., 2019. 25 Figure 7: Lifetime of heterozygous donor. YPet (yellow fluorescent protein for energy transfer) donor at C-terminus (blue) or N-terminus (green). Unpaired t-test (** p = 0.0011). ns – nanoseconds. 26 Results Talin is present and necessary for perineurial glial focal adhesions The focal adhesion (FA) protein, talin, was previously observed to localize throughout the peripheral nerve, and its knockdown in the perineurial glia (PG) caused a failure to wrap phenotype similar to β-integrin knockdown (Xie and Auld, 2011). However, previous observations of the loss of talin phenotype were qualitative and so we sought to quantify and characterize these phenotypes. To better understand the role that talin plays in facilitating communication between the perineurial glia (PG) and extracellular matrix (ECM), talin was knocked down using RNA interference (RNAi) in the PG. To assess distribution, talin endogenously tagged with YPet at the C-terminus (Talin::C-YPet; Figure 8A-B) and at the N-terminus of the rod domain (Talin::1-YPet; Figure 8C-D) were tested for localization. In controls, talin was observed throughout the peripheral nerve and concentrated in the PG and SPG. Using a PG-specific Gal4 driver (46F-Gal4) to drive talin-RNAi led to the loss of signal within the PG membranes as well as phenotypes of varying severity (Figure 8B, D, G). Specifically, the knockdown of talin in the PG led to failure of the PG to fully ensheath the nerve (Figure 8B’’, F’’; orange arrowheads), similar to the phenotype seen with loss of the integrin beta-subunit βPS (aka myospheroid; mys) (Xie and Auld, 2011). This phenotype was seen on average in 90.78% of all nerves with talin-RNAi and was consistently observed along the length of the affected nerves (Figure 8G). These data suggest that PG that lacks talin is likely unable to bind to ligand in the ECM when wrapping the nerve during development. As well, we observed a less common phenotype of an asymmetric and uneven accumulation of PG membrane, often surrounding a PG nucleus, on one side of the nerve in 28.45% of nerves (Figure 8B’’, D’’, G; white arrowheads). Both phenotypes were significantly greater with talin knockdown compared 27 to controls, which never displayed these phenotypes. In controls, talin colocalized with βPS in the PG (Figure 8E), while knockdown of talin caused a loss of the βPS signal in the PG (Figure 8E’’’-F’’’). These results indicate the importance of talin expression in the PG for proper FA formation and that FAs are needed for the PG to fully ensheath nerves. Basigin co-localizes to glial focal adhesions and knockdown causes a ruffle phenotype Basigin was previously shown to colocalize with βPS in the PG (Hunter et al., 2020), so we sought to confirm that this colocalization between Basigin and βPS occurs at mature FAs also containing talin. Previously, Basigin was endogenously tagged with mCherry using Recombinant-Mediated Cassette Exchange; Bsg::mCherry is a homozygous viable insertion (Hunter et al., 2020). In wildtype peripheral nerves, Basigin::mCherry colocalized with both talin and βPS, presumably at PG focal adhesions (Figure 9 A-A’’’, arrowheads). To confirm the importance of Basigin, we knocked down Basigin in the PG using 46F-Gal4 and observed the expected ruffle phenotype (Hunter et al., 2020) where PG membranes appear compressed and protrude from the nerve (Figure 9C’’, D’’, arrowheads), and the ECM mirrors these protrusions (Figure 9 C’, D’). This phenotype has been previously characterized and quantified (Hunter et al., 2020), where the ruffle phenotype is observed in 40-50% of nerves per animal on average. The ruffle phenotype was normally observed only once per affected nerve (in a region spanning 15-30µm) (Figure 9E-E’’; showing morphologically normal region). Since the Bsg-RNAi phenotype can be suppressed with a reduction in either of two key focal adhesion proteins, the integrin beta-subunit (mys) and talin (rhea) (Hunter et al., 2020), we hypothesized that the Bsg-RNAi phenotype is caused by an increase in integrin activation at FAs connecting the PG to the 28 ECM. An increase in integrin activation leads to an increase in adhesion between the PG and ECM cell layers, resulting in the ruffle phenotype. FRET sensors in talin are expressed in peripheral nerves To test the hypothesis that knockdown of Basigin in the PG leads to an increase in integrin activation, we investigated the effect on talin using previously developed tension sensors (Lemke et al., 2019). Specifically, Förster resonance energy transfer (FRET) sensors utilizing yellow fluorescent protein for energy transfer (YPet; donor) and mCherry (acceptor) connected by a linker peptide were inserted into the endogenous talin gene either at the C-terminus or the N-terminus of the rod domain (Figure 10A) (Lemke et al., 2019). The C-terminus sensor is a zero-force control whereas the insertion at the N-terminus of the rod domain is susceptible to stretching forces (Austen et al., 2015; Kumar et al., 2016; Ringer et al., 2017). Two sensors of varying sensitivity to force – the F40 sensor (1-6pN) and the HPst sensor (9-11pN) were used (Grashoff et al., 2010; Austen et al., 2015). Single fluorophore controls with either just the donor or just the acceptor at one of the two positions in the rod domain were also employed. All insertions are viable and do not disrupt talin function (Lemke at al., 2019). As outlined above (Figure 8), talin tagged with only the donor colocalized with βPS in the PG, confirming that the insertions do not disrupt talin localization in nerves. In addition, Talin::YPet colocalized with Talin::mCherry in peripheral nerves expressing the HPst sensor at the C-terminus and showed the expected talin expression pattern (Figure 10B-B’’, arrowheads). 29 Knockdown of Basigin in the perineurial glia leads to a decrease in intramolecular FRET The F40 and HPst sensors were expressed as heterozygotes in the background of either UAS-LacZ (control) or Basigin RNAi with 46F-Gal4 as the PG specific driver (Figure 11, Figure 12). We performed fluorescence lifetime imaging microscopy (FLIM) using these sensors to assess the fluorescence lifetime of the donor (YPet), and draw conclusions regarding its association with the acceptor (mCherry). Close association of the two fluorophores (1-10nm) leads to a reduction in the fluorescence lifetime of the donor due to FRET, while increasing distances between the fluorophores (>10nm) leads to an unaffected donor lifetime (Ishikawa-Ankerhold et al., 2012). We first assessed the most sensitive sensor (F40, 1-6-pN) at both the C-terminus and N-terminus of the rod domain of talin (Figure 11A-G). In controls, FLIM-FRET analysis recorded longer donor lifetimes in the outer layers of glia at the exterior of the nerve (within the PG layer where the majority of FAs reside) and this is indicated by warmer colours (Figure 11A’’, D’’). There is talin within the interior of the nerve (Xie and Auld, 2011) and analysis of donor FLIM within the interior of the nerve recorded shorter lifetimes indicated by cooler colours (Figure 11A’’, D’’). These data suggest that within the PG the talin sensor is normally under some degree of tension whereas talin is not as engaged in the interior of the nerve. FLIM-FRET was then measured in nerves where Basigin was knocked down using the PG specific driver 46F-Gal4. There was an increase in the fluorescence lifetime using the F40 sensor with Basigin-RNAi indicated by the prevalence of warmer colours compared to controls, particularly within the ruffle regions (Figure 11B”, E”). Quantification of FLIM-FRET was carried out in regions of interest that spanned the width of control nerves where we measured a FRET efficiency of 27.7% for the C-terminus sensor and 35.6% for the N-terminus sensor (Figure 11H). FLIM-FRET of the F40 sensors was quantified in nerves with knockdown of 30 Basigin. Two regions of interest were assessed: one spanning the width of the nerve (same as control) in a morphologically normal region and one within a ruffle area (Figure 11B”, E’’). Bsg-RNAi nerves in both regions had significantly reduced FRET efficiency compared to controls (Figure 11H). For Bsg-RNAi line 1, the FRET efficiency of morphologically normal nerves was 15.77% with the C-terminus sensor but 21.9% with the N-terminus sensor. In ruffled areas, the C-terminus sensor showed 6.29% efficiency and 8.43% at the N-terminus. Similar values were recorded for Bsg-RNAi line 2 (Figure 11H). The decrease in FLIM-FRET efficiency with Bsg-RNAi signifies an increase in stretching of the talin sensor, as shown by the higher donor lifetime compared to controls. The above experiments used heterozygous copies of the talin sensors and represented a mixture of tagged and non-tagged talin protein. To check for the influence of potential interactions between sensor and non-sensor talin, homozygous expression of the F40 sensor was checked for FLIM-FRET efficiency (Figure 11I). Controls nerves showed 15.28% FLIM-FRET efficiency at the C-terminus and 20.04% at the N-terminus, whereas knockdown of Basigin led to significant reductions in FLIM-FRET (Figure 11I). With Bsg-RNAi, morphologically normal nerves showed 8.79% efficiency at the C-terminus and 10.87% at the N-terminus, whereas ruffled areas showed 1.11% efficiency at the C-terminus and 3.07% at the N-terminus. Therefore, significant reductions in FLIM-FRET efficiency with knockdown of Basigin were observed with both the C-terminus and N-terminus insertion of the sensor. Since the same trends were observed for both homozygous and heterozygous insertions of the sensor, this data validated that heterozygous expression of the sensor is sufficient to measure FLIM-FRET in nerves. However, we noticed that roughly half the amount of FLIM-FRET efficiency was observed for any given genotype for the homozygous sensor as compared to the heterozygous F40 sensor (Figure 11H 31 compared to 11I). We hypothesize that since the homozygote possessed approximately double the amount of sensor as compared to the heterozygote, but was imaged using the same level of laser power and imaging time, this led to excitation of fewer donor fluorophores in the homozygotes leading to less measured FRET in the homozygotes as compared to the heterozygotes. We next assessed the least sensitive sensor (HPst, 9-11pN) as heterozygotes at both the C-terminus and N-terminus of the rod domain (Figure 12) and observed similar results to the F40 sensors. Controls had a FLIM-FRET efficiency of ~35% and knockdown of Basigin led to significant reductions in FRET. Bsg-RNAi showed 25% FLIM-FRET efficiency at the C-terminus and 31.1% at the N-terminus where nerves appeared morphologically normal. Ruffled areas showed 7.3% efficiency at the C-terminus and 8.3% at the N-terminus (Figure 12F). Overall, the decrease in FLIM-FRET with knockdown of Basigin indicated for both sensors an increase in the distance between fluorophores (or a decrease in the ability of the donor to transfer energy to the acceptor), to the point that in ruffled areas, the donor lifetime was no longer influenced by the acceptor mCherry. However, when we compared the FRET efficiency between the F40 and the HPst sensors, we noted significant differences in the apparent FRET efficiencies (Figure 12G) in the regions outside of the ruffles. Within the ruffles there were no differences between sensors since they were equally affected. However, in the morphologically normal regions, the F40 sensors in Bsg-RNAi nerves had significantly lower FRET efficiencies compared to the less sensitive HPst sensors (compare dark triangles to lighter triangles). In addition, loss of Basigin had a greater effect at the C-terminus (C-sensor) than the N-terminus (1-sensor) (compare blue triangles to green) for both F40 and HPst. Interestingly, wildtype controls had no significant differences between sensors with the exception of the C-terminus F40 sensor 32 which showed significantly lower FRET efficiency. Overall, these data signify that loss of Basigin causes talin to be stretched at both its N-terminus and C-terminus, presumably from pulling forces of proteins binding at either end of talin due to FA activation. The knockdown of Basigin in the PG causes a decrease in intermolecular FLIM-FRET We next hypothesized that the ruffle phenotype seen with Bsg-RNAi could also be caused by an increase in clustering of FAs. Clustering allows for an increase in proteins like talin to be recruited to FAs (Ellis et al., 2014; Goult et al., 2021). To test if the knockdown of Basigin increases the clustering of focal adhesions, the levels of intermolecular FLIM-FRET between talin proteins were measured. We used single fluorophore insertions of either the donor or the acceptor with both fluorophores at the C-terminus, both at the N-terminus of the rod domain, or with the donor at the N-terminus and the acceptor at the C-terminus. Since talin dimerizes at its C-terminus (Gingras et al., 2008), we predicted high FRET efficiency with both the donor and acceptor at the C-terminus. In contrast, we predicted low FRET efficiency with the donor and acceptor both at the N-terminus, since typical talin length is approximately 50-60nm without full extension of rod domains (Yao et al., 2016), and FRET can only occur between fluorophores up to 10nm apart (Ishikawa-Ankerhold et al., 2012) (Figure 13C, F, I). We also performed FRET between the donor at the N-terminus and the acceptor at the C-terminus and hypothesized an intermediate level of FRET compared to the C- and N-terminus combinations. The PG-specific driver 46F-Gal4 was used to drive Bsg-RNAi or UAS-LacZ as the control. We also performed a donor-only control at both termini to ensure that changes in the donor lifetime are dependent on the presence of the acceptor. Nerves from control (UAS-LacZ) showed normal morphology with higher levels of FRET in the interior of the nerve and lower levels of FRET at the exterior, 33 similar to the intramolecular FRET pattern (Figure 13A’’, D’’, G’’). Control nerves with the C-terminus donor/acceptor showed 18.29% efficiency, the N-terminus donor/acceptor showed 20.23% and the N-terminus donor/C-terminus acceptor showed 22.56% efficiency (Figure 13J). For these experiments, we also tested the responses of donor-only controls (as heterozygotes) with YPet inserted at the C-terminus and N-terminus and each showed between 20-23% efficiency (Figure 13J; circles with yellow middle). Of note, the apparent FLIM-FRET efficiency in the UAS-LacZ control for each fluorophore pairing (C-terminus donor/acceptor (blue); N-terminus donor/acceptor (green); N-terminus donor/C-terminus acceptor (pink)) was not significantly different from the respective donor alone (C-terminus (blue); N-terminus (green)) (Figure 13J). In addition, control animals showed no substantial differences in FLIM-FRET efficiencies between the different fluorophore combinations (Figure 13H). This suggests that each combination of fluorophore in control nerves was not sufficiently close to allow for FRET between the donor and the acceptor. However, as outlined in a following section, it is possible that our measurements with the donor only FLIM-FRET were false positives and thus our conclusions on intermolecular FRET may need to be revised. Due to the large size of the talin proteins and the estimated length of the unstretched rod domain being 50 nm (Yao et al., 2016), any observed FRET is not likely to be within the dimer as the donor and acceptor must be a minimum of 10nm apart (Ishikawa-Ankerhold et al., 2012). Any observed FRET is more likely caused by interactions between talin proteins within the dense meshwork created as the FA matures (Golji and Mofrad, 2014; Legerstee and Houtsmuller, 2021). Therefore, we tested the effect that loss of Basigin has on the intermolecular FRET pairs and surprisingly found that the knockdown of Basigin within the PG led to a decrease in FRET with each of the intermolecular donor/acceptor combinations. We assessed regions of nerves that 34 appeared morphologically normal as well as ruffled areas, and observed a decrease in the observed FLIM-FRET (Figure 13J). Ruffled areas in particular showed significant reductions in FRET with averages of ~2% FLIM-FRET efficiency with all donor/acceptor combinations. When visualized, the ruffle regions displayed warm colours indicating a longer donor lifetime and lower levels of FRET (Figure 13 B’’, E’’, H’’). For each of the pairs tested there were significant differences in the FRET efficiency in the morphological normal regions of the nerve with the N-terminus donor/C-terminus acceptor (pink) compared to the C-terminus donor/acceptor (blue) and the N-terminus donor/acceptor group (green) (Figure 13J). Within the ruffles, there were no significant differences between each fluorophore pairings with each displaying a significant reduction in FRET. Prior observations have found that there is a dense and extensive meshwork of proteins created in FAs under increased tension (Sigaut et al., 2018; Goult et al., 2021) including the recruitment of vinculin and extensive branched actin networks (Hu et al., 2016; Burridge and Guilluy, 2016). Thus, it is possible that the lifetime of each YPet donor in Bsg-RNAi animals is unaffected by the acceptor due to the tighter packing of proteins causing obstruction of energy transfer. Investigating the higher-than-expected donor alone FRET efficiencies The degree of FRET observed with the heterozygous donors alone was unexpected, ~20-23% efficiency with both at the C-terminus and N-terminus YPet insertions. Given that the intermolecular FRET combinations in UAS-LacZ control animals also showed this level of FRET efficiency, this led us to conclude that there is no intermolecular FLIM-FRET occurring in control animals. However, we wanted to further understand why each YPet donor was demonstrating substantial FRET efficiency in the absence of any acceptor. In reviewing our 35 imaging protocol, we discovered an issue in the experimental set-up of the donor and acceptor lasers when collecting FLIM-FRET data. Previously, when collecting FLIM-FRET data for both the intra- and inter-molecular FRET experiments (Figures 11-13), both the donor (YPet – 517 excitation) and acceptor lasers (mCherry – 587 excitation) were turned on. This meant that the samples were being excited with both lasers at the same time which led to the excitation of both the donor and the acceptor with consequences on the donor lifetime and the acceptor’s ability to receive energy. To test the impact on the measured FLIM-FRET, we collected new FLIM data on donor-only controls at the C-terminus and N-terminus with only the donor laser turned on and observed a consistent lifetime of 3.15ns for both YPet donors (Figure 14A), similar to the value observed by others using YPet inserted in talin (Austen et al., 2015; Ringer et al., 2017). Of note, when both the lasers were turned on, this significantly decreased the lifetime to 2.6ns (Figure 14A). Using the same samples, we then calculated the FLIM-FRET efficiency using the new protocol and observed very low baseline levels of FRET with each YPet donor (~2-3%) with only the donor laser on compared to when both lasers were on (~11.5%) (Figure 14B). This likely explains why high levels of FRET were observed with donor-only controls in our intermolecular experiments (Figure 13J). This also suggests that the intermolecular experiments need to be repeated with the new imaging protocol to ensure that our conclusions of no intermolecular FRET are based on the true donor lifetime. To test the effect of the revised imaging protocol on the intramolecular FRET measurements, we performed FLIM-FRET in wildtype animals using the F40 sensor at the C-terminus and N-terminus. These tests were carried out using homozygous animals, using a donor lifetime of 3.15ns with only the donor laser turned on. We observed ~13-15% FRET efficiency with both talin sensors, which closely matched the ~13-18% FRET efficiency when both lasers 36 were turned on (Figure 14C). These FRET efficiencies also closely matched those observed with the F40 sensors in the UAS-LacZ control (Figure 11I). Thus, the presence of the acceptor laser did not greatly affect the ability of the system to detect the proper FLIM-FRET efficiency, however, it did lead to a false positive signal in animals only expressing the donor. Overall, our conclusions that the loss of Basigin reduces the intramolecular FRET efficiency due to increased separation of the donor and acceptor pair in the sensors is likely true. However, the inter- and intra-molecular FLIM-FRET experiments should be repeated using the revised imaging protocol with utilizing only the donor laser. The knockdown of Basigin leads to an increase in vinculin As an alternate method to test for an increase in FA activation with the knockdown of Basigin, we imaged nerves to check for an increase in vinculin recruitment since the addition of vinculin signifies a mature, active focal adhesion (Humphries et al., 2007). Peripheral nerves were immunolabelled with an anti-Drosophila vinculin antibody along with the ECM using collagen IV (Viking) tagged with GFP and the perineurial glial membranes marked with mCD8::RFP. Control nerves showed low baseline levels of vinculin in the PG (Figure 15A-A’’’, white arrowheads) and a uniform distribution of collagen-IV (Viking) along the nerve. The ruffled areas seen with the knockdown of Basigin showed an increase in recruitment of vinculin (Figure 15B’’’-D’’’, white arrowhead) along with an increase in the presence of collagen-IV. We quantified these increases in the morphologically normal areas and ruffled areas. There was a significant increase in the fluorescence integrated density for collagen-IV for both Basigin-RNAi 1 and RNAi 2 (Figure 15 B’-E’, F, yellow arrowheads) compared to control (UAS-LacZ). An increase in the integrated density of vinculin was observed with RNAi 1, with an increase where 37 nerves appeared morphologically normal and a larger increase where nerves appeared ruffled (Figure 15G). However, Bsg-RNAi 2 did not show a significant difference, but as this experiment had only one replicate, further repeats are needed to determine if this trend holds true. Overall, ruffled nerves observed with Basigin RNAi showed an increase in collagen-IV and vinculin integrated density, suggesting an increase in focal adhesion activation and demonstrating the presence of mature FAs in the PG. 38 39 Figure 8: Talin is needed for proper formation of focal adhesions in the perineurial glia. A-B: Peripheral nerves with perineurial glia (PG) driver 46F-GAL4 driving expression of UAS-LacZ control (A) or talin-RNAi (B). Talin is endogenously tagged with YPet at the C-terminus of the protein (Talin::C-YPet, green) along with the PG membrane (UAS-mCD8::mCherry, red) and axons immunolabelled with mAB 22C10 (blue). Yellow arrowheads show localization of talin in the PG layer in UAS-LacZ, which is absent in talin RNAi. White arrowheads indicate asymmetric accumulation of PG membrane around PG nuclei on one side of the nerve whereas orange arrowheads demonstrate failure of PG to fully wrap the nerve. C-D: Peripheral nerves with perineurial glia (PG) driver 46F-GAL4 driving expression of UAS-LacZ control (C) or talin-RNAi (D). Talin is endogenously tagged with YPet at the N-terminus of the talin rod domain (Talin::1-YPet, green) along with the PG membrane (UAS-mCD8::mCherry, red) and axons immunolabelled with mAB 22C10 (blue). Talin is localized to the PG in control but is absent with talin RNAi (yellow arrowheads). White arrowheads indicate asymmetric accumulation of PG membrane around PG nuclei on one side of the nerve. E-F: Peripheral nerves with perineurial glia (PG) driver 46F-GAL4 driving expression of UAS-LacZ control (E) or talin-RNAi (F). Talin is endogenously tagged with YPet at the C-terminus of the protein (Talin::C-YPet, green) along with the PG membrane (UAS-mCD8::mCherry, red) and the integrin beta subunit βPS immunolabelled with an anti-βPS antibody (blue). Yellow arrowheads show co-localization of talin with βPS in the PG. Talin RNAi leads to absence of both talin and βPS in the PG. Orange arrowheads indicate failure of PG to fully wrap the nerve. G: Quantification of percent nerves showing failure of PG to wrap the nerve (purple) or an asymmetric accumulation of PG membrane on one side of the nerve (green) in either LacZ control or talin RNAi. Kruskal-Wallis test, Dunn’s multiple comparison test were performed (**** p<0.0001). n value shows number of animals. Scale bars 15µm. 40 41 Figure 9: Basigin co-localizes with focal adhesions in peripheral nerves and knockdown causes a ruffle phenotype. A: Peripheral nerve showing Basigin endogenously tagged with mCherry (Bsg::mCherry, red) along with talin tagged with YPet (Talin::YPet, green) and the integrin beta subunit βPS immunolabelled with an anti-βPS antibody (blue). Co-localization of the three proteins is indicated with white arrowheads. B-E: Peripheral nerves with the perineurial glia (PG) driver 46F-GAL4 driving expression of UAS-LacZ (B) or Bsg-RNAi (C-E). The ECM is labelled with Collagen IV (Viking::GFP, green) along with the PG membrane (mCD8::RFP, red) and the axons immunolabelled with 22C10 (blue). C-D show the ruffle phenotype observed with loss of Bsg (Hunter et al., 2020), including ruffling of the ECM and extension of PG protrusions off of the membrane (yellow arrowheads). E shows an example of a Bsg RNAi nerve region with normal nerve morphology. Scale bars 15µm. 42 Figure 10: FRET sensors inserted endogenously into talin are expressed in peripheral nerves. A: Two FRET (Förster resonance energy transfer) sensors, F40 and HPst are inserted into the talin gene, either at the C-terminus or at the N-terminus of the talin rod domain (Lemke et al., 2019). The F40 sensor linker between the donor (D – YPet) and acceptor (A – mCherry) is composed of a flagelliform protein and is sensitive to 1-6pN of force, whereas the HPst linker is a stable villin headpiece sensitive to 9-11pN of force (Lemke et al., 2019). Single fluorophore controls contain either the donor or acceptor at the C-terminus or N-terminus. Made in BioRender, adapted from Lemke et al., 2019. B: Peripheral nerve showing expression of the HPst sensor donor at the C-terminus of talin (Talin::YPet – green) and acceptor (Talin::mCherry – red). Axons are immunolabelled with mAB 22C10 (blue). White arrowheads indicate colocalization of the donor and the acceptor. Scale bar 15µm. 43 44 Figure 11: Basigin RNAi decreases intramolecular FLIM-FRET with F40 sensor in peripheral nerves. A-C: Peripheral nerve expression of the donor (YPet), the acceptor (mCherry) and FLIM (fluorescence lifetime imaging microscopy) of donor with C-terminus insertion of F40 sensor. C: C-terminus insertion of heterozygous FRET sensor in talin. D-F: Peripheral nerve expression of the donor (YPet), the acceptor (mCherry) and FLIM (fluorescence lifetime imaging microscopy) of donor with N-terminus insertion of F40 sensor. F: N-terminus insertion of heterozygous FRET sensor in talin. Examples of regions of interest (ROIs) assayed are illustrated in D’’-E’’ with dotted white lines where circle represents ROI for control (D”), star represents ruffle ROI and triangle is ROI of morphologically normal region in Bsg-RNAi (E”). The same symbols represent these ROIs in graphs H and I. G: Colour look up table: warmer colours indicate a longer donor lifetime (decrease in FRET signal) and cooler colours indicate a shorter donor lifetime (increase in FRET signal). H: Quantification of apparent FLIM-FRET efficiency (%) measured in ROIs with either the C-terminus (blue) or N-terminus (green) sensors in UAS-LacZ (control) or UAS-Bsg-RNAi lines driven with 46F-GAL4. In Bsg-RNAi nerves, ROIs were selected where nerves looked normal (triangle shape) and where the ruffle phenotype was seen (star shape). One-way ANOVA with Tukey multiple comparison test (** p<0.01; **** p< 0.0001). I: Quantification of FLIM-FRET apparent efficiency (%) with homozygous F40 sensor. Peripheral nerves expressing the C-terminus (blue) or the N-terminus (green) sensor in UAS-LacZ control) or in Bsg-RNAi driven by 46F-GAL4. In Bsg-RNAi nerves, ROIs were selected where nerves looked normal (triangle shape) and where ruffle phenotype was seen (star shape). One-way ANOVA with Tukey multiple comparison test (* p <0.05; ** p<0.01; *** p<0.01; **** p < 0.0001). Scale bar 15µm. 45 46 Figure 12: Basigin RNAi differentially affects the two FRET sensors. A-F: FLIM-FRET with heterozygous HPst sensor in peripheral nerves. A-D: Peripheral nerve expression of the donor (YPet), the acceptor (mCherry) and FLIM of donor. A-B: nerves expressing the C-terminal sensor. C-D: nerves expressing the N-terminus sensor. Examples of regions of interest (ROIs) assayed are illustrated in C’’-D’’ with dotted white lines where circle represents ROI for control (C”), star represents ruffle ROI and triangle is ROI of morphologically normal region in Bsg-RNAi (D”). The same symbols represent these ROIs in graphs F and G. E: Colour look up table: warmer colours indicate a longer donor lifetime (decrease in FRET signal) and cooler colours indicate a shorter donor lifetime (increase in FRET signal). F: Quantification of apparent FLIM-FRET efficiency (%) measured in ROIs across peripheral nerves with the C-terminus (light blue) or the N-terminus (light green) sensors in UAS-LacZ (control) or in UAS-Bsg-RNAi lines driven with 46F-GAL4. ROIs were selected where nerves looked normal (triangle shape) and where ruffle phenotype was seen (star shape). One-way ANOVA with Tukey multiple comparison test (**** p < 0.0001; ns – not significant). G: Comparison of the apparent FLIM-FRET efficiency (%) of F40 and HPst sensors with C-terminus (blue) or N-terminus (green) sensors in UAS-LacZ (control) or UAS-Bsg-RNAi lines driven with 46F-GAL4. Darker colours correspond to F40 and lighter hues to HPst. In Bsg-RNAi nerves, ROIs were selected where nerves looked normal (triangle shape) and where the ruffle phenotype was seen (star shape). One-way ANOVA with Tukey multiple comparison test (****p< 0.0001; *** p<0.001; ** p<0.01; * p<0.05; ns – not significant). Scale bar 15µm. 47 48 Figure 13: Basigin RNAi may decrease intermolecular FLIM-FRET in peripheral nerves. A-I: Peripheral nerve with UAS-LacZ control or Bsg-RNAi driven by 46F-GAL4. Channels include talin tagged with donor (YPet) or the acceptor (mCherry) and the FLIM channel showing the donor lifetime. In FLIM channel, warmer colours signify a longer donor lifetime indicating a decrease in FRET signal, whereas cooler colours signify a shorter donor lifetime leading to an increase in FRET signal (see colour lookup table below H). Cartoons illustrate the tagged-talin combinations. D – donor, A – acceptor. Talin diagrams made in BioRender. A-C: The donor and acceptor inserted independently at the C-terminus of talin. D-F: The donor and acceptor at the N-terminus of the talin rod domain. G-I: The donor at the N-terminus of the rod domain and the acceptor at the C-terminus of talin. J: Quantification of apparent FLIM-FRET efficiency (%) in peripheral nerve regions of interest in donor alone controls (open circle with yellow interior; blue exterior – C-terminus insertion; green exterior – N-terminus insertion), UAS-LacZ control (circle) and in Bsg-RNAi in ROIs with normal nerve morphology (triangle) or in ruffle regions (star). Blue data points indicate C-terminus donor and acceptor, green indicates N-terminus donor and acceptor and pink indicates N-terminus donor with C-terminus acceptor. One-way ANOVA and Tukey multiple comparison tests were performed (**** p< 0.0001; ns – not significant). Scale bars 15µm. 49 50 Figure 14: FLIM-FRET experiments should be performed by only exciting the donor. A: Donor lifetime data was collected for animals homozygous for the donor at the C-terminus (C-YPet; dark blue) or N-terminus (1-YPet; dark green) with only the donor laser turned on (YPet – 517 excitation) and the N-terminus donor was collected with both the donor and acceptor lasers turned on (light green) (mCh – mCherry; 587 excitation). (Kruskall-Wallis test, Dunn’s multiple comparisons; ** p-value = 0.0014; ns – not significant) B: FLIM-FRET efficiency was calculated with only the donor laser on for animals homozygous for the donor at the C-terminus (C-YPet; dark blue) or at the N-terminus (1-YPet; dark green), as well as with both the donor and acceptor lasers on with the donor at the N-terminus (light green) as both a homozygote and a heterozygote. The C-terminus donor was also collected as a heterozygote with the acceptor laser on (light blue). One-way ANOVA and Tukey’s multiple comparisons were performed (*** p-value <0.001; **** p-value <0.0001; ns – not significant). C: FLIM-FRET efficiency was calculated for animals homozygous for the F40 sensor with only the donor laser on at the C-terminus (C-F40; dark blue) or at the N-terminus (1-F40; dark green), as well as with both the donor and acceptor lasers on at the C-terminus (light blue) and at the N-terminus (light green). One-way ANOVA and Tukey’s multiple comparisons were performed (*p-value = 0.0016; ns – not significant). 51 52 Figure 15: Basigin RNAi causes increased recruitment of vinculin at focal adhesions in peripheral nerves. A-E: Perineurial glial driver 46F-GAL4 driving the expression of UAS-lacZ (A), Bsg-RNAi 1 (B-C), Bsg-RNAi 2 (D-E). The ECM is labeled with Collagen IV endogenously tagged with GFP (Viking::GFP, green), the perineurial glial membrane with mCD8::RFP (blue) and immunolabeling for vinculin (red). White arrowheads show expression of vinculin on perineurial glial membrane while yellow arrowheads show ruffling of the ECM and protrusions of perineurial glial membrane in Bsg RNAi. F: Quantification of integrated density of Viking::GFP (Collagen IV) fluorescence in control (UAS-LacZ, black circle) or in Bsg RNAi lines in regions that appeared phenotypically normal (red or purple circles) or where the ruffle phenotype was observed (star shape). One-way ANOVA with Tukey’s multiple comparisons were performed (**** p <0.0001). G: Quantification of integrated density of vinculin antibody fluorescence in control (UAS-LacZ, black circles) or in Bsg RNAi lines in regions that appeared phenotypically normal (red or purple circles) or where the ruffle phenotype was observed (star shape). One-way ANOVA with Tukey’s multiple comparisons were performed (***p=0.0004; * p=0.045). Scale bars 15µm. 53 54 Figure 16: Basigin RNAi leads to increased activation of focal adhesion complexes. Comparison between wildtype (top) and loss of Basigin (bottom) peripheral nerve focal adhesions connecting the perineurial glia (PG) to the extracellular matrix (ECM). Knockdown of Basigin in the PG leads to ruffling of the membrane caused by over-activation of integrin-based focal adhesions. Opposing forces acting on the membrane from excessive ECM binding as well as increased actin recruitment intracellularly lead to deformation of the membrane. The likely role for basigin in wildtype PG is to negatively regulate integrin activation to modulate interaction between the ECM and PG. Made in BioRender, diagram not to scale. 55 Discussion In the nervous system, the role of the conserved perineurial glia (PG) is still being elucidated, even though these cells are critical to animal survival. In peripheral nerves, the PG contact the extracellular matrix (ECM), otherwise known as the epineurium in vertebrates and the neural lamella in Drosophila. Focal adhesions are conserved protein complexes that connect the PG to the ECM in Drosophila, and facilitate communication through the cell layers to maintain proper nerve morphology through mechanotransduction and integrin signalling (Xie and Auld, 2011). My thesis investigated the role that the glycoprotein, Basigin, plays in modulating integrin activation in the PG and specifically, the hypothesis that loss of Basigin leads to an over-activation of integrin resulting in deformation of the PG membrane. Loss of Basigin in the perineurial glia causes an increase in integrin activation To test the hypothesis that the loss of Basigin phenotype is caused by an increase in integrin activation, we used FRET sensors to measure stretching of the talin protein as an indirect measure of focal adhesion activation (Cost et al., 2015). Under normal conditions we observed high levels of FLIM-FRET efficiency in wildtype animals with two sensors: the F40 sensor sensitive to 1-6pN and the HPst sensor sensitive to 9-11pN. With both sensors, we observed high levels of FRET in the interior of the nerve, likely due to low engagement of talin from a lack of ECM components. In contrast, low levels of FRET were consistently observed at the outer membranes of the nerve. Our results point to a strong PG/ECM engagement through integrins in vivo and mirrors results in vitro that found talin experiences an average of forces ranging from 5-10pN during cell adhesion (Austen et al., 2015; Yao et al., 2016). Prior work found that loss of the integrin beta subunit (mys) leads to disruption of the PG wrap surrounding the peripheral 56 nerve (Xie and Auld, 2011), and we confirmed that loss of talin leads to similar phenotypes (along with the presence of abnormal PG membrane processes). Talin and integrin complexes were distributed throughout the PG layers and loss of talin lead to the loss of integrin within the PG. Thus, FAs and in particular, talin, is engaged in wildtype nerves and is necessary for PG/ECM interactions. We confirmed that Basigin is expressed within the PG, and that loss of Basigin leads to changes in PG membrane including ruffling and increased recruitment of collagen-IV. Using both the F40 and HPst sensors, we observed a significant reduction in intramolecular FLIM-FRET with loss of Basigin. This indicated that the talin protein is being stretched, leading to an increase in distance between the donor and acceptor and thus a decrease in FRET signal. The resting talin protein is approximately 60nm long, but can be stretched up to hundreds of nanometers when activated and recruited to FAs (Margadant et al., 2011; Yao et al., 2016). In nerves that appeared morphologically normal, there was a significant increase in the stretching of engaged talin proteins, as indicated by the reduction in FLIM-FRET efficiency. Furthermore, ruffled areas showed the greatest reduction in FLIM-FRET efficiency, to the point where there was little to no FRET observed, suggesting that the donor and acceptor were separated beyond the 10nm FRET limit. This was also the case for the HPst sensor, which requires more force for separation. When we compared the FLIM-FRET efficiencies of the F40 and HPst sensors, we observed a significant difference between the two sensors with Bsg-RNAi even in regions where nerves appeared morphologically normal. At the C-terminus, the F40 sensor detected 15.77% FRET efficiency while the HPst was significantly higher with 25%. At the N-terminus, the F40 sensor detected 21.9% FRET efficiency while the HPst measured 31.1%. Since the F40 sensor is sensitive to forces ranging from 1-6pN whereas the HPst sensor is sensitive to greater forces of 57 9-11pN, a greater decrease in FLIM-FRET efficiency with the F40 sensor means that the talin proteins are more often experiencing small forces ranging from 1-6pN. This finding supports that the increase in integrin activation at FAs produces minute changes in force experienced by the PG membrane. As well, we observed significant decreases at both termini with either sensor, however larger decreases were observed at the C-terminus compared to the N-terminus, which indicated that the talin proteins are experiencing greater forces at the C-terminus. Furthermore, all sensors experienced the same significant decrease in FRET in the ruffled areas compared to controls, implying that both sensors maxed out, and the ruffled areas are experiencing a much larger range of forces of up to 11pN compared to where nerves appeared morphologically normal. We were surprised to observe the significant reduction in intramolecular FLIM-FRET with the C-terminus sensors since previous studies in talin tension using FRET sensors have shown lower tension at the C-terminus compared to the N-terminus (Austen et al., 2015; Kumar et al., 2016; Ringer et al., 2017; Lemke et al., 2019). We wondered if the heterozygous expression of the sensors were impacting the data due to interactions between sensor and non-sensor talin. However, the same trend was observed with animals homozygous for the tension sensor with significant decreases in FLIM-FRET with loss of Basigin. Therefore, we concluded that the significant reduction in FLIM-FRET observed with the C-terminal sensor was not caused by interactions between sensor and non-sensor talin. We hypothesize that the decrease in intramolecular FLIM-FRET observed with loss of Basigin at the C-terminus is likely caused by stretching forces but perhaps through a different means than the N-terminal sensors. The N-terminus sensor was inserted in the linker region connecting the head and rod domains of talin whereas the C-terminus sensor was inserted at the end of the rod domain where the dimerization 58 domain is found (Lemke et al., 2019). The linker between the head and rod domains stretches when external forces cause talin proteins to switch from autoinhibited states to active states (Goult et al., 2013; Yao et al., 2016; Kumar et al., 2016). In addition, the N-terminal region of the rod domain will experience stretching and tension as the head domain responds to conformational changes in integrin when the FA complex is active (Calderwood et al., 2013; Kanchanawong and Calderwood, 2023). Along the rod domain are multiple vinculin-binding domain sites where vinculin binds and recruits actin to the complex, leading to tension in the range of approximately 2.5pN on vinculin (Grashoff et al., 2010). Vinculin binding stabilizes talin in its unfolded conformation, contributing to the stretching of talin by up to 400nm (Margadant et al., 2011; Yao et al., 2014). Conversely, the C-terminus of talin possesses a dimerization domain that also contains a THATCH domain that can only bind actin when talin is dimerized (Gingras et al., 2008; Klapholz and Brown, 2017). Indeed, loss of ABS3 and the dimerization domain led to the loss of peripheral actin bundles localized to FAs (Austen et al., 2015). As well, FRET studies have shown that the C-terminus is more sensitive to myosin activity compared to the N-terminus (Ringer et al., 2017). High levels of myosin activity are associated with mature FAs with associated actin bundles (Geiger et al., 2009), and higher forces at the C-terminus of talin has been associated with increased myosin activity (Ringer et al., 2017). Therefore, talin experiences stretching across the entire length of the protein, not just at the N-terminus. Prior work on the same talin tension sensors (Lemke et al. 2019) observed similar levels of FRET efficiency at muscle attachment sites in Drosophila as we recorded in wildtype peripheral nerves. However, they observed consistently higher levels of FRET with the C-terminus sensors as compared to those at the N-terminus, which indicated that the talin protein was being stretched less at the C-terminus compared to the N-terminus, whereas we observed 59 greater stretching of the C-terminus compared to the N-terminus sensor. It is possible that the stronger reduction in FRET efficiency with our experiments at the C-terminus is caused by the same effects observed with the intermolecular FRET (i.e. the potential increased recruitment of FA associated proteins). However, with loss of Basigin, our data supports a role for stretching of the C-terminal sensor given the differences between the F40 and HPst sensors suggesting that at least in the morphologically normal regions of the nerve that the C-terminal sensor is more sensitive to changes in FA activation. Overall, the change in FRET efficiency is likely a product of changes in forces at both ends of the protein, with potential changes in talin dimerization or actin recruitment at the C-terminus playing a larger role. To address the involvement of the C-terminal actin binding in mediating talin tension in the PG, we propose to perform an experiment where Basigin is knocked down in the background of talin mutations with alternations in actin binding. There is a mutation in the actin binding site 3 (ABS3) at the C-terminus of talin that inhibits the binding of actin. The rhea[KVK] mutation affects residues 2450-2452 within the dimerization domain and has been shown to lead to a decrease in actin binding of up to 40% in sedimentation assays and decreased actin binding at muscle attachment sites by 30% (Franco-Cea et al., 2010). As well, point mutations in the ABS3 of talin in cell culture assays have significant reductions in actin binding (Gingras et al., 2008). Austen and others performed an experiment in cell culture using the YPet/mCherry FRET pair to measure tension across talin at FAs (Austen et al., 2015). They mutated the C-terminal ABS of talin and found a slight reduction in the forces experienced across talin as well as a reduced ability to form actin bundles, although integrin activation was unaffected. As well, other FRET-based measurements in talin found that the ABS3 is important for maintaining tension at the C-terminus of the protein (Ringer et al., 2017). Therefore, we hypothesize that expression of the 60 rhea[KVK] mutation will rescue the decrease in FLIM-FRET with the C-terminus sensors when Bsg-RNAi is expressed, but will not impact to the same extent the decrease observed with the N-terminus sensors. Intermolecular FLIM-FRET measurements If loss of Basigin causes an increase in clustering of FAs at the membrane, we hypothesized that there would be an increase in intermolecular FLIM-FRET with Basigin knockdown as more talin proteins are recruited to the FA. Since talin is a large protein that spans up to 60nm in its resting state and up to 800nm when active (Yao et al., 2016), we hypothesize that any degree of intermolecular FRET we observed must be between neighboring talin proteins, and not within a talin dimer. However, we observed a decrease in intermolecular FLIM-FRET with loss of Basigin in regions that appeared phenotypically normal, and a significant reduction in FLIM-FRET where the ruffle phenotype was observed. This trend was observed with the three donor/acceptor combinations used: C-terminus donor and acceptor, N-terminus donor and acceptor, and N-terminus donor with C-terminus acceptor. Prior work in muscle attachment sites (Lemke et al., 2019) observed very low baseline levels of intermolecular FRET at the C-terminus and at the N-terminus. We observed ~18-23% FLIM-FRET efficiency in control nerves, however we also observed significant FRET with the donor-only control that showed ~20-23% efficiency at both termini. Our subsequent revision of the imaging protocol revealed that the significant FRET observed with the donor control was an artefact and our intermolecular FRET experiments will need to be repeated to make clear conclusions. However, there are two potential outcomes: 1) We observe clear intermolecular FRET between the donor and the acceptor tagged talin proteins. Given the large size of the talin domains, this becomes a potential measure of the 61 clustering of talin in the meshwork formed at the strongly activated FA. We hypothesize that the N-terminus donor/acceptor pair will have the least degree of FRET since the donor and acceptor will be separated by the greatest distance in the talin dimers. 2) We observe no intermolecular FRET suggesting that the distances between the fluorophore tagged proteins are too large. In this instance, we hypothesize that the decrease in the FRET signal with Basigin-RNAi is caused by the increased recruitment of FA proteins, leading to changes in the fluorescence lifetime of the donor. This is supported by the observation of similar decreases in FLIM-FRET efficiency with each of the donor/acceptor combinations regardless of the placement of the fluorophores. Indeed, hundreds of talin proteins can be recruited to a single FA (Margadant et al., 2011), which also bind hundreds of vinculin and actin proteins. Using cluster-detecting simulations, the predicted distance between talin proteins is 20 to 25 nm with an approximate density of 600 talin molecules/um2 in a mature FA (Fischer et al., 2021). Furthermore, fluorescence lifetimes of fluorophores can be influenced by various micro-environmental factors such as pH, temperature, local viscosity, refractive index and fluorophore density (Lakowicz, 2006; Vu and Arai, 2023). At high concentrations, fluorescence lifetime can increase due to photon re-absorption where the fluorescence lifetime remains unchanged but the measurable lifetime value is increased (Berezin and Achilefu, 2020). Thus, for our intermolecular FRET analysis in a hyper-active FA, as more proteins are recruited, the donor would be unable to see any acceptor, or as talin concentrations increase, the donor lifetime could increase and measurement of FRET efficiency decrease. We propose to repeat this experiment under our revised imaging conditions, with a donor-only control animal with and without Bsg-RNAi in the background to normalize the FRET signal. As well, FLIM data can be collected of the donor alone with Bsg-RNAi in the background to ensure that loss of Basigin does not influence the microenvironment of FAs leading to changes in the 62 donor’s lifetime. We can use this approach to test if the recruitment of untagged associated FA proteins or the increase in donor proteins alone affects the donor lifetime independent of the presence of the acceptor. Loss of Basigin promotes vinculin recruitment and ECM binding at glial FAs To further support the finding that loss of Basigin leads to an increase in FA activation and increased recruitment of FA associated proteins, we observed a significant increase in the integrated density of vinculin at phenotypically normal areas and ruffled areas with one of two Bsg-RNAi lines. Of note, the expression pattern of vinculin in loss of Basigin animals was more diffuse throughout the PG membrane at ruffled areas with overall brighter intensity, but in controls, formed discrete bright puncta at the surface of the membrane. We hypothesize that the increase in FA activation with loss of Basigin caused stretching on talin which led to decreases in FRET, which also consequently led to an increase in exposure of vinculin binding sites. When talin is unfolded under mechanical stress, this increases exposure of vinculin binding sites allowing for an increase in vinculin binding to stabilize talin in its unfolded conformation (del Rio et al., 2009). Vinculin also recruits actin to stabilize the FA under tension. This prediction is supported by findings in cell culture using FRET sensors in talin where tension across the protein was reduced in vinculin deficient cells, and the presence of vinculin supports a high-tension state in talin (Austen et al., 2015; Ringer et al., 2017). To test the recruitment of vinculin to talin and the influence of Basigin loss, we propose to perform FLIM-FRET between talin and vinculin under Bsg-RNAi conditions, with the prediction that there will be higher FRET between the two proteins with loss of Basigin compared to wildtype animals. We will use vinculin tagged with mCherry at its N-terminus (Kale et al., 2018) and test the association of vinculin with the YPet 63 tagged talin (in the N- and C-terminal domains). It is possible that the fluorophores will not be sufficiently close given the large size of the talin rod domain, so as an alternative approach, we propose to carry out a PLA analysis that can detect protein-protein proximity of 40nm or less. An alternative approach would be to test for a rescue of the ruffle phenotype with a loss of function mutation in vinculin. Although, preventing the increased recruitment of vinculin to ruffled areas could lead to its own phenotype, since the recruitment is likely a compensatory response and needed to stabilize the active adhesions under loss of Basigin conditions. With Basigin knockdown, we also observed an increase in the integrated density of collagen IV, which may represent an increase in ECM binding by integrin or an increase in ECM deposition due to the increase in FA activation. Given the increase in intensity, more ECM ligand may be recruited and deposited due to the activation of integrins (Kim et al., 2014; Kechagia et al., 2019). Alternatively, the increase in collagen could be caused by an increase in the surface area of the membrane caused by the ruffling. The ruffled areas show protrusions of the PG membrane enveloped by ECM. How loss of Basigin leads to the ruffles and the changes to the overlying ECM is not known. One possibility is that loss of Basigin leads to changes in ECM deposition given its involvement in ECM remodelling as an MMP inducer in tumor cells (Guindolet and Gabison, 2020). The stiffness and bulk of the ECM (fibronectin, collagens) has been shown to affect integrin clustering and the promotion of mature FAs, where stiffer substrates support stable, mature adhesions (Roca-Cusachs et al., 2012; Kim et al., 2014; Paszek et al., 2014; Yamada et al., 2022). As well, talin FRET sensors in cell culture have shown increased tension on stiffer substrates (Kumar et al., 2016; Ringer et al., 2017). Furthermore, ECM ligands including perlecan have been shown to decrease ECM stiffness, whereas collagen-IV and integrins promote stiffness of Drosophila basement membranes (Pastor-Pareja and Xu, 64 2011). Since we did not observe a significant increase in collagen deposition in the morphologically normal areas, it is more likely that the ruffles are a consequence of increased activation of integrin independent of changes in the ECM ligands. Prior work found that it was the extracellular domain of Basigin that interacted with integrin and was sufficient to rescue the Basigin loss of function phenotype (Hunter et al., 2020) and thus the ruffle could represent a disruption of tightly regulated integrin signalling. However, to test if increases in collagen deposition are the trigger (rather than the consequence) of ruffle formation, we propose to use a Viking loss of function mutant as a heterozygote to test if reducing collagen levels can rescue the ruffle phenotype. Furthermore, the ECM plays a key role during the embryonic morphogenesis of Drosophila trachea, where sinusoidal buckling of the membrane is observed when there is too much membrane expansion and/or decreased ECM elasticity (Dong and Hayashi, 2015). Tubular shapes including nerves, trachea, arteries, etc. rely on constant membrane stability to maintain their shape; disruption via changes in growth, surface tension, or pressure can lead to compromised stability (Hannezo et al., 2012). Therefore, disruption of nerve morphology with loss of Basigin is likely a consequence of changes in PG-ECM interaction mediated through FAs. The overexpression of Basigin has been linked to many cancerous cell types because of its ties to the degradation of ECM, leading to compromised basement membranes and unregulated cell proliferation (Muramatsu, 2016; Guindolet and Gabison, 2020). Interestingly, the overexpression of Basigin in the PG does not lead to a phenotype (Petley-Ragan, 2016), but the loss of Basigin leads to the ruffle phenotype in peripheral nerves, as well as shortening of the VNC (Hunter et al., 2020). We propose that Basigin negatively regulates integrin activation, either directly or indirectly, since a reduction in myospheroid rescues the ruffle phenotype (Hunter et al., 2020). In contrast, in hepatoma cells, Basigin interacts with and activates β1 65 integrins through trans interaction with the MIDAS (Li et al., 2012; Wu et al., 2015). However, Basigin-integrin interaction has not yet been solved in cis. Although, Basigin has been shown to interact in cis with many transport proteins such as MCTs and PMCA (Kirk et al., 2001; Schmidt et al., 2017). Furthermore, Basigin has been shown to chaperone CD98hc (CD98 heavy chain, SLC3A2; solute carrier) to the membrane in hepatoma cells, and together they play a role in the activation of β1 integrins as well as increasing cell proliferation through the PI3K/Akt signalling pathway via FAK phosphorylation (Feral et al., 2004; Li et al., 2012; Fei et al., 2014; Wu et al., 2015). Conserved interactions of Basigin may exist in Drosophila. For example, Drosophila has been shown to possess an orthologue of CD98hc that is associated with the growth of brain tumors in flies and is expressed in surface glia where it plays a role in regulating sleep patterns (Reynolds et al., 2009; Rebelo and Homem, 2023; Li et al., 2023). Therefore, it is a possibility that the ruffle phenotype observed with loss of Basigin in the PG is caused by failure of Basigin to deliver transporters to the membrane such as SLCs, MCTs or PMCAs, and loss of Basigin leads to a disruption in ion balance and/or signaling pathways that produce compounding effects that disrupt cell morphology (Kirk et al., 2001; Wu et al., 2015; Schmidt et al., 2017). A possible hypothesis is that lack of Basigin leads to failure to form the PMCA transporter, for which Basigin/Neuroplastin have been shown to be essential auxiliary subunits (Schmidt et al., 2017). Since PMCA moves calcium ions to the extracellular space, and extracellular Ca2+ has been shown to negatively regulate integrin activation (Luo et al., 2007), a lack of extracellular calcium could lead to integrin overactivation. An alternative explanation is that negative regulation of integrin activation represents a new role for Basigin specifically at focal adhesions in Drosophila. In a screen of mouse antibodies to proteins that associate with α3β1 integrins, it was observed that EMMPRIN and 66 integrin colocalized with one another, but mostly at cell-cell contacts and not at focal adhesions (Berditchevski et al., 1997). However, evidence in Drosophila suggests a potential involvement of Basigin at focal adhesions where it associates with integrins and plays a role in the rearrangement of the cytoskeleton (Curtin et al., 2005). As well, we have previously shown that Basigin co-localizes with integrins in the perineurial glia, and loss of a key FA protein, talin, partially rescues the ruffle phenotype (Hunter et al., 2020). Furthermore, the expression of Basigin and myospheroid mutations lead to strong dominant genetic interactions causing defects during dorsal closure in Drosophila (Reed et al., 2004). The evidence in this thesis suggests that Basigin indeed plays a role in regulating the activation of integrins, however, the mechanisms underlying this interaction are still unknown. Since Basigin plays a variety of roles at the cell membrane, it is possible that Basigin differentially regulates integrins depending on their subcellular localization. Overall, we propose a model based on increased integrin activation (Figure 16) that leads to changes in talin and FA protein complexes. We propose that loss of Basigin in the perineurial glia leads to an increase in the activation of integrin-based focal adhesions connecting the glia to the ECM. In the absence of Basigin, the integrin complex binds an excess of ligand in the ECM or becomes hyperactivated (or both), which results in the activation of talin and the recruitment of a dense meshwork of proteins including activated vinculin and branched actin filaments. With our talin sensors we have evidence that loss of Basigin leads to increased tension in talin leading to the increased recruitment of vinculin and ultimately leading to a ruffled phenotype of the PG membrane. It was previously shown that loss of function mutants in the beta integrin gene (mys) or talin (rhea) rescues the Basigin loss of function phenotype (Hunter et al., 2020). We propose to perform FLIM-FRET using the talin sensors with Bsg-RNAi and a mys loss of function 67 mutant. Our hypothesis is that a decrease in βPS will rescue the decrease in FLIM-FRET observed with loss of Basigin. As well, we plan to perform FRET between tagged βPS (YFP) and tagged Basigin (cherry), which will test for an interaction of 10nm or closer, compared to the 40nm distance previously demonstrated with PLA (Hunter et al., 2020). 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Thesis/Dissertation
eng
Zoology
Vancouver : University of British Columbia Library
Attribution-NonCommercial-NoDerivatives 4.0 International
Graduate
Auld, Vanessa
Understanding the role of Basigin in regulating Integrin activation at glial focal adhesion complexes in the peripheral nerves of Drosophila melanogaster
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