Science, Faculty of
Science, Irving K. Barber Faculty of (Okanagan)
DSpace
UBCO
Tabatabaee, Mitrasadat
Doctor of Philosophy - PhD
University of British Columbia
Growing evidence indicates that dysregulation of glutamate signaling between neurons and astrocytes leads to the onset and progression of brain tumors (glioma). The aberrant morphology of glioma cells from astrocytic origin (astrocytoma cells) with elongated processes is also emerging as an underlying cause of progression and therapy resistance of gliomas. Whether glutamate dysregulation conducts this atypical morphology of astrocytoma cells has not been explored. Since normal astrocytes respond to glutamate by filopodiagenesis and processes extension, observing aberrantly elongated processes in response to excessive glutamate in astrocytoma cells is conceivable; yet it requires detailed empirical support to make it translatable to therapies. In this dissertation, the role of glutamate receptors and contribution of L-type voltage-gated calcium channels (CaVs) in elongation and directional protrusion of the astrocytoma cell processes toward glutamate were investigated. Protrusion of cellular processes toward glutamate and an intracellular calcium rise were observed in U118-MG astrocytoma cells. α-Amino-3-hydroxy-5-methyl-4-isoxazolepropionic acid (AMPA), whose role in neuro-gliomal interactions has been reported, did not trigger filopodiagenesis in these cells. In contrast, use of selective agonists (kainate and phenyl-kainate) and an antagonist (CNQX) indicated that kainate receptors (KARs) contribute to the glutamate-induced morphological response. However, precisely releasing a KAR caged-agonist (DECM-PhKA) to the 3D-cultured cells showed that despite their role in processes extension, KARs do not contribute to the pathfinding and the directional motility towards glutamate. CaVs were also found to play a role in the glutamate-induced morphological response in U118-MG cells by using selective antagonists (nifedipine, verapamil and diltiazem). Interestingly, a real-time imaging probe (FluoBar1) revealed that the membranar density of CaV1.2 rises immediately upon exposure to glutamate. Results from application of selective ligands (gabapentinoids and antibody) suggested that an auxiliary subunit of CaVs, α2δ1, links glutamate sensing to the intracellular calcium rise and filopodiagenesis. Strong evidence for the role of α2δ1 was found in heterologous expression systems that acquired the ability to respond to glutamate upon transfection. Altogether, determining the involvement of ligand-gated KARs and voltage-gated CaVs for the first time in the morphology of astrocytoma cells paves the way for therapeutical purposes to prevent progression and malignancy of gliomas.
i GLUTAMATE INDUCED MORPHOLOGICAL RESPONSE IN ASTROCYTOMA CELLS by Mitrasadat Tabatabaee Ph.D. ISLAMIC AZAD UNIVERSITY (Tehran Science and Research) Tehran, Iran, 2010 A THESIS SUBMITTED IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF DOCTOR OF PHILOSOPHY in THE COLLEGE OF GRADUATE STUDIES (Biochemistry and Molecular Biology) THE UNIVERSITY OF BRITISH COLUMBIA (Okanagan) October 2021 © Mitrasadat Tabatabaee, 2021 ii The following individuals certify that they have read, and recommend to the College of Graduate Studies for acceptance, a thesis/dissertation entitled: Studying the morphological response induced by glutamate in astrocytoma cells submitted by Mitrasadat Tabatabaee in partial fulfillment of the requirements of the degree of Doctor of philosophy. Dr. Frederic Menard; I.K. Barber Faculty of science, University of British Columbia (Okanagan) Supervisor Dr. Mark Rheault; I.K. Barber Faculty of science, University of British Columbia (Okanagan) Supervisory Committee Member Dr. Andis Klegeris; I.K. Barber Faculty of science, University of British Columbia (Okanagan) Supervisory Committee Member Dr. Michael A Russello; I.K. Barber Faculty of science, University of British Columbia (Okanagan) University Examiner Dr. Keith Murai; Faculty of medical and health sciences, University of McGill External Examiner iii Abstract Growing evidence indicates that dysregulation of glutamate signaling between neurons and astrocytes leads to the onset and progression of brain tumors (glioma). The aberrant morphology of glioma cells from astrocytic origin (astrocytoma cells) with elongated processes is also emerging as an underlying cause of progression and therapy resistance of gliomas. Whether glutamate dysregulation conducts this atypical morphology of astrocytoma cells has not been explored. Since normal astrocytes respond to glutamate by filopodiagenesis and processes extension, observing aberrantly elongated processes in response to excessive glutamate in astrocytoma cells is conceivable; yet it requires detailed empirical support to make it translatable to therapies. In this dissertation, the role of glutamate receptors and contribution of L-type voltage-gated calcium channels (CaVs) in elongation and directional protrusion of the astrocytoma cell processes toward glutamate were investigated. Protrusion of cellular processes toward glutamate and an intracellular calcium rise were observed in U118-MG astrocytoma cells. α-Amino-3-hydroxy-5-methyl-4-isoxazolepropionic acid (AMPA), whose role in neuro-gliomal interactions has been reported, did not trigger filopodiagenesis in these cells. In contrast, use of selective agonists (kainate and phenyl-kainate) and an antagonist (CNQX) indicated that kainate receptors (KARs) contribute to the glutamate-induced morphological response. However, precisely releasing a KAR caged-agonist (DECM-PhKA) to the 3D-cultured cells showed that despite their role in processes extension, KARs do not contribute to the pathfinding and the directional motility towards glutamate. CaVs were also found to play a role in the glutamate-induced morphological response in U118-MG cells by using selective antagonists (nifedipine, verapamil and diltiazem). Interestingly, a real-time imaging probe (FluoBar1) revealed that the membranar density of CaV1.2 rises immediately upon exposure to glutamate. Results from application of selective ligands (gabapentinoids and antibody) suggested that an auxiliary subunit of CaVs, α2δ1, links glutamate sensing to the iv intracellular calcium rise and filopodiagenesis. Strong evidence for the role of α2δ1 was found in heterologous expression systems that acquired the ability to respond to glutamate upon transfection. Altogether, determining the involvement of ligand-gated KARs and voltage-gated CaVs for the first time in the morphology of astrocytoma cells paves the way for therapeutical purposes to prevent progression and malignancy of gliomas. Keywords: Astrocytoma, Glutamate signaling, Morphological response, Kainate receptors, Voltage-gated calcium channels, α2δ1 v Lay Summary Astrocytes are supportive cells in the brain that are essential for neuronal wiring. Growing evidence suggest that their uncontrolled reshaping is associated to the progression of brain tumors. This thesis describes how two membranal channels are involved in the morphology of transformed astrocytes in the progression of the brain tumors, which will pave the way for therapeutic approaches. We found that kainate receptors, that receive the neuronal signals, can trigger membranal protrusions in astrocytoma cells. Results also indicated the contribution of an electrically active protein (CaV) in this phenomenon. A component of CaVs (α2δ1) was revealed to be crucial to the cells reshaping in response to neurotransmitters. Taken together, our findings begin to explain the mechanism by which transformed astrocytes help tumors progression by their aberrant morphology. vi Preface This research was conducted at the University of British Columbia, Okanagan campus, under the supervision of Dr. Frederic Menard. Portions of the results described herein, have been submitted to peer-reviewed journals and presented at academic conferences. A majority of the work was performed in the chemical biology laboratory of Dr. Frederic Menard. Some experiments were conducted in the laboratory of Dr. Mark Rheault or Dr. Philip Barker, as well as at the Facility for Environmental and Biological Imaging (FEBI). A version of Chapter 1 of this thesis is in preparation as a review journal article for the journal of Molecular Neurobiology “Mitra Tabatabaee, Frederic Menard; Morphology of astrocytoma cells in the pathology of glioma; a review”. The results from Chapter 2 were submitted as two journal articles to ACS Chemical Neuroscience and Neurochemical Research and their preprints are available online: 1) “Zhenlin Tian#, Mitra Sadat Tabatabaee#, Simon Edemann, Julien Gibon, Frederic Menard (2020) Optical control of Ca2+-mediated morphological response in glial cells with visible light using a photocaged kainoid. ChemRxiv. https://doi.org/10.26434/chemrxiv.11888925.v1” and 2) “Mitra Sadat Tabatabaee, Zhenlin Tian, Julien Gibon, Frederic Menard, Aminooxadiazolyl kainic acid reveals that kainic acid receptors contribute to astrocytoma glutamate signaling, BioRxiv, 2021.01. 16.426948 https://doi.org/10.1101/2021.01.16.426948” Parts of Chapter 2 have been presented at the 12th Annual Neuroscience Meeting of Canadian Association of Neuroscience, 2018, Vancouver, Canada: “Mitra Tabatabaee, Kabilan Sakthivel, Keekyoung Kim, Frederic Menard, “Using 3D cell-printing to study astrocytes morphology changes in real time”. Findings of Chapter 3 have been published as two journal articles in Cellular and Molecular Neurobiology, and Molecular Biology and Imaging: 1) “Mitra Sadat Tabatabaee, Jeff Kerkovius, Frederic Menard* (2021) Design of an imaging probe to monitor real-time redistribution of L-type vii voltage-gated calcium channels in astrocytic glutamate signaling, Molecular Imaging and Biology. https://doi.org/10.1007/s11307-020-01573-x“ and 2) “Mitra Sadat Tabatabaee, Frederic Menard (2020) L-type voltage-gated calcium channel modulators inhibit glutamate-induced morphology changes in U118-MG astrocytoma cells, Cellular and Molecular Neurobiology, https://doi.org/10.1007/s10571-020-00828-z”. 3) Based on these publications, an invited commentary manuscript is in preparation for the journal of Cellular Signaling (Elsevier). A version of Chapter 3, was presented in the 13th Annual Canadian Neuroscience Meeting, 2019, Toronto, Canada: “Mitra Tabatabaee, Frederic Menard*, The role of voltage-gated calcium channels in astrocytic glutamate signaling.” Findings of Chapter 4 were presented at Cold Spring Harbor Lab (CSHL), Glia in health and disease 2020: “Mitra Tabatabaee, Fred Menard*, The α2δ1, a Calcium Channel Subunit or a Neuronal Signal Transducer?”. This chapter is in preparation for submission as a research article. viii Table of Contents Abstract ............................................................................................................................. iii Lay Summary ..................................................................................................................... v Table of Contents ............................................................................................................ viii List of Tables ................................................................................................................... xiv List of Figures ................................................................................................................... xv List of Abbreviations ....................................................................................................... xvii Acknowledgements ......................................................................................................... xix Chapter 1. Introduction .................................................................................................... 1 1.1 Morphology of astrocytes .......................................................................................... 4 1.1.1 Morphology of astrocytoma cells ................................................................. 5 1.2 Glutamate signaling in astrocytes ............................................................................. 7 1.2.1 Glutamate uptake by astrocytes .................................................................. 8 1.2.2 Glutamate-induced calcium signaling in astrocytes ..................................... 8 1.2.3 Glutamate release from astrocytes .............................................................10 1.3 Astrocytic glutamate signaling in health and disease ...............................................10 1.3.1 Glutamate and the biology of astrocytoma .................................................11 1.4 Glutamate receptors in astrocytes ...........................................................................12 1.5 Voltage-gated calcium channels in astrocytes .........................................................16 1.5.1 Voltage-gated calcium channels in pathology of astrocytoma .....................18 1.6 Glioma .....................................................................................................................18 1.7 Thesis overview .......................................................................................................21 1.7.1 Knowledge gap ..........................................................................................21 1.7.2 Hypotheses ................................................................................................21 1.7.3 General questions ......................................................................................21 ix 1.7.4 Significance ................................................................................................22 1.7.5 Methodology ..............................................................................................22 1.7.5.1 Cell lines.................................................................................................23 Chapter 2. Kainate Receptors Contribute to the Glutamate-induced Morphological Response in Astrocytoma Cells ............................................................................................. 25 2.1 Background .............................................................................................................26 2.2 Material and methods ..............................................................................................28 2.2.1 Cell culture .................................................................................................28 2.2.2 Transfection ...............................................................................................28 2.2.3 Chelating free Ca2+ .....................................................................................29 2.2.4 Stimulation of U118-MG cells with glutamate receptor agonists .................30 2.2.5 Treatment with a Kainate receptor antagonist ............................................30 2.2.6 Confocal live imaging .................................................................................31 2.2.7 Time-lapse monitoring: effect of KA versus AMPA .....................................32 2.2.8 Quantification of filopodia extension ...........................................................32 2.2.9 Ca2+ imaging with Fluo4-AM ........................................................................32 2.2.10 3D cell culture ............................................................................................33 2.2.11 Uncaging MNI-Glu optically to stimulate U118-MG astrocytoma cells ........34 2.2.12 Optimization of irradiation time to uncage Ph-KA and to elicit processes extension in U118-MG cells ...........................................................................................34 2.2.13 Uncaging DECM-PhKA optically to stimulate U118-MG astrocytoma cells .35 2.2.14 Locally releasing KAR agonists to stimulate 3D cultured cells ...................36 2.2.15 SDS PAGE and Western blotting ...............................................................36 2.2.16 Statistical analysis ......................................................................................37 2.3 Results ....................................................................................................................38 x 2.3.1 Glutamate induced processes extension in U118-MG cells but not U138-MG cells ...............................................................................................................................38 2.3.2 Glutamate-induced process extension in U118-MG astrocytoma cells depends on extracellular calcium. ..................................................................................39 2.3.3 U118-MG cells respond morphologically to glutamate and kainate but not to AMPA ……………………………………………………………………………………………41 2.3.4 CNQX inhibits the morphological response of U118-MG astrocytoma cells caused by kainate and glutamate. .................................................................................44 2.3.5 Phenyl kainic acid (Ph-KA) causes a morphological response similar to KA in U118-MG cells ...........................................................................................................46 2.3.6 Photocleavage of DECM-PhKA promotes filopodia extension and increases intracellular calcium in U118-MG astrocytoma cells .......................................................48 2.3.7 Locally releasing glutamate induces directional extension of filopodia in 3D cultured U118-MG cells, but not PhKA. .........................................................................51 2.4 Discussion ...............................................................................................................53 Chapter 3. L-Type Voltage-Gated Calcium Channels Contribute to Glutamate-Induced Morphological Response in Astrocytoma Cells ..................................................... 59 3.1 Background .............................................................................................................60 3.2 Material and methods ..............................................................................................64 3.2.1 Cell culture .................................................................................................64 3.2.2 Transfection ...............................................................................................64 3.2.3 Treatment with voltage-gated calcium channel antagonists ........................65 3.2.4 Monitoring the processes extension in time-lapse confocal imaging ...........66 3.2.5 Quantification of filopodia extension ...........................................................67 3.2.6 Statistical analysis ......................................................................................67 xi 3.2.7 SDS PAGE and Western Blotting ...............................................................68 3.2.8 Chemical synthesis of FluoBar1 .................................................................69 3.2.9 Immunocytochemistry ................................................................................69 3.2.10 Protein labeling with FluoBar1 ....................................................................69 3.2.11 Staining for colocalization test ....................................................................70 3.2.12 Image analysis for colocalization ................................................................70 3.2.13 Real-time monitoring of LTCC’s redistribution using FluoBar1 ....................71 3.3 Results ....................................................................................................................71 3.3.1 Blocking the pore-forming subunit of CaVs in U118-MG astrocytoma cells caused a significant decrease in their processes extension upon exposure to glutamate 71 3.3.2 Ligands for the α2δ1 subunit suppressed the glutamate-induced processes extension in U118-MG cells ...........................................................................................73 3.3.3 Imaging probe FluoBar1 is selective for CaV1.2 ..........................................74 3.3.4 Membranar density of CaV1.2 changed when U118-MG cells were exposed to glutamate ......................................................................................................................76 3.4 Discussion ...............................................................................................................78 Chapter 4. The α2δ1 Contributes to the Glutamate-Induced Morphological Response and is Independent from Voltage-Gated Calcium Channels ................................................ 83 4.1 Background .............................................................................................................84 4.2 Material and methods ..............................................................................................86 4.2.1 Cell culture .................................................................................................86 4.2.2 Transfection ...............................................................................................86 4.2.3 Treating cells with 2δ1 ligands ...................................................................87 4.2.4 Confocal imaging .......................................................................................87 xii 4.2.5 Calcium imaging with Fluo4-AM .................................................................88 4.2.6 Quantification of processes extension ........................................................88 4.2.7 Immunocytochemistry (ICC) to label α2δ1 ..................................................88 4.2.8 FluoBar1 staining to label CaV1.2 ...............................................................89 4.2.9 Image analysis ...........................................................................................89 4.2.10 SDS-PAGE and Western blotting ...............................................................90 4.2.11 Statistical analysis ......................................................................................90 4.3 Results ....................................................................................................................91 4.3.1 Chemical and immunological ligands for α2δ1 inhibited the glutamate-induced morphological response in U118-MG astrocytoma cells ...................................91 4.3.2 Glutamate induced a change in the distribution and localization of α2δ1 on the membrane. ..............................................................................................................92 .................................................................................................................................93 4.3.3 Overexpressing α2δ1 significantly enhanced the response of U138-MG cells to glutamate stimulation .................................................................................................94 4.3.4 COS-7 cells responded to glutamate when they transiently expressed α2δ1 ……………………………………………………………………………………..95 4.1 Discussion ...............................................................................................................98 Chapter 5. Conclusion ................................................................................................. 102 5.1 Summary of findings .............................................................................................. 103 5.1.1 U118-MG astrocytoma cells demonstrate a directional motility towards glutamate ……………………………………………………………………………………104 5.1.2 KARs contribute to the glutamate-induced morphological response in U118-MG astrocytoma cells .................................................................................................. 104 xiii 5.1.3 CaVs contribute to the glutamate-induced morphological response in U118-MG astrocytoma cells .................................................................................................. 105 5.1.4 The α2δ1 controls filopodiagenesis upon exposure to glutamate ............. 106 5.2 Future research direction ....................................................................................... 106 5.3 Implications and significance ................................................................................. 108 Bibliography ..................................................................................................................... 110 Appendices ..................................................................................................................... 122 Appendix A: Scheme of synthesis of DECM-PhKA ..................................................... 122 Appendix B: Processes extension in response to glutamate is a faster response compared to KA, in U118-MG astrocytoma cells. ............................................................ 123 Appendix C: Synthesis of MNI-Glu .............................................................................. 125 Appendix D: Synthesis of FluoBar1............................................................................. 126 Appendix E: Rapid dot blot assay with FluoBar1 ......................................................... 127 Appendix F: Ligands for α2δ1 blocks the response to kainoids ................................... 130 xiv List of Tables Table 3.1 Calcium channel inhibitors and chemicals ............................................................66 Table 3.2 Colocalization analysis of confocal images.. .........................................................76 xv List of Figures Figure 1.1 Glia and the tripartite synapse……………………………………………………….….2 Figure 1.2 The role of perisynaptic astrocyte processes (PAPs) in the synapse formation……………………………………………………………………………………………….….3 Figure 1.3 Astrocytic glutamate signaling………………………………………………………..…9 Figure 1.4 Glutamate receptors…………………………………………………………………….14 Figure 1.5 Voltage-gated calcium channels (CaVs)……………………………………………...17 Figure 2.1 U118-MG cells respond to glutamate but not U138-MG cells…………………...…38 Figure 2.2 The morphological response of U118-MG cell to glutamate depends on extracellular calcium…………………………………………………………………………………….40 Figure 2.3 KA, but not AMPA, triggers filopodiagenesis in U118-MG astrocytoma cells…………………………………………………………………………………………………….…42 Figure 2.4 An antagonist for kainate receptors inhibited the morphological response of U118-MG cells both to KA and glutamate………………………………………………………………...…45 Figure 2.5 Ph-KA induced a morphological response in U118-MG astrocytoma cells comparable to KA………………………………………………………………………………...……..47 Figure 2.6 Photocleavage of DECM-PhKA promotes filopodia extension in U118-MG astrocytoma cells…………………………………………………………………………………...…...49 Figure 2.7 Photocleavage of DECM-PhKA causes an intracellular Ca2+ rise in U118-MG astrocytoma cells……………………………………………………………………………………......50 Figure 2.8 Optically releasing glutamate but not PhKA induces a directional extension of the processes in 3D cultured U18-MG cells………………………………………………………………52 Figure 3.1 Voltage-gated calcium channel subunit topology……………………………………62 Figure 3.2 Blocking the L-type calcium channels decreases the glutamate-induced processes extension in U118-MG cells……………………………………………………………….72 xvi Figure 3.3 Ligands for α2δ1 subunit of CaVs inhibited glutamate-induced filopodia extension in U118-MG cells………………………………………………………………………………………..73 Figure 3.4 Representative confocal fluorescence images of cells labeled with an anti-CaV1.2/DyLight550 antibody and FluoBar1…………………………………………………………..75 Figure 3.5 The density and distribution of CaV1.2 changed when U118-MG cells were exposed to 100 µM glutamate………………………………………………………………………....77 Figure 4.1 Structure of the α2δ1……………………………………………………………………85 Figure 4.2 antibody for α2δ1 inhibited the morphological response of U118-MG cells to glutamate………………………………………………………………………………………………...91 Figure 4.3 Exposure to glutamate changed the membranar density and localization of α2δ1 in U118-MG astrocytoma cells………………………………………………………………………...93 Figure 4.4 Morphological response and the calcium influx in U138-MG cells raised significantly upon transiently transfection with α2δ1………………………………………………...94 Figure 4.5 Transiently transfecting COS-7 cells with α2δ1 causes processes extension and intracellular Ca2+ rise in response to glutamate……………………………………………………...96 Figure appendix.1 Synthesis of DECM-PhKA scheme………………………………………...122 Figure appendix.2 In U118-MG astrocytoma cells, processes extension in response to glutamate is a faster response compared to KA……………………………………………………123 Figure appendix.3 Synthesis of MNI-Glu scheme………………………………………………125 Figure appendix.4 Synthesis of imaging probe FluoBar1………………………………….…..126 Figure appendix.5 FluoBar1 shows affinity for both CaV1.2 and CaV1.3………………..……129 Figure appendix.6 Ligands against α2δ1 blocked the morphological response that is induced by kainoids in U118-MG astrocytoma cells………………………………………………………....131 xvii List of Abbreviations [Ca2+]i Intracellular calcium concentration AMPA α-amino-3-hydroxy-5-methyl-4-isoxazolepropionic acid ANOVA Analysis of variance BAPTA-AM (1,2-bis(o-aminophenoxy)ethane-N,N,N′,N′-tetra acetic acid (acetoxymethyl ester) BK channel Big potassium channel Cache domain Calcium-chemotactic domain CaV Voltage-gated calcium channels CNQX Cyanquixaline (6-cyano-7-nitroquinoxaline-2,3-dione) CNS Central nervous system DB Dot Blot DECM-PhKA (2S,3S,4S)-3-(Carboxymethyl)-1-(((7-(diethylamino)-2-oxo-2H-chromen-4-yl)methoxy)carbonyl)-4-phenylpyrrolidine-2-carboxylic acid DMEM Dulbecco’s modified essential medium DMSO Dimethylsulfoxide EDTA Ethylenediaminetetraacetic acid EGTA Ethylene glycol-bis(β-aminoethyl ether)-N,N,N′,N′-tetraacetic acid ELFN1 Extracellular Leucine Rich Repeat and Fibronectin type III domain containing 1 Fluo4-AM Fluorescent calcium indicator 4- acetoxymethyl ester GABA Gamma aminobutyric acid GelMA Gelatin methacrylamide GIMR Glutamate-induced morphological response Glu Glutamate xviii GluK2 glutamate ionotropic receptor kainate type subunit 2 GluR Glutamate receptor GluT Glutamate transporter GPCR G protein-coupled receptor HBSS Hank's Balanced Salt Solution HEPES 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid ICC Immunocytochemistry iGluR Ionotropic glutamate receptor KA kainic acid LTCC L-type voltage-gated calcium channels mGluR Metabotropic glutamate receptor MNI-Glu (S)-2-Amino-5-(4-methoxy-7-nitroindolin-1-yl)-5-oxopentanoic acid NMDA N-methyl-D-aspartate (NMDA) NP 40 nonyl phenoxypolyethoxylethanol 40 PAPs Perisynaptic astrocytic processes PBS Phosphate-Buffered Saline PhKA Phenylkainic acid PKC Protein Kinase C PLC Phospholipase C SDS-PAGE Sodium dodecyl-sulfate polyacrylamide gel electrophoresis SEM Standard Error of Mean TSP Thrombospondin VGCC Voltage-gated calcium channels VWF-A Von Willebrand Factor A WB Western Blot xix Acknowledgements First and foremost, I must express my profound gratitude to my advisor Dr. Frederic Menard for his continuous support and his valuable guidance of this research work. I am also grateful to Dr. Mark Rheault and Dr. Andis Klegeris, members of my PhD advisory committee, for their valuable advices, useful critiques, and their support at each and every step of my Ph.D research. I would also like to thank all the current and past members of the Menard lab, particularly Dr. Zhenlin Tian, Jeff kerkovius and Dr. Vojtec Kapras, who without their hard work the synthetic chemistry side of the project, doing this research was much more challenging. I also need to thank members of the biofabrication lab of Dr. Kim for helping in optimization of the 3D cell culture technique. A special thanks to Dr. Julien Gibon for his advices and his help in calcium imaging experiments. I would also like to thank Dr. Andis Klegris, Dr.Philip Barker and Dr. Terry Snutch for generously gifting the cell lines and the plasmids, as well as Dr. Milani lab for creating and adapting the heating stage for the confocal microscopy. I would like to acknowledge the help of Jessica Lowry from the Center of Scholarly Writing for her assistance in the writing and structuring of this thesis. At the end, I would like to thank my family members for their patience, continues support and encouragements.1 Chapter 1. Introduction The central nervous system (CNS) is characterized by its rapid signal transduction and data processing. Neurons in the CNS use electrical signals in the form of action potentials to relay information rapidly. All other cells in the CNS that lack this property are categorized as glia. Neurons communicate with each other via specialized intercellular adhesion sites called synapses. Neuronal signaling typically starts with an action potential that propagates down a neuron's axonal process to a presynaptic terminal. Depolarization of this terminal leads to the release of neurotransmitters into the interstitial space between neurons: the synaptic cleft. Those chemical molecules bind to receptors on the postsynaptic membrane of another neuron. Subsequent depolarization of the second neuron propagates the signal further [1, 2]. Glia compose about 90% of the human brain and do not fire action potentials like neurons, instead they surround and ensheath neuron’s or synapses throughout the nervous system [1, 2]. Glia are classified based on their morphology, function, and location in the nervous system. In mammal’s CNS, glia are categorized as: microglia, astrocytes, oligodendrocytes and ependymal cells (Figure 1.1). They were thought for long to only provide the environment for neuronal-cell function [3]. However, accumulating data now suggests that glia, particularly astrocytes, are also involved in the active control of neuronal activity, synapse formation and synaptic neurotransmission ̶both during development and in adulthood [4, 5]. In a healthy brain a functional synapse consists of a pre- and postsynaptic neuronal element, as well as an astrocytic projection that envelops the synapse. This close spatial connection has led to the term tripartite synapse to acknowledge the astrocyte's contribution in a functional synapse (Figure 1.1) [3, 5]. Astrocytes are perfectly placed their fine processes at synapses to monitor — and respond to — synaptic activity; they express many of the same transmitters, receptors, and channels as neurons. 2 Perisynaptic astrocyte processes (PAPs) ensheath neuronal connections and assist synapse formation. They initially release some factors that form silent synapses [2]. Silent synapses contain all the structural elements of a synapse (such as presynaptic vesicles, active release sites, and postsynaptic density) and are presynaptically active because they release and reuptake synaptic vesicles. However, they are postsynaptically silent and cannot trigger action potentials [2, 4]. By releasing other signaling factors, PAPs activate postsynaptic glutamate receptors and control the functional synapse formation as well [2, 6, 7] As has been shown in hippocampus, the PAPs are also required for long-term potentiation to stabilize synapses (Figure 1.2) [8] Figure 1.1 Glia and the tripartite synapse. Microglia are the immune cells of the brain that interact with neurons and surrounding blood vessels. Oligodendrocytes create the wrap myelin sheath which enwraps axons and accelerate neuronal transmission. Astrocytes extend processes that contact blood vessels and ensheath synapses. Ependymal cells line the ventricles in the brain, which become filled with cerebrospinal fluid. A tripartite synapse is consisting of a presynaptic terminal, a postsynaptic terminal, and a presynaptic astrocyte process (PAP). 3 In fact, astrocytes not only interact with neurons but also with other cells such as neighboring astrocytes and vascular cells. At the neuronal interface, astrocytic fine processes regulate the precise neurotransmission of synaptic signals [1, 6, 9]. At the vascular interface, these processes control blood flow to deliver oxygen and glucose to the neurons in active regions of the brain [1]. They can also create a wide astrocytic network by connecting physically and chemically with other astrocytes (Figure 1.1) [10, 11] . Astrocytes owe their ability to communicate with different type of cells and several synapses simultaneously to their specific morphology [12]. They have numerous fine and highly motile processes, which can extend and reach the points of connection where their presence is required to modulate cellular activities [7, 12]. This specific morphology of astrocytes is very diverse and is determined by the cytoarchitecture of a given brain region [12]. Researchers believe that the complex morphology of human astrocytes is one of the main underlying reasons for the unique cognitive abilities of the human brain [13]. The mice that were transplanted with human astrocytes Figure 1.2 The role of perisynaptic astrocyte processes (PAPs) in the synapse formation. A tripartite synapse consists of the pre-synaptic and post-synaptic neurons as well as astrocytic processes, PAPs (green). PAPs respond to long-term potentiation (LTP) with structural changes. LTP increases the motility and retraction of PAPs and allows the postsynaptic dendritic spine to grow. While a dendritic spine remodels to create a strong connection, PAPs cover the synapses more tightly and stabilize LTP. 4 have been reported to display an improved long-term potentiation (LTP) and a better learning abilities [14]. These human astrocytes retain their size and morphology when proliferate in the mouse brain [13, 14]. The unique morphology of astrocytes not only plays an essential role for the brain’s health and function, it is also emerging to contribute to pathological states of the brain in conditions such as brain tumors [15–17] In the brain, glutamate is a common signal that both neuron and astrocytes can use in autocrine or paracrine manner to crosstalk [3]. Since glutamate is the major mediator of excitatory signals, there is a delicate regulation for its release, removal and recycling [18]. Astrocytes play the key role in this regulation. The role of astrocytes in glutamate regulation has been extensively studied and will be discussed in Section 1.2. Furthermore, glutamate can influence the morphology of both neurons and astrocytes. In neurons, glutamate regulates pathfinding by either inducing filopodiagenesis which steers direction of neuronal processes (neurites), or lamellipodiagenesis which leads to the whole cell migration and chemotaxis [19–22]. While processes extension in response to glutamate in astrocytes has also been documented a lot remains to be elucidated to complete our understanding about the molecular mechanism of this phenomenon [23–26]. Emerging roles of the morphology of astrocyte in neurological diseases such as brain cancers, is highlighting the importance of understanding the molecular mechanisms that direct this morphological plasticity. 1.1 Morphology of astrocytes Like other motile cells, astrocytes display membrane protrusions in the form of lamellipodia and filopodia [8, 21, 27]. Filopodia are rod-like cell surface projections filled with bundles of parallel actin filaments. They are found in many cell types and are involved in sensory or exploratory functions [21, 22, 24]. Filopodium formation is associated with Ca2+ elevations and protrusion of sheet-like actin filament arrays. The molecular details underpinning the initiation and maintenance of filopodia are just beginning to emerge [21, 28–30]. 5 Studies using live confocal imaging have demonstrated that astrocytes frequently extend and retract their perisynaptic processes, and subsequently enwrap spines tightly to form a functional synapse [11, 23, 26, 31]. This filopodial extension of astrocytes has been reported at glutamatergic synapses in the hippocampus, the brainstem and the cortex [17]. In the brainstem, two different motility modes have been reported. First, non-directed gliding of thin lamellipodia‐like process endings along neuronal surfaces. Second, directed filopodia‐like extensions budding off from primary or secondary processes. These filopodia with oriented motility are < 1 µm in diameter and 2–6 µm long and stayed elongated from 3 minutes to > 15 minutes [26]. These extensions and retractions have only been recorded at synaptic sites. They have never been observed in regions where astrocytes are in direct contact with neuronal soma or blood vessels. These directional motilities at synapses suggest their dependence on the signals that are released at the synaptic cleft [9, 32]. Since glutamate is known to induce processes extension in astrocytes [25], glutamate can be conceived as a shared signal between neurons and astrocytes at the synaptic sites that directs the motility of the astrocytic processes. In contrast to normal astrocytes, the extension and retraction of the processes in transformed astrocytes (astrocytoma cells) in brain cancers is not restricted to the synaptic sites. The brain cancers that are rose from glia are called glioma. When the glioma is originated from astrocytes, it is called astrocytoma [33]. Unlike normal astrocytes, astrocytoma cells can extend their processes to the greater distances and can make direct connections with other distant astrocytes or neurons [15–17]. 1.1.1 Morphology of astrocytoma cells Astrocytoma cells show ramified membrane and motile processes similar to normal astrocytes. In addition to the normal cellular projections, they also display an aberrant cellular morphology that plays a significant role in the pathology of the gliomas. Typically, the elongated astrocytoma cells and their projections create a fibrillary background in the brain tumors, that can keep the 6 tumor’s microenvironment secure from interventions and therapies like chemo or radiotherapy [15, 17, 34]. Moreover, the extension of astrocytoma processes into brain tissue facilitates the infiltration of malignant cells [16, 17, 35–37]. Astrocytoma cells make long and highly dynamic membrane protrusions that extend to distances far from their soma [16, 17]. During development, cell migration depends on long cellular protrusions that act as “pathfinders” in response to guidance cues [38, 39]. This phenomenon is similar to what has been shown during the progression of astrocytomas in the mouse brain [15]. The elongated membrane protrusions of astrocytoma cells have received different names, including membrane nanotubes, tunneling nanotubes, cytonemes and tumor microtubes [15–17]. Tumor cells use these long processes for invasion, proliferation, recurrence and interconnection of the single tumor cells to a functional communicating network [15]. In addition, these protrusions has been shown to play a role in creating complex and interconnected cellular communities which make the tumor resistant to the conventional therapies [16]. Interestingly, these long processes of astrocytoma cells have been recently shown to create a direct connection with neurons and to form functional glutamatergic synapses [17]. The neurogliomal synapses typically contain a cluster of neuronal presynaptic vesicles, a synaptic cleft, and a postsynaptic astrocytoma process, showing all the hallmark features of a glutamatergic chemical synapse [17]. The molecular mechanisms that control the formation of these long processes in astrocytoma cells as well as their guidance cue has not been explored. It is likely that astrocytes and astrocytoma cells use the same molecular mechanism for their process’s extension, but dysregulation in the signaling pathways in glioma has led to developing abnormal morphologies in astrocytoma cells. Identifying the molecules and receptors that conduct morphological changes in astrocytoma cells can introduce new therapeutical targets to intervene the tumor growth and its infiltration within the brain. 7 In fact, dysregulation in glutamate signaling in the brain, which will be detailed in Section 1.3.1, is emerging to be one of the main underlying mechanisms of glioma progression. Whether this uncontrolled glutamate signaling in gliomas causes to the aberrant morphology of astrocytoma cells is not known. Therefore, we hypothesized that the elongated extensions of astrocytic processes in glioma, which leads to the microtube formation, may be caused by wrong signals from dysregulated glutamate homeostasis. In this dissertation, we tested whether glutamate can induce a directional extension in the processes of astrocytoma cells. Throughout this dissertation, processes formation (filopodiagenesis) and extension in astrocytoma cells is addressed as morphological response. However, it should be noted that when astrocytoma cells migrate, they show a polarized morphology with a leading (front) and a lagging (back) poles in which lamellipodia play a more significant role than filopodia [21, 27, 28]. Lamellipodia are sheet-like membrane protrusions at the leading edge (front) of motile cells [28, 29]. Both filopodia and lamellipodia are composed of a dense and dynamic network of actin filaments and are devoid of cellular organelles [24, 28, 29]. 1.2 Glutamate signaling in astrocytes The nonessential amino acid, glutamate is the main excitatory neurotransmitter in the CNS and is a major bioenergetic substrate that is involved in biosynthetic, metabolic, and signaling pathways [40, 41]. At glutamatergic synapses, there is a precise control over glutamate signaling. Since glutamate is the major mediator of excitatory signals in the CNS, it should be released at right concentrations with a spatiotemporal precision [41]. Astrocytes govern the precision of glutamate signaling at a synapse by placing their fine processes and enwrapping the neuronal contacts [3, 8]. Astrocytes employ their membranal glutamate transporters and glutamate receptors as well as proteins that metabolize glutamate and links glutamate with calcium cascades to manage different aspects of glutamate signaling [4, 18, 42]. In general, glutamate 8 uptake, glutamate-induced calcium signaling, and glutamate release are the three main characteristics of glutamate signaling in astrocytes (Figure 1.3). 1.2.1 Glutamate uptake by astrocytes Fine processes of astrocytes use their glutamate transporters (GluTs) to control the diffusion of glutamate and prevent its spillover to seal neural connections [40, 43]. In addition, scavenging of glutamate at synaptic clefts by astrocytic GluTs prevents synaptic overstimulation and cytotoxicity [18, 41, 44]. Excessive glutamate stimulation causes further Ca2+ influx into neurons which leads to neuronal death and is called excitotoxicity [45]. In an astrocyte, the transported glutamate is converted to glutamine by glutamine synthetases [41, 46, 47]. The glutamine will be exported via glutamine transporters and returned to the nearby neurons [46]. In neurons, glutamine is converted back to glutamate by glutaminase enzymes and loaded into synaptic vesicles for another round of presynaptic release. This glutamate-glutamine cycle is an essential step in neurotransmission in the CNS. Glutamate transport in astrocytes is well-studied and its mechanism has been demonstrated [12, 46, 48]. 1.2.2 Glutamate-induced calcium signaling in astrocytes When glutamate binds to glutamate receptors (GluRs) in astrocytes (Section 1.4), it triggers a form of excitability that manifests by an elevation of intracellular Ca2+ concentration ([Ca2+]i) and follows by a morphological response [41, 42, 46, 49]. This Ca2+-dependent response to glutamate in astrocytes, which is called glioexcitability, is a relatively slow response to glutamate when is compared to the rapid signal transmission observed in neurons [4, 50]. Upon sensing glutamate, a Ca2+ transient spreads within the cell and is followed by a long-range Ca2+ waves within the astrocytic network that are conducted through gap junctions [25, 42]. Gap junctions are direct connections between the cytoplasm of two cells that allows various molecules ions and electrical impulses pass directly through a regulated gate between cells [51]. 9 Local application of glutamate to cultured astrocytes from both human and rat brain, has shown that Ca2+ waves can proceed up to a millimeter through the astrocytic network [42]. This [Ca2+]i cascade is triggered by the glutamate receptors that are either calcium-permeable (ionotropic receptors) or can release Ca2+ from internal stores (metabotropic receptors). Astrocytes have been reported to employ different types of glutamate receptors at different stages of their cellular development in distinct regions of the brain [32, 52–55]. However, the contribution of other Figure 1.3 Astrocytic glutamate signaling. Glioexcitability: Astrocytes demonstrate a form of excitability in their fine processes in response to neuronal glutamate by a calcium influx and processes extension. This Ca2+ wave can propagate through astrocytic network. Glutamate-glutamine cycle: Astrocytes uptake extra glutamate from the synaptic cleft to prevent excitotoxicity and recycle it to non-toxic glutamine, which is then released back to neurons to serve as precursor for the next round of neurotransmission. Gliotransmission: Transient elevations of calcium concentration in astrocytes lead to release of neuroactive chemicals, called gliotransmitters, that signal neurons or other astrocytes. 10 molecules and channels that may be involved in coupling glutamate signaling with calcium signaling in astrocytes has been underestimated. Section 1.5 of this chapter will describe how voltage-gated calcium channels were conceived to play a role in the Ca2+ dependent response that is induced by glutamate. This research focuses on the role of these channels in the morphological response caused by glutamate stimulation in astrocytoma cells. 1.2.3 Glutamate release from astrocytes Multiple independent studies have reported that signal transmission from astrocytes to neurons comprises several mechanisms. They include: gliotransmitter release via vesicular mechanisms, channel- and transporter-mediated mechanisms, as well as changes in neurotransmitter and ion uptake by the astrocytes [56]. Glutamate release from astrocytes was one of the first indications of astrocytic signal transmission, and it is still an actively studied topic [3, 4, 57]. Astrocytic signals such as glutamate, ATP, and D-serine have been reported to translate into different types of synaptic and network modulation. These astrocytic signals are involved in controlling presynaptic transmitter release, postsynaptic excitability, different forms of activity-dependent and tonic synaptic plasticity as well as in complex oscillatory network states [4, 31, 56]. The process of releasing chemical transmitters from astrocytes is collectively called gliotransmission [58–60]. Many of the above-mentioned mechanisms of gliotransmission are Ca2+-dependent (or at least sensitive to Ca2+ manipulations) and are involved vesicular neurotransmitter releases [56]. Due to longstanding debates over gliotransmission and the focus of this research, which is on the morphological response of astrocytes to glutamate, we will not discuss this phenomenon further in this dissertation. 1.3 Astrocytic glutamate signaling in health and disease As mentioned, in the CNS, neurons and astrocytes both release and receive glutamate as a common signal [5, 8, 51]. In a healthy brain, glutamate acts as the main excitatory 11 neurotransmitter that controls electrical activity of neurons by an ion influx and relays the signal to the adjacent neuron which eventually transmits throughout the neuronal network [40]. But in an astrocyte, glutamate triggers Ca2+ signaling which can initiate a cascade of intracellular events and can diffuse via gap junctions the astrocytes that are connected to its processes of that astrocyte [3, 4, 61]. A single mouse cortical astrocyte is estimated to contact over 100,000 synapses, whereas a human astrocyte can contact up to 2,000,000 synapses [2]. Consequently, a signal triggered by glutamate at a PAP can propagate throughout an astrocytic network which may transduce to the neurons that astrocyte processes are in contact with [61]. This fact suggests the range of impact that astrocytic glutamate signaling may have on the brain integrity and data processing. On the other hand, growing evidence show that the dysregulation of glutamate signaling in astrocytes play a significant role in brain disorders such as n epilepsy, neurodegenerative diseases and brain cancers [33, 36, 57, 62]. The transformation of astrocytes into malignant cells (astrocytoma) is known to be initiated by extracellular stimuli from surrounding microenvironments that sustain the function of both glia and neurons. However, the role of dysregulation in astrocytic glutamate signaling in shaping the microenvironments that support progression of glioma has been shown and will be discussed in the next section [33, 36, 62]. 1.3.1 Glutamate and the biology of astrocytoma As explained in Section 1.2 healthy astrocytes uptake the excessive glutamate at synapses to prevent spillover and excitotoxicity. In addition, astrocytes can also release glutamate to regulate the activities of adjacent neurons and synchronize the cellular events of nearby glial cells. Unlike normal astrocytes, astrocytoma cells do not efficiently uptake glutamate and their glutamate release rises significantly [36, 63–65]. The intracellular Ca2+ rise following activation of glutamate receptors in astrocytoma cells enhances their proliferation [17, 63, 66]. This glutamate-induced intracellular Ca2+ rise creates a positive feedback loop which amplifies the glutamate 12 release further from astrocytoma cells [36, 63]. On the other hand, the elevated glutamate concentration in the tumor’s microenvironment is fatal for peritumoral neurons. Overstimulation of glutamate receptors in peritumoral neurons can induce seizure as well as invasive excitotoxicity, which destroys neuronal tissues [43, 45, 63, 64]. The vacated spaces provide extra room for expansion of the proliferating tumor cells which results in tumor progression. Endeavors to explore the mechanisms in the pathology of glioma in which dysregulation of glutamate signaling in astrocytoma cells is connected to Ca2+ signaling are increasingly growing [17, 36, 66, 67]. However, it has not yet been translated to therapies for patients and highlights the remaining gaps in our knowledge. 1.4 Glutamate receptors in astrocytes Receiving an extracellular glutamate signal either in paracrine or in autocrine manner is followed by an intracellular Ca2+ rise in astrocyte. Literature studies shows that, studies have mainly focused on glutamate receptors that can either conduct Ca2+ or release it from internal stores to identify the link between glutamate sensing and [Ca2+]i rise in astrocytic glutamate signaling [2–4, 24, 52, 54]. It is now known that glutamate receptors that contribute to Ca2+ events following glutamate stimulation are heterogenous in different regions of the brain, and differ in the adult brains from the ones that are active during cellular development [12, 52, 55, 68]. All subfamilies of ionotropic glutamate receptors (iGluRs) have been reported to be expressed in astrocytes from different regions of the brain [3, 12, 18, 55, 68]. Ionotropic glutamate receptors are ligand-gated ion channels [40, 41]. They mediate excitatory neurotransmission and are essential to neuronal wiring. This family includes α-amino-3-hydroxy-5-methyl-4-isoxazolepropionic acid (AMPA), kainic acid (KA), and the N-methyl-D-aspartate (NMDA) receptor subfamilies—so termed for the chemical agonists that selectively activates them (Figure 1.4) [69, 70]. 13 AMPA and kainate receptors are usually pooled together into what is called non-NMDA receptors due to the structure and lack of distinctive antagonists for KARs to distinguish them from AMPARs [71–73]. The active role of AMPARs in astrocytic glutamate signaling has been demonstrated [3, 54]. Calcium-permeable AMPARs have been found to be expressed in astrocytes from the forebrain, the neocortex and in Bergmann glial cells of the cerebellar cortex [3]. The expression of calcium-permeable AMPARs has also been reported in glioma cells and seems to play an important role in glutamate-mediated proliferative signals [17, 33, 74]. Despite several indications about the activity and functions of AMPARs, very little information is available about the role of KARs in astrocytic glutamate signaling, although their expression both in astrocytes and astrocytoma cells have been reported [25, 74, 75]. Interestingly, hippocampal astrocytes do not express calcium-permeable AMPARs, instead, they express metabotropic glutamate receptors (mGluRs) [48, 52, 53]. Astrocytes from hippocampus exhibit metabotropic glutamate receptor 5 (mGluR5)–dependent rise in [Ca2+]i in response to glutamatergic transmission [53, 76]. mGluR5 is a type of G protein-coupled receptor (GPCR) that activates Gq and phospholipase C to release calcium from internal stores via an IP3-mediated pathway [48, 53]. While the involvement of mGluR5 holds true in the juvenile hippocampus, it was called into question in the adult brain, where mGluR5 agonists failed to induce calcium signaling in the astrocyte soma [3, 52]. In addition, glutamate release from mossy fibers of the mature hippocampus, can only partially induce calcium signals via mGluR5 in astrocytes [77]. The expression of mGluRs have also been also reported in gliomas, however, their specific role and function are still unknown [74]. It should be noted that astrocytes have been shown to express NMDARs at the mRNA and protein level as well [12, 55]. However, NMDARs are mainly inactive in astrocytes due to their voltage-dependent nature [3, 12, 78]. NMDARs are naturally blocked by Mg2+ and depolarization of the membrane is required to unblock the channel to permit Na+ and Ca2+ to pass through [18, 14 69]. But the membrane of astrocytes is highly polarized (resting potential ~ –80 mV) and NMDARs remain inactive [18, 78]. In many of the brain regions, the activity of NMDARs has been questioned and has not been recorded in the cultured astrocytes [3, 12]. However, NMDA receptors that are functional have been reported in cortical astrocytes [79]. Interestingly, glioma cells have also been shown to lack the NR1 subunit of the NMDA receptor, suggesting that NMDARs are not functional in glioma cells [74]. To date, research about glutamate signaling in astrocytes has tended to focus on glutamate receptors that increase cytosolic calcium ions in response to glutamatergic transmission. This cytosolic Ca2+ rise triggers several cellular events such as gliotransmission and cell proliferation have been extensively studied [4, 50, 80]. One of the consequences of glutamate stimulation in astrocytes is filopodiagenesis and processes extension [11, 81, 82], which has been less studied compared to the other Ca2+-Figure 1.4 Glutamate receptors. (A) Most glutamate receptors are ligand-gated ion channels (ionotropic glutamate receptors). AMPA and KA receptors conduct ions upon binding glutamate, but NMDA receptors require glycine or D-serine as well as glutamate to be activated. In addition, NMDARs are blocked by a Mg2+ in their inactive state, which requires an initial membrane depolarization to unblock the pore. (B) Fewer channels in the CNS are metabotropic glutamate receptors (mGluRs). mGluRs are GPCRs that conduct a secondary messenger cascade upon binding of glutamate. 15 dependent cellular events. The [Ca2+]i rise following glutamate stimulation can regulate the activity of actin-binding proteins that are involved in the dynamic turnover and reconstruction of the cytoskeleton in fine processes of astrocytes [24, 25, 83]. However, not much is known about the molecular mechanism of this response. In contrast to astrocytes, the molecular mechanism by which glutamate controls elaboration and protrusion of neurites have been studied in detail. For example, it has been shown in cultured Xenopus spinal neurons that neurites can turn and extend toward the source of glutamate to make a connection and form a synapse [20]. This morphological response to glutamate depends on the activation of the NMDA subtype of glutamate receptors and requires the presence of extracellular Ca2+ [20]. In another report in postnatal ventral midbrain cultures, dopaminergic axons have been shown to respond rapidly to glutamate stimulation with accelerated growth and growth cone splitting when NMDA and AMPA/kainate receptors were activated [84]. Interestingly, it was demonstrated that when AMPA/kainate receptors were selectively activated, axon growth rate was decreased. AMPA/kainate receptor activation alone elicited calcium signals that were solely attributable to calcium influx through L-type calcium channels. In contrast, stimulation of NMDARs, activates the ryanodine receptor-induced calcium release from intracellular stores. Thus, the authors suggested that a balance between NMDA and AMPA/kainate receptor activation is required to regulate the axonal arborization pattern of dopamine axons through the activation of competing calcium-dependent signaling pathways [84]. Understanding the mechanisms of dopaminergic axonal arborization is essential to understand molecular mechanisms that are involved in the Parkinson's disease for therapeutical approaches. In fact, the molecular mechanism of processes extension and elaboration in astrocytes has not attracted as much attention as in neurons because its pathophysiological significance has not been highlighted. But the emerging role of elongated processes of transformed astrocytes in 16 progression of the gliomas shed light in the importance of understanding the molecular mechanisms of this phenomenon. The activity of AMPARs at elongated processes of astrocytoma cells has been reported by Venkataramani et.al [17, 33]. However, that study focused on the cytosolic Ca2+ rise rather than the morphology of the astrocytoma cells upon exposure to glutamate. Whether the same glutamate receptors also control the aberrant morphology of these cells has not been examined. One of the objectives of this dissertation is identifying the glutamate receptors that are involved in the initiation of the morphological response in astrocytoma cells in response to glutamate. 1.5 Voltage-gated calcium channels in astrocytes Astrocytes express voltage-gated calcium channels (CaVs), despite being non-excitable [85, 86]. The first evidence for the expression of voltage-gated calcium channels (CaVs) in astrocytes was the discovery of Ca2+-dependent action potential in cultured cells [4, 85, 87]. Following the identification of CaV action potentials in cultured astrocytes, the propagation of calcium waves throughout the astrocytic network upon stimulation with the neurotransmitter glutamate was demonstrated [42, 87]. These two groundbreaking findings raised the possibility that CaVs contribute to the glutamate-induced Ca2+ waves in astrocytes, which could be considered as an extraneuronal signaling system in the CNS [4, 10, 42, 50]. However, failure to record the same type of calcium influx in astrocytes in vivo, combined with the complexity of astrocytic calcium signaling, hampered the interest to scrutinize the role of CaVs in the astrocytic glutamate signaling [52, 88, 89]. Typically, CaV channels couple changes in the membrane potential to a Ca2+ influx [90]. This [Ca2+]i rise serves in cellular ion homeostasis, and acts as a pivotal secondary messenger in events such as proliferation, cellular motility, and secretion [91]. L-type voltage-gated calcium channels (LTCCs), known as the CaV1 calcium channel family, were long considered to be exclusive to excitable cells [92]. However, they are now known to be the predominant type of CaV 17 in astrocytes [85, 86]. CaVs are classified based on the molecular variation in their pore forming subunit, α1, which associates with an internal, β, and a transmembrane, α2δ subunit (Figure 1.5) [67]. In fact , LTCCs have been shown to contribute to non-electrical phenomena that require a high volume of Ca2+ input, such as in the dynamic morphology of the cells. In neurons, the calcium hotspots caused by clustering of LTCCs have been shown to promote the morphological changes in neuronal growth cones [93]. In addition, voltage-gated calcium channels have been shown to Figure 1.5 Voltage-gated calcium channels (CaVs). (A) CaVs are classified based on their current conductance into five major groups: L, P/Q, N, R and T. Molecular analysis of the pore forming subunit, α1, classified them further into 10 subtypes. (B) Schematic of L-type CaVs (LTCC) assembly made of three subunits in neural cells. (C) Transmembrane topology of LTCC α1 subunit. It consists of 24 transmembrane segments which make four homologous domains of six segments each. The β subunit is bound to one of the to the internal loops of VGCCs and regulates trafficking of the channels to the membrane. α2δ subunit is tethered extracellularly. It modulates the channel’s conduction and its localization. Recently it has also been shown to play active role in and synaptogenesis. 18 contribute to the directed motility and pathfinding of the neurites and their antagonists inhibit this morphological response [94, 95]. Whether LTCCs play the same role in supplying Ca2+ to participate in morphological responses in the fine processes of astrocytes is not known. In this dissertation, we hypothesize that CaVs play the same role in extension of the processes of astrocytes as has been reported in neurites. Specifically, this hypothesis was tested in astrocytoma cells whose their processes formation and extension plays a critical role in the progression of tumors. 1.5.1 Voltage-gated calcium channels in pathology of astrocytoma The altered regulation of Ca2+ signaling, which actively modulates cellular proliferation, is one of the hallmarks of brain cancers [96, 97]. Over the last decades, therapeutical approaches have been used to adjust dysregulated Ca2+ signaling in glioma cells [98]. CaV antagonists are used as one of the effective drugs in treatment of gliomas. They have been shown to control their proliferation and infiltration within the brain [96, 97]. However, the molecular mechanism by which CaVs contribute to the progression and invasiveness of glioma is not fully understood. While a decrease in CaV expression levels has been reported in brain cancers [98], blocking CaVs has been shown to reduce glioma cell proliferation and infiltration in the brain [99–102]. However, CaV agonists neither enhanced the tumor growth nor induced a significant increase in the intracellular Ca2+ concentration [101]. How CaVs contribute to the tumor growth and progression is not known. Identifying the molecular mechanism of their effect will provide a better understanding about the biology of glioma and may pave the way fortherapeutical approaches. 1.6 Glioma Gliomas are the most prevalent primary tumors in the central nervous system [103, 104]. Histologically, gliomas share characteristics of normal glia and are typically named based on these similarities. The World Health Organization (WHO) has classified glial tumors into: 19 astrocytic tumors, oligodendroglial tumors, oligoastrocytic tumors, ependymal tumors, and neuronal and mixed neuronal-glial tumors (such as gangliogliomas) [105]. The more circumscribed tumors are classified under grade I tumors such as pilocytic astrocytomas, pleomorphic xanthoastrocytomas, and subependymal giant cell astrocytomas. The infiltrating gliomas include grade II (oligodendrogliomas and astrocytomas), grade III (anaplastic oligodendrogliomas, anaplastic astrocytomas, anaplastic oligoastrocytomas, anaplastic ependymomas), and grade IV (glioblastomas (GBM)). Grade I glioma have low proliferative potential, but the grades II through IV glioma are malignant tumors [36]. Due to aggressive diffusion features and primary and adaptive resistance against chemo- and radiotherapy and local recurrence after surgery, the average survival time in high-grade gliomas is 12–16 months [36] Invasiveness is a major determinant for malignant behavior in human gliomas. The classical way to determine the invasiveness of a tumor is injection of glioma cells into athymic mice, where histotypic tumors develop after a latency period [106]. In the last couple of years, large-scale sequencing analyses have provided an understanding of molecular changes in gene expression, copy number, somatic mutations, and epigenetic signatures in malignant gliomas [104]. Together, these studies have identified the role of the most common cancer-associated genes in the progression of glioma: TP53, PTEN, NF1, EGFR, ERBB2, RB1, PIK3R1, and PIK3CA [36, 104]. These genes are mainly involved in the dysregulation of growth factor signaling e.g. receptor tyrosine kinase (RTK)/phosphatidylinositide 3-kinase (PI3K)/Ras), p53, and Rb signaling pathways which lead to aberrant signaling in proliferation, cell cycle regulation, senescence, and apoptosis [36, 104]. In addition to the genetic transformations of glioma cells, disturbance in the glutamate homeostasis of the brain has also been shown to be one of the main factors in glioma progression [17, 62, 67, 107]. An almost complete absence of Na+-dependent glutamate uptake into gliomas has been reported [36, 63, 108]. This effect is due to a complete absence of EAAT2 transporters 20 in glioma cells as well as the mislocalization of EAAT1 to the nuclear membrane of these cells [108]. Interestingly, glioma cells release significant and excitotoxic quantities of glutamate into the extracellular space [107]. These findings suggests that glioma cells behaved opposite to astrocytes and increase the concentration of glutamate rather than sequestering it. The excessive glutamate release from glioma cells activates neuronal glutamate receptors and causes a sustained Ca2+ influx which results in excitotoxic cell death [36, 63]. These findings suggest that glioma cells use a programmed release of glutamate to promote expansion of tumors [63]. Another more recent discovery about the aberrant morphology of glioma cells with ultra-long, and highly functional membrane protrusions, particularly in astrocytomas, has expanded our knowledge about the mechanisms by which these tumors progress in the brain and how they resist conventional therapies [15, 109]. These membrane protrusions direct the brain invasion, proliferation, and interconnection of single cells to a network of astrocytoma cells that is resistant to therapies [15, 16, 109]. In fact, disconnecting astrocytoma cells by targeting their long protrusions has been conceived as a new avenue to reduce the treatment resistance of these tumors [109]. Therefore, understanding the molecular mechanisms that contribute to this specific morphology of astrocytoma cells would pave the way for therapeutical purposes. While the effect of dysregulation in glutamate signaling and the aberrant morphology of astrocytoma cells have both been established, whether glutamate directs the elongated processes of astrocytoma cells is not known. In this dissertation, two astrocytoma cell lines with different tumorigenic effect have been used to study this notion. These astrocytoma cell lines are immortal cells that have been derived from human malignant tumors and will be described in Section 1.7.5. 21 1.7 Thesis overview 1.7.1 Knowledge gap There has been tremendous progress in our understanding of glutamate signaling in astrocytes over the past few decades [3, 5, 25]. Growing evidence indicates that dysregulation in neuro-astrocytic glutamate signaling can trigger the onset and progression of glioma cancers [17, 63, 110]. Glutamate normally regulates the electrical activity of neurons and the Ca2+ signaling of astrocytes, yet can become a fatal factor for peritumoral neurons and a proliferative factor for cancerous cells (astrocytoma cells) in glioma [36, 62, 63, 66, 110]. On the other hand, astrocytoma cells exploit their atypical long processes to create an interconnected network of transformed cells which enhances their proliferation and infiltration [17, 35]. These elongated processes can also make the tumors resistant to conventional therapies such as radio and chemotherapies [15–17]. Whether these two phenomena, the dysregulated glutamate signalinFtreatg and the specific morphology of astrocytoma cells are related is unknown. This dissertation aims to identify whether glutamate serves as a signal that directs the extension of fine processes of astrocytoma cells. 1.7.2 Hypotheses • Glutamate serves as a cue for processes extension in astrocytoma cells and exerts its effect via the activity of a type of glutamate receptor in a Ca2+-dependent manner. • CaVs contribute to the morphological response that is induced by glutamate in astrocytoma cells. 1.7.3 General questions • Is glutamate a guiding cue for processes extension in astrocytoma cells? • Does glutamate exert its effect via a type of glutamate receptor? 22 • Do glutamate receptors play a role in the directional motility of the processes of astrocytoma cell towards glutamate? • Do L-type CaVs contribute to the morphological response that is induced by glutamate in astrocytoma cells? 1.7.4 Significance One of the factors that determines the rate of mortality in malignant gliomas is their resistance to chemotherapy, radiotherapy, and local recurrence after surgery. Elongated processes of astrocytoma cells have been shown to contribute to the resistance against standard treatment modalities in gliomas. Preventing aberrant processes extension in astrocytoma cells can control their infiltration into the brain tissues as well as the resistance of the tumors to the conventional therapies for cancer. Elucidating whether there is an interplay between glutamate signaling and this atypical morphology of astrocytoma cells, helps to identify previously overlooked physiological targets for therapeutical purposes. Determining the molecules that contribute to the aberrant morphology of astrocytoma cells can introduce target proteins to manage the invasiveness of glioma. 1.7.5 Methodology To simulate glutamate signaling, synthetic L-glutamate was used to mimic an incoming signal from adjacent cells to stimulate astrocytoma cells. The morphological response to glutamate was analyzed by measuring the change in the processes length and intracellular calcium rise upon exposure to glutamate using confocal and calcium imaging. An established astrocytoma cell line, derived from a glioma tumor, U118-MG cells, was used in this research. Other cell lines were also used as heterologous or control systems e.g. COS-7, U138-MG, MCF7 and HEK293. When applicable, synthetic chemical biology techniques were developed or optimized to create new tools to answer the questions of this dissertation. 23 1.7.5.1 Cell lines In this dissertation, two established human glioblastoma cells—U118-MG and U138-MG—were studied for their morphological response to glutamate. U118-MG and U138-MG were originally derived from different individuals (47-year old and 50-year old Caucasian men). They are very similar cytogenetically and share several derivative marker chromosomes [111, 112]. They are frequently used as astrocyte model cells in neuroscience and as cancer cells in oncology research. These two cell lines have been derived from malignant gliomas by J. Ponten and associates in 1966 [113]. The main differences between these two cells are their shape, doubling time and tumorigenesis. U118-MG cells are typical astrocytoma cells that demonstrate a mixed morphology with 33 hours doubling time and are tumorigenic. It induces tumor formation in nude mice inoculated subcutaneously within 21 days in 100% of the cases [111, 114, 115]. U138-MG cells are known as astrocytoma cells that are derived from glioblastoma and demonstrate a polygonal morphology with 50 hours doubling time and they are not tumorigenic, even in immunosuppressed mice. However, they can develop a tumor-like structure in a semisolid medium [112, 114]. Another cell line that has been used as a control cell line in this dissertation was COS-7 cells [116]. In contrast to U118-MG and U138-MG cells, COS-7 cells are not originally isolated from the human brain and are not naturally immortal cells. The COS-7 (CV-1 in Origin with SV40 genes) cell line has been developed by Prof. Yakov Gluzman in 1980s. It has been derived from the CV-1 African green monkey (Cercopithecus aethiops) kidney fibroblast and has been transformed by a mutant strain of Simian Virus 40 (SV40) [117]. The COS-7 cell line is a commonly used research tool and is an excellent expression system for DNA plasmid transfection experiments [117]. To the best of our knowledge, there is no report that addresses their glutamatergic events. COS-7 are CaV-free cells and are frequently used in studies about CaVs as heterologous expression system [118–121]. Therefore, these cells are good control cells for experiments presented in 24 Chapter 3 and Chapter 4 of this dissertation, where CaVs will be studied and their potential role in glutamatergic events will be investigated. HEK-293 cells and MCF7 cells that respectively express only CaV1.2 and CaV1.3 subtypes of CaVs were other cell types that used as a control for these channel’s expression in Chapter 3. The properties of these cell lines will be discussed further in the experiments that they have been employed. 25 Chapter 2. Kainate Receptors Contribute to the Glutamate-induced Morphological Response in Astrocytoma Cells Binding of glutamate to the glutamate receptors (GluRs) initiate intracellular signaling both in neurons and astrocytes [3, 8, 26]. Dysregulation of glutamatergic events in neurons and astrocytes is known to be associated with the development and progression of brain cancers [36, 62, 63]. In astrocytoma, glutamate drives the progression of tumors by inducing neuronal death while enhancing the glioma cells’ proliferation [33, 36, 63, 66, 100]. Excessive glutamate stimulation causes excitotoxicity and neuronal death and vacates space for the glioma invasion [17, 36]. In addition, glutamate has been shown to play a role in migration of astrocytoma cell and infiltration of cancerous cells within the brain [16, 64, 67, 110]. In addition to the significant role of glutamate signaling in initiation and progression of gliomas, morphological features of these cells are also emerging to play role in the progression and recurrence of glioma [16]. The specific morphological features in astrocytic tumors were reported first by Bailey and Cushing in 1926 [15]. However, until recently the significance of the cellular morphology of astrocytoma cells with their thin and abundant cellular processes had not gained much attention in neurooncological studies. These morphological characteristics aid the diffusion of gliomas throughout the brain and save them from conventional therapeutical approaches [15, 16, 67]. However, the molecules and pathways that are involved in these morphological features in astrocytoma cells has remained unexplored. As mentioned, it is known that glutamate induces processes extension and filopodiagenesis in normal astrocytes [3, 8, 25, 42]. On the other hand, aberrantly extended processes of astrocytoma cells are one the main features in invasiveness of brain tumors [109]. In addition, dysregulation in the glutamate homeostasis in the brain is another feature in progression of these tumors [36, 63, 66]. Therefore, we hypothesized that glutamate serves as a cue for processes extension in astrocytoma cells and exerts its effect via the activity of a type glutamate receptor in a Ca2+-26 dependent manner. Identifying the glutamate receptor that contributes to the morphological response of astrocytoma cells in response to glutamate can suggest pharmacological targets in controling progression and diffusion of malignant astrocytomas. 3D culture techniques combined with photo-pharmacology which gives control over precise spatiotemporal release of compounds with light indicated that U118-MG cells respond to and display a directional motility toward glutamate. However, a related type of glioma cell line, U138-MG did not demonstrate any significant morphological response to glutamate. Using selective agonist and antagonist, kainic acid receptors (KARs) were found to be required for initiation of the morphological response of U118-MG astrocytoma cells to glutamate. A novel caged agonist for KARs (DECM-PhKA) was created, to examine whether KARs contribute to this glutamate-induced oriented motility of the processes of astrocytoma cells, Photo-uncaging of the agonist triggered a glutamate-induced calcium signaling followed by processes formation and extension in U118-MG astrocytoma cells. However, no clear directional motility in the filopodia of 3D cultured cells in the Gel-MA matrix was detected, which suggests that KARs are not involved in the pathfinding of fine processes of U118-MG cells toward glutamate. 2.1 Background Astrocytoma stemms from astrocytes and is the prevalent malignant glioma in the brain [100]. Aggressive infiltration of malignant astrocytoma in the central nervous system (CNS) produces progressive and profound disability—ultimately leading to death in nearly all cases [33, 100, 122]. In fact, transformed astrocytes (astrocytoma cells) are found in almost all types of gliomas [103]. Glioma cells manipulate different elements of normal synapses to enhance their progression [33]. Alterations in glutamate signaling both in neurons and astrocytes are increasingly reported as an underlying cause of development and progression of gliomas [33, 36, 66, 100]. Glutamate has been shown to induce seizure and excitotoxicity by binding to peritumoral glutamate receptors of neurons [36, 66]. Neuronal death allows tumor cells to expand into the 27 space vacated by the damaged tissue [33, 79]. In addition, glutamate activates ionotropic and metabotropic glutamate receptors on glioma cells and initiates a cascade of intracellular events that can result in the proliferation of glioma cells [81, 123]. It has been shown in cortical astrocytes that metabotropic glutamate receptor 5 stimulates astrocyte proliferation via a protein kinase C-phospholipase D-phosphatidic acid-dependent pathway [124]. In addition, in glioma cells blocking AMPARs has also been shown to inhibit the proliferation of these cells, which suggest the contribution of AMPARs in proliferation of astrocytoma cells [125]. Glutamate can also create a positive feedback loop for further tumor growth by enhancing the Ca2+ signaling that leads to the extra glutamate release from astrocytoma cells [63]. Normal astrocytes from different regions of the brain are known to respond physiologically to glutamate via their ionotropic and metabotropic glutamate receptors, specifically, calcium-permeable AMPAR and mGluR5 [3, 18, 54]. However, whether transformed astrocytes—astrocytoma cells—employ the same receptors as normal astrocytes to initiate the Ca2+ transients and its consequent processes extension in the glutamate signaling is not known. In this Chapter, employing a combination of imaging, pharmacology, photochemistry, and 3D culture techniques, contribution of glutamate receptors in glutamate-induced morphological response of astrocytoma cells has been investigated. 3D cell culturing is a technique that allows one to observe the cellular responses that are limited in conventional 2D cultures. In this technique cells can freely move their projections and display their normal morphology [126, 127]. Caged compounds were used to stimulate 3D-cultured cells to examine the role of glutamate receptors in the directed motility of astrocytoma cells. Caged compounds are a type of photochemical tools in which a biologically active molecule is protected with a photochromic group that is cleaved upon light irradiation [60, 128, 129]. The activation of glutamate receptors with photocaged ligands was first demonstrated by Ellis-Davies—initially with the indoline MNI-Glu uncaged by UV light, then with coumarin DECM-Glu 28 uncaged with visible light [130]. While they are powerful tools, these molecules all release glutamate and do not allow the selective investigation of glutamate receptor subgroups [129]. Therefore, to examine a subgroup of ionotropic glutamate receptors, a selective caged agonist is required [128–130]. Several examples have been reported about caged NMDA and AMPA [128–133]. However, no study has been reported on developing caged KA. 2.2 Material and methods 2.2.1 Cell culture Human U118-MG cell line (ATCC HTB-15TM), and U138-MG (ATCC HTB-16TM) cells were cultured in Dulbecco’s modified essential medium (DMEM, Gibco 11995-065) supplemented with 10% v/v heat-inactivated fetal bovine serum (HyClone 12483020) 1% v/v penicillin 10,000 U/ml and streptomycin 10,000 μg/ ml (Gibco 15140-163). Cells were incubated at 37 ºC in a humidified atmosphere containing 5% CO2. For experiments, cells at 80-90% confluence were suspended using a pre-warmed 0.25% trypsin-EDTA solution for 5 minutes maximum (Gibco SH30236.02) and transferred to 35 mm glass bottom culture dishes (Mutsunami D1130H) in phenol red-free, high glucose DMEM supplemented with 25 mM HEPES (Gibco 21063-029). Enough cells were transferred to make a ~50% confluent culture dish after adhesion. The cells were incubated for less than their doubling time [84] for statistical analysis (see statistics section). This incubation period is long enough for cells to adopt a flattened shape after adherence which is required to obtain defined filopodia for measurements. 2.2.2 Transfection Fine processes of astrocytes are formed by the arrangement of the actin cytoskeleton [81, 83] . To observe morphological response of astrocytoma cells to glutamate, U118-MG and U138-MG cells were transfected with the F-actin marker mCherry-LifeAct-7 plasmid (gift from Michael 29 Davidson, Addgene #54491). Lifeact is a 17 amino acid peptide that binds to the actin cytoskeleton, and Lifeact does not interrupt cellular processes and actin polymerization [134]. When Lifeact is conjugated to a fluorophore molecule, e.g. mCherry, it allows us to observe and record the microfilaments rearrangement in live cells. Calcium phosphate co-precipitation technique was used to transfect the astrocytoma cells [135]. The DNA (1200 ng of plasmid) was mixed with calcium chloride (2.5 M, Sigma Aldrich, anhydrous 499609) in a buffered saline/phosphate solution (2X HEPES) to generate a calcium-phosphate–DNA co-precipitate, which will be engulphed by the cultured cells. The 2X HEPES buffer solution was prepared by mixing 0.8 g of NaCl, 0.027 g of Na2HPO4•2 H2O, and 1.2 g of HEPES in a total volume of 90 ml of distilled H2O. pH was adjusted to 7.05 with 0.5 N NaOH, and then adjust the volume to 100 ml with distilled H2O. The mCherry-LifeAct-7 plasmid employs the transcription machinery of the transfected cells and binds to G and F actin reversibly. To increase the expression level and the intensity of the fluorescence for imaging cells were transfected at least 72 hours before imaging. This overexpression caused saturation of mCherry signals in the cell's body in some images, however, it was necessary to observe the motility of the fine processes of astrocytes with adequate resolution. 2.2.3 Chelating free Ca2+ The free Ca2+ was chelated to identify whether the Ca2+ that supports the morphological response of U118-MG cells to glutamate is supplied from internal or external sources. Understanding the source of Ca2+ would help to determine whether the Ca2+ that is conducted through ionotropic glutamate receptors from the extracellular fluid supports the processes extension in response to glutamate; or it is the released Ca2+ from internal stores (via activation of metabotropic glutamate receptors) that contribute to the processes extension in U118-MG cells. 30 A set of cells were incubated for 3 hours with BAPTA-AM (Cayman chemical 15551) at the final concentration of 10 μM to chelate the internal free Ca2+ [87]. While, the other set of cells were treated with EGTA (Biobasic ED0077) at final concentration of 2 mM to chelate free Ca2+ in the media for 5 minutes before stimulation with glutamate [136]. Stock solutions of chelators were prepared in DMSO (vehicle) before each experiment. Solutions were filter-sterilized using a 0.22 μm syringe filter. To avoid the negative effect of DMSO on the cells, stock solutions were highly concentrated to maintain the final amount of DMSO 0.1% v/v in the media. To test the morphological response of the cells in absence of calcium, U118-MG cells were first incubated with a 10 µM BAPTA-AM in phenol-red free DMEM for 3 hours (intracellular calcium chelator). Then, 2 mM EGTA solution was added to the same culture dish for 5 minutes (extracellular calcium chelator) prior to time-lapse imaging morphological response of the cells to glutamate [137]. 2.2.4 Stimulation of U118-MG cells with glutamate receptor agonists On the microscope stage, U118-MG cells were stimulated with glutamate (Alfa Aesar A15031-30), AMPA (Sigma-Aldrich A6816-1MG), kainic acid (Tocris 0222), Phenyl kainate (Ph-KA, prepared in house) (E)-7-(Diethylamino)-4-(2-(dimethylamino)vinyl)-2H-chromen-2-one (DECM) and(2S,3S,4S)-3-(Carboxymethyl)-1-(((7-(diethylamino)-2-oxo-2H-chromen-4yl)methoxy) carbonyl)-4 phenylpyrrolidine-2-carboxylic acid (DECM-PhKA, made in-house). The stock solution for each compound was prepared freshly in deionized millipore water and filter-sterilized through a 0.22 μm syringe filter. Final concentration of all compounds was 100 µM, which consistent with the known effective concentration reported in the literature [25, 138]. 2.2.5 Treatment with a Kainate receptor antagonist U118-MG cells were treated with a selective antagonist against KARs to examine whether KARs participate in glutamate-induced morphological response. As a control, U118-MG cells in phenol red-free/high glucose DMEM supplemented with 25 mM HEPES were stimulated with 31 glutamate (Alfa Aesar A15031-30) and KA (Tocris 0222) on the confocal microscope stage. The stock solutions of L-glutamate and KA were prepared in deionized millipore water and filter-sterilized through a 0.22 μm syringe filter. Agonists were delivered to the medium four minutes after the beginning of imaging. The KAR antagonist CNQX (Abcam ab120017) was delivered to glass-bottom culture dishes (Mutsunami D11130H) for a final concentration of 10 μM. Stock solutions of CaV inhibitors were prepared in water (vehicle) before each experiment. Solutions were filter-sterilized using a 0.22 μm syringe filter. Cells were incubated with CNQX for 1 hour incubated at 37 ºC in a humidified atmosphere containing 5% CO2 before stimulation with KA and glutamate as described for control experiments. Cellular morphology and processes motility were considered as phenotypic indicators for viability of U118-MG cells. The viability of U118-MG cells after treatments with the antagonist was assessed based on the high motility of their processes as well the extended and flattened soma of the cells [111]. The cells that were under detrimental conditions displayed a shrinking and spherical soma as well as highly ramified membrane and non-motile processes. However, other viability assays could provide further information about the general metabolic health of the cells [139]. 2.2.6 Confocal live imaging Filopodiagenesis and extension of cells' processes were recorded on an Olympus FV1000 fluorescence confocal microscopy equipped with a Plan-ApoN 60x/i.4 oil-immersion objective. Changes in cell morphology were recorded at 60X maginification. To track the process's extension in time, an image was acquired (one second) at 20 seconds intervals. Using intervals and short exposure to laser reduced the photobleaching of the fluorescent signals and maximized cell survival during the course of the experiments. 32 2.2.7 Time-lapse monitoring: effect of KA versus AMPA Transfected U118-MG cells with mCherry-LifeAct7 were imaged with an Olympus FV1000 fluorescence confocal microscope equipped with a Plan-ApoN 60X/i.4 oil-immersion objective. The changes in process extension were recorded in time. To reduce photo bleaching, images were acquired at 20 seconds intervals with short 1 second acquisition time. U118-MG cells were imaged for 15 minutes prior to stimulation. The cells were first stimulated with AMPA (Sigma-Aldrich A6816-1MG) at a final concentration of 100 µM and imaged for 15 minutes; they were then further stimulated with KA at a final concentration of 100 µM and imaged for an additional 15 minutes. 2.2.8 Quantification of filopodia extension The length of twenty filopodia per cell was measured manually using ImageJ software as described by Zatkova et al. (NIH ImageJ, FIJI) [95]. Briefly, only filopodia whose entire arbor remained in the field of view throughout the imaging period and were in straight line were selected. A filopodium’s length was measured from the edge of the cell membrane to its apical end. The length of each filopodia was manually measured three times. To calculate the change in length (Δ length), a process's length from the image prior to stimulation was subtracted from the length of the same filopodia after 20 minutes. The reported Δ length for each cell is the averaged change in length of the 20 selected filopodia per cell. Measurements were done by an investigator blinded to the experimental conditions and to the expected results, while the experiments have been performed by the researcher who had designed the experiments. 2.2.9 Ca2+ imaging with Fluo4-AM U118-MG and U138-MG cells grown on a glass-bottom dish were incubated with 1.8 μM Fluo-4 AM (Invitrogen F14201 dissolved in less than 0.1% v/v of DMSO) for 10 min in a 5% CO2 incubator at 37 °C. Cells were then washed twice with phenol red-free DMEM (Gibco 11995-065) 33 and kept in the same incubator for 20 minutes to allow de-esterification. Before imaging, the media was replaced with HBSS (Lonza 10547F) supplemented with 2 mM Ca2+. Fields of view showing at least 10 cells were acquired using a 20x objective. Images were recorded at a rate of one frame every 5 seconds using an inverted epifluorescence microscope (Zeiss, Axio ObserverZ.1, controlled by ZEN2 software). Baseline fluorescence was recorded for 60 seconds and averaged (F0), followed by stimulation with kainoids or glutamate each at the final concentration of 100 μM. ImageJ software (NIH ImageJ, FIJI) was used to correct for background fluorescence for each frame: briefly, pixel intensity values of three background regions were averaged and subtracted from the mean pixel intensity of the whole frame. Changes in Fluo-4 fluorescence (F) as a function of time were expressed as F/F0. The change in the fluorescence intensity of Fluo4-AM upon stimulation with chemicals was calculated by subtracting the mean of the fluorescence intensity right before stimulation from the mean of the peak of the curve after stimulation. 2.2.10 3D cell culture The Gel-MA matrix for 3D culturing cells was synthesized in the Kim lab at the Department of Engineering UBC Okanagan as described by Wang et al [126, 127]. Gelatin methacrylate (GelMA), also referred to as gelatin methacryloyl, methacrylated gelatin and gelatin methacrylamide, is a natural hydrogel that is widely used in biofabrication. It is cost-effective, easy to synthesize and photo-crosslink, and biocompatible that allows cells to adhere and proliferate [126, 127, 140]. GelMA was used to 3D-culture U118-MG cells transfected with mCherry-Lifeact7 in a 35 mm glas- bottom culture dish (Mutsunami D1130H). The cell somas were encapsulated in porous structure of the hydrogel and their processes were free to move in response to environmental signals. Briefly, cells at 80-90% confluence were suspended using a pre-warmed 0.25% trypsin-EDTA solution for 5 minutes maximum (Gibco SH30236.02). Trypsin was quenched with DMEM and the suspended cells were centrifugated. 34 The cell pellet was mixed with warmed GelMA (37 °C) and transferred to 35 mm glass bottom culture dishes (Mutsunami D1130H). A 405 nm light was irradiated to crosslink the matrix. The 3D-cultured cells were incubated in Dulbecco’s modified essential medium (DMEM, Gibco 11995-065) supplemented with 10% v/v heat-inactivated fetal bovine serum (HyClone 12483-020) 1% v/v penicillin 10,000 Unit/ml streptomycin 10,000 μg/ ml (Gibco 15140-163) at 37 ºC in a humidified atmosphere containing 5% CO2. 2.2.11 Uncaging MNI-Glu optically to stimulate U118-MG astrocytoma cells (S)-2-Amino-5-(4-methoxy-7-nitroindolin-1-yl)-5-oxopentanoic acid, MNI-Glu, was prepared in our lab by Zhenlin Tian according to modified literature procedures (Appendix C) [60, 70, 131, 141]. MNI-caged glutamate rapidly and efficiently releases glutamate when photolysed (300 - 380 nm excitation). Before imaging, the media of the cells was replaced by HBSS (Lonza 10547F, 2 mM Ca2+) supplemented with 2 mM concentration of MNI-Glu. The final concentration of MNI-Glu was adapted from literature [128, 131, 133]. To release glutamate locally from MNI-Glu, an optic fiber (550 μm internal diameter) was used to irradiate 450 nm light. The light was produced by a laser pointer. The optical fiber was precisely aimed just outside a cell either 3D cultured (in GelMA as described in Section 2.2.9) or 2D cultured, with the aid of a micromanipulator. The light was kept on for 20 seconds and the motility of the processes of transfected U118-MG with mCherry-Lifeact-7 was recorded for 15 minutes. 2.2.12 Optimization of irradiation time to uncage Ph-KA and to elicit processes extension in U118-MG cells The caged kainoid (DECM-Ph-KA) was synthesized by Zhenlin Tian as described by Tian et al [60, 142]. The KAR agonist was released in the cell’s media by irradiating 405 nm light to DECM-PhKA [60]. An illumination-concentration curve was plotted to find the minimum irradiation time to release enough kainoid (Ph-KA) from DECM-PhKA that elicits the maximum cellular 35 response, Control experiments showed that 100 μM of Ph-KA induces processes extension in U118-MG cells, however, measuring the exact amount of released KA at the tip of the optic fiber in the media was impossible. The illumination-concentration curve was plotted based on processes extension in cells to optimize the irradiation time. HBSS solution (Lonza 10547F, 2 mM Ca2+) containing different concentrations (0.2 mM, 0.5 mM, 1 mM, 1.5 mM and 2 mM) of the DECM-PhKA was prepared. Before imaging, the media of the plated cells was replaced by various concentrations of DECM-PhKA supplemented HBSS. To release the kainoid locally in close proximity of target cell(s), the optical fiber was precisely aimed just outside a cell. With the aid of a micromanipulator 405 nm blue light was irradiated precisely to the media containing the caged-kainoid (DECM-PhKA) at the end of an optic fiber (550 μm internal diameter). The 405 nm light was produced by a commercial laser pointer (Eleksmark model FA02). In each field of view the successfully transfected U118-MG cells with mCherry-Lifeact, that expressed high fluorescence signal at their filopodia, were selected for imaging. To release the kainoid, the 405nm light was kept on for 10 seconds. At each of the EDCM-PhKA concentrations, if the process's formation and/or extension was not observed within 15 minutes after turning off the light, the irradiation time was increased by 10 seconds. The light was kept on till the process's formation and/or extension was observed. The duration of the light irradiation before initiation of filopodiagenesis (illumination time period) was recorded for the illumination-concentration curve. 2.2.13 Uncaging DECM-PhKA optically to stimulate U118-MG astrocytoma cells The kainoid (Ph-KA) was released precisely in proximity of the target cell as described in Section 2.2.10. Based on illumination-concentration curves, cells in HBSS solution (with 2 mM Ca2+) supplemented with 2 mM of DECM-PhKA were irradiated for 10 seconds. The 36 morphological response and Ca2+ influx in U118-MG after uncaging Ph-KA have been recorded. Change in the process’s length was quantified and reported as explained in Section 2.2.8. The change in the [Ca2+]i in U118-MG cells upon release of Ph-KA by light was recorded as described in Section 2.2.8. The baseline fluorescence was recorded before the application of light (405 nm) for 10 seconds and average (F0). ImageJ software (NIH ImageJ, FIJI) was used to correct for background fluorescence in each frame: the pixel intensity value of three background cell-free regions in each image was averaged, then subtracted from the mean pixel intensity of each uncorrected frame. The changes in Fluo-4 fluorescence as a function of time are expressed as F/F0, with F being the Fluo-4 fluorescence. 2.2.14 Locally releasing KAR agonists to stimulate 3D cultured cells To study directional motility of the processes, 3D cultured U118-MG astrocytoma cells were stimulated by locally photo-releasing PhKA and glutamate. The U118-MG cellstransfected with mCherry-Lifeact were 3D-cultured in GelMA as described in Section 2.2.10. The synthesis of the caged-KA (DECM-Ph-KA) and caged-Glu (MNI-Glu) has been described by Zhenlin Tian (Appendix C) [91, 92]. Glutamate and kainoid were released locally as described in Section 2.2.11 and 2.2.12 using an optic fiber. For the images presented in Figure 2.8.B, cells were fixed 15 minutes after the photorelease of glutamate and PhKA, and then imaged. The HBSS solution (supplemented with 2 mM concentration of MNI-Glu) was replaced with 4% paraformaldehyde in PBS solution for 20 min at the room temperature. The fixed cells were imaged with an Olympus FV1000 fluorescence confocal microscope equipped with a Plan-ApoN 60X/i.4 oil-immersion objective. 2.2.15 SDS PAGE and Western blotting Total protein content was collected from cultured U118-MG, U138-MG, and COS-7 cells. Cells were resuspended with prewarmed 0.25% trypsin-EDTA solution (Gibco SH30236.02), then transferred to a 15 mL Falcon tube, and centrifugated at 1000 rpm for 5 min at 4ºC. The pellet 37 was mixed with 50 μL of 1% protease inhibitor cocktail (Amresco, mammalian M250) in sterile Millipore deionized water. Then, 50 μL of NP40 lysis buffer (50 mM Tris-HCl, NaCl 150 mM, 1% NP40-Triton 100x pH 7.4) was added to the solution and incubated for 15 minutes. The mixture was centrifuged at 3000 rpm for 5 min at 4 ºC. The pellet was discarded, and the supernatant was stored at –20 ºC. Presence of a subunit of KARs, GRIK2, was determined by Western blotting. Electrophoresis at 120 mV for 60 minutes separated the native proteins in a 4% Tris-glycine gel based on their charge and the length of the polypeptide. The protein band were transferred electrophoreticly onto a Nitrocellulose membrane (Amersham Protran pore size 0.45 μm) and followed by an immunostaining procedure to visualize GRIK2 protein on the blotted membrane. The primary antibody used for Western blots was against GRIK2 (1:1000; AGC-009 Alomone). GRIK2 band was detected by chemiluminescence using the Amersham ECL kit (GE Healthcare) with horseradish peroxidase-conjugated secondary antibodies (GE Healthcare) [143]. The blots were analyzed solely qualitatively, and no normalization or quantification was done. 2.2.16 Statistical analysis Each value represents the average change measured from five individual cells, each from a separate culture dish (n = 5); the results are presented as the mean ± SEM. Significant differences among groups were determined using one-way or two-way analysis of variance (ANOVA) and post hoc Tukey’s multiple comparison was used to analyze the data analysis with GraphPad prism 7.04. A p value < 0.05 was considered significant. Statistically non-significant data are labeled the same on the plots. To minimize technical variations within samples for ANOVA analysis, a “completely randomized block design” was employed for testing each compound [144, 145]. The incubation time was kept shorter than the doubling time of U118-MG cells (33 hours) [114], which minimizes variation of cells within passages before exposing them to different treatments. 38 2.3 Results 2.3.1 Glutamate induced processes extension in U118-MG cells but not U138-MG cells U118-MG and U138-MG cells were used, to test if glutamate induces morphological response in astrocytoma cells Despite the high similarity between these two glioma cells. U118-MG cells extend their processes upon exposure to 100 µM glutamate but not U138-MG cells. When U118-Figure 2.1 U118-MG cells respond to glutamate but not U138-MG cells. (A) Representative confocal images of U138-MG and U118-MG astrocytoma cells exposed to 100 μM glutamate for 20 minutes. The cells were transfected with mCherry-lifeact, for live imaging of actin cytoskeleton (red). 60X magnification, 2X zoom; scale bar 5 μm (B) Change in length (Δ) was calculated by subtracting the length of a filopodium immediately prior to stimulation from that of the same filopodium after 20 minutes. Glutamate was dissolved in water (Vehicle). Data were analyzed using two-way ANOVA (F 1,24 treatment = 101.5, F 1,24 cell type = 105.2, for both factors p < 0.0001); Data were compared by a post hoc Tukeys’s multiple comparison. *p < 0.0001 significantly different from U138-MG cells that were stimulated with glutamate and non-stimulated cells. n = 5 (C) Normalized fluorescence of Ca2+ imaging dye Fluo4-AM in U138-MG and U118-MG cells stimulated with 100 μM glutamate, n = number of cells / number of dishes. Data are shown 39 MG cells were imaged in their medium without stimulation, their filopodia length changed by 0.28 ± 0.24 µm over the course of 20 minutes. Upon exposure to 100 µM glutamate, their processes grew by 4.82 ± 0.95 µm (17.2-fold) (p > 0.0001) (Figure 2.1). Unexpectedly, the change in the length of the processes of U138-MG cells after exposure to glutamate was not significantly different from the cells that were not stimulated with glutamate (Figure 2.1.B). It should be stressed that the only concentration that that the astrocytoma cells were exposed to was 100 µM, which is an established concentration at which cultured astrocytes extend their processes [25, 138]. While an intracellular Ca2+ rise was observed in both U118-MG and U138-MG cells, the change in intracellular Ca2+ rise in U118-MG cells was 33 folds more than what was recorded in U138-MG cells when both were exposed to 100 µM glutamate (Figure 2.1.C). The change in the fluorescence intensity of Fluo4-AM was 1.33 ± 0.08 AU/µm2 when U118-MG cells exposed to glutamate while this change was 0.04 ± 0.02 AU/µm2 in U138-MG upon glutamate stimulation. 2.3.2 Glutamate-induced process extension in U118-MG astrocytoma cells depends on extracellular calcium. To identify the source of Ca2+ that correlates with a morphological response to glutamate, U118-MG, intracellular and extracellular calcium were chelated seperately. The change in the filopodial length in unstimulated U118-MG cells were 0.28 ± 0.24 µm over the course of 20 minutes. When they were exposed to 100 µM glutamate, the change in their processes increased to 4.82 ± 0.95 µm which was significant (17.2-fold) (p < 0.0001) (Figure 2.2). Both process formation and extension were detectable within the first one to five minutes after glutamate exposure and many of the processes kept extending during the imaging period. However, the cells that their intracellular Ca2+ was sequestered with 10 μM BAPTA-AM before stimulation with glutamate demonstrated a significant decrease in their processes Δ length compared to untreated cells (p < 0.0001). The change in the length of the processes of BAPTA-40 AM pretreated cells upon exposure to glutamate was 1.7 ± 0.9 µm, compare to 4.82 ± 0.95 µm in the cells that were exposed to glutamate without any treatment. Yet, this extension was significantly different and more than the cells that were not stimulated with glutamate (0.28 ± 0.24 µm) (p < 0.05) (Figure 2.2.B). The only concentration at which intracellular Ca2+ was sequestered was 10 μM BAPTA-AM, that has been repeatedly reported to efficiently chelates intracellular Ca2+. Figure 2.2 The morphological response of U118-MG cell to glutamate depends on extracellular calcium. (A) Representative confocal micrographs of U118-MG astrocytoma cells exposed 100 μM glutamate. The cells were transfected with mCherry-lifeact, for live imaging of actin cytoskeleton (red). Cells were pretreated with chelators of intracellular Ca2+ (10 μM PABTA_AM) or extracellular Ca2+ (2 mM EGTA). 60X magnification, 2X zoom; scale bar 5 μm. (B) Change in length (Δ) was calculated by subtracting the length of a filopodium immediately prior to stimulation from that of the same filopodium after 20 minutes. Data were analyzed using two-way ANOVA (F 2,24 glutamate treatment = 34.98, F 2,24 Ca2+ chelators = 71.38, for both factors p < 0.0001); All the data sets were compared with a post hoc Tukey’s multiple comparison. *p < 0.0001 significantly different from the cells not exposed to glutamate, and the cells pretreated with Ca2+ chelator when exposing to glutamate. #p < 0.001 significantly different from the cells in the media with chelated extracellular Ca2+ when exposed to glutamate, as well as all the cells not exposed to glutamate. Data from the cells that were not exposed to glutamate and the cells that were pretreated with EGTA before exposure to glutamate were not significant. n = 5. Data are shown mean ± SEM. 41 It should be noted that higher concentration of BAPTA-AM might have more significant effect on the extent of processes extension in U118-MG cells. In contrast, the processes of the cells in the media with chelated Ca2+ (with 2 mM EGTA) did not demonstrate a significant change in their length compared to unstimulated cells. As expected, when both intra and extracellular Ca2+ were chelated and there was no free Ca2+ available, the change in the processes Δ length upon exposure to glutamate was comparable to unstimulated cells. The cell bodies shrunk continuously in time, and neither process extension nor motility was observed. 2.3.3 U118-MG cells respond morphologically to glutamate and kainate but not to AMPA The inhibitory effect of extracellular Ca2+ chelation on the process’s extension of U118-MG cells upon exposure to glutamate (Section 2.3.2) suggested that ionotropic glutamate receptors are more active in glutamate-induced response. Since in cortical astrocytes, NMDA cannot induce processes extension, and activity of NMDARs in astrocytoma cells is arguable due to the lack of its main subunit (NR1) [25, 146], the role AMPA/KA ionotropic glutamate receptors in glutamate-induced morphological response were examined. KA induced a significant change in the length of the processesof U118-MG cells’ comparable to glutamate. AMPA did not elicit the same response. KA and AMPA were both used at 100 μM to be consistent with effective concentration of glutamate. it should be noted that, higher concentrations of AMPA might have different effect, which remains for future investigation. When U118-MG cells were stimulated with 100 μM glutamate, the change in their processes length (Δ length) over 20 minutes was 4.82 ± 0.58 μm. This change was 21 times longer than t the cells that were not stimulated over the same time period (0.22 ± 0.09 μm). Application of 100 μM KA also induced a 15-fold increase (3.49 ± 0.42 μm) within 20 minutes in the processes length 42 of U118-MG cells compared to non-stimulated cells. However, the change in the processes length in the cells that were exposed to 100 μM of AMPA was 0.33 ± 0.17 µm, which was comparable Figure 2.3 KA, but not AMPA, triggers filopodiagenesis in U118-MG astrocytoma cells. (A) Cells extend their processes significantly in response to 100 μM of glutamate and KA. AMPA does not have the same effect. Change in length (Δ) was calculated by subtracting the length of a filopodium immediately prior to stimulation from that of the same filopodium after 20 minutes. Compounds were dissolved in water (Vehicle). The change in length is the average data of 5 cells, each from a separate culture dish; measurements of at least 20 filopodia were averaged for each cell (n = 5). Comparisons were done with one-way ANOVA. All the data sets were compared by a post hoc Tukey’s multiple comparison; *p < 0.0001 significantly different from cells exposed to AMPA. The change in the length of the cells that were exposed to AMPA was not significantly different from the control cells (vehicle); Data are shown mean ± SEM (B) Representative confocal micrographs of a U118-MG cell before and after consecutive exposure to 100 M of AMPA and then to KA. The cell was transfected with mCherry-lifeact, for live imaging of actin cytoskeleton (red). 60X magnification. 5X zoom. Scale bar 1 μm. (C) Averaged length of processes during a time-lapse imaging. 100 µM AMPA was added 15 minutes after beginning of imaging, and KA was added after 30 minutes to the same culture dish; The average size of filopodia of cells at the minute 30 (when KA was added to the media) was compared with their size at the minute 35 with an unpaired two-tailed T test. *p < 0.05 significantly different from the data point at the minute 30. n = 3, Data are shown mean ± SEM. (D) Western blot of U118-MG and U138-MG astrocytoma cells compared to COS-7 (as negative control) for GluK2 expression 43 with the non-stimulated cells (0.22 ± 0.09 μm). This comparison between the cells that were stimulated with glutamate receptors agonists and the cells that were not stimulated suggested that glutamate and KA can trigger filopodiagenesis in U118-MG cells but not AMPA (Figure 2.3.A). Data points on Figure 2.3.A show the change in the length as explained in Section 2.2.8. To confirm that 100 μM AMPA was not cytotoxic and to reduce the variation between samples, cells were stimulated sequentially, first with 100 μM AMPA and then with 100 μM KA. A time-lapse live imaging of U118-MG cells indicated that the cells that had not shown significant change in their processes length upon exposure to 100 μM AMPA for 15 minute (from 1.83 ± 1.0 μm to 2.0 ± 1.7 μm), extended their processes significantly and rapidly when exposed to 100 μM of KA (from 2.0 ± 1.7 μm to 8.33 ± 2.0 μm) (Figure 2.3.B and C). Data points on Figure 2.4.B show the average size of the filopodia at the indicated minutes on the X axis after stimulation. The average size of filopodia at minute 15:00 (when AMPA was added to the media) was compared with their size at minute 20:00 with an unpaired two-tailed T test. The size of the filopodia, 5 minutes after exposure to AMPA, was not significantly different from the point of stimulation. In contrast, when the same comparison was done between the data point at the minute 30 (when KA was added to the media) with the average size of filopodia at the minute 35, a significant rise in the size of the filopodia in 5 minutes after exposure to KA was observed (P value < 0.05). This significant data is shown as an asterisk in Figure 2.3.B. KARs are tetramers that consist of different combinations of subunits (GluK1-5). GluK2 has been reported to be required for Ca2+-permeability of KARs in normal astrocytes [95]. To confirm that U118-MG astrocytoma cells follow the same expression pattern as normal astrocytes, Western blotting was used and expression of GluK2 in this astrocytoma cell line was observed (Figure 2.3.D). COS-7 cells have been used as negative control. The properties of this cell line will be discussed further in Chapter 4. 44 2.3.4 CNQX inhibits the morphological response of U118-MG astrocytoma cells caused by kainate and glutamate. To examine whether the activity of KARs is essential for the morphological response to glutamate, and to rule out contribution of mGluRs, U118-MG cells were pretreated with a KAR antagonist before exposure to glutamate. Since there are several types of ionotropic and metabotropic receptors for glutamate, inhibition of the morphological response to glutamate by an antagonist against one type of these receptors (non-NMDA) suggests the critical role of that specific receptor. 100 μM glutamate induced 4.82 ± 0.58 μm lengthening of the cell’s processes, which was significantly (p <0.0001) different from the change that was observed in non-stimulated cells (0.22 ± 0.09 μm). Likewise,100 μM KA induced 3.49 ± 0.01 μm change in the length of the processes that was also significantly (p < 0.0001) different from the change in the non-stimulated cells. In contrast, the cells that had been pretreated with 10 μM of CNQX before stimulation with both KA and glutamate show a processes extension comparable with non-stimulated cells (0.12 ± 0.26 μm for KA; 0.16 ± 0.29 μm for glutamate; non-stimulated cells 0.22 ± 0.09 μm) (Figure 2.4.C). CNQX is a common antagonist for KAR and AMPARs, however, AMPARs were shown to be inactive in the process extension of U118-MG cells, which suggest that CNQX is exerting its effect of KARs (Figure 2.4). U118-MG cells have highly motile processes, which demonstrates their viability. Cytotoxic concentration of drugs causes hypotrophy in the cell’s body and inhibits the motility in the of the cell’s processes. Since our imaging was time-lapse and with 60 X magnification, the motility of the processes were detectable throughout the imaging processes, therefore, the viability test (MTT assay) was not done after each imaging. 45 Figure 2.4 An antagonist for kainate receptors inhibited the morphological response of U118-MG cells both to KA and glutamate. (A) Representative confocal micrographs of U118-MG astrocytoma cells exposed 100 μM of KA (top row) and glutamate (bottom row). Cells were transfected with mCherry-lifeact, for live imaging of actin cytoskeleton (red). 60X magnification; 2X zoom; scale bar is 5 μm (B) Representative confocal micrographs of U118-MG astrocytoma cells transfected with mCherry-LifeAct7 that were pretreated with 10 μM CNQX before stimulation with 100 μM of KA (top row) and glutamate (bottom row). cells at 60X magnification, 2X zoom; scale bar is 5 μm (C) Change in a cell's processes length when stimulated with 100 µM glutamate and KA compared to the cells pre-treated with 10 μM CNQX antagonist before stimulation. Compounds were dissolved in water (vehicle). The change in length is the average data of 5 cells, each from a separate culture dish; measurements of at least 20 filopodia were averaged for each cell (n = 5). Data was analyzed by two-way ANOVA (F 1,24 KAR antagonist = 112.9, F 2.24 KAR agonists = 29), All the data sets were compared by Tukey’s multiple comparison post hoc; *p < 0.0001 significantly different from the control (vehicle) and cells that were pretreated with CNQX before exposure to glutamate. The change in processes length of the cells that were pretreated with CNQX pretreated cells before exposure to glutamate and KA were not significantly different with control. n = 5; Data are shown mean ± SEM 46 2.3.5 Phenyl kainic acid (Ph-KA) causes a morphological response similar to KA in U118-MG cells Phenylkainic acid (Ph-KA) was created as a precursor for designing the caged kainoid to have spatiotemporal control over stimulating KARs. The crystal structure and its chemical properties, Ph-KA has potentially a high affinity to the binding site of KARs [142]. Ph-KA was developed in the chemistry lab of Dr. Menard and the processes of its synthesis and validation has been detailed by Tian et al [60, 147]. To use Ph-KA in a biological system, whether it exerts morphological response in astrocytoma cells was tested. Ph-KA induced similar morphological response in U118-MG astrocytoma cells as KA. Within 300 ± 20 seconds after exposure to both KA and PhKA, cells started protruding their processes (Figure 2.5.B). The change in the processes length in U118-MG cells exposed to kainic acid was 3.27 ± 0.4 μm, and was 3.52 ± 0.4 μm upn exposure to Ph-KA which was significantly higher than non-stimulated cells (p <0.0001) (Figure 2.5.C). A KAR antagonist, CNQX, inhibited the morphological response to KA and Ph-KA. When U118-MG cells were pretreated with CNQX before stimulation with KA and Ph-KA, the change in their processes length was statistically non-significant and was comparable with the non-stimulated cells (Figure 2.5.C). Calcium imaging using Fluo4-AM demonstrated that U118-MG cells show a simillar response by a [Ca2+]i rise to KA and Ph-KA. When Ph-Ka was employed, the intracellular Ca2+ reached its maximum concentration 10 seconds later than the KA-induced peak (with 20 seconds lag from stimulation) (Figure 2.5.D). The change in intracellular Ca2+ rise in U118-MG cells exposed to 100 µM KA was 1.5 folds greater than what was recorded in the cells exposed to 100 µM Ph-KA (Figure 2.5.C). The change in the fluorescence intensity of Fluo4-AM was 0.27 ± 0.05 AU/µm2 when U118-MG cells exposed to KA. While this change was 0.18 ± 0.03 AU/µm2 in they were stimulated with Ph-KA (Figure 2.5.C). 47 Figure 2.5 Ph-KA induced a morphological response in U118-MG astrocytoma cells comparable to KA. (A) Representative confocal micrographs of U118-MG cells before and after exposure to 100 M Ph-KA. Cells were transfected with mCherry-lifeact, for live imaging of actin cytoskeleton (red). 60X magnification, Scale bar 5 μm. (B) Time elapsed before the initiation of process extension after stimulation with KA and Ph-KA, n=3. Data are shown mean ± SEM (C) The change in a cell's processes length when stimulated with 100 µM of kainoids compared to cells pre-treated with 10 μM CNQX before stimulation. Compounds were dissolved in water (Vehicle). The change in length is the average data of 5 cells, each from a separate culture dish; measurements of at least 20 filopodia were averaged for each cell (n = 5). Data was analyzed by two-way ANOVA (F 1,32 KAR antagonist = 203.7, F 1, 32 KAR agonists = 23.36), All the data sets were compared by Tukey’s multiple comparison post hoc; *p < 0.0001 significantly different from cells that were pretreated with CNQX before exposure to glutamate and KA. n = 5. (D) Normalized fluorescence of Ca2+ imaging dye Fluo4-AM in U118-MG cells stimulated with 100 μM kainoids or glutamate, n = number of cells / number of dishes. Data are shown mean ± SEM 48 2.3.6 Photocleavage of DECM-PhKA promotes filopodia extension and increases intracellular calcium in U118-MG astrocytoma cells The structure of KA does not allow to be chemically attached of caging molecules for photochemistry purposes [147, 148]. Caged compounds are one type of the photochemical compounds in which the bioactive compound is released upon irradiation of light [129]. Based on the chemical structure of PhKA which suggest a high affinity to KARs as well as its ability to elicit the morphological response in U118-MG cells, DECM-PhKA was envisaged as a “turn-on” probe for the optical control of kainate receptors using visible blue light (Figure 2.6.A). The photochemical properties of the caged DECM-PhKA were characterized by measuring the progressive photolysis of DECM-PhKA by NMR. The photolysis experiments were carried out with two types of blue LED photochemical reactors (custom-made in our lab). Time-dependent photolysis of the caged compound was performed in aqueous solutions to mimic physiological conditions (0.1% DMSO/H2O). NMR spectra and the experimental validation of DECM-PhKA have been detailed by Tian et al [60, 147]. Photocleavage of DECM-PhKA triggered the process extension in U118-MG cells (Figure 2.7.B). Optimization of the 405 nm illumination time based on the DECM-PhKA concentration, showed that with a single 10-second illumination of 405 nm blue light at 2 mM, DECM-PhKA can induce a measurable morphological response in U118-MG cells (Figure 2.6.C). Uncaging DECM-PhKA locally caused the extension of the targeted cell’s processes, comparable to the whole-bath perfusion of a PhKA solution (3.58 ± 0.01 µm and 3.52 ± 0.01 µm, respectively), which was significantly different from not stimulated cells (Figure 2.6.D). The caging compound (DECM-OH), the caged compound (DECM-PhKA in the absence of light), and their vehicle (DMSO at 0.1% v/v concentration) did not cause a significant change in length of the processes of astrocytoma cells compared with non-stimulated cells (Figure 2.6.D). 49 Figure 2.6 Photocleavage of DECM-PhKA promotes filopodia extension in U118-MG astrocytoma cells. (A) Chemical scheme of optical cleavage DECM-PhKA, which causes processes extension in U118-MG astrocytoma cells. (B) Confocal micrographs of U118-MG cells in the cell culture solution (HBSS containing 2 mM Ca2) pretreated with an intracellular Ca2+ dye, Fluo4-AM. The media were then supplemented with either 2mM of DECM-OH (the kainoid’s cage) as a control (top row) or with DECM-PhKA (bottom row) before imaging. 405nm light was irradiated for 10 seconds to the media. Cells were transfected with mCherry-lifeact, for live imaging of actin cytoskeleton (red). 60X magnification, 2X zoom, scale bar 5 μm. (C) DECM-PhKA-irradiation curve: Irradiation time to initiate filopodia extension in U118-MG cells as a function of the initial concentration of the DECM-PhKA in the cell culture solution (HBSS containing 2 mM Ca2+). Mean ± SD, n = 3. (D) The change in the cellular processes length (Δ length) without stimulation (blank) and in response to 405nm light, DMSO (0.01% v/v, vehicle), DECM-OH (the caging compound), DECM-PhKA without and after light exposure as well as KA and PhKA . The final concentration of glutamate, KA, Ph-KA, DECM-OH and DECM-PhKA were 100 µM. For experiments with light irradiation, 405 nm light was irradiated for 10 S via an optical fiber. The change in length is the average data of 5 cells, each from a separate culture dish; measurements of at least 20 filopodia were averaged for each cell (n = 5). All the data sets were compared with Tukey’s multiple comparison post hoc (one-way ANOVA); *p < 0.0001 significantly different from controls (blank, light, vehicle, DECM-OH and EDCM-PhKA without light). n = 5; Data are shown mean ± SEM 50 When U118-MG cells were pretreated with an intracellular calcium sensor (Fluo4-AM) and their media was replaced by HBSS (supplemented with 2 mM Ca2+) containing 2 mM caged kainoid (DECM-PhKA), irradiating 405 nm blue light for 10 seconds induced a significant increase in the fluorescence signals from Fluo4-AM (Figure 2.7.A and B). Photocleavage of DECM-PhKA produces the KAR agonist, Ph-KA, and the DECM-OH, which is the molecular cage for Ph-KA. Figure 2.7 Photocleavage of DECM-PhKA causes an intracellular Ca2+ rise in U118-MG astrocytoma cells. A) Epi-fluorescent micrographs of U118-MG cells in the cell culture solution (HBSS containing 2 mM Ca2) pretreated with an intracellular Ca2+ dye, Fluo4-AM. The media were then supplemented with either 2mM of DECM-OH (the kainoid’s cage) as a control (top row) or with DECM-PhKA (bottom row) before imaging. 405nm light was irradiated for 10 seconds to the media. 20X magnification; scale bar: 100 μm. (B) Normalized Fluo4-AM fluorescence of Ca2+ imaging of U118-MG cells with 2 mM of DECM-PhKA (red curve) and DECM-OH (gray curve) before and after exposure to 405 nm light; blue bar: 10 s blue light irradiation. (C) Normalized Fluo4-AM fluorescence of Ca2+ imaging of U118-MG cells with (2 mM) DECM-PhKA and DECM-OH, 100 μM glutamate, KA and PhKA, DMSO (0.01% v/v) and 10 seconds irradiation of 405 nm light. n = number of cells / number of dishes 51 Control experiments with DECM-OH did not induce a change above the level of the background fluorescent from calcium within the cells (marked as blank in Figure 2.6.B). A rise in the [Ca2+]i was observed after turning off the light as the result of the DECM-PhKA photocleavage, and reached its maximum in 10 seconds. However, unlike Ph-KA instead of a single sharp peak of [Ca2+]i, (Figure 2.7.C) a continuous plateau was observed (Figure 2.7.B). The change in intracellular Ca2+ concentration upon exposure to 100 μM KA, PhKA, glutamate, DMSO (vehicle) and caged Kainate (DECM-PhKA) in absence of light were also recorded as control experiments (Figure 2.7.C). 2.3.7 Locally releasing glutamate induces directional extension of filopodia in 3D cultured U118-MG cells, but not PhKA. To study the directional motility of the processes of astrocytoma cells toward glutamate, U118-MG cells were cultured in a 3D matrix, where their processes could freely move in three dimensions. Culturing cells in 2D conventional technique was allowing us to observe and quantify the processes extension but was not able to provide any data about the direction of the motility of the cellular processes because they were mainly attached on the surface. Therefore, we optimized culturing U118-MG cells in a 3D matrix (GelMA) and monitor the spatial motility of its processes. However, quantifying the directional motility of the processes required designing a specific algorithm for image processing and measurements with ImageJ, which remained for future investigations. Optically releasing glutamate and PhKA, respectively, from MNI-Glu and EDCM-PhKA using an optical fiber induced a rapid process extension in U118-MG cells with a single 10-second illumination of 405 nm blue light both in 2D and 3D cultured cells. In 2D cultured cells, no directional motility was observed upon exposure to both PhKA and glutamate at the focal plane of view, where the cellular processes were attached on the surface of the culture dish. In both 52 treatments, the processes were protruding toward different directions (marked by white arrows in Figure 2.8.A). Figure 2.8 Optically releasing glutamate but not PhKA induces a directional extension of the processes in 3D cultured U18-MG cells. Confocal micrograph of a U118-MG. Cells were transfected with mCherry-lifeact, for live imaging of actin cytoskeleton (red). 60x magnification. Scale bars 20 μM (A) Cell were encapsulated in GelMA (3D-cultured ) and imaged in MNI-Glu (2 mM) supplemented culture solution (HBSS containing 2 mM Ca2+). Glutamate was released by 10 s irradiation of 450 nm light via an optical fiber to media; Ph-KA was released in the culture solution of 3D cultured cells that was supplemented with 2mM of DECM-PhKA and released by 10 S irradiation of 405 nm light. Directional motility was observed upon the glutamate release from MNI-Glu but not in response to the release of PhKA from DECM-PhKA. (shown by white arrows). Some of the cellular processes have been highlighted in yellow on the surface of the cells in magnified images to show the direction of cellular processes. Arrows show the tip of the processes. (B) cells were cultured in glass bottom imaging dishes (2D). Glutamate was released by 10 s irradiation of 450 nm light via an optical fiber to the culture solution (HBSS containing 2 mM Ca2+) supplemented with 2 mM of MNI-Glu; and Ph-KA was released by irradiating 450 nm light via an optical fiber to the culture solution supplemented with 2 mM of DECM-PhKA . Cells show processes extension upon exposure to both compounds with random direction (shown by white arrows). 53 in contrast, the processes of U118-MG cells that were cultured in 3D GelMA matrix demonstrated a directional motility toward the tip of the optical fiber where glutamate was being released. In fact, glutamate can activate different types of glutamate receptors. However, such a directional motility was not observed when PhKA, that only activates KARs, was released optically. 2.4 Discussion A growing body of evidence indicates that dysregulation of glutamate signaling in the brain is one of the main underlying factors in the onset and progression of brain tumors [63, 64, 67]. The detrimental effect of dysregulated glutamate signaling is mainly attributed to the consequences of its associated Ca2+ signaling, which causes neuronal death and proliferation of glioma cells [36, 62, 63, 149]. In addition to the glutamate homeostasis, emerging evidence suggest that the alteration in the morphology of astrocytoma cells is also an important driving factor in gliomas [15–17]. While the effect of glutamate in shaping polarized morphology in astrocytoma cells, which leads to their migration and infiltration through the brain tissues, has been studied [67, 110, 150], it is not known if glutamate regulates the aberrant morphology of cellular projections of astrocytoma cells. Highly motile and protruding projections can form a growing network of astrocytoma cells that can protect the tumor microenvironment from its surrounding and can also penetrate into the healthy tissues of the brain [15, 16]. These processes have also been shown to make abnormal connections with peritumoral neurons, which enhances the growth of the tumors and protects it from conventional cancer therapies [17]. Yet, the mechanisms through which glutamate may influence the morphology of the projections of astrocytoma cell is less explored. In this dissertation, two astrocytoma cell lines, U118-MG and U138-MG, were selected for studying the response of their cellular processes to glutamate. Unexpectedly, the processes 54 extension in response to glutamate was only observed in U118-MG cells and not U138-MG cells. As has been discussed in Section 1.1, astrocytes naturally extend their processes upon exposure to glutamate in a Ca2+-dependent manner [2, 25]. In fact, many similarities in terms of genetics and protein expression has been reported between U118-MG and U138-MG cell lines [112]. However, there are few major differences between these two cell lines, that has been discussed in Section 1.9.1. One of their main differences is their tumorigenesis potential. U118-MG cells are highly tumorigenic, while U138-MG cells cannot induce tumor formation in immunosuppressed mice [112]. This difference in tumorigenesis might be related to the ability of U118-MG cells in extending their processes in response to glutamate as we have shown. In fact, the ability to extend long processes has been demonstrated to be one the main features of astrocytoma cells in progression and therapy resistance of glioma tumors [15, 16]. However, whether this processes extension is in governed glutamate or other factors in not known. It should be noted that , to be consistent, the only concentration of glutamate that has been used in this research was 100 μM, which its effectiveness in inducing protrusions to astrocytes has been established [3, 25, 42, 138]. The difference in the morphological response between U118-MG and U138-MG can be due to their sensitivity to different concentrations of glutamate. The peak in intracellular Ca2+ rise upon exposure to 100 μM, glutamate was 1.62 ± 0.10 times more in U118-MG cells compared to U138-MG cells. It is possible that U138-MG cells respond to other concentrations of glutamate, which remains to be tested. Interestingly, using photochemistry to release glutamate locally and stimulate 3D cultured cells revealed that processes of U118-MG cells protrude toward the source of glutamate (Figure 2.8.A). The directional migration of astrocytoma cells toward glutamate and its role in infiltration of glioma through the brain has been reported [67, 110]. Our observations suggest that glutamate can also serve as a guidance cue for free moving processes of astrocytoma cells similar to what has been shown in neurons [20]. 55 The contribution of ionotropic glutamate receptors (iGluRs) in this morphological response of U118-MG cells to glutamate was investigated, based on the results from Section 2.3.2, which showed that processes extension depends on the extracellular Ca2+. iGluRs including NMDARs, AMPARs and KARs are ligand gated ion channels and conduct Ca2+ upon binding glutamate. Therefore, their function depends on the free extracellular Ca2+. In contrast to iGluRs, metabotropic glutamate receptors (mGluRs), release Ca2+ from internal stores and their activity is independent from the free Ca2+ in the media, which makes their contribution in this glutamate-induced morphological response less likely [76, 151]. In addition, among iGluRs, there is no consensuses about the activity of NMDARs in astrocytes. This uncertainty about the activity of NMDARs in astrocytes is due to the mode of function of NMDARs because their activity depends on voltage. NMDARs display a degree of voltage dependence due to Zn2+ or Mg2+ blockade in the pore but astrocytes are electrically inactive and cannot unblock the channel [18, 69, 79]. It should be noted that a rare modified class of NMDARs that are devoid of Mg2+ blockade have been reported in a subset of neocortex and cerebral astrocytes [78, 79]. The NMDARs found in astrocytoma cells and glioma cell lines are usually not functional because they lack the essential NR1 subunit [74]. AMPA and kainate receptors are usually pooled together as non-NMDA receptors due to their structure similarities and the lack of selective antagonists to discriminate them from each other [71–73]. 100 μM of both AMPA and KA (but not NMDA) are known to efficiently cause filopodiagenesis and processes extension in cultured astrocytes [25]. But in this chapter, the consecutive application of AMPA followed by KA on U118-MG astrocytoma cells demonstrated that only KA induces the morphological response (Figure 2.3.B and 2.3.C). It should be noted that the only concentration tested was 100 μM, and if other concentrations of AMPA can elicit a different response in U118-MG cells remains to be studied. 56 KARs are tetramers that consist of different subunit combinations (GluK1-5) [40, 152, 153]. Based on these combinations, they respond to different selective antagonists [40, 152, 153]. CNQX is the only effective antagonist against all of the KARs with different arrangements in their subunits [71]. It should be noted that CNQX can non-selectively block AMPARs as well [71]. However, as shown in Section 2.2.3, AMPA did not activate the processes extension in U118-MG cells, therefore the effect of CNQX on AMPARs should not change the interpretation of the results about the role of KARs. Applying CNQX prior to stimulating U118-MG cells with KA blocked their normal process's extension. Intriguingly, CNQX also inhibited the morphological response to the glutamate (Figure 2.5.B and 2.5.C). Glutamate is a common ligand for different types of glutamate receptors (3 types of ionotropic and 8 types of metabotropic receptors) [41]. The inhibitory effect of a KAR antagonist on the processes extension in response to glutamate suggests that while different types of glutamate receptors (iGluRs or mGluRs) may receive the signal from glutamate, the activity of KARS is essential for the initiation of morphological response of U118-MG cells to glutamate. In fact, a comparison between the peak of the Ca2+ rise upon stimulation with KA and glutamate (Figure 2.7.C) suggests that other glutamate receptors rather than KARs, may have contributed to the rise of intracellular calcium upon stimulation with glutamate. The [Ca2+]i peak induced by glutamate was 1.2 - 1.4 folds higher than when cells were stimulated by KAR agonists. This higher peak of [Ca2+]i upon exposure to glutamate may also explain why U118-MG cells initiate protruding their processes in a shorter time gap in response to glutamate compared to kainoids (Figure 2.5.B and the Figure of appendix. 2). Ca2+ is a required element in actin cytoskeleton rearrangement which constructs the fine processes of astrocytoma cells; an increase in Ca2+ may have supported faster assembly of actin cytoskeleton when cells were exposed to glutamate. 57 KARs were examined to determine whether they control the directional motility of the filopodia in astrocytoma cells toward glutamate. The same technique to test directional motility of the processes of U118-MG cells to glutamate was used, but instead cells were stimulated with a selective agonist for KARs. To create a KAR-specific activator, a KA analog made in house: phenylkainic acid (PhKA) was caged (Appendix A) [60]. The “free” PhKA demonstrated the same level of morphological response as KA in U118-MG astrocytoma cells (Figure 2.5). KA was caged with a DECM group that can be cleaved at a visible wavelength which is harmless to the cells. Uncaging the kainoid induced process extension and a [Ca2+]i rise in astrocytoma U118-MG cells (Figure 2.6), but no directional motility in the processes of the cells was observed (Figure 2.8). This result suggests that glutamate but not PhKA can direct the pathfinding of the cellular processes of U118-MG cells. In general, these findings can suggest that KARs are required for initiation of the morphological response of astrocytoma cells to glutamate but other proteins or receptors sensitive to glutamate are responsible for pathfinding. The chemical structure of caged-Ph-KA may have affected the response of the cells, since we observed a different pattern in the Ca2+ influx following the photorelease of Ph-KA. Other experimental designs or novel tools in future may provide more information about the role of KARs in the directional motility of astrocytoma cells. Molecular studies could provide details about the specific subunits of KARs, their combinations and how they are contributing to the glutamate signaling of U118-MG astrocytoma cells. In this research, Western blotting showed the expression of GluK2 (Figure 2.3.D). Selective blockers for different combinations of KAR agonists and silencing KAR genes would help to identify the type of active KARs in astrocytoma cells. This study needs to be repeated using primary cells and organotypic slices to provide a clear understanding about the role of KARs in glutamate-induced morphological response and the pathology of astrocytoma tumors. 58 It should also be noted that the high motility of the processes of astrocytoma cells being imaged in real time was a indicating the cell viability in this research. However, viability assays could provide further information about the toxicity of pharmacological and chemical reagents in each imaging dish [139]. In summary, two astrocytoma cell lines were tested to determine if they respond to glutamate by a directional extension of processes via the activity of a glutamate receptor. U118-MG cells were found to respond to glutamate by significant processes extension and a Ca2+ influx. KARs were found to be essential for the initiation of the morphological response to glutamate in U118-MG cells, but do not control the directional motility of the processes of these cells toward glutamate. Further studies are required to investigate the role of KARs in pathology of astrocytoma cells and glioma. 59 Chapter 3. L-Type Voltage-Gated Calcium Channels Contribute to Glutamate-Induced Morphological Response in Astrocytoma Cells Voltage-gated calcium channels (CaVs) are ion channels whose gating depends on the change in the membrane potential [154]. High voltage CaVs were thought for long to be exclusive to excitable cells [154–157]. However, astrocytes, which are classified as non-excitable members of the CNS have been shown to express high voltage CaVs (L-type) [85, 86]. The function of L-type calcium channels (LTCCs) in complex and interconnected calcium signaling pathways in astrocytes is still not known. However, they have been shown to participate in different pathological conditions of brain such as in neurodegeneration and brain cancers [86, 98, 99, 158]. In brain tumors, LTCC antagonists have been reported to control the proliferation and migration of transformed astrocytoma cells [98–100]. However, the molecular mechanism through which LTCCs contribute to the growth and progression of astrocytoma tumors is unknown. On the other hand, dysregulation in glutamate homeostasis is emerging to be one of the underlying mechanisms of the onset and progression of astrocytomas [62, 63]. Typically, there is a direct correlation between glutamate and calcium events in cells [4, 50, 64]. Whether L-type calcium channels contribute to the tumor progression by involving in the Ca2+ events following glutamate signaling in astrocytoma cells has remained unexplored. In this chapter, we hypothesized that LTCC contribute to the glutamate-induced morphological response in astrocytoma cells. The objectives of this chapter are (1) to identify whether the calcium influx through CaVs contributes to the glutamate-induced morphological response in astrocytoma cells (U118-MG), and (2) to determine if glutamate can influence the dynamics of LTCCs in U118-MG astrocytoma cells. Using a pharmacological approach, different subunits of LTCCs were found to play a role in glutamate-induced morphological response in U118-MG astrocytoma cells, in this research. 60 Pharmacological agents, that are known to block the pore-forming subunit of CaV’s, decreased the process's extension in response to glutamate in U118-MG cells. While pharmacological ligands for their extracellular auxiliary subunit (α2δ1) inhibited this response entirely. By employing a custom fabricated imaging probe, glutamate was shown to induce a rapid rise in the membranal expression of LTCCs in U118-MG cells. FluoBar1 is a molecular imaging tool that we developed to monitor the real-time dynamics of LTCCs in living cells. 3.1 Background Growing evidence shows that glutamate and calcium signaling provide a positive feedback to promote glioma progression [17, 33, 36]. Glutamate induces an intracellular Ca2+ rise in glioma cells and the elevation of intracellular Ca2+ has been suggested to upregulate cytoskeletal proteins and to increase glutamate release; both of which can facilitate formation of synaptic-like connections with surrounding cells and accelerate tumor progression [36, 103]. In tumor cells, altered Ca2+ homeostasis often involves dysregulated expressions, cellular localization and activity of Ca2+-transporting proteins which leads to the tumor specific characteristics such as unlimited growth, metastasis and resistance to apoptosis. The role of different receptors and proteins that are involved in calcium signaling of astrocytoma cells such as Store-Operated Ca2+ Entry (SOCE), TRP channels, IP3 receptors, P2X receptors and CaVs have been studied [36, 159]. In primary human glioma cells, higher levels of SOCE have been observed in comparison to non-malignant human primary astrocytes [159]. The SOCE is a process in which a Ca2+ influx refills the endoplasmic reticulum’s (ER) after Ca2+ depletion [160]. Silencing SOCE has been shown to reduces invasion and proliferation of glioma cells [161]. On the other hand, a number of studies have shown the involvement of TRP channels in proliferation, invasion and angiogenesis of glioma cells [162]. TRP channels are calcium channels that will be activated upon chemical 61 and physical stimuli. Different studies have shown the contribution of TRP channels in the glioma progression [159]. Interestingly, some studies have shown that inhibition of IP3 receptors IP3R3 by caffeine reduces cell invasion and migration in glioma, and it increases survival of subject animals [163]. IP3R convert external stimuli to intracellular Ca2+ signals. In addition, contribution of P2X receptors, to the cell migration and expression of pro-inflammatory factors in glioma cells has also been reported [159]. P2X channels are membrane ion channels that open upon binding of ATP [96, 159]. Voltage-gated calcium channels (CaVs) are the other group of calcium channels that their contribution to pathology of glioma tumors have been shown. The role of low voltage CaVs (T-type) in proliferation and survival of glioma cells has been repeatedly reported and reviewed [159]. T-type Ca2+ channels typically control processes such as cell proliferation and differentiation by regulating Ca2+ levels at low voltage through creation of resting inward calcium currents in the cells [98, 120]. In addition, antagonists against L-type CaVs (LTCC) have been shown to inhibit glioma progression [83, 99, 100]. However, the mechanism through which LTCC antagonists exert their effect to control glioma progression has remained to be elucidated. In this chapter, the potential contribution of LTCCs in the morphological response of astrocytoma cells to glutamate will be examined. In most cells, LTCCs couple the change in membrane potential to a Ca2+ influx, where Ca2+ ions act as secondary messengers in different cellular signaling cascades [164]. Structurally, LTCCs consist of three main subunits that associate in a 1:1:1 stoichiometry: a pore-forming α1 subunit, an external α2δ subunit, and an intracellular β subunit (Figure 3.1) [90, 91, 154, 165]. The intracellular β subunit does not contribute to the channel’s conductance, but targets the channel to the membrane [166]. The α1 transmembrane subunit forms the channel’s pore and contains the voltage sensors [154, 156]. The α1 subunit also contains the binding site for calcium 62 channel blockers (Figure 3.1) [67, 167]. The extracellular α2δ subunit assists both the α1 and β subunits by modulating α1 subunit conductance [168, 169] and directing the channels’ localization density, and dynamics on the membrane [90, 170]. In fact, the two main subunits of CaVs that affect Ca2+ conductance are α1 and α2δ and will be studied in this dissertation. CaV antagonists or Ca2+ blockers inhibit the Ca2+ influx through the pore-forming subunit of the channel [167]. They are categorized into two different classes, dihydropyridines and non-dihydropyridines (non-dihydropyridines consist of phenylalkylamines and benzothiazepines subclasses). The most common representative of each class of the Ca2+ blockers are Nifedipine (dihydropyridine), Verapamil (phenylalkylamine) and Diltiazem (benzothiazepine) [91, 92, 171]. Figure 3.1 Voltage-gated calcium channel subunit topology. Channel inhibition can be induced by modification of channel gating (interaction with voltage-sensing domains shown in blue) and by interaction within the activation gates of the pore domain channel, shown in brown. The α2δ ligands can modify the channel trafficking. The antibody that was used in this research to stain CaV1.2 (Alomone ACC-003) binds to the second internal loop of the pore-forming subunit. Created with biorender.com 63 These blockers typically bind to the α1 subunit and by either blocking or changing the configuration of the channel’s pore interrupt the Ca2+ conductance. In addition to the α1 subunit, the extracellular α2δ has its own pharmaceutical ligands called gabapentinoids. As their name imply, gabapentinoids were originally designed as a GABA mimetic, but they do not affect GABAA or GABAB receptor activity [170, 172, 173]. Gabapentin and pregabalin bind to the α2δ subunit of CaVs [174]. Gabapentin (IC50 = 17 mM), and pregabalin (IC50 = 37 nM) are approved anti-epileptic and anti-allodynic class of drugs [174–179]. Gabapentinoids demonstrate high efficacy in pain management, which has been their most extensively prescribed application to date [180]. However, how they carry out their effects through α2δ has been a long question. LTCCs are highly mobile channels that move in clusters on the cell membrane [181]. This motility is tightly regulated both in time and space to align with all the other cellular components that contribute to a specific signal transduction pathway [169, 182]. There is a precise spatiotemporal control over the Ca2+ influx through CaVs within the crowded dynamic environment of biological membranes due to the biophysical properties of these channels [156, 169]. Thus, observing their membranar motility would provide important data about the contribution of CaVs in a particular cellular event. However, current conventional techniques cannot efficiently show the translocation of CaVs on the membrane. Current techniques are either based on immunostaining that requires fixed cells or is based on genetically fusing fluorescent proteins with the target protein (CaVs) that affect the protein’s size. Emerging chemical tools such as small molecule imaging probes are growing and promising techniques to monitor the movement of these channels in real time. In this research, we developed and used one of these chemical imaging tools to monitor the CaVs translocation in real time by time-lapse confocal imaging. The imaging probe, FluoBar1, was developed based on a well-studied unsymmetrical barbiturate that is a selective ligand for astrocytic LTCCs (CaV1.2 and CaV1.3) and does not block 64 the activity of CaV1.2 [183, 184]. Detailed electrophysiology studies on the selectivity of FluoBar1's parent barbiturate for LTCCs have been reported by the Silverman, Striessnig and Soong labs [165, 183, 184]. The process of its chemical synthesis and validation has been detailed by Tabatabaee et al [185] and will be discussed briefly in this chapter. 3.2 Material and methods 3.2.1 Cell culture Human astrocytoma cells, U118-MG, were cultured in Dulbecco’s modified essential medium (DMEM, Gibco 11995-065) supplemented with 10% v/v heat-inactivated fetal bovine serum (HyClone 12483020) 1% v/v penicillin 10,000 U/ml and streptomycin 10,000 μg/ ml (Gibco 15140-163). Cells were incubated in a humidified atmosphere containing 5% CO2 at 37 ºC and passaged at 80% confluence using a pre-warmed 0.25% trypsin-EDTA solution for a maximum of 5 minutes (Gibco SH30236.02). For the experiments, the cells from a 10 cm culture dish at ~80% confluence were resuspended with a pre-warmed 0.25% trypsin-EDTA solution and transferred to 35 mm glass-bottom culture dishes (Mutsunami D1130H) in phenol red-free/high glucose DMEM supplemented with 25 mM HEPES (Gibco 21063-029). Enough cells were transferred to obtain a ~50% confluent 35 mm culture dish after adhesion. Cells were incubated for less than the cells’ doubling time, typically 30-33 hours, to minimize imaging variation between samples (Section 3.2.6) [84]. This period allows cells to adopt a flattened shape that helps define the filopodia for measurements. Identical incubation times were respected to ensure cells were at the same growing stage in all treatments. 3.2.2 Transfection U118-MG astrocytoma cells were transfected with 1200 ng/ μl of the F-actin marker mCherry-LifeAct-7 plasmid using a calcium phosphate precipitation protocol as described in Section 2.2.2 [85]. mCherry-Lifeact-7 was a gift from Michael Davidson (Addgene plasmid 54491). 65 COS-7 cells were co-transfected with ~1200 ng each of two plasmids: pcDNA3-α1c (rat CaV1.2) generously gifted by Terry P. Snutch (UBC Vancouver), and pcDNA3-EGFP was a gift from Doug Golenbock (Addgene plasmid 13031) using a calcium phosphate precipitation protocol [85]. pcDNA3-EGFP is used as a reporter to identify cells that have taken pcDNA3-α1c (CaV1.2). 3.2.3 Treatment with voltage-gated calcium channel antagonists The α1 and the α2δ1 subunits of CaVs were treated with antagonists to examine whether they participate in glutamate-induced morphological response in astrocytoma cells. As a control, U118-MG cells in phenol red-free/high glucose DMEM supplemented with 25 mM HEPES were stimulated with glutamate (Alfa Aesar A15031-30) on the confocal microscope stage. Briefly: 10.0 μL of a 20X glutamate stock solution was delivered to the medium (2.0 mL in a 35 mm dish) at the outer edge of the dish for a final concentration of 100 µM. The stock solution of L-glutamate was prepared in deionized millipore water and filter-sterilized through a 0.22 μm syringe filter. Glutamate was delivered to the medium four minutes after the beginning of imaging. CaV Blockers: Nifedipine (N7634, sigma Aldrich), Verapamil hydrochloride (14288, Cayman chemical) and Diltiazem hydrochloride (AC329080010, Acros organic) were delivered to glass-bottom culture dishes (Mutsunami D11130H) for a final concentration of 10 μM. Stock solutions of CaV inhibitors were prepared in DMSO (vehicle) before each experiment. Solutions were filter-sterilized using a 0.22 μm syringe filter. To avoid the negative effect of DMSO on the cells, stock solutions were highly concentrated (200X), thereby maintaining the final amount of DMSO below 1% v/v in the culture medium. Gabapentinoids: Gabapentin (G154, Sigma Aldrich) and pregabalin (made in house) were prepared directly in deionized millipore water (vehicle). They were also prepared as a 200X stock solution for consistency and to avoid osmotic shock to cultured cells. They were delivered to the medium at final concentration of 10 μM. After adding the pharmacological agents, cells were incubated at 37 ºC under a humidified 5% CO2 atmosphere 66 for 30 minutes. The incubation time and final concentration were adopted from the product instruction from the suppliers and controlled with the literature [95, 98, 137, 186]. Cellular morphology and processes motility were considered as indicators for cellular viability in U118-MG cells. The viability of U118-MG cells after treatments with pharmacological reagents was assessed based on the high motility of their processes as well the extended and flattened soma of the cells. The cells that were under detrimental conditions displayed a shrinking and spherical soma as well as highly ramified membrane and non-motile processes. However, viability assays could provide further information about the general metabolic health of the cells after treatments with each compounds [139]. Table 3.1 Calcium channel inhibitors and chemicals Reagents Type Source Concentration Treatment duration References Nifedipine Dihydropyridine L-type channel blocker Sigma-Aldrich 10 μM 30 min Subbarao et al. 1995; Maneshi et al. 2015; Zatkova et al. 2018, Sinnegger-Brauns et al. 2009 [95, 186–188] Verapamil hydrochloride Phenylalkylamine L-type channel blocker Cayman chemical company 10 μM 30 min Wick et al. 2004; Maneshi et al. 2015, Kunert‐Radek et al. 1989, Lee et al. 1994 [101, 102, 187, 189] Diltiazem hydrochloride Benzothiazepine L-type channel blocker Acros organic 10 μM 30 min Maneshi et al. 2015, Lee at al. 1994 [101, 187] Gabapentin Gabapentinoid α2δ subunit blockers Sigma-Aldrich 10 μM 30 min Fehrenbacher et al. 2003; Sun et al. 2013 [52, 190] Pregabalin Gabapentinoid α2δ subunit blockers Prepared in house, Menard Lab 10 μM 30 min Fehrenbacher et al. 2003; Chen et al. 2018 [190, 191] 3.2.4 Monitoring the processes extension in time-lapse confocal imaging Process extension was monitored as the morphological response of glutamate signaling in astrocytoma cells. mCherry-Lifeact-7 transfected U118-MG cells were imaged on an Olympus FV1000 fluorescence confocal microscope equipped with a Plan-ApoN 60x/i.4 oil-immersion objective. mCherry-Lifeact-7 was observed at 590 nm (Olympus filter set Cy3.5), with 559 nm excitation from a He–Ne laser. Changes in the cell morphology were recorded at 60X 67 magnification. To measure filopodia extension, an image of a single cell was acquired every 20 seconds for a total of 60 frames. Photobleaching was minimized by irradiating samples for 2.0 seconds at 20 seconds intervals with the laser power set at 3%. 3.2.5 Quantification of filopodia extension To quantify the morphological response in astrocytoma cells, the change in the length of cellular processes upon exposure to glutamate was reported. The length of twenty filopodia per cell was measured manually using ImageJ software as described by Zatkova et al. (NIH ImageJ, FIJI) [95]. Briefly, only the filopodia that their entire arbor was in the field of view throughout the imaging period and were in straight line were selected. A filopodium’s length was measured from the edge of the cell membrane to its apical end. The length of each filopodia was manually measured three times. To calculate the change in length (Δ length), a process's length from the image prior to stimulation was subtracted from the length of the same filopodia after 20 minutes. The reported Δ length for each cell is the averaged change in the length of 20 selected filopodia per cell. One cell was chosen from each dish; and the average length of 5 dishes has been reported. Measurements were done by an investigator blinded to the experimental conditions or to the expected results while the experiments have been performed by the researcher who had designed the experiments. 3.2.6 Statistical analysis Each value represents the averaged changes in the length of processes measured from at least 5 cells, each from a separate culture dish (n ≥ 5); the results are presented as the mean ± SEM. Significant differences among groups were determined using one-way or two-way analysis of variance (ANOVA) and followed be a Tukey’s post hoc analysis multiple comparison analysis as well as unpaired T test using Prism 7.04 from GraphPad. A p value < 0.05 was considered significant. To minimize technical variations within our samples for ANOVA analysis, we employed 68 a completely randomized block design [192]. Briefly: cells of a dish at 80-95% confluency were trypsinized and simultaneously re-plated in several 35 mm imaging dishes and incubated in a humidified atmosphere containing 5% CO2 at 37 ºC for less than 33 hours to avoid variation between samples. 3.2.7 SDS PAGE and Western Blotting Total protein was collected from cultured U118-MG, COS-7 and CaV1.2 transfected COS-7 cells. Cells were resuspended with prewarmed 0.25% trypsin-EDTA solution (Gibco SH30236.02), then transferred to a 15 mL falcon tube, and spun down at 1000 rpm for 5 min at 4 ºC. The pellet was mixed with 50 μL of 1% protease inhibitor cocktail (Amresco, mammalian M250) in sterile Millipore deionized water. Then, 50 μL of NP40 lysis buffer (50 mM Tris-HCl, NaCl 150 mM, 1% NP40-Triton 100x pH 7.4) was added to the solution and incubated for 15 min at 4ºC. The mixture was centrifuged at 3000 rpm for 5 min at 4 ºC. The pellet was discarded, and the supernatant was stored at –20 ºC. Using electrophoresis at 400 mA for 90 minutes on ice separated the native proteins in a 4% Tris-glycine gel based on their charge and the length of the polypeptide. The protein band were transferred electrophoreticly onto a Nitrocellulose membrane (Amersham Protran pore size 0.45 μm) and followed by an immunostaining procedure to visualize the proteins on the blotted membrane. The primary antibody used for Western blots was against GRIK2 (1:1000; AGC-009 Alomone). The primary antibodies used for Western blots were against: CaV1.2 (1:1000; ACC-003 Alomone), CaV1.3 (1:1000; ACC-005 Alomone). The protein bands were detected by chemiluminescence using the Amersham ECL kit (GE Healthcare) with horseradish peroxidase-conjugated secondary antibodies (GE Healthcare) The blots were analyzed solely qualitatively and no normalization or quantification was done. 69 3.2.8 Chemical synthesis of FluoBar1 The convergent synthesis consists of ten reactions with six steps for the longest linear sequence (14% overall yield). Starting materials are: cyclopentane carboxylic acid 2-(3-chlorophenyl) acetonitrile, and 2-aminoethanol. Procedures for the synthesis of the imaging probes, reagents, instruments, and chemical characterization are detailed in appendix D [185]. 3.2.9 Immunocytochemistry To identify the change in the distribution of CaVs after exposure to glutamate, U118-MG cells were fixed and stained with selective antibody for the pore-forming subunit of CaV1.2. Briefly, control cells were washed with phosphate buffered saline (PBS) solution. Cells were fixed in 4% paraformaldehyde in PBS solution for 20 min at room temperature without stimulus; experimental cells were exposed to glutamate for 10 minutes, then fixed with the same procedure. Fixed cells were incubated in a blocking solution (10% BSA in PBS), followed by overnight incubation at 4 °C with the anti-CaV1.2 (rabbit anti-rat/human polyclonal, Alomone ACC-003) primary antibody in 3% BSA in PBS (1:200 dilution). After three rinses with PBS, cells were treated with a 1:200 dilution of the goat anti-rabbit IgG (H+L) conjugated to DyLight 550 (Thermoscientific PA5-84541) secondary antibody at room temperature on a rocking platform for 2 hours. The same procedure was used to immunostain COS7 and MCF7 cells for CaV1.2 to serve as controls for colocalization assays. Selectivity of the CaV1.2 antibody and its isotypes were evaluated by dot blotting assays based on the differential expression of native LTCCs in U118-MG, COS7and MCF7 cells (Appendix E). 3.2.10 Protein labeling with FluoBar1 Distribution of CaV1.2 proteins in the live cell was monitored using FluoBar1 fluorescent labeling. Incubation time and the probe concentration were optimized for the least signal to noise ratio at the shortest time (to avoid passive diffusion). Live cells were incubated with 50 nM of FluoBar1 in PBS solution at room temperature for 2-5 minutes. Short incubation time minimizes 70 passive diffusion of the probe to the cell membrane. The FluoBar1 solution was replaced by Hank's balanced salt solution (HBSS, Lonza 10547F) supplemented with 2 mM CaCl2 (Sigma Aldrich, C1016) before imaging. 3.2.11 Staining for colocalization test Fixed cells immunostained for CaV1.2 (secondary antibody conjugated to Dylight 550) were incubated with 50 nM of FluoBar1 in PBS at room temperature on the confocal microscope stage. The probe solution was replaced with PBS solution after 3 minutes, and cells were imaged immediately thereafter. Images were acquired sequentially using fluorescence filters in separate channels: AlexaFluor405 for the probe (emission at ~ 450 nm, blue) and Cy3 for antibody–DyLight 550 (emission at ~620 nm, red). The images were acquired at 60x magnification, with Kalman factor 4, and analyzed for colocalization. 3.2.12 Image analysis for colocalization Confocal microscopy images from AlexaFluor405 and Cy3 channels were processed with ImageJ software (NIH ImageJ, FIJI). using the colocalization analysis function to obtain the Pearson’s r value, the Costes P value, and the Mander’s coefficient. The changes in fluorescence density of FluoBar1 were also measured with FIJI [193]. Briefly: each cell's contour was outlined manually, the total fluorescence signal for this area was integrated, and the fluorescence density was calculated. Background fluorescence was measured by selecting three cell-free areas (approximately half a cell’s size each, from the same image) and their signal was integrated, and their density was averaged. The area conversion value for calculations was 3.02 pixels / μm. The corrected total cell fluorescence density (CTCF) was calculated according to: CTCF = (Integrated density of a cell) – (Integrated density of background) All data were processed and plotted using Prism 7.04 from GraphPad. Each value represents the averaged changes measured from three separate culture dishes; the results are presented as the mean ± SEM. 71 3.2.13 Real-time monitoring of LTCC’s redistribution using FluoBar1 Live U118-MG cells labeled with 50 nM of FluoBar1 for 5 minutes were imaged every 20 seconds for a total number of 70 frames on an Olympus FV1000 fluorescence confocal microscope equipped with a Plan-ApoN 60x/i.4 oil-immersion objective. Glutamate was added four minutes after beginning of imaging for signal stability at a final concentration of 100 μM. Photobleaching was minimized by irradiating samples for 1 second at 20 second intervals with the laser power set at 3%. Fluorescence of the chemical probe was excited with a 405 nm He–Ne laser (Olympus filter set AlexaFluor 405). Confocal microscopy images from AlexaFluor405 channel. Reported data is the average of signals from three cells of separate dishes analyzed with ImageJ software (NIH ImageJ, FIJI). 3.3 Results 3.3.1 Blocking the pore-forming subunit of CaVs in U118-MG astrocytoma cells caused a significant decrease in their processes extension upon exposure to glutamate U118-MG astrocytoma cells (ATCC HTB-15TM) were treated with known Ca2+ blockers to identify whether CaVs contribute to the glutamate-induced morphological response. The glutamate-induced process extension in U118-MG astrocytoma cells decreased significantly (50 – 65 %) in cells that were pretreated with LTCC blockers before exposure to glutamate. The change in the process’s length in U118-MG cells when cells exposed to glutamate was 4.82 ± 0.92 μm. While the cells that were pre-treated with LTCC antagonists for 30 minutes prior to glutamate stimulation demonstrated significantly smaller change in their process’s length compared to untreated cells. The filopodia outgrowth was 2.36 ± 0.77 µm, when cells were pretreated with 10 µM nifedipine before exposure to glutamate, 1.82 ± 0.60 µm when pretreated with 10 µM verapamil, and 1.70 ± 72 0.79 µm when pretreated with 10 µM diltiazem. While filopodia protrusion upon exposure to glutamate decreased significantly in the cells that were pretreated with LTCC blockers compared Figure 3.2 Blocking the L-type calcium channels decreases the glutamate-induced processes extension in U118-MG cells. (A) Representative images of U118-MG astrocytoma cells stimulated with 100 μM glutamate that were pretreated with LTCC blockers (10 μM) for 30 minutes prior to stimulation. Cells were transfected with mCherry-lifeact, for live imaging of actin cytoskeleton (red). 60X magnification, 2X zoom; scale bar 5 μm. (B) Glutamate induced a significant change in processes length compared to unstimulated cells. Processes extension upon exposure to glutamate was decreased significantly when cells were pre-treated with LTCC blockers. DMSO (0.01% v/v) was the vehicle. Change in length (Δ) was calculated by subtracting the length of a filopodium immediately prior to stimulation from that of the same filopodium after 20 minutes. The change in length is the average data of 5 cells, each from a separate culture dish; measurements of at least 20 filopodia were averaged for each cell (n = 5). Data were analyzed by two-way ANOVA (F 4,40 glutamate treatment = 16, F 1,40 Ca2+ blockers = 227.3, for both factors p < 0.0001). All data sets were compared with each other by post hoc Tukey’s multiple comparison. *p <0.0001 significantly different from the cells that were pretreated with LTCC blockers before exposure to glutamate. #p <0.01 significantly different from the cells that has not been stimulated with glutamate. Data are shown mean ± SEM. (C) Western blots show that U118-MG cells express CaV1.2 but not CaV1.3.; Cell lysates were blotted and imaged for CaV1.2 and CaV1.3 with chemiluminescence. CaV peptides (Alomone) and the rat brain lysate were used as positive controls and COS-7 cell lysate was used as a negative control. 73 to the control cells that were not pretreated with the antagonists (4.82 ± 0.95 µm), the processes extension was still significantly greater (6.0-8.4 folds) than the cells that were not exposed to glutamate (p < 0.0001). SDS-PAGE and Western blotting showed detectable expression of CaV1.2 but not CaV1.3 in these cells Figure 3.2.C. 3.3.2 Ligands for the α2δ1 subunit suppressed the glutamate-induced processes extension in U118-MG cells To identify whether the extracellular subunit (α2δ1) modulates the function of CaVs in morphological response of astrocytoma cells to glutamate, U118-MG cells were pretreated with Figure 3.3 Ligands for α2δ1 subunit of CaVs inhibited glutamate-induced filopodia extension in U118-MG cells. (A) Representative micrographs of U118-MG cells that were stimulated with 100 µM glutamate without treatment (control) and pretreated with 10 μM α2δ ligands. Cells were transfected with mCherry-lifeact, for live imaging of actin cytoskeleton (red). 60X magnification, 2X zoom; scale bar 5 μm. (B) Gabapentinoids (α2δ1 ligands) inhibited processes extension in response to 100 µM glutamate. Change in length (Δ) was calculated by subtracting the length of a filopodium immediately prior to stimulation from that of the same filopodium after 20 minutes. The change in length is the average data of 5 cells, each from a separate culture dish; Measurements of at least 20 filopodia were averaged for each cell (n = 5). Data were analyzed by two-way ANOVA (F 1,24 glutamate treatment = 38.75, F 2,24 gabapentinoids = 48.75, for both factors p < 0.0001); All data sets were compared with each other by post hoc Tukey’s multiple comparison, *p < 0.0001 statistically different with the cells that were not stimulated with glutamate or the cells that have been pretreated with gabapentinoids before stimulation with glutamate. Results are presented as mean ± SEM. 74 chemical ligands for α2δ1 before stimulation with glutamate. Unexpectedly, no significant change in the processes length of the cells that were pretreated with α2δ1 ligands (gabapentinoids) before exposure to glutamate was observed which suggest that α2δ1 ligands had an inhibitory effect on glutamate stimulation (Figure 3.3.B). Glutamate in the cells that were not pretreated with gabapentinoids induced 4.82 ± 0.92 μm change in the length while this change in the cells that were pretreated with gabapentinoids were 0.04 ± 0.5 μm (pregabalin) and 0.4 ± 0.5 μm (gabapentin). 3.3.3 Imaging probe FluoBar1 is selective for CaV1.2 The selectivity of FluoBar1 was tested for CaV1.2 in U118-MG cells. Confocal fluorescence micrographs of fixed U118-MG cells showed that FluoBar1 has labeled CaV1.2 and colocalized with the anti-CaV1.2 antibody (Figure 3.4). Colocalization analyses between fluorescence signals observed in a blue (FluoBar1, Olympus AlexaFluor405 filter) and the red channel (anti-CaV1.2 / DyLight 550, Olympus Cy3 filter) returned a Pearson’s r value of 0.87 and a Costes P value of 1.00 (a P value above 0.95 confirms the accuracy of Pearson’s coefficient) (Table 3.1) [194, 195]. The Mander’s coefficient indicated > 99 % colocalization for both the blue and the red channels; this coefficient is the ratio of pixels from a channel that colocalized over the total signal detected from the same channel for a given cell area [196]. The analyses suggest that unselective labeling by FluoBar1 is negligible or similar to the CaV1.2 antibody. COS-7 cells served as a negative control for colocalization analysis because Western blotting demonstrated the absence of CaV1.2 and CaV1.3 in these cells (Figure 3.2.C). The results of colocalization were consistent with Western blots. Negative controls with COS-7 cells showed no fluorescence signal above background with neither anti-CaV1.2 nor FluoBar1 (Figure 3.4 second row). In contrast, transfected COS-7 cells with the gene encoding CaV1.2 pore-forming subunit showed fluorescence signals with both FluoBar1 and the CaV1.2 / DyLight550 antibody (Figure 3.4 third column). The cells were co-transfected with EGFP and CaV1.2 plasmids. COS-7 cells 75 displaying green fluorescence were chosen for image analysis. The level of colocalization in transfected COS-7 cells was lower than for the astrocyte model cells: a Pearson r value of 0.64 was calculated with high confidence (Costes P = 1.00). It shows, in transfected COS-7 cells, 64% of blue signals form FluoBar1 colocalized with red signals from the antibody against CaV1.2. Figure 3.4 Representative confocal fluorescence images of cells labeled with an anti-CaV1.2/DyLight550 antibody and FluoBar1. Imaging probe FluoBar1 is highly colocalized with the CaV1.2 antibody in U118-MG cells that natively express LTCCs, as well as in COS-7 cells transfected with CaV1.2. Nontransfected COS-7 do not express LTCCs and showed no signals either from the probe or the antibody. Immunostained fixed cells were treated with 50 nM FluoBar1 for 3 minutes, then imaged in HBSS; 60x magnification, Scale bar 10 µm. 76 Table 3.2 Colocalization analysis of confocal images. The fluorophores of FluoBar1 and anti-CaV1.2/DyLight550 colocalize with one another with high correlation. Cell lines Pearson’s r value Coste’s P value Mander’s coefficient 405 nm excite. 559 nm excite. U118 0.87 1.00 0.997 0.998 Transfected COS-7 0.64 1.00 1.00 0.677 3.3.4 Membranar density of CaV1.2 changed when U118-MG cells were exposed to glutamate Using FluoBar1, glutamate was shown to have an immediate effect on the density and distribution of CaV1.2 on the membrane in U118-MG astrocytoma cells. The novel imaging probe, FluoBar1 was applied to the live U118-MG astrocytoma cells which made the monitoring the localization of CaV1.2 proteins in real time possible (Figure 3.5). When the labelled cells were exposed to 100 µM glutamate, time-lapse and Z-stack confocal imaging revealed a gradual increase in the fluorescence density that leveled to a plateau after 10 minutes (from 5,297 ± 1,474 to 25,905 ± 2,085 AU/µm2, n = 3, Figure 3.5.A and 3.5.B). Images were acquired from the surface of the FluoBar1 stained cells and show the density of CaV1.2 on the membrane before and after exposure to glutamate (Figure 3.5.A and 3.5.C). Figure 3.5.A represent confocal images that were acquired from the lateral (x and y; in the specimen plane) of the cells surface, where the membrane over the nuclei was not in the focal plane of the view. U118-MG cells plated in culture dishes display a flattened soma compared to their nuclei in the axial (z; along the optical axis) images (Figure 3.5.B). The distribution of fluorescent signals on the membrane of the cells stained with either FluoBar1 (Figure 3.5.A) or with the selective antibody for CaV1.2 (Figure 3.5.D) changed from a compartmentalized 77 arrangement to a widespread distribution on the membrane when the cells exposed to glutamate (Figure 3.5.A and 3.5.D). Figure 3.5 The density and distribution of CaV1.2 changed when U118-MG cells were exposed to 100 µM glutamate. (A) Representative front view of confocal micrographs (from Z stack images) of a live cell labelled with 50 nM FluoBar1: prior to glutamate stimulation (left), and after signal stabilization 20 minutes post-stimulation (right). (B) Representative confocal Z Stack micrographs of a U118-MG cell. The front view of the cells (2D) on the left; and the side view of the cells based on constructed Z stack images (3D) on the right. These cells have a large nucleus and flat soma. Cells were exposed to glutamate for 15 minutes and stained with FluoBar1. (C) The data that were acquired from live cell with Z stack images were used for analyzing the change in the fluorescent intensity upon glutamate stimulation. Quantified fluorescence from FluorBar1 observed in living cells as a function of time; fluorescence density was measured in each cell and corrected with averaged background density (n = 3). (D) Representative confocal micrographs of two sets of fixed cells immunostained with an anti-CaV1.2/DyLight550 antibody. U118-MG cells were fixed without stimulation (control), or 10 min after glutamate exposure. All images were acquired at 60X magnification; All the scale bars are 20 μm. 78 Using immunocytochemistry, the U118-MG astrocytoma cells that had been fixed without any stimulation were compared to cells that had been fixed after exposure to glutamate. The same patterns of distribution were observed as in live cells stained with FluoBar1 before and after being stimulated with glutamate (Figure 3.5.C). U118-MG cells that were not exposed to glutamate showed a more localized expression of CaV1.2 channel than the cells that were exposed to 100 µM glutamate for 10 minutes prior to fixation. The stimulated cells consistently displayed a widespread distribution and denser exofacial expression of CaV1.2. Signals from CaV1.2 were localized with lower fluorescence density in U118-MG cells, while upon exposure to glutamate, signals were observed throughout the cell with increased fluorescent density. 3.4 Discussion LTCCs are voltage-dependent ion channels that are typically studied with electrophysiological approaches [90, 154]. Given their electrical activity, the role of LTCCs in normal function of non-excitable astrocytes in the brain has remained an enigma. However, their contribution in different neuropathological conditions have been reported [86, 97, 165, 177, 197, 198] Therefore, we tested the hypothesis that LTCCs participate in the voltage-independent phenomenon of glutamate-induced morphological response and results suggested that they contribute to this phenomenon in U118-MG astrocytoma cells. Four different isoforms of LTCCs have been identified (Figure 1.5). They possess similar pharmacological properties, but they differ when it comes to their tissue distribution and biophysical properties. In most electrically excitable cells, CaV1.2 and/or CaV1.3 are expressed and both isoforms are often even expressed in the same cell, such as in neurons and atrial cardiomyocytes [154, 156, 165, 176]. Both CaV1.2 and CaV1.3 are required for the normal function of the CNS [154], but, in the brain, CaV1.2 outnumbers CaV1.3 (90% : 10%) [188]. Unexpectedly, astrocytes that are non-excitable members of the CNS also express both CaV1.2 and CaV1.3 79 isoforms [85, 155, 197]. The role and function of LTCCs in astrocytes is unknown and remained to be elucidated. Interestingly, the role of CaV1.2 and CaV1.3 in migration and proliferation of cancer cells is emerging [99, 199]. Western blotting shows that, U118-MG astrocytoma cells that have been derived from glioma tumor and have been used in this study express CaV1.2 but not CaV1.3 (Figure 3.2 C). This expression profile of U118-MG cells is consistent with the expression profile of fully sequenced astrocytoma cells—U138-MG and U87-MG cells—in the “Human Atlas Proteome” database (https://www.proteinatlas.org/) that they also only express CaV1.2 (CACNA1C) [200, 201]. Pretreatment of U118-MG cells with nifedipine, verapamil and diltiazem (10 μM) at certain concentration, that its efficiency in blocking the calcium influx have been reported [91, 92, 171], decreased the process’s extension significantly, when U118-MG cells were stimulated with glutamate (Figure 3.2 A and B). The ability of the antagonists of LTCCs to modulate the extent of morphological response to glutamate suggest participation of the Ca2+ influx through LTCCs in supplying the required Ca2+ for subsequent cytoskeletal rearrangement. However, whether this effect of the blockers is dose-dependent remained to be determined. The concentrations that were used in this study were based on the concentrations that have been repeatedly reported as an effective concentration to block CaV1.2 currents (Table 3.1) [95, 98, 137, 186]. In fact, the role of the concentration of the blockers would not affect the interpretation about the participation of LTCCs in glutamate-induced morphological response in U118-MG astrocytoma cells but the extent of this response could be different at higher concentrations. It should also be stressed that the activity of LTCCs in response to glutamate might be the consequence of initial activation of a glutamate receptor(s), that has been led to a change in the membrane potential of the cells and has triggered gating of LTCCs. Typically, when LTCCs are involved in a cellular event they translocate on the cell membrane instantly to align with the other elements that are involved in that specific cellular response [169, 80 185]. Since with the conventional laboratory techniques real-time tracking of LTCC on the membrane without manipulating their activity was not possible, the FluoBar1 imaging tool was developed. As mentioned above, the synthetic process and chemical validation of FluoBar1 has been detailed by Tabatabaee et al [185] and has briefly shown in appendix D. FluoBar1 was created for optical imaging applications, therefore, its selectivity was validated by a colocalization analysis of confocal images that were stained with both FluoBar1 and selective antibodies. The fluorescence signals from FluoBar1 were highly colocalized with immunocytochemical labels in U118-MG cells that natively express CaV1.2 (Figure 3.4, Table 3.2). As expected, no signal from either channels were detectable in the CaV-free COS-7 cells [119, 202]. This negative control shows that (1) no non-selective CaV1.2 labeling occurs with the FluoBar1 and the anti-CaV1.2 antibody; and (2) no passive diffusion of FluoBar1 takes place through the cell membrane within the incubation time used in this experiment (Figure 3.4). However, colocalization analysis of transfected COS-7 cells with CaV1.2 suggest that FluoBar1 binds only to properly assembled CaV1.2s. This is supported by a Mander’s analysis in which 100% of signals from FluoBar1 colocalized (405 nm excitation) with signals from anti-CaV1.2 fluorescent antibodies (559 nm excitation). While one third of the signals acquired from the antibody-labeled proteins were not labeled with FluoBar1. It shows that a signal from antibody presents wherever FluoBar1 has been labeled; but there are peptides that were labeled by an antibody where FluoBar1 could not bind to. The difference between the recorded signals from the immunostains and the chemical imaging probe is likely due to the different binding modes of these two imaging tools. The anti-CaV.1.2 antibody used herein has affinity to a short peptide sequence between second and third internal loop of the channel (residues 848-865, Alomone, Figure 3.1); this epitope is the same regardless to the protein's folding, location, or activity of the channel. In contrast, FluoBar1 requires a fully folded and assembled channel on the membrane to bind, as it is designed based on a pore-forming subunit blocker. In COS-7 cells, the exogenous expression of the large CaV1.2 protein is 81 subjected to misfolding and can lead to improper assembly or trafficking [203]. This may explain why less fluorescence from FluoBar1 was observed compared to the signals from anti-CaV1.2 antibody in transfected COS-7 cells. To monitor the dynamic of CaV1.2 on the membrane of U118-MG cells, the selectivity of FluoBar1 for only CaV1.2 has been validated. These cells express CaV1.2 and not CaV1.3 (Figure 3.2.C). The parent barbiturate for FluoBar1 has been reported to bind to both CaV1.2 and CaV1.3 [183, 184]. In addition, in a complementary experiment, where FluoBar1 was used as a preliminary screening tool to rapidly detect the presence of LTCCs in a non-denatured cell lysate (Appendix E), FluoBar1 was found to label CaV1.3 as well in MCF7 cells, that only express CaV1.3 [200]. The selectivity of FluoBar1 for CaV1.3 for future applications needs further investigation. In fact, a rapid increase in the membranar density of CaV1.2 upon exposure to glutamate was observed using FluoBar1. However, the main function of Ca2+ influx through CaVs after the rise in their surface expression in the glutamate signaling of U118-MG cells remains to be identified. To further study the role of CaVs (CaV1.2 in U118-MG cells) in glutamate signaling, participation of the extracellular subunit, α2δ, was also examined. Thus far, four subtypes of α2δ has been identified (α2δ1-4) [90, 204]. The α2δ1 subtype associates with CaV1.2 calcium channels [177, 205]. Unexpectedly, pretreatment with selective ligands for α2δ1, the gabapentinoids, inhibited the morphological response to glutamate (Figure 3.3.). The inhibitory effects of the ligands for α2δ1 were not expected, because one of the established functions of α2δ1 is modulating the Ca2+ conductance through the pore-forming subunit [164, 170, 184, 206] and blocking Ca2+ conductance by CaV antagonists did not inhibit the filopodiagenesis, Alternatively, filopodial outgrowth was probable due to the Ca2+ influx by nonselective effect of gabapentinoids, which could support polymerization of the actin cytoskeleton and consequent filopodiagenesis. Yoshizumi et al. have proposed that gabapentinoids can enhance sodium/glutamate cotransport when cells are exposed to glutamate [44]. To adjust the consequent intracellular excessive 82 sodium, the Na+/Ca2+ exchangers impose an intracellular calcium rise to the astrocytes [44]. This unexpected inhibitory effect of gabapentinoids on glutamate-induced processes extension in U118-MG cells suggests a different and currently undefined mechanism of action for gabapentinoids. This notion has been investigated further in the next chapter. In summary, the results of this chapter suggest that (1) LTCCs contribute to U118-MG glutamate signaling, (2) glutamate has an immediate effect on CaV1.2 distribution and localization and (3) α2δ1, the CaV1.2’s extracellular subunit, is essential to the glutamate-induced filopodia extension. These findings led me to investigate the role of α2δ1 further in the morphological features of glutamate signaling events in astrocytoma cells, which will be discussed in the next chapter. 83 Chapter 4. The α2δ1 Contributes to the Glutamate-Induced Morphological Response and is Independent from Voltage-Gated Calcium Channels In the previous chapter the selective ligands for the extracellular subunit of CaVs, α2δ1, were found to inhibit the morphological response of astrocytoma cells to glutamate. In addition, real-time imaging in U118-MG astrocytoma cells with FluoBar1 probe revealed that glutamate causes an instant change in the density and distribution of CaV1.2 on the membrane of U118-MG astrocytoma cells. It is known that the membranal expression of the CaVs is directed by α2δ1 subunit [170, 204]. Therefore, this chapter is focused on the role of α2δ1 and aims to test the hypothesis that α2δ1 is an essential component in the morphological response that is induced by glutamate in astrocytoma cells. A selective antibody for α2δ1 had the same inhibitory effect on the processes extension of U118-MG astrocytoma cells as had been observed with the chemical ligands for α2δ1 (gabapentinoids). These observations suggested the participation and role of α2δ1 in initiation of the morphological response to glutamate. Immunocytochemistry demonstrated that upon exposure to glutamate, α2δ1 proteins translocate to the filopodia. Interestingly, in heterologous expression systems α2δ1 endow the ability to respond morphologically to glutamate to transfected cells. U138-MG cells are a type of astrocytoma cells who do not exhibit a significant extension in their processes upon exposure to glutamate. Overexpressing α2δ1 made these cells responsive to glutamate. These cells demonstrated a large peak in Ca2+ influx and subsequent process extensions after transfection with α2δ1. Since α2δ1 is known as an auxiliary subunit of CaVs, to identify whether α2δ1 can initiate the morphological response to glutamate independent from CaVs, a CaV-free cell line, COS-7, was used. Transfection with α2δ1, transformed COS-7 to the cells that respond to glutamate and show both filopodiagenesis and rise in the intracellular Ca2+. 84 These results suggest a significant role for α2δ1 protein in filopodiagenesis regarding glutamate signaling. 4.1 Background The α2δ proteins were initially described as auxiliary subunits for CaVs [180, 207]. When associated with CaVs, the α2δs has been reported to have three main functions: (1) it assists trafficking of the channels complex to the plasma membrane, which is a key step in regulating calcium influx [147, 148]; (2) it dictates the localization of CaVs into the lipid rafts of the membrane [170, 207, 208]; and (3) it mediates increased peak of Ca2+ influx through L-type CaVs and CaV1.2 in particular, both natively and in heterologous expression systems [177, 180, 209]. The α2δ protein is translated from a single gene product, which gets post-translationally cleaved into an α2 and a δ fragment that remain associated via disulfide bridges [208, 210]. It is categorized into four classes based on the δ subunit (α2δ1-4) [208]. The α2 part of the protein (~950 amino acids) is entirely extracellular and contains a Von Willebrand Factor A (VWF-A)—a protein-protein interaction domain—along with four bacterial chemotactic domains (Cahe) [156, 211, 212]. The presence of these domains suggests that α2 may act as a receptor for extracellular ligands (Figure 4.1). The first identified synthetic ligands for α2δ proteins were gabapentinoids. Gabapentin and pregabalin suppress seizures and are approved as a class of anti-epileptic and anti-allodynic drugs [174–179]. However, how they carry out their effects through α2δ is not fully known. Thrombospondins (TSPs) are endogenous soluble ligands for the α2δ proteins [211]. TPS are signals from astrocytes that promote synaptic adhesion at excitatory synapses and initiate the events that lead to the establishment of pre- and postsynaptic specializations [213, 214]. In 85 addition to the soluble TSPs, some membrane proteins, such as the N-terminus region of BK potassium channels [175] and synaptogenic mediator, ELFN1, also bind to α2δ [215] . At a tripartite synapses, where an astrocytic process, a presynaptic and a postsynaptic neuron are active, the α2δ1 isoform contributes dynamically to the regulation of functionality of the synapse [2, 6, 207, 216]. Presynaptically, it modulates neurotransmitter release via interacting with α-neurexins and by increasing intracellular calcium concentration [163]. Postsynaptically, it promotes surface expression of the α2δ1-NMDA receptor complexes that are required to initiate the action potential [191, 217]. It is also essential for formation and/or maturation of dendritic spines [209]. However, the role that α2δ1 plays in the astrocytes at a tripartite synapse has remained unexplored. In addition, whether α2δ1 play any role in astrocytoma cells and the glioma progression in the brain is also unknown. While the contribution of α2δ1 in the lung and gastric cancer is emerging and has been shown to have a significant role in the invasion and progression of these cancer, their role and function in brain cancers has not been investigated [205, 218, 219]. Figure 4.1 Structure of the α2δ1. α2δ is translated from a single gene product, which gets post-translationally cleaved into an α2 and a δ fragments that remain associated via disulfide bridges. The α2 part of the protein (~950 amino acids) is entirely extracellular and contains a Von Willebrand Factor A (VWF-A) along with four bacterial chemotactic domains (Cache). Gabepentin (GB) and pregabalin (PGB) are proposed to bind to the Argenin repeats upstream VWA and thrombospondins (TSB) bind to the metal ion-dependent adhesion site (MIDAS) of the VWA domain. Created with Biorender.com 86 The inhibitory effect of gabapentinoids on the morphological response that is induced by glutamate in the U118-MG astrocytoma cells, as shown in Chapter 3 [137], suggested the contribution of α2δ1 in glutamate signaling events. In this chapter, using pharmacology, imaging, and heterologous expression systems, further evidence for involvement of α2δ1 in morphological response to glutamate has been provided [30]. Results suggest that α2δ1 is the link between glutamate sensing, Ca2+ influx and processes extension in the glutamate signaling pathway. 4.2 Material and methods 4.2.1 Cell culture Human astrocytoma U118-MG cell line (ATCC HTB-15TM), COS-7 cell line (ATCC CRL-1651TM) and U138-MG glioblastoma (ATCC HTB-16TM) cells were cultured in Dulbecco’s modified essential medium (DMEM, Gibco 11995-065) supplemented with 10% v/v heat-inactivated fetal bovine serum (HyClone 12483020) 1% v/v penicillin 10,000 U/ml and streptomycin 10,000 μg/ ml (Gibco 15140-163). Cells were incubated in a humidified atmosphere containing 5% CO2 at 37 ºC. They typically passaged at 80% confluency using a pre-warmed 0.25% trypsin-EDTA solution (Gibco SH30236.02) for a maximum of 5 minutes. 4.2.2 Transfection To study the role of α2δ1 in the initiation of calcium signaling, COS-7 and U138-MG cells were transfected with α2δ1 plasmid. The α2δ1 plasmid was a gift from Diane Lipscombe (Addgene 26575) [166]. To monitor the effect of α2δ1 in real time morphological response, cells were co-transfected with both mCherry-Lifeact-7 (Addgene plasmid 54491) and CaV-α2δ1 plasmids (1200 ng/ μl of each). The mCherry-Lifeact-7 plasmid was a gift from Michael Davidson (Addgene 54491) and used to stain actin to monitor the fine movements and extensions of astrocytic processes in live cells. Transfected cells that had ramified membrane and high intensity of mCherry fluorescent signals expression were chosen for imaging. Calcium phosphate 87 precipitation transfection technique was used for both experiments, as explained in the previous chapter (Section 3.2.2). 4.2.3 Treating cells with 2δ1 ligands The effect of 100 μM of glutamate (Alfa Aesar A15031-30) on the morphological response of U118-MG cells was studied when cells were pretreated with 2δ1 ligands and compared to the control. A set of mCherry-Lifeact-7 transfected U118-MG cells were pretreated with 10 μM of gabapentinoids before exposure to glutamate (as described in Section 3.2.3). Gabapentinoids: gabapentin (G154, Sigma Aldrich), pregabalin (made in house) were prepared directly in deionized millipore water (vehicle). Stock solutions were prepared concentrated (200X) to avoid osmotic shock. Before imaging, cells were incubated at 37 ºC under a humidified 5% CO2 atmosphere for 30 minutes. Another set of cells were pretreated with antibodies selective for α2δ1. Cells were incubated with 1:50 Anti-CACNA2D1 (CaV-α2δ1 extracellular Antibody, Alomone ACC-015) in HBSS (Lonza 10-547F) supplemented with 2 mM CaCl2 (Sigma Aldrich C1016) for 15 minutes prior to exposure to glutamate. Glutamate (Alfa Aesar A15031-30) was delivered to the antibody-treated cells four minutes after beginning the imaging. The same procedure was used for COS-7 and U138-MG cells, both for α2δ1-transfected and non-transfected cells. 4.2.4 Confocal imaging Morphological response of the cells was recorded by confocal time-lapse imaging. Cells were imaged every 20 second for a total number of 60 frames on an Olympus FV1000 fluorescence confocal microscope equipped with a Plan-ApoN 60x/i.4 oil-immersion objective Photobleaching was minimized by irradiating samples for 1.0 seconds at 20 second intervals with the laser power set at 3%. Fluorescence of mCherry-Lifeact-7 was observed at 590 nm (Olympus filter set Cy3.5), 88 with 559 nm excitation from a He–Ne laser. FluoBar1 (CaV1.2 labeling) labeled cells were imaged at 405 nm excitation using a He–Ne laser (Olympus filter set Alexa-Fluor 405). 4.2.5 Calcium imaging with Fluo4-AM To identify the role of α2δ1 on intracellular Ca2+ rise upon glutamate stimulation, the Ca2+ transients were recorded using Fluo4-AM as described in Section 2.2.8 in U138-MG and COS-7 cells both non-transfected and α2δ1-transfected as well as in U118-MG cells as a control. 4.2.6 Quantification of processes extension The change in a cell process length was measured as described in Section 3.2.5. 4.2.7 Immunocytochemistry (ICC) to label α2δ1 The α2δ1 was immunostained to identify its localization in U118-MG cells that were fixed without or after exposure to glutamate. U118-MG cells were labeled with anti-α2δ1 antibody (Alomone ACC-015) in two different sets. In one set, the cellular actin cytoskeleton was initially labeled by transfecting cells with mCherry-Lifeact-7 (red) as described (Section 4.2.2). To have a contrast between fluorescent signals in these transfected cells, goat anti-rabbit IgG Alexafluor488 conjugated (Abcam 150077) (green) was used for labeling α2δ1, as the secondary antibody. While in non-transfected cells goat anti-rabbit IgG (H+L) conjugated to DyLight 550 (Thermoscientific PA5-84541) (red) was used as the secondary antibody to label the α2δ1 proteins. For ICC, plated cells were washed with phosphate buffer saline (PBS) solution, then fixed in 4% paraformaldehyde in PBS solution for 20 min at room temperature. The fixed cells were incubated in a blocking solution (10% BSA in PBS), followed by overnight incubation at 4 °C with the primary antibody, extracellular Anti-CACNA2D1 (α2δ1) (Alomone ACC-015) in 3% BSA in PBS (1:200 dilution). After three rinses with PBS: non-transfected cells were incubated with a 1:200 dilution of the secondary antibody goat anti-rabbit IgG (H+L) conjugated to DyLight 550 89 (Thermoscientific PA5-84541) at room temperature on a rocking platform for 2 hours. Transfected cells were treated with Goat anti-rabbit IgG Alexafluor488 conjugated (Abcam 150077). Cells were rinsed with PBS for 3 times and imaged in PBS. 4.2.8 FluoBar1 staining to label CaV1.2 Live U118-MG astrocytoma cells were incubated with 50 nM of FluoBar1 in PBS solution at room temperature for 2-5 minutes [123]. FluoBar1 was replaced by 2.0 ml Hank's balanced salt solution (HBSS, Lonza 10547F) supplemented with 2 mM CaCl2 (Sigma Aldrich, C1016) before imaging. 4.2.9 Image analysis The change in the membranar density of α2δ1 upon exposure to glutamate was acquired from confocal images of mCherry-Lifeact-7 transfected U118-MG cells that were immunostained for α2δ1 protein. Images were analyzed with ImageJ software (NIH ImageJ, FIJI). To report the change in the density of fluorescent signals from α2δ1, the data were normalized to the fluorescent signals from actin (mCherry-Lifeact-7). The cells' bodies were outlined manually, and their total fluorescence signal was integrated to calculate the fluorescence density of α2δ1. Background fluorescence was measured separately for mCherry-Lifeact-7 and the anti-α2δ1 fluorescent signals by selecting three cell-free areas from the same image and integrating their signals. Then their densities were averaged. The corrected total cell fluorescence density (CTCF) was calculated for actin—mCherry-Lifeact-7—and the anti-α2δ1 (AlexaFluor 488) separately as follows. 𝐶𝑇𝐶𝐹 = 𝑖𝑛𝑡𝑒𝑔𝑟𝑎𝑡𝑒𝑑 𝑑𝑒𝑛𝑠𝑖𝑡𝑦 𝑜𝑓 𝑎 𝑐𝑒𝑙𝑙 − 𝑖𝑛𝑡𝑒𝑔𝑟𝑎𝑡𝑒𝑑 𝑑𝑒𝑛𝑠𝑖𝑡𝑦 𝑜𝑓 𝑏𝑎𝑐𝑘𝑔𝑟𝑜𝑢𝑛𝑑 Eventually, the α2δ1 fluorescence density was normalized to actin for each image in two steps. First, an image normalization factor was calculated from the red channel (actin) using equation A. Then, signals for the anti-α2δ1 (green) channel were normalized using equation B. The data from 90 equation B were processed and plotted using Prism 7.04 from GraphPad. Each value represents the averaged changes measured from five cells in five separate culture dishes; the results are presented as the mean ± SEM. 𝐴: 𝐼𝑚𝑎𝑔𝑒 𝑁𝑜𝑟𝑚𝑎𝑙𝑖𝑧𝑎𝑡𝑖𝑜𝑛 𝐹𝑎𝑐𝑡𝑜𝑟 (𝐼𝑁𝐹) =𝑂𝑏𝑠𝑒𝑟𝑣𝑒𝑑 𝐶𝑇𝐶𝐹 𝑜𝑓 𝑎𝑐𝑡𝑖𝑛 (𝑟𝑒𝑑) 𝑓𝑜𝑟 𝑒𝑎𝑐ℎ 𝑖𝑚𝑎𝑔𝑒𝐻𝑖𝑔ℎ𝑒𝑠𝑡 𝑜𝑏𝑠𝑒𝑟𝑣𝑒𝑑 𝐶𝑇𝐶𝐹 𝑜𝑓 𝑎𝑐𝑡𝑖𝑛 𝑜𝑓 𝑎𝑙𝑙 𝑖𝑚𝑎𝑔𝑒𝑠 𝐵: 𝑁𝑜𝑟𝑚𝑎𝑙𝑖𝑧𝑒𝑑 𝐸𝑥𝑝𝑒𝑟𝑖𝑚𝑒𝑛𝑡𝑎𝑙 𝑆𝑖𝑔𝑛𝑎𝑙 (𝑁𝐸𝑆) =𝑂𝑏𝑠𝑒𝑟𝑣𝑒𝑑 𝑒𝑥𝑝𝑒𝑟𝑖𝑚𝑒𝑛𝑡𝑎𝑙 𝑠𝑖𝑔𝑛𝑎𝑙 (𝑔𝑟𝑒𝑒𝑛)𝐼𝑁𝐹 To assess if there is any correlation between the change in fluorescence density of α2δ1 and CaV1.2, images of non-transfected U118-MG cells that were stained with FluoBar1 for real-time monitoring were used. The change in fluorescence density of FluoBar1 (CaV1.2) was also measured with FIJI calculating CTCF before and after exposure to glutamate [193] and plotted using Prism 7.04 from GraphPad. Each value represents the averaged changes measured from three separate culture dishes; the results are presented as the mean ± SEM. 4.2.10 SDS-PAGE and Western blotting SDS-PAGE and Western blotting was done as described in Section 2.2.15. The primary antibody was extracellular Anti-CACNA2D1 (α2δ1) (Alomone ACC-015) and horseradish peroxidase-conjugated GE Healthcare) was used as the secondary antibody. The blots were analyzed solely qualitatively, and no normalization or quantification was done. 4.2.11 Statistical analysis Significant differences among groups were determined using one-way or two-way analysis of variance (ANOVA) and Tukey’s post hoc analysis as well as unpaired two-tailed T-test using Prism 7.04 from GraphPad. A p value < 0.05 was considered significant. Each value of the change in the process's length represents the averaged changes measured from at least 5 cells, each from a separate culture dish, independently transfected (n ≥ 5); the results are presented as the mean ± SEM. 91 The change in fluorescence density before and after exposure to glutamate was compared by two-tail T test (paired for live cells or unpaired for fixed cells) using Prism 7.04 from GraphPad. Differences between the process length of transfected and non-transfected COS-7 cells were also compared by two-tailed unpaired T Test. A p value < 0.05 was considered significant. 4.3 Results 4.3.1 Chemical and immunological ligands for α2δ1 inhibited the glutamate-induced morphological response in U118-MG astrocytoma cells As shown in Figure 3.3, when U118-MG c ells were pretreated with α2δ1 ligands (gabapentinoids) prior to stimulation with glutamate, no significant difference in the process's Δ Figure 4.2 antibody for α2δ1 inhibited the morphological response of U118-MG cells to glutamate. (A) Representative confocal micrographs of U118-MG cells, before and after exposure to 100 M glutamate. Cells were transfected with mCherry-lifeact, for live imaging of actin cytoskeleton (red) and pretreated with 1:50 anti- α2δ1 before stimulation with glutamate. 60X magnification, 2X zoom. Scale bar 5 μm. (B) Change in the processes length of the cells that were not treated (control) or pre-treated with α2δ1 ligands was compared when cells were stimulated with 100 µM glutamate. The data were compared using Tukey’s post hoc multiple comparison (two-way ANOVA F1,16 α2δ1 antibody = 8.05 ; F 1.16 glutamate treatment = 56.72); n = 5; *p<0.0001 significantly different from stimulated cells with glutamate while pretreated with α2δ1 ligands. #p<0.0001 significantly different from untreated and unstimulated cells Data are shown ± SEM. (C) Western blots show that both U118-MG cells and U138-MG cells express α2δ1; Cell lysates were blotted and imaged for α2δ1 with chemiluminescence. α2δ1 peptides (Alomone) was used as positive controls and COS-7 cell lysate was used as a negative control. (C) The α2δ1 peptide served as a positive control for Western blots was provided by Alomone. Western blots show that α2δ1 is detectable in U118-MG cells and U138-MG cells but not COS-7 cells. 92 length was observed (Figure 4.2.B). That result suggested the inhibitory effect of α2δ1 ligands on glutamate-induced morphological response in U118-MG astrocytoma cells. Likewise, anti-α2δ1 antibody inhibited the further extension of the processes by glutamate and only 0.04-fold change in length was observed (Figure 4.2.A). The selectivity of the Anti-CACNA2D1 extracellular Antibody (CaV-α2δ1) (Alomone ACC-015) was confirmed by Western blotting (Figure 4.2.C). It should be noted that, applying an antibody against α2δ1 (Alomone ACC-015) solely, induced a change in the Δ length of the processes in U118-MG astrocytoma cells by 5.5 times compared to untreated cells (from 0.28 μm to 1.54 μm). 4.3.2 Glutamate induced a change in the distribution and localization of α2δ1 on the membrane. Glutamate induced a change to the distribution of α2δ1 on the membrane as well as a translocation from soma to the filopodia. Exposure to 100 µM glutamate for 15 minutes before immunocytochemistry (ICC) caused the α2δ1 proteins to be redistributed from the membrane to the peripheries of the cells (Figure 4.3.A). A closer examination of the processes of U118-MG cells that transfected with mCherry-Lifeact7 showed that the α2δ1 proteins translocated to the processes of the cells when they were exposed to glutamate for 15 minutes before ICC. α2δ1 was absent on the U118-MG cells filopodia, which formed in the absence of glutamate before fixation (Figure 3.4.B first column). Exposure to glutamate for 15 minutes before fixation translocated α2δ1 on both the formation and the extension of the filopodia (Figure 4.3.B second column). Fluorescent signals from anti-α2δ1 antibody (Alexa 448) were 13.15 times more intense on the periphery of U118-MG cells that were treated with 100 μM glutamate for 15 minutes in comparison to the control cells. The fluorescent signals from the anti-α2δ1 antibody were 83.48 ± 25.71 AU/μm2 in untreated cells compared to 1098 ± 169.30 AU/ μm2 in glutamate-treated cells (mean difference of 1015 AU/ μm2) (Figure 4.3.C). 93 Figure 4.3 Exposure to glutamate changed the membranar density and localization of α2δ1 in U118-MG astrocytoma cells. (A) Immunostained micrographs of U118-MG cells for α2δ1 with secondary antibody conjugated with Dylight 550 (red). α2δ1 distributes evenly on the cell membrane of U118-MG cells (a). Membranar distribution of α2δ1 changed when the cells were exposed to 100 μM of glutamate for 15 minutes before fixation (b). sale bars 10 μm (B) Confocal micrographs of U118-MG cells immunostained for α2δ1. Cells were transfected with mCherry-lifeact, for live imaging of actin cytoskeleton (red). The U118-MG cells were immunostained for α2δ1 with secondary antibody conjugated with Alexafluor488 (green). The cell ‘b’ has been fixed 15 minutes after exposure to glutamate. Blue arrows show the filopodia. There is no α2δ1 in the filopodia of the cell ‘a’. In the cell ‘b’ that has been exposed to glutamate for 15 minutes, α2δ1 has translocated to the filopodia. 100X magnification 5X zoom. Scale bar 1μM. (C) change in the fluorescent density of immunostained α2δ1 (secondary antibody is conjugated with Alexafluor488 in green) in the cells with mCherry lifeact7 labeled actin (in red) that were exposed to glutamate for 15 minutes before fixation compared to control. Fluorescence density was measured in each cell and corrected with averaged background density (n = 3). Results are normalized to actin protein expression. Data were compared with unpaired two tailed T test. *p < 0.05 significantly different from the control. Data are shown mean ± SEM 94 4.3.3 Overexpressing α2δ1 significantly enhanced the response of U138-MG cells to glutamate stimulation U138-MG cell line is a type of astrocytoma cell that is regularly used in neuroscience and neurooncological studies [137, 167]. However, the results in Chapter 2 revealed that 100 μM glutamate does not induce a measurable morphological response in U138-MG cells. Intriguingly, Figure 4.4 Morphological response and the calcium influx in U138-MG cells raised significantly upon transiently transfection with α2δ1. (A) Representative confocal micrographs of cells transfected with α2δ1before and after exposure to 100 M glutamate. Cells were co-transfected with mCherry-lifeact, for live imaging of actin cytoskeleton (red). 60X magnification; Scale bar 10 μm, (B) Comparing normalized fluorescence of Ca2+ imaging with Fluo4-AM dye in U118-MG and U138-MG cells stimulated with 100 glutamate. n = number of cells / number of dishes (C) Change in a processes’ length of U118-MG, COS-7 and α2δ1-transfected COS-7 cells stimulated with 100 µM glutamate. Tukey’s multiple comparison post hoc was used to compare the glutamate treated cells with untreated cells (two-way ANOVA F 1, 24 glutamate treatment = 131.0; F 2,21 Transfected and non-transfected U138-MG= 34.4); n = 5; *p < 0.0001 significantly differ from U138-MG cells and the cells that were not stimulated with glutamate. Data are shown mean ± SEM. (D) Normalized fluorescence of Ca2+ imaging by Fluo4-AM in U118-MG, COS-7 and α2δ1-transfected COS-7 cells stimulated with 100 glutamate. n = number of cells / number of dishes 95 transiently transfecting these cells with α2δ1 significantly enhanced their process extension and Ca2+ influx in response to glutamate. U138-MG expresses both CaV1.2 and α2δ1 (Figure 3.2.C and 4.2.C). Overexpression of α2δ1 by transiently transfecting U138-MG cells caused a 4.84-fold increase in the Δ length of their processes in response to glutamate. The mean difference in the change in the length of the processes before and after glutamate stimulation in non-transfected U138-cells was only 0.18 μM compared to 4 μM when they were transfected with α2δ1 (Figure 4.4.A and B). In addition, the peak of the calcium rise was significantly higher when U138-MG cells were transfected with α2δ1 compared to the native response of these cells to glutamate. The Ca2+ rise in U118-MG cells there was a rapid calcium transient upon glutamate stimulation, but α2δ1 transfected U138-MG cells demonstrated a slower retention time (Figure 4.4.D). 4.3.4 COS-7 cells responded to glutamate when they transiently expressed α2δ1 To define whether α2δ1 can induce a morphological response to glutamate in a cell that is not differentiated for glutamate signaling and does not express CaVs, COS-7 cells were selected. Transfected COS-7 cells with α2δ1 displayed a more ramified membrane and extended their processes with a Ca2+ influx upon exposure to glutamate. COS-7 is a kidney fibroblast cell line from Cercopithecus aethiops. It is a calcium channel free cell line and is regularly used as a heterologous expression system for calcium channel studies [166, 202]. They do not have a ramified membrane and show no response to glutamate (Figure 4.5 A). They do not express CaV1.2 and CaV1.3 (Figure 3.2.C). The extracellular subunit of these channels, α2δ1, is also absent (Figure 4.2.C). When COS-7 cells were transiently transfected with α2δ1, 52.4% of the cells showed more processes on their membrane compared to normal COS-7 cells. This percentage of response in the cells after transfection is more likely due to the 96 efficiency of Ca2+ precipitation transfection technique. The average length of the processes was Figure 4.5 Transiently transfecting COS-7 cells with α2δ1 causes processes extension and intracellular Ca2+ rise in response to glutamate. (A) Representative confocal micrographs of the COS-7 cells. Cells were transfected with mCherry-lifeact, for live imaging of actin cytoskeleton (red). The second row of the cells were co-transfected with α2δ1. Cells are shown before and after exposure to 100 M glutamate. 60X magnification, 2X zoom. Scale bar 10 μm, (B) Transfection with α2δ1 has raised the average length and the number of processes in COS-7 cells compared to non-transfected COS-7 cells. Transfection reagent contained CaCl2 2.5 M (5% v/v) and HEPES (50% v/v) (without the plasmid). The COS-7 cells that were not transfected were compared with α2δ1-transfected cells in the absence of glutamate. Comparison was done using two-tailed unpaired T test for transfected and non-transfected COS7. n = 5; *p < 0.05 is significantly different from non-transfect cells. Data are shown mean ± SEM . 97 0.84 ± 0.08 in non-transfected cells and was 3.34 ± 0.37 in transfected cells (4-folds increase) (Figure 4.4.E). When α2δ1 transfected cells were exposed to glutamate, the increase in the average length of their processes was 10.6 times, from 0.24 ± 0.16 µM to 2.56 ± 0.3 µM over the course of 15 minutes (Figure 4.5.A and Figure 4.5.B). This response was blocked by selective ligands for α2δ1: gabapentin and pregabalin (Figure 4.5.C). Unexpectedly, the transfected COS-7 cells with α2δ1 demonstrated a Ca2+ spike upon exposure to glutamate (Figure 4.5.D). The pattern of the calcium transients in transfected COS-7 cells with α2δ1 differ from what was observed in U118-MG cells. In U118-MG cells, there was a large intracellular calcium rise upon exposure to glutamate; α2δ1-COS-7 cells showed a peak followed by two smaller peaks within 20 seconds from each other. It should be noted that, only 46% of transfected α2δ1-COS-7 cells reacted to glutamate by a calcium spike, which indicates the efficiency of calcium phosphate precipitation transfection technique. Only the cells that have shown the Ca2+ peak have been reported in Figure 4.5.D. The effect of co-expression of α2δ1 with CaV1.2 in COS-7 cells was also tested (data not shown). The co-expression of α2δ1 with CaV1.2 caused an extremely ramified membrane with abnormal morphology and highly motile processes that prevented collecting any conclusive data. Continue figure 4-5 (C) Change in a processes’ length of U118-MG, COS-7 and α2δ1-transfected COS-7 cells before and after stimulation by 100 µM glutamate. Tukey’s multiple comparison post hoc (two-way ANOVA; F 1,24 glutamate treatment = 156.1; F 2,24 cell type = 61.23); # p < 0.0001 significantly different from COS_7/ α2δ1 upon exposure to glutamate, *p < 0.0001 significantly different from the COS-7 cells that were exposed to glutamate and the cells that were not exposed to glutamate. n = 5 (D) Normalized fluorescence of Ca2+ imaging dye Fluo4-AM in U118-MG, COS-7 and α2δ1-transfected COS-7 cells stimulated with 100 glutamate. n = number of cells/ number of dishes (E) 10μM of gabapentin (GP) and pregabalin (PG) inhibited the processes extension of α2δ1-transfected COS-7 cells in response to glutamate. Data were compared by Tukey’s multiple comparison post-hoc test (two-way ANOVA; F 2,24 treatments = 53.89; F 2,24 cells = 65.85); *p < 0.0001 significantly different from COS-7 cells and COS-7/ α2δ1 pretreated with gabapentinoids; n = 5. Data are shown mean ± SEM 98 4.1 Discussion In this chapter, α2δ1 was found to participate in glutamate-induced cellular responses both in a native system and in heterologous expression systems. Results suggest that α2δ1 links the sensing of glutamate with the intracellular Ca2+ rise and processes extension in glutamate signaling. Selective ligands for α2δ1 inhibited these cellular responses to glutamate in U118-MG astrocytoma cells that normally extend their processes in a Ca2+-dependent manner upon exposure to glutamate. The glutamate-induced response in astrocytes is usually attributed to glutamate receptors that can either conduct Ca2+ or release it from internal stores [4]. In normal astrocytes contribution of AMPARs and mGluR5 to glutamate signaling have been well studied and demonstrated [3, 18, 53, 54]. In fact, early studies have shown the effect of KA in inducing processes extension in cultured astrocytes but the role of KARs in this phenomenon has not been studied in detail [25, 75]. The results of Chapter 2 of this thesis added kainate receptors to the list of active glutamate receptors that can participate in the morphological response of the glutamatergic events in astrocytoma cells [60, 70]. Nevertheless, the robust and unexpected effect α2δ1 antagonists in inhibiting processes formation in response to glutamate suggests either its previously unknown interaction with one of the known glutamate receptors or another unidentified pathway through which α2δ1 contributes to the glutamate signaling. As we reported in Chapter 2, 100 μM glutamate did not induce any significant change in the process extension and the Ca2+ influx in U138-MG cells that are a type of astrocytoma cell line and are well-established model cells for astrocytic studies (Figure 4.4). 100 μM is a concentration of glutamate that has repeatedly been shown that triggers morphological response in astrocytes in vitro and we have shown that it induces processes extension in U118-MG cells [3, 25, 42, 137, 138]. U138-MG cells express active AMPARs, Ca2+-permeable KARs and different types of CaVs, as the Human Protein Atlas shows [200, 201, 220]. They also express a low level of α2δ1 (Figure 99 4.2.C) [201, 220]. Interestingly, overexpression of α2δ1 in these cells enhanced their Ca2+ influx and caused a significant change in their process extension in response to glutamate (Figure 4.4). The initial interpretation about the enhanced Ca2+ influx upon transfection with α2δ1 in U138-MG cells was prototypical association of α2δ1 as an auxiliary subunit with CaVs that could increase the Ca2+ conductance. This consequent Ca2+ elevation can lead to the actin polymerization and the processes extension, which was observed in transfected U138-MG cells with α2δ1.However, induction of an intracellular Ca2+ rise and processes extension by glutamate in the CaV-free COS-7 cells upon transfection with α2δ1 questioned this interpretation. COS-7 cells showed an intracellular calcium peak followed by two other smaller peaks in response to glutamate when transfected with α2δ1. These cells do not express CaVs, thus, they are the most commonly used expression system to study these channels [119, 202]. Therefore, in COS-7 cells association of α2δ1 with CaVs that leads to an intracellular Ca2+ rise and its consequent processes extension is unlikely. In fact, this intracellular Ca2+ rise and filopodiagenesis following glutamate stimulation suggest a pivotal role for α2δ1 in transforming a cell that shows no reaction to glutamate to a cell that responds to glutamate. However, the mechanism through which expression of α2δ1 causes an intracellular Ca2+ rise and filopodiagenesis in COS-7 cells remains to be identified. Recently, interaction of 2δ1 with NMDARs has been proposed to be the underlying mechanism of contribution of 2δ1 in glutamate signaling [191]. Therefore, its association with other glutamate receptors and proteins is likely. Whether α2δ1 mediates the intracellular Ca2+ rise and filopodiagenesis in association with a Ca2+-permeable glutamate receptor such as KAR (in astrocytoma cells) or it independently can sense glutamate and use the properties of associated proteins to rise the intracellular Ca2+ such as CaVs, or a combination of both should be identified in future studies. 100 Interestingly, COS-7 cells showed a ramified membrane and more processes upon transfection with α2δ1. These observations were consistent with reports from Wang et al. [215], and Risher et al. about the role of α2δ1 in cell ramification and filopodiagenesis [209]. Translocation of α2δ1 to the filopodia in U118-MG cells after exposure to glutamate (Figure 4.3.B) evidence the contribution of α2δ1 to the filopodiagenesis pathways. Further research should be undertaken to investigate the molecular mechanism of this ramification and process extension that was imposed by α2δ1 to COS-7 cells. In this research, to determine whether α2δ1 conducts directional motility of the cellular processes remains to be tested for future reports. Glutamate triggers several pathways in astrocytoma cells and the same experimental design that we have used to study KARs, which have a selective agonist (KA) in addition to glutamate, could not provide conclusive results about the contribution of α2δ1 in glutamate-induced morphological response. However, stimulating 3D cultured astrocytoma cells that their α2δ1 has been blocked with precisely photoreleasing glutamate could help to examine this notion further. Indicating that α2δ1 contributes to glutamate sensing and activating downstream pathways in heterologous expression systems with different molecular properties opens a new avenue of research in the field. There is abundant room for further progress in identifying the molecular mechanism by which α2δ1 senses glutamate and how it can initiate downstream pathways such as: determining the mechanism by which α2δ1 links the glutamate signaling to the Ca2+ signaling, the mechanisms that α2δ1 translates the extracellular signals into intracellular downstream pathways and describing the roles of the molecular domains that are expressed in the extracellular α2 fragment of α2δ (Cache and VWA). Abundance of VWA in human proteins has paved the way to describing its role in α2δ1 protein [178, 211, 221]. However, the role and function of Cache domains in α2δ1 has remained unexplored. Cache is a ligand-binding domain common to bacterial chemoreceptors, that senses 101 variety of chemoattractant such as small amino acids [222, 223]. Therefore, it is possible that it serves as a binding side for glutamate in α2δ1 as well. Exploiting the knowledge from bacterial Cache signaling will help to decipher their function in human. In addition, the mechanism of action of gabapentinoids has been a long-standing puzzle in managing pain in the CNS. Despite decades has been passed from its development and widespread application, how these α2δ1 ligands can interrupt neuropathic and chronic pain is not still known [177, 178, 180, 190, 224–226]. Recent studies have revealed that 2δ1 interacts with several proteins in addition to CaVs, and these associated proteins have been proposed to involve in the gabapentinoid pharmacology [224]. For instance, gabapentinoids have been proposed to inhibit forward trafficking of 2δ1-NMDAR complexes, and thus reduces the detrimental effect of NMDAR-dependent glutamatergic events in neuropathic pain [191]. The role of α2δ1 in increasing extension and number of the cellular process in a Ca2+-dependent manner (independent to CaVs) in response to glutamate, that had not been previously shown, may help to describe the molecular mechanisms through which gabapentinods are involved in glutamatergic events in the brain. In summary, in this chapter, α2δ1 was shown to contribute to both filopodiagenesis and calcium responses upon exposure to glutamate. Testing this role of α2δ1 in neurons and astrocytes may redefine our understanding about the role of α2δ1 in glutamatergic events and synaptogenesis in the CNS. 102 Chapter 5. Conclusion Growing evidence indicates that dysregulation of glutamate signaling between neuron and astrocytes leads to the onset and progression of the brain tumors [36, 51, 62, 63]. Glutamate is a common signal in central nervous system that both astrocytes and neurons use to communicate [3, 18, 25]. Intercellular glutamate signaling between neurons and astrocytes was the first breakthrough to support concept of a tripartite synapse [5, 25, 227]. Ever since, there has been tremendous progress in our understanding about different aspects of glutamate signaling in astrocytes [3, 12, 48, 58]. However, our appreciation of glutamate signaling in pathology of the brain tumors (gliomas) has just started to emerge. The excitotoxic effect of excessive glutamate on the neurons and its proliferative effect on the glioma cells in the brain tumors, as well as its role as a chemotactic signal in migration and infiltration of cancer cells throughout the brain, can result in the progression and malignancy of gliomas [33, 34, 36, 63, 66, 67, 100, 110, 149]. In addition to the detrimental effect of dysregulation in glutamate homeostasis in glioma progression, the aberrant morphology of astrocytoma cells in plays an important role in the progression and therapy-resistance of brain tumors [15–17, 63]. Since astrocytes from normal brain respond to glutamate by filopodiagenesis and processes extension, observing the morphological response to the glutamate dysregulation in astrocytoma cells is conceivable, but needs further empirical support. In normal astrocytes, the fine cellular processes extend in a Ca2+-dependent manner that is mediated by glutamate receptors [25, 52–54]. The same type of morphological response has also been observed in neurons where their neurite protrude toward glutamate via a Ca2+-dependent mechanism that is mediating by glutamate receptors [93, 95, 228, 229]. In the neurons, the contribution of voltage-gated calcium channels (CaVs) to this phenomenon has also been shown, but has not still been tested in the astrocytes [93, 94, 229]. This dissertation attempts to identify the correlation between glutamate signaling and processes extension in transformed astrocytes 103 (astrocytoma cells) which can be critical in the onset and progression of the brain tumors. Therefore, astrocytoma cells were tested to determine whether formation and extension of their fine processes can be induced and directed by glutamate; and then the possible contribution of glutamate receptors and CaVs in this phenomenon were studied. Unexpectedly, among the two representative types of astrocytoma cells—U118-MG and U138-MG cells—that have been used in this research, only U118-MG cells extend their processes significantly in response to glutamate. The processes of U118-MG cells showed a directional motility toward glutamate when cultured in a 3D matrix (GelMA). 5.1 Summary of findings Results from Chapter 2 of this thesis suggested that the activity of kainite-sensitive glutamate receptors (KARs) is crucial for the initiation of the process extension in response to glutamate in U118-MG cells. However, precisely releasing a caged agonist for KARs using optical fibers in 3D-cultured cells suggested that KARS are not responsible for the pathfinding of the protruding processes of U118-MG cells toward glutamate. In Chapter 3, studying the possible contribution of CaVs in a voltage-independent morphological response using a novel imaging probe, showed that CaVs translocate immediately upon exposure to glutamate. In addition, Ca2+ blockers decreased the extent of the processes extension that was induced by glutamate. Interestingly, the extracellular and non-essential auxiliary subunit of CaVs, α2δ1, was found to participate in filopodiagenesis and processes extension in glutamate signaling. To investigate the role of α2δ1 further, in Chapter 4, the natural properties of different cell lines were exploited. Overexpressing α2δ1 made U138-MG cells responsive to glutamate morphologically. In COS-7 cells, which do not express α2δ1 natively, process extension and a calcium influx was observed after their transfection with α2δ1. These observations confirm that 104 role of α2δ1 can initiate morphological responses upon exposure to glutamate. However, whether α2δ1 is playing any role in pathfinding of the processes toward glutamate remains to be identified. 5.1.1 U118-MG astrocytoma cells demonstrate a directional motility towards glutamate The first hypothesis that was tested in this research was the role of glutamate in inducing processes extension in astrocytoma cells and directing their protrusion. Using natural properties of two known glioma cell lines which are frequently used as representative of astrocytoma cells, in Chapter 2, showed that not all the astrocytoma cells extend their processes upon exposure to 100 μM of glutamate. U118-MG cells demonstrated a change in their intracellular calcium concentration that was 33-folds more than what was observed in U138-MG cells and a significantly greater processes extension upon exposure to 100 μM. In addition, using 3D cultured cells and photo-release of glutamate revealed that the processes of U118-MG cells can protrude toward glutamate with a directional motility. 5.1.2 KARs contribute to the glutamate-induced morphological response in U118-MG astrocytoma cells In Chapter 2 of this dissertation the contribution of glutamate receptors in the glutamate-induced morphological response in astrocytoma cells was verified. The results suggest that KARs contribute to the glutamate-induced morphological response in U118-MG astrocytoma cells. The active role of KARs in glutamate signaling of normal astrocytes is not fully understood [3], however, their overexpression in astrocytoma cells has been recently shown which can imply their role in pathogenesis of astrocytoma cells [36, 62, 63]. Demonstrating the role of KARs in the morphology of U118-MG astrocytoma cells paves the way for identifying their role in organotypic slices of the tumors and primary astrocytoma cells. In fact, if KARs conduct processes motility in 105 tumoral astrocytoma cells, these receptors can be targeted at early stages of glioma to control its progression and reduce its invasion. The optimization of a 3D cell culturing method for astrocytes as well as validating Ph-KA as a modifiable kainoid for research about KAR and introducing DECM-PhKA for studying KARs precisely were other hallmarks of Chapter 2. These chemical tools can pave the way to explore undermined KARs in CNS. 5.1.3 CaVs contribute to the glutamate-induced morphological response in U118-MG astrocytoma cells Despite decades of attempts to identify the functional role of high voltage CaVs in non-excitable astrocytes [85–88, 197], their role and function in normal astrocytes as well as in the transformed astrocytoma cells is still an enigma. Therefore, in Chapter 3 their contribution to a glutamate-induced morphological response, which is a voltage-independent cellular event, was examined. Specifically, the role of L type CaVs (LTCC) in the processes formation and extension in U118-MG astrocytoma cells in response to glutamate was studied. The results from a pharmacological approach indicated that Ca2+ channel blockers had a controlling effect on the extent of process extension upon exposure to glutamate. In addition, glutamate has an immediate effect on the redistribution and translocation of CaV1.2 in U118-MG cells. It should be noted that U118-MG cells, which were used in this research, only express CaV1.2 among L-type CaVs [185, 200, 220], which may differ from the expression profile of the astrocytoma cells in the fresh tumors. Further study may reveal the possible contribution of other CaVs in addition to CaV1.2 to the glutamate-induced processes extension in astrocytoma cells. Altogether, the findings presented in Chapter 3 in an astrocytoma cell line can be the first step towards describing the mechanism of inhibitory effect of CaV blockers in the progression of the malignant gliomas, which has been a longstanding question in cancer studies [98–100]. 106 Creating and validating a real-time imaging tool, FluoBar1, was another outcome of Chapter 3. No simple and efficient tool for real-time imaging, which does not alter the activity of these channels, was available, despite the significance of the dynamic of CaVs in several cellular events have been highlighted, FluoBar1 can be used to monitor the immediate effect of glutamate on membranar distribution of CaVs in primary astrocytoma cells in the next step of this research. 5.1.4 The α2δ1 controls filopodiagenesis upon exposure to glutamate Results presented in Chapter 4 suggest that α2δ1 plays a role in linking calcium signaling with glutamate signaling and filopodiagenesis. Both in native and heterologous expression systems, α2δ1 was shown to contribute to both filopodiagenesis and intracellular Ca2+ rise upon exposure to glutamate. Selective ligands for α2δ1 inhibited the morphological response of U118-MG, which are the astrocytoma cells that respond to glutamate natively. In the U118-MG cells that were stimulated with glutamate, α2δ1 translocated to the filopodia. These results suggest a role for α2δ1 in filopodiagenesis in U118-MG cells upon exposure to glutamate. This observation was confirmed further when U138-MG astrocytoma cells, that do not natively respond morphologically response to glutamate, extended their processes upon exposure to glutamate when transfected with α2δ1. Intriguingly, the α2δ1 also transformed COS-7 cells, which are CaV free cells, to the cells that can respond to glutamate by process extension and intracellular Ca2+ rise. In fact, showing the direct involvement of α2δ1 in glutamate signaling can redefine our understanding about glutamatergic events in the central nervous system. In addition, this finding suggest co ntribution of α2δ1 in the aberrant morphology of astrocytoma cells which can result in developing therapy-resistant tumors. 5.2 Future research direction In this research, KARs were shown to contribute to the morphological response to glutamate using an astrocytoma cell line. However, this notion needs to be cinfirmed in primary astrocytoma 107 cells and organotypic slices of astrocytoma tumors. The role of KARs in morphological response can also be examined in normal astrocytes. Testing different types and different concentrations of KAR agonists and antagonists can provide more detailed information about the role and function of KARs in astrocytoma cells. Creating more efficient tools and techniques to study a morphological response will pave the way for better understanding about the role of these receptors. The caged-kainoid that was created and used to activate KARs, in this dissertation caused a different form of Ca2+ rise in U118-MG cells compared to its parent molecule PhKA. More studies is needed to explain this difference and chemical modification might be required for the next generation caged kainoids. In addition to KARs, participation of extracellular subunit of CaVs, α2δ1, in glutamate-induced morphological response has also been found in this dissertation. Molecular studies is required to explore and identify the role of each of the domains of α2δ1 in defining the functions of α2δs in CNS. The structure of α2δ consists of a protein-protein interaction domain (Von Willebrand Factor A) and four evolutionarily conserved chemotactic domains (Cache). Each of these domains might be a receptor for a specific ligand which trigger an intracellular downstream pathway. The downstream pathway(s) by which α2δ1 relates to Ca2+ signaling independent to CaVs to exert its role in response to glutamate remains to be explored. Overall, the findings of this dissertation in U118-MG cells showed that both KARs and α2δ1 contribute to the initiation of the morphological response to glutamate. Blocking each of them separately, inhibited the morphological response entirely. Our preliminary results show the requirements of both proteins (Appendix F) to initiate the morphological response to glutamate. More research can closely examine whether this effect is due to a physical interaction and association of α2δ1 and KARs, or it is through other previously unidentified pathways. In addition, U138-MG cells express both KARs and α2δ1, yet they did not naturally extend their processes upon exposure to glutamate. But upon transfection with α2δ1 and its overexpression, 108 U138-MG cells started reacting to glutamate by a significantly more intracellular Ca2+ rise and filopodiagenesis, which may suggest the crucial role of α2δ1. However, overexpression of KARs is also required in this cells line to draw an inclusive conclusion about the role of each channel in this phenomenon. 5.3 Implications and significance The findings of this dissertation provide insights for the effect of glutamate dysregulation in shaping specific morphology of astrocytoma cells. Describing the underlying mechanisms that link these two main features of gliomas, paves the way for therapeutic purposes. Given the fact that ramification of the membrane and elongated projections of astrocytoma cells causes resistant of the tumor to the therapies [15, 16], finding the proteins that contribute to this morphology introduces previously overlooked pathophysiological targets. Results of this dissertation indicated that KARs and CaVs contribute to the filopodiagenesis and processes extension of astrocytoma cells when they were exposed to glutamate. Therefore, these two ion channels can be investigated more to be considered as target molecules for preventing the progression of gliomas. In addition, these findings provide more support to the growing body of knowledge about emerging roles for KARs and CaVs in the health and disease. Specifically, identifying a function for KARS in a morphological feature, that has not been reported previously, help to appreciate the unique properties of these receptors in the CNS further. Although KARs display both ionotropic and metabotropic modes of action as glutamate receptors, their function in the CNS may have been underestimated. In fact, KARS have been shown to signal via a less explored non-canonical G-protein coupled (GPCR) metabotropic cascade beside their classical function as an ion channel [230–232]. This research also found that a voltage-operated channel, CaV1.2, contributes to morphological responses, which are non-electrical cellular events. A role CaV1.2 in the progression of cancers 109 is emerging, however, all previous published studies focused on the role of the these channels in supplying intracellular Ca2+ that supports the cell proliferation [96, 98, 99]. One intriguing finding of this dissertation was participation of a non-essential auxiliary subunit of CaVs, α2δ1, in the morphological response of astrocytoma cells to glutamate. While the involvement of α2δ1 in filopodiagenesis, independent to CaVs, has recently been reported in a different context about synaptogenesis [209], their contribution to the morphological response of astrocytoma cells to glutamate was not expected. This role of α2δ1 may introduce a new target for chemotherapeutical approaches in glioma. In fact, identifying the role of α2δ1 in filopodiagenesis and processes extension complement those of emerging studies that unravel the characteristics of α2δ in neural wiring. Exploring the functions of α2δ1 is one of the fast-growing topics in neuroscience. Its association with different membranal proteins and significant in synaptogenesis makes it one of the frontier topics in the field [175, 191, 209, 211, 233]. Emerging evidence reveals the association of α2δ1 with different proteins that are involved in synaptogenesis such as with NMDARs. Indicating that both KARs and α2δ1 contribute to the morphological response of astrocytoma cells may also imply these two proteins are associating with each other which remains to be indicated. 110 Bibliography 1. Allen NJ, Barres BA (2009) Glia — more than just brain glue. Nature 457:675–677. https://doi.org/10.1038/457675a 2. Allen NJ, Eroglu C (2017) Cell Biology of Astrocyte-Synapse Interactions. Neuron 96:697–708. https://doi.org/10.1016/j.neuron.2017.09.056 3. Rose CR, Felix L, Zeug A, et al (2018) Astroglial Glutamate Signaling and Uptake in the Hippocampus. Front Mol Neurosci 10:1–20. https://doi.org/10.3389/fnmol.2017.00451 4. Bazargani N, Attwell D (2016) Astrocyte calcium signaling: The third wave. Nat Neurosci 19:182–189. https://doi.org/10.1038/nn.4201 5. Araque A, Parpura V, Sanzgiri RP, Haydon PG (1999) Tripartite synapses: glia, the unacknowledged partner. Trends Neurosci 22:208–215. https://doi.org/10.1016/S0166-2236(98)01349-6 6. Chung W-S, Allen NJ, Eroglu C (2015) Astrocytes Control Synapse Formation, Function, and Elimination. Cold Spring Harb Perspect Biol 7:a020370. https://doi.org/10.1101/cshperspect.a020370 7. Stogsdill JA, Ramirez J, Liu D, et al (2017) Astrocytic neuroligins control astrocyte morphogenesis and synaptogenesis. Nature 551:192–197. https://doi.org/10.1038/nature24638 8. Schiweck J, Eickholt BJ, Murk K (2018) Important shapeshifter: Mechanisms allowing astrocytes to respond to the changing nervous system during development, injury and disease. Front. Cell. Neurosci. 12 9. Allen NJ, Barres BA (2005) Signaling between glia and neurons: Focus on synaptic plasticity. Curr. Opin. Neurobiol. 15:542–548 10. Hirase H, Qian L, Barthó P, Buzsáki G (2004) Calcium dynamics of cortical astrocytic networks in vivo. PLoS Biol 2:494–499. https://doi.org/10.1371/journal.pbio.0020096 11. Santello M, Toni N, Volterra A (2019) Astrocyte function from information processing to cognition and cognitive impairment. Nat Neurosci 22:154–166. https://doi.org/10.1038/s41593-018-0325-8 12. Matyash V, Kettenmann H (2010) Heterogeneity in astrocyte morphology and physiology. Brain Res Rev 63:2–10. https://doi.org/10.1016/j.brainresrev.2009.12.001 13. Zhang Y, Barres BA (2013) A smarter mouse with human astrocytes. BioEssays 35:876–880. https://doi.org/10.1002/BIES.201300070 14. Han X, Chen M, Wang F, et al (2013) Forebrain engraftment by human glial progenitor cells enhances synaptic plasticity and learning in adult mice. Cell Stem Cell 12:342–353. https://doi.org/10.1016/j.stem.2012.12.015 15. Osswald M, Solecki G, Wick W, Winkler F (2016) A malignant cellular network in gliomas: Potential clinical implications. Neuro Oncol 18:479–485. https://doi.org/10.1093/neuonc/now014 16. Weil S, Osswald M, Solecki G, et al (2017) Tumor microtubes convey resistance to surgical lesions and chemotherapy in gliomas. Neuro Oncol 19:1316–1326. https://doi.org/10.1093/neuonc/nox070 17. Venkataramani V, Tanev DI, Strahle C, et al (2019) Glutamatergic synaptic input to glioma cells drives brain tumour progression. Nature 573:532–538. https://doi.org/10.1038/s41586-019-1564-x 18. Verkhratsky A, Kirchhoff F (2007) Glutamate-mediated neuronal-glial transmission. In: Journal of Anatomy. John Wiley & Sons, Ltd (10.1111), pp 651–660 19. Barnes GN, Slevin JT (2012) Effect of Genetic Background on Ionotropic Glutamate Receptor and Neurotrophin Biology: Synaptic Connectivity and Function in Neurological Disease. Front Med Chem - (Volume 2) 111–132. https://doi.org/10.2174/978160805205910502010111 20. Zheng JQ, Wan JJ, Poo MM (1996) Essential role of filopodia in chemotropic turning of nerve growth cone induced by a glutamate gradient. J Neurosci 16:1140–1149. https://doi.org/10.1523/jneurosci.16-03-01140.1996 21. Heckman CA, Plummer HK (2013) Filopodia as sensors. Cell Signal 25:2298–2311. https://doi.org/10.1016/J.CELLSIG.2013.07.006 22. Chien C-B, Rosenthal DE, Harris WA, Holt CE (1993) Navigational errors made by growth cones without filopodia in the embryonic xenopus brain. Neuron 11:237–251. https://doi.org/10.1016/0896-6273(93)90181-P 23. Haber M, Zhou L, Murai KK (2006) Cooperative Astrocyte and Dendritic Spine Dynamics at Hippocampal Excitatory Synapses. J Neurosci 26:8881–8891. https://doi.org/10.1523/JNEUROSCI.1302-06.2006 24. Lavialle M, Aumann G, Anlauf E, et al (2011) Structural plasticity of perisynaptic astrocyte processes involves ezrin and metabotropic glutamate receptors. Proc Natl Acad Sci U S A 111 108:12915–12919. https://doi.org/10.1073/pnas.1100957108 25. Cornell‐Bell AH, Prem TG, Smith SJ (1990) The excitatory neurotransmitter glutamate causes filopodia formation in cultured hippocampal astrocytes. Glia 3:322–334. https://doi.org/10.1002/glia.440030503 26. Hirrlinger J, Hülsmann S, Kirchhoff F (2004) Astroglial processes show spontaneous motility at active synaptic terminals in situ. Eur J Neurosci 20:2235–2239. https://doi.org/10.1111/j.1460-9568.2004.03689.x 27. Mortimer D, Fothergill T, Pujic Z, et al (2008) Growth cone chemotaxis. Trends Neurosci 31:90–98. https://doi.org/10.1016/J.TINS.2007.11.008 28. Faix J, Rottner K (2006) The making of filopodia. Curr Opin Cell Biol 18:18–25. https://doi.org/10.1016/j.ceb.2005.11.002 29. Mejillano MR, Kojima S-I, Applewhite DA, et al (2004) Lamellipodial Versus Filopodial Mode of the Actin Nanomachinery: 30. Tabatabaee M, Menard F (2020) The α2δ1 protein―Voltage-gated calcium channel subunit or neuronal signal transducer? In: Cold Spring Harbor symposia on Glia in health and disease. Virtual meeting 31. Perez-Alvarez A, Navarrete M, Covelo A, et al (2014) Structural and functional plasticity of astrocyte processes and dendritic spine interactions. J Neurosci 34:12738–44. https://doi.org/10.1523/JNEUROSCI.2401-14.2014 32. Bernardinelli Y, Randall J, Janett E, et al (2014) Activity-dependent structural plasticity of perisynaptic astrocytic domains promotes excitatory synapse stability. Curr Biol 24:1679–88. https://doi.org/10.1016/j.cub.2014.06.025 33. Lim-Fat MJ, Wen PY (2020) Glioma progression through synaptic activity. Nat. Rev. Neurol. 16:6–7 34. Weingart JD, McGirt MJ, Brem H (2010) High-grade astrocytoma/glioblastoma. In: Oncology of CNS Tumors. Springer Berlin Heidelberg, pp 147–161 35. Vargová L, Homola A, Zámečník J, et al (2003) Diffusion parameters of the extracellular space in human gliomas. Glia 42:77–88. https://doi.org/10.1002/glia.10204 36. Pei Z, Lee KC, Khan A, et al (2020) Pathway analysis of glutamate-mediated, calcium-related signaling in glioma progression. Biochem. Pharmacol. 176:113814 37. Heller JP, Rusakov DA (2015) Morphological plasticity of astroglia: Understanding synaptic microenvironment. Glia 63:2133. https://doi.org/10.1002/GLIA.22821 38. Fulga TA, Rørth P (2002) Invasive cell migration is initiated by guided growth of long cellular extensions. Nat Cell Biol 4:715–719. https://doi.org/10.1038/ncb848 39. Yee KT, Simon HH, Tessier-Lavigne M, O’Leary DDM (1999) Extension of long leading processes and neuronal migration in the mammalian brain directed by the chemoattractant netrin-1. Neuron 24:607–622. https://doi.org/10.1016/S0896-6273(00)81116-2 40. Willard SS, Koochekpour S (2013) Glutamate, glutamate receptors, and downstream signaling pathways. Int J Biol Sci 9:948–959. https://doi.org/10.7150/ijbs.6426 41. Zhou Y, Danbolt NC (2014) Glutamate as a neurotransmitter in the healthy brain. J Neural Transm 121:799–817. https://doi.org/10.1007/s00702-014-1180-8 42. Cornell-Bell AH, Thomas PG, Caffrey JM (1992) Ca2+ and filopodial responses to glutamate in cultured astrocytes and neurons. Can J Physiol Pharmacol 70 Suppl:S206-18 43. Choi DW (1985) Glutamate neurotoxicity in cortical cell culture is calcium dependent. Neurosci Lett 58:293–297. https://doi.org/10.1016/0304-3940(85)90069-2 44. Yoshizumi M, Eisenach JC, Hayashida K (2012) Riluzole and gabapentinoids activate glutamate transporters to facilitate glutamate-induced glutamate release from cultured astrocytes. Eur J Pharmacol 677:87–92. https://doi.org/10.1016/J.EJPHAR.2011.12.015 45. Mattson MP (2019) Excitotoxicity. In: Stress: Physiology, Biochemistry, and Pathology. Elsevier, pp 125–134 46. Hayashi MK (2018) Structure-function relationship of transporters in the glutamate–glutamine cycle of the central nervous system. Int J Mol Sci 19:. https://doi.org/10.3390/ijms19041177 47. Hassinger TD, Atkinson PB, Strecker GJ, et al (1995) Evidence for glutamate-mediated activation of hippocampal neurons by glial calcium waves. J Neurobiol 28:159–170. https://doi.org/10.1002/neu.480280204 48. Bernardinelli Y, Muller D, Nikonenko I (2014) Astrocyte-synapse structural plasticity. Neural Plast 2014:232105. https://doi.org/10.1155/2014/232105 49. Kucukdereli H, Allen NJ, Lee AT, et al (2011) Control of excitatory CNS synaptogenesis by astrocyte-secreted proteins Hevin and SPARC. 112 Proc Natl Acad Sci 108:E440–E449. https://doi.org/10.1073/pnas.1104977108 50. Shigetomi E, Patel S, Khakh BS (2016) Probing the Complexities of Astrocyte Calcium Signaling. Trends Cell Biol. 26:300–312 51. Nedergaard M (1994) Direct signaling from astrocytes to neurons in cultures of mammalian brain cells. Science (80- ) 263:1768–1771. https://doi.org/10.1126/science.8134839 52. Sun W, McConnell E, Pare J-F, et al (2013) Glutamate-dependent neuroglial calcium signaling differs between young and adult brain. Science 339:197–200. https://doi.org/10.1126/science.1226740 53. Panatier A, Robitaille R (2016) Astrocytic mGluR5 and the tripartite synapse. Neuroscience 323:29–34. https://doi.org/10.1016/J.NEUROSCIENCE.2015.03.063 54. Iino M, Goto K, Kakegawa W, et al (2001) Glia-Synapse Interaction Through Ca2+-Permeable AMPA Receptors in Bergmann Glia. Science (80- ) 292:926–929. https://doi.org/10.1126/science.1058827 55. Farmer WT, Murai K (2017) Resolving Astrocyte Heterogeneity in the CNS. Front Cell Neurosci 11:1–7. https://doi.org/10.3389/fncel.2017.00300 56. Savtchouk I, Volterra A (2018) Gliotransmission: Beyond Black-and-White. J Neurosci 38:14–25. https://doi.org/10.1523/JNEUROSCI.0017-17.2017 57. Trudler D, Sanz-Blasco S, Eisele YS, et al (2021) α-Synuclein Oligomers Induce Glutamate Release from Astrocytes and Excessive Extrasynaptic NMDAR Activity in Neurons, Thus Contributing to Synapse Loss. J Neurosci 41:JN-RM-1871-20. https://doi.org/10.1523/jneurosci.1871-20.2020 58. Parpura V, Basarsky TA, Liu F, et al (1994) Glutamate-mediated astrocyte-neuron signalling. Lett to Nat 369:744–747. https://doi.org/10.1038/369744a0 59. Malarkey EB, Parpura V (2008) Mechanisms of glutamate release from astrocytes. Neurochem. Int. 52:142–154 60. Tian Z, Tabatabaee MS, Edemann S, et al (2020) Optical Control of Ca2+-Mediated Morphological Response in Glial Cells with Visible Light Using a Photocaged Kainoid. ChemRxiv. https://doi.org/10.26434/CHEMRXIV.11888925.V1 61. Bohmbach K, Schwarz MK, Schoch S, Henneberger C (2018) The structural and functional evidence for vesicular release from astrocytes in situ. Brain Res. Bull. 136:65–75 62. Takano T, Lin JHC, Arcuino G, et al (2001) Glutamate release promotes growth of malignant gliomas. Nat Med 7:1010–1015. https://doi.org/10.1038/nm0901-1010 63. de Groot J, Sontheimer H (2011) Glutamate and the biology of gliomas. Glia 59:1181–1189. https://doi.org/10.1002/glia.21113 64. Hamadi A, Giannone G, Takeda K, Rondé P (2014) Glutamate involvement in calcium-dependent migration of astrocytoma cells. Cancer Cell Int 14:1–10. https://doi.org/10.1186/1475-2867-14-42 65. Turner DA, Adamson DC (2011) Neuronal-astrocyte metabolic interactions: Understanding the transition into abnormal astrocytoma metabolism. J. Neuropathol. Exp. Neurol. 70:167–176 66. Rosati A, Marconi S, Pollo B, et al (2009) Epilepsy in glioblastoma multiforme: Correlation with glutamine synthetase levels. J Neurooncol 93:319–324. https://doi.org/10.1007/s11060-008-9794-z 67. van Lith SAM, Navis AC, Verrijp K, et al (2014) Glutamate as chemotactic fuel for diffuse glioma cells: Are they glutamate suckers? Biochim. Biophys. Acta - Rev. Cancer 1846:66–74 68. Oberheim NA, Wang X, Goldman S, Nedergaard M (2006) Astrocytic complexity distinguishes the human brain. Trends Neurosci 29:547–553. https://doi.org/10.1016/J.TINS.2006.08.004 69. Swanson GT, Sakai R (2009) Ligands for ionotropic glutamate receptors. Prog Mol Subcell Biol 46:123–57. https://doi.org/10.1007/978-3-540-87895-7_5 70. Tabatabaee MS, Tian Z, Gibon J, Menard F (2021) Aminooxadiazolyl kainic acid reveals that kainic acid receptors contribute to astrocytoma glutamate signaling. bioRxiv 2021.01.16.426948. https://doi.org/10.1101/2021.01.16.426948 71. Alt A, Weiss B, Ogden AM, et al (2004) Pharmacological characterization of glutamatergic agonists and antagonists at recombinant human homomeric and heteromeric kainate receptors in vitro. Neuropharmacology 46:793–806. https://doi.org/10.1016/j.neuropharm.2003.11.026 72. Matute C (2011) Therapeutic potential of kainate receptors. CNS Neurosci Ther 17:661–669. https://doi.org/10.1111/j.1755-5949.2010.00204.x 73. Lerma J (2003) Roles and rules of kainate receptors in synaptic transmission. Nat Rev Neurosci 4:481–495. https://doi.org/10.1038/nrn1118 113 74. Stepulak A, Rola R, Polberg K (2014) Glutamate and its receptors in cancer. J Neural Transm 121:933–944. https://doi.org/10.1007/s00702-014-1182-6 75. Vargas JR, Koji Takahashi D, Thomson KE, Wilcox KS (2013) The Expression of Kainate Receptor Subunits in Hippocampal Astrocytes After Experimentally Induced Status Epilepticus 76. Panatier A, Vallée J, Haber M, et al (2011) Astrocytes are endogenous regulators of basal transmission at central synapses. Cell 146:785–798. https://doi.org/10.1016/j.cell.2011.07.022 77. Haustein MD, Kracun S, Lu X-H, et al (2014) Conditions and constraints for astrocyte calcium signaling in the hippocampal mossy fiber pathway. Neuron 82:413–29. https://doi.org/10.1016/j.neuron.2014.02.041 78. Verkhratsky A, Kirchhoff F (2007) NMDA receptors in glia. Neuroscientist 13:28–37. https://doi.org/10.1177/1073858406294270 79. Lalo U, Pankratov Y, Kirchhoff F, et al (2006) NMDA receptors mediate neuron-to-glia signaling in mouse cortical astrocytes. J Neurosci 26:2673–2683. https://doi.org/10.1523/JNEUROSCI.4689-05.2006 80. Rusakov DA (2015) Disentangling calcium-driven astrocyte physiology. Nat Rev Neurosci 16:226–233. https://doi.org/10.1038/nrn3878 81. Molotkov D, Zobova S, Arcas JM, Khiroug L (2013) Calcium-induced outgrowth of astrocytic peripheral processes requires actin binding by Profilin-1. Cell Calcium 53:338–348. https://doi.org/10.1016/J.CECA.2013.03.001 82. Khakh BS, Sofroniew M V (2015) Diversity of astrocyte functions and phenotypes in neural circuits. Nat Neurosci 18:942–52. https://doi.org/10.1038/nn.4043 83. Derouiche A, Geiger KD (2019) Perspectives for Ezrin and Radixin in Astrocytes: Kinases, Functions and Pathology. Int J Mol Sci 20:3776. https://doi.org/10.3390/ijms20153776 84. Schmitz Y, Luccarelli J, Kim M, et al (2009) Glutamate Controls Growth Rate and Branching of Dopaminergic Axons. J Neurosci 29:11973. https://doi.org/10.1523/JNEUROSCI.2927-09.2009 85. Latour I, Hamid J, Beedle AM, et al (2003) Expression of voltage-gated Ca2+ channel subtypes in cultured astrocytes. Glia 41:347–353. https://doi.org/10.1002/glia.10162 86. Cheli VT, Santiago González DA, Smith J, et al (2016) L-type voltage-operated calcium channels contribute to astrocyte activation In vitro. Glia 64:1396–415. https://doi.org/10.1002/glia.23013 87. MacVicar BA (1984) Voltage-Dependent Calcium Channels in Glial Cells Author ( s ): B . A . MacVicar Published by : American Association for the Advancement of Science Stable URL : http://www.jstor.org/stable/1693335 . Science (80- ) 226:1345–1347 88. Carmignoto G, Pasti L, Pozzan T (1998) On the role of voltage-dependent calcium channels in calcium signaling of astrocytes in situ. J Neurosci 18:4637–4645. https://doi.org/10.1523/jneurosci.18-12-04637.1998 89. Rungta RL, Bernier L-P, Dissing-Olesen L, et al (2016) Ca 2+ transients in astrocyte fine processes occur via Ca 2+ influx in the adult mouse hippocampus. Glia 64:2093–2103. https://doi.org/10.1002/glia.23042 90. Zamponi GW, Striessnig J, Koschak A, Dolphin AC (2015) The Physiology, Pathology, and Pharmacology of Voltage-Gated Calcium Channels and Their Future Therapeutic Potential. Pharmacol Rev Pharmacol Rev 67:821–870. https://doi.org/10.1124/pr.114.009654 91. Zuccotti A, Clementi S, Reinbothe T, et al (2011) Structural and functional differences between L-type calcium channels: crucial issues for future selective targeting. Trends Pharmacol Sci 32:366–75. https://doi.org/10.1016/j.tips.2011.02.012 92. Striessnig J, Bolz HJ, Koschak A (2010) Channelopathies in CaV1.1, CaV1.3, and CaV1.4 voltage-gated L-type Ca2+ channels. Pflugers Arch Eur J Physiol 460:361–374. https://doi.org/10.1007/s00424-010-0800-x 93. Silver RA, Lamb AG, Bolsover SR (1990) Calcium hotspots caused by L-channel clustering promote morphological changes in neuronal growth cones. Nature 343:751–754. https://doi.org/10.1038/343751a0 94. Tam T, Mathews E, Snutch TP, Schafer WR (2000) Voltage-gated calcium channels direct neuronal migration in Caenorhabditis elegans. Dev Biol 226:104–117. https://doi.org/10.1006/dbio.2000.9854 95. Zatkova M, Reichova A, Bacova Z, et al (2018) Neurite Outgrowth Stimulated by Oxytocin Is Modulated by Inhibition of the Calcium Voltage-Gated Channels. Cell Mol Neurobiol 38:371–378. https://doi.org/10.1007/s10571-017-0503-3 96. Morrone FB, Gehring MP, Nicoletti NF (2016) Calcium Channels and Associated Receptors in Malignant Brain Tumor Therapy. Mol Pharmacol Mol Pharmacol 90:403–409. https://doi.org/10.1124/mol.116.103770 97. Nicoletti NF, Erig TC, Zanin RF, et al (2017) Pre-clinical evaluation of voltage-gated calcium 114 channel blockers derived from the spider P. nigriventer in glioma progression. Toxicon 129:58–67. https://doi.org/10.1016/j.toxicon.2017.02.001 98. Phan NN, Wang CY, Chen CF, et al (2017) Voltage-gated calcium channels: Novel targets for cancer therapy. Oncol Lett 14:2059–2074. https://doi.org/10.3892/ol.2017.6457 99. Martínez-Delgado G, Felix R (2017) Emerging Role of CaV1.2 Channels in Proliferation and Migration in Distinct Cancer Cell Lines. Oncology 93:1–10. https://doi.org/10.1159/000464293 100. Reardon DA, Rich JN, Friedman HS, Bigner DD (2006) Recent Advances in the Treatment of Malignant Astrocytoma. J Clin Oncol 24:1253–1265. https://doi.org/10.1200/JCO.2005.04.5302 101. Lee YS, Sayeed MM, Wurster RD (1994) Inhibition of cell growth and intracellular Ca2+ mobilization in human brain tumor cells by Ca2+ channel antagonists. Mol Chem Neuropathol 22:81–95. https://doi.org/10.1007/BF03160097 102. Kunert‐Radek J, Stepien H, Radek A, et al (1989) Inhibitory effect of calcium channel blockers on proliferation of human glioma cells in vitro. Acta Neurol Scand 79:166–169. https://doi.org/10.1111/j.1600-0404.1989.tb03731.x 103. Chen R, Smith-Cohn M, Cohen AL, Colman H (2017) Glioma Subclassifications and Their Clinical Significance. https://doi.org/10.1007/s13311-017-0519-x 104. Alcantara Llaguno SR, Parada LF (2016) Cell of origin of glioma: biological and clinical implications. Br J Cancer 2016 11512 115:1445–1450. https://doi.org/10.1038/bjc.2016.354 105. Louis DN, Ohgaki H, Wiestler OD, et al (2007) The 2007 WHO Classification of Tumours of the Central Nervous System. Acta Neuropathol 114:97. https://doi.org/10.1007/S00401-007-0243-4 106. De Ridder LI, Laerum OD, Murk SJ, Bigner DD (1987) Invasiveness of human glioma cell lines in vitro: Relation to tumorigenicity in athymic mice. Acta Neuropathol 72:207–213 107. Ye ZC, Sontheimer H (1999) Glioma cells release excitotoxic concentrations of glutamate. Cancer Res 59:4383–4391 108. ZC Y, JD R, H S (1999) Compromised glutamate transport in human glioma cells: reduction-mislocalization of sodium-dependent glutamate transporters and enhanced activity of cystine-glutamate exchange. J Neurosci 19:10767–10777. https://doi.org/10.1523/JNEUROSCI.19-24-10767.1999 109. Osswald M, Jung E, Sahm F, et al (2015) Brain tumour cells interconnect to a functional and resistant network. Nature 528:93–98. https://doi.org/10.1038/nature16071 110. Lyons SA, Chung WJ, Weaver AK, et al (2007) Autocrine glutamate signaling promotes glioma cell invasion. Cancer Res 67:9463–9471. https://doi.org/10.1158/0008-5472.CAN-07-2034 111. U-118 MG ATCC ® HTB-15TM Homo sapiens brain classified as gr. https://www.atcc.org/products/all/HTB-15.aspx. Accessed 24 Jan 2020 112. U-138 MG ATCC ® HTB-16TM Homo sapiens brain glioblastoma; cl. https://www.atcc.org/products/all/HTB-16.aspx. Accessed 20 Apr 2021 113. PONTEN J (1975) Neoplastic Human Glia CellsCells in Culture. In: Fogh J (ed) Human Tumor Cells in Vitro. New York, pp 175–206 114. Westermark B (1973) The deficient density‐dependent growth control of human malignant glioma cells and virus‐transformed glia‐like cells in culture. Int J Cancer 12:438–451. https://doi.org/10.1002/ijc.2910120215 115. Fredman P, Månsson JE, Bigner SH, et al (1990) Gangliosides in the human glioma cell line U-118 MG grown in culture or as xenografts in nude rats. Biochim Biophys Acta - Lipids Lipid Metab 1045:239–244. https://doi.org/10.1016/0005-2760(90)90126-I 116. COS-7 | ATCC. https://www.atcc.org/products/crl-1651. Accessed 23 Oct 2021 117. Nozaki M, Haraguchi S, Miyazaki T, et al (2018) Expression of steroidogenic enzymes and metabolism of steroids in COS-7 cells known as non-steroidogenic cells. Sci Rep 8:. https://doi.org/10.1038/S41598-018-20226-2 118. Meir A, Dolphin AC (1998) Known Calcium Channel α1 Subunits Can Form Low Threshold Small Conductance Channels with Similarities to Native T-Type Channels. Neuron 20:341–351. https://doi.org/10.1016/S0896-6273(00)80461-4 119. Meir A, Dolphin AC (1998) Known calcium channel α1subunits can form low threshold small conductance channesl with similarities to native T-type channels. Neuron 20:341–351. https://doi.org/10.1016/S0896-6273(00)80461-4 120. Robinson P, Etheridge S, Song L, et al (2010) Formation of N-type (CaV2.2) voltage-gated calcium channel membrane microdomains: Lipid raft association and clustering. Cell Calcium 48:183–194. https://doi.org/10.1016/J.CECA.2010.08.006 121. Douglas L, Davies A, Wratten J, Dolphin AC (2006) Do voltage-gated calcium channel α 2 δ 115 subunits require proteolytic processing into α 2 and δ to be functional? Biochem Soc Trans 34:894–898. https://doi.org/10.1042/BST0340894 122. Boone M, Roussel M, Chauffert B, et al (2016) Prevalence and profile of cognitive impairment in adult glioma: a sensitivity analysis. J Neurooncol 129:123–130. https://doi.org/10.1007/s11060-016-2152-7 123. Takano T, Wallace JT, Baldwin KT, et al (2020) Chemico-genetic discovery of astrocytic control of inhibition in vivo. Nature 588:296–302. https://doi.org/10.1038/s41586-020-2926-0 124. Kanumilli S, Roberts PJ (2006) Mechanisms of glutamate receptor induced proliferation of astrocytes. Neuroreport 17:1877–1881. https://doi.org/10.1097/WNR.0B013E3280102EE6 125. Savaskan NE, Seufert S, Hauke J, et al (2011) Dissection of mitogenic and neurodegenerative actions of cystine and glutamate in malignant gliomas. Oncogene 30:43–53. https://doi.org/10.1038/onc.2010.391 126. Wang Z, Abdulla R, Parker B, et al (2015) A simple and high-resolution stereolithography-based 3D bioprinting system using visible light crosslinkable bioinks. Biofabrication 7:45009. https://doi.org/10.1088/1758-5090/7/4/045009 127. Wang Z, Tian Z, Menard F, Kim K (2017) Comparative study of gelatin methacrylate hydrogels from different sources for biofabrication applications. Biofabrication 9:044101. https://doi.org/10.1088/1758-5090/aa83cf 128. Palma-Cerda F, Auger C, Crawford DJ, et al (2012) New caged neurotransmitter analogs selective for glutamate receptor sub-types based on methoxynitroindoline and nitrophenylethoxycarbonyl caging groups. Neuropharmacology 63:624–634. https://doi.org/10.1016/j.neuropharm.2012.05.010 129. Ellis-Davies GCR (2007) Caged compounds: photorelease technology for control of cellular chemistry and physiology. Nat Methods 4:619–28. https://doi.org/10.1038/nmeth1072 130. Shembekar VR, Chen Y, Carpenter BK, Hess GP (2005) A protecting group for carboxylic acids that can be photolyzed by visible light. Biochemistry 44:7107–7114. https://doi.org/10.1021/bi047665o 131. Wang X, Gao Y, Xu Y, et al (2009) Efficient preparation of photolabile agent mni-glu by regioselective nitration of 4-methoxyindoline derivative. Synth Commun 39:4030–4038. https://doi.org/10.1080/00397910902883645 132. Olson JP, Kwon H-B, Takasaki KT, et al (2013) Optically Selective Two-Photon Uncaging of Glutamate at 900 nm. J Am Chem Soc 135:5954–5957. https://doi.org/10.1021/ja4019379 133. Maier W, Corrie JET, Papageorgiou G, et al (2005) Comparative analysis of inhibitory effects of caged ligands for the NMDA receptor. J Neurosci Methods 142:1–9. https://doi.org/10.1016/J.JNEUMETH.2004.07.006 134. Riedl J, Crevenna AH, Kessenbrock K, et al (2008) Lifeact: a versatile marker to visualize F-actin. Nat Methods 5:605–607. https://doi.org/10.1038/nmeth.1220 135. Kingston RE, Chen CA, Rose JK (2003) Calcium Phosphate Transfection. Curr Protoc Mol Biol 63:9.1.1-9.1.11. https://doi.org/10.1002/0471142727.mb0901s63 136. Sabanov V, Nedergaard J (2007) Ca2+-independent effects of BAPTA and EGTA on single-channel Cl− currents in brown adipocytes. Biochim Biophys Acta - Biomembr 1768:2714–2725. https://doi.org/10.1016/J.BBAMEM.2007.07.003 137. Tabatabaee MS, Menard F (2020) L-type Voltage-Gated Calcium Channel Modulators Inhibit Glutamate-Induced Morphology Changes in U118-MG Astrocytoma Cells. Cell Mol Neurobiol 1–9. https://doi.org/10.1007/s10571-020-00828-z 138. Aumann G, Friedländer F, Thümmler M, et al (2017) Quantifying Filopodia in Cultured Astrocytes by an Algorithm. Neurochem Res 42:1795–1809. https://doi.org/10.1007/s11064-017-2193-0 139. Kumar P, Nagarajan A, Uchil PD (2018) Analysis of cell viability by the MTT assay. Cold Spring Harb Protoc 2018:469–471. https://doi.org/10.1101/pdb.prot095505 140. Murphy S V., Atala A (2014) 3D bioprinting of tissues and organs. Nat. Biotechnol. 32:773–785 141. Fedoryak OD, Sul JY, Haydon PG, Ellis-Davies GCR (2005) Synthesis of a caged glutamate for efficient one- and two-photon photorelease on living cells. Chem Commun 3664–3666. https://doi.org/10.1039/b504922a 142. Tian Z, Clark BLM, Menard F (2019) Kainic Acid-Based Agonists of Glutamate Receptors: SAR Analysis and Guidelines for Analog Design. ACS Chem Neurosci 10:4190–4198. https://doi.org/10.1021/acschemneuro.9b00349 143. Stott DI (1989) Immunoblotting and dot blotting. J. Immunol. Methods 119:153–187 144. Krzywinski M, Altman N (2014) Comparing samples—part I. Nat Methods 11:215–216. 116 https://doi.org/10.1038/nmeth.2858 145. Krzywinski M, Altman N (2014) Comparing samples—part II. Nat Methods 11:355–356. https://doi.org/10.1038/nmeth.2900 146. Stepulak A, Luksch H, Gebhardt C, et al (2009) Expression of glutamate receptor subunits in human cancers. Histochem Cell Biol 2009 1324 132:435–445. https://doi.org/10.1007/S00418-009-0613-1 147. Tian Z (2019) Synthesis of kainic acid derivatives and their application as kainate receptors probes. University of British Colombia 148. Tian Z, Menard F (2018) Synthesis of Kainoids and C4 Derivatives. J Org Chem 83:6162–6170. https://doi.org/10.1021/acs.joc.8b00179 149. Mega A, Hartmark Nilsen M, Leiss LW, et al (2020) Astrocytes enhance glioblastoma growth. Glia 68:316–327. https://doi.org/10.1002/glia.23718 150. Roussos ET, Condeelis JS, Patsialou A (2011) Chemotaxis in cancer. Nat. Rev. Cancer 11:573–587 151. Centuries ST, Principles AT (2019) Pharmacology Rang and Dale 152. Jin R, Banke TG, Mayer ML, et al (2003) Structural basis for partial agonist action at ionotropic glutamate receptors. Nat Neurosci 6:803–810. https://doi.org/10.1038/nn1091 153. Huettner JE (2003) Kainate receptors and synaptic transmission. Prog. Neurobiol. 70:387–407 154. Simms BA, Zamponi GW (2014) Neuronal voltage-gated calcium channels: Structure, function, and dysfunction. Neuron 82:24–45 155. Cheli VT, Santiago González DA, Namgyal Lama T, et al (2016) Conditional Deletion of the L-Type Calcium Channel CaV1.2 in Oligodendrocyte Progenitor Cells Affects Postnatal Myelination in Mice. J Neurosci 36:10853–10869. https://doi.org/10.1523/JNEUROSCI.1770-16.2016 156. Shaw RM, Colecraft HM (2013) L-type calcium channel targeting and local signalling in cardiac myocytes. Cardiovasc Res 98:177–186. https://doi.org/10.1093/cvr/cvt021 157. Hille B (2001) Ion channel of excitable membranes, 3rd ed. Sinauer 158. Daschil N, Obermair GJ, Flucher BE, et al (2013) CaV1.2 calcium channel expression in reactive astrocytes is associated with the formation of amyloid-β plaques in an Alzheimer’s disease mouse model. J Alzheimers Dis 37:439–51. https://doi.org/10.3233/JAD-130560 159. Maklad A, Sharma A, Azimi I (2019) Calcium Signaling in Brain Cancers: Roles and Therapeutic Targeting. Cancers (Basel) 11:. https://doi.org/10.3390/CANCERS11020145 160. Prakriya M (2013) Store-Operated Orai Channels: Structure and Function. Curr Top Membr 71:1–32. https://doi.org/10.1016/B978-0-12-407870-3.00001-9 161. RK M, MC H-G, X Z, et al (2013) STIM1 and Orai1 mediate CRAC channel activity and are essential for human glioblastoma invasion. Pflugers Arch 465:1249–1260. https://doi.org/10.1007/S00424-013-1254-8 162. J C, Y L, R Y, et al (2014) Transient receptor potential (TRP) channels, promising potential diagnostic and therapeutic tools for cancer. Biosci Trends 8:1–10. https://doi.org/10.5582/BST.8.1 163. SS K, KS H, BM K, et al (2010) Caffeine-mediated inhibition of calcium release channel inositol 1,4,5-trisphosphate receptor subtype 3 blocks glioblastoma invasion and extends survival. Cancer Res 70:1173–1183. https://doi.org/10.1158/0008-5472.CAN-09-2886 164. Woolley CS, Gould E, Frankfurt M, McEwen BS (1990) Naturally occurring fluctuation in dendritic spine density on adult hippocampal pyramidal neurons. J Neurosci 10:4035–9. https://doi.org/10436047 165. Liao P, Soong TW (2010) CaV1.2 channelopathies: From arrhythmias to autism, bipolar disorder, and immunodeficiency. Pflugers Arch Eur J Physiol 460:353–359. https://doi.org/10.1007/s00424-009-0753-0 166. Lao QZ, Kobrinsky E, Liu Z, Soldatov NM (2010) Oligomerization of Ca v β subunits is an essential correlate of Ca 2+ channel activity. FASEB J 24:5013–5023. https://doi.org/10.1096/fj.10.165381 167. Eisenberg MJ, Brox A, Bestawros AN (2004) Calcium channel blockers: An update. Am. J. Med. 116:35–43 168. Striessnig J, Koschak A (2008) Channels Exploring the function and pharmacotherapeutic potential -of voltage-gated Ca 2+ channels with gene-knockout models. https://doi.org/10.4161/chan.2.4.5847 169. Heine M, Heck J, Ciuraszkiewicz A, Bikbaev A (2020) Dynamic compartmentalization of calcium channel signalling in neurons. Neuropharmacology 169:107556 170. Robinson P, Etheridge S, Song L, et al (2011) Targeting of voltage-gated calcium channel α2δ-1 subunit to lipid rafts is independent from a GPI-anchoring motif. PLoS One 6:. https://doi.org/10.1371/journal.pone.0019802 117 171. Harrison PJ, Tunbridge EM, Dolphin AC, Hall J (2020) Voltage-gated calcium channel blockers for psychiatric disorders: Genomic reappraisal. Br J Psychiatry 216:250–253. https://doi.org/10.1192/bjp.2019.157 172. Justin S. Bryans *, Natasha Davies, Nicolas S. Gee, et al (1998) Identification of Novel Ligands for the Gabapentin Binding Site on the α2δ Subunit of a Calcium Channel and Their Evaluation as Anticonvulsant Agents. https://doi.org/10.1021/JM970649N 173. Wang M, Offord J, Oxender DL, Su TZ (1999) Structural requirement of the calcium-channel subunit alpha2delta for gabapentin binding. Biochem J 342 ( Pt 2):313–20 174. Taylor CP, Angelotti T, Fauman E (2007) Pharmacology and mechanism of action of pregabalin: The calcium channel α2–δ (alpha2–delta) subunit as a target for antiepileptic drug discovery. Epilepsy Res 73:137–150. https://doi.org/10.1016/J.EPLEPSYRES.2006.09.008 175. Zhang FX, Gadotti VM, Souza IA, et al (2018) BK Potassium Channels Suppress CaVα2δ Subunit Function to Reduce Inflammatory and Neuropathic Pain. Cell Rep 22:1956–1964. https://doi.org/10.1016/j.celrep.2018.01.073 176. Alles SRA, Garcia E, Balasubramanyan S, et al (2018) Peripheral nerve injury increases contribution of L-type calcium channels to synaptic transmission in spinal lamina II: Role of α2δ–1 subunits. Mol Pain 14:174480691876580. https://doi.org/10.1177/1744806918765806 177. Ablinger C, Geisler SM, Stanika RI, et al (2020) Neuronal α2δ proteins and brain disorders. Pflugers Arch. Eur. J. Physiol. 472:845–863 178. Dolphin AC (2010) Calcium channel α2δ subunits in epilepsy and as targets for antiepileptic drugs. Epilepsia 51:82. https://doi.org/10.1111/j.1528-1167.2010.02868.x 179. Alvarez-Laviada A, Kadurin I, Senatore A, et al (2014) The inhibition of functional expression of calcium channels by prion protein demonstrates competition with α2δ for GPI-anchoring pathways. Biochem J 458:365–374. https://doi.org/10.1042/BJ20131405 180. Alles SRA, Cain SM, Snutch TP (2020) Pregabalin as a Pain Therapeutic: Beyond Calcium Channels. Front. Cell. Neurosci. 14:83 181. Ghosh D, Nieves-Cintrón M, Tajada S, et al (2018) Dynamic L-type CaV1.2 channel trafficking facilitates CaV1.2 clustering and cooperative gating. Biochim Biophys Acta - Mol Cell Res 1865:1341–1355. https://doi.org/10.1016/J.BBAMCR.2018.06.013 182. Pravettoni E, Bacci A, Coco S, et al (2000) Different localizations and functions of L-type and N-type calcium channels during development of hippocampal neurons. Dev Biol 227:581–594. https://doi.org/10.1006/dbio.2000.9872 183. Kang S, Cooper G, Dunne SF, et al (2012) CaV1.3-selective L-type calcium channel antagonists as potential new therapeutics for Parkinson’s disease. Nat Commun 3:1146–1147. https://doi.org/10.1038/ncomms2149 184. Ortner NJ, Bock G, Vandael DHF, et al (2014) Pyrimidine-2,4,6-triones are a new class of voltage-gated L-type Ca2+ channel activators. Nat Commun 5:1–10. https://doi.org/10.1038/ncomms4897 185. Tabatabaee MS, Kerkovius J, Menard F (2021) Design of an Imaging Probe to Monitor Real-Time Redistribution of L-type Voltage-Gated Calcium Channels in Astrocytic Glutamate Signaling. Mol Imaging Biol 1–10. https://doi.org/10.1007/s11307-020-01573-x 186. Subbarao K V., Stolzenburg J-U, Hertz L (1995) Pharmacological characteristics of potassium-induced, glycogenolysis in astrocytes. Neurosci Lett 196:45–48. https://doi.org/10.1016/0304-3940(95)11834-J 187. Maneshi MM, Sachs F, Hua SZ (2015) A Threshold Shear Force for Calcium Influx in an Astrocyte Model of Traumatic Brain Injury. J Neurotrauma 32:1020–1029. https://doi.org/10.1089/neu.2014.3677 188. Sinnegger-Brauns MJ, Huber IG, Koschak A, et al (2009) Expression and 1,4-dihydropyridine-binding properties of brain L-type calcium channel isoforms. Mol Pharmacol 75:407–414. https://doi.org/10.1124/mol.108.049981 189. Wick A, Wick W, Hirrlinger J, et al (2004) Chemotherapy-induced cell death in primary cerebellar granule neurons but not in astrocytes: in vitro paradigm of differential neurotoxicity. J Neurochem 91:1067–1074. https://doi.org/10.1111/j.1471-4159.2004.02774.x 190. Fehrenbacher JC, Taylor CP, Vasko MR (2003) Pregabalin and gabapentin reduce release of substance P and CGRP from rat spinal tissues only after inflammation or activation of protein kinase C. Pain 105:133–141. https://doi.org/10.1016/S0304-3959(03)00173-8 191. Chen J, Li L, Chen SR, et al (2018) The α2δ-1-NMDA Receptor Complex Is Critically Involved in Neuropathic Pain Development and Gabapentin Therapeutic Actions. Cell Rep 22:2455–2468. https://doi.org/10.1016/j.celrep.2018.02.021 192. Krzywinski M, Altman N (2014) Analysis of variance and blocking. Nat Methods 11:699–700. 118 https://doi.org/10.1038/nmeth.3005 193. Waters JC (2007) Live-Cell Fluorescence Imaging. Methods Cell Biol. 81:115–140 194. Dunn KW, Kamocka MM, McDonald JH (2011) A practical guide to evaluating colocalization in biological microscopy. Am. J. Physiol. - Cell Physiol. 300:C723 195. Costes S V., Daelemans D, Cho EH, et al (2004) Automatic and quantitative measurement of protein-protein colocalization in live cells. Biophys J 86:3993–4003. https://doi.org/10.1529/biophysj.103.038422 196. Manders EMM, Verbeek FJ, Aten JA (1993) Measurement of co‐localization of objects in dual‐colour confocal images. J Microsc 169:375–382. https://doi.org/10.1111/j.1365-2818.1993.tb03313.x 197. Zamora NN, Cheli VT, Santiago González DA, et al (2020) Deletion of voltage-gated calcium channels in astrocytes during demyelination reduces brain inflammation and promotes myelin regeneration in mice. J Neurosci 40:3332–3347. https://doi.org/10.1523/JNEUROSCI.1644-19.2020 198. Anekonda TS, Quinn JF (2011) Alzheimer ’ s disease : the case for isradipine. Biochim Biophys 2011 1812:1584–1590. https://doi.org/10.1016/j.bbadis.2011.08.013.Calcium 199. Hao J, Bao X, Jin B, et al (2015) Ca2+ channel subunit 1D promotes proliferation and migration of endometrial cancer cells mediated by 17 -estradiol via the G protein-coupled estrogen receptor. FASEB J 1–11. https://doi.org/10.1096/fj.14-265603 200. Thul PJ, Åkesson L, Wiking M, et al (2017) A subcellular map of the human proteome. Science 356:eaal3321. https://doi.org/10.1126/science.aal3321 201. Uhlen M, Oksvold P, Fagerberg L, et al (2010) Towards a knowledge-based Human Protein Atlas. Nat Biotechnol 28:1248–1250. https://doi.org/10.1038/nbt1210-1248 202. Xu W, Lipscombe D (2001) Neuronal Ca(V)1.3alpha(1) L-type channels activate at relatively hyperpolarized membrane potentials and are incompletely inhibited by dihydropyridines. J Neurosci 21:5944–5951. https://doi.org/21/16/5944 [pii] 203. Stirling PC, Lundin VF, Leroux MR (2003) Getting a grip on non-native proteins. EMBO Rep. 4:565–570 204. Dolphin AC (2018) Voltage-gated calcium channel α 2δ subunits: an assessment of proposed novel roles. F1000Research 7:. https://doi.org/10.12688/f1000research.16104.1 205. Zhang Z, Zhao W, Lin X, et al (2019) Voltage-dependent calcium channel α2δ1 subunit is a specific candidate marker for identifying gastric cancer stem cells. Cancer Manag Res 11:4707–4718. https://doi.org/10.2147/CMAR.S199329 206. Song L, Espinoza-Fuenzalida I, Etheridge S, et al (2015) The R-Domain: Identification of an N-terminal Region of the α2δ-1 Subunit Which is Necessary and Sufficient for its Effects on Cav2.2 Calcium Currents. Curr Mol Pharmacol 8:169–179. https://doi.org/10.2174/1874467208666150507104555 207. Risher WC, Eroglu C (2020) Emerging roles for α2δ subunits in calcium channel function and synaptic connectivity. Curr Opin Neurobiol 63:162–169. https://doi.org/10.1016/j.conb.2020.04.007 208. Dolphin AC (2012) Calcium channel auxiliary α 2 δ and β subunits: Trafficking and one step beyond. Nat. Rev. Neurosci. 13:542–555 209. Christopher Risher W, Kim N, Koh S, et al (2018) Thrombospondin receptor α2δ-1 promotes synaptogenesis and spinogenesis via postsynaptic Rac1. https://doi.org/10.1083/jcb.201802057 210. Dolphin AC (2013) The α2δ subunits of voltage-gated calcium channels. Biochim Biophys Acta - Biomembr 1828:1541–1549. https://doi.org/10.1016/J.BBAMEM.2012.11.019 211. Eroglu Ç, Allen NJ, Susman MW, et al (2009) Gabapentin Receptor α2δ-1 Is a Neuronal Thrombospondin Receptor Responsible for Excitatory CNS Synaptogenesis. Cell 139:380–392. https://doi.org/10.1016/j.cell.2009.09.025 212. Wu J, Yan Z, Li Z, et al (2016) Structure of the voltage-gated calcium channel Cav1.1 at 3.6 Å resolution. Nature 537:191–196. https://doi.org/10.1038/nature19321 213. Eroglu C, Allen NJ, Susman MW, et al (2009) Supplemental data, Gabapentin receptor alpha2delta-1 is a neuronal thrombospondin receptor responsible for excitatory CNS synaptogenesis. Cell 139:380–92. https://doi.org/10.1016/j.cell.2009.09.025 214. Christopherson KS, Ullian EM, Stokes CCA, et al (2005) Thrombospondins are astrocyte-secreted proteins that promote CNS synaptogenesis. Cell 120:421–433. https://doi.org/10.1016/j.cell.2004.12.020 215. Wang Y, Fehlhaber KE, Sarria I, et al (2017) The Auxiliary Calcium Channel Subunit α2δ4 Is Required for Axonal Elaboration, Synaptic Transmission, and Wiring of Rod Photoreceptors. 119 Neuron 93:1359-1374.e6. https://doi.org/10.1016/j.neuron.2017.02.021 216. Hill DR, Suman-Chauhan N, Woodruff GN (1993) Localization of [3H]gabapentin to a novel site in rat brain: autoradiographic studies. Eur J Pharmacol Mol Pharmacol 244:303–309. https://doi.org/10.1016/0922-4106(93)90156-4 217. Caldwell KA, Tucci ML, Armagost J, et al (2009) Investigating Bacterial Sources of Toxicity as an Environmental Contributor to Dopaminergic Neurodegeneration. PLoS One 4:e7227. https://doi.org/10.1371/journal.pone.0007227 218. Yu J, Wang S, Zhao W, et al (2018) Mechanistic exploration of cancer stem cell marker voltage-dependent calcium channel a2d1 subunit-mediated chemotherapy resistance in small-cell lung cancer. Clin Cancer Res 24:2148–2158. https://doi.org/10.1158/1078-0432.CCR-17-1932 219. Sui X, Geng JH, Li YH, et al (2018) Calcium channel α2δ1 subunit (CACNA2D1) enhances radioresistance in cancer stem-like cells in non-small cell lung cancer cell lines. Cancer Manag Res 10:5009–5018. https://doi.org/10.2147/CMAR.S176084 220. The Human Protein Atlas. https://www.proteinatlas.org/. Accessed 20 Apr 2021 221. Whittaker CA, Hynes RO (2002) Distribution and evolution of von Willebrand/integrin A domains: widely dispersed domains with roles in cell adhesion and elsewhere. Mol Biol Cell 13:3369–87. https://doi.org/10.1091/mbc.e02-05-0259 222. Zhulin IB, Nikolskaya AN, Galperin MY (2003) Common extracellular sensory domains in transmembrane receptors for diverse signal transduction pathways in bacteria and archaea. J Bacteriol 185:285–94. https://doi.org/10.1128/JB.185.1.285-294.2003 223. Salah Ud-Din AIM, Roujeinikova A (2017) Methyl-accepting chemotaxis proteins: a core sensing element in prokaryotes and archaea. Cell Mol Life Sci 74:3293–3303. https://doi.org/10.1007/s00018-017-2514-0 224. Taylor CP, Harris EW (2020) Analgesia with gabapentin and pregabalin may involve N-methyl-D-aspartate receptors, neurexins, and thrombospondins. J. Pharmacol. Exp. Ther. 374:161–174 225. Supasitthumrong T, Bolea-Alamanac BM, Asmer S, et al (2019) Gabapentin and pregabalin to treat aggressivity in dementia: a systematic review and illustrative case report. Br. J. Clin. Pharmacol. 85:690–703 226. Hundehege P, Fernandez-Orth J, Römer P, et al (2018) Targeting Voltage-Dependent Calcium Channels with Pregabalin Exerts a Direct Neuroprotective Effect in an Animal Model of Multiple Sclerosis. Neurosignals 26:77–93. https://doi.org/10.1159/000495425 227. Glaum SR, Holzwarth JA, Miller RJ (1990) Glutamate receptors activate Ca2+ mobilization and Ca2+ influx into astrocytes. Proc Natl Acad Sci U S A 87:3454–8. https://doi.org/10.1073/pnas.87.9.3454 228. Joseph DJ, Williams DJ, MacDermott AB (2011) Modulation of neurite outgrowth by activation of calcium-permeable kainate receptors expressed by rat nociceptive-like dorsal root ganglion neurons. Dev Neurobiol 71:818–35. https://doi.org/10.1002/dneu.20906 229. Kamijo S, Ishii Y, Horigane SI, et al (2018) A Critical Neurodevelopmental Role for L-Type Voltage-Gated Calcium Channels in Neurite Extension and Radial Migration. J Neurosci 38:5551–5566. https://doi.org/10.1523/JNEUROSCI.2357-17.2018 230. Petrovic MM, Viana Da Silva S, Clement JP, et al (2017) Metabotropic action of postsynaptic kainate receptors triggers hippocampal long-term potentiation. Nat Neurosci 20:529–539. https://doi.org/10.1038/nn.4505 231. Rodríguez-Moreno A, Sihra TS (2007) Metabotropic actions of kainate receptors in the CNS. J Neurochem 103:2121–2135. https://doi.org/10.1111/j.1471-4159.2007.04924.x 232. Evans AJ, Gurung S, Henley JM, et al (2019) Exciting Times: New Advances Towards Understanding the Regulation and Roles of Kainate Receptors. Neurochem Res 44:572–584. https://doi.org/10.1007/s11064-017-2450-2 233. Kurshan PT, Oztan A, Schwarz TL (2009) Presynaptic α2δ-3 is required for synaptic morphogenesis independent of its Ca2+-channel functions. Nat Neurosci 12:1415–1423. https://doi.org/10.1038/nn.2417 122 Appendices Appendix A: Scheme of synthesis of DECM-PhKA Chemical probe (2S,3S,4S)-3-(Carboxymethyl)-1-(((7-(diethylamino)-2-oxo-2H-chromen-4-yl)methoxy)carbonyl)-4-phenylpyrrolidine-2-carboxylic acid, DECM-PhKA (1) was developed to control the activity of kainic acid (KA) receptors in cells. Phenylkainic acid (PhKA) is photo-released with a half lifetime of 61 seconds upon irradiation with blue light. Precise spatio-temporal control of cellular stimulation was achieved using an optic fiber using a physiologically compatible wavelength. The caged kainoid 1 (DECM-PhKA) acts as a “turn-on” probe to study KAR activity in living cells. The secondary amine of kainoids is essential to their binding, thus, protecting it with a photocleavable carbamate makes 1 biologically inactive. The synthesis, characterization, and application of DECM-PhKA (1) to control KAR-mediated filopodia extension in glial cells has been described by Tian et.al in detail [60, 70, 148]. Figure appendix.1 Synthesis of DECM-PhKA scheme. 123 Appendix B: Processes extension in response to glutamate is a faster response compared to KA, in U118-MG astrocytoma cells. To determine whether there is a difference in the time scale of morphological response to kainoids compared to glutamate mCherry-LifeAct7 transfected U118-MG cells were imaged by an Olympus FV1000 fluorescence confocal microscopy equipped with a Plan-ApoN 60x/i.4 oil-immersion objective. Changes in cell morphology were recorded with 60X objective lenses. To track the process's extension in time, an image was acquired (one second) at 20 seconds intervals. Using intervals and short exposure to laser reduced the photobleaching of the fluorescent signals and maximized cell survival during the course of the experiments. Processes motility was recorded for 15 minutes after stimulating cells with glutamate and kainoids. Images were analyzed for filopodiagenesis. The time span from stimulating the cells with the agonists till the processes extension was recorded and plotted. Each value represents the average time in seconds from three individual cells, each from a separate culture dish (n = 3). The results were Figure appendix.2 In U118-MG astrocytoma cells, processes extension in response to glutamate is a faster response compared to KA. Time elapsed before the initiation of process extension after chemical stimuli, Data are shown mean ± SEM 124 plotted using GraphPad prism 7.04 and are presented as the mean ± SEM. Upon exposure to glutamate, cells started protruding their processes 70 ± 15 seconds after stimulation while it took 300 ± 20 seconds to initiate the change in the length of processes upon exposure to KA and PhKA (230 seconds time gap) (Figure Appendix. 2). 125 Appendix C: Synthesis of MNI-Glu (S)-2-Amino-5-(4-methoxy-7-nitroindolin-1-yl)-5-oxopentanoic acid, MNI-Glu was prepared by Zhenlin Tian according to modified literature procedures. Characterization data matched literature precedents [60, 70, 131, 141]. Figure appendix.3 Synthesis of MNI-Glu scheme. 126 Appendix D: Synthesis of FluoBar1 FluoBar1, an imaging probe for L-type voltage-gated calcium channels, was created from a barbiturate ligand modified with a fluorescent coumarin moiety. An overview of the synthesis is presented in Figure appendix. 4. The convergent synthesis consists of ten reactions with six steps for the longest linear sequence (14% overall yield). Starting materials are: cyclopentane carboxylic acid (2), 2-(3-chlorophenyl) acetonitrile, and 2-aminoethanol. All procedures for the synthesis of the imaging probes, reagents, instruments, and chemical characterization are detailed in the Tabatabaee et.al report [185]. Figure appendix.4 Synthesis of imaging probe FluoBar1 (12). The fluorescent moiety is introduced at the last step to allow maximal flexibility in the type of desired reporter 127 Appendix E: Rapid dot blot assay with FluoBar1 Since the results presented in Chapter 3 has shown the selectivity of FluoBar1 for CaV1.2, I envisaged FluoBar1 protein as a simple and rapid staining tool to substitute antibody labeling in immuno-dot blotting. Dot blotting (or dot immunobinding) provides a quick way to detect the presence of a protein of interest and/or to assess whether an antibody is not selective (often the secondary antibody) [143]. It is a variant of Western blotting in which antigens are detected in samples spotted directly on to a membrane without prior separation. Herein, I compared antibodies and the imaging probe, FluoBar1, to selectively label L-type voltage-gated calcium channels in a cell lysate—CaV1.2 and CaV1.3 in particular. Methods: Cell lysate. Cells were resuspended with prewarmed 0.25% trypsin-EDTA solution (Gibco SH30236.02), then transferred to a 15 mL falcon tube, and spun down at 1000 rpm for 5 min at 4 ºC. The pellet was mixed with 50 μL of 1% protease inhibitor cocktail (Amresco, mammalian M250) in sterile Millipore deionized water. Then, 50 μL of NP40 lysis buffer (50 mM Tris-HCl, NaCl 150 mM, 1% NP40-Triton 100x pH 7.4) was added to the solution and incubated for 15 min at 4ºC. The mixture was centrifuged at 3000 rpm for 5 min at 4 ºC. The pellet was discarded, and the supernatant was stored at –20 ºC. Rat brain lysate. Hippocampal brain tissue from eight weeks rats was homogenized in an ice-cooled buffer containing: Triton100x 1%, 50 mM Tris-HCl (pH 7.4), NaCl 150 mM, EDTA 10 mM, EGTA 10 mM, and 1% protease inhibitor cocktail. The homogenized lysate was centrifuged at 14,000 rpm at 4 ºC, the pellet was discarded, and the supernatant was stored at –20 ºC until needed. Immuno-dot blotting. Cell lysate aliquots (1 μl) were blotted on two dry nitrocellulose membranes (GE Healthcare, Amersham Protran Sandwich 0.45 μm, GE10600114). Membranes were then blocked for 30 minutes on a rocking plate by blocking buffer: 3% bovine serum albumin 128 (BSA, Alfa aesar J64777, New Zealand origin) in 1X Tris-buffered saline + 0.1% Tween (TBS-T). TBS consists Tris 20 mM, NaCl 150 mM. The membranes were washed three times with 1X TBS-T for 5 min each, then followed by incubation with the primary antibody. One membrane was treated with anti-CaV1.2 primary antibody (rabbit anti- cacna1C, Alomone ACC-003), and the other with anti-CaV1.3 primary antibody (rabbit anti-cacna1D, Alomone ACC-005); both were incubated for 2 hours at room temperature. The membranes were washed 3 times with TBS-T for 5 minutes each at room temperature on a rocking platform. Both membranes were then incubated with the same secondary antibody (goat anti-rabbit IgG HRP-conjugated, HAF008) for one hour at room temperature. Membranes were washed three times with TBS-T each for 5 minutes; the blots were visualized by a 1:1 solution of luminol + enhancer (Pierce ECL Western blotting substrate, 32106) using a quantitative Western blot imaging system (Alpha-Innotech FluorChem HD2). Fluobar1 dot-labeling. Cell lysate aliquots (1 μl) were blotted on a dry nitrocellulose membrane. The dried membrane was soaked in a 50 nM FluoBar1 solution in sterile millipore dionized water for 1 minute. The membrane was rinsed twice with sterile millipore deionized water, then immediately placed on a fluorescence confocal microscope slide, and imaged at 10x magnification and under laser excitation at 405 nm (Olympus FV1000, AlexaFluor-405 filter). The membrane was kept wet during the imaging processes. Results and discussion: Membranes stained with the imaging probe FluoBar1 show fluorescent signals in both MCF7 cells and U118-MG cells when irradiated at 405 nm, suggesting that FluoBar1 binds to both CaV1.2 and CaV1.3 proteins (Figure Appendix. 5). This result is consistent with the reports from the Silverman and the Striessnig labs, which shows the affinity of FluoBar1 parent molecule to both MCF7 and U118-MG cells [183, 184]. 129 Immunodot blotting showed the presence of CaV1.2 in U118-MG cells and the rat brain lysate, but not in MCF7 cells and COS-7 cells. (Figure Appendix. 5). The presence of CaV1.3 was observed in rat brain and the MCF7 cell lysate. COS-7 cells were used as a negative control. The immunostainings of MCF7 cells shows that these cells express CaV1.3 but not CaV1.2. When the blotted lysates were stained with FluoBar1 probe, the rat brain lysate showed a robust fluorescent signal. These signals were also observable in U118-MG and MCF7 lysates. Only background levels of fluorescence were present with COS-7 cells, suggesting that minimal nonselective binding occurs with FluoBar1, where the membrane was blotted (Figure Appendix. 5). Figure appendix.5 FluoBar1 shows affinity for both CaV1.2 and CaV1.3. (a) Immuno-dot blotting of cell lysates from MCF7 cells, U118-MG cells, COS-7 cells and brain lysate, imaged by chemiluminescence. (b) FluoBar1 dot-labeling imaged by fluorescence confocal microscopy using AlexaFluor 405 filter, 10x magnification 130 Appendix F: Ligands for α2δ1 blocks the response to kainoids The results from Chapter 2 and Chapter 4 shows that antagonists against KARs and α2δ1 both inhibit the processes extension of astrocytoma cells upon exposure to glutamate. However, how these two receptors interplay to conduct the processes extension is not known. To identify the possible crosstalk between KARs and α2δ1, the cells that were treated with α2δ1 were stimulated with KAR agonists. Interestingly, α2δ1 antagonists blocked the effect of kainoids in processes extension. Methods: The effect of 100 μM of glutamate (Alfa Aesar A15031-30), kainic acid (Tocris 0222) and Ph-KA (made in-house) on the morphological response of U118-MG cells was studied when cells were pretreated with 2δ1 ligands and compared to the control. mCherry-Lifeact-7 transfected U118-MG cells were pretreated with 10 μM of gabapentinoids before exposure to glutamate (as described in Section 3.2.3). Gabapentinoids: gabapentin (G154, Sigma Aldrich), pregabalin (made in house) were prepared directly in deionized millipore water (vehicle). Stock solutions were prepared concentrated (200X) to avoid osmotic shock. Before imaging, cells were incubated at 37 ºC under a humidified 5% CO2 atmosphere for 30 minutes. Results and discission: The antagonists for α2δ1 blocked the effect of the agonists for KARs. KAR agonists normally induce processes extension in U118-MG cells (Figure 2.4). The change in the processes extension in U118-MG cells upon exposure to 100 μM glutamate, KA or PhKA were significantly different from the cells that were pretreated with gabapentinoids before stimulation by KAR agonists (p < 0.0001). While the cells that were pretreated with gabapentinoids before stimulation by KAR agonists demonstrated no significant difference with the cells that were not stimulated at all. 131 This result suggests either 1) a nonselective crosstalk between agonists and antagonists of these two proteins, or 2) a cooperation between KARs and α2δ1 to initiate the morphological response of astrocytoma cells to glutamate. If α2δ1 is the only essential protein for initiation of glutamate-induced morphological response (GIMR) (based on the results from Section 4.3.5) it is possible that kainoids can non-selectively activate α2δ1. Therefore, the α2δ1 antagonists might have blocked the off-target effect of KAR agonists on α2δ1. However, the results presented in Figure 2.4.B Show that blocking KARs can also blocked the GIMR. Therefore, this interpretation requires further assessment. On the other hand, If KARs are the essential proteins for the initiation of GIMR, it is possible that α2δ1 antagonists—gabapentinoids—have non-selectively blocked KARs. Using a heterologous expression system that only expresses KARs and not α2δ1, could help to test this Figure appendix.6 Ligands against α2δ1 blocked the morphological response that is induced by kainoids in U118-MG astrocytoma cells. U118-MG cells that were pre-treated with 10 μM gabapentinoids were stimulated with 100 µM of KAR agonists: glutamate, KA and Ph-KA and. Comparisons were done by two-way ANOVA (F 3,64 α2δ1 antagonists = 100.1; F 3,64 KAR agonists= 13.6). The data from the cells stimulated with KAR agonists were compared with the data from non-stimulated cells using Sidak’s multiple comparison post hoc (labeled as vehicle of the graph). Then all the bars were compared together using Tukey’s post hoc multiple comparison; n = 5; * p<0.0001 significantly different from the cells that were not stimulated or were pretreated with α2δ1 ligands before stimulation with KAR agonists. Data are shown mean ± SEM. 132 possible nonselective effect of α2δ1 antagonists. Stimulation of the cells, that were pretreated with gabapentinoids, with the agonists for KAR could indicate whether gabapentinoids can non-selectively block KARs. In fact, it is also possible that cooperation and presence of both α2δ1 and KARs is required for the morphological response to glutamate. Both U118-MG and U138-MG cells express KARs. Therefore, expression and overexpression of α2δ1 could have made their response to glutamate possible and measurable. On the other hand, COS-7 cells lack α2δ1, but they may express KARS (other subunits except GluK2 as has been shown in Figure 2.3.D). If they express some combinations of KAR subunits (which we have not tested), it is possible that the exogenously introducing α2δ1 has triggered their morphological response to glutamate as a result of the cooperation of α2δ1 with KARS. Indeed, further investigation is required to demonstrate and identify the mechanism of the interplay between α2δ1 and KARs in glutamate-induced morphological response.
Thesis/Dissertation
eng
Biochemistry and Molecular Biology
Vancouver : University of British Columbia Library
Attribution-NonCommercial-NoDerivatives 4.0 International
Graduate
Menard, Frederic
Glutamate induced morphological response in astrocytoma cells
Text