Medicine, Faculty of
DSpace
UBCV
Long, Rory Kirk Mackenzie
Master of Science - MSc
University of British Columbia
The Zika virus (ZIKV) is a positive-sense RNA flavivirus that has been determined as a causative agent of severe neurological diseases including Guillain-Barré syndrome and microcephaly. A common characteristic of flavivirus infection is the re-organization of host cell membranes, usually endoplasmic reticulum (ER) derived, to promote viral replication. As ZIKV-induced ER structures, being vesicular replication factories (RF) and convoluted membranes (CM), are smaller than the diffraction limit of light, studies to characterize these structures and their protein compositions have been limited. In this thesis, Stimulated Emission Depletion (STED) super resolution microscopy is utilized to study the whole cell organization of ZIKV-induced ER morphologies and several viral proteins involved in replication. Generally in cells, the ER can be divided into two regions, central ER sheets and peripheral ER tubules. Here we report the formation of an ZIKV-induced, organized crescent-shaped, dense central ER region of ER tubules. ZIKV RFs localize to this dense, central ER region in a similar crescent-shaped organization. Computational 3D reconstruction of these 3D STED imaged ZIKV-induced ER structures, along with validation by electron microscopy of ZIKV infected cerebral organoids, revealed that these dense ER regions are composed of novel smooth ER tubular matrices. Moreover, we have found that ZIKV NS4B, a viral encoded integral membrane protein predicted to play a role in ER membrane re-organization during ZIKV replication, is enriched in ZIKV-induced tubular matrices. Additionally, a subpopulation of ZIKV NS4B localizes to active RFs suggesting that NS4B plays a role in the formation of both ZIKV-induced tubular matrices and RFs among other functions. Overall, the application of super resolution microscopy to study ZIKV replication has led to the identification of novel ZIKV-induced, RF-containing tubular matrices and characterization of a viral protein involved in this process.
3D Stimulated Emission Depletion super resolution microscopy reveals novel association between replication factories and organized smooth tubular matrices during Zika virus infection by Rory Kirk Mackenzie Long B.Sc., The University of British Columbia, 2017 A THESIS SUBMITTED IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF MASTER OF SCIENCE in THE FACULTY OF GRADUATE AND POSTDOCTORAL STUDIES (Cell and Developmental Biology) THE UNIVERSITY OF BRITISH COLUMBIA (Vancouver) December 2019 © Rory Kirk Mackenzie Long, 2019 ii The following individuals certify that they have read, and recommend to the Faculty of Graduate and Postdoctoral Studies for acceptance, the thesis entitled: 3D Stimulated Emission Depletion super resolution microscopy reveals novel association between replication factories and organized smooth tubular matrices during Zika virus infection submitted by Rory Long in partial fulfillment of the requirements for the degree of Master of Science in Cell & Developmental Biology Examining Committee: Robert Nabi, Cellular & Physiological Sciences Co-supervisor Francois Jean, Microbiology and Immunology Co-supervisor Supervisory Committee Member Lacey Samuels, Botany Additional Examiner Additional Supervisory Committee Members: Wayne Vogl, Cellular & Physiological Sciences Supervisory Committee Member Eric Jan, Biochemistry and Molecular Sciences Supervisory Committee Member iii Abstract The Zika virus (ZIKV) is a positive-sense RNA flavivirus that has been determined as a causative agent of severe neurological diseases including Guillain-Barré syndrome and microcephaly. A common characteristic of flavivirus infection is the re-organization of host cell membranes, usually endoplasmic reticulum (ER) derived, to promote viral replication. As ZIKV-induced ER structures, being vesicular replication factories (RF) and convoluted membranes (CM), are smaller than the diffraction limit of light, studies to characterize these structures and their protein compositions have been limited. In this thesis, Stimulated Emission Depletion (STED) super resolution microscopy is utilized to study the whole cell organization of ZIKV-induced ER morphologies and several viral proteins involved in replication. Generally in cells, the ER can be divided into two regions, central ER sheets and peripheral ER tubules. Here we report the formation of an ZIKV-induced, organized crescent-shaped, dense central ER region of ER tubules. ZIKV RFs localize to this dense, central ER region in a similar crescent-shaped organization. Computational 3D reconstruction of these 3D STED imaged ZIKV-induced ER structures, along with validation by electron microscopy of ZIKV infected cerebral organoids, revealed that these dense ER regions are composed of novel smooth ER tubular matrices. Moreover, we have found that ZIKV NS4B, a viral encoded integral membrane protein predicted to play a role in ER membrane re-organization during ZIKV replication, is enriched in ZIKV-induced tubular matrices. Additionally, a subpopulation of ZIKV NS4B localizes to active RFs suggesting that NS4B plays a role in the formation of both ZIKV-induced tubular matrices and iv RFs among other functions. Overall, the application of super resolution microscopy to study ZIKV replication has led to the identification of novel ZIKV-induced, RF-containing tubular matrices and characterization of a viral protein involved in this process. v Lay Summary The Zika virus is an RNA virus that is transmitted largely by mosquitoes and is currently present in 86 countries. During the 2015 Zika virus outbreak in Brazil, Zika virus infection was characterized by the development of serious neurological diseases such as Guillain-Barré syndrome in adults and microcephaly in infants. When RNA viruses infect a host cell, they must hijack the cell’s proteins and organelles to create the optimal environment to replicate. The main cellular organelle hijacked by the Zika virus is the endoplasmic reticulum (ER), which is turned into a factory that produces new virus particles. By investigating Zika virus-induced ER morphology and the mechanism behind this process in brain cells at the nanoscale level, we look to shed light onto how the Zika virus replicates in brain cells. Additionally, by better understanding Zika virus replication in the ER, antiviral compounds that inhibit this process can be created. vi Preface Thesis Contributions: Long, R.K.M., Jean F., and Nabi, I.R. conceived and designed this thesis project. Long, R.K.M performed all experiments and analysis of the resultant data. Cardoen, B. wrote a Jupyter notebook to aid in data analysis. Chapter 2 is based in part on a manuscript in preparation for publication: “Deep learning of STED super resolution microscopy identifies Zika virus-induced reorganization of the endoplasmic reticulum” by Long, R.K.M., Moriarty, K., Cardoen, B., Gao, G., Vogl, A.W., Hamarneh, G., Jean, F., and Nabi, I.R. Contributions: Long, R.K.M., Jean, F., and Nabi, I.R. conceived, designed, and analyzed all experiments. Long, R.K.M., Moriarty, K., Nabi, I.R., and Hamarneh, G., designed the deep learning analysis of the data. Long, R.K.M., Moriarty, K., and Nabi, I.R. wrote the manuscript. Long, R.K.M., and Moriarty, K. performed all experiments and prepared all figures. Vogl, A.W. performed the electron microscopy imaging. Cardoen, B. wrote a Jupyter notebook to aid in data analysis. vii Table of Contents Abstract ………………………………………………………………………..…. iii Lay Summary....………………………………………………………………....….v Preface ……………………………………………………………………………. vi Table of Contents ………………………………………………………...…….... vii List of Figures ………………………………………………………………....….. ix List of Abbreviations …………………………………………………………..…. xi Acknowledgements …………………………………………………………...… xiv Chapter 1:Introduction ……………………………………………………….….... 1 1.1 The endoplasmic reticulum (ER) ……………………………….….... 1 1.1.1 Protein synthesis and trafficking ……………………..……..… 1 1.1.2 Ca2+ regulation …………………………………………...….... 3 1.1.3 Lipid synthesis ………………………………………………... 5 1.1.4 Unfolded Protein Response (UPR) …………………………… 6 1.1.5 ER-Associated Degradation (ERAD) ……………………....… 8 1.2 ER structure …………………………………………………..……... 9 1.2.1 Peripheral ER and ER tubules ……………………………….. 11 1.2.2 Central ER sheets ………………………………………….… 13 1.2.3 Cooperation of ER proteins to form the peripheral ER network ……………………………………………………………….. 15 1.2.4 Role of the cytoskeleton in ER morphology ……………..….. 17 1.2.5 Drug-induced central ER smooth convoluted tubules …….… 18 1.2.6 Membrane contact sites ……………………………..………. 19 1.3 The role of the ER in flaviviral replication ………………………… 22 1.3.1 The Flaviviridae family …………………………..….……… 22 1.3.2 The role of lipid droplets in flaviviral replication ……….….. 25 1.3.3 Flaviviral hijacking of the ER …………………………....…. 26 1.3.4 Convoluted membranes in viral replication …………….....… 27 1.3.5 Replication factories …………………………………....…… 28 1.3.6 Hijacked host proteins involved in membrane re-organization ……………………………………………………………….. 31 1.4 Zika virus (ZIKV) manipulation of host ER membranes …….….… 34 1.4.1 Brief history of the ZIKV …………………………..……..… 34 1.4.2 ZIKV lifecycle ………………………………………....……. 35 1.4.3 ZIKV-induced ER re-organization ……………………….…. 38 1.5 Super resolution microscopy ………………………………….…… 40 1.5.1 Structured Illumination Microscopy (SIM) ……………….… 41 viii 1.5.2 Stimulated Emission Depletion Microscopy (STED) ……..… 42 1.5.3 Single Molecule Localization Microscopy (SMLM) ……..… 43 1.5.4 Super resolution microscopy in the study of ER morphology and dynamics ………………………………………………….…. 44 1.5.5 Use of super resolution microscopy to study viruses ……..… 47 1.6 Objective ……………………………………………………..…….. 49 Chapter 2: 3D STED super resolution microscopy reveals novel association between replication factories and organized smooth tubular matrices during Zika virus infection ……………………………………………………………...………….. 51 2.1 Introduction ……………………………………………..…….…… 51 2.2 Results ………………………………………………………..….… 55 2.2.1 3D Stimulated Emission Depletion (STED) microscopy reveals ZIKV-induced central ER morphological changes in U87 Glioblastoma cells ……………………………………...…… 55 2.2.2 Zika virus infection induces the formation of a dense ER tubular matrix within the central ER …………………..………….… 57 2.2.3 The ZIKV NS4B and NS2B proteins localize differently within the ER ……………………………………………………….. 58 2.3 Discussion ………………………………………………...………... 60 2.4 Materials and Methods ……………………………………...……… 79 2.4.1 Antibodies, plasmids, chemicals, and viruses ……………….. 79 2.4.2 Cell culture and stable cell line generation ………………...... 79 2.4.3 Infection and immunofluorescence labeling ………………… 80 2.4.4 3D STED microscopy …………………………………....….. 81 2.4.5 Transmission electron microscopy ………………………….. 81 2.4.6 Quantification and statistical analysis ……………………….. 82 Bibliography ………………………………………………………………...….... 83 Appendix ………………………………………………………………..……… 101 ix List of Figures Figure 1-1 Central ER and peripheral ER domains in mammalian cells ………… 10 Figure 1-2 Cooperation of ER shaping proteins to shape the ER ………………... 17 Figure 1-3 Cellular functions of ER-mitochondria contact sites …………….....… 21 Figure 1-4 General ER re-organization during flaviviral infection …………….… 24 Figure 1-5 General ZIKV lifecycle and polyprotein organization …………….…. 37 Figure 1-6 STED microscopy utilizes a secondary doughnut-shaped depletion laser to achieve sub-diffraction limited resolution …………………………………….. 43 Figure 1-7 3D-SIM super resolution microscopy reveals classically described peripheral sheet structures to instead be dense tubular matrices …………….…… 47 Figure 2-1 3D STED microscopy reveals the presence of ZIKV-induced ER re-organization in U87 Glioblastoma cells …………………………………………. 68 Figure 2-2 Defining the central and peripheral ER of ZIKV and mock infected cells by Imaris segmentation to better characterize ZIKV-induced ER re-organization …………………………………………………………………...………………. 70 Figure 2-3 Central ER (CER) and Peripheral ER (PER) masks reveal changes in ER domain volume ………………………………………………………….……….. 72 Figure 2-4 The majority of dsRNA localizes to a central ER subdomain …….…. 73 x Figure 2-5 The ZIKV-induced perinuclear region of dense ER is a network of closely packed, dsRNA labeled tubular ER that encompasses a region of lower ER density …………………………………………………………….……….… 74 Figure 2-6 Labeling of the ZIKV NS4B and NS2B proteins reveal that both proteins localize to the central ER while NS4B also displays peripheral ER localization with an enrichment in the crescent-shaped dense ER region …………………...……... 76 Figure 2-7 dsRNA colocalizes with distinct subpopulations of NS4B and NS2B in the crescent-shaped dense, tubular ER region. ………………………………….... 78 Figure A-1 Ultrastructural analysis of ZIKV infected cerebral brain organoids reveal perinuclear, smooth convoluted ER membranes that contain ZIKV induced vesicles ………………………………………………………………………………..… 101 Figure A-2 Labeling of the ZIKV NS4B protein reveals both central and peripheral Sec61β-GFP ER localization with an enrichment in the dense ER region …...…. 103 Figure A-3 ZIKV infected cells displaying dsRNA is largest at 48 hours ……… 104 Figure A-4 Both ZIKV NS2B and NS4B antibodies produce significant mean fluorescence intensity in ZIKV infected cells at concentrations of 1:500 …...… 105 xi List of Abbreviations +ssRNA, positive sense single-stranded RNA ATF6, activating transcription factor 6 ATLs, Atlastins bHLH, basic Helix-Loop-Helix C protein, capsid protein CDP, cytidine diphosphate CER, central ER CERT, ceramide transport protein CHOP, CCAAT-enhancer-binding protein homologous protein CLEM, correlative light and electron microscopy Climp-63, cytoskeleton-linking membrane protein 63 CM, convoluted membrane DC-SIGN, dendritic cell-specific intercellular adhesion molecule-3-grabbing non-integrin DENV, dengue virus DP1, depleted in polyposis 1 dsRNA, doubled-stranded RNA dSTORM, direct stochastic optical reconstruction microscopy DUBs, deubiquitinases E protein, envelope protein EB1, end-binding protein 1 EGF, epithelial growth factor EM, electron microscopy ER, endoplasmic reticulum ERAD, ER-associated degradation ERMES, ER-mitochondria encounter structure FAM134, family with sequence similarity 134 FASN, fatty acid synthase FGF, fibroblast growth factor FIB, focus ion beam GBV-B, GB virus-B GI-SIM, grazing incidence structured illumination microscopy GPCR, G protein-coupled receptor GRP94, glucose-regulated protein 94 HCV, hepatitis C virus HIV-1, Human immunodeficiency virus type 1 hNPCs, human neuronal progenitor cells HRD1, hydoxymethylglutaryl reductase degradation protein 1 Hsp40, heat stock protein 40 IP3R, inositol 1,4,5-triphosphate (IP3) receptors IRE1, inositol requiring enzyme 1 JEV, Japanese encephalitis virus LD, lipid droplet xii Lnp, Lunapark M protein, membrane protein MAVs, mitochondrial antiviral signaling protein MARCH6, membrane-associated ring-CH-type finger 6 MCS, membrane contact sites MDA5, melanoma differentiation-associated protein 5 MFN2, mitofusin2 mRNA, messenger RNA NADPH, nicotinamide adenine dinucleotide phosphate NS protein, non-structural protein OBP oxysterol-binding protein OSBP oxysterol binding protein PAT, Perilipin, Adilofilin/ADRP and the Tail interacting protein PC, paracrystalline array PDI, protein disulfide isomerase PDMP, phenyl-2-decanoyl-amino-3-morpholino-1-propanol-hydrochloride PDZD8, PDZ domain containing 8 PER, peripheral ER PERK, double-stranded RNA-activated protein kinase-like ER kinase PIP2 phosphatidylinositol 4,5 bisphosphate pRM protein, precursor membrane protein PtdCho, phosphatidylcholine PtdEtn, phosphatidylethanolamine PtdIns, phosphatidylinositol PtdSer, phosphatidylserine RF, replication factory RHD, reticulon homology domain RIG1, retinoic acid-inducible gene 1 ROI, region of interest RTN, reticulon SEL1L, suppressor/enhancer of Lin-12-like SERCAs, sarcoendoplasmic reticulum Ca2+ ATPases SIM, structured illumination microscopy SMLM, single molecule localization microscopy SMP, synaptotagmin-like mitochondrial-lipid binding protein SRP, signal recognition particle STED, STimulated Emission Depletion STIM1, stromal interaction molecule 1 TAG, triacylglycerides TBEV, tick-borne encephalitis virus, TIM-1, T-cell immunoglobulin and mucin domain 1 UPR, unfolded protein response VAPB-PTPIP51 vesicle-associated membrane protein-associated protein B- protein tyrosine phosphatase interacting protein 51 VCP, valosin containing protein VDAC, voltage-dependent anion channel xiii WNV, West Nile virus XBP1, X-box binding protein 1 YFV, yellow fever virus ZIKV, Zika virus xiv Acknowledgements Firstly, I would like to thank both Dr. Robert Nabi, and Dr. Francois Jean for their wonderful support throughout both my Master’s project and my applications for various scholarships, conferences and future endeavours. By both agreeing to step outside of their area of expertise to co-supervise me, we were able to explore an area of research that neither lab could study on its own. After many hours of helpful discussion and experimental planning, I believe this collaboration was very fruitful. I hope to apply what I learned during this time to succeed in the future research I conduct. Secondly, I would like to thank all of lab members, collaborators and technicians that I worked with or besides during my time in the Nabi and Jean Labs. I am grateful for role models such as Dr. Bharat Joshi, Dr. Sharlene Eivemark, Dr. Libin Abraham and Dr. Guang Gao, whom thoroughly taught me the different techniques utilized during my Master’s and were always available for consultation when I had questions. Thirdly, I would like to thank my Master’s committee, Dr. Wayne Vogl and Dr. Eric Jan, for their guidance and advice both during and outside of committee meetings. Their critical analysis of my results and experiments aided me in designing better experiments and presenting my results in a clearer, more concise manner. Lastly, I would like to thank my family and friends for supporting me throughout this time. I was faced with many challenges since starting my Master’s project and my life has taken many unexpected turns along the way, however, the support I have received from family, friends, and lab members has been exceptional. Thank you. 1 Chapter 1: Introduction 1.1 The endoplasmic reticulum (ER) The endoplasmic reticulum (ER) is a continuous membrane network that spans the cytoplasm from the nucleus to the plasma membrane. It is the largest cellular membrane bound organelle and is composed of multiple ER domains defined initially by the presence of ribosomes under electron microscopy (EM): sheet-like rough ER or, the lack thereof, being tubular smooth ER (Palade and Porter, 1954, Shibata et al., 2006). In addition to being morphologically complex, the ER plays a critical role in a multitude of cellular processes including membrane and secreted protein synthesis and trafficking, Ca2+ regulation, lipid synthesis, the formation of membrane contact sites with other organelles, and several cellular stress response pathways (Walter and Ron, 2011; Rowland and Voeltz, 2012; Schwarz and Blower, 2016). 1.1.1 Protein synthesis and trafficking A major function of the ER is to serve as a platform where the biosynthesis of secreted and integral membrane proteins takes place (Jan, Williams, and Weissman, 2014). Once the translation of the amino terminus of secretory or integral membrane proteins on cytoplasmic ribosomes begins, these ribosomes are recruited to the ER membrane. The binding of the signal peptide within the translated amino terminus to the signal recognition particle (SRP) pauses peptide elongation and functions to target the messenger RNA (mRNA): ribosome:nascent polypeptide complexes to the ER (Walter, Ibrahimi, and Blobel, 1981). At the 2 ER, the newly formed mRNA:ribosome:nascent polypeptide:SRP complex docks to the ER membrane localized SRP receptor (Gilmore, Blobel, and Walter, 1982). Binding to the SRP receptor provides the signal for translation to continue and the protein enters the ER lumen co-translationally through the Sec61 translocon (Park and Rapoport, 2012). Co-translational or post-translational cleavage of the signal peptide allows the release of the protein into the ER. From there the protein either remains in the ER lumen or becomes anchored into the ER membrane as an integral protein through the intrinsic presence of at least one hydrophobic transmembrane domain (Evans, Gilmore, and Blobel, 1986). In the ER lumen the newly formed protein interacts with ER-resident chaperone and folding proteins, such as calnexin, calreticulin, immunoglobulin heavy chain-binding protein (BiP), and protein disulfide isomerase (PDI) to facilitate proper protein folding and post-translational modifications required for correct protein trafficking and function (Braakman and Hebert, 2013). Such modifications include N-linked glycosylation, disulfide bond formation and oligomerization. In addition to proteins destined for the ER, a subset of cytosolic proteins has been characterized as being translated on ER-bound ribosomes. While it is not clear how the mRNAs of these proteins are recruited to the ER it has recently been elucidated that the ER-resident protein p180 may play a role in the process (Cui, Zhang, and Palazzo, 2012). Following correct folding and post-translationally modification within the ER, secretory and non-ER integral membrane proteins are then packaged into COPII-coated transport vesicles at ER exit sites (Orci et al., 1993). Here the vesicles bud off of the ER and travel to the Golgi where these proteins undergo further modification. After traveling through the Golgi cisternae, the proteins are loaded into vesicles at the trans-Golgi network and travel to the plasma membrane if they are to be secreted or to their target organellar membrane. Therefore, the ER acts as the first step of the 3 secretory pathway involved in the production and transport of secreted and integral membrane proteins to the Golgi. 1.1.2 Ca2+ regulation Another major role of the ER is functioning as the major store of Ca2+ within the cell (Clapham, 2007; Samtleben et al., 2013). The pumping of sarcoendoplasmic reticulum Ca2+ ATPases (SERCAs) maintains a Ca2+ concentration in the ER lumen of 100-800 µM while cytosolic concentration of Ca2+ is ~100 nM (Clapham, 2007, Samtleben et al., 2013). To regulate the cytoplasmic concentration of Ca2+, the ER possesses calcium receptors, inositol 1,4,5-triphosphate (IP3) receptors (IP3R), and ryanodine receptors that release Ca2+ into the cytoplasm upon detection of low intracellular Ca2+ levels (Clapham, 2007). Ca2+ release begins with the activation of the G protein-coupled receptor (GPCR) which then stimulates phospholipase C to cleave phosphatidylinositol 4,5 bisphosphate (PIP2) into diacyl-glycerol and IP3 (Schwarz and Blower, 2016). The binding of free IP3 to IP3R leads to initial Ca2+ release into the cytoplasm, which is then increased by the opening of Ca2+-induced ryanodine receptors (Endo, 2009). Additional ryanodine receptor Ca2+ release can occur upon conformational changes in other intracellular voltage-dependent Ca2+ channels such as dihydropyridine receptors (Fill and Copello, 2002). Another source of Ca2+ for the cell is the extracellular environment, which contains a Ca2+ concentration of ~2 mM (Samtleben et al., 2013). Extracellular Ca2+ can be used to alter both cytoplasmic and ER luminal Ca2+ concentrations allowing the cell many mechanisms by which Ca2+ concentration can be further regulated to maintain homeostasis. As Ca2+ is an important cellular signalling molecule spatial and temporal regulation is required. 4 A number of essential cell functions are controlled by cytoplasmic Ca2+ levels such as cellular proliferation, differentiation, secretion, contraction, metabolism, trafficking, gene transcription, and apoptosis (Berridge, Lipp, and Bootman, 2000). An area of research in which the role of Ca2+ signaling has been extensively explored is during neural development. Throughout mammalian neural development, neural progenitor cells must proliferate, migrate and ultimately differentiate into the many distinct cellular populations that form the brain (Toth, Shum and Prakriya, 2016). Ca2+ signaling has been shown to play crucial roles in each of these processes. During neural development, many extrinsic growth factors, such as epithelial growth factor (EGF) and fibroblast growth factor (FGF), utilize Ca2+ channels to promote and regulate neural progenitor proliferation (Weissman et al., 2004). In addition, precise migration of these highly proliferating neural progenitor cells has been shown to be both facilitated and inhibited depending on changes in intracellular Ca2+ levels (Bando et al., 2014). Moreover, once neural progenitor cells reach their destination, they must undergo differentiation into their respective cell type. Ca2+ signaling plays a role in neural cell differentiation in two ways. Intracellular Ca2+ levels are responsible for regulating the activity of the basic Helix-Loop-Helix (bHLH) gene, NeuroD, a transcription factor, along with other transcription factors, that drives the survival and differentiation of many neuronal cell lineages in the brain (Gaudilliere et al., 2004). Once neuronal differentiation has been activated, increases in voltage-gated Ca2+ channel expression lead to a greater intrinsic ability of Ca2+ signaling in these cells, promoting neuronal excitability, neurotransmitter specification, axonal pathfinding, and dendritic outgrowth which further shapes the final neuronal phenotype of these cells (Toth, Shum and Prakriya, 2016). Overall, the mammalian brain is one example of the 5 role that Ca2+ signaling plays each step of the development of complex, heterogenous cellular populations. 1.1.3 Lipid synthesis In addition to protein synthesis, the ER is a major site of membrane lipid and lipid precursor synthesis. ER resident membrane proteins cytidine diphosphate (CDP)-choline:1,2-diacylglycerol cholinephosphotransferase, and CDP-choline:1,2-diacylglycerol choline/ethanolamine phosphotransferase synthesize the two most abundant lipid components found in cellular membranes: phosphatidylcholine (PtdCho) and phosphatidylethanolamine (PtdEtn) respectively (van der Veen et al., 2017). Other ER resident enzymes are involved in the synthesis of less abundant membrane lipids such as ceramide, an important component of sphingolipids, cholesterol, phosphatidylserine (PtdSer), and phosphatidylinositol (PtdIns) (Tidhar and Futerman, 2013, Jacquemyn, Cascalho, and Goodchild, 2017). The ER is also involved in the formation of triacylglycerides (TAG), important components of cellular energy storage (Lange et al., 1999). Several lipids require enzymes from different organelles to be produced, therefore these lipids such as cardiolipin are not produced in the ER. However, as almost all of these lipids require ER-synthesized precursor lipids as upstream building blocks, the ER acts as the central regulator of lipids across all cellular membranes (Jacquemyn, Cascalho, and Goodchild, 2017). Besides playing a major role in lipid synthesis, the ER is responsible for bringing enzymes involved in lipid synthesis and lipid transfer, such as the ceramide transport protein (CERT) and oxysterol binding protein (OSBP) that mediate transfer of sterols and sphingolipids, into close proximity to other cellular membranes (Petit et al., 2019). While studies elucidating where lipid 6 enzymes are localized in the ER are still in their infancy, there is evidence that several of these transmembrane enzymes are localized to sites of the ER that are positioned near the membrane of their target organelle. Such examples include the positioning of enzymes involved in TAG synthesis to regions of lipid droplet formation and enzymes involved in PtdCho synthesis, PtdCho being the most common lipid in the plasma membrane, to sites of ER contact with the plasma membrane (Wilfling et al., 2013, Kaplan and Simoni, 1985, Jain and Holthuis, 2017). Additionally, PtdSer has been shown to be synthesized at the mitochondria-associated membrane allowing for rapid entry into the mitochondria where it is decarboxylated into PtdEtn by mitochondrial PtdSer decarboxylase 1 (Schumacher, Choi, and Voelker, 2002). Further, the enrichment of ER lipid transfer proteins oxysterol-binding protein5/8 (ORP5/ORP8) at ER-mitochondria contact sites, through interaction with the outer mitochondrial protein, protein tyrosine phosphatase interacting protein 51 (PTP1P5), allow for the rapid transport of PtdSer to the mitochondrial membrane (Petit et al., 2019). By localizing ER lipid synthesizing and transfer enzymes to sites of organellar contact, lipids can be rapidly moved around the cell maintaining organelle and cell homeostasis. 1.1.4 Unfolded Protein Response (UPR) The lumen of the ER is an oxidative environment which, together with the many ER chaperone proteins, is highly conducive to proper protein folding and disulfide bond formation. Many different disturbances, such as hypoxia, viral infection, and glucose deprivation, however, can result in the accumulation of unfolded proteins within the ER causing ER stress (Kim, Xu, and Reed, 2008). In the event that the protein-folding capacity of the ER lumen decreases or is overwhelmed leading to the build-up of misfolded proteins there are a collection of three 7 conserved, metazoan signalling pathways known collectively as the unfolded protein response (UPR) to sense the insufficiency and alter gene expression to increase the protein-folding capacity of the ER lumen (Walter and Ron, 2011). The three branches are inositol requiring enzyme 1 (IRE1), double-stranded RNA-activated protein kinase (PKR)-like ER kinase (PERK), and activating transcription factor 6 (ATF6) (Walter and Ron, 2011). The activation of each of these branches leads to the production of b-ZIP transcription factors which activate UPR target genes, however, the mechanism by which each pathway relieves ER stress is different. By splicing the mRNA of the transcription factor X-box binding protein 1 (XBP1), IRE1 promotes lipid synthesis and the expansion of the ER lumen to provide more space for protein folding, increases expression of ER chaperone proteins such as BiP and glucose-regulated protein 94 (GRP94) and ER-associated degradation proteins, and promotes cellular mRNA decay to decrease the ER protein folding load (Reimold et al., 2001). On the other hand, the PERK oligomerizes and indirectly inactivates eIF2a, which inhibits mRNA translation and prevents the further accumulation of unfolded proteins in the ER (Hughes and Mallucci, 2019). Furthermore, PERK indirectly activates CCAAT-enhancer-binding protein homologous protein (CHOP) which controls genes involved in apoptosis (Nishitoh, 2012). In this way prolonged PERK activation due to failure in resolving ER stress leads to apoptosis. Additionally, as with IRE1, PERK also promotes expression of ER chaperone proteins (Walter and Ron, 2011). Lastly, the activation of the third branch of the UPR, ATF6, directly promotes the expression of chaperone proteins to counteract the buildup of unfolded proteins (Walter and Ron, 2011). Together, IRE1, PERK, and ATF6 act synergistically to the counteract the accumulation of unfolded proteins in the ER lumen. Clinically, the dysregulation of the UPR, particularly the PERK pathway, has been closely associated with 8 neurodegenerative disease leading a field of research into therapeutic strategies that can restore a functional UPR in these individuals (Hughes and Mallucci, 2019). 1.1.5 ER-associated degradation (ERAD) The ER possesses a rigorous quality control system, that works in tandem with IRE1 of the UPR, to continuously assess proteins within its lumen and remove those that are terminally misfolded (Hwang and Qi, 2018). This process is known as ER-associated degradation (ERAD). By the degradation of misfolded proteins, ERAD maintains the function of the ER by preventing the buildup of toxic aggregates that underlie many human diseases such as Parkinson’s disease, autonomic neuropathy, cardiomyopathy, and nephropathy (Tanaka et al., 2004; Yamamoto et al., 2007). Through ERAD, misfolded proteins are removed from the ER and escorted to the 26S proteasome for degradation (Smith, Ploegh, and Weissman, 2011). The ERAD pathway begins first by the recognition of misfolded substrates. While this process is largely unknown, it has been shown that ER chaperone proteins such as BiP play a dual role in protein folding and ERAD targeting, identifying aberrantly exposed hydrophobic regions of misfolded substrates (Okuda-Shimizu and Hendershot, 2007). In mammals, the most conserved ERAD machinery is the suppressor/enhancer of Lin-12-like (SEL1L)- hydroxymethylglutaryl reductase degradation protein 1 (HRD1) protein complex, where HRD1, a E3 ligase and dislocation channel, together with its cofactor SEL1L, mediate the transfer of activated ubiquitin to either the N-terminus or a lysine residue of misfolded substrates from a E2 ubiquitin-conjugating enzyme. Additional E3 ligases such as GP78 or membrane-associated ring-CH-type finger 6 (MARCH6) have also been determined to play roles in ubiquitin transfer (Stevenson, Huang, and 9 Olzmann, 2016). Once ubiquitinated misfolded substrates cross ERAD dislocation channels, such as HRD1, into the cytoplasm by utilizing energy derived from ATP hydrolysis by the ubiquitin-dependent AAA-ATPase valosin containing protein (VCP) (Meyer, Bug, and Bremer, 2012). In the cytoplasm chaperone complexes maintain the solubility of the dislocated, misfolded proteins as they travel to the 26S proteasome where the polyubiquitin tag is removed by deubiquitinases (DUBs) and the misfolded protein is degraded. 1.2 ER structure The advent of high-resolution electron microscopy (EM) and confocal microscopy techniques have revealed the ER to be a morphological complex but interconnected organelle consisting of a peripheral ER tubular network, nuclear envelope, and perinuclear, central ER sheets (Figure 1-1). The function of these distinct regions largely remain to be elucidated, however, by EM it is known that central ER sheets contain ribosomes and therefore are the sites of protein synthesis while the peripheral ER tubules lack ribosomes and are believed to be where lipid synthesis occurs (Shibata, Voeltz, and Rapoport, 2006). In addition to being morphologically complex, the ER is also a very dynamic organelle where ER tubules can slide on microtubules and rapidly fuse with each other to create the peripheral ER tubular network. An example of the dynamic nature of the ER occurs during mitosis in metazoans where the network of peripheral ER tubules collapses into ER sheets (Lu, Ladinsky, and Kirchhausen, 2009; Wang et al., 2013; Wang et al., 2016). ER morphology is maintained and regulated according to the local balance of different ER-shaping proteins (Wang et al., 2016). Therefore, different cell types express ER-shaping proteins at different levels to meet their functional needs. Secretory cells require large amounts of central ER sheets for protein 10 secretion while cells that require high amounts of lipid biogenesis, such as muscle cells, have more peripheral ER tubules. Figure 1-1 Central ER and peripheral ER domains in mammalian cells. ER network of Cos-7 cells labeled with mCherry-KDEL (red) and ER junctions labeled with Lunapark 1-GFP (green). The ER network is a continuous membrane system composed of ER tubules which make up the peripheral ER and ER sheets (denoted here as ER cisternae) that populate the perinuclear central ER region. Nucleus is denoted by N. (Adapted from ER structure and function by Chen, S., Novick, P., and Ferro-Novick, S., 2013 © 2013 by Current Opinions in Cell Biology) 11 1.2.1 Peripheral ER and ER tubules The peripheral ER consists largely of ER tubules, yet peripheral ER sheet-like structures that contain holes, known as fenestrated ER sheets, have been shown by EM to also be present (Puhka et al., 2012). ER tubules are structures of high cross-sectional membrane curvature with a diameter of 60-100 nm (Figure 1-2A). The proteins involved in inducing the high membrane curvature of ER tubules have been identified as those of the reticulon (RTN) protein family (Figure 1-2C). Proteins in the RTN family share a highly conserved C-terminal reticulon homology domain (RHD) which contains two hydrophobic motifs separated by a 66 amino-acid hydrophilic loop (Ramo et al., 2016). It has been proposed that the two hydrophobic regions of the RHD form two hairpin loops that act as wedges in the outer membrane leaflet of the ER membrane. By producing an increase in local surface area on the cytoplasmic side of the membrane while not impacting the lumenal side high membrane curvature can be created. Members of the RTN family including RTN1, RTN2, RTN3, and two isoforms of RTN4, being RTN4A and RTN4B, have been reported to generate curved ER structures by forming homo-oligomers and hetero-oligomers together with depleted in polyposis 1 (DP1) (Hu et al., 2008; Shibata et al., 2009). DP1, also known as REEP5 in metazoans and Yop1p in Saccharomyces cerevisiae, like the RTN proteins contains a conserved domain with two hydrophobic hairpins that can form a wedge-like structure in membranes. It has been determined by computer modeling that these oligomer arcs only need to occupy approximately 10% of the total ER membrane surface to generate an ER tubule and therefore do not impede the motility of other ER membrane proteins and lipids (Hu et al, 2008). The peripheral ER consists of sheet-like ER regions and ER tubules that fuse at three-way junctions, small triangular sheets with negatively curved edges, to produce a tubular matrix. The 12 protein components involved in the homotypic fusion of ER tubules are the Atlastins (ATLs), a group of membrane-anchored GTPases, and Rab GTPases (Figure 1-2C). Currently, there are four different ATL proteins being ATL1, ATL2, ATL3, and ATL4, which all are known to mediate homotypic fusion of ER tubules (Wang et al., 2016). When an ER tubule makes contact with another tubule ATL molecules on both membranes form trans-dimers (Orso et al., 2009; Bian et al., 2011). After multiple dimerization events occur, conformation changes of these dimers triggered by GTP-hydrolysis pull the two opposing membranes together and fusion occurs (Liu et al., 2015). After fusion, the ATL proteins diffuse away from the resulting three-way junction to participate in future homotypic fusion events. Interestingly, Rab GTPases are required for ER tubule extension likely by interacting with molecular motors such as kinesin-1 or dynein and play a role in homotypic fusion that is separate from that of ATL yet the exact function of Rab in this process remains unknown (English and Voeltz, 2013). The importance of the ATLs in maintaining the peripheral ER tubular matrix in addition to forming three-way junctions has been determined through the expression of non-functional ATL proteins in U2OS cells. When ATL mutants that cannot dimerize are introduced long, unbranched ER tubules are seen while large regions of the ER tubular network also disassemble into small ER vesicles (Wang et al., 2016). Thus, it has been purposed that the free ends of ER tubule produced by the inhibition of homotypic fusion are highly unstable, energetically favouring the formation of vesicles by the curvature induced by RTN and DP1 (Wang et al., 2016). Therefore, ATL could play a role in stabilizing free tubule ends and three-way junctions to prevent the destabilization of the ER into vesicles. In addition to ATLs, Lunapark (Lnp) a conserved membrane protein that contains two closely spaced transmembrane domains and a Zn2+-finger domain is found to preferentially localize to 13 three-way junctions (Chen et al., 2012). While Lnp is not essential for the generation or maintenance of three-way junctions or the ER tubular matrix as seen with ATL, it localizes to the sheet-like region of the three-way junction where it prevents this region from expanding (Wang et al., 2016). Further, during mitosis Lnp is both degraded and phosphorylated, preventing it from localizing to three-way junctions, leading to expansion of the three-way junctions into peripheral ER sheets similar to that seen during Lnp depletion studies (Lu et al., 2009; Shemesh et al., 2014; Wang et al., 2016). 1.2.2 Central ER sheets ER sheets consist of two closely apposing, low curvature ER membranes, with exception to high curvature sheet edges (Shibata, Voeltz, and Rapoport, 2006) (Figure 1-2D). The thickness of the sheets is consistently between 60-100 nm, similar to that of ER tubules, allowing for the movement of ER resident proteins, such as calnexin and BiP, through the ER lumen. To identify the cellular components that play a role in the formation of ER sheets Shibata et al., performed mass spectrometry on rough microsomes produced by proliferated ER sheet containing dog pancreatic cells (Shibata, Voeltz, and Rapoport, 2006). This study, along with immunofluorescence, determined that in addition to the enrichment of components of the translocon, such as the Sec61 complex, ER sheets also are enriched with the coiled-coil domain proteins ribosome-binding protein 1 or p180, cytoskeleton-linking membrane protein 63 (Climp-63), and kinectin (Shibata et al., 2010). Such findings have led to the current model of ER sheet formation where the luminal domains of coiled-coil domain ER membrane proteins located on the closely apposed membranes interact and form a bridge that holds ER sheets to a constant thickness, acting as ER luminal spacers (Shibata et al., 2010). 14 Climp-63 is a type 2 transmembrane protein found in humans, yet possesses no apparent homologues in bacteria, yeast, C. elegans or insects. In addition to the single transmembrane region of Climp-63, this protein consists of a cytosolic C terminus that contains a microtubule binding domain and a 91 nm rod-like luminal domain. This luminal domain of Climp-63 can form homo-oligomeric complexes with Climp-63 molecules located on the opposing ER membrane (Sandoz and van der Goot, 2015). The role of the microtubule binding domain of Climp-63 in ER sheet formation is unknown, however, by homo-dimerization of Climp-63 molecules on the two membranes of ER sheets Climp-63 has been shown to act as a luminal ER spacer (Figure 1-2D). Evidence regarding the importance of Climp-63 as an ER spacing protein has come from RNAi knock-down experiments showing that while the depletion of Climp-63 does not lead to a loss of ER sheets, it results in a decrease in ER sheet luminal width from 50 nm down to 25-30 nm (Shibata et al., 2010). Interestingly, while Climp-63 is not required for ER sheet formation, overexpression studies reveal Climp-63 to induce a proliferation of ER sheets at high protein concentrations (Shibata et al., 2010). Additional knock-down studies were conducted on the other ER enriched coiled-coil proteins p180 and kinectin, which both possess coiled-coil domains on the cytoplasmic side of the protein rather than on the luminal side as with Climp-63. The depletion of these two proteins alone did not result in a change in ER luminal width nor was there a synergistic effect when Climp-63, p180, and kinectin were all knocked down at once (Shibata et al., 2010). Hence, p180 and kinectin do not have a role in the spacing of the ER lumen, however, the cytoplasmic nature of their coiled-coil domains may form rod-like scaffolds that keep the central ER sheets flattened (Shibata et al., 2010)(Figure 1-2D). Overall, while the role p180 and kinectin play in generating flat ER sheets is unknown, Climp-63 has been shown to both regulate the luminal width 15 of the ER sheets and upon increased levels of expression lead to the formation of additional ER sheets. While little is still known about how the formation of the flat surface of ER sheets occurs, the high curvature sheet edges are better understood. Similar to ER tubules, the curvature of ER sheets is generated by the oligomerization of ubiquitous reticulon proteins and DP1 into arc-like scaffolds (Hu, Prinz, and Rapoport, 2011, Ramo et al., 2016). Localization studies have revealed these proteins to clearly localize to ER sheet edges stabilizing the region (Shibata et al., 2009; Schuck et al., 2009) (Figure 1-2D). Once central ER sheets are formed, they often stack and are connected by twisted, helicoidal membranes similar to that of a parking garage (Terasaki et al., 2013). By stacking together, the ER sheets can be densely packed while optimizing membrane surface area upon which protein synthesizing ribosomes can dock (Terasaki et al., 2013). 1.2.3 Cooperation of ER proteins to form the peripheral ER network Overexpression and depletion studies have revealed that the ER tubular matrix is formed and maintained by the cooperation of RTN, ATL, Rab, and to a smaller extent Lnp proteins (Wang et al., 2016). Overexpression of RTN leads to the formation of long, unbranched tubules, possibly due to the exclusion of ATL from tubular tips by RTN oligomers (Wang et al., 2016). When additional co-overexpression of ATL is performed, these long, unbranched tubules are again able to form a tubular matrix (Wang et al.,2016). Further, the depletion of RTN results in the proliferation of peripheral ER sheets and loss of tubular ER similar to that seen during Climp-63 16 overexpression or Rab depletion (Voeltz et al., 2006; Shibata et al., 2010, English and Voeltz, 2013). It has been predicted that this proliferation of sheets in the place of ER tubules occurs due to the exclusion of RTNs by Climp-63 overexpression or the depletion of membrane curvature generating RTNs while Rab depletion hinders the formation of new ER tubules causing the formation of peripheral ER sheets (Shibata et al., 2010; English and Voeltz, 2013). Overall, a balance of high expression levels of membrane curvature proteins, RTNs and DP1, and three-way junction and branching tubule forming proteins, ATL, Rab, and Lnp, along with low expression levels of luminal spacing proteins, such as Climp-63, must be maintained by the cell to produce a tubular ER matrix and a functional sheet/tubule ratio. Deviation from this balance allows cells of different functional physiological types or at different life stages such as mitosis to manipulate the sheet/tubule ratio to best meet cellular or physiological needs. 17 Figure 1-2 Cooperation of ER shaping proteins to shape the ER. (A) Systematic showing the presence of both ER tubules and sheets within the peripheral ER. (B) (C) Oligomerization of reticulons along with DP1 (not shown) to form positive curvature inducing arc-like scaffolds. ER tubules form a connected network after undergoing atlastin dependent homotypic fusion. (D) Climp-63 homo-oligomers regulate the space between ER sheet membranes while other ER proteins, p180 and kinectin likely for scaffolds that keep the membranes flattened. Reticulons stabilize the high curvature edges of the ER sheets. (Adapted from Untangling the web: Mechanisms underlying ER network formation by Goyal, U., and Blackstone, C., 2013 © 2013 by Biochim Biophys Acta) 1.2.4 Role of the Cytoskeleton in ER morphology Microtubules work together with ER shaping proteins to regulate the morphology of the ER. By binding to the plus end of growing tips of microtubules or interacting with molecular motors that move along microtubules the ER tubular matrix is able to span the entire cytoplasm of the cell (Du et al., 2004). Stromal interaction molecule 1 (STIM1), an integral ER protein, interacts with end-binding protein 1 (EB1) at the plus end of the microtubule to bind ER tubules to growing microtubule tips, however, which ER proteins are involved in the interaction with molecular motors are unknown (Grigoriev et al., 2008) (Figure 1-2B). While STIM1 is a peripheral ER tubule element, several microtubule binding proteins such as Climp-63, p180, and kinectin have been suggested to play a role in the interaction of central ER sheets with cytoskeleton elements. Further studies based on the disruption of microtubules in mammalian cells reveal that while the depolymerization of microtubules leads to a collapse of the ER network there is a period of delay before the collapse (Terasaki et al., 1986). Therefore, microtubules play a role in the ability of the ER tubular matrix to spread throughout the cell, however, additional cellular factors may participate in the stabilization of the ER network. 18 1.2.5 Drug-induced central ER smooth convoluted tubules Classically, the central ER is thought to be composed of ER sheets, however, the addition of certain drugs to cells can create perinuclear, random tubular networks know as smooth ER patches (Remmer and Merker, 1963; Porter and Bruni, 1959; Tyson and Bulger, 1974; Feldman, Swarm, and Becker, 1980; Sprocati et al., 2006). It has been shown that these smooth ER patches contain ER resident enzymes involved in drug metabolism and therefore, are likely to serve a protective role upon drug uptake (Orrenius and Ericsson, 1966). One such drug is phenyl-2-decanoyl-amino-3-morpholino-1-propanol-hydrochloride (PDMP) which induces the formation of smooth ER patches, up to 5 microns in size, in the rough ER independent of its classical mode of action as a sphingolipid synthesis inhibitor (Sprocati et al., 2006). Interestingly, the protein and lipid compositions of these patches differ from that of other central ER regions and they are not formed from pre-existing ribosome-studded central ER as ribosome detachment was not observed in the study (Sprocati et al., 2006). Therefore, it has been suggested that smooth ER patches are formed from regions of peripheral ER tubules; however, how tubules are incorporated into the largely sheet-like central ER and the mechanism behind their convoluted organization remain unknown (Sprocati et al., 2006). Other drugs such as vinblastine sulfate, an inhibitor of microtubule formation that binds to tubulin; phenobarbital; and 3’-methyl-4-dimethyl aminoazobenzene induce the formation of smooth ER patches that are reversible, as with PDMP, likely due to autophagy of excess SER (Remmer and Merker, 1963; Porter and Bruni, 1959; Tyson and Bulger, 1974; Feldman, Swarm, and Becker, 1980). Further studies into the mechanism behind the proliferation of perinuclear smooth ER membranes upon drug addition could lead to better understanding of how dynamic changes to ER morphology occur and provide better understanding into the role of the ER during pharmacological stress. 19 1.2.6 ER membrane contact sites To maintain proper cell functioning under changing environmental conditions each of the organelles within a cell must communicate with each other. Communication can occur slowly by the use of cargo carrying vesicles, vesicular trafficking, or more rapidly when two communicating organelles have closely apposed membranes. These regions of membranes in close proximity, typically between 15-30 nm, are known as membrane contact sites (MCS) (Helle et al., 2013). The formation of MCSs occurs when two organelles come into close enough proximity that proteins or protein complexes known as tethers can be formed between the two membranes, maintaining the close positioning of the two organelles (English and Voeltz, 2013, Helle et al., 2013). Functionally, MCSs play important roles in lipid and Ca2+ transport, organelle trafficking, and intracellular signaling (Prinz, 2014). The ER acts as a unique organelle as it has been identified to possess membrane contact sites with many of the other organelles including the mitochondria, Golgi, peroxisomes, endosomes, lipid droplets, lysosomes, and the plasma membrane (Levine, 2004). Of these various contacts, ER-mitochondria MCSs are the best characterized (Rowland and Voeltz, 2012). ER-mitochondria MCSs are typically 10-30 nm wide and hence exclude bulky ER-bound ribosomes, leading to the belief that only areas of smooth ER can move into close enough proximity to become tethered to the outer mitochondrial membrane. Several protein complexes have been reported to tether the two membranes at the ER-mitochondrial interface in mammalian cells. Currently, these include mitofusin2 (MFN2) dimers, ER vesicle-associated membrane protein-associated protein B-outer mitochondrial membrane protein tyrosine phosphatase interacting protein 51 (VAPB-PTPIP51) 20 complexes, calnexin-phosphofurin acidic cluster sorting protein 2 (PASC-2) complex, IP3R-voltage-dependent anion channel (VDAC) complex, and PDZ domain containing 8 (PDZD8), a mammalian ER-mitochondria tethering protein with a synaptotagmin-like mitochondrial-lipid binding protein (SMP) domain that is orthologous to that of the yeast ER-mitochondria encounter structure (ERMES) complex (Simmen et al., 2005; Szabadkai et al., 2006; Myhill et al., 2008; Gomez-Suaga et al., 2018; Hirabayashi et al., 2017). Smooth ER-mitochondria contacts have been shown to play a role in lipid biosynthesis, Ca2+ signaling involved in mitochondrial biogenesis, cell death, autophagosome formation and the innate immune response (Rowland and Voeltz, 2012; Filadi, Theurey, and Pizzo, 2017) (Figure 1-3). Interestingly, recent studies have shown regions of close contact, ~50 nm, between rough ER and the mitochondria by EM (Wang et al., 2015). These rough ER-mitochondria contacts have been reported to be regulated by Mfn2, yet the protein composition and function of these contact sites are still undetermined (Wang et al., 2015). 21 Figure 1-3 Cellular functions of ER-mitochondria contact sites. ER-mitochondria contact sites offer regions of interaction between ER and mitochondrial localized proteins to play roles in mitochondrial dynamics, Ca2+ mediated biogenesis, autophagy, and lipid composition. Besides aiding in mitochondrial health, ER-mitochondria contact sites play roles in the initiation of the innate immune response, activation of interleukins IL1b and IL18 for inflammation, and apoptosis. (Adapted from The endoplasmic reticulum-mitochondria coupling in health and disease: Molecules, functions and significance by Filadi, R., Theurey, P., and Pizzo, P., 2017 © 2017 by Cell Calcium) 22 1.3 The role of the ER in flaviviral replication 1.3.1 The Flaviviridae Family Flaviviridae is a viral family of positive sense single-stranded RNA (+ssRNA) arthropod-borne viruses (arboviruses) composed of four closely related viral genera, the flaviviruses, pestiviruses, pegiviruses and hepaciviruses (Stapleton et al., 2011; Romero-Brey and Bartenschlager, 2014). The best characterized genera is the flaviviruses, the largest clade containing almost 70 different viral species and hepaciviruses (Mackenzie, Gubler, and Petersen, 2004; Mazeaud, Freppel, and Chatel-Chaix, 2018). The flaviviruses include members such as the West Nile virus (WNV), dengue virus (DENV), Japanese encephalitis virus (JEV), tick-borne encephalitis virus (TBEV), yellow fever virus (YFV), and Zika virus (ZIKV). The hepaciviruses consist of two members: the hepatitis C virus (HCV) and GB virus-B (GBV-B). Flaviviruses have small genomes, roughly 11 kb in length, that encode 10 proteins that are translated as one polyprotein including 3 structural proteins: Capsid (C), Membrane (M), and Envelope (E) proteins and 7 non-structural (NS) proteins: NS1, NS2A, NS2B, NS3, NS4A, NS4B, and NS5 (Bartenschlager and Miller, 2008; Nitatpattana et al., 2018). Several NS proteins have largely conserved functions such as NS3, along with its cofactor NS2B, which cleave both the viral polyprotein and host proteins by its serine protease domain. NS3 also aids in genomic RNA production through the activity of its RNA helicase and nucleotide triphosphatase domains (Bartenschlager and Miller, 2008). NS5 has conserved RNA-dependent RNA polymerase activity and is also involved in capping newly formed RNA genomes allowing their translation. Despite high degrees of genomic similarity, other viral NS and structural proteins often play distinct roles in different virus lifecycles and therefore 23 clinical manifestations can widely vary from neurological pathogenesis in ZIKV, WNV, and JEV to severe hemorrhage during DENV, and YFV infection (Martin-Acebes, Vazquez-Calvo, and Saiz, 2016). While these viruses also differ in how they hijack host cell pathways, all members of Flaviviridae induce large re-organizations of cellular membranes and lipids to promote their own replication after entry into the cell (Romero-Brey and Bartenschlager, 2014; Hamel et al., 2015) (Figure 1-4). 24 Figure 1-4 General ER re-organization during flaviviral infection. (I) Various flaviviral non-structural proteins, NS4A, NS4B, dimers of NS1, replicase (NS2B, NS3, and NS5), are predicted to be involved in the formation and function of spherical invaginations of the ER known as replication factories. Genomic replication occurs within replication factories and newly synthesized RNA genomes exit through the pore into the cytoplasm to be incorporated by adjacent viral particles budding into the ER lumen. Convoluted membranes form from pre-existing ER membranes and remain connected to the ER, yet spatially distinct from replication factories. (II) Several flaviviruses, such as the Zika virus, accumulate immature virus particles in ER membrane multivesicular body-like structures before being transported to the Golgi and ultimately secreted. (III) Besides ER re-organization and viral particle formation, the ER is also utilized to begin the transport of NS1 hexamers throughout the secretory pathway. Once secreted NS1 plays a role the suppression of the host immune response. (Adapted from Dengue virus non-structural protein 1 modulates infectious particle production via interaction with the structural proteins by Scaturro, P., Cortese, M., Chatel-Chaix, L., Fischl, W., and Bartenschlager, R., 2015 © 2015 by PLoS Pathogen) 25 1.3.2 Flaviviral re-organization of lipid droplets Several members of Flaviviridae, such as ZIKV, HCV and DENV, interact with and promote the formation of cellular lipid droplets (LDs), resulting in clinical liver steatosis during HCV and DENV infection (Bach, Thung and Schaffner, 1992, Coyaud et al., 2018). LDs are organelles derived from the ER that are composed of a core of neutral lipids, mostly triacylglyercols and sterol esters, surrounded by a monolayer of phospholipids that can contain a variable profile of cellular proteins (Martin and Parton, 2006; Thiele and Spandl, 2008). Perilipin, Adilofilin/ADRP and the Tail interacting protein (PAT) proteins are the most common LD associated proteins, however, lipid metabolism enzymes, Rab GTPases, SNARE proteins, and coat proteins I and II have all been identified on LD membranes (Brasaemle, 2007). LDs have been shown to be essential in the replication of various members of Flaviviridae. In DENV, the viral C protein uses the GBF1/Arf/COP1 system to accumulate on LDs where it is believed to modulate host lipid metabolism and utilize LDs as a storage area to sequester its nucleic acid binding domain away from newly synthesized viral genomes before nucleocapsid formation on the ER occurs (Iglesias et al., 2015). In HCV, the core protein accumulates on LDs recruiting NS5A, which further recruits NS3 and NS4AB viral proteins to LDs to form replication complexes. Additionally, NS5A also directly interacts with genomic RNA to promote viral nucleocapsid formation, in contrary to DENV where nucleocapsid formation occurs in the ER (Miyanari et al., 2007; Welsch et al., 2009). The C protein of ZIKV similarly localizes to lipid droplets, however, the functional role of this association remains unknown (Shang et al., 2018). Besides hijacking LDs, it has also been determined that both the number and size of LDs increase early on following DENV and HCV infection (Miyanari et al., 2007; Samsa et al., 2009; Assuncao- 26 Miranda et al., 2010). This increase in LDs has been shown to be induced by the viral C or core protein in both DENV and HCV respectively (Miyanari et al., 2007; Samsa et al., 2009). Conversely as DENV-induced membrane re-organization and replication begins, the number of LDs in the cell decrease (Heaton et al., 2010; Hyrina et al., 2017). The reduction of LDs is driven by the formation of DENV NS4A-NS4B protein complexes (Zhang et al., 2018). By recruiting AUP1 to autophagophores, NS4A-NS4B protein complexes trigger lipophagy to free fatty acids from LDs for membrane re-organization, beta-oxidation, and ATP release. Overall, as LDs play multiple roles in Flaviviridae replication the inhibition of LD formation with various compounds, such as PF-429242, has recently been shown to severely reduce the production of infectious virus particles (Olmstead et al., 2012; Hyrina et al., 2017; Martin-Acebes, de Oya, and Saiz, 2019). 1.3.3 Flaviviral hijacking of the ER +ssRNA viruses possess an RNA genome which can be directly utilized as a template for translation, similar to cellular mRNA (Cortese et al., 2017). Therefore, upon entry into the host cell +ssRNA viruses release their genome into the cytoplasm where it can interact with ribosomes to produce viral proteins required for viral replication (White, Enjuanes, and Berkhout, 2011). Viral RNA and its double stranded intermediate are the target of cellular pattern recognition receptors such as retinoic acid-inducible gene 1 (RIG1) and melanoma differentiation-associated protein 5 (MDA5), which will activate the cell’s antiviral state through mitochondrial antiviral signaling protein (MAVs) (Aktepe and Mackenzie, 2018). Therefore, to sequester viral RNA away from host immune receptors, a common feature of the lifecycle of possibly all +ssRNA viruses is the formation of virus-induced membrane structures during genomic replication (Aktepe and Mackenzie, 2018). Viruses of the Flaviviridae family induce the formation of some of the most 27 well characterized virus-induced organelle-like membranous structures (Aktepe and Mackenzie, 2018). Initial EM studies conducted on a strain of the WNV called the Kunjin virus revealed that several ultrastructural changes occurred during virus replication in monkey kidney Vero and C6/36 mosquito cell lines. It was determined that during viral replication the formation of virus-induced smooth convoluted membranes (CM) among microtubule paracrystalline arrays (PC) were located in close proximity to virus-induced vesicular replication factories (RF) and proliferated rough ER (Ng and Hong, 1989; Mackenzie, Jones and Young, 1996). Since, then both RFs and CM/PCs have been identified during the replication of many other flaviviruses such as DENV, TBEV, and JEV (Hase, Summers, and Dubois, 1990; Grief et al., 1997; Miorin et al., 2013) (Figure 1-4). 1.3.4 Convoluted Membranes in viral replication CMs are electron dense, often punctate in shape, branching networks of membrane that are highly dynamic and formed following infection by flaviviruses and various coronaviruses (Mackenzie, Jones and Young, 1996; Ulasli et al., 2010; Hagemeijer et al., 2014; Chatel-Chaix et al., 2016; Arakawa and Morita, 2019). Recent studies on DENV have shown that ER proteins such as PDI, Climp-63, Syntaxin17, and calnexin specifically label CMs while markers of the Golgi and endosomes are absent (Steegmaier et al., 2000, Miller et al., 2006; Welsch et al., 2009). These studies have shown that CMs are formed from ER derived membranes. In addition to being ER derived, 3D EM tomography has shown that CMs are continuous with the ER and therefore are not novel organelles but virally induced morphological changes of a pre-existing organelle (Welsch et al., 2009). Functionally, the enrichment of DENV or WNV NS3 proteins in CMs, along with the connection of CMs to other flaviviral-induced ER derived structures involved in replication, suggest a role in flaviviral protein processing and viral protein transport. Yet, as CMs 28 appear to be devoid of ribosomes by EM alternate roles for these virally induced structures are likely to exist (Westaway et al., 1997; Welsch et al., 2009; Chatel-Choix et al., 2016). 1.3.5 Replication factories RFs are vesicular structures that form during replication of most +ssRNA viruses (Mackenzie et al., 1996; Welsch et al., 2009; Cortese et al., 2017). The host membranes utilized to create RFs range from a diverse range of organelles including the ER, mitochondria, peroxisomes, and the trans-Golgi network, however, for members of Flaviviridae, the cellular origin of the RF is the ER (Froshauer, Kartenbeck, and Helenius, 1988; Magliano et al., 1998; Miller, Schwartz, and Ahlquist, 2001; Panavas et al., 2005; Kopek et al., 2007; Gillespie et al., 2010). The size of RFs varies depending on the cell type from 60-100 nm, suggesting cell-type dependent host factors play a part in their formation (Cortese et al., 2017; Liu et al., 2018). The majority of studies on RFs have been conducted using EM and EM tomography allowing for high resolution visualization yet sparse protein immunolabeling of these structures (Mackenzie et al., 1996; Welsch et al., 2009; Cortese et al., 2017). Such studies on DENV and WNV have shown that RFs are ribosome devoid, vesicle-shaped invaginations within the rough ER lumen that are open to the cytoplasm by a pore-like opening (Welsch et al., 2009; Gillespie et al., 2010). The size of this pore is constant, ~11 nm, among different cell lines and flaviviruses suggesting conserved viral and host proteins involved in its formation (Welsch et al., 2009; Cortese et al., 2017). Additionally, by immunolabeling ER proteins such as Climp-63, Calnexin, and PDI are found to localize to RFs more sparsely than to the surrounding rough ER suggesting that while RFs are ER derived, ER luminal proteins are excluded from RFs likely by viral proteins (Welsch et al., 2009). On the other hand, electron dense thread-like structures have been identified within RFs (Ng and Hong, 1989). Further studies 29 elucidated that these thread-like structures consist of a viral RNA genome intermediate product, double stranded RNA (dsRNA) and are largely devoid of the flaviviral +ssRNA genome (Gillespie et al., 2010). Functionally, RFs are believed to be the site of RNA genomic replication. Immunolabeling of dsRNA and various viral proteins involved in genomic replication such as NS3, NS2B, NS5, NS1, and NS4A at RFs further confirms their role in genomic replication (Welsch et al., 2009; Gillespie et al., 2010). By replicating the viral genome in vesicular RFs, it is believed that local enrichment of viral and co-opted host proteins required for genomic replication, together with required metabolites such as ribonucleotides and ATP, can occur leading to highly efficient genomic replication (Paul and Bartenschlager, 2013). In addition to spatial enrichment, by having genomic replication occur on the surface of a membrane each intermediate in genomic replication is subject to two-dimensional diffusion rather than three-dimensional diffusion, as is the case in the cytosol, greatly increasing local reaction efficiency (Monine and Haugh, 2005). During the lifecycle of flaviviruses the RNA genome is utilized for three major yet diverse processes: viral protein translation, genomic replication, and virus particle formation (White, Enjuanes, and Berkhout, 2011) (Figure 1-4). The RNA genome can be translated into the viral polyprotein often due to the possession of an internal ribosome entry site, used as a template for genomic replication, or packaged into immature virus particles. Therefore, conducting genomic replication within RFs isolates this process from that of translation and genomic packaging as all three are conducted within the ER lumen, preventing any interference between these different stages of the viral lifecycle (Paul and Bartenschlager, 2013). When the +ssRNA genome is 30 synthesized from dsRNA in the RF it can then exit through the pore into the cytoplasm and be translated or packaged (Paul and Bartenschlager, 2013). Recent studies have revealed that immature virus particles bud into the ER adjacent to regions of the ER that contain RFs suggesting that newly synthesized RNA genomes can be efficiently delivery and incorporated into the capsid of these immature virus particles before they bud into the ER (Welsch et al., 2009; Cortese et al., 2017). Temporally the utilization of the RNA genome is believed to be regulated to promote successful progression through the different viral life stages, however, the mechanisms which govern this are largely unknown. Lastly, the formation of the membranous RF plays a protective role (Paul and Bartenschlager, 2013). The cellular innate immune response consists of cytoplasmic sensors, Rig1-like receptors, that efficiently recognize foreign 5’ triphosphorylated RNAs and dsRNA in a length-dependent manner (Kato, Takahashi, and Fujita, 2011; Binder et al., 2011) Therefore, by isolating genomic replication to membrane enclosed RFs, the exposure of dsRNA to innate immune surveillance cytoplasmic proteins can be minimized (Paul and Bartenschlager, 2013). Additionally, by forming viral replicase protein and genomic RNA complexes in RFs, cytoplasmic cellular proteases and nucleuses such as endoribonuclease L (RNase L), can be avoided (Whelan et al., 2019). Overall, EM has led to much insight into RF and CM structure, however, the characterization of viral and host proteins involved in the formation of these virally induced structures remain largely incomplete (Aktepe et al., 2017). 31 1.3.6 Hijacked host proteins responsible for membrane re-organization Viruses are obligate, intracellular parasites and therefore, require host metabolites, lipids, and proteins throughout all stages of replication. No better example is the flaviviral re-organization of the ER membrane to create CMs and RFs. One example of a host protein utilized by most flaviviruses for membrane re-organization is fatty acid synthase (FASN) (Aktepe and Mackenzie, 2018). FASN is a cytoplasmic, multi-enzyme protein that catalyses the conversion of acetyl-CoA and malonyl-CoA into the fatty acid palmitate in a nicotinamide adenine dinucleotide phosphate (NADPH)-dependent reaction (Jones and Infante, 2015). Once palmitate is formed it is used as a building block to create the more complex lipids utilized in cellular lipid membranes. WNV and DENV both recruit and localize FASN to the site of RF and CM formation (Heaton et al., 2010; Martin-Acebes et al., 2011). In DENV the interaction of FASN with viral NS3 stimulates FASN activity leading to the increased synthesis of fatty acids and a resultant expansion of the ER membrane (Heaton et al., 2010). In concordance with increased fatty acid synthesis function, ER reticulophagy, used to selectively recycle ER membrane, is also inhibited through the cleavage of the reticulon homology domain of family with sequence similarity 134 (FAM134) by the flaviviral NS3 protease (Lennemann and Coyne, 2017). Overall, by facilitating the synthesis of ER membranes while inhibiting their degradation through NS3, flaviviruses acquire lipids that are likely added onto pre-existing ER membranes during the biosynthesis of RFs and CMs. Besides proteins involved in the regulation of lipid levels and composition, very little is known about host proteins involved in the formation and stabilization of CMs and RFs. The best characterized protein involved in the formation of RFs is RTN3 (Aktepe and Mackenzie, 2018). Originally identified as a necessary protein in the formation of brome mosaic virus and enterovirus 32 71 RFs, RTN3 has since been demonstrated to be involved in flaviviral replication (Aktepe et al., 2017). During WNV infection, the depletion of RTN3.1A results in a disappearance of CMs and RFs, while with DENV, elongation of RFs was observed (Aktepe et al., 2017). Additionally, the depletion of RTN3.1 during ZIKV infection resulted in a decrease in RF number and overall ER membrane curvature (Aktepe et al., 2017). At least part of the effect RTN3.1A depletion has on RFs and WNV CMs comes from its ability to stabilize WNV and DENV NS proteins. Two proteins stabilized by RTN3.1A are NS4A, a viral protein involved in membrane curvature of CMs and RFs, and NS4B, a viral integral membrane protein that likely plays a role in host membrane shaping (Aktepe et al., 2017). Overall, while RTN3.1A is important for the formation of RFs in DENV, WNV, and ZIKV, and CMs in WNV, it is utilized differently by each virus, showing that ER re-organization is induced differently among closely related flaviviruses. Furthermore, studies exploring the roles of the other three RTNs and other well characterized ER-shaping proteins have not been undertaken in terms of RF and CM formation. Besides ER-shaping proteins, ER lumenal chaperone proteins have been determined to be involved in flaviviral RF formation. DnaJC14, a chaperone molecule within the heat shock protein 40 (Hsp40) isoform family of protein chaperones, has been shown to play a role in bringing YFV NS proteins to detergent-resistant membranes. This process leads to the formation of a NS protein derived scaffold that maintains RF vesicular structure (Yi et al., 2012). Whether DnaJC14 plays a role in the formation of RFs in other flaviviruses remains to be seen. Additionally, DNAJB11, another Hsp40 isoform, binds to both positive and negative viral RNA strands within DENV RFs (Taguwa et al., 2015). DNAJB11 is suggested to act in bringing these RNA complexes to Hsp70-stabilized NS5 complexes to promote RNA replication (Taguwa et al., 2015). 33 In addition to viral and host proteins, the formation of RFs has also been shown to require unique compositions of lipids such as ceramide and cholesterol that differ from that of the native host ER membrane (Aktepe, Pham and Mackenzie, 2015). Ceramide has been shown to be enriched within WNV RFs, however, in the case of DENV replication, this lipid plays an inhibitory role (Aktepe, Pham and Mackenzie, 2015). An explanation for the contradictory roles of ceramide in WNV and DENV RFs could be that the two flaviviruses require different ratios of sphingomyelin, a more complex form of ceramide that possesses a phosphorylcholine head group, and negative curvature generating ceramide to maintain RF structure (Goni and Alonso, 2002). Another important lipid in the flavivirus lifecycle is cholesterol (Mackenzie et al., 2007; Osuna-Ramos et al., 2018). By acting as a rigid, space filling lipid element cholesterol is able to maintain membrane fluidity (Smith, 1994). It has been determined that WNV up-regulates cholesterol biosynthesis leading to a redistribution of cholesterol to RFs (Mackenzie et al., 2007). Depletion of cholesterol greatly reduced WNV genomic replication, however, similar studies on DENV failed to show an impact on RFs (Osuna-Ramos et al., 2018). Overall, as seen with host proteins, different flaviviruses utilize lipids for genomic replication differently. Likely, the formation of RFs and CMs require unique regulation of fatty acids, cholesterol and sphingolipids depending on both hijacked host proteins and viral proteins encoded by the specific virus. 34 1.4 Zika virus (ZIKV) manipulation of host ER membranes 1.4.1 Brief history of ZIKV ZIKV is an enveloped, flavivirus that was first identified from the serum of rhesus monkeys in the Zika forest in 1947 (Dick, Kitchen and Haddow, 1952). Comparable to DENV, a genetically close relative, ZIKV particles are ~50 nm in size and consist of a +ssRNA genome containing nucleocapsid within a host membrane-derived lipid bilayer with viral glycoproteins arranged in icosahedral symmetry (Barreto-Vieira et al., 2016, Kostyuchenko et al., 2016; Sirohi et al., 2016). In 1952 the first human cases were identified and following sporadic cases throughout Africa and Asia were classified as largely self-limiting infections (Hayes, 2009). Until recently, ZIKV was considered a largely unknown tropical arbovirus of little importance to public health (Sager et al., 2018). In 2007, an outbreak of the Asian strain of ZIKV on Yap Island and French Polynesia were the first recorded cases of ZIKV outside of Africa and Asia, raising global interest in ZIKV (Duffy et al., 2009). In 2015 an epidemic of the Asian strain of ZIKV was reported throughout South and Central America and the Caribbean (Hennessey, 2016). The association and later causative nature of this epidemic with a major increase in the incidence of congenital microcephaly, the development of an abnormally small head due to damage to developing neural tissue, lead to the World Health Organization to declare ZIKV as a global health concern (WHO, 2015; Mlakar et al., 2016). Currently, the Brazilian strain of ZIKV while largely asymptomatic, has been determined to cause Guillain-Barre syndrome, an autoimmune neurological disorder, meningoencephalitis, transverse myelitis, and potential male infertility due to viral shedding through semen and transient reduction in sperm count (Carteaux et al., 2016; Fauci and Morens, 35 2016; Mecharles et al., 2016; Slavov et al., 2017, Avelino-Silva et al., 2018). Comparative studies between the African and microcephaly causing Brazilian strains of ZIKV have determined a single serine-to-asparagine substitution in the prM protein to be responsible for the neurological virulence of the Brazilian strain (Yuan et al., 2017). When attempting to manufacture a vaccine against ZIKV, recent studies have shown that ZIKV antibodies promote subsequent enhanced dengue infection in dengue virus-naïve individuals confirming that ZIKV antibodies are cross-reactive and can cause DENV antibody-dependent enhancement (Valiant et al., 2018; Valiant and Mattapallil, 2018). Therefore, the possibility of antibody-dependent enhancement due to circulating ZIKV antibodies from a vaccine in areas endemic to both ZIKV and DENV has hindered the development of an effective vaccine against ZIKV glycoproteins (Dejnirattisai et al., 2016; Valiant et al., 2018; Valiant and Mattapallil, 2018). Further, no current FDA approved antivirals against ZIKV exist (Saiz and Martin-Acebes, 2017). 1.4.2 Zika virus life cycle To gain entry into a host cell, ZIKV must bind to one of several cellular glycoproteins to trigger clathrin-dependent endocytosis (Meertens et al., 2017). In neural tissue, it was elucidated that ZIKV requires interaction with the tyrosine kinase receptor Axl and Tyro3AxlMer ligand Gas6 or phosphatidylserine to induce receptor-mediated endocytosis and therefore, enter into the host cell (Hamel et al., 2015; Meertens et al., 2017) (Figure 1-5A). ZIKV can also enter and infect immune cells such as dendritic cells, macrophages and monocytes by binding to the innate immune receptor dendritic cell-specific intercellular adhesion molecule-3-grabbing non-integrin (DC-SIGN) or to a lesser extent phosphatidylserine and T-cell immunoglobulin and mucin domain 1 (TIM-1) (Hamel et al., 2015). Once internalized into the acidic environment of the endosome, the viral E protein 36 undergoes an irreversible conformational change leading to the transition from a dimer to a trimer (Modis et al., 2004) (Figure 1-5A). This transition exposes the highly conserved fusion peptide which integrates into the host cell endosomal membrane and facilitates the fusion of the viral envelope with the endosomal membrane leading to the release of the ZIKV genome into the host cell cytoplasm (Zhang et al., 2017). Once in the cytoplasm the genome is translated on ER localized ribosomes into a polyprotein that is subsequently cleaved by both viral and host cell proteases into 3 structural proteins, being the capsid (C) protein, precursor membrane (prM) protein, and the envelope (E) protein, and 7 non-structural (NS) proteins being NS1, NS2A, NS2B, NS3, NS4A, NS4B, and NS5 (Figure 1-5A,B). ER reorganization then occurs creating RFs where the +ssRNA genome is replicated and utilized for translation or loaded into cytoplasmic capsids to form nucleocapsids. These nucleocapsids then bud into the ER lumen, adjacent to RFs, picking up the envelope which contains both prM and E proteins as was previously reported with DENV (Welsch et al., 2009; Cortese et al., 2017) (Figure 1-5A). The immature virus particles then bud from the ER, together in large multivesicular bodies and travel to the Golgi where cleavage of the prM protein into mature M protein driven by furin, and likely other protease convertases, begins (Sager et al., 2018; Jean lab, unpublished results). Viral maturation is completed as the pr fragment disassociates from the virion leaving the mature, fusogenic M protein upon secretion from the cell (Yu et al., 2009) (Figure 1-5A). 37 Figure 1-5 General ZIKV lifecycle and polyprotein organization. (A) The ZIKV lifecycle begins with an infectious Zika virus particle binding to its host receptor, triggering receptor-mediated endocytosis. Next, upon exposure of the acidic environment of the endosome the viral E glycoprotein undergoes an irreversible conformational change that triggers fusion of the viral envelope with the endosome membrane. The ZIKV RNA genome is released into the cytoplasm and is translated by ER bound ribosomes. ER re-organization occurs, and RNA replication occurs in ER-derived replication factories (not shown here). Newly synthesized viral genomes are released into the cytoplasm and are either translated or incorporated into immature viral particles that bud into the ER lumen. These immature virus particles then proceed through the secretory pathway to the Golgi where furin cleavage of the prM glycoprotein begins. Upon secretion from the cell, the pr fragment disassociates from the viral particle producing a mature, infectious virion. (B) The ZIKV encodes 10 distinct proteins that are translated as one polyprotein and then cleaved by the NS3 protease and host proteins. 3 of these proteins are structural proteins that constitute the virus particle: The C protein, prM protein, and E protein. The other 7 viral proteins are involved A B 38 in viral replication, host cell hijacking, and innate immune evasion and are known as non-structural (NS) proteins. ZIKV NS proteins include NS1, NS2A, NS2B, NS3, NS4A, NS4B, and NS5. (Adapted from Dengue virus life cycle: viral and host factors modulating infectivity by Rodenhuis-Zybert, I.A., Wilschut, J., and Smit, J.M. 2010 © 2010 by Cellular and Molecular Life Sciences) 1.4.3 ZIKV-induced ER re-organization As with DENV and WNV, ZIKV infected cells undergo a massive re-organization of the ER to form both RFs and CMs (Hamel et al., 2015; Hanners et al., 2016; Cortese et al., 2017; Offerdahl et al., 2017; Liu et al., 2018). Vesicular RF structure varies among ZIKV strains as the African MR 766 strain induces ovoid RFs compared to spheroid RFs formed following infection by the BeH819015 Brazilian strain (Liu et al., 2018). Additionally, cell type has an impact on RF size as ZIKV RFs in human neural progenitor cells (hNPCs) are ~20 nm smaller than those observed in human liver Huh7 cell lines when infected with either the African MR 766 or Asian H/PF/2013 ZIKV strains (Cortese et al., 2017; Liu et al., 2018). Similar to DENV and WNV, ZIKV RFs are found within the rough ER lumen of dilated ER sheets and contain electron dense structures believed to be protein and dsRNA replication complexes (Offerdahl et al., 2017; Cortese et al., 2017). The recent use of correlative light and electron microscopy (CLEM) with focus ion beam (FIB)-scanning EM or transmission EM has revealed that the African MR766 and Asian H/PF/2013 strains of ZIKV induce the formation of large CMs with RFs located at the periphery that are located outside of a RF enriched perinuclear region, upon infection of Huh7 cells (Cortese et al., 2017). A series of ZIKV exclusive ultrastructural changes have also been observed such as microtubules crossing CMs and the formation tightly juxtaposed and collapsed ER cisternae with restricted luminal space referred to as zippered ER (Cortese et al., 2017). Additionally, a close relationship between the formation of RFs and viral RNA levels has been noted supporting that 39 RFs are the site of viral genomic replication for ZIKV (Cortese et al., 2017). Similar to other flaviviruses, ZIKV RFs possess the 11 nm pore that connects the lumen of the RF to the cytoplasm (Welsch et al., 2009; Cortese et al., 2017). Besides allowing for the transfer of molecules and proteins required for RNA replication, this pore facilitates the exit and incorporation of newly synthesized viral genomes into nearby nucleocapsids which then bud into the ER lumen to form an immature virus. In contrast to ZIKV infected Huh7 cells, ZIKV infected hNPCs revealed a convoluted tangle of perinuclear rough ER cisternae with RFs clustered inside and mitochondria accumulated at the periphery (Cortese et al., 2017). Interestingly, previously described zippered rough ER and smooth CMs were observed to be absent in ZIKV infected hNPCs (Cortese et al., 2017). Taken together, this suggests that both ZIKV strain and cell line specific host factors largely influence the ZIKV-induced ER morphologies observed. ZIKV ER re-organization also displaces microtubules and intermediate filaments from the perinuclear region (Cortese et al., 2017). While the mechanism is unknown, ZIKV replication has been shown to require cytoskeleton flexibility as the stabilization of microtubules by paclitaxel, an antitumor drug that binds to tubulin enhancing its polymerization into stable microtubules, leads to strongly reduced viral titers (Cortese et al., 2017). 40 1.5 Super resolution microscopy Fluorescence microscopy is an invaluable technique utilized for the visualization of discrete biological processes at the cellular or subcellular level (Lichtman and Conchello, 2005). By specifically labeling molecules of interest with fluorescent proteins or antibodies their localization can be observed in fixed or real time. While this technique is widely applicable to the study of many cellular processes, subcellular molecular interactions or dynamics in the nanoscale level often go undetected due to the 200-250 nm xy resolution limit, known as the Abbe diffraction limit, of fluorescence microscopy (Schermelleh et al., 2019). Therefore, several techniques have been developed to achieve xy and z resolution below that of the Abbe diffraction limit of light known as super resolution microscopy (Gustafsson et al., 1997; Hell and Wichmann, 1994). Super resolution microscopy techniques can be grouped into two distinct categories: super resolution structured illumination microscopy that utilizes the moire effect and mathematical reconstruction to generate super resolution images, or deterministic super resolution microscopy that by use of additional high powered laser beams exploit the modulation or stochastic blinking properties of fluorophore emission to obtain super resolution images (Schermelleh et al., 2019). Super resolution structured illumination microscopy techniques include Structured Illumination Microscopy (SIM) based techniques, while deterministic super resolution microscopy techniques include STimulated Emission Depletion microscopy (STED) and single molecule localization microscopy (SMLM) (Schermelleh et al., 2019). 41 1.5.1 Structured Illumination Microscopy SIM consists of a wide-field fluorescence microscope that possesses a movable diffraction grating (Gustafsson et al., 1999). By having the laser excitation beam travel through a grating, SIM utilizes the moire effect that occurs when two fine patterns are superposed multiplicatively to produce moire fringes (Gustafsson et al., 1997; Gustafsson, 2001). In the case of SIM, the unknown fluorescent dye pattern of the sample is one pattern while structured excitation light emitted from the microscope is the known pattern which allows the user to obtain super resolution information about the sample from the resultant moire fringes (Gustafsson, 2001). To reconstruct a super resolution image by this technique, several images must first be acquired at different orientations of the diffraction grating, both translational and rotational orientation changes, which produce different, known structured illumination patterns. To produce a single 2D SIM image 3 translations and 3 rotations are required for a total of 9 raw images while a single 3D SIM image requires 5 translations and 3 rotations for a total of 15 raw images (Gustafsson et al., 1999; Gustafsson, 2001). By combining these raw images, the resultant increase in resolution is ~100 nm in the xy plane and ~400 in the z plane (Gustafsson et al., 2001). An advantage of SIM over other super resolution techniques is the increase in image acquisition speed and lack of photobleaching due to the lack of additional high-powered lasers (Gustafsson et al., 2001). Therefore, SIM is very informative in live cell applications, especially when imaging highly dynamic structures. Recently, a novel technique known as grazing incidence SIM (GI-SIM) has further optimized SIM as a technique to study cellular dynamics (Li et al., 2015; Nixon-Abell et al., 2016). By the use of counterpropagating light beams located just above the sample one is able to image cellular features at frequencies of up to 40 Hz, a 4 to 10 times increase in acquisition 42 speed relative to the commonly used spinning disk confocal microscopy (Nixon-Abell et al., 2016). 1.5.2 Stimulated Emission Depletion Microscopy STED microscopy utilizes a second, high-powered doughnut shaped depletion laser that possesses a zero-intensity center to supress low intensity fluorescence from the excitation laser point spread function (PSF) (Figure 1-6) (Hell and Wichmann, 1994; Klar et al., 2000). A PSF is the three-dimensional distribution of intensity emitted from a photon source. By de-excitation of low intensity fluorophores only, the high intensity center of the excitation PSF remains allowing for precise excitation of high intensity fluorophores and a resolution of 50 nm in the xy plane and 300 nm in the z plane (Figure 1-6). As a point-scanning form of microscopy, the speed of STED microscopy imaging is restricted to the speed in which it can scan an area and the size of the area (Vicidomini, Bianchini, and Diaspro, 2018). Additionally, as the resolution of STED microscopy depends on the use of a high-powered depletion laser, photobleaching of samples occurs during image acquisition (Vicidomini, Bianchini, and Diaspro, 2018). Therefore, while useful for fixed cell imaging due to the increase in resolution, for use in live cell imaging a small field of ~4.5 µm2 must be imaged in 2D to reach a speed in which cell dynamics can be elucidated (Shao et al., 2011). 43 Figure 1-6 STED microscopy utilizes a secondary doughnut-shaped depletion laser to achieve sub-diffraction limited resolution. (A-B) The xy plane intensity distributions for the excitation PSF and STED depletion beam PSF. (C) De-excitation of the lower intensity fluorescence of the excitation beam results in a sub-diffraction limit effective PSF composed of only high intensity fluorescence. (Adapted from Review of super-resolution fluorescence microscopy for biology by Leung, B.O., and Chou, K.C. 2011 © 2011 by Applied Spectroscopy) 1.5.3 Single Molecule Localization Microscopy Single molecule localization microscopy (SMLM) utilizes stochastic blinking of discrete fluorophores to separate fluorophore labeled molecules temporally and spatially (Endesfelder and Heilemann, 2014). By acquiring many images, each revealing different subpopulations of activated fluorophores, SMLM is able to merge and apply a Gaussian fit to the blinks obtained from single molecules precisely localizing single labeled protein molecules (Endesfelder and Heilemann, 2014). Due to the large number of image stacks and blinks obtained during imaging, Gaussian fitting of the data creates a super resolution image with a xy resolution of ~20 nm and a Z resolution of ~50 nm, theoretically even in structures which contain densely organized proteins (van de Linde, Sauer, and Heilemann, 2008). SMLM consists of several different imaging techniques that all utilize stochastic blinking including direct Stochastic Optical Reconstruction Microscopy (dSTORM), Photoactivation Localization Microscopy (PALM), Ground State 44 Depletion Individual Molecule Return (GSDIM), and more recently, a hybrid of dSTORM and STED known as Miniflux microscopy (Betzig et al., 2006; Rust, Bates, and Zhuang, 2006; Balzarotti et al., 2016). Each of these SMLM techniques acquire 3D data that consists of point clouds, the distribution of points that represent molecules in 3D space, however, the development of computation methods to analyze these distributions have been lacking (Khater et al., 2018). These methods commonly utilize point density, or more recently by Khater et al. network analysis, to cluster and analysis the distribution and interactions of these points among each other (Rubin-Delanchy et al., 2015; Caetano et al., 2015; Khater et al., 2018). 1.5.4 Super resolution microscopy in the study of ER morphology and dynamics The ER is a morphologically complex organelle that ranges from central sheets to peripheral tubules with a diameter of 60-100 nm. In addition to being comprised of morphologically distinct central ER and peripheral ER regions, the tubular network of the peripheral ER is also highly dynamic (Wang et al., 2016). Previously, microscopy techniques were limited by the Abbe diffraction limit of light or the stable “snapshot” nature of EM and therefore, the spatiotemporal resolution to study ER morphology and dynamics were unavailable (Abbe, 1873; Nixon-Abell et al., 2016). The recent advent of super resolution techniques including SIM, and STED microscopy, among others, that break the diffraction limit of light while offering the ability to image rapid cellular dynamics have led to much insight into the ER. 45 The use of super resolution microscopy has drastically altered the field of ER dynamics and morphology (Nixon-Abell et al., 2016; Gao et al., 2019). Previously, ER tubules were known to slide along cytoskeletal elements and to fuse in a homotypic fashion (Du et al., 2004; Liu et al., 2015). The recent use of high-speed GI-SIM has shown that other forms of movement exist such as tubular and three-way junction oscillations, that are driven by more than merely thermal energy (Nixon-Abell et al., 2016). The source and importance of these oscillations on ER function remain to be determined. Additionally, GI-SIM and FIB-SEM led to the observation that previously described peripheral ER sheet-like structures contain spaces less than 50 nm in size and are actually composed of densely clustered tubules that can rapidly assemble and disassemble into isolated tubules (Nixon-Abell et al., 2016) (Figure 1-7). By using a plethora of different super resolution techniques, Nixon-Abell and coworkers (2016) go on to determine that conventional techniques and even several super resolution techniques lack the spatial resolution to resolve the newly termed peripheral ER tubular matrices (Nixon-Abell et al., 2016). The presence of three-way tubular junction forming ATL1 in these structures further confirmed the presence of tubular matrices (Nixon-Abell et al., 2016). Recently, STED microscopy has revealed much about ER morphology that was previously obscured by the diffraction limit of light. Live cell STED microscopy has shown for the first time by light microscopy that peripheral ER sheets exist that contain distinct holes known as nanoholes, in addition to tubular matrices (Schroeder et al., 2019). These nanoholes, predicted to be the fenestrated sheets described by EM tomography, are highly transient and contain RTNs that are involved in the formation of these holes (Schroeder et al., 2019). The dynamic nature of nanoholes has led to the speculation that they function as reservoirs of RTNs that can be utilized to rapidly 46 facilitate the formation of new ER tubules allowing the ER to maintain its many functions under changing cellular conditions. Hence, super resolution microscopy has allowed for characterization of the morphology and dynamics of two ER structures, being peripheral ER sheets with nanoholes and tubular matrices, that under diffraction-limited light microscopy both appear as uniform sheet-like structures (Nixon-Abell et al., 2016; Schroeder et al., 2019). Fixed cell STED microscopy has been utilized to better understand the organization of recently observed periodic peripheral ER tubular nanodomains. These periodic nanodomains consist of ER luminal maxima that associate with ER lumenal proteins and are separate from areas of ER membrane proteins, known as minima (Gao et al., 2019). The manipulation of Climp-63 and RTN4a expression levels alter the length and organization of each of these nanodomains and disrupt the nanodomain segregation of ER proteins (Gao et al., 2019). Through the use of live cell STED, it was further determined that overexpression levels of Climp-63 disrupt while RTN4A overexpression enhance the stability of lumenal ER nanodomains (Gao et al., 2019). Overall, the use of multiple super resolution techniques has allowed for much insight into how the ER is morphological organized at nanoscopic levels and of the dynamics that occur as the ER fulfills its many roles in the cellular environment. 47 Figure 1-7 3D-SIM super resolution microscopy reveals classically described peripheral sheet structures to instead be dense tubular matrices. (A) Left side of panel: Fixed COS-7 cell expressing ER membrane marker Sec61b-GFP, imaged by 3D-SIM. Heatmap represents different position in the z axis. Regions of interest (i, ii, iii) correspond to zoomed images. Scale bar= 10 microns. Right side of panel: Zoomed images displaying diffraction limited epifluorescence images of peripheral ER sheets, which are resolved as tubular matrices by 3D-SIM (Adapted from Increased spatiotemporal resolution reveals highly dynamic dense tubular matrices in the peripheral ER by Nixon-Abell, J., Obara, C., Weigel, A.V., Li, D., Legant, W., Xu, S., Pasolli, A., Harvey, K., Hess, H.F., Betzig, E., Blackstone, C., and Lippincott-Schwartz, J., 2016 © 2016 by Science) 1.5.5 Use of super resolution microscopy to study viruses Several super resolution microscopy techniques have been utilized in the field of virology (Chojnacki et al., 2012; Horsington et al., 2013; Li et al., 2016; Liu et al., 2018; Barlett et al., 2018). Human immunodeficiency virus type 1 (HIV-1) requires maturation of its viral envelope surface glycoprotein, gp120, to facilitate entry into host cells (Rizzuto et al., 1998). Previously, it was unknown how the distribution of HIV-1 gp120 differed between mature and immature virus particles as diffraction limited imaging of the two populations revealed no differences (Chojnacki 48 et al., 2012). The use of 2D fixed cell STED microscopy provided the resolution to determine that the HIV-1 gp120 protein clusters to one focal point upon maturation while the Env protein on immature virus particles remains distributed among multiple foci (Chojnacki et al., 2012). Similar studies using 2D fixed cell STED microscopy and distance analysis have been used to reveal that the viral proteins used in viral genomic replication of the herpes simplex virus type 1 (HSV-1) are spatially separated from host proteins involved in viral protein transcription on the viral genome (Li et al., 2016). Therefore, STED microscopy has also been used to separate two distinct biological processes that occur on the same nanoscale structure. While HIV-1 is the best characterized virus by super resolution microscopy, structural analysis of the organization of other viruses and their glycoproteins throughout the cell has been undertaken for the vaccinia and Nipah viruses by 3D-SIM and, a method of single molecule localization microscopy termed, direct stochastic optical reconstruction microscopy respectively (dSTORM) (Horsington et al., 2013; Liu et al., 2018). For Nipah virus, dSTORM provided the resolution to reveal that several glycoproteins required for viral particle formation stochastically distribute throughout the plasma membrane rather than at the assembly site of the virus (Liu et al., 2018). Besides the use of super resolution microscopy to study virus structural organization and egress, a more recent study has begun to determine the organization of viral proteins involved in genomic replication (Bartlett et al., 2018; Suarez et al., 2019). By utilizing dSTORM, it has been observed that two proteins involved in HCV genomic replication NS5A and NS3 form clusters of different sizes suggesting different organization of the two proteins at RFs (Bartlett et al., 2018). However, the use of super resolution microscopy to study viral replication is still in its infancy (Bartlett et al., 2018; Suarez et al., 2019). 49 1.6 Objective The ultrastructure of ZIKV RFs and closely surrounding ER regions have been extensively characterized by EM and EM tomography, however, much is still unknown regarding the 3D ultrastructure and protein composition of CMs. Additionally, as EM tomography of viral-induced structures have typically only focused on defining RFs, studies on 3D whole cell ER re-organization during ZIKV replication have yet to be conducted. EM provides unrivaled resolution; however, limitations exist when studying the organization and interactions of multiple tagged proteins at once (Humbel et al., 1998). Hence, other studies have turned to diffraction-limited fluorescence microscopy techniques that offer simultaneous labeling of usually up to three different cellular or viral proteins, yet, do not possess the resolution to study protein localization to these nanoscale viral-induced ER morphologies. Consequently, this study looks to utilize the increased resolution of STED microscopy to provide insight into ZIKV-induced ER re-organization and the proteins that are involved in their formation. We hypothesize that 3D STED microscopy will allow for novel characterization of the spatial relationship between different ZIKV-induced ER morphologies and in addition, viral proteins. In this thesis I used STED microscopy to characterize ZIKV-induced ER re-organization. As it has previously been shown that ZIKV-induced ER structures are largely cell and viral strain dependent, I have studied this process in cellular models related to microcephaly. Therefore, I infected the human U87 glioblastoma cell line with the ZIKV strain PRVABC59 (Puerto Rico 2015), a strain of ZIKV closely related to the clinically relevant microcephaly-causing strains in Brazil (Lanciotti et al., 2016). Additionally, ZIKV preferentially infects AXL expressing glial cells 50 and NPCs in the developing human brain, and astrocytes later on in development, to induce microcephaly (Retallack et al., 2016). Therefore, U87 cells, which express high levels of astrocyte marker genes and AXL and have been shown to support strong ZIKV production at 48 hours and robust cytopathic effect at 72 hours were chosen over NPCs, which have been shown to undergo variably levels of apoptosis by several studies (Hanners et al., 2016; Onorati et al., 2016; Retallack et al., 2016). By use of this viral strain and cell line I report a novel ZIKV-induced ER morphology, a perinuclear ZIKV-induced smooth ER tubular matrix that contains RFs. In addition, I have used a computational approach to characterize ZIKV-induced ER re-organization in terms of central ER and peripheral ER domains to examine the 3D organization of ZIKV-induced tubular matrices and replication factories. Lastly, as the viral proteins involved in ZIKV-induced membrane re-organization are largely unknown, I characterize the cellular localization of the ZIKV NS4B protein, a multifunctional integral membrane protein predicted to shape host membranes upon ZIKV infection, to multiple ZIKV-induced ER structures. Overall, I demonstrate that viral infection induces NS4B enriched smooth tubular matrices within the classical rough ER sheet domain of the cell that are utilized as the location of viral genomic replication. 51 Chapter 2: 3D STED super resolution microscopy reveals novel association between replication factories and organized smooth tubular matrices during Zika virus infection 2.1 Introduction The endoplasmic reticulum (ER) is the most morphologically complex organelle in the cell. As one continuous network it spreads from the nucleus to the plasma membrane as either central ER sheets or a peripheral tubular network. In addition, it plays a role in a multitude of cellular processes including protein and lipid synthesis, Ca2+ regulation, protein trafficking, and is known to form membrane contacts with all other cellular organelles (Walter and Ron, 2011; Rowland and Voeltz, 2012; Schwarz and Blower, 2016). Due to the diversity of roles played by the ER in the cell, it is often utilized by pathogens to successfully subvert the cellular immune response and promote replication. Bacteria, such as Legionella pneumophila and Brucella abortus, RNA viruses, and some intracellular protist pathogens, such as Toxoplasma gondii, all utilize the unique functions of the ER to promote their own replication (Roy, Salcedo, and Gorvel, 2006; Romero-Brey and Bartenschlager, 2016; Tsai, English, and Tsolis, 2019). Often ER membranes are re-organized into novel structures to better facilitate intracellular pathogen replication (Mackenzie, Jones and Young, 1996; Romero-Brey and Bartenschlager, 2016). One particularly well-studied group of intracellular pathogens that re-organizes the ER is the viral family of Flaviviridae. This group, which includes members such as the Zika virus (ZIKV), dengue 52 virus (DENV), West Nile virus (WNV), yellow fever virus (YFV), hepatitis C virus (HCV), and the Japanese encephalitis virus (JEV), induce the formation of several different ER re-organized structures to aid in replication (Ng and Hong, 1989; Hase, Summers, and Dubois, 1990; Mackenzie, Jones and Young, 1996; Grief et al., 1997; and Miorin et al., 2013). The two best characterized of these structures are replication factories (RFs) and convoluted membranes (CMs). Extensive studies involving the use of electron microscopy (EM) and EM tomography have described DENV, WNV, and ZIKV RFs to be 60-100 nm spherical vesicular invaginations that form within the rough ER lumen and are connected to the cytoplasm by a conserved, ~11 nm pore (Mackenzie et al., 1996; Welsch et al., 2012; Cortese et al., 2017). These vesicular RF structure vary among both ZIKV strains, as the African MR 766 strain induces ovoid RFs compared to spheroid RFs formed following infection by the BeH819015 Brazilian strain, and cell type as ZIKV RFs in human neural progenitor cells (hNPCs) have been shown to be ~20 nm smaller than those observed in human liver Huh7 cell lines (Cortese et al., 2017; Liu et al., 2018). Yet, by EM immunolabeling of double-stranded RNA (dsRNA), an intermediate in NS5 RNA-dependent RNA polymerase dependent genomic replication, RFs have been determined to be the site of viral RNA genomic replication (Welsch et al., 2009; Gillespie et al., 2010). Functionally, the replication of the viral genome within RFs is suggested to locally enrich viral and host proteins together with required metabolites allowing for more efficient replication and further, for products of genomic replication such as dsRNA and the viral genome to be sequestered away from cytoplasmic innate immune response receptors (Kato, Takahashi, and Fujita, 2008; Binder et al., 2011, Paul and Bartenschlager, 2013). 53 Apart from RFs, studies conducted by EM have also identified virally induced, dense, branching networks of smooth ER tubules known as convoluted membranes (CM) (Steegmaier et al., 2000, Miller et al., 2006; Welsch et al., 2009). Interestingly, the presence of CMs during ZIKV replication vary depending on the cell type being studied as CMs were observed in liver derived human Huh7 cells, however, not in human neural progenitor cells (Cortese et al., 2017). Functionally, the presence of NS3 in CMs suggests that ZIKV protein translation and proprotein cleavage occur here, however, the lack of ribosomes suggests other roles for CMs in replication (Westaway et al., 1997; Welsch et al., 2009; Chatel-Choix et al., 2016). In more thoroughly characterized flaviviruses such as DENV, it has been shown that host factors involved in the cellular innate immune response such as STING and TBK1 are enriched in CMs where they are then cleaved or inhibited by NS3 and NS4B (Aguirre et al., 2012; Yu et al., 2012). While, these structures have been studied by high resolution EM and EM tomography, limitations involving the achievement of dense labeling of intracellular proteins without inducing substantial structural changes to the sample when conducting EM immunolabeling have hindered studies on these viral-induced structures (Humbel et al., 1998; Welsch et al., 2009). Therefore, while the ZIKV membrane proteins NS4A, and NS4B are both predicted to play a role in ER membrane re-organization, precise spatial localization of these proteins to nanoscale RFs or CMs has not previously been shown (Zou et al., 2015; Aktepe et al., 2017). Additionally, in the case of ZIKV, all studies to our knowledge have only utilized EM tomography to study small regions with the goal of obtaining high resolution images of RFs (Gillespie et al., 2010; Romero-Brey and Bartenschlager, 2014). Therefore, much is still unknown regarding the spatial relationship between CMs and RFs throughout the whole cell upon ZIKV infection, and the viral membrane and host 54 ER shaping proteins that are involved in the formation of these structures (Aktepe and Mackenzie, 2018). Studies on the characterization of flaviviral and host protein localization during viral genomic replication have largely been conducted by diffraction limited confocal microscopy, capable of resolving structures larger than ~200-250 nm (Welsch et al., 2009; Aktepe et al., 2017). Therefore, the characterization of proteins localized to RFs and tubules of CMs, shown by EM to have diameters of ~60-100 nm, have been limited. STimulated Emission Depletion (STED) microscopy is a form of super resolution microscopy that utilizes a doughnut-shaped depletion laser to supress low intensity fluorescence from the excitation laser point spread function (PSF) (Hell and Wichmann, 1994; Klar et al., 2000). The resultant xy resolution of STED is ~50 nm while in 3D imaging the z resolution is ~300 nm. The use of STED microscopy to study the ER has previously revealed distinct periodic ER tubule nanodomains and the presence of nanoholes containing ER sheets within the peripheral ER that by conventional microscopy cannot be observed (Schroeder et al., 2018; Gao et al., 2019). Here, application of 3D STED microscopy is used to characterize dense, ZIKV-induced tubular matrices found within a cellular model of ZIKV-induced microcephaly, human U87 glioblastoma cells, and the spatial relationship of these structures with RFs and various ZIKV non-structural proteins. 55 2.2 Results 2.2.1 3D Stimulated Emission Depletion microscopy reveals ZIKV-induced central ER morphological changes in U87 Glioblastoma cells Human U87 glioblastoma cells were stably transfected with either the ER lumenal marker ERmoxGFP, which contains a KDEL retention sequence, or the ER membrane-associated Sec61b-GFP, a non-functional GFP tagged Sec61 protein that labels all domains of the ER, to study the effect on ZIKV infection on ER organization (Shibata et al., 2008; Gao et al., 2019). An initial time course analysis of infection with the ZIKV strain PRVABC59 (Puerto Rico 2015) at a MOI of 1 showed that the largest number of ERmoxGFP and Sec61b-GFP transfected U87 cells displaying viral replication is at 48 hours (Figure A-3). Therefore, all subsequent ZIKV infections were conducted with an infection time of 48 hours. By 3D STED, ER labeling of uninfected cells showed a high intensity central ER region with low intensity peripheral ER tubules (Figure 2-1A) (Shibata et al., 2008; Constantini et al., 2015; Gao et al., 2019). ZIKV infection of ERmoxGFP or Sec61b-GFP transfected U87 cells induced a vast re-organization of the ER into a crescent-shaped, perinuclear region of high intensity ER seemingly on the edge of the central ER with a lower intensity interior central ER region (Figure 2-1A). To determine if this virally re-organized region of the ER is spatially associated with another ZIKV-induced ER structure, RFs, we additionally labeled for dsRNA (Cortese et al., 2017). dsRNA labeling also produced a crescent-shaped pattern of punctate localization (Figure 2-1A). As dsRNA labeling resembled the pattern of high intensity re-organized ER, we utilized the 3D image segmentation software Imaris x64 version 9.2.1 to analyze where in the ER dsRNA was located. Segmentation of the ER and partial overlap 56 colocalization analysis revealed that dsRNA was localized to the crescent-shaped ZIKV-reorganized, perinuclear ER region (Figure 2-1A,B). As suggested by the intensity distribution of ZIKV-infected cells, segmentation analysis revealed the dsRNA containing crescent-shaped ER region to have higher density than the rest of the ER (Figure 2-1A). To determine whether ZIKV induces proliferation of central ER in U87 cells, and to further characterize where in the ER dsRNA and the high intensity ZIKV-induced ER region are located, we looked to utilize Imaris to define central ER and peripheral ER by 3D STED microscopy. By categorizing the peripheral ER regions of mock infected cells as regions of previously described tubular networks, we were able to determine the average max Z-height, 1.23 µm, of the peripheral ER segments obtained from Imaris segmentation (Figure 2-2A). Using this method, any ER segments above 1.23 µm were classified as central ER (Figure 2-2A). In addition, unsegmented frames from the deconvoluted 3D STED images revealed peripheral ER tubules to appear in as many as 6 frames or 1.23 µm, similar to the Imaris derived segments (Figure 2-2A). The resultant Z-height classification of central ER and peripheral ER was applied to both ZIKV and mock infected cells and was determined to provide central ER and peripheral ER masks of average Z-heights of 1.75 µm and 0.8 µm respectively (Figure 2-2B, C). ZIKV infected ERmoxGFP cells were shown to have a smaller total ER volume than mock ERmoxGFP cells and utilizing CER and PER masks, both ZIKV infected ERmoxGFP or Sec61b-GFP transfected cells had a smaller percentage of central ER than that of mock infected cells (Figure 2-3A,B). Additionally, the overall ER density of ZIKV and mock infected cells were not significantly different (Figure 2-3C). 57 Utilizing central ER and peripheral ER masks, we looked to determine which ER domain contained the crescent-shaped dsRNA containing, high-density region of ER. On average, regions of the ER that contain dsRNA make up ~27% of total ER volume (Figure 2-4A, B). Quantification revealed that while 65% of total ER volume is in the CER, only ~22.8% of total ER volume is dsRNA-associated central ER for both ERmoxGFP and Sec61b-GFP transfected U87 cells (Figure 2-4A,B). Additionally, the peripheral ER makes up 35% of total ER volume with 4.3% of total ER volume consisting of dsRNA-associated peripheral ER, again for both ER markers (Figure 2-4A,B). Further quantification showed that ~84.9% of dsRNA-associated ER was located within the central ER (Figure 2-4C). 2.2.2 Zika virus infection induces the formation of a dense ER tubular matrix within the central ER As 3D STED microscopy allows for the opportunity to study cellular components in high resolution 3D space, we next zoomed into the central ER to further determine its structure in regard to the presence of dsRNA. 2D regions of interest (ROI) of central ER that spatially associate with dsRNA show densely packed ER tubules, areas of high ER density, for both ERmoxGFP and Sec61b-GFP transfected U87 cells (Figure2-5A,B). In contrast, 2D ROIs of ZIKV infected central ER regions not associated with dsRNA and the central ER of mock cells both show ER tubules with pronounced spaces between them, leading to lower ER density than what is seen in dsRNA-associated central ER regions (Figure2-5A,B). Again, this was seen for both ERmoxGFP and Sec61b-GFP transfected U87 cells. 3D voxel-based visualization and quantification further show a higher density of ER tubular structures associated with dsRNA than what is seen in central ER regions that lack the presence of dsRNA and mock infected central ER (Figure2-5C,D). 58 As 3D STED microscopy suggested the presence of dense ER tubules in close proximity to dsRNA labeled RFs, we analysed previously obtained high resolution EM images, by members of the Jean Lab, to better characterize this region in the microcephaly relevant cerebral brain organoid model (Qian et al., 2017). Mock infected cells displayed an organized, perinuclear network of stacked rough ER sheets/cisternae (Figure A-1A). On the contrary, ZIKV infected brain organoids reveal the formation of perinuclear, circular region of convoluted smooth ER tubules, similar to CMs, with a less tubule dense center (Figure A-1B). ZIKV-induced vesicles are observed to be clustered within these dense regions of smooth ER tubules. While it has previously been reported that arrays of microtubules are often observed with ZIKV-induced smooth convoluted tubules no cytoskeleton elements were observed here (Cortese et al., 2017). 2.2.3 The ZIKV NS4B and NS2B proteins localize differently within the ER After observing the presence of the crescent-shaped, ZIKV-induced region of convoluted smooth ER tubules, we then undertook to study which ZIKV proteins localize to this structure and RFs. Signal to noise analysis of commercially available ZIKV NS protein antibodies determined that only the ZIKV NS2B and NS4B antibodies give significant signal to noise ratios with STED imaging and therefore were used in this study (Figure A-4). Previously, NS4B, a transmembrane protein with five integral transmembrane domains, has been observed in the CMs of DENV infected Huh7 cells, so we first characterized this protein in terms of ZIKV-induced ER reorganization in U87 cells (Chatel-Chaix et al., 2016). 3D STED labeling of NS4B, along with dsRNA and either ERmoxGFP or Sec61b-GFP, showed NS4B to localize to either ER marker with 59 an apparent increase in labeling intensity in the dense dsRNA associated, crescent-shaped ER region (Figure 2-6A, A-2A,B,C). Quantification of the distribution of NS4B into our described central ER, peripheral ER and dsRNA-associated regions determined that NS4B colocalizes with ~92% of the total ER volume including 98.2% of central ER volume and 79.8% of peripheral ER volume (Figure2-6A,B,C,D). Through comparison to the distribution of dsRNA which was previously determined to localize to 28% of total ER volume, 35.5% of the central ER, and 13.5% of the peripheral ER, NS4B occupies a larger volume of the ER (Figure2-6A,B,C,D). To determine if NS4B was enriched in areas of ZIKV-induced ER reorganization density analysis was undertaken. NS4B density was determined to be higher in dsRNA-associated ER as opposed to central ER and peripheral ER that does not localize to dsRNA (Figure2-6E, A-2C). For further comparison we labeled for NS2B, a ZIKV integral membrane protein which functions as a cofactor for the protease and helicase activity of NS3, which localizes to both CMs and RFs by EM (Welsch et al., 2009; Chatel-Chaix et al., 2016; Cortese et al., 2017). Therefore, labeling of NS2B should be localized more tightly to dsRNA and dense ER regions than what was seen for NS4B. NS2B localized to 41.5% of the total ER volume, largely in the central ER, 53.3% of the central ER compared to 19% of the peripheral ER (Figure2-6A,B,C,D). Additionally, NS2B appeared to be denser in dsRNA-associated ER regions, however, was not found to be significantly denser following statistical analysis (Figure2-6F) NS4B was found to be enriched within the ZIKV re-organized crescent-shaped region of dense ER, however, it is unknown whether ZIKV NS4B localizes with dsRNA. Partial overlap colocalization analysis of 3D STED images determined that NS4B overlaps with 62% of dsRNA 60 puncta while dsRNA overlaps with 11.5% of NS4B puncta (Figure2-7A,B,C). For comparison, analysis of ZIKV NS2B determined that NS2B overlaps with 52.6% of dsRNA puncta while dsRNA overlaps with 24.9% of NS2B puncta (Figure 2-7A,B,C) Therefore, distinct subpopulations of NS4B and NS2B interact with dsRNA. 2.3 Discussion The re-organization of the ER during ZIKV infection has been characterized by the formation of RFs and either rough or smooth convoluted ER membranes by EM (Cortese et al., 2017; Aktepe et al., 2017). Yet, most studies focus on small regions of the cell to characterize RFs and CMs and therefore, the spatial organization of these ZIKV-induced re-organized ER membranes is largely unknown. In this study, 3D STED microscopy reveals the formation of a novel perinuclear, crescent-shaped region of dense tubular ER that encompasses a less dense interior region of ER in both ERmoxGFP, an ER lumen marker, and Sec61b-GFP, an ER membrane marker, transfected U87 glioblastoma cells. As both ER markers are enriched in this dense tubular ER region, it suggests that rather than an enlargement of lumenal ER to facilitate RF formation, a local increase of ER tubules occurs as seen previously (Pena and Harris, 2012; Lenneman and Coyne, 2017; Aktepe et al., 2017). There is no significant difference in total ER density between mock and ZIKV infected cells further suggesting that ER tubules are displaced from the less dense regions of central ER into the region of dense ER tubules hence causing a localized expansion of the ER (Lenneman and Coyne, 2017). It is important to note however that our labeling approach did not allow us to conclude whether an overall volume increase in ER membranes occurred. Additionally, this crescent-shaped dense region of ER was observed to be organized along the edge of the central ER region. Classically, virally induced regions of dense ER membranes correspond to smooth 61 CMs or more recently with ZIKV-induced convoluted rough ER membranes, yet all description of these structures by EM show perinuclear, circular inclusions with no particular organization (Welsch et al., 2009; Chatel-Chaix et al., 2016; Cortese et al., 2017). Therefore, we identify the first, to our knowledge, organized region of virally induced, dense ER membranes that encompasses a region of lower ER density. In addition to characterizing the 3D, cellular organization of regions of virally-induced dense ER, we looked to determine where RFs localize and what, if any, their spatial association with the described dense ER region is by 3D STED microscopy. Punctate active RFs, viewed by dsRNA labeling, displayed a crescent-shaped distribution throughout the cell that was spatially associated with dense ER. The organization of ZIKV RFs within dense ER tubules observed here has also been seen in hNPCs, where ZIKV RFs were often clustered within convoluted rough ER membranes (Cortese et al., 2017) It is of interest to note that this morphology has not been observed in non-neuronal cell types. Therefore, ZIKV re-organizes ER membranes similarly between U87 cells and hNPCs yet differences the organization of these dense ER regions suggest further complexity in the utilization of cell specific host factors. Next, by utilizing Z-height of the ER we were able to classify RFs as largely central ER specific, in concordance with numerous previous EM studies (Mackenzie et al., 1996; Welsch et al., 2009; Gillespie et al., 2010; Cortese et al., 2017). Interestingly, areas of the ER that contain RFs comprise of only a small volume of the central ER further suggesting that ER re-organization occurs within a unique subdomain of the central ER. Whether the dense ER region or RFs form first and the role each of these viral-induced re-organized structures have in the formation of the other is still 62 unknown. Additionally, the determination of a unique subdomain of central ER that contains RFs suggests that this region possesses unique combinations of host factors or concentrations thereof that are required for ER re-organization. As RTNs play a major role in ER membrane curvature of both ER tubules and sheet edges and have been shown to induce ER sheet expansion following depletion, it is likely these proteins play a role in ZIKV-induced ER reorganization. Indeed, the role of RTN3.1A has been determined in WNV CM and RF formation while in terms of ZIKV infection the depletion of RTN3.1A leads to a decrease in the number of RFs and overall membrane curvature in HEK293T kidney cells (Aktepe et al., 2017). No impact on CM formation was noted in that study (Aktepe et al., 2017). Therefore, further studies into the role of ER shaping proteins such as other RTN proteins, DP1, and rough ER/central ER proteins such as CLIMP-63 in ZIKV ER re-organization must be undertaken. High-resolution EM images of cerebral brain organoids, obtained previously by members of the Jean Lab, revealed the formation of a region of closely packed, perinuclear smooth convoluted tubules that contain ZIKV-induced vesicles and surround a less membrane dense interior region, similar to what was suggested by 3D STED microscopy. Therefore, although cerebral brain organoids are a model of the developing fetal brain that are composed of a heterogenous population of cell types, this EM data appears to be in agreement with what was determined by 3D STED of U87 glioblastoma cells. Combining the results of the two different forms of microscopy it is interesting to note the presence of closely packed ER tubules within the normally ER sheet dominated central ER. Studies on the morphologically similar drug-induced perinuclear smooth ER convoluted membranes suggest that these smooth ER membranes are derived from the peripheral ER, possibly localized to central ER regions by cytoskeletal changes (Sprocati et al., 63 2006). The same cellular mechanism could be responsible for ZIKV-induced ER morphologies as re-organization of intermediate filaments and ER-interacting microtubules into a birdcage-like structure that encompasses dense RF containing ER regions, and is necessary for genomic replication, has been shown previously (Cortese et al., 2017). Re-organization of microtubules may act as a scaffold which brings and stabilizes dense regions of ER tubules into the central ER. While convoluted membranes have been observed during the replication of other flaviviruses these structures have yet to be defined in terms of 3D ER morphology. Through observation of the crescent-shaped, densely packed ER tubules by STED and EM imaging, we suggest that the morphology of these structures resembles that of the recently described sub-diffraction limited ER tubular matrices, closely packed ER tubules first described in the peripheral ER (Nixon-Abell et al., 2016). Functionally, the formation of these structures is suggested to stabilize high curvature regions of the ER while acting as a reservoir for excess membrane proteins and lipids due to possessing increased membrane surface area (Nixon-Abell et al., 2016). In this study and others, CMs have been shown to be enriched with viral NS proteins and various host proteins involved in innate immune response, therefore acting as a protein reservoir. Additionally, as the formation of ZIKV RFs require the stabilization of high curvature membranes and an increase in ER membrane surface area, as it is this membrane that forms the interior of the RF, along with an enrichment of viral and host membrane proteins involved in genomic replication, it would therefore be favourable to induce the formation of ER tubular matrices. Besides functional similarity, the dynamics of tubular matrices and CMs in the related DENV share resemblance, rapidly growing and shrinking in size (Chatel-Chaix et al., 2016; Nixon-Abell et al., 2016). This further suggests that virally induced rough or smooth CMs are a variant of peripheral ER tubular matrices (Chatel-Chaix et al., 64 2016; Nixon-Abell et al., 2016). Therefore, further studies should focus on determining whether ER proteins involved in the formation of tubular matrices, such as three-way tubular junction forming atlastins, localize to regions of ZIKV-induced dense ER tubules. Interestingly, the only previous study that has characterized the formation of convoluted, ZIKV RF-containing ER membranes in brain derived tissue, hNPCs, observed this region to be composed of rough ER while we show this region to be smooth ER convoluted tubules (Cortese et al., 2017). This is surprising as the presence of RFs within rough ER membranes is believed to facilitate transfer of viral and host proteins utilized for genomic replication to RFs from nearby ribosomes (Gillespie et al., 2010). Additionally, RFs have been described to be found in rough ER in numerous other flavivirus including DENV, WNV, and JEV (Leary and Blair, 1980; Welsch et al., 2009; Gillespie et al., 2010). Considering that the bulk of enzymes associated with lipid synthesis are located within the smooth ER, it is likely that RF formation is enhanced by occurring in smooth ER (Heaton et al., 2010; Martin-Acebes, 2011). Additionally, as smooth CMs in other viruses and cell types are known to be enriched with viral proteins involved in genomic replication such as NS3, NS2B or NS4B, as seen in this thesis, the formation of RFs within close proximity to ribosomes might not be as important as previously believed (Welsch et al., 2009; Gillespie et al., 2010; Chatel-Chaix et al., 2016). As the ER is a continuous organelle in which RFs, the rough ER, and smooth CMs are in direct lumenal contact proteins from the rough ER can travel to the dense smooth ER and RFs facilitating genomic replication without being spatially linked. Additionally, previous studies on DENV reveal RFs within rough ER to encompass smooth CMs showing that a close spatial relationship, that until now was not seen with ZIKV, can exist (Welsch et al., 2009). 65 Interestingly, another proposed functional role of CMs is based on the presence of enriched DENV or WNV NS3 protein, a helicase and protease involved in the cleavage of the flaviviral proprotein and different host proteins along with cleaved STING and other antiviral proteins in these structures (Westaway et al., 1997; Aguirre et al., 2012; Yu et al., 2012). Therefore, CMs have been suggested to play a role in the subversion of the cellular innate immune response (Aguirre et al., 2012; Yu et al., 2012). As our results show RFs to be organized within these structures further studies exploring whether cleaved antiviral proteins are located here and whether cleavage of these proteins occurs before the production of dsRNA and genomic RNA would be of interest. Again, ZIKV-induced ER-organization was shown to vary even among closely related cell lineages, cerebral brain organoids and hNPCs, with the presence of RFs within smooth convoluted membranes representing another unique virally induced ER morphology (Cortese et al., 2017). All of the flaviviral non-structural (NS) proteins are believed to play a role in the formation of the genomic replication machinery and RFs. However, studies to confirm the localization of these proteins under ZIKV infection are largely absent other than by diffraction limited confocal microscopy that does not possess the resolution to observe distinct RFs. Such studies have identified ZIKV NS3 and NS4A to localize to both CMs and RFs (Aktepe et al., 2017, Cortese et al., 2017). Here we determine by 3D STED microscopy that ZIKV NS4B is localized throughout the ER and enriched within the novel crescent-shaped, dense ER region that contains dsRNA, as opposed to the largely central ER localization of the NS3 cofactor, NS2B. Comparisons with dsRNA and NS2B labeling reveal that while NS4B occupies a much larger area of the ER, only a subpopulation of NS4B colocalizes with dsRNA. Furthermore, this subpopulation of NS4B 66 comprises of a smaller percentage of total NS4B labeling than that of dsRNA colocalized NS2B. While NS2B has largely been shown to function as a cofactor for NS3 protease activity, NS4B has been shown to be a multifunctional, membrane protein involved in many processes of ZIKV infection and pathogenesis. Such functions of NS4B include the inhibition of the Akt-mTOR pathway leading to defective neurogenesis, the inhibition of the interferon pathway by interacting with TANK-binding kinase 1, upregulation of the ATF6/IRE-1 pathways during DENV infection, likely to cause further proliferation of ER membranes and aid in the folding of viral proteins, and has been suggested to play a role in the induction of curvature in ER membranes to produce RFs (Zmurko, Neyts, and Dallmeier, 2015; Chatel-Chaix et al., 2016; Liang et al., 2016). In addition, flaviviral NS4B serves as a mutational hotspot where small changes in its amino acid sequence lead to large changes in replication capacity and pathogenesis suggesting that this protein can acquire additional functions after several rounds of infection cycles (Zmurko, Neyts, and Dallmeier, 2015). Similarly, our quantification of NS4B colocalization with dsRNA also suggests a multifunctional role of NS4B in ZIKV infection, as NS4B is found at several ER regions associated with different functions: the dsRNA associated region, the dsRNA devoid central ER, and the peripheral ER. Previous studies on DENV have highlighted the localization of NS4B to NS2B/NS3 rich CMs (Chatel-Chaix et al., 2016). It has been shown that by inactivation of DRP1, a mitochondrial fission factor, NS4B is able to induce mitochondria elongation that is required for CM formation (Chatel-Chaix et al., 2016). Further, NS4B-induced mitochondria elongation plays a functional role in the increased disruption of MAMs allowing the virus to successfully subvert the host innate immune response (Chatel-Chaix et al., 2016). Whether NS4B plays a similar role in the formation of the crescent-shaped, dense ER region we describe remains unclear. Alternatively, only 50-60% of dsRNA, representing sites of active genomic replication, 67 colocalized with NS4B and NS2B suggesting that these proteins do not localize at RFs during all stages of active replication. Given that NS4B is a multifunctional protein that localizes to functionally distinct regions of the ER, the determination of whether the colocalization of NS4B to RFs and specific regions of the ER change during the course of infection must be undertaken by future super resolution live cell or time course studies. Overall, this study utilizes 3D STED microscopy together with computationally derived central ER and peripheral ER masks to introduce and characterize a novel ZIKV-induced ER morphology: organized, RF-containing, crescent-shaped central ER tubular matrices in U87 cells. This region was further determined to be enriched in both ZIKV NS4B, similar to what has been observed in RF devoid CMs induced by DENV infection (Chatel-Chaix et al., 2016). Additionally, electron microscopy of related ZIKV-infected cerebral brain organoids revealed RF-containing, smooth CMs in agreement with the ZIKV-induced tubular matrices described by STED. Taken together, these results suggest that CMs in U87 cells and cerebral brain organoids are organized tubular matrices that contain RFs. Further studies into the host proteins hijacked to create these novel viral-induced ER morphologies and whether their formation plays a role in RF formation remain to be elucidated. In conclusion, a novel ZIKV-induced morphology is characterized adding to the diverse array of ER derived structures produced during flaviviral infection. 68 69 Figure 2-1 3D STED microscopy reveals the presence of ZIKV-induced ER re-organization in U87 Glioblastoma cells. (A) ERmoxGFP or Sec61β-GFP stable transfected U87 cells were mock or infected for 48 hours with the PRVABC59 2015 ZIKV strain (MOI= 1) prior to 0.2 % Glutaraldehyde + 3% PFA fixation. Either the lumenal or membrane domains of the ER were visualized through expression of ERmoxGFP or Sec61β-GFP respectively (white). ZIKV replication factories were visualized by immunostaining for dsRNA (white). Cells were then imaged by 3D STED microscopy utilizing a step size of 200 nm. Mock-infected cells serve as the control. Fluorescence intensity of the ER of both infected and mock cells is depicted using a spectrum heat map alongside 3D STED maximum projections both generated on Imaris x64 9.2.1 (Imaris). A segmentation mask of the ER that colocalizes with dsRNA was also generated using Imaris (grey). Yellow squares in the panels indicate the magnified ROIs shown in the corresponding panels below. Scale bar= 10 microns. (B) Quantification of the mean normalized ER density ((Intensity sum of mask/total cell intensity sum)/ (volume sum of mask/ total cell volume sum)) by Imaris segmentation. 5 cells per biological replicate were analyzed for a total of N=3. Statistics were done using unpaired Student’s T tests where **= P<0.01, Error bars represent SEM. 70 71 Figure 2-2 Defining the central and peripheral ER of ZIKV and mock infected cells by Imaris segmentation to better characterize ZIKV-induced ER re-organization. (A) Mock infected ERmoxGFP stable transfected U87 cells were imaged by 3D STED microscopy with a step size of 200 nm. Yellow boxes indicate magnified ROIs distinguished by red Roman numerals. Yellow arrows mark persistent ER tubules in the peripheral ER and ER sheets in the central/nuclear ER regions. The maximum Z height of peripheral ER Imaris segmentation derived objects of each cell, either ERmoxGFP or Sec61β-GFP stably transfected, is indicated by a dot in the graph (B) ER labeling from 48 hour mock or ZIKV infected ERmox-GFP or Sec61β-GFP stable transfected U87 cells (MOI= 1) was segmented by Imaris and height in the Z dimension was visualized by a Z height spectrum heat map. Central (green) and peripheral ER (red) regions were separated by Z height (central ER> 1.26 microns and peripheral ER< 1.26 microns) and were used to create masks for further analysis. (C) Quantification of average Z bounding box length between the central and peripheral ER masks. 5 cells per biological replicate were analyzed for a total of N=3. Statistics were done using unpaired Student’s T tests where ***= P<0.001. Error bars represent SEM. Scale bar= 10 microns. 72 Figure 2-3 Central ER (CER) and Peripheral ER (PER) masks reveal changes in ER domain volume. (A) Total ER volume between mock and ZIKV infected ERmoxGFP or Sec61β-GFP stably transfected cells. (B) Volume percentage of CER and PER masks between mock and ZIKV infected cells (C)Quantification of CER and PER mean normalized density between mock and ZIKV infected cells. 5 cells per biological replicate were analyzed for a total of N=3. Statistics were done using ANOVA with post-hoc Tukey HSD (A, B) and unpaired Student’s T tests (C), where NS= nonsignificant, *= P<0.05; and **= P<0.01. Error bars represent SEM. 73 Figure 2-4 The majority of dsRNA localizes to a central ER subdomain. (A) Representative Imaris central (green) and peripheral (red) ER masks from 48-hour infected ERmoxGFP and Sec61b-GFP stable transfected U87 cells are overlaid with the dsRNA-associated CER or PER mask (white). Scale bar= 10 microns. (B) Quantification of the average volume percentage of dsRNA-associated ER, CER, dsRNA-associated CER, PER, and dsRNA-associated PER for either ER marker. (C) Quantification of the average volume percentage of dsRNA that resides in the CER mask. 5 cells per biological replicate were analyzed for a total of N=3. Statistics were done using repeated measures ANOVA with post-hoc Tukey HSD where **= P<0.01, and ***= P<0.001. Error bars represent SEM. 74 75 Figure 2-5 The ZIKV-induced perinuclear region of dense ER is a network of closely packed, dsRNA labeled tubular ER that encompasses a region of lower ER density. (A) 3D STED images of 48-hour mock or ZIKV infected ERmoxGFP or Sec61β-GFP stable transfected U87 cells (MOI= 1) immunostained for dsRNA are shown here as Imaris generated maximum projections. Red (dsRNA associated CER), yellow (dsRNA devoid CER), and green (mock CER) boxes indicate ROIs used on the corresponding panel of figure 5B. Scale bar= 10 microns. (B) 2D 3-micron by 3-micron panels displaying dsRNA (red), ER (white), and merge images from each ROI. Scale bar= 500 nm. (C) Imaris 3D surface rendering of the ER (grey) of 1-micron by 1-micron by 1-micron regions of the above ROIs. (D) Quantification of mean normalized ER density for each of the three distinct CER regions by Imaris segmentation and masking. All quantification involves the analysis of ~5 cells per biological replicate for a total of N=3. Statistics were done using ANOVA with post-hoc Tukey HSD where ***= P<0.001. Error bars represent SEM. 76 77 Figure 2-6 Labeling of the ZIKV NS4B and NS2B proteins reveal that both proteins localize to the central ER while NS4B also displays peripheral ER localization with an enrichment in the crescent-shaped dense ER region. (A) 48-hour mock or ZIKV infected ERmoxGFP (red) stable transfected U87 cells (MOI= 1) were immunostained for ZIKV NS4B or NS2B (green) and dsRNA (white) and imaged by 3D STED. The distribution of NS4B or NS2B in the CER, PER or dsRNA associated ER (white) was overlaid with NS4B or NS2B labeling to highlight the presence of NS4B or NS2B in these three ER regions. All images shown here are Imaris maximum projections. Scale bar= 10 microns (B) Quantification comparing the percentage of total ER volume that NS4B, NS2B and dsRNA occupy. (C) Quantification comparing the percentage of central ER volume that NS4B, NS2B and dsRNA occupy. (D) Quantification comparing the percentage of peripheral ER volume that NS4B, NS2B and dsRNA occupy. (E) Quantification of mean normalized NS4B density in dsRNA-associated ER, dsRNA-devoid CER, and dsRNA-devoid PER by Imaris segmentation and masking. (F) Quantification of mean normalized NS2B density in dsRNA-associated ER, dsRNA-devoid CER, and dsRNA-devoid PER by Imaris segmentation and masking. 5 cells per biological replicate were analyzed for a total of N=3. Statistics were done using ANOVA with post-hoc Tukey HSD (B,C,D) and repeated measures ANOVA with post-hoc Tukey HSD (E,F), where NS= nonsignificant, *=P<0.05, **=P<0.01, and ***=P<0.001. Error bars represent SEM. 78 Figure 2-7 dsRNA colocalizes with distinct subpopulations of NS4B and NS2B in the crescent-shaped dense, tubular ER region. (A) Mock or 48-hour ZIKV infected ERmoxGFP stable transfected U87 cells (MOI= 1) were immunostained for ZIKV NS4B or NS2B (green) and dsRNA (red) and imaged by 3D STED. Images shown are Imaris maximum projections. White squares represent the zoom ROIs used to depict colocalization. White arrowheads represent spots of colocalization (B) Quantification of dsRNA that are partially overlapping with NS4B or NS2B by Imaris segmentation. (C) Quantification of NS4B or NS2B that are partially overlapping with dsRNA by Imaris segmentation. Each dot represents a cell for both ER markers. ~5 cells per biological replicate were analyzed for a total of N=3. Scale bar = 10 microns. ROI scale bar = 2 microns. Statistics were done using unpaired Student’s T tests where NS= nonsignificant, and ***= P<0.001. Error bars represent SEM. 79 2.4 Materials and methods 2.4.1 Antibodies, plasmids, chemicals, and viruses ERmoxGFP was a gift from Dr. Erik Snapp (Albert Einstein College of Medicine, present Howard Hughes Medical Institute Janelia Research Campus, Virginia) (Addgene plasmid # 68072), and Sec61b-GFP from Dr. Gia Voeltz (University of Colorado, Boulder). Mouse anti-dsRNA antibody (1:500) was purchased from Scicons English and Scientific Consulting Kft. (Cat#: J2-1904), Rabbit anti-NS4B (1:500) from Genetex (Cat#: GTX133321), Rabbit anti-NS2B (1:500) from Genetex (Cat#: GTX133308), Goat anti-rabbit Alexa Fluor 532 (1:300) from ThermoFisher Scientific (Cat#: A-11009), Goat anti-mouse Alexa Fluor 568 (1:300) from ThermoFisher Scientific (Cat#: A-11031), and goat serum from Thermo Fisher Scientific (Cat#: 16210-064). 16% paraformaldehyde (Cat#: 15710) and 25% glutaraldehyde (Cat#: 16220) were from Electron Microscopy Sciences, USA. All other chemicals were obtained from Sigma. Zika virus strain (PRVABC59), a microcephaly causing Asian strain isolated from Puerto Rico in 2015, was obtained from ATCC (Cat#: VR-1843). 2.4.2 Cell culture and stable cell line generation U87 MG cells from ATCC (Cat#: HTB-14) were grown at 37°C with 5% CO2 in Gibco Minimum Essential Media (MEM) (Cat#: 11095080) containing 10% FBS, 1% MEM non-essential amino acids (Cat#: 11140050), and 1% sodium pyruvate (Cat#: 11360070) (all from ThermoFisher Scientific) unless otherwise stated. Plasmids were transfected in U87 cells with Effectene (Qiagen, 80 Germany) according to the manufacturer’s protocols. Following 24 hours of plasmid transfection cells were incubated in MEM with 800 µg/mL of Geneticin (Cat#: 10131027). Media was changed every 24 hours for a week and then every 2 days. Once colonies grew to confluency the resultant GFP positive cells were sorted by FACS. Plasmid stable lines were maintained in MEM with 400 µg/mL of Geneticin where media was changed every two days. 2.4.3 Infection and Immunofluorescence labeling Cells grown on #1.5H coverslips (Paul Marienfeld, Germany) for 24 hours were washed 1X with PBS and incubated with Zika virus, diluted in MEM, at a MOI of 1 for 48 hours. Then cells were: 1) fixed with 3% paraformaldehyde with 0.2% glutaraldehyde at room temperature for 15 minutes and washed with PBS-CM (phosphate buffer solution supplemented with 1 mM CaCl2 and 10 mM MgCl2) (all PBS-CM washes consist of two quick washes and then two 5 minute washes unless further specified); 2) permeabilized with 0.2% Triton X-100 for 5 minutes followed by PBS-CM washes; 3) quenched of glutaraldehyde aldehydes with 1mg/mL of NaBH4 for 10 minutes and washed with PBS-CM; 4) blocked with 10% Goat serum (ThermoFisher Scientific) and 1% bovine serum albumin (Sigma) in PBS-CM for 1 hour; 5) incubated with primary antibodies in the same blocking solution as described above for 1 hour at room temperature (RT) then washed quickly with PBC-CM once followed by three 5 minute washes with Antibody Wash Buffer (20x SSC, 0.05% Triton-X100 in Milli-Q H2O); 6) incubated with secondary antibodies in Antibody Buffer (1% BSA, 2% goat serum, 0.05% Triton-X100, 20X sodium/sodium citrate buffer in Milli-Q H2O) for 1 hour at RT then washed quickly with PBS-CM once followed by six 10 minute washes with Antibody Wash Buffer on a rocker; 7) rinsed with Milli-Q H2O and mounted with ProLong Diamond (ThermoFisher Scientific) and cured for 24-48 hours at RT. 81 2.4.4 3D STED microscopy 3D STED imaging was performed with the 100X/1.4 Oil HC PL APO CS2 objective of a Leica TCS SP8 3X STED microscope (Leica, Germany) using white light laser excitation with HyD detectors, and Leica Application Suite X (LAS X) software. Sample acquisition was done at a scan speed of 600Hz with a line average of 6. Time-gated fluorescence detection was used to further improve lateral resolution. GFP was excited at 488 nm and depleted using the 592 nm depletion laser. Alexa Fluor 532 was excited at 528 nm, and Alexa Fluor 568 was excited at 577 nm. Both were depleted using the 660 nm depletion laser. Each channel was acquired sequentially, in the order of Alexa Fluor 568/ Alexa Fluor 532/GFP, where all the frames of one channel were imaged before moving on to the next channel to avoid crosstalk. A step size of 210 nm was utilized for 3D STED. Images were then deconvoluted using Huygens Professional software (Scientific Volume Imaging). 2.4.5 Transmission electron microscopy 18 and 63-day old cerebral organoids were infected with the PRVABC59 Zika virus strain, MOI of 1, for 48 hours. Organoids were then processed by replacing media with the primary fixative [0.1 M sodium cacodylate), 1.5% paraformaldehyde, 1.5% glutaraldehyde, pH 7.3, room temperature] (Lyon, et al. 2017) and allowed to sit for 1 hour before placing them into a fridge (4oC) overnight. The organoids were washed three times (10 minutes each wash) with buffer (0.1M sodium cacodylate) and then post-fixed for 1 hour on ice in 0.1M osmium tetroxide in 0.1M sodium cacodylate (pH7.3). They were then washed three times with distilled water (10 min each wash, room temperature) and then stained en bloc with 1% aqueous uranyl acetate for 1 hour at 82 room temperature. They were washed again for three times with distilled water and then dehydrated through an ascending concentration series of ethyl alcohol (30%, 50%, 70%, 95%, 100%), passed through two changes of propylene oxide, and then infiltrated overnight in a 1:1 solution of propylene oxide:EMBED 812 resin. The organoids were then passed through two changes of 100% EMBED resin, embedded and the resin polymerized at 60°C for 48 hours. Thick sections (0.8-1.0 micron) were cut using newly prepared glass knives mounted on a Leica Ultracut ultramicrotome, collected on glass slides, and then stained with toluidine blue. Cover slips were fixed to the slides using EMBED resin and the sections viewed and images collected using a Ziess Imager A1 photomicroscope. Thin sections were cut using a diamond knife and sections were collected on copper grids. The sections were then stained with uranyl acetate and lead citrate, and images collected using a FEI tecnai G2 Spirit transmission electron microscope. 2.4.6 Quantification and statistical analysis 3D STED ER segmentation and ER region mask generation were done using Imaris x64 9.2.1. dsRNA associated ER and NS4B/NS2B associated ER regions were generated by partial overlap analysis after surface generation. 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Mock cells also exhibit isolated mitochondria (Mito). Zoomed images highlight rough ER (rER) in mock cells and tubular matrices with Ve in infected cells. Red arrows mark tubular matrices and Ve. Scale bars for overview images are 500 nm and 100 nm for zoomed image ROIs. 103 Figure A-2 Labeling of the ZIKV NS4B protein reveals both central and peripheral Sec61β-GFP ER localization with an enrichment in the dense ER region. (A) 48-hour mock or ZIKV infected Sec61β-GFP (red) stable transfected U87 cells (MOI= 1) were immunostained for ZIKV NS4B (green) and dsRNA (white) and imaged by 3D STED. The distribution of NS4B in the CER, PER or dsRNA associated ER (white) was overlaid with NS4B labeling to highlight the presence of NS4B in these three ER regions. All images shown here are Imaris maximum projections. Scale bar= 10 microns (B) Quantification comparing the percentage of total ER volume that NS4B and dsRNA occupy in the total ER, CER and PER for both ER markers. (C) Quantification of mean normalized NS4B density in dsRNA-associated ER, dsRNA-devoid CER, and dsRNA-devoid PER by Imaris segmentation and masking. 5 cells per biological replicate were analyzed for a total of N=3. Statistics were done using repeated measures ANOVA with post-hoc Tukey HSD where *= P<0.05, **=P<0.01, and ***= P<0.001. Error bars represent SEM. 104 Figure A-3 ZIKV infected cells displaying dsRNA is largest at 48 hours. ZIKV infected ERmoxGFP, Sec61b-GFP transfected and non-transfected U87 cells were fixed and immunostained for dsRNA at 12, 24, or 48 hours. Cells containing dsRNA puncta were counted and the percentage of cells with dsRNA was calculated. ~ 100 cells per biological replicate were analyzed for a total of N=3. Statistics were done using unpaired Student’s T tests where *= P<0.05. Error bars represent SEM. 105 Figure A-4 Both ZIKV NS2B and NS4B antibodies produce significant mean fluorescence intensity in ZIKV infected cells at concentrations of 1:500. (A and B) 48-hour mock or ZIKV infected ERmoxGFP transfected U87 cells were labeled for either Genetex ZIKV NS2B or NS4B antibodies at concentrations of 1:100, 1:300, and 1:500. Cells were imaged by 2D STED microscopy, ROIs were drawn around cells and fluorescence intensity was obtained. 5 cells per biological replicate were analyzed for a total of N=1. Statistics were done using unpaired Student’s T tests where *= P<0.05, **= P<0.01. Error bars represent SEM. A B
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Cell and Developmental Biology
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3D Stimulated Emission Depletion super resolution microscopy reveals novel association between replication factories and organized smooth tubular matrices during Zika virus infection
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