Medicine, Faculty of
DSpace
UBCV
Zou, Yuanjie
Doctor of Philosophy - PhD
University of British Columbia
Diabetes affects more than 500 million people worldwide and results in serious health complications that contribute to millions of deaths annually. Diabetes is characterized by high blood glucose levels resulting from insufficient insulin production due to dysfunction and loss of insulin-producing pancreatic β-cells. The success of islet transplantation as a functional cure for autoimmune type 1 diabetes is also hindered by dysfunction and death of transplanted β-cells. Autophagy is a conserved cellular process that helps maintain cell function and survival by lysosomal degradation and recycling of damaged cellular components under stress. However, the role of autophagy in β-cell survival under hypoxia and following islet transplantation remains unclear. Furthermore, recent evidence suggests that failure of β-cell autophagy may contribute to β-cell dysfunction and the development of both type 1 and type 2 diabetes, but the underlying mechanisms are not understood. This dissertation investigates the importance of autophagy and lysosomal homeostasis in pancreatic β-cells under diabetes- and transplantation-relevant stress. We created a mouse model with β-cell-specific deletion of the autophagy-related gene Atg5 and demonstrated that basal autophagy is essential for maintaining glucose tolerance and β-cell survival. We further found that autophagy protects β-cells under hypoxic stress and during islet transplantation, as loss of autophagy exacerbated β-cell death and islet graft failure. Prolonged hypoxia was found to impair autophagic flux, and this was associated with lysosomal dysfunction and decreased β-cell levels of Tfeb, a transcriptional master regulator of lysosomes and autophagy. By pharmacologically activating Tfeb, we were able to restore lysosomal function and improve β-cell viability under stress. To further understand the role of Tfeb in β-cells, we generated a β-cell-specific Tfeb knockout mouse. Loss of β-cell Tfeb made the mice more susceptible to streptozotocin-induced hyperglycemia but did not worsen glucose tolerance under high-fat diet-induced stress, possibly due to compensation by the related transcription factor Tfe3. The research in this dissertation highlights the protective role of autophagy in β-cells under stress and identifies Tfeb dysregulation as a factor in pathogenic β-cell autophagy loss. Targeting Tfeb may offer a therapeutic strategy to improve clinical islet transplantation outcomes and enhance β-cell resilience in diabetes.
ROLES OF AUTOPHAGY AND TRANSCRIPTION FACTOR EB (TFEB) IN β-CELL VIABILITY AND FUNCTION by Yuanjie Zou B.Sc., University of Science and Technology of China, 2012 M.Sc., Dalhousie University, 2015 A THESIS SUBMITTED IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF DOCTOR OF PHILOSOPHY in THE FACULTY OF GRADUATE AND POSTDOCTORAL STUDIES (Cell and Developmental Biology) THE UNIVERSITY OF BRITISH COLUMBIA (Vancouver) December 2024 © Yuanjie Zou, 2024 ii The following individuals certify that they have read, and recommend to the Faculty of Graduate and Postdoctoral Studies for acceptance, the dissertation entitled: Roles of Autophagy and Transcription Factor EB (TFEB) in β-Cell Viability and Function submitted by Yuanjie Zou in partial fulfilment of the requirements for the degree of Doctor of Philosophy in Cell and Developmental Biology Examining Committee: Dr. Dan S. Luciani, Associate Professor, Surgery, UBC Supervisor Dr. Francis C. Lynn, Associate Professor, Surgery, UBC Supervisory Committee Member Dr. Brian Rodrigues, Professor, Pharmaceutical Sciences, UBC University Examiner Dr. James D. Johnson, Professor, Cellular & Physiological Sciences, UBC University Examiner Additional Supervisory Committee Members: Dr. C. Bruce Verchere, Professor, Pathology and Laboratory Medicine, UBC Supervisory Committee Member Dr. Elizabeth Conibear, Professor, Medical Genetics, UBC Supervisory Committee Member Dr. Sharon M. Gorski, Professor, Molecular Biology & Biochemistry, Simon Fraser University Supervisory Committee Member iii Abstract Diabetes affects more than 500 million people worldwide and results in serious health complications that contribute to millions of deaths annually. Diabetes is characterized by high blood glucose levels resulting from insufficient insulin production due to dysfunction and loss of insulin-producing pancreatic β-cells. The success of islet transplantation as a functional cure for autoimmune type 1 diabetes is also hindered by dysfunction and death of transplanted β-cells. Autophagy is a conserved cellular process that helps maintain cell function and survival by lysosomal degradation and recycling of damaged cellular components under stress. However, the role of autophagy in β-cell survival under hypoxia and following islet transplantation remains unclear. Furthermore, recent evidence suggests that failure of β-cell autophagy may contribute to β-cell dysfunction and the development of both type 1 and type 2 diabetes, but the underlying mechanisms are not understood. This dissertation investigates the importance of autophagy and lysosomal homeostasis in pancreatic β-cells under diabetes- and transplantation-relevant stress. We created a mouse model with β-cell-specific deletion of the autophagy-related gene Atg5 and demonstrated that basal autophagy is essential for maintaining glucose tolerance and β-cell survival. We further found that autophagy protects β-cells under hypoxic stress and during islet transplantation, as loss of autophagy exacerbated β-cell death and islet graft failure. Prolonged hypoxia was found to impair autophagic flux, and this was associated with lysosomal dysfunction and decreased β-cell levels of Tfeb, a transcriptional master regulator of lysosomes and autophagy. By pharmacologically activating Tfeb, we were able to restore lysosomal iv function and improve β-cell viability under stress. To further understand the role of Tfeb in β-cells, we generated a β-cell-specific Tfeb knockout mouse. Loss of β-cell Tfeb made the mice more susceptible to streptozotocin-induced hyperglycemia but did not worsen glucose tolerance under high-fat diet-induced stress, possibly due to compensation by the related transcription factor Tfe3. The research in this dissertation highlights the protective role of autophagy in β-cells under stress and identifies Tfeb dysregulation as a factor in pathogenic β-cell autophagy loss. Targeting Tfeb may offer a therapeutic strategy to improve clinical islet transplantation outcomes and enhance β-cell resilience in diabetes. v Lay Summary Diabetes is a serious disease that occurs when pancreatic beta-cells stop functioning or die, preventing the production of sufficient insulin to lower blood sugar levels. Beta-cell failure and loss also limit the effectiveness of islet transplantation, a promising treatment for type 1 diabetes. In this dissertation, we investigated a cellular process called autophagy (self-eating), where cells remove damaged components by delivering them to lysosomes, the cell’s ‘garbage cans.’ Research shows that beta-cell lysosomes and autophagy stop functioning in diabetes, but the reasons for this remain unclear. We demonstrated that autophagy is essential for beta-cell health and found that TFEB, a protein regulating genes for lysosomes and autophagy, is lost in stressed beta cells. Through drug treatments and gene manipulation, we discovered that restoring or activating TFEB could be a promising strategy to preserve autophagy and protect beta-cells. This knowledge may ultimately help develop improved treatments for diabetes. vi Preface The studies presented in this thesis were conceived and designed by the author, Yuanjie Zou, and the supervisor Dr. Dan S. Luciani. Experiments were carried out and analyzed by Yuanjie Zou, with assistance and input as outlined in the following. Studies from Chapters 3 and 4 are being prepared for submission in a manuscript: Zou Y, Pasula DJ, Tang B, Tang M, Dai DL, Soukhatcheva G, Verchere CB & Luciani DS. Loss of autophagy, Transcription Factor EB and lysosomal homeostasis limit β-cell function and survival under hypoxic stress in vitro and in vivo. Studies were conceived and designed by YZ and DSL. Islet transplantation experiments were designed in collaboration with BV, DLD and GS. YZ carried out the majority of the experiments and data analysis. DJP, BT, MT, DLD and GS helped with experiments as follows - DJP helped with the qPCR experiments in Figs. 4.4, 4.6, 4.9a. BT assisted with qPCR and western blot experiments. MT assisted with mouse husbandry, STZ administrations, in vivo assessments, tissue extractions, and whole pancreas immunohistochemistry. DLD and GS carried out intraductal AAV injections and islet transplantation surgeries. Additional technical assistance for mouse husbandry, islet isolation and tissue preparation was provided by R. Shumiatcher, M. Komba and W.Y. Tam. Studies in Chapter 5 were conceptualized and designed by YZ and DSL. The viral vectors presented in Figure 5.10a were designed by YZ and DSL with input from Dr. Robert Baker. YZ carried out the majority of the experiments and data analysis with help from DJP, MT and WYT. DJP carried out qPCR experiments for Figs. 5.8 & 5.10c, and islet Seahorse experiments for Fig. vii 5.7. MT helped with mouse husbandry, genotyping, in vivo assessments, qPCR and western blot. WYT helped with western blots. Y. Zou wrote this thesis with input and edits from supervisor Dr. D.S. Luciani, and committee members Drs. B. Verchere, F. Lynn, E. Conibear and S. Gorski. All animal studies were approved by the University of British Columbia (UBC) Animal Care Committee under Breeding Protocol A20-0144, as well as Research Protocols A16-0102 and A20-0141. Human islet work was approved by the BC Children's and Women's Research Ethics Board under certificate H13-00554. Regarding gene and protein nomenclature: In the following, gene names are in italic while protein names are not. Specific references to mouse genes and proteins have the first letter of the name capitalized (e.g. Tfeb and Tfeb) while references to human genes and proteins are in all caps (e.g. TFEB and TFEB). The human nomenclature is used in discussion that does not specifically refer to mouse findings. viii Table of Contents Abstract ......................................................................................................................................... iii Lay Summary ................................................................................................................................ v Preface ........................................................................................................................................... vi Table of Contents ....................................................................................................................... viii List of Tables ................................................................................................................................ xi List of Figures .............................................................................................................................. xii List of Abbreviations ................................................................................................................. xiv Acknowledgements .................................................................................................................. xviii Dedication .................................................................................................................................... xx Chapter 1: Introduction ............................................................................................................... 1 1.1 Pancreatic β-cell physiology and glucose homeostasis ..................................................... 1 1.2 β-Cell stress, failure and death in diabetes and islet transplantation ................................. 4 1.2.1. Stress-induced β-cell failure in diabetes and islet transplantation .......................... 4 1.2.2. Causes and mechanisms of β-cell failure ................................................................ 7 1.2.3. β-Cell hypoxia in islet transplantation and T2D ....................................................11 1.2.4. ER stress and the unfolded protein response in β-cells ........................................ 12 1.3 Autophagy in β-cell homeostasis and adaptation to stress ............................................... 14 1.3.1. Introduction to autophagy ..................................................................................... 14 1.3.2. Autophagosome formation and degradation ......................................................... 17 1.3.3. Nutrient- and stress-dependent regulation of autophagy ...................................... 22 1.3.4. Roles of autophagy in pancreatic β-cells .............................................................. 25 1.4 Regulation of autophagy and lysosomes by TFEB .......................................................... 28 1.4.1. TFEB and the MiTF family of transcription factors ............................................. 28 1.4.2. Mechanisms of TFEB regulation .......................................................................... 30 1.4.3. Roles for TFEB in cellular stress-adaptation and metabolism .............................. 33 1.4.4. TFEB in the pancreatic β-cell ............................................................................... 36 1.5 Thesis Overview .............................................................................................................. 38 Chapter 2: Materials and Methods ........................................................................................... 40 ix 2.1 Mice ................................................................................................................................. 40 2.2 Streptozotocin (STZ) injection models ............................................................................ 42 2.3 Islet transplantation .......................................................................................................... 43 2.4 Body weight and glucose metabolism ............................................................................. 44 2.5 Islet isolation, culture, transduction and dispersion ......................................................... 45 2.6 MIN6 cell culture and transfection .................................................................................. 47 2.7 Human islets..................................................................................................................... 47 2.8 Hypoxic culture ................................................................................................................ 48 2.9 In vitro glucose-stimulated insulin secretion ................................................................... 48 2.10 Calcium imaging ............................................................................................................ 49 2.11 Seahorse oxygen consumption measurements & analysis ............................................. 50 2.12 Cell death assays ............................................................................................................ 50 2.13 RNA extraction and RT-qPCR analysis ......................................................................... 51 2.14 Western blotting ............................................................................................................. 53 2.15 Lysosomal cathepsin B activity ..................................................................................... 54 2.16 Lysosomal labelling ....................................................................................................... 54 2.17 Immunofluorescent staining........................................................................................... 55 2.18 3D live-cell autophagy flux imaging ............................................................................. 56 2.19 Statistics ......................................................................................................................... 57 Chapter 3: Autophagy is essential for β-cell function and viability under hypoxia and following islet transplantation ................................................................................................... 58 3.1 Background ...................................................................................................................... 58 3.2 Results .............................................................................................................................. 61 3.2.1. Hypoxia leads to loss of adaptive UPR- and autophagy-related transcripts in pancreatic β-cells. ........................................................................................................... 61 3.2.2. Endogenous autophagic flux is impaired in hypoxic β-cells ................................ 64 3.2.3. Autophagy is important for β-cell survival under hypoxia-induced stress ........... 67 3.2.4. β-Cell-specific deletion of Atg5 causes early-onset glucose intolerance in male and female mice ..................................................................................................................... 70 3.2.5. β-Cell autophagy-deficiency accelerates failure of syngeneic islet grafts ............ 72 x 3.3 Discussion ........................................................................................................................ 77 Chapter 4: Dysregulation of TFEB and lysosomes in β-cells under stress ............................ 82 4.1 Background ...................................................................................................................... 82 4.2 Results .............................................................................................................................. 84 4.2.1. Hypoxia perturbs lysosomal homeostasis in β-cells ............................................. 84 4.2.2. Hypoxia impairs the expression of β-cell Tfeb and its target genes ..................... 87 4.2.3. Inhibition of mTORC1 restores Tfeb, improves lysosomal function and reduces β-cell death under hypoxia ................................................................................................. 89 4.2.4. Glucolipotoxicity- and ER stress-induced β-cell death involves loss of Tfeb ...... 97 4.3 Discussion ........................................................................................................................ 99 Chapter 5: β-Cell-specific Tfeb knockout has minor and sex-specific effects on glucose tolerance: Possible compensation by Tfe3. ............................................................................. 107 5.1 Background .................................................................................................................... 107 5.2 Results ............................................................................................................................ 109 5.2.1. β-Cell-specific Tfeb knockout mice do not develop glucose intolerance ........... 109 5.2.2. β-Cell Tfeb knockout sensitizes to MLD STZ-induced glucose intolerance .......113 5.2.3. β-Cell Tfeb knockout does not worsen glucose tolerance in HFD-fed mice .......114 5.2.4. Sex-dependent transcriptional changes and potential Tfe3-mediated compensation in HFD-fed TfebβKO mice ........................................................................................... 120 5.3 Discussion ...................................................................................................................... 125 Chapter 6: Conclusion .............................................................................................................. 130 6.1 Research summary ......................................................................................................... 130 6.2 Limitations and caveats.................................................................................................. 131 6.3 Future directions ............................................................................................................ 135 References .................................................................................................................................. 140 xi List of Tables Table 2.1 Mouse strains ................................................................................................................ 40 Table 2.2 Mouse diets ................................................................................................................... 42 Table 2.3 Primers used for SYBR Green qPCR ........................................................................... 51 Table 2.4 Antibodies used for Westen Blot ................................................................................... 53 Table 2.5 Antibodies used for immunofluorescent staining .......................................................... 55 xii List of Figures Figure 1.1 The general process of autophagic flux ....................................................................... 20 Figure 1.2 – TFEB-dependent regulation of autophagy and lysosomal homeostasis ................... 30 Figure 3.1 – Time-dependent expression of known hypoxia-responsive transcripts in male C57Bl/6J islets under normal and 1% O2 culture. ........................................................................ 62 Figure 3.2 – Hypoxia-induced changes to islet expression of select UPR / ER stress response genes in male C57Bl/6J islets under normal and 1% O2 culture. ........................................................... 62 Figure 3.3 – Hypoxia-induced changes to islet expression of select autophagy-related genes in male C57Bl/6J islets under normal and 1% O2 culture................................................................. 63 Figure 3.4 – Correlation analysis of hypoxia-induced changes to UPR- and autophagy-related genes ............................................................................................................................................. 64 Figure 3.5 – Principle of autophagic flux measurements by RFP-EGFP-LC3 biosensor ............ 65 Figure 3.6 - Prolonged hypoxia interrupts autophagic flux in mouse islet cells........................... 67 Figure 3.7 – Efficient recombination and Atg5 KO by adenovirus Cre transduction in vitro. ..... 68 Figure 3.8 - Autophagy is essential for β-cell survival under hypoxia. ........................................ 69 Figure 3.9 – Basal Atg5-dependent β-cell autophagy is essential for in vivo glucose tolerance. . 71 Figure 3.10 - Schematic outline of workflow for marginal mass islet transplantation experiment........................................................................................................................................................ 72 Figure 3.11 – dsAAV6-RIP1-Cre induced β-cell Atg5 deletion and autophagy deficiency in vivo........................................................................................................................................................ 73 Figure 3.12 – Characterization of islet graft recipient and donor mice pre-transplantation. ........ 74 Figure 3.13 – β-cell autophagy-deficient islet grafts are sensitized to failure. ............................. 76 Figure 4.1 – Effects of hypoxia on LysoTracker and Lamp-1 staining in Atg5Ad.Ctrl and Atg5Ad.Cre β-cells. ........................................................................................................................................... 86 Figure 4.2 – Individual and combined effects of β-cell Atg5 knockout and lysosomal inhibition on hypoxia-induced islet-cell death. .................................................................................................. 87 Figure 4.3 – Hypoxia-induced changes to islet expression of select lysosome-related genes and lysosomal master regulators Tfeb and Tfe3 in C57BL/6J islets under normal and 1% O2 culture........................................................................................................................................................ 88 Figure 4.4 – Tfeb overexpression increases expression of lysosome- and autophagy-related genes in hypoxic islets. ........................................................................................................................... 89 xiii Figure 4.5 - Hypoxia reduces TFEB immunoreactivity in mouse and human β-cells but can be restored by torin-1. ........................................................................................................................ 91 Figure 4.6 – Effects of Torin-1 treatment on expression of select Tfeb target genes in hypoxic islets........................................................................................................................................................ 92 Figure 4.7 – Hypoxia and Torin-1 treatment increase β-cell Lamp-1 and insulin immunoreactivity........................................................................................................................................................ 93 Figure 4.8 – Hypoxia impairs lysosomal cathepsin B activity in β-cells, but it can be restored by torin-1. ........................................................................................................................................... 95 Figure 4.9 – Torin-1 stimulates the UPR and reduces islet cell death under hypoxia. ................. 96 Figure 4.10 – Tfeb promotes MIN6 β-cell survival and is reduced under lipotoxicity and ER stress........................................................................................................................................................ 98 Figure 5.1 – Confirmation of Tfeb knockout and C57BL/6 Nnt genotype ................................. 109 Figure 5.2 – Body weight, IPGTT and ITT in chow-fed male and female TfebβWT & TfebβKO mice ..............................................................................................................................................112 Figure 5.3 – TfebβKO mice have reduced islet-cell CTSB activity but normal islet function ....113 Figure 5.4 – TfebβKO mice are sensitized to multiple low-dose STZ-induced hyperglycemia .114 Figure 5.5 – β-Cell Tfeb knockout does not worsen high-fat diet-induced glucose intolerance .117 Figure 5.6 – Ca2+ signaling and insulin secretion in islets from HFD-fed TfebβWT & TfebβKO mice ..............................................................................................................................................118 Figure 5.7 – Mitochondrial respiration in islets from HFD-fed TfebβWT & TfebβKO mice. ....119 Figure 5.8 – Tfe3 up-regulation and sex-specific effects on lysosomal and mitochondrial transcriptional adaptation to diet-induced stress in islets from HFD-fed TfebβKO mice. ......... 121 Figure 5.9 –Tfe3 and Lamp1 immunostaining in β-cells from HFD-fed TfebβKO & TfebβKO mice...................................................................................................................................................... 122 Figure 5.10 – Tfe3 upregulation and partial compensatory gene expression in the absence of Tfeb..................................................................................................................................................... 124 xiv List of Abbreviations AAV adeno-associated virus dsAAV double-stranded adeno-associated virus ADP adenosine diphosphate AKT protein kinase B AMPK AMP-activated protein kinase ATF4 activating transcription factor 4 ATF6 activating transcription factor 6 ATG autophagy-related proteins ATP adenosine triphosphate AUC area under the curve BCCHR British Columbia Children’s Hospital Research CaN calcineurin CD control diet CHQ chloroquine CLEAR coordinated lysosomal expression and regulation CMA chaperone-mediated autophagy CTSB cathepsin B DDIT3 DNA damage inducible transcript 3 DMEM Dulbecco’s modified Eagle’s medium xv ER endoplasmic reticulum ERK1/2 the extracellular signal-regulated kinase 1/2 ETC electron transport chain FBS fetal bovine serum FFA free fatty acids GLP1R glucagon-like peptide 1 receptor GSIS glucose-stimulated insulin secretion GSK3β glycogen synthase kinase 3β HFD high fat diet hIAPP human islet amyloid polypeptide HIF-1α hypoxia-inducible factor 1α HSPA5 heat shock protein family A (Hsp70) member 5 IPGTT intraperitoneal glucose tolerance test IRE1 inositol-requiring enzyme-1 ITT insulin tolerance tests KRB Krebs-ringer buffer LAMP lysosome-associated membrane protein LC3 microtubule-associated protein 1A/1B-light chain 3 MCOLN1 mucolipin-1 MITF microphthalmia-associated transcription factor xvi MLD multiple low dose MOI multiplicity of infection mTORC1 mechanistic target of rapamycin kinase complex 1 NADH nicotinamide adenine dinucleotide hydrogen NRF2 nuclear factor erythroid 2-related factor 2 OCR oxygen consumption rate OPTI-MEM optimized minimal essential medium PE phosphatidylethanolamine PERK protein kinase RNA-like ER kinase PFA paraformaldehyde PGC-1α proliferator-activated receptor γ coactivator-1 alpha PI propidium iodide PI3K phosphatidylinositol 3-Kinase qPCR quantitative PCR Rab7 Ras-associated binding protein 7 RIP1 rat insulin promoter-1 ROS reactive oxygen species RPMI Roswell Park Memorial Institute 1640 media STZ streptozotocin T1D type 1 diabetes xvii T2D type 2 diabetes TCA cycle citric acid cycle (tricarboxylic acid cycle) TFE3 transcription factor E3 TFEB transcription factor EB TFEC transcription factor EC Tg thapsigargin ULK1 Unc-51 like autophagy activating kinase 1 UPR unfolded protein response v-ATPase vacuolar-type proton ATPases XBP1 X-box binding protein 1 XBP1s spliced X-box binding protein 1 ZKSCAN3 zinc finger with KRAB and SCAN domains 3 xviii Acknowledgements My PhD training has been a journey filled with both challenges and rewards. I could not have reached this milestone and completed my degree without the support of those around me. Your kindness and advice have guided me through this long journey and will be cherished deeply. First, I am grateful to my PhD supervisor, Dr. Dan S. Luciani, for your guidance throughout my research. Your generosity and patience over the past years have been invaluable. Your discussions about science consistently inspired my work, encouraging me to think boldly while also urging me to approach my research with careful consideration. You taught me the importance of setting clear goals and objectives, which greatly enhanced the efficiency of my training. Additionally, I appreciate your support as I adapted to life in a new country and unfamiliar city. I extend my gratitude to the members of my supervisory committee: Dr. Sharon Gorski, Dr. C. Bruce Verchere, Dr. Francis Lynn, and Dr. Elizabeth Conibear. Your knowledge in the fields of autophagy, diabetes, β-cell biology, and cellular stress response greatly informed the design of my research. I also want to thank Dr. Jim Johnson for your insightful questions during my comprehensive exam, which significantly improved my research proposal. I feel incredibly fortunate to have worked with such a great team and am grateful to the past and present members of the Luciani Lab for your expertise and support. Specifically, I would like to thank Mei Tang for assisting with many in vivo experiments, W.Y. Tam for your help in preparing and staining pancreas sections, Daniel J. Pasula for sharing your knowledge in gene xix transcription and mitochondrial biology as well as the culture of Canada, and Rocky Shi for showing me around this beautiful city. I truly enjoyed my time in the Luciani Lab, and the experiences shared with all of you were an essential part of my training. I am also deeply thankful to our collaborators from the Verchere Lab, Derek Dai and Galina Soukhatcheva, for your professional advice and technical support in critical animal experiments. Many thanks to Meg Hughes and the BCCHR support team for organizing outstanding research events, allowing me to engage with Vancouver's vibrant diabetes and islet biology research community. I also acknowledge the animal care staff at BCCHR and CDM for their excellent support in managing my animal colonies. Finally, my deepest thanks go to my family in China and to my wife, Jingyi Li, here in Vancouver. Your unwavering support and understanding have helped me through the toughest times, especially during the pandemic. Without you, I could not have overcome the challenges. You have made my life meaningful and filled it with love. xx Dedication To my parents & my wife for your love and understanding 1 Chapter 1: Introduction 1.1 Pancreatic β-cell physiology and glucose homeostasis Diabetes is a prevalent health issue that poses a significant threat to public health and the global economy. According to the International Diabetes Federation, the number of adults aged 20 to 79 living with diabetes reached 537 million in 2021, triple the estimated number from 2000 (1). The complications associated with diabetes have led to approximately 6.7 million deaths annually. Projections from the IDF indicate that the prevalence of diabetes could rise to 783 million by 2045, highlighting an urgent need for effective global management strategies (1,2). Diabetes is characterized by high blood glucose levels resulting from insufficient insulin release and ineffective insulin action, a condition known as insulin resistance. Insulin, a peptide hormone produced by pancreatic β-cells, promotes glucose uptake by fat and muscle cells to maintain proper glucose control (3). Impaired insulin production and secretion due to loss or dysfunction of β-cells is a critical pathological event in developing hyperglycemia and associated complications (4,5). Proper insulin production and secretion in β-cells is a complex multistep process involving several cellular compartments, especially the rough endoplasmic reticulum (ER) and mitochondria. Processing and folding of insulin mainly take place in the ER. In mice, insulin is encoded by two genes, Ins1 and Ins2; in humans, it is encoded by INS. Initially, insulin is translated into pre-proinsulin with a single peptide with A- and B- chains, an N-terminal signal peptide, and a C-peptide (3). The signal peptide directs pre-proinsulin to the ER, where it is processed into 2 proinsulin by cleaving the signal peptide and forming multiple disulfide bonds between the A- and B-chains, stabilizing insulin’s three-dimensional structure. Insulin is continuously produced and folded in the ER to regulate blood glucose levels. Consequently, pancreatic β-cells have highly efficient ER networks to meet the high-volume insulin folding task. Once properly folded and processed in the ER, proinsulin is transported to the Golgi apparatus for further modification and sorting into immature secretory granules. During granule maturation, proinsulin has its C-peptide removed, finally forming an active insulin peptide, which is crystalized in hexamers ready for glucose-induced secretion. Excessive insulin demand, such as under conditions of nutrient excess or insulin resistance, can induce ER stress and activate the unfolded protein response (UPR), potentially disrupting insulin synthesis (6). Furthermore, disulfide bond formation is a redox-dependent process, and high levels of proinsulin processing and folding can contribute to oxidative stress. Therefore, it is necessary to have a delicate balance in insulin production to avoid detrimental stress in β-cells. While ER is fundamental in the production of insulin, mitochondria play a pivotal role in insulin secretion by metabolizing glucose and generating ATP, as well as other key signals that trigger and shape the release of insulin granules. Generally, insulin is secreted biphasically. The first phase involves the exocytotic release of readily available granules upon glucose stimulation, and the second phase involves ATP-dependent mobilization of stored granules to the plasma membrane for release (7). Granule exocytosis is calcium-dependent, relying on the opening of voltage-dependent calcium channels on the plasma membrane, and this process is closely tied to 3 glucose metabolism by mitochondria. When β-cells take up glucose, it is converted into pyruvate through glycolysis in the cytoplasm. In the canonical model for glucose-stimulated insulin secretion (GSIS) mitochondrial metabolism then drives subsequent events that trigger and amplify insulin granule release. Pyruvate enters the mitochondria, fueling the citric acid cycle (TCA cycle) that generates nicotinamide adenine dinucleotide hydrogen (NADH), which enters the electron transport chain (ETC) as an electron carrier. The ETC is a series of proteins bound on the mitochondrial inner membrane, in which the electrons delivered by NADH go through redox reactions and release energy, which promotes the ETC to pump protons from the matrix into the mitochondrial intermembrane space. The accumulation of protons in the intermembrane space establishes a proton gradient across the inner mitochondrial membrane that powers the ATP synthase, which converts adenosine diphosphate (ADP) to adenosine triphosphate (ATP). The resulting increase in the cytosolic ATP/ADP ratio inhibits ATP-sensitive potassium channels on the plasma membrane, leading to membrane depolarization, activation of voltage-dependent calcium channels, calcium entry and insulin granule exocytosis in the first phase (8). Additionally, other metabolites, such as citrate, produced in the TCA cycle, contribute to further increasing the exocytosis of secretory granules in the second phase via mechanisms known as the amplifying pathways (9). Notably, an alternative model was recently proposed where the initial ATP that triggers membrane depolarization and GSIS is generated by pyruvate kinase near the plasma membrane, but in this model mitochondria-derived ATP and metabolites are still essential for sustained insulin release (10). 4 In summary, insulin-producing pancreatic β-cells are essential for glucose homeostasis. Their ability to synthesize and release insulin, which relies heavily on the function of ER and mitochondria, is crucial for lowering blood glucose levels after a meal. Therefore, conditions that cause β-cell failure or loss will result in the development of hyperglycemia leading to diabetes. 1.2 β-Cell stress, failure and death in diabetes and islet transplantation 1.2.1. Stress-induced β-cell failure in diabetes and islet transplantation Diabetes can be classified into different types and the major forms are type 1 diabetes (T1D) and type 2 diabetes (T2D). Loss and dysfunction of β-cells are major pathogenic events in the progression of both forms of diabetes, but the causes are different. T2D, which accounts for about 90% of the total diabetes cases, is characterized by a progressive loss of adequate β-cell function and insulin secretion. Insulin resistance, which can be caused by physical inactivity and obesity, initially prompts pancreatic β-cells to adapt during the prediabetic stage in the development of T2D by increasing their mass and activity (11). However, this increased workload and activity can lead to ER- and oxidative stress-induced damage to the viability and function of a portion of β-cells (6,12). Prolonged hyperglycemia becomes prevalent when the adaptation mechanisms in β-cells fail to compensate for the increased demand for insulin, further stressing the remaining β-cells and accelerating disease progression (4). T1D is characterized by immune cell-mediated killing of pancreatic β-cells, which results in insufficient insulin production and hyperglycemia. In addition to the death caused by direct 5 interaction with activated β-cell specific macrophages and T-cells, exposure to increased inflammatory cytokines, including interleukin-1β, interferon-γ and tumor necrosis factor-α, induces transcriptional and functional changes in β-cells and leads to the impairment of insulin secretion (13). Inflammatory cytokines also cause ER calcium depletion and mitochondria membrane disruption and contribute to the loss of functional β-cells. Islet transplantation has emerged as a promising β-cell replacement treatment for T1D patients. According to the current standard protocol, islets are infused into the hepatic portal system, where the transferred β-cells can work effectively in insulin secretion. When successful, this procedure can help patients regain the ability to control blood glucose levels without exogenous insulin (14). However, a major limiting factor is the rapid loss of graft viability and insulin production due to conditions such as hypoxia, glucotoxicity and inflammation (15). These risk factors also activate the intracellular pathways of ER and oxidative stress, further limiting the success rate of the islet transplantation (16,17). The progression of stress-induced β-cell function and loss may be affected by internal factors, such as sex hormones. It is known that the incidence of diabetes varies between the sexes, being more prevalent and occurring earlier in males. The influence of sex differences on β-cell failure is increasingly recognized as an important topic of research (18-20). Female sex hormones have been implicated in both T1D and T2D and are suggested to enhance β-cell regeneration in rodent models, although this has not been established in human β-cells (21). Meta-analyses of human single β-cell RNA-Seq data indicate a male-specific loss of 6 transcriptional integrity related to β-cell identity and function with age (22). On the other hand, female rats are protected from age-dependent loss of β-cell insulin secretion, reflecting increased mitochondrial function and expression of mitochondrial biogenesis genes (23). A recent study by Noelia Jacobo-Piqueras et al. suggests female islets have better glucose-induced electrical activity, higher intracellular calcium amplification, or enhanced insulin granule exocytosis, providing further evidence for better function of female β-cells and the underlying cellular mechanisms (24). The role of sex differences in β-cells stress responses is less studied. Recently, a single-cell analysis study of human β-cells revealed differential enrichment of UPR pathway-associated genes, with higher expression of genes linked with protein synthesis, folding, and processing in females, which may contribute to a higher resilience in female islets against chemically induced ER stress (19). Although it is not the primary focus, this thesis therefore also considers possible sex difference in some instances, mostly related to the in vivo effects of β-cell-specific gene deletion. Overall, the progression of diabetes and the failure of islet transplantation involve β-cell dysfunction and loss. Multiple stress-inducing conditions and downstream pathways are involved, and these may show sex differences. Therefore, it is important to identify the types of stress causing β-cell dysfunction under specific conditions, so that potential interventions or preventative measures can be developed accordingly. 7 1.2.2. Causes and mechanisms of β-cell failure As briefly introduced in the previous section, external inducers of stress, including glucolipotoxicity, inflammation and hypoxia, are associated with β-cell failure in diabetes and islet transplantation. The cellular responses to these inducers converge on shared intracellular pathways that can negatively impact β-cell health, e.g. ER and oxidative stress. Glucolipotoxicity: Glucolipotoxicity describes the combined deleterious effects caused by excess glucose and free fatty acids (FFA). Excessive nutrient uptake, lack of physical exercise and obesity are believed to be significant inducers of glucolipotoxicity, which contributes to insulin resistance and β-cell dysfunction (25). Pancreatic β-cells initially respond to glucolipotoxicity and insulin resistance with a compensatory expansion in mass and increased insulin secretion, but chronic exposure to increased levels of circulating glucose and FFA leads to a phase of decompensation where β-cell failure and death promote the progression of T2D (11). The consequences of glucolipotoxicity include increased ER stress and activation of the UPR in β-cells due to more demand for insulin processing and disruption of ER calcium homeostasis (25). In addition, it stimulates excess formation of reactive oxygen species (ROS) and leads to oxidative stress (25). Both forms of stress cause β-cell dysfunction and apoptosis and contribute to β-cell decompensation. Inflammation: Inflammation plays distinct roles in β-cell failure in different types of diabetes. Islet inflammation during T1D is marked by the infiltration of various leukocytes. Among these immune cells, β-cell specific CD4+/CD8+ T-cells contribute predominantly to the 8 failure and dysfunction of β-cells (26). In T2D, a low-level, chronic inflammatory state contributes to insulin resistance and dysfunction or death of β-cells (27). Various stress inducers including elevated glucose/ FFA and pro-inflammatory cytokines lead to recruitment of pro-inflammatory macrophages which destroy β-cells (28). Additionally, inflammation acts as a risk factor in islet transplantation, as the allogenic islet graft from donors will likely be recognized as exogenous antigens and attacked by the host immune system. Therefore, to reduce the host immune response, immunosuppression treatment is applied in the islet transplant protocol to preserve the function of the islet graft (29). Islet Amyloid: Human islet amyloid polypeptide (hIAPP) is released alongside insulin, and generally acts as a signaling molecule in controlling food intake and glucose levels by delaying gastric emptying. However, high concentrations of incompletely processed hIAPP lead to islet amyloid formation, a hallmark of T2D development. hIAPP aggregates also appear quickly post-islet transplantation (30). It has also been suggested that hIAPP is an autoantigen during the development of T1D (31), but the exact role in the disease development requires further investigations. hIAPP cytotoxicity is linked to the dysregulation of ER and mitochondria, inflammation, and impaired autophagy, which all contribute to β-cell failure (32-34). Hypoxic stress: β-Cells receiving reduced oxygen face hypoxic stress. As will be discussed further in Section 1.2.3, hypoxia is commonly seen after islet transplantation prior to graft revascularization and can also be induced in T2D when β-cells are under high metabolic demand and mitochondrial respiration consumes a large amount of oxygen. Hypoxia is associated with 9 the stabilization and activation of hypoxia-inducible factor 1α (HIF-1α), a key transcriptional regulator of the cellular response to hypoxia. HIF-1α-dependent reprogramming of β-cell gene transcription can lead to impaired glucose-stimulated insulin secretion (35), in part by stimulating metabolism through lactate dehydrogenase A, which redirects pyruvate toward lactate formation and away from the TCA cycle and mitochondrial ATP generation. Lack of oxygen also directly reduces the production of ATP molecules through the ETC, further impairing glucose-stimulated insulin release. Because oxygen is required to form disulfide bonds, hypoxia also impairs β-cells insulin folding, which can cause harmful ER stress and result in β-cell apoptosis through the upregulation of DNA damage-inducible transcript 3 (DDIT3), also known as C/EBP homologous protein (CHOP). Hypoxia also leads to ROS formation, which both contributes to the stabilization of HIF-1α and is directly related to oxidative stress (36). ER stress: The ER is central to insulin synthesis and is critical for β-cell function. ER stress occurs whenever the protein-folding capacity of the ER is overwhelmed, for instance under conditions of excessive nutrient supply. Exposure to various harmful conditions, including glucolipotoxicity and hypoxia, can cause the accumulation of misfolded proteins in the ER. Once the accumulation of unfolded proteins reaches a critical threshold, an adaptive signal transduction pathway, the UPR, is induced. To alleviate ER stress, the UPR will downregulate general transcription and protein translation but selectively increase transcription and translation of proteins that help ER biogenesis, protein folding, and degradation of misfolded proteins (37). As β-cells are highly active in insulin synthesis, a constitutively active adaptive UPR is required, 10 and the margin for adjustment is limited. Thus, challenged β-cells may more quickly cross from ER stress adaptation to apoptosis, which leads to loss of insulin secretion, increased workload for insulin production in the remaining β-cells, and accelerated progression T1D or T2D. In addition to the adaptive UPR, emerging studies suggest ER-phagy, a selective form of autophagy, removes damaged organelles and helps maintain ER quality control (38). Mitochondrial stress: Mitochondria consume a large quantity of O2 to generate the energy supply for insulin production and secretion, but during respiration, electrons leak from the mitochondrial ETC, leading to the generation of ROS (12). Although studies have shown that ROS acts as a signaling molecule in insulin secretion, excessive ROS generation is a major inducer of mitochondria dysfunction and oxidative stress. Increasing metabolic demand due to elevation of blood glucose and FFA levels contributes to prolonged exposure to ROS in β-cell mitochondria, leading to defective gene expression, reduced glucose secretion and increased β-cell apoptosis (12). Antioxidant enzymes such as superoxide dismutase, glutathione peroxidase, and catalase can help mitigate the effects of increasing ROS production, but their activities and levels are relatively low in β-cells, which makes β-cells sensitive to oxidative stress. Recently, some other antioxidant enzymes, e.g. peroxiredoxin/thioredoxin, are suggested to be upregulated in β-cell in an attempt to alleviate the harmful consequences of ROS accumulation (39). Moreover, mitophagy, the selective autophagy of mitochondria, contributes significantly to maintaining mitochondria health (38,40). 11 1.2.3. β-Cell hypoxia in islet transplantation and T2D Although the causes of hypoxia differ in islet transplantation and T2D, it can contribute to β-cell failure in both situations. In the pancreas, β-cell insulin production is supported by a highly developed vascular network, which is destroyed during the preparation of donor islets for graft transplantation (14). Therefore, the isolated islets lack sufficient oxygen supply, leading to hypoxia in β-cells. It takes approximately two weeks to revascularize the donor islets in the recipient (41). Since the failure of islet graft is correlated with the level of hypoxia (42), before revascularization occurs, oxygenation treatments were used as a strategy to alleviate islet hypoxia (41). However, oxygen application level and duration must be optimized carefully, as excessive oxygen is also toxic to the isolated islets. The production and secretion of insulin also consume a large amount of oxygen, so when β-cells are overstimulated by elevated glucose and FFA levels during the development of T2D, intracellular oxygen consumption can activate β-cell hypoxia signaling (35). Although the hypoxia induced by excessive nutrient metabolism is relatively moderate and less abrupt compared to devascularization, it also contributes to the dysfunction and failure of β-cells. In addition to activating HIF-1α signaling, moderate hypoxia causes HIF-1α independent suppression of genes that are important for β-cell function, including Mafa, Pdx1 and Ins1 (43), likely via activation of the transcriptional repressor BHLHE40 (44). β-cells counter the negative impact of hypoxia by activating adaptive and protective mechanisms, and failure of these can worsen β-cell loss. For example, impaired ER-Golgi trafficking, suppression of adaptive UPR 12 signaling, and upregulation of the pro-apoptotic ER stress mediator DDIT3/CHOP may contribute significantly to hypoxia-induced β-cell death (45,46). 1.2.4. ER stress and the unfolded protein response in β-cells Insulin production imposes a significant burden on the β-cell ER and can lead to a state of basal ‘ER stress’ with induction of the adaptive UPR to meet demand. Therefore, the margin for β-cell UPR adaptation under ER stress is narrow, making β-cell more susceptible to pro-apoptotic ER stress (47). Activating transcriptional factor 6 (ATF6), inositol-requiring enzyme-1 (IRE1), and protein kinase RNA-like ER kinase (PERK) are three ER stress sensors that initiate parallel transcriptional responses when there is an accumulation of misfolded proteins in the ER lumen (48). The ER chaperone, HSPA5, also known as immunoglobulin heavy-chain-binding protein (BIP) binds with these three sensors to keep them inactive under non-stress conditions (49). Under ER stress, it is released to interact with unfolded proteins and facilitate their proper folding, which results in activation of ATF6, IRE1 and PERK signaling. UPR mediated by different pathways plays complex roles in β-cells and can be either adaptive or harmful (50). PERK is involved in the regulation of proinsulin folding capacity. Inhibition of PERK by gene knockout, dominant-negative mutation, or chemical inhibition results in abnormal proinsulin folding (51-53). In contrast, PERK activation suppresses insulin production under low glucose conditions (52). IRE1 achieves much of its protective role by cleaving X-box binding protein 1 (XBP1) mRNA into a spliced form (XBP1s). The translated 13 protein of XBP1s acts as a transcription factor that stimulates protective pathways by upregulating chaperones, folding enzymes and ER-associated protein degradation components to reduce ER stress. Reduction of IRE1-XBP1 signaling is associated with upregulation of DDIT3/CHOP and cell death in NOD T1D mice and obese insulin resistant ob/ob mice, while induction of IRE1 signaling protects INS-1 cells against glucolipotoxicity (54). Notably, the XBP1s-mediated transcriptional changes must be finely regulated, as overexpression of XBP1s via adenovirus suppresses insulin secretion in rats (55). Although the role of ATF6 in insulin production is less understood, its deletion reduces insulin content in mice under HFD (56). Importantly, all three UPR arms are involved in regulating β-cell apoptosis. Increased β-cell death occurs in multiple models with PERK deletion or inhibition (57-59). Otherwise, PERK can mediate autophagy and maintain β-cells under stress (60). Knockdown of IRE1 increased β-cell death and (61) this could be rescued by overexpression of XBP1s, suggesting the protective role of IRE1 is XBP1s dependent (61). However, the UPR pathways can also signal β-cell apoptosis in some conditions. In several animal models, PERK, and possibly ATF6 as well, mediated β-cell death through induction of DDIT3/CHOP (62,63). IRE1 can mediate β-cell death activation of JNK (54). Interestingly, although XBP1s is generally considered adaptive for β-cells, hyperactivation of IRE1 and sustained expression of XBP1s can cause β-cell death (55). In summary, a balance between adaptive and maladaptive UPR signaling is critical for β-cell survival. The adaptive UPR fine tunes the stress levels in β-cells, especially through its direct effects on the protein processing in ER. UPR signaling also affects redox homeostasis and other 14 protective mechanisms, including autophagy, that may cooperate with the UPR to affect the health of β-cells. 1.3 Autophagy in β-cell homeostasis and adaptation to stress 1.3.1. Introduction to autophagy Cells employ different adaptation mechanisms to survive and function during stress. An essential part of cellular adaptation is to remove unwanted substances and recycle them for valuable molecules. Autophagy, which derives its name from ancient Greek meaning ‘self-eating’, is a process characterized by lysosomal digestion of a cell’s own materials, which are delivered to the lysosomes via different routes (64). Unlike the ubiquitin-proteasome system, which is another important intracellular degradation pathway that mainly clears smaller, short-lived proteins, autophagy is responsible for the removal and recycling of longer-lived proteins, protein aggregates and damaged cellular organelles (65). Through autophagic clearance, cells achieve quality control of their organelles, which is critical for proper cellular function (64). In addition, autophagic degradation recycles essential biological molecules, including amino acids, FFA and adenosine phosphates, that can be used as building blocks and fuels to support cell survival and metabolism under starvation and other stress (66). Autophagy is a highly conserved process in eukaryotic cells. Studies of autophagy started in yeast and have since expanded to provide a wealth of information regarding the roles and importance of autophagy in most mammalian cells and many human disorders (67). In 2016, the 15 importance of autophagy was recognized by the Nobel Prize in Physiology or Medicine being awarded to Dr. Yoshinori Ohsumi for his seminal discoveries of its mechanisms (68). Three major types of autophagy have been described, each with a distinct mechanism to deliver the cargos to lysosomes: i) microautophagy, ii) chaperone-mediated autophagy (CMA) and iii) macroautophagy. In microautophagy, the targets for degradation are directly internalized by lysosomes and late endosomes (69). The membrane deformation processes during microautophagy - including the internalization of cytosolic cargo in late endosomal multivesicular bodies and the final membrane scission of lysosomes - rely on a series of cytosolic protein complexes, the endosomal sorting complexes required for transport (70). Unlike microautophagy, CMA does not require invaginations of the lysosomal membrane but rather engages a series of proteins that help the direct translocation of cargo across the membrane (71). CMA operates in a highly selective way, digesting only specifically modified and folded soluble proteins that are recognized by the cytosolic chaperone heat shock protein of 70 kD, but not sub-cellular organelles (72). Macroautophagy is by far the most studied and best-characterized form of autophagy. In this thesis, I will mainly focus on macroautophagy and will refer to this simply as autophagy in the following. As will be described in more detail in Section 1.3.2, removal by autophagy requires the formation of a double-membrane autophagosome, which encloses the intended cargos and delivers them to the lysosomes for degradation (73). Autophagy can be a mechanism for bulk degradation of cytosolic proteins. In addition, multiple mechanisms exist by which autophagy 16 can selectively recognize and degrade distinct cargos – including, but not limited to, protein aggregates, external pathogens, and various organelles such as mitochondria, lysosomes, peroxisomes, and ER (74). Selective cargo recognition involves various autophagy receptors, often via poly-ubiquitin tags, that occurs prior to the autophagosome’s formation. Among the various organelle-targeted forms of autophagy, mitophagy is the most studied and best understood (75). This process involves the isolation and selective clearance of damaged mitochondria via the PINK1-Parkin system. Briefly, the PINK1 kinase accumulates on the outer membrane of depolarized mitochondria, where it recruits and phosphorylates Parkin, an E3 ubiquitin ligase (76). Parkin then ubiquitinates mitochondrial proteins in the outer membrane, causing autophagic adaptor proteins (e.g. p62) to bind and deliver the organelles to the forming autophagosome via their interaction with microtubule-associated protein 1A/1B-light chain 3 (LC3) (77). Mitophagy can also be regulated by another E3 ubiquitin ligase CLEC16A, which is also a T1D susceptibility gene (78). Moreover, several mitochondria-located proteins - including FUNDC1, BNIP3 and BNIP3L – also act as important mitophagy receptors to help maintain mitochondrial homeostasis (75). Overall, autophagy is also an essential mechanism for organelle quality control. It is now widely recognized for its key role in maintaining organelle function and cellular health, including in β-cells and diabetes (38,67). 17 1.3.2. Autophagosome formation and degradation Studies on the mechanisms and machinery of autophagy, often in yeast, have shown that it involves the formation of an autophagosome, followed by its delivery to and fusion with a lysosome (79). This fusion process results in the formation of autolysosomes in which the cargo is degraded and recycled. The knowledge of autophagy has expanded since its discovery, and the following chapters will mainly focus on the machinery and molecules in mammalian cells. 1.3.2.1. Autophagosome biogenesis and ATG proteins The first step in forming an autophagosome involves assembling a double-membrane structure, known as the phagophore, usually at a site on the ER membrane (80). The phagophore expands and grows into a spherical shape, which, with the help of autophagy adaptors like p62, encloses and isolates a part of intracellular materials – i.e. the autophagic cargo (81). Autophagosome biogenesis completes when the pore structure on the phagophore closes, and the organelle is split from the initiation site. The details of phagophore enclosure are still unclear, although recent studies suggested the ESCRT complexes may mediate this process (82). A subset of autophagy-related (ATG) proteins are key players in the formation of autophagosomes (83). These proteins, along with multiple metabolism-associated kinases and receptors for selective autophagy targets, initiate the signal that activates the assembly of autophagy initiation complexes. This is achieved through the autophosphorylation of Unc-51-like kinases 1 (ULK1, Atg1 in yeast) and their binding with ATG13-FIP200-ATG101 (Atg17–18 Atg31–Atg29 in yeast) in mammalian cells (84). The formed complexes consistently stimulate the kinase activity of ATG1/ULK and serve as a platform to recruit other components for autophagy (85). Some studies suggest that initiating the phagophore membrane requires a membrane enriched with ATG9A (Atg9 in yeast), which may deliver membrane materials to autophagosome precursors (86). During the elongation process, the phagophore membrane is suggested to be tethered to the initiation site on the ER. This tethering is facilitated by ATG2–WD-repeat protein, a crucial player that interacts with phosphoinositide (87). The recruited components bind with phagophore membranes to regulate phosphatidylinositol 3-kinase (PI3K) complexes. For instance, one of the components of PI3K complexes, BECLIN1 (Atg6 in yeast), can be phosphorylated by ULK1 and contribute to the production of Phosphatidylinositol 3-phosphate on the membrane and autophagosome maturation (88). During autophagosome biogenesis, the elongation process involves the conjugation of phosphatidylethanolamine (PE) with LC3 (Atg8 in yeast), which then mediates the integration of the ULK complex with the phagophore membrane. Conjugation of LC3 relies on two ubiquitin-like conjugation systems, the ATG5-ATG12-ATG16L system and the LC3-PE lipidation system (89). ATG5 first interacts with ATG12 and then localizes on the autophagosome outer membrane after forming the complex with ATG16L (89). This complex catalyzes the lipidation of LC3 that has been activated and transferred by ATG7 and ATG3 (89). In addition to its role in membrane elongation, LC3 participates in cargo selection and fusion processes, both before and after 19 autophagosome membrane closure, making it a widely accepted and used marker of autophagosomes (74) (Figure 1.1). After autophagosome-lysosome fusion, Atg8/LC3 is recycled for the next round of autophagosome biogenesis via delipidation by ATG4 (74). 1.3.2.2. Autophagosome-lysosome fusion and cargo degradation Autophagosomes deliver their cargo to lysosomes following an autophagosome-lysosome fusion process. The fused organelle is known as an autolysosome, and the cargo is broken down by various lysosomal hydrolases in the acidified lumen of this vesicle (90). The dynamic processes of autophagosome formation, fusion with lysosomes, and continuous cargo degradation are collectively known as autophagic flux; its measurement is widely accepted to assess the overall activity of successful autophagy in a cell or tissue (73) (Figure 1.1). Autophagosomes and lysosomes are first primed for further fusion steps as multiple small GTPases - such as Ras-associated binding protein 7 (Rab7) - appear on the vesicle membrane and regulate the interaction between organelle membranes and the motor proteins (91). Binding motor proteins with small GTPases or the vesicle membranes enables microtubule trafficking and the encounter of autophagosomes and lysosomes. After proper positioning through the microtubular network, autophagosomes are tethered to the lysosomal surface by adaptor proteins interacting with GTP-bound (i.e. active) small GTPases (92). The subsequent fusion between autophagosome-lysosome membranes is executed by multiple SNARE complexes (92). 20 Autolysosomal cargos are broken down into small molecules by more than 50 hydrolases found in the lysosomal lumen. Cathepsins are the primary group of proteases, divided into three sub-types: serine proteases (cathepsins A and G), cysteine proteases (cathepsins B, C, L, F, H, K, O, S, V, X and W) and aspartic proteases (cathepsins D and E) (93). The interaction between protons and the active sites of cathepsins is critical for their structural stability, thus an acidic lysosomal pH is required for optimal cathepsin function. For instance, lysosomal alkalization leads to inactivation and irreversible structural changes of human cathepsins B (CTSB) and L. Acidic lumens are separated from the cytosol by the limiting lysosomal membrane, where lysosome-associated membrane proteins (LAMP) are aligned and serve to sequester the hydrolytic enzymes in the lumen from the phospholipid bilayer (94). The most abundant LAMPs, LAMP1 and LAMP2, constitute up to 50 percent of the lysosomal membrane proteins and act as the fundamental elements constructing the lysosomal membrane. In addition, they play Figure 1.1 The general process of autophagic flux Schematic summarizing key stages of autophagic flux. Autophagosomes grow from a double-membrane phagophore and engulf cellular aggregates or dysfunctional organelles. ATG5 forms a complex with ATG12 and ATG16L to facilitate the lipidation of LC3, which is critical for autophagosome formation and function. In the last part of autophagic flux the autophagosome fuses with a lysosome to generate an autolysosome in which the autophagic cargos are degraded. See text for further details. Created with BioRender.com. 21 important roles during the fusion of autophagosomes and lysosomes (95). Interestingly, the deletion of LAMP1 does not directly destabilize the lysosomal limiting membrane, possibly due to compensation from LAMP2 (96). However, double knockout of both LAMPs leads to lethal phenotypes and defects in the trafficking of cholesterols. Double knockout also leads to the accumulation of autophagic vacuoles and altered lysosomal distribution, indicating their roles in lysosomal trafficking and degradative function (97). LAMP2A, an isoform of LAMP2, acts as a receptor of the target proteins and then forms the complexes that facilitate the transport of the CMA targets across the lysosomal membrane (98). Acidification of the lysosomal lumen is mainly maintained by vacuolar-type proton ATPases (v-ATPase) in an ATP-dependent manner (99). By catalyzing the hydrolysis of ATP, V-ATPase transfers protons across plasma or organelle membranes. The V-ATPase is a multi-subunit complex comprising the soluble V1 and the membrane-associated V0 domains. The V1 domain assembles from eight types of subunits (A-H) critical for ATP hydrolysis. Six subunits comprise the V0 domain, which mediates the proton transport through membranes (100). Traditionally, acidification is considered to occur before autolysosome formation and contributes to the regulation of autophagosome-lysosome fusion. However, a recent investigation suggested that acidification is not required and can be achieved after the formation of autolysosomes (101). Regulatory roles of calcium have also been characterized in autophagosome biogenesis and autophagosome-lysosome fusion (102). By releasing lysosomal calcium storage and mediating an elevation of cytosolic calcium, lysosomal calcium channels, like MCOLN1, can trigger 22 autophagy by activating calcium-dependent kinases that increase ULK1 activity (103) and by promoting calcineurin (CaN)-dependent dephosphorylation and activation of transcriptional regulators, such as transcription factor EB (TFEB), which I will discuss in more detail later (104). Recent studies also suggest the critical role of several lysosomal calcium channels in autophagosome-lysosome fusion steps (105-107). 1.3.3. Nutrient- and stress-dependent regulation of autophagy Autophagy is known to be activated by multiple conditions of cellular stress, of which starvation is the best studied. Starvation- or stress-induced autophagy has been reported in various cell types and is generally regarded as an adaptive and cell protective mechanism (64). Signaling pathways that sense changes in nutrient availability and stress factors have been characterized and linked to autophagy initiation through the ULK1 complex and other mechanisms. Nutrient deprivation is the most common inducer of autophagy in mammalian cells, and it is mainly detected by the mechanistic target of rapamycin kinase complex 1 (mTORC1) - a protein complex that locates on lysosomes and acts as a master regulator of cell growth and metabolism (108). Free amino acids derived from lysosomes activate Rag GTPases anchored to the lysosomal membrane, leading to the recruitment and activation of mTOR, the core protein kinase in the mTORC1 complex. Active mTOR phosphorylates and inhibits ULK1, preventing the formation of the ULK1 complex and autophagosome biogenesis (109). ATG13, another component of the ULK1 complex, can also be inhibited by active mTOR (110). During 23 starvation, lack of free amino acids attenuates mTOR activity, leading to the dissociation of ULK1, which translocate to the ER membrane and nucleates the ULK1 complex at the autophagosome initiation site (86). Conversely, activated ULK1 also inhibits the mTOR by phosphorylation, ensuring the continuous induction of autophagy (111). Other than the direct modulation of ULK1 activity, mTORC1 also inhibits autophagy indirectly by phosphorylation and inhibition of transcriptional regulators, such as TFEB and the related family member Transcription factor E3 (TFE3), which will be discussed further in Section 1.4.1 (112,113). When nutrients are sufficient, TFEB and TFE3 are phosphorylated by mTORC1 and retained in their inactive form via binding with 14-3-3 proteins (114). Under nutrient deprivation, reduced activity of mTORC1 allows the nuclear translocation of TFEB/TFE3 and transcription of genes involved in lysosomal function and autophagy (115,116). Starvation can also promote autophagy by inhibiting a mTORC1-dependent process in which - m6A RNA methylation leads to the degradation of autophagy-related mRNA (117). Another important nutrient- and energy-sensing kinase involved in the regulation of autophagy is AMP-activated protein kinase (AMPK), which can respond to fuel change by sensing the change of ATP/ADP (118). During the state of nutrition abundance, ATP-bound AMPK is inactivated, while the decreasing ratio of ATP/ADP during starvation activates AMPK with ADP/AMP loading. Activated AMPK suppresses mTORC1 activity by phosphorylating RAPTOR, a molecule in the mTORC1 protein complex and can directly phosphorylate and increase the activity of key autophagy regulators, including ULK1, ATG9A and BECLIN1 (109). 24 Autophagy can be induced upon sensing various stress conditions, including ER and oxidative stress (66). ER stress can stimulate autophagy via UPR signaling. XBP1s can mediate the transcription of a series of adaptive genes to relieve ER stress, and one of its targets is BECLIN1 (119). Furthermore, IRE1 signaling can also induce free BECLIN1 by disrupting its complexing with BCL-2 protein (120). In contrast, ATF6 can induce autophagy by regulating the death-associated protein kinase 1, which phosphorylates BECLIN1 (121). Activating Transcription Factor 4 (ATF4), a downstream target of PERK, is a regulator of many autophagy-related genes, and its activation increases autophagy proteins, such as LC3, BECLIN1, ATG12, ATG16L and ATG3 (122). Interestingly, some reports also suggest a role for DDIT3/CHOP in the upregulation of several autophagy proteins (122), while it also potentiates ER stress-induced apoptosis (123). As mentioned in section 1.3.2, calcium is also an important autophagy regulator. Since ER stress often involves an increased leak or depletion of ER calcium storage, it can increase cytosolic calcium levels, which may contribute to ER stress-induced autophagy (124). One of the mechanisms by which autophagy can be activated during conditions of oxidative stress is via the Nuclear factor erythroid 2-related factor 2 (NRF2), which acts as a sensor of ROS accumulation (125). Under normal conditions, NRF2 binds to Kelch-like ECH-associated protein 1 and stays in the cytoplasm in the inactive form. However, when the levels of ROS increase, they translocate to the nucleus, where they activate several antioxidant responses and up-regulate multiple autophagy-related transcripts. 25 1.3.4. Roles of autophagy in pancreatic β-cells Several findings indicate that autophagy has roles in β-cell development and normal functions. In a study of porcine stem cells, the differentiation and formation of insulin-producing cells required the participation of autophagy (126). In mice with β-cell deletion of Atg7, the architecture of islets is disturbed, possibly due to the increase of β-cell apoptosis and the reduction of β-cell proliferation (127,128). Lack of Atg7 also leads to an accumulation of protein aggregates and the reduction of insulin secretion in mouse primary islets and further induces a loss of glucose homeostasis in vivo (127,128). Further, autophagy appears to affect insulin secretion by controlling the availability of proinsulin in the secretory pathway (129,130). In INS-1 β-cells, autophagy was found to degrade proinsulin and chemical inhibition of autophagy, or genetic knockout of Atg7/Atg5, increased granule proinsulin content and secretion (129). Autophagy plays important roles in the adaptation and quality control of β-cell mitochondria and ER (38). As discussed, the ER is essential for insulin production but sensitive to the stress associated with an excessive protein folding workload. Active autophagy helps clears accumulated misfolded proinsulin to reduce ER stress (131,132). In contrast, defective autophagy is associated with impaired ER adaptation and promotes the development of diabetes and obesity in animal models (133). Disruptions of mitochondrial homeostasis also impair insulin secretion and β-cell survival. Multiple factors, including genetic/transcriptional abnormalities and acute elevation of mitochondrial workload, increase the chances of mitochondrial damage (134). In conjunction with mitochondrial biogenesis, mitophagy acts as a 26 quality control mechanism by which β-cells can recycle damaged mitochondria to maintain both mass and function of their mitochondrial population under inflammation, as well as metabolic and oxidative stress (33,40,135,136). In agreement with its roles in organelle homeostasis and adaptation, studies have demonstrated that autophagy is important for β-cell survival and function under conditions of glucolipotoxicity, islet amyloid accumulation and inflammatory cytokine exposure. In different models, high levels of FFA induce autophagy in vitro and in vivo, and increased apoptosis occurs in these β-cells when autophagy is disrupted chemically or genetically, demonstrating the protective role of autophagy under gluco- or lipo-toxicity (137-142). Consistent with this, animals with β-cell Atg7 deficiency showed a loss of β-cell mass, impaired secretory function and induced apoptosis when they were fed a high fat diet, or a combined high fat and high glucose diet (128,140). Autophagy and mitophagy also protect β-cells from dysfunction and apoptosis caused by inflammatory cytokines, and several models with combined β-cell autophagy deficiency and hIAPP overexpression have demonstrated that autophagy helps counter the toxicity associated with accumulation of hIAPP oligomers (32,143-145). It is therefore noteworthy that prolonged glucolipotoxicity, inflammation and hIAPP accumulation all have been reported to disrupt β-cell autophagic flux in vitro, suggesting that negative feedback effects may ultimately limit autophagy-dependent β-cell adaptation (143,146,147). Loss of β-cell viability and function under these conditions may be related to unmitigated oxidative and ER stress, as discussed above (127,132). 27 Importantly, impaired β-cell autophagy has been described in both T1D and T2D. In a recent study of NOD mice and pancreas samples from human donors with T1D, Muralidharan et al. detected an accumulation of autophagosomes and a reduction in the number of β-cell autolysosomes, which likely reflects dysfunction of the lysosomal clearance step of autophagic flux (148). Abnormal double-membrane vesicles and a decrease in autophagy-related transcripts have been observed β-cell and islet samples from patients with T2D, which may be linked with the loss of β-cell mass (149). Interestingly, CLEC16A, a risk gene in T1D, acts as a mitophagy regulator, and its deficiency leads to the accumulation of defective mitochondrial populations in β-cells (78,136). These findings together suggest that the disruption of autophagy might contribute to the failure of β-cells during the development of diabetes. While much has been learned about the role of autophagy in β-cells, many questions remain to be answered. For instance, it is still controversial how starvation affects autophagy in β-cells. Some reports suggested nutrient deprivation induces β-cell autophagy, as in most other cell types (150,151). Interestingly, however, others have reported that starvation induces a selective lysosomal degradation of insulin granules, which then provides amino acids to activate mTOR and suppress β-cell macroautophagy (152). Questions also remain regarding the regulation and role of β-cells autophagy under specific types of stress. For instance, the role of autophagy is poorly understood in β-cells under hypoxia and in islet transplantation, and this is one of the primary questions that is addressed in this thesis. It is also an important unresolved question why β-cell autophagic flux fails under different forms of stress and in diabetes. The work in this thesis 28 will shed light on a possible role for dysregulation of TFEB and lysosomal function. 1.4 Regulation of autophagy and lysosomes by TFEB 1.4.1. TFEB and the MiTF family of transcription factors TFEB is a master transcriptional regulator of lysosomal biogenesis and function and plays a major role in the activation and modulation of autophagy (153). TFEB exerts its regulatory effects on lysosomal biology and autophagy through the so-called Coordinated Lysosomal Expression and Regulation (CLEAR) network, supported by TFE3 (154). Phosphorylation at multiple sites is crucial for controlling TFEB’s cellular localization and transcriptional activity (114,155,156). Specifically, changes in mTORC1 or CaN-mediated phosphorylation status link TFEB to the protective role of autophagy under nutrient deprivation and other stress conditions (40,112,157). TFEB belongs to the Microphthalmia-associated transcription factor (MiTF) family, which includes MiTF, TFEB, TFE3, and transcription factor EC (TFEC) (158). These proteins feature a basic region crucial for DNA binding and conserved helix–loop–helix-leucine zipper regions that facilitate homo- or hetero-dimerization among MiTF family proteins. This dimerization is essential for DNA recognition and transcription initiation (159). MiTF family members often have overlapping regulatory targets, as seen with TFEB and TFE3, in processes like innate immune response, stress adaptation, and whole-body energy metabolism (160-162). Unlike MiTF, TFEB and TFE3, which typically act as transcriptional activators, TFEC functions as a 29 repressor and counters the transcriptional regulation effects of TFE3 (163). Proteins that contribute to lysosomal clearance during autophagy, such as LAMPs, cathepsins, and V-ATPase, are all target genes of TFEB (115). The promoters of these lysosome-related genes commonly share a ‘GTCACGTGAC’ binding site for TFEB (CLEAR motif) (115,164). Pharmacological activation and overexpression of TFEB activates these genes to promote a number of biological processes, including lysosomal exocytosis, lysosomal biogenesis and autophagy (115,165-167). Specific examples include increases in the expression and activity of TFEB-targeted cathepsins (115) and induction of LAMPs to support lysosomal function and autophagy in various tissues and conditions (165,168,169). TFEB also affects V-ATPase-dependent lysosomal acidification as starvation-induced inhibition of mTORC1 induces TFEB-dependent V-ATPase expression (170). In summary, TFEB supports lysosomal autophagy by controlling the transcription of different CLEAR genes that regulate autophagosome formation, as well as lysosomal hydrolytic activity, membrane stability and lumen acidification (Figure 1.2). This emphasizes the importance of understanding the external factors and cellular mechanisms that regulate TFEB activity and levels, including in β-cells. 30 1.4.2. Mechanisms of TFEB regulation TFEB achieves its regulatory function when it enters the nucleus and binds to CLEAR motifs in the promoter of its target genes. Therefore, the subcellular localization of TFEB is closely associated with its functional activities. Various post-translational modifications, such as phosphorylation and acetylation, regulate the subcellular localization, binding targets, and overall Figure 1.2 – TFEB-dependent regulation of autophagy and lysosomal homeostasis Schematic summarizing how dephosphorylation-induced activation of TFEB, via nutrient-dependent inhibition of mTORC1 and/or activation of CaN, supports autophagic flux via the expression of genes related to autophagy and lysosomal homeostasis. See text for details. Created with BioRender.com. 31 activity of TFEB (156). Among these modifications, phosphorylation resulting in TFEB’s cytoplasmic retention and inactivation is the most well-studied (112,157). Specific serine residues, including S122, S134, S138, S142, S211 and S467 regulate TFEB’s protein-protein interaction and cytoplasmic localization (156). Mutation of S142, S138 or S211 to alanine results in constitutively nuclear localization and activation of TFEB (114). Phosphorylation at S142 and S138 may promote TFEB’s nuclear export efficiency (114), and the phosphorylation at S211 has been suggested to determine TFEB’s binding and sequestration by 14-3-3 scaffolding proteins in the cytoplasm (112,157). Multiple kinases, including mTORC1, glycogen synthase kinase 3β (GSK3β), the extracellular signal-regulated kinase 1/2 (ERK1/2), and protein kinase B (AKT), have been shown to phosphorylate TFEB directly and regulate its activity (156,168). Most notably, mTORC1 links the sensing of nutrients with the activity of TFEB by phosphorylation at S211, S142, and S122 under conditions of nutrient abundance (166). It has also been suggested that TFEB phosphorylation by GSK3β or ERK1/2 is at least partially dependent on mTORC1, further indicating the essential role of mTORC1 in regulating TFEB activity (114). Kinase inhibitors effectively regulate TFEB; mTORC1, GSK3β, and AKT inhibitors have been found to promote TFEB nuclear localization and transcriptional activity (155,168,171). Pharmacological manipulation of these kinases holds potential therapeutic value, as enhancing lysosomal performance and autophagy can help treat various diseases. An inhibitor of mTORC1 and mTORC2, torin-1, has been reported to rescue TFEB-mediated lysosomal biogenesis and 32 autophagy in Thiel-Behnke corneal dystrophy (172). Another mTORC1 inhibitor, rapamycin, is used as an immunosuppressant in islet transplantation and has been suggested to have beneficial effects in tuberous sclerosis via TFEB-dependent lysosomal biogenesis and autophagy (173). However, in some earlier reports, rapamycin did not affect TFEB phosphorylation and subcellular localization (155), indicating its effects on TFEB might be complex and cell type-dependent. Trehalose promotes cellular clearance in neurodegenerative storage disease by inhibiting AKT-dependent TFEB phosphorylation at S467, which induces TFEB nuclear translocation and activity (168). Together, these findings suggest that TFEB-regulating kinases could be potential targets for pharmacological development. TFEB’s phosphorylation status is also regulated by the calcium-activated phosphatase CaN (104). Cytosolic calcium signals, which regulate TFEB activity, can arise via calcium release from various intracellular organelles. For instance, MCOLN1-mediated calcium release from lysosomes plays a crucial role in CaN-dependent dephosphorylation and activation of TFEB (102,103). In addition, inositol triphosphate-triggered ER calcium release promoted TFEB-mediated lysosomal biogenesis in HeLa cells (174). Therefore, calcium-mediated and CaN-dependent activation of TFEB represents additional avenues for identifying pharmacological TFEB agonists for therapeutic purposes. Notably, a newly identified small molecule, MSL-7, activates TFEB independently of mTORC1 via CaN activation and has been suggested to enhance lysosomal clearance capacity and improve the metabolic profiles in genetic- or diet-induced obese mice (175). Interestingly, Exendin-4, a glucagon-like peptide 1 receptor (GLP1R) 33 agonist used to control hyperglycemia in diabetes, has been found to activate TFEB through lysosomal calcium release and CaN stimulation (176). The activity of TFEB is also partly dependent on other transcriptional regulators that target similar genes. As mentioned, TFE3 cooperates with TFEB to enhance autophagy and lysosomal biogenesis in stress responses, immune responses and metabolic regulation (160-162). Evidence also suggests that Peroxisome proliferator-activated receptor γ coactivator-1 alpha (PGC-1α), a target of TFEB and a transcriptional regulator of mitochondria biogenesis, can promote autophagy via activation of TFEB through a positive feedback loop (177,178). On the other hand, the transcriptional regulator zinc finger with KRAB and SCAN domains 3 (ZKSCAN3) targets many of the same lysosome-related genes as TFEB but functions as a transcriptional repressor that opposes TFEB’s activity (179). In summary, TFEB activity is regulated by post-translational modifications, especially phosphorylation on multiple serine residues, and key kinases and phosphatases that modulate TFEB phosphorylation status have been suggested for therapeutic uses. In addition, TFEB often coordinates with other transcriptional regulators to achieve fine-tuning in gene transcription. 1.4.3. Roles for TFEB in cellular stress-adaptation and metabolism Since it was identified as a master transcriptional regulator of cellular clearance mechanisms, numerous roles have been identified for TFEB in stress adaptation and metabolic regulation of various tissues and organs. Initially, TFEB was studied mostly for its involvement in nutrient 34 sensing and starvation-induced autophagy. It was later found that TFEB and TFE3 can also be activated in response to ER stress and not only induce transcription of genes related to lysosomes and autophagy but also the expression of ER stress regulators such as ATF6b, IRE1 and ATF4 (161). Recent findings by Cinque et al. also linked TFEB and TFE3 to the regulation of ER homeostasis via transcriptional induction of the ER-phagy receptor FAM134B (180). Further highlighting their general functions in cellular adaptation to stress, it was reported that oxidative stress can activate TFEB and TFE3 via Protein Phosphatase 2A-dependent, including when mTORC-1 is active (181). Roles of TFEB in homeostasis and metabolism have been studied in different tissues, such as bone, liver, muscle, and kidney. Using overexpression and knockout models, Ferron et al. examined the roles of TFEB in bone and found that it is critical for osteoclast function and lysosomal collagen degradation during bone matrix resorption (182). Specifically, RANKL, a key cytokine in osteoclast differentiation, recruits PKCβ to phosphorylate serine residues S461/S462 or S465/S466 of TFEB, which promotes stabilization of the TFEB protein and increases its transcriptional activity. Other studies demonstrated that TFEB-mediated ER-phagy in chondrocytes is important during skeletal growth in medaka fish models (180). TFEB has been implicated in several aspects of liver homeostasis (183). TFEB-mediated autophagy is important for alleviating alcohol-induced liver injury in both acute and chronic models (184,185). Chao et al. found that TFEB levels were compromised in alcohol-treated mice and patients with alcoholic hepatitis and further demonstrated that TFEB overexpression 35 protected liver cells under alcohol treatment while suppression of TFEB increased the risk of liver injury (186). TFEB-mediated autophagy also protects hepatocytes against alpha-1 antitrypsin deficiency and diclofenac-induced toxicity (187,188). Of note, TFEB has been linked to liver metabolic functions and regulates gene transcription related to lipid metabolism via directly targeting PGC-1α, and TFEB knockout mouse models exhibit impaired lipid oxidation under starvation conditions (167). Lipid droplet accumulation is observed in mice with liver-specific deletion of TFEB on a high-fat diet, further indicating that TFEB plays an important role in regulating hepatic lipid metabolism. Additionally, overexpression of TFEB rescues lipid storage in wild-type but not Atg7-deficient mice, indicating that TFEB regulates liver lipid homeostasis through autophagic pathways. Other studies revealed that TFEB and TFE3 play cooperative and interchangeable roles in regulating liver and whole-body metabolism (162). TFEB also plays a significant role in muscle energy metabolism (162,189,190). Overexpression of TFEB increases the levels of multiple mitochondrial regulators and induces mitochondrial biogenesis (189,190). Conversely, loss of TFEB is associated with abnormal mitochondrial dynamics and function, leading to exercise-deficient phenotypes in vivo (189). Interestingly, this study suggested that the beneficial metabolic effects of TFEB in muscle cells are independent of PGC-1α (189), whereas others have reported that exercise-induced TFEB expression and nuclear translocation was attenuated in PGC-1α KO mice (190). Irrespective of the mechanistic details, and possible tissue-specific differences, these findings all support the important role of TFEB in lipid and glucose metabolism. 36 TFEB has been associated with many human diseases. Examples include neurodegenerative and kidney diseases, where its canonical functions in cellular clearance have protective effects. For instance, overexpression of PGC-1α improved neurological function in a model of Huntington’s disease by inducing TFEB expression and TFEB-dependent clearance of mutant huntingtin protein aggregates (191), and AKT-dependent TFEB activation and lysosomal clearance is crucial for protecting against neurodegenerative storage diseases (168). In kidney cells affected by cystinosis, a type of lysosomal storage disease, activation of TFEB can improve cell physiology and viability, including by effects on LAMP2A-dependent CMA (192). In summary, various models from different tissues and disease contexts have highlighted critical roles for TFEB in metabolism and stress adaptation. Given that pancreatic β-cells are highly metabolic endocrine cells, this raises the important question of how TFEB is involved in their function, adaptation and survival. 1.4.4. TFEB in the pancreatic β-cell A small but growing body of work suggests that TFEB is important in both β-cell physiology and pathophysiology. A recent study demonstrated that mTORC1 regulates insulin release in an autophagy-dependent way in response to different nutrient levels, possibly through controlling the phosphorylation and activity of TFEB (193). Of note, Ballabio’s group recently made the discovery that TFEB and TFE3 act as nutrient-sensitive regulators of insulin transcription; specifically, they help suppress insulin expression under nutrient-depleted conditions (194). 37 In the context of β-cell stress, it has been demonstrated that chronic glucolipotoxicity impairs β-cell autophagic flux and lysosomal homeostasis in vitro (146), and cathepsin release from defective lysosomes contributes to glucolipotoxicity-induced β-cell death (195). Importantly, treatment with the GLP1R agonist exendin-4 improved both lysosomal homeostasis and autophagic flux in these cells; this depended on intracellular calcium release and CaN-dependent activation of TFEB and contributed to the anti-apoptotic effects of exendin-4 (176). Recent work by Park et al. demonstrated that lysosomal calcium release activates TFEB and stimulates β-cell mitophagy (40). Investigation of mice with β-cell Tfeb knockout further indicated that this protects β-cells from metabolic and oxidative stress under HFD feeding (40). Combined with the finding that GLP1R agonists can signal β-cell survival through TFEB, this suggests that TFEB is a potential drug target for β-cell protection and diabetes treatment. In support of this, Lim et al. identified a small molecule, MSL-7, that activates TFEB through CaN and found that MSL-7 accelerates cellular lipid clearance in vitro and improves the metabolic profiles in mice with genetic and diet-induced obesity (175). More specific to β-cells, autophagy activation by MSL-7 stimulates TFEB-dependent clearance of toxic hIAPP oligomers in vitro and in vivo, demonstrating beneficial effects of TFEB activation (196). Despite the increasing number of studies, relatively little is known about the roles of TFEB in β-cell function and failure. As part of this thesis, I investigate whether dysregulation of endogenous TFEB contributes to β-cell death and failure under stress and use β-cell-specific gene deletion to clarify the importance of TFEB for in vivo β-cell adaptation to metabolic stress. 38 1.5 Thesis Overview As a cellular recycling mechanism, autophagy is important for β-cell integrity and has been recognized as an adaptive pro-survival mechanism for β-cells under various forms of stress. However, it remains an important question if autophagy can promote β-cell survival under hypoxia or the stresses associated with pancreatic islet transplantation. The final step of autophagic flux relies on functional lysosomes, which are also increasingly recognized as hubs for nutrient sensing and signaling. TFEB plays an important role in regulating the transcription of genes that promote lysosomal function, as well as the biogenesis of both lysosomes and autophagosomes. Increasing evidence suggests lysosomes are compromised in β-cells under stress, but the mechanisms of this failure remain to be determined. Finally, better understanding the roles of TFEB in β-cells may provide important new insights into β-cell physiology and pathophysiology, to ultimately help identify new targets for future clinical research and pharmaceutical development for prevention or treatment of diabetes. The research in this thesis was done with the goals of shedding new light on the importance of autophagy for β-cell function and survival in vitro and in vivo, and provide a focused investigation of TFEB, and TFEB-dependent lysosomal homeostasis, in β-cells. Overall hypothesis: Disruption of TFEB and lysosomal clearance contribute to failure of autophagic flux, and exacerbates β-cell dysfunction and death under diabetes-related stress. 39 Objective 1: Determine the importance of autophagy for β-cell function and survival under hypoxia and transplantation-induced stress. This objective is addressed in Chapter 3. Objective 2: Determine if dysregulation of TFEB and lysosomal homeostasis are fundamental events in stressed β-cells. This objective is primarily addressed in Chapter 4. Objective 3: Determine the importance of TFEB for β-cell function and survival under diet-induced stress. This objective is primarily addressed in Chapter 5. 40 Chapter 2: Materials and Methods 2.1 Mice All mice (Table 2.1) were housed and bred with a 12-hour light-dark cycles at the British Columbia Children’s Hospital Research Institute (BCCHR) Animal Care Facility or at the Centre for Disease Modelling on the UBC Vancouver Campus. Table 2.1 Mouse strains Common name used Strain name Strain Number Original Source B6 C57BL/6J #000664 The Jackson Laboratory Ins1Cre B6(Cg)-Ins1tm1.1(cre)Thor/J #026801 The Jackson Laboratory CAG-RFP-EGFP-LC3 C57BL/6-Tg(CAG-RFP/EGFP/Map1lc3b)1Hill/J #027139 The Jackson Laboratory mTmG Gt(ROSA)26Sortm4(ACTB-tdTomato,-EGFP)Luo/J #007576 The Jackson Laboratory Atg5flox/flox B6.129S-Atg5tm1Myok RBRC02975 RIKEN BRC, through National Bio-Resource Project of the MEXT, Japan Tfebflox/flox N/A Dr A Ballabio, TIGEM Italy Atg5βKO N/A In-house breeding TfebβKO N/A In-house breeding 41 The following animals were originally purchased from The Jackson Laboratory (Bar Harbor, ME, USA) and then further bred in-house: C57BL/6J (#000664), B6(Cg)-Ins1tm1.1(cre)Thor/J (common name: Ins1Cre, #026801), C57BL/6-Tg(CAG-RFP/EGFP/Map1lc3b)1Hill/J (common name: CAG-RFP-EGFP-LC3, #027139) and Gt(ROSA)26Sortm4(ACTB-tdTomato,-EGFP)Luo/J (common name: mTmG, #007576). B6.129S-Atg5tm1Myok (common name: Atg5flox/flox) and Tfebflox/flox mice were kindly gifted from Dr. Noboru Mizushima (via RIKEN) (197) and Dr Andrea Ballabio (155), respectively. Beta cell-specific Atg5 knockout mice (Atg5βKO) were established by crossing female Atg5flox/flox and male Ins1Cre/wt mice. Male heterozygous knockout (Atg5flox/wt;Ins1Cre/wt) offspring were further crossed with female Atg5flox/flox mice to generate Atg5flox/flox;Ins1Cre/wt homozygous knockout (Atg5βKO) and Atg5flox/flox;Ins1wt/wt control (Atg5βWT) mice, which were bred with each other to maintain the colony. Beta cell-specific Tfeb knockout mice (TfebβKO) were established by crossing female Tfebflox/flox and male Ins1Cre/wt mice. Male heterozygous knockout (Tfebflox/wt;Ins1Cre/wt) offspring were further crossed with female Tfebflox/flox mice to generate Tfebflox/flox;Ins1Cre/wt homozygous knockout (TfebβKO) and Tfebflox/flox;Ins1wt/wt control (TfebβWT) mice, which were bred with each other to maintain the colony. 2.1.1 Mouse diets To examine the role of β-cell Tfeb under diabetes-relevant stress in vivo, a subset of TfebβKO and TfebβWT mice were fed a high fat diet known to induce obesity and glucose intolerance 42 phenotypes. Specifically, 8–10-week-old mice were fed either a high fat (HFD, 60%KcalFat, 20%KcalProtein, 20%KcalCarbohydrate; #D12492, Research Diets, New Brunswick, NJ, USA) or a control diet (CD, 10%KcalFat, 20%KcalProtein, 70%KcalCarbohydrate; #D12450B, Research Diets) for up to 20 weeks, as indicated. All other mice were fed a chow diet (18%KcalFat, 24%KcalProtein, 58%KcalCarbohydrate; #2918, Envigo, Indianapolis, IN, USA). Diets are summarized in Table 2.2. Table 2.2 Mouse diets Common Name used Diet Description Cat # Source High fat diet (HFD) Rodent Diet With 60 kcal% Fat D12492 Research Diets Control diet (CD) Rodent Diet With 10 kcal% Fat D12450B Research Diets Chow diet Irradiated Teklad Global 18% Protein Rodent Diet 2918 Envigo 2.2 Streptozotocin (STZ) injection models A single high-dose STZ injection induces rapid loss of β-cell mass and was used to generate diabetic syngeneic recipients for marginal mass islet transplantation surgeries in Chapter 3. A multiple low dose (MLD) STZ injection model (198,199), was used to study the role of Tfeb under more gradual conditions of in vivo inflammation- and ER-stress in Chapter 5. Single high dose STZ model - Male Atg5flox/flox mice of 8-12 weeks of age were injected i.p. with a single high STZ dose (175 mg/kg; #S0130, Sigma-Aldrich, St Louis, MO, USA) or acetate 43 buffer (control). Body weight and blood glucose were monitored before and every day after the STZ injection. Four days after injection, male mice showing hyperglycemia (two consecutive days with blood glucose >20 mM) were used as islet transplant recipients. Multiple low dose STZ model - 12–15-week-old littermate male TfebβWT and TfebβKO mice were injected i.p. with STZ (50 mg/kg) or acetate buffer (control) for 5 consecutive days. Body weight and blood glucose were monitored before the first STZ injection and every 2 days after the last injection. Glucose tolerance was assessed two weeks after the first injection, or at the time animals became hyperglycemic. 2.3 Islet transplantation A marginal mass syngeneic islets transplantation model was used to study the role of autophagy under post-transplantation stresses that occur independent of autoimmune graft rejection (200). Islet donors and recipients - Female Atg5flox/flox mice of 10-14 weeks of age were used as transplant donors. Two weeks before islet isolation, double-stranded adeno-associated viruses (dsAAV) encoding Rat Insulin Promoter-1 (RIP1)-driven Cre recombinase (dsAAV6-RIP1-Cre) or empty dsAAV control viruses (dsAAV6-Empty) were delivered into the pancreas through an intra-ductal injection to create Atg5 knockout and control donor islets, respectively (201,202). Prior to donor islet isolation, intraperitoneal glucose tolerance tests (IPGTT) were performed on 44 the mice to ensure similar glucose tolerance between the dsAAV6-Empty and dsAAV6-RIP1-Cre injected groups. After isolation, Atg5 deletion was verified by qPCR and glucose-stimulated Ca2+ and insulin secretion responses were compared in aliquots of control and Atg5 KO donor islets. We pooled islets from littermate donor mice before preparing equal-sized grafts. Male Atg5flox/flox mice of 8-12 weeks of age were administered a single high dose STZ and used as islet transplant recipients if they developed hyperglycemia (blood glucose >20 mM for 2 consecutive days). The amount of STZ was established in prior tests using mice of the same sex and age. Islet graft preparation and transplantation surgery – Four days after STZ injection, the diabetic mice were transplanted with a marginal mass of islets under the kidney capsule, essentially as described (203). Donor islets were isolated one day prior and hand-picked for equal size grafts on the morning of islet transplantation. Each recipient received a graft containing 300-350 islets under the kidney capsule. The marginal number of islets was established in prior tests on mice of the same sex and age. The recipients were monitored for body weight and blood glucose every day for 2 weeks post-surgery, and IPGTTs were performed 2 weeks after the transplant, or at any earlier time if the recipients became hyperglycemic due to graft failure. 2.4 Body weight and glucose metabolism Non-fasting body weight and glucose measurements were recorded in the late morning (10 am-12 pm), using OneTouch Ultra® or OneTouch Verio Flex® blood glucose meters and test strips 45 (LifeScan Inc., Malvern, PA, USA). During the dietary intervention, the non-fasting body weight and glucose were monitored weekly. Glucose tolerance was assessed by IPGTTs. Prior to the IPGTT, mice were fasted for 6 hours, except mice that had received multiple-low dose STZ injections, which were fasted overnight (~14 hours). The IPGTTs were performed in the afternoon, starting at 3-4 pm. Unless otherwise indicated, mice were injected with 2 g glucose/kg body weight. Some mice on late-stage HFD were injected with 0.5 g glucose/kg of body weight to avoid severe hyperglycemia and saturating the glucose meter. Blood glucose levels were measured at 0, 15, 30, 60, 90 and 120 min after glucose injection. Insulin tolerance tests (ITT) were performed in the afternoon, starting at 3-4 pm after a 3-hour fast. The shorter fasting time was to prevent the mice from becoming hypoglycemic during the tests. Unless otherwise specified, the mice were injected i.p. with 0.75 U/kg insulin (Novolin, DIN 02024233, Novo Nordisk, Bagsværd, Denmark). Some mice on late-stage HFD were injected with 1 U/kg insulin to account for their diet-induced insulin resistance. Blood glucose levels were measured at 0, 15, 30, 60, 90 and 120 min after the insulin injection. 2.5 Islet isolation, culture, transduction and dispersion Islet isolation - After anesthesia and euthanasia, mouse pancreatic islets were isolated by a collagenase digestion/filtration method as described before (204,205). In brief, collagenase (1000 U/mL, #C7657; Sigma-Aldrich) in Hank’s solution (137 mM NaCl, 5.4 mM KCl, 4.2 mM 46 NaH2PO4, 4.1 mM KH2PO4, 10 mM HEPES, 1 mM MgCl2, and 5 mM glucose) was administered via the pancreatic duct to distend the pancreas. The pancreas was removed and incubated in collagenase solution and washed with Hank’s solution containing 1 mM CaCl2. After centrifugation at 2000g, the islets were re-suspended in Roswell Park Memorial Institute 1640 media (RPMI, #11875-093, Thermo Fisher Scientific, Waltham, MA, USA) and picked by hand for purification. Unless otherwise indicated, mouse islets were cultured at 37°C in 5% CO2 in RPMI completed with 10% fetal bovine serum (FBS, #12483-020, Thermo Fisher Scientific) and 2% Penicillin-Streptomycin (#15140-122, Thermo Fisher Scientific). Islet dispersion - For single cell studies, mouse islets were dispersed with 0.01% trypsin (#25300-054, Thermo Fisher Scientific) onto glass coverslips (#16004-310, VWR international, LLC, Radnor, PA, USA) or ibidi 8-well chamber-slides with polymer (#80806/#80826, ibidi, Munich, Germany) or glass bottom (#80827, ibidi) [2, 3]. Islet cells were cultured at 37°C in 5% CO2 in complete RPMI 1640 media for 1-2 days after dispersion before staining or imaging. Adenoviral transduction – Intact islets were transduced within 2-3 hours of isolation while dispersed cells were allowed to recover for 1-2 days in ibidi chamber slides before transduction. The dispersed cells or intact islets were incubated in completed RPMI 1640 media containing adenovirus at the indicated multiplicity of infection (MOI), which represents the number of viral particles added per cell. For intact islets, medium size islets were picked and the cell number 47 estimated as 1,000 cells per islet on average. Following overnight transduction, the media was replaced by fresh RPMI without virus and the cells/islets were cultured for additional 3-4 days before being used for imaging or RNA analysis. 2.6 MIN6 cell culture and transfection MIN6 cell culture - MIN6 cells were cultured at 37°C / 5% CO2 in Dulbecco’s modified Eagle’s medium (DMEM, #11995-065, Thermo Fisher Scientific) completed with 10% FBS and 2% Penicillin-Streptomycin. Culture media was replaced at least every 48 hours, and the cells were passaged when at 70-80% confluence. MIN6 cell transfection and adenoviral transduction - MIN6 cells were transfected using a Lipofectamine 2000 kit (#11668019, Thermo Fisher Scientific) at a confluence of 60-70%. Cells were pre-cultured in optimized Minimal Essential Medium (OPTI-MEM, #31985070, Thermo Fisher Scientific) for 20-30 min before the transfection and cultured in incomplete DMEM with no FBS and Penicillin-Streptomycin. Transfected cells were used for experiments 24-48 hours after transfection. 2.7 Human islets Human islets were provided by Clinical Islet Laboratory through Alberta Islet Distribution Program. The islets were purified by hand-picking and then dispersed onto ibidi 8-well chamber 48 slides (#80806/#80826, ibidi) for microcopy. The dispersed islet cells were cultured in CMRL media (#11530037, Thermo Fisher Scientific) completed with 10% FBS, 2% Penicillin-Streptomycin, at 22°C in 5% CO2 for 1-2 days before experiments. 2.8 Hypoxic culture Hypoxic culture (1% O2, 5% CO2 and 94% N2) of intact mouse islets, as well as dispersed mouse and human islet cells to be used for immunofluorescent imaging and lysosomal function assays, was done in an incubator sub-chamber (C-Chamber) with CO2 and O2 levels maintained by ProCO2 120 and ProOx 110 controllers (Biospherix, Parish NY, USA). After hypoxic culture, whole islets were immediately processed for qPCR or Western blot analysis. Hypoxic islet cells were immediately fixed in paraformaldehyde (PFA, #AC41678, Fisher Scientific) - 0.5% PFA was used for Magic Red Cathepsin B activity assays and 4% PFA for immunostaining. In some experiments, hypoxic signaling pathways were chemically induced in dispersed mouse islets by treatment with 200 µM CoCl2 for 5-8 hours, as indicated. For imaging-based analysis of cell survival and autophagic flux (detailed below), dispersed islet cells were cultured at 37oC under hypoxia (1% O2, 5% CO2 and 94% N2) or normoxia (20% O2, 5% CO2 and 75% N2) in a Tokai Hit INUBTFP-WSKM microscope stage-top incubator. 2.9 In vitro glucose-stimulated insulin secretion To assess glucose stimulated insulin secretion, 15 size-matched islets were placed into low-49 retention microcentrifuge tubes and cultured in Krebs-ringer buffer (KRB) with 3 mM glucose for 1 hour prior to the experiments. The islets were then transferred to 300 μl fresh KRB containing 3 mM glucose, 15 mM glucose, and finally a combination of 3 mM glucose and 30 mM KCl, each for 1 hour sequentially. The supernatants from each stage were collected and assayed for secreted insulin using a Mouse Ultrasensitive Insulin ELISA (#80-INSMSU-E10, ALPCO, Salem, NH, USA) following the manufacturer’s instructions. 2.10 Calcium imaging Cytosolic Ca2+ was imaged in intact islets using Fura-2 AM (#F1221, Thermo Fisher Scientific) (204). After isolation, 10-15 islets were picked and seeded onto a 25 mm glass coverslip (#16004-310, VWR) placed in 35 mm culture dishes (#353001, Corning Incorporated, Corning, NY, USA). Islets were incubated for 3 days to attach before imaging. Islets were loaded with 5 µM Fura-2 for 30 min and then perifused with a modified Ringer’s Buffer (5.5 mM KCl, 2 mM CaCl2, 1 mM MgCl2, 20 mM HEPES, 144 mM NaCl2) containing 3 mM glucose for at least 30 min to reach a stable baseline. Glucose responsiveness was tested by exposing the islets to a ramp of Ringer’s containing 3 mM, 5 mM, 10 mM and 15 mM glucose, followed by 30 mM KCl in 3 mM glucose. Fura-2 was excited at 340 nm and 380 nm and the emission were collected at 502-538 nm every 10 seconds in wide-field settings on a Leica DMI6000 inverted microscope (Leica, Wetzlar, Germany) equipped with a Leica HCX Plan FLUOTAR L 10x objective (#506505), Fura-2 filter cube (#79001, Chroma, Bellows Falls, VT, USA). 50 2.11 Seahorse oxygen consumption measurements & analysis Whole islets were cultured in completed RPMI media for 16-32 hours before 70-80 islets were loaded into Seahorse XF24 islet capture microplates (#101122-100, Agilent, Santa Clara, CA, US) containing Seahorse XF RPMI Media pH 7.4 (#103576, Agilent) with 1% FBS, 3 mM glucose (#G7528, Sigma-Aldrich), 2 mM glutamine (#25030081, Thermo Fisher Scientific) and 2 mM Sodium Pyruvate (#11360070, Thermo Fisher Scientific) and incubated at 37oC in a non-CO2 incubator for 60 minutes. Oxygen consumption rate (OCR) was then measured using a Seahorse XFe24 Analyzer (Agilent). Injection ports were loaded with the following compounds in Seahorse XF RPMI Media pH 7.4 with 3 mM glucose, 2 mM glutamine and 2 mM sodium pyruvate: A) glucose (15 mM), B) oligomycin (2 μM, #O4876, Sigma-Aldrich), C) FCCP (2 μM, # C2920, Sigma-Aldrich), D) antimycin A1 and rotenone (both 1.5 μM, #A8674 & #R8875, Sigma-Aldrich). OCR measurements were normalized to the total DNA content of the assayed islets of each well, as determined by Qubit HS DNA Assay (Q32854, Thermo Fisher Scientific) using a QubitTM Flurometer (Q32857, Invitrogen) after lysis in RIPA buffer. 2.12 Cell death assays For quantification of cell death, mouse islet cells were dispersed and seeded onto 25 mm glass coverslips or ibidi 8-well chamber-slides (#80806/#80826). The islet cells were pre-loaded for 30 min with 0.05 μg/ml Hoechst 33342 (#H3570, Thermo Fisher Scientific) and 5 μM propidium iodide (PI, #P1304MP, Thermo Fisher Scientific). To quantify cell death, the cells were imaged 51 during culture in normoxia or hypoxia within a Tokai Hit INUBTFP-WSKM Stage-top incubator at 37℃ and images were taken at different time intervals using the wide-field setting on a Leica DMI6000 inverted microscope using a Leica HCX Plan FLUOTAR L 20x objective (#506242), DAPI ET and DsRed ET filter cubes (#49005, Chroma). The percentage of dead cells (PI+ / Hoechst+) was quantified using ImageJ. 2.13 RNA extraction and RT-qPCR analysis Total RNA was extracted from mouse islets using RNeasy Micro Kit (#74104, Qiagen, Hilden, Germany) according to the manufacturer’s guidelines. RNA yield was measured on a NanoDropTM 2000c (#ND2000CLAPTOP, Thermo Fisher Scientific) spectrophotometers. cDNAs were generated through reverse transcription by qScript cDNA synthesis kits (#95047, Quanta Bioscience, Gaithersburg, MD, USA). Quantitative PCR (qPCR) was performed using PerfeCTa SYBR Green SuperMix (#95053, Quanta Bioscience) in the ViiA7 Real-Time PCR System (Thermo Fisher/Applied Biosystems) using primers listed in Table 2.3. Table 2.3 Primers used for SYBR Green qPCR Gene Target Forward Primer (5` to 3`) Reverse Primer (5` to 3`) Actb GATCTGGAAACATTGGAGAGG TGCGGCCTCTGAACATCTCAT Aldoa AGGAACCAATGGCGAGACAAC AAGAGAGATTCACTGGCTGCGG Atf3 GCTGCCAAGTGTCGAAACAAG CAGTTTTCCAATGGCTTCAGG Atg5 CCTCGGTTTGGCTTTGGTTG CGTCAAATAGCTGACTCTTGG 52 Atp5e CAGGCTGGACTCAGCTACATC GTTCGCTTTGAACTCGGTCTT Atp6v1h CCAAGATGGACATTCGAGGTG CACTTTGTTGGCACGAACTTC Becn1 ATGGAGGGGTCTAAGGCGTC TGGGCTGTGGTAAGTAATGGA Bnip3 GCTCCCAGACACCACAAGATA TGAGAGTAGCTGTGCGCTTC Cox4l1 ATTGGCAAGAGAGCCATTBTCTAC CACGCCGATCAGCGTAAGT Ctsb TCCTTGATCCTTCTTTCTTGCC ACAGTGCCACACAGCTTCTTC Ctsd GCTTCCGGTCTTTGACAACCT CACCAAGCATTAGTTCTCCTCC Ddit3 (Chop) CTGCCTTTCACCTTGGAGAC CGTTTCCTGGGGATGAGATA Gapdh AGGTCGGTGTGAACGGATTTG TGTAGACCATGTAGTTGAGGTCA Hmox1 CAGAAGGGTCAGGTGTCCA CTTCCAGGGCCGTGTAGAT Hspa5 (Bip) TCATCGGACGCACTTGGAA CAACCACCTTGAATGGCAAGA Ern1(Ire1) CCGAGCCATGAGAAACAAGAA GGGAAGCGGGAAGTGAAGTAG Lamp1 CAGCACTCTTTGAGGTGAAAAAC ACGATCTGAGAACCATTCGCA Lamp2 ATATGTGACAACAAAGAGCAGGT TCGCAATTAGGTAAGCAATCACT Map1lc3a CGTCCTGGACAAGACCAAGT CTCGTCTTTCTCCTGCTCGT Map1lc3b AGCAGCATCCAACCAAAATC CTGTGTCCGTTCACCAACAG Mcoln1 CTGACCCCCAATCCTGGGTAT GGCCCGGAACTTGTCACAT Sdhb AATTTGCCATTTACCGATGGG AGCATCCAACACCATAGGTCC Tfeb GCTCCAACCCCGAGAAAGAG CAGCGTGTTAGGCATCTGC Tfe3 TGCGTCAGCAGCTTATGAGG AGACACGCCAATCACAGAGAT Xbp1s GAGTCCGCAGCAGGTG GTGTCAGAGTCCATGGGA Xbp1t TGGCCGGGTCTGCTGAGTCCG GTCCATGGGAAGATGTTCTGG 53 2.14 Western blotting Isolated islets were transferred to a 1.5-ml microcentrifuge tube. After brief centrifugation, culture medium was discarded and 50 μl of RIPA lysis buffer (150 mM NaCl, 50 mM Tris-HCl (pH 8.0), 0.1% Triton X-100, 0.5% sodium deoxycholate, 0.1% SDS, 1 x Halt Protease and Phosphatase inhibitor (#78440, Thermo Fisher Scientific) was added. The cells were lysed by sonication (Misonix S-4000) with 40% power output, 6-10 cycles of 10-sec-ON – 20-sec-OFF. Protein lysates were then separated by 4-20% SDS-PAGE (#4561095, Bio-Rad, Hercules, CA, USA) and transferred to a PVDF membrane (#1620177, Bio-Rad). The membrane was blocked with 5% non-fat dry milk in TBST and incubated overnight with primary antibody at 4°C. After three washes with TBST, the membrane was incubated with HRP-conjugated secondary antibody at room temperature for 1 hour. After three washes with TBST, the membrane was then incubated with SuperSignal West Pico or Femto ECL Substrates (#34579/#34094, Thermo Fisher Scientific) and the chemiluminescent signals were detected by X-ray film. Table 2.4 Antibodies used for Westen Blot Antibody Source Cat. No Dilution Βeta Actin Antibody Novus® NB600-501 1 to 10000 Atg5 Rabbit Antibody Cell Signaling® #12994 1 to 1000 Anti-Cre Antibody Novagen® 60950-3 1 to 10000 LC3 Antibody Novus® NB100-2220 1 to 900 Cleaved Caspase-3 Antibody Cell Signaling® #9661 1 to 1000 54 TFEB Antibody Bethyl® A303-673A 1 to 10000 Anti-mouse IgG HRP-linked Antibody Cell Signaling® #7076 1 to 3000 Anti-Rabbit IgG HRP-linked Antibody Cell Signaling® #7074 1 to 3000 2.15 Lysosomal cathepsin B activity Lysosomal cathepsin B activity was examined in dispersed mouse islet cells using the Magic RedTM assay Kit (#ICT937, Bio-Rad), following the manufacturer’s guidelines. For this assay, islet cells were fixed with 0.5% PFA for 30 min before staining. Images were taken in wide-field settings using a Leica DMI inverted microscope with a Leica HCX Plan FLUOTAR L 10x objective (#506505) and DsRed ET filter cube (#49005, Chroma). Magic Red fluorescence intensities were quantified in ImageJ. 2.16 Lysosomal labelling In live cells, lysosomes were labelled with 50 nM LysoTrackerTM DeepRed (#L12492, Thermo Fisher Scientific) for 30 minutes and washed with fresh medium before being imaged in 2D on a Leica SP8 Laser Scanning Confocal Microscope equipped with a Leica HC PL APO oil 63x objective (#50634). LysoTracker DeepRed was excited using a 633 nm laser and emission collected at wavelengths 650-710. Images were analyzed using ImageJ. 55 2.17 Immunofluorescent staining Mouse or human islet cells were fixed in 4% PFA, rinsed with PBS (#L14190-136, Thermo Fisher Scientific), permeabilized using 0.05% Triton X-100 (X100, Sigma-Aldrich) for 5 min, and blocked for 1 hour with Serum Free Protein Block (#X0909, Agilent-DAKO) at room temperature. Primary antibodies were added at 4℃ overnight and secondary antibodies for ~1 hour at room temperature. Primary and secondary antibodies are listed in Table 2.5. Table 2.5 Antibodies used for immunofluorescent staining Antibody Source Cat. No Dilution Anti-Lamp1 antibody Abcam® Ab25245 1 to 500 Anti-TFEB antibody Mybiosource® MBS7604860 1 to 200 TFEB antibody Cell Signaling® #4240 1 to 200 Anti-Insulin antibody Dako® A0564 1 to 400 Anti-TFE3 antibody Sigma® HPA023881 1 to 1000 Alexa Fluor® 488 Goat anti GP IgG Cross-Adsorbed Antibody Thermo Fisher-Invitrogen A11073 1 to 2000 Alexa 555 Fluor® Goat anti Rabbit IgG antibody Thermo Fisher-Invitrogen A27039 1 to 2000 Alexa 633 Fluor® Goat anti GP IgG Highly Cross-Adsorbed Antibody Thermo Fisher-Invitrogen A21105 1 to 2000 56 Alexa 594 Fluor® Goat anti Rat Cross-Absorbed Antibody Thermo Fisher-Invitrogen A11007 1 to 2000 Alexa 680 Fluor® Goat anti Rabbit Highly Cross-Adsorbed Antibody Thermo Fisher-Invitrogen A32734 1 to 2000 LysoTracker staining and Immunocytochemistry images were analyzed for their mean intensity and/or puncta counts using ImageJ software. Masks for the cellular nuclear area, cytosolic area and/or the whole cell area were generated using built-in threshold function. Puncta were identified and analyzed for size and fluorescence intensity using ImageJ Auto Local Threshold and Particle Analysis functions. 2.18 3D live-cell autophagy flux imaging Dispersed islet cells from CAG-RFP-EGFP-LC3 mice were seeded in ibidi 8-well glass bottom chamber slides coated with 804G cell-derived extracellular matrix to improve islet cell attachment and spreading (206). The islet cells were imaged after 24 hours culture in a Tokai Hit INUBTFP-WSKM stage-top incubator under normoxia or hypoxia. Stack images were taken using the high-speed (8,000 Hz) resonance scanner of a Leica SP8 Laser Scanning Confocal Microscope and collected using an HC PL APO oil 63x objective. Signals were collected at the spectrum of EGFP (excitation: 488 nm, emission: 493-550 nm) and mRFP (excitation: 561 nm, 57 emission 650-765 nm). The resulting image stacks were deconvolved in Huygens Pro® deconvolution wizard and then reconstructed 3-dimensionally in ImageJ using the “3D object count” plugin. Colocalization of GFP- and RFP-positive signal and puncta volumes were quantified using the ImageJ “Coloc 2” and “3D object count” plugins, respectively. 2.19 Statistics Data are represented as mean ± SEM and were analyzed in GraphPad Prism 9.0 (La Jolla, California) using Student’s t-test, one-way ANOVA, or two-way ANOVA followed by multiple comparison tests, as appropriate. Statistical significance was set at p < 0.05. 58 Chapter 3: Autophagy is essential for β-cell function and viability under hypoxia and following islet transplantation 3.1 Background In this chapter, we examine the importance of autophagy for β-cell survival and adaptation under hypoxia. Hypoxia is a known stress factor that can negatively affect β-cell function and survival - mostly in the setting of clinical islet transplantation efforts to treat T1D, but also during the development of T2D (36). With the development of the Edmonton Protocol islet transplantation was established as an efficient T1D treatment, but its widespread use is still limited by donor islet availability and long-term graft failure (207). Major progress has been made toward generating insulin-secreting β-cells from human embryonic stem cells for transplantation (208). While this may soon provide an abundant supply of human β-cells, transplanted stem cell-derived islets will be vulnerable to stress-induced dysfunction and failure, similar to primary donor derived islets. Among the factors that negatively affect islet transplantation outcomes, hypoxia affects the function and viability of pancreatic β-cells both pre- and post-transplantation (15). Oxygen deprivation of transplanted islets is largely due to a lack of graft re-vascularization for weeks after the surgery (42,209); it has been estimated that O2 levels are as low as 1% in islets transplanted under the renal capsule of mice (210). Evidence also suggests that β-cell hypoxia is an etiological factor in T2D (46,211). As hyperglycemia develops, β-cell workload and mitochondrial oxygen consumption increase markedly, which can induce a state of local hypoxia in the islets (35). This activity-dependent reduction in local O2 levels may possibly be 59 worsened by disruptions of islet vascularization and blood flow (212,213), and ultimately contribute to β-cell failure and loss in T2D. For all these reasons, it is important to understand why and how β-cell hypoxic stress causes β-cell dysfunction, and how this may be mitigated. The mechanisms of hypoxia-induced cell dysfunction and death involve oxidative, inflammatory, and ER stress. Hypoxia leads to an increase in ROS , and β-cells are particularly vulnerable to this oxidative stress because of their low expression levels of anti-oxidant enzymes (36). Moreover, the pro-inflammatory cytokine IL-1β is produced locally in hypoxic and transplanted islets (214). In the setting of islet transplantation, it is interesting to speculate that hypoxia-induced islet ROS might also act as a signal that can recruit immune cells to further amplify inflammation and provoke graft failure (215). Finally, hypoxic β-cells undergo maladaptive ER stress (46) that contributes to dysfunction and DDIT (i.e. CHOP)-dependent death of transplanted β-cells (16). To understand how hypoxic β-cell failure and death might be prevented, it is beneficial to consider which β-cell-intrinsic mechanisms are being activated to help offset the detrimental effects. Known adaptive responses to hypoxia include HIF-1α-mediated expression of angiogenic factors, metabolic reprogramming and upregulation of antioxidant defenses (216-218). Because hypoxia triggers ER stress, the adaptive UPR is also engaged to restore homeostasis. However, prolonged β-cell hypoxia may ultimately result in failure of the adaptive UPR (45). Autophagy is also known to be activated under hypoxia. Mechanistically, AMPK and mTORC1 regulate hypoxia-induced autophagy (219) and hypoxia increases BNIP3 expression in 60 a HIF-1α-dependent manner (43,220). BNIP3 functions as a mitophagy receptor and it can also bind to Bcl-2 to release Bcl-2-bound BECLIN-1, which then participates in autophagy initiation (220). Whether autophagy, like the adaptive UPR, acts as a protective β-cell response to hypoxia remains unclear. Autophagy is generally considered to be cell-protective, but the relationship between autophagy and apoptosis is complex, and autophagy has also been reported to promote hypoxic cell death (221). Relatively little is known about the role of autophagy in hypoxic β-cells. In MIN6 β-cells, 24 hours of hypoxia (1% O2) caused Ddit3 (i.e. Chop)-dependent apoptosis and increased autophagy (indicated by increased LC3-II levels). The increase in autophagy was speculated - but not demonstrated - to contribute to the observed cell death (46). In INS1 β-cells, intermittent hypoxia (a model of hypoxia-re-oxygenation induced by sleep apnea) induced protective autophagy that depended on PERK/eIF2α/ATF4 signaling (60). This suggests a close relationship between the UPR and autophagy in β-cells, but this relationship has not been examined in the context of sustained hypoxia. It also remains to be conclusively established whether endogenous autophagy promotes or prevents β-cell apoptosis under continued hypoxia, and if it protects or kills β-cells in transplanted islets. In this study, we investigated the impact of sustained hypoxia on islet expression of UPR- and autophagy-related genes, as well as autophagic function. Further, we tested the hypothesis that the endogenous autophagy machinery promotes β-cell function and survival under hypoxic stress in vitro, and in vivo following islet transplantation. For this, we established a novel model 61 of β-cell-specific Atg5 knockout by crossing Atg5flox/flox and Ins1Cre mice, and further achieved time-controlled Atg5 deletion in vivo and in vitro by intraductal delivery of Cre-expressing AAV and adenoviral vectors, respectively. 3.2 Results 3.2.1. Hypoxia leads to loss of adaptive UPR- and autophagy-related transcripts in pancreatic β-cells. Hypoxia is known to induce both Hif-1α dependent and independent changes to gene expression. To investigate how hypoxia affects primary mouse islets, we first cultured isolated C57BL/6J islets under hypoxia (1% Oxygen) or normoxia (20% Oxygen) and quantified the time-dependent effects on gene expression by qPCR. As expected, the known hypoxia-induced Hif-1α target transcripts Hmox1, AldoA, Gapdh and Bnip3, were rapidly upregulated under hypoxic culture (Figure 3.1). 62 Hypoxia also significantly induced the expression of pro-apoptotic ER stress genes Ddit3 and Atf3, whereas the expression of Xbp1s and Hspa5, which are part of the adaptive UPR, on average were reduced or unchanged (Figure 3.2). These findings agree with a previous report of selective hypoxia-induced impairment of the adaptive UPR (45). Autophagy is well-known for its protective role in β-cells under various stress conditions, including glucolipotoxicity (128,137) and ER stress (132), but the impact of hypoxia on the β-cell autophagy pathways is not clear. To explore this, we quantified the mRNA levels of the key autophagy-regulators Atg5, Map1lc3b and Becn1 and found that these on average were Figure 3.1 – Time-dependent expression of known hypoxia-responsive transcripts in male C57Bl/6J islets under normal and 1% O2 culture. Data represent mean ± SEM of 4-6 independent experiments. *p < 0.05, **p < 0.01, ***p < 0.001, ****p < 0.0001 by two-way ANOVA followed by Tukey’s multiple comparison test. Figure 3.2 – Hypoxia-induced changes to islet expression of select UPR / ER stress response genes in male C57Bl/6J islets under normal and 1% O2 culture. Data represent mean ± SEM of 4-6 independent experiments. *p < 0.05, **p < 0.01, ***p < 0.001, ****p < 0.0001 by two-way ANOVA followed by Tukey’s multiple comparison test. 63 downregulated similarly to the adaptive UPR transcripts, suggesting that autophagy may also be negatively affected by hypoxic stress (Figure 3.3). To investigate the relationship between the UPR and autophagy (60), we performed correlation analysis of the changes to gene expression across all time-points. Notably, this revealed that the hypoxia-induced changes to autophagy-related transcripts were correlated with changes to the adaptive UPR transcript Xbp1s, but not to the pro-apoptotic ER stress genes Ddit3 and Atf3 (Figure 3.4). These results highlight that gene expression in the adaptive UPR and autophagy pathways may be correlated, and are similarly suppressed, by hypoxia in mouse islets. Figure 3.3 – Hypoxia-induced changes to islet expression of select autophagy-related genes in male C57Bl/6J islets under normal and 1% O2 culture. Data represent mean ± SEM of 4-6 independent experiments. *p < 0.05, **p < 0.01, ***p < 0.001, ****p < 0.0001 by two-way ANOVA followed by Tukey’s multiple comparison test. 64 3.2.2. Endogenous autophagic flux is impaired in hypoxic β-cells Since hypoxia impaired the transcription of several autophagy-related genes, we further assessed how endogenous autophagy is affected by prolonged hypoxia. For this, we cultured pancreatic islet-cells from RFP-EGFP-LC3 autophagy reporter mice under normoxia and hypoxia and quantified autophagic flux using 3D live-cell imaging. This fluorescent autophagy biosensor combined with chemical inhibitors of lysosomes provide a readout of autophagic flux (Figure 3.5a) (222). As illustrated in Figure 3.5b, treatment with lysosomal inhibitors such as Figure 3.4 – Correlation analysis of hypoxia-induced changes to UPR- and autophagy-related genes a) Examples of linear regression fits and correlation analysis of hypoxia-induced changes to gene expression across all time-points examined in Figures 3.1-3.3. b) Summary of correlation between the expression of UPR transcripts (adaptive Xbp1s, Hspa5; maladaptive Ddit3, Atf3) and autophagy-related transcripts (Atg5, Map1lc3a, Map1lc3b, Becn1, Bnip3). The data were analyzed using Pearson’s Correlation. Numbers in the table indicate the Pearson’s correlation coefficient and the statistical significance is indicated by asterisks; *p < 0.05, **p < 0.01, ***p < 0.001. 65 bafilomycin-A1 or chloroquine (CHQ), which cause a loss of autophagosome clearance and accumulation of swollen autophagosomes and/or autolysosomes with impaired acidification (yellow puncta) (223). Using this approach, we assessed autophagic flux in live islet cells after 24 hours culture under normoxia or hypoxia. As illustrated in images from 3D reconstructions of reporter islet cells (Figure 3.6a) and quantified by the EGFP/RFP colocalization (Figure 3.6b), prolonged hypoxia led to an accumulation of autophagosomes and/or dysfunctional autolysosomes. This accumulation resembles what was seen when lysosomal flux was blocked with CHQ. When comparing the two conditions (Figure 3.6b; 2nd and 3rd bars) it appears that autophagic flux is Figure 3.5 – Principle of autophagic flux measurements by RFP-EGFP-LC3 biosensor a) Schematic illustrating the principles of the RFP-EGFP-LC3 fluorescent autophagy biosensor. RFP-EGFP-double-tagged LC3 accumulates on autophagosomes, which can be visualized as green and red (i.e. yellow) puncta. After fusion with a lysosome, EGFP fluorescence is quenched by the low pH in the lysosomal lumen and the resulting autolysosome can be visualized as a red-only vesicle (created with BioRender.com). b) Example illustrating the fluorescent readout of autophagic flux and its inhibition by bafilomycin-A1 (BafA) treatment in islet cells from RFP-EGFP-LC3 reporter mouse. In control culture, there is a predominance of autolysosomes (red fluorescence only). Following inhibition of lysosomal function by BafA there is a loss of autophagosome-lysosome fusion, resulting in accumulation of autophagosomes and/or non-acidic autolysosomes (red + green fluorescence = yellow in overlay). 66 almost completely halted in a significant fraction of hypoxic β-cells. An increase in the volume of the individual autophagosomes and autolysosomes further supports that hypoxia disrupts normal homeostatic control of these organelles (Figure 3.6c). 67 3.2.3. Autophagy is important for β-cell survival under hypoxia-induced stress Atg5 is one of the autophagy-related mRNAs that was significantly down-regulated by hypoxia (Figure 3.3). It is a key player in the initiation of autophagosome formation and lack of Atg5 leads to loss of LC3-II and autophagy-deficiency in animal models (197,224). To investigate the role of autophagy in β-cells under hypoxia, we used adenovirus-mediated expression of Cre recombinase (Ad.Cre) to delete Atg5 in dispersed islet cells from Atg5flox/flox mice. To examine any potential virus-induced stress on the islet cells, and to optimize the transduction efficiency, we transduced the dispersed islet cells with different Ad.Cre titers (MOI 10, 40 and 100). At an MOI of 40, Cre recombination without signs of cell loss was observed in nearly 100% of dispersed islet cells from ROSAmTmG Cre reporter mice (Figure 3.7a,b), and resulted in ~90% loss of Atg5 mRNA expression in Atg5flox/flox islet cells (Figure 3.7c). Figure 3.6 - Prolonged hypoxia interrupts autophagic flux in mouse islet cells. Autophagic flux was analyzed by 3D confocal analysis of islet cells from male CAG-LC3-RFP-EGFP reporter mice, cultured for 24 hours under 20% O2 or 1% O2 with and without addition of 10 µM chloroquine (CHQ). a) Representative 2D cross-sectional confocal images of RFP-EGFP-LC3-expressing islet cells showing autolysosomes in red (RFP single-positive) and autophagosomes and/or non-acidic autolysosomes in yellow (RFP/EGFP double-positive) under the various culture conditions. b) Colocalization of EGFP and RFP fluorescence was quantified on a per-cell basis using the Manders’ Overlap Coefficient, which ranges from 0 (no overlap) to 1 (complete overlap). Increased colocalization coefficients indicate autophagosome accumulation with preserved EGFP fluorescence in cells exposed to hypoxia and/or CHQ treatment. The combined number of autophagosomes and autolysosomes were quantified from the number of RFP-positive (RFP+) puncta in 3D reconstructions of individual cells. c) Quantification of individual RFP+ puncta shows that hypoxia- and CHQ-induced loss of autophagic flux is associated with increased autophagosome/autolysosome volume. Data represent mean ± SEM of 3 independent experiments. *p < 0.05, **p < 0.01, ***p < 0.001 by two-way ANOVA followed by Tukey’s multiple comparison test. 68 Assessment of Atg5flox/flox islet cell viability 4 days after transduction with either a control adenovirus (Atg5Ad.Ctrl) or an adenovirus expressing Cre (Atg5Ad.KO) showed that autophagy deficient Atg5Ad.KO cells underwent significantly quicker and more severe death than Atg5Ad.Ctrl cells under hypoxia (Figure 3.8). This demonstrates that autophagy is a critical pro-survival mechanism for pancreatic β-cells under hypoxic stress. Figure 3.7 – Efficient recombination and Atg5 KO by adenovirus Cre transduction in vitro. a) Representative images of dispersed islet cells from ROSAmTmG reporter mice following transduction by adenovirus Cre (Ad.Cre) at MOI 40. Loss of tdTomato and gain of EGFP fluorescence shows the degree of LoxP recombination over 4 days. b) The ratio of tdTomato/EGFP reporter fluorescence intensities was quantified and used as an indicator of recombination over time at different MOIs of virus. ****p < 0.0001 by two-way ANOVA followed by Tukey’s multiple comparison test. c) Atg5 mRNA levels in Atg5flox/flox mouse islet 4 days after transduction with 40 MOI Ad.Cre virus (Atg5Ad.KO cells) or Ad.Ctrl virus (Atg5Ad.Ctrl cells) n = 3; *p < 0.05 by paired Student’s t-test. 69 Figure 3.8 - Autophagy is essential for β-cell survival under hypoxia. a) Representative images comparing hypoxia-induced cell death (propidium iodide incorporation) of Atg5Ad.KO and Atg5Ad.Ctrl mouse islet cells at 0 and 16 hrs (scale bar 100 µm). b) Time-course quantification of hypoxia-induced cell death in 3 independent cultures of Atg5Ad.KO and Atg5Ad.Ctrl mouse islet cells. c) Total cell death (area under the curve; AUC) of each experiment in panel (b). *p < 0.05 by Student’s t-test. 70 3.2.4. β-Cell-specific deletion of Atg5 causes early-onset glucose intolerance in male and female mice We next aimed to determine if Atg5-dependent autophagy is important for β-cell viability in vivo and under the compound stresses associated with islet transplantation. To this end, we established a mouse with β-cell-specific Atg5 knockout and autophagy-deficiency. Previous work using RIP-Cre transgenic mice to delete the core autophagy protein, Atg7 showed an inability for β-cell adaptation to diet-induced obesity and impaired clearance of toxic IAPP oligomers (32,128,145). These findings strongly support an important role of autophagy in sustaining β-cell stress adaptation and glucose tolerance. They are, however, confounded by possible RIP-Cre-mediated recombination in the hypothalamus (225,226), where Atg7 affects lipid metabolism and glucose homeostasis (227-229). Moreover, none of these prior studies examined possible sex differences, in the requirement for β-cell autophagy. To address these limitations, we crossed the Ins1Cre knock-in mice (230) with our Atg5flox/flox mice to induce β-cell-specific deletion of Atg5, without disturbing Atg5 expression in the hypothalamus. Islets from Atg5flox/flox:Ins1Cre (i.e. Atg5βKO) mice showed the expected ~85% loss of Atg5 mRNA (Figure 3.9a). Although both male and female Atg5βKO mice gained the same amount of weight as their Atg5flox/flox (Atg5βWT) littermates (Figure 3.9b), they rapidly developed significant glucose intolerance by 5 weeks and overt diabetes with concomitant increases in fasting blood glucose by 10 weeks (Figure 3.9c). 71 These results demonstrate that, independent of any brain effects, chronic loss of basal β-cell autophagy results in severe glucose intolerance and show that the overall progression of the glucose intolerance phenotype is similar between the sexes. Figure 3.9 – Basal Atg5-dependent β-cell autophagy is essential for in vivo glucose tolerance. β-Cell Atg5 knockout (Atg5βKO) mice were established by crossing Atg5flox/flox and Ins1Cre knock-in animals. a) qPCR confirmation of Atg5 mRNA loss in pancreatic islets from 10–12-week-old Atg5βKO and Atg5βWT littermates. Residual Atg5 mRNA in KO islets is likely from non-β-cells. b) Body weights of male and female Atg5βWT and Atg5βKO mice at different ages show no genotype-dependent differences. c) Intraperitoneal glucose tolerance tests of 3-, 5- and 10-week-old male (top) and female (bottom) Atg5βWT and Atg5βKO mice show significant glucose intolerance as early as 5 weeks of age. Rightmost panels summarize the data as the area under the curve (AUC) of the blood glucose measurements. Data are shown as mean ± SEM. Panel (a) was analyzed by paired Student’s t-test; n=3; *p < 0.05. Panels (b) & (c) were analyzed by repeated measures two-way ANOVA and Sidak’s multiple comparison tests; n = 4-11 littermates at the different ages. *p < 0.05, **p < 0.01, ***p < 0.001, ****p < 0.0001. 72 3.2.5. β-Cell autophagy-deficiency accelerates failure of syngeneic islet grafts The early-onset diabetes in our Atg5βKO mice prevented us from isolating autophagy-deficient islets that were not already stressed and dysfunctional. We consistently observed that Atg5βKO islets were pale, fragile and difficult to keep viable in culture. To examine the role of β-cell autophagy in islet graft function, we therefore induced time-controlled Atg5 knockout by injecting Atg5flox/flox mice with dsAAV6 expressing Cre recombinase under the rat Ins1 promoter (dsAAV6-RIP1-Cre; Atg5AAV.βKO mice), or empty dsAAV6 control virus (dsAAV6-Empty; Atg5AAV.Ctrl mice). To ensure pancreas-specific expression, the virus was injected via the pancreatic duct 2 weeks before the islets were isolated and transplanted into syngeneic recipients with STZ-induced diabetes (Figure 3.10). As expected from previous work with this dsAAV6 vector (201,202), Cre was expressed in insulin-positive β-cells, but not glucagon-positive α-cells (Figure 3.11a), and qPCR analysis and western blot confirmed the loss of Atg5 mRNA and protein in Atg5AAV.βKO mouse islets (Figure 3.11b). Reduced protein levels of LC3-II relative to LC3-I further showed that Atg5AAV.βKO donor islets had a basal autophagy deficiency, as expected in the absence of Atg5 (Figure 3.11c). Figure 3.10 - Schematic outline of workflow for marginal mass islet transplantation experiment. 73 Blood glucose measurements confirmed that all the STZ-treated recipient mice were diabetic before receiving islet transplants (Figure 3.12a). We further confirmed that both Atg5AAV.Ctrl and Atg5AAV.βKO islet donor mice showed normal glucose tolerance 1 day before islet isolation (Figure 3.12b). The isolated donor islets also had similar glucose-stimulated Ca2+ and insulin secretion responses prior to transplantation (Figures 3.12c-e). Overall, the virus-induced short-term Atg5 knockout does not seem to impair β-cell function in vivo. Figure 3.11 – dsAAV6-RIP1-Cre induced β-cell Atg5 deletion and autophagy deficiency in vivo. a) Illustrative immunostaining showing Cre expression in β-cells, but not α-cells, 2 weeks after intraductal dsAAV6-RIP1-Cre injection. Top insert: Contrast enhancement reveals low levels of Cre expression in seemingly negative β-cells. b) Western blot and qPCR quantification demonstrating loss of Atg5 mRNA and protein in islets from dsAAV6-RIP1-Cre-injected (Atg5AAV.βKO) mice compared to islets from mice injected with dsAAV6-Empty virus (Atg5AAV.Ctrl) (n = 3 islet preparations of each genotype, **p < 0.01, ***p < 0.001 by t-test). c) Western blot showing that islets from Cre-expressing Atg5AAV.βKO islets have reduced levels of LC3-II, indicating loss of basal autophagy. 74 We monitored blood glucose and body weight in the islet transplant recipients until the grafts failed and the mice became diabetic, or the mice reached our pre-defined endpoint at 14 days Figure 3.12 – Characterization of islet graft recipient and donor mice pre-transplantation. a) Random blood glucose measurements confirming that Atg5flox/flox mice that were intended as islet transplant recipients developed STZ-induced diabetes. Prior to STZ injection, the mice were randomly assigned to receive islets from either Atg5AAV.Ctrl (black) or Atg5AAV.βKO (red) donor mice. b) IPGTT demonstrating that the Atg5AAV.βKO donor mice had normal glucose tolerance 2 weeks after AAV injection (n=12 mice in each group). c) Fura-2 fluorescence imaging demonstrating similar glucose-stimulated Ca2+ responses in aliquots of Atg5AAV.Ctrl and Atg5AAV.βKO donor islets (14 Ctrl and 13 KO islets from 3 separate and matched donor preparations). d) Area under the curve (AUC) summary of Ca2+ imaging results from panel (c). e) Static incubation demonstrating robust and comparable glucose-stimulated insulin secretion in aliquots of Atg5AAV.Ctrl and Atg5AAV.βKO donor islets (n=4 islet preparations). 75 post-transplantation (Figure 3.10). Both before and after islet transplantation, the body weights did not differ between the two recipient groups, and the transplantation of a marginal mass of islets reversed diabetes in both groups (Figure 3.13a,b). Importantly, however, autophagy-deficient grafts failed at a faster rate than the similar-sized Atg5AAV.Ctrl grafts (Figures 3.13b,c). Mice that had received transplants of Atg5AAV.βKO islets also had worse glucose tolerance 14 days post-transplantation (Figure 3.13d). At the experimental endpoint, 5 control and 1 KO graft recipients had random blood glucose measurements below the 20 mM cutoff we used to define the onset of diabetes. After removal of the graft bearing kidney all of these mice became diabetic within 4 days, confirming that their normoglycemia was maintained by the transplanted islets (Figures 3.13b,e). Together, these data show that β-cell autophagy plays an important role in sustaining islet viability under the compound hypoxic, inflammatory and glucotoxic stress experienced in syngeneic grafts immediately post-transplantation (231). 76 Figure 3.13 – β-cell autophagy-deficient islet grafts are sensitized to failure. a) Average body weights of STZ-induced diabetic Atg5flox/flox mice before and after receiving marginal mass transplants of 350 Atg5AAV.Ctrl or Atg5AAV.βKO islets (n=7 recipients of each graft genotype). b) Random blood glucose measurements from individual Atg5flox/flox mice made diabetic by STZ and receiving marginal mass islet grafts from Atg5AAV.Ctrl or Atg5AAV.βKO donors. Endpoint was defined by 2 consecutive days with BG > 20 mM or 2-weeks post-transplantation for mice with no graft failure. c) Survival curve showing diabetes incidence in islet recipients (n=7 recipients of each graft genotype). d) Endpoint IPGTT of a subset of graft recipients (n=4 recipients of each graft genotype, **p < 0.01, ***p < 0.001 by repeated measures two-way ANOVA followed by Sidak’s multiple comparison tests). e) Blood glucose levels of normoglycemic islet graft recipients following nephrectomy of the graft-bearing kidney. The return to hyperglycemia confirms that normoglycemia depended on the islet graft. 77 3.3 Discussion In this chapter, we investigated the importance of endogenous β-cell autophagy under hypoxic stress in vitro, and in animal models in vivo. We characterized glucose tolerance in male and female mice with β-cell-specific knockout of Atg5 and demonstrated that basal Atg5-dependent β-cell autophagy is required for glucose homeostasis in both sexes. We further showed that β-cell autophagy-deficiency sensitizes to hypoxia-induced death in vitro and islet graft failure in vivo. Moreover, prolonged hypoxic stress was associated with a correlated disruption of UPR- and autophagy-related gene expression and led to an impairment of β-cell autophagic clearance. Mice with global homozygous deletion of Atg5 or Atg7 die within a day of birth (224,232), so knockout studies are primarily done using Cre-mediated tissue-specific deletion. Our mice with β-cell-specific Atg5 deletion developed glucose intolerance, which began manifesting as early as 3 weeks of age. Loss of glucose homeostasis has also been reported in other mouse lines with genetic β-cell autophagy-deficiency (32,127,128,133,145,233), but these previous models are all confounded by possible gene deletion in non-β-cells. Specifically, RIP-Cre transgenic mice (234,235) were used to delete Atg7 and study the importance of β-cell autophagy in normal in vivo physiology (127), and under the stresses induced by genetic obesity (ob/ob) (133), high fat diet feeding (128) or accumulation of toxic IAPP oligomers (32,145). In all these settings, β-cell Atg7 was found to be critical for maintaining glucose homeostasis. More recently, Atg5 knockout was also reported to cause β-cell failure and glucose intolerance in the absence of external stressors (233). However, these Atg7 and Atg5 mouse models all expressed Cre under 78 the rat Ins2 promoter, which is now known to cause off-target recombination in hypothalamic neurons (225,226). This leaves some uncertainty regarding the observed phenotypes, particularly because hypothalamic autophagy is important for metabolic control (227-229). For these reasons, we used the more recent Ins1Cre mouse in which Cre recombinase is knocked into the endogenous Ins1 locus, replacing the coding sequence (230). Ins1 expression is more restricted to the pancreatic β-cells than Ins2 (236), so the resulting gene deletion is more β-cell-specific. To our knowledge there have been no reports of recombination in the brain with the Ins1Cre model. Another question we addressed is if β-cell autophagy-deficiency results in sex-specific phenotypes. Most β-cell Atg7 knockout studies have used male mice (128,133,144) or mice of both sexes without reporting the results separately (127). Kim et al. did present IPGTT data from 12–15-week-old male and female mice with RIP2-Cre-driven Atg7 knockout, but their statistical comparisons focused on the impact of transgenic hIAPP expression. It is therefore unclear what effects Atg7 deletion, per se, had in their mice, and how this compared between the sexes (145). More recently, RIP2-Cre driven Atg5 deletion was shown to cause glucose intolerance in 8-week-old male and female mice. The effects of biological sex were not compared directly, but the phenotype was not obviously worse in either sex (233). Overall, this agrees with what we observed in our β-cell-specific Atg5 KO mice. Although the progression of i.p. glucose intolerance did not differ markedly between our male and female Atg5βKO mice, it is still possible that there are sex-differences in other measures such as random blood glucose levels, β-cell stress responses, organelle function and insulin secretion. 79 In addition to the importance of basal constitutive β-cell autophagy, we investigated the more acute roles of autophagy under transplantation-relevant hypoxic stress. Our finding that hypoxia up-regulated islet expression of Hif-1α target genes and pro-apoptotic UPR effectors, but reduced the expression of adaptive UPR genes, is consistent with previous reports (45,211). ER stress is known to activate autophagy and specific ER-phagy, which, as an example, can facilitate cancer cell survival in the hypoxic tumor environment (237). Our data are compatible with a connection between the adaptive UPR and autophagy at the transcriptional level. As mentioned earlier, signaling in the PERK/eIF2α/ATF4 axis can drive expression of autophagy genes, including in β-cells (60,122). It is therefore possible that loss of UPR function could impair Atf4-dependent autophagy gene expression in our hypoxic islets. It is less established if impaired β-cell autophagy positively or negatively affects UPR signaling – islets from RIP2-Cre Atg7 KO mice have reduced expression of UPR genes (133), but for reasons that are unclear, many of the same UPR transcripts are increased in islets from RIP2-Cre Atg5 KO mice (233). The nature of this relationship therefore remains to be resolved. It is also not evident how loss of core ATG proteins causes changes to the expression of UPR genes, but it seems likely to be an indirect effect of impaired cellular homeostasis. Another fundamental question we addressed is whether β-cell autophagy is pro- or anti-apoptotic under hypoxia. Previous reports have provided conflicting data in this regard. We tested this directly using our Atg5 knockout islet cells and established that endogenous β-cell autophagy has a protective role under culture in 1% O2. Autophagy does promote cell death in 80 some situations, including neuronal death related to ischemia and hypoxia, so the role of autophagy likely depends on the cell-type, as well as the severity of the hypoxic stress (238). In cultured β-cells, autophagy is protective under CoCl2-induced Hif-1 stabilization and in response to intermittent hypoxia (60,239). However, neither of these conditions are likely to model the sustained oxygen deprivation experienced by transplanted islets. In rodent studies where islet grafts are placed beneath the renal capsule, the oxygen tension has been estimated to fall below 2% the day after surgery, and drop to 1% O2 after one month, despite the fact that graft revascularization is considered to have happened at this point (210). In comparison, the level of islet hypoxia induced by the O2 consumption in ‘hyperactive’ β-cells is estimated to be in the low-to-moderate range of 1.4%-6.3% (35), which will activate the UPR, as well as both Hif-1α-dependent and independent transcriptional changes (35,43). Our results using the 1% O2 model therefore demonstrate that autophagy is important for β-cell survival at the levels of hypoxia they experience in islet grafts, and possibly also in particularly low-oxygen islets in T2D (35,41). Little is known about the role of β-cell autophagy in islet transplantation. Previous work has shown both positive and negative effects of autophagy in islets pre- and post-transplantation, but this has all relied on pharmacological stimulation by rapamycin or carbon monoxide exposure (240-242). Our comparison of Atg5AAV.Ctrl and Atg5AAV.βKO islet grafts is the first direct test of whether endogenous β-cell autophagy is protective or detrimental in this setting. That transplanted islets from our Atg5AAV.βKO donor mice failed faster than those from Atg5AAV.Ctrl control mice establishes that β-cell autophagy is a protective response in syngeneic islet grafts. In 81 agreement, our in vitro experiments showed that autophagy initiation is critical for β-cell viability under hypoxic stress. However, our autophagy reporter data also revealed that prolonged hypoxia impairs lysosomal clearance and causes autophagosomes to accumulate. A similar phenomenon of autophagosome buildup is observed in β-cells from patients with T2D (149,243) and T1D (148). We do not yet know if this happens in transplanted β-cells, but it seems likely considering the hypoxic stress they are under. Not only are the benefits of autophagic degradation then lost, but the accumulation of autophagosomes has additional cytotoxic effects (244) that could conceivably worsen graft failure or diabetes pathogenesis. This also implies that stimulating autophagy might have unintended toxic effects if it is done at a point where autophagosome removal is insufficient. Interestingly, rapamycin is toxic to β-cells, in part, because it stimulates autophagy (242). The authors attributed this to excessive autophagic self-digestion (242), but that was not directly demonstrated and could conceivably involve toxic autophagosome hyper-accumulation. As discussed further in Chapter 4, we believe lysosomal clearance needs to be carefully considered if we want to harness autophagy for β-cell protection. In summary, we used specific Cre-deleter models to establish that β-cell autophagy is essential for basal glucose homeostasis and promotes β-cell survival under the hypoxic islet stress associated with the immediate post-transplantation period. Under chronic hypoxic stress, however, β-cell adaptation may become compromised by failure of autophagosome clearance. Further investigations of the mechanisms are warranted and may identify ways to improve β-cell stress resilience after transplantation of donor- or stem cell-derived islets and in pre-diabetes. 82 Chapter 4: Dysregulation of TFEB and lysosomes in β-cells under stress 4.1 Background In Chapter 3 we found that prolonged in vitro hypoxia impedes autophagic flux, and this appears to be associated with failure of the lysosomal clearance step. In this chapter we shed light on the underlying mechanisms by further examining the impact of hypoxia, and other stress conditions, on β-cell lysosomal homeostasis. β-Cell lysosomes have long been known to mediate degradation and quality control of β-cell insulin granules through the autophagy-independent mechanism of crinophagy (245). In recent years, β-cell lysosomes have had renewed attention for their role in autophagy, which we now know is involved in both stress adaptation and (pro)insulin homeostasis (129,246,247). Of note, lysosomes appear to be dysregulated when autophagy is impaired in β-cells exposed to pro-inflammatory cytokines (147) or glucolipotoxic stress in vitro (146,248,249). Prolonged β-cell lipotoxicity reduces lysosomal acidification (249). Because luminal hydrolases – including cathepsins – need a low pH environment to function, this will impair autophagic degradation. It was shown that pharmacological inhibition of lysosomal cathepsins worsens β-cell death under glucose stress (250). If lysosomes become dysfunctional in hypoxic β-cells - as our autophagy flux data indicate - it would thus likely contribute significantly to β-cell failure and loss. Lysosomal defects have been implicated in several human diseases. The most direct examples are lysosomal storage diseases/disorders (LSD), which are a category of genetic disorders that result in defective lysosomal degradation and consequent organ dysfunction (251). 83 There is also evidence to suggest that lysosomal dysfunction plays a role in diabetes. β-Cells from donors with T2D show an accumulation of autophagosomes, and this is accompanied by impaired lysosomal homeostasis and loss of lysosome-related transcripts, such as LAMP2, and the cathepsins CTSB and CTSD (149,252). In the context of T1D, a recent study revealed that β-cells from NOD mice and human pancreas donors with T1D have impaired autophagic flux, characterized by accumulation of autophagosomes and reduced LC3-LAMP1 co-localization (i.e. autophagosome-lysosome fusion) (148). Of note, they also found ultrastructural evidence for lysosomal abnormalities in β-cells from pre-diabetic autoantibody-positive donors, which suggests that lysosomal clearance mechanisms may start failing quite early in the progression of T1D. Overall, there is thus a growing body of evidence associating lysosomal dysfunction with autophagy impairment and β-cell failure in the development of diabetes. However, the underlying causes and mechanisms of this lysosomal failure remain unclear. Lysosomal biogenesis is largely processed in ER and Golgi-networks (253), so disruption of ER homeostasis in stressed β-cells could conceivably affect the number and quality of lysosomes. But, perhaps more importantly, the above-mentioned changes to islet gene expression in T2D islets suggest β-cell stress may disrupt the transcriptional regulation of lysosomes, possibly involving dysregulation of TFEB or other MITF transcription factors (252,254). In this study, we explore β-cell lysosomal function and regulation in the context of hypoxia and other in vitro conditions related to ER stress. We first investigate the relative roles of Atg5-dependent autophagy and lysosomal function on mouse β-cell survival under hypoxia. Moreover, 84 we test the hypothesis that hypoxia, and other forms of diabetes-relevant stress, impair β-cell lysosomes and Tfeb-dependent gene expression. We further investigate the effects of pharmacologically and genetically manipulating Tfeb activity on β-cell lysosomal function and survival under stress. 4.2 Results 4.2.1. Hypoxia perturbs lysosomal homeostasis in β-cells In Chapter 3, we observed that the number of β-cell autophagosomes (RFP+/EGFP+ double-positive vesicles) increased due to hypoxia or CHQ treatment, while the number of autolysosomes was reduced (Figure 3.6). This suggests that autophagosomes are accumulating because their lysosomal clearance fails. To examine this possibility, we investigated lysosomal function in hypoxic β-cells. In a preliminary test, we used LysoTracker staining to visualize lysosomes (acidic organelles) in live Atg5Ad.Ctrl and Atg5Ad.KO islet cells. Because LysoTracker dye cannot be added to cells under 1% O2 culture without reoxygenating the chamber, we instead mimicked hypoxic signaling chemically using cobalt chloride (CoCl2), which increases Hif-1α expression and reduces its proteasomal degradation (255). In both Atg5Ad.Ctrl and Atg5Ad.KO islet cells, the number of LysoTracker labeled organelles was reduced by CoCl2, and there was a tendency for this loss to be exacerbated in the Atg5-deficient cells (Figure 4.1a,b). In another preliminary experiment, we cultured Atg5Ad.Ctrl and Atg5Ad.KO islet cells under 1% O2 and investigated the lysosomal population using Lamp-1 immunostaining. Surprisingly, this 85 experiment suggested that Lamp-1 protein levels and the number of lysosomes (Lamp-1 positive puncta) were additively increased by hypoxia and Atg5 knockout (Figure 4.1c). Together, this indicates that lysosomal pH regulation, but not their numbers, is impaired in islet cells under hypoxic stress, and further suggests that autophagy may play a role in counteracting hypoxia-related lysosomal impairment. 86 Autophagosome formation and lysosomal degradation are both critical components of autophagic flux. We used Atg5 knockout and lysosomal inhibition to evaluate their individual and combined importance for islet-cell survival under hypoxia. As expected from our experiments in Chapter 3, Atg5 knockout amplified hypoxia-induced islet cell death. Interestingly, inhibition of lysosomal function by CHQ markedly worsened this loss of Atg5-deficient islet cells, but on its own it did not significantly increase death of Atg5Ad.Ctrl cells during the 24-hour period (Figure 4.2). The synergistic effect of Atg5 knockout and CHQ treatment could suggest that deletion of Atg5 and/or the resulting autophagy deficiency may not be complete, or that there are Atg5-independent autophagy pathways active in β-cells that then get inhibited upon CHQ treatment (256). It is also possible that the combined insults of Atg5 deficiency and impaired lysosomal function amplifies hypoxia-induced cell death by mechanisms such as lysosomal membrane permeabilization (LMP) (257). Irrespective of the exact mechanism, these results demonstrate a combined importance of sustained autophagy and lysosomal function for β-cell viability under hypoxic stress. Figure 4.1 – Effects of hypoxia on LysoTracker and Lamp-1 staining in Atg5Ad.Ctrl and Atg5Ad.Cre β-cells. a) Representative LysoTrackerTM staining in Atg5Ad.Ctrl and Atg5Ad.Cre islet cells with and without 5 hours CoCl2 treatment. b) Count of LysoTrackerTM positive puncta in Atg5Ad.Ctrl and Atg5Ad.Cre islet cells with and without 5 hours CoCl2 treatment. Data points represent individual cells and lines indicate the mean ± SEM (n=1, preliminary data). c) Lamp-1 immunostaining of Atg5Ad.Ctrl and Atg5Ad.Cre β-cells cultured 24 hours under 20% O2 (Normoxia) or 1% O2 (Hypoxia). Left: Mean Lamp-1 immunofluorescence intensity. Right: Count of Lamp-1 positive vesicles per cell area. Data points represent individual cells from one experiment (preliminary data set) and lines indicate the mean ± SEM. *p < 0.05, **p < 0.01, ***p < 0.001, ****p < 0.0001 by two-way ANOVA followed by Tukey’s multiple comparison test. 87 4.2.2. Hypoxia impairs the expression of β-cell Tfeb and its target genes To further probe if β-cell hypoxia causes lysosomal dysregulation, we quantified the effects of hypoxia on the expression of some key lysosome-related genes. Hypoxic culture reduced the islet mRNA levels of Lamp1, Lamp2 and Atp6v1h, which are known for having important roles in lysosomal membrane formation and luminal acidification (95) (Figure 4.3a). These genes are all known targets of Tfeb and Tfe3, and we found a similar tendency for both Tfeb and Tfe3 expression to be reduced (Figure 4.3b). As will be discussed in section 4.2.3, there is also a significant loss of Tfeb protein in hypoxic β-cells. Figure 4.2 – Individual and combined effects of β-cell Atg5 knockout and lysosomal inhibition on hypoxia-induced islet-cell death. a) Quantification of cell death progression based on Propidium Iodide incorporation into Atg5Ad.Ctrl and Atg5Ad.Cre β-cells under 20% O2 (Normoxia) or 1% O2 (Hypoxia) culture with or without CHQ treatment. Cell death is presented as the percentage of cell death above the Atg5Ad.Ctrl normoxia group at 24 hrs. b) Cumulative cell death (area under the curve; AUC) of each group shown in panel (b). Data represent mean ± SEM. n=3, *p < 0.05, **p < 0.01, ***p < 0.001, ****p < 0.0001 by one-way ANOVA followed by Tukey’s multiple comparison test. 88 To assess the role of Tfeb for lysosomal and autophagy gene expression under hypoxia, we used an adenoviral vector (Ad.Ins2>mTFEB:T2A:TagBFP2) to increase Tfeb levels specifically in islet β-cells. Compared to islets transduced with control virus (Ad.Ins2>TagBFP2), β-cell overexpression of Tfeb successfully restored, and in some instances further amplified, the expression of lysosome- and autophagy-related Tfeb targets in islets cultured under 1% O2 (Figure 4.4). Taken together, our data suggest that hypoxia may impair β-cell lysosomal function by disrupting Tfeb-dependent transcription of targets in the CLEAR gene network (116). Figure 4.3 – Hypoxia-induced changes to islet expression of select lysosome-related genes and lysosomal master regulators Tfeb and Tfe3 in C57BL/6J islets under normal and 1% O2 culture. Data represent mean ± SEM. n=4-6 independent experiments, *p < 0.05, **p < 0.01 by Student’s t test. 89 4.2.3. Inhibition of mTORC1 restores Tfeb, improves lysosomal function and reduces β-cell death under hypoxia To further explore the loss of Tfeb and its transcriptional targets, we used immunostaining to assess the changes in endogenous Tfeb protein. This revealed a significant loss of Tfeb immunoreactivity in both mouse and human β-cells under prolonged (24 hours) hypoxia (Figure 4.5a,b). Phosphorylation by mTORC1 keeps TFEB tethered in the cytosol, and mTORC1 inhibitors - including torin-1 and rapamycin - are known to allow TFEB dephosphorylation and Figure 4.4 – Tfeb overexpression increases expression of lysosome- and autophagy-related genes in hypoxic islets. Islets from male C57BL/6J mice were cultured for 16 hours under 20% O2 (normoxia), or under 1% O2 (hypoxia) with and without adenoviral Tfeb expression in their β-cells. Data represent mean ± SEM of 4 independent experiments. *p < 0.05, **p < 0.01, ***p < 0.001 by one-way ANOVA followed by Tukey’s multiple comparison test. Each gene was analyzed separately. 90 transcriptional activation (181) (Figure 4.5c,d). We tested the effects of torin-1 on hypoxic mouse β-cells and found that it rescued the loss of Tfeb (Figure 4.5a). This striking outcome is possibly the result of transcriptional autoregulation by activation of residual Tfeb (167). 91 Torin-1-induced restoration of Tfeb protein levels was paralleled by an upregulation of several Tfeb target genes in hypoxic islets (Figure 4.6). However, some of the transcripts we examined (Lamp1, Lamp2 and Atg5) were resistant to this effect, and even showed a tendency for down-regulation (Figure 4.6) despite being induced by Tfeb overexpression (Figure 4.4). The reason for this difference is not clear, but studies have suggested that other mTORC1 independent transcriptional regulators, like FOXO, could co-regulate gene transcription under certain stress conditions (258). It has also been reported that TFEB target genes are differentially sensitive to the amount of nuclear TFEB, and that high levels of TFEB activation can trigger a negative feedback mechanism that only affects some of its target genes (259). Figure 4.5 - Hypoxia reduces TFEB immunoreactivity in mouse and human β-cells but can be restored by torin-1. a) Left: Representative Tfeb immunostaining. Right: Quantified Tfeb immunofluorescence intensity in male C57BL/6J β-cells cultured under 20% O2 and under 1% O2 with and without torin-1 treatment (500 nM) for 24 hours. Grey markers represent the mean TFEB fluorescence values of individual β-cells (identified by insulin staining, not shown). Black markers and lines show the mean ± SEM of three independent experiments. b) Left: Representative TFEB immunostaining. Right: Quantified TFEB immunofluorescence intensity in human β-cells cultured under 20% O2 and 1% O2 for 24 hours. Grey markers represent mean TFEB fluorescence values of individual β-cells (identified by insulin). Black markers and lines show the mean ± SEM of cells preparations from 3 different donors. c) Schematic showing the role of phosphorylation and dephosphorylation on TFEB nuclear translocation and activation. Torin-1 inhibits mTORC1 and activates TFEB. d) Left: Representative images of torin-1-induced TFEB translocation in MIN6 cells transfected with TFEB-GFP. Right Quantified ratio of nuclear-to-cytosolic TFEB:GFP fluorescence. Data represent mean ± SEM of 42 Control and 29 torin-1 treated cells in a single test. *p < 0.05, ***p < 0.001 ****p < 0.0001. Data in panel (a) were analyzed by one-way ANOVA followed by Tukey’s multiple comparison test. Data in panels (b) and (d) were analyzed by Student’s t-test. 92 Interestingly, we found that Lamp-1 immunoreactivity tended to be stronger in the same hypoxic β-cells that we showed have Tfeb loss in Figure 4.5, and the Lamp-1 levels increased further when hypoxic cells were treated with torin-1 (Figure 4.7a). The elevated Lamp-1 protein levels did not reach statistical significance but are in striking contrast to the reduction in islet Lamp1 mRNA, and these experiments do agree with the preliminary data in Figure 4.1. Closer inspection showed clear localization of Lamp-1 immunoreactivity to vesicular membranes (Figure 4.7a; magnifications), which strongly suggests the staining is specific. It therefore appears that Lamp-1 protein levels in hypoxic β-cells are determined mainly by post-transcriptional mechanisms, such as altered turnover of lysosomal membranes. Figure 4.6 – Effects of Torin-1 treatment on expression of select Tfeb target genes in hypoxic islets. Islets from male C57BL/6J mice were cultured for 16 hours under 20% O2 (normoxia), or under 1% O2 (hypoxia) with and without torin-1 treatment (500 nM). Data represent mean ± SEM of 4 independent experiments. *p < 0.05, **p < 0.01, ***p < 0.001, ****p < 0.0001 by one-way ANOVA followed by Tukey’s multiple comparison test. Each gene was analyzed separately. 93 Unexpectedly, the same cell cultures also showed a pattern of hypoxia- and torin-1-dependent increases in insulin staining (Figure 4.7b). Autophagy and crinophagy can degrade proinsulin and insulin granules, and it was recently reported that TFEB and TFE3 suppress β-cell insulin expression (260). As such, the hypoxia-induced increase in insulin staining could result from the reduced autophagic flux and lower Tfeb levels. However, the further increase in response to torin-1 suggests other factors are also involved, including possible effects on insulin secretion. Figure 4.7 – Hypoxia and Torin-1 treatment increase β-cell Lamp-1 and insulin immunoreactivity. Islet cells from male C57BL/6J mice were cultured for 24 hours under 20% O2 (normoxia), or under 1% O2 (hypoxia) with and without torin-1 treatment (500 nM). a) Left: Representative Lamp-1 immunostaining. Images in the right column show zooms of the areas indicated by white squares in the left column. Right: Quantified Lamp-1 immunofluorescence intensities. Grey markers represent the mean Lamp-1 intensity values of individual β-cells. Black markers and lines show the mean ± SEM of three independent experiments. Scale-bars = 10 µm in the original images and 5 µm in zoomed images. b) Left: Representative Insulin immunostaining. Right: Quantified Insulin immunofluorescence intensities. Grey markers represent the mean Insulin intensity values of individual β-cells. Black markers and lines show the mean ± SEM of three independent experiments. Scale-bars = 10 µm. Increases in immunostaining intensities did not reach statistical significance by one-way ANOVA followed by Tukey’s multiple comparison test. 94 To determine if the reductions in Tfeb and target gene expression impaired lysosomes, we investigated the capacity for lysosomal degradation by assaying the activity of the lysosomal cysteine protease CTSB. As can be seen in representative images of cell populations in Figure 4.8a, and from the quantifications in Figures 4.8b and 4.8c, C57BL/6J mouse islet cells that were cultured for 24 hours in 1% O2, or exposed to CoCl2-induced chemical hypoxia for 8 hours, had reduced lysosomal hydrolase activity compared to control cells. Treatment with 250 nM torin-1 alleviated the 1% O2-induced loss of CTSB activity, which suggests that the torin-1-induced restoration of β-cell Tfeb and Ctsb expression leads to physiological improvements. Interestingly, CTSB activity was not improved in response to another mTOR inhibitor, rapamycin (Figure 4.8b). When we later obtained Tfebflox/flox mice (studied further in Chapter 5), we tested if Tfeb mediates the effect of torin-1. Culture in 1% O2 robustly decreased CTSB activity in Tfebflox/flox islet cells transduced with control adenovirus (TfebAd.Ctrl), and this loss was prevented by 500 nM torin-1 (Figures 4.8d,e). Notably, the restoration of CTSB activity by torin-1 was absent in cells with Tfeb knocked out by adenoviral expression of Cre (TfebAd.KO), demonstrating that it depends on Tfeb (Figure 4.8d,e). Together, these data show that hypoxic stress reduces β-cell Tfeb and impairs the expression of lysosomal and autophagy genes. This impairs lysosomal function, but nonetheless leaves the lysosomal population intact. 95 Figure 4.8 – Hypoxia impairs lysosomal cathepsin B activity in β-cells, but it can be restored by torin-1. a) Representative images showing Magic Red® fluorescent staining of cathepsin B activity in male C57BL/6J islet cells cultured under 20% O2 (normoxia control), or under hypoxia (1% O2 for 24 hours or CoCl2 for 8 hours) with and without torin-1 or rapamycin treatment. Scale bars = 100 µm. b) Normalized Magic Red® fluorescence intensity in groups with hypoxia induced by 1% O2 ± treatment with 250 nM torin-1 or 100 nM rapamycin. c) Normalized Magic Red® fluorescence intensity in groups treated with CoCl2 ± 1000 nM torin-1. d) Representative images showing the effects of 24 hours hypoxia (1% O2) ± 500 nM torin-1 on Magic Red® fluorescence in Tfebflox/flox islet cells 4 days after transduction with control adenovirus (TfebAd.Ctrl) or adenovirus expressing Cre (TfebAd.KO). Scale bars = 100 µm. e) Quantified Magic Red® fluorescence intensity in the experimental groups shown in panel (d). Cells from 4-6 randomly chosen fields from each group were analyzed and averaged. Data show the mean ± SEM of three experiments. *p < 0.05 by one-way ANOVA followed by Tukey’s multiple comparison test. 96 Our experiments in Chapter 3 suggested a possible interdependency of the adaptive UPR and autophagy, including at the transcriptional level. Intriguingly, TFEB and TFE3 also act as transcriptional inducers of the UPR in response to ER stress (161). This raises the possibility that loss of Tfeb might contribute to the impaired adaptive UPR in our hypoxic mouse β-cells. Indeed, we found that torin-1 increased Ern1 expression (i.e. Ire1α) and restored Xbp1 mRNA splicing in hypoxic islets (Figure 4.9a), which is compatible with this possibility. Moreover, torin-1 treatment successfully prevented islet cell death induced by 48-hours culture in 1% O2 (Figure 4.9b). It therefore appears that mTORC inhibition has favorable effects on Tfeb levels, lysosomal function and gene expression, which together have protective roles in hypoxic β-cells. Figure 4.9 – Torin-1 stimulates the UPR and reduces islet cell death under hypoxia. a) qPCR quantification of UPR-related gene expression in male C57BL/6J islets cultured under 20% O2 (normoxia) and 1% O2 (hypoxia) with and without 500 nM torin-1 for 16 hours. Data represent mean ± SEM of 4 independent experiments. b) Quantification of the fold increase in islet cell death during 48 hours culture under 20% O2 (normoxia), and 1% O2 (hypoxia) with and without 500 nM torin-1. Data represent mean ± SEM of 3 independent experiments. *p < 0.05, **p < 0.01 by one-way ANOVA followed by Tukey’s multiple comparison test. 97 Collectively, our experiments demonstrate that prolonged hypoxia impairs lysosomal function and disrupts protective autophagic clearance mechanisms in pancreatic β-cells. This dysregulation is associated with a reduction in β-cell Tfeb levels and in the expression of lysosomal and autophagic Tfeb target genes, both of which can be rescued by Tfeb activation. 4.2.4. Glucolipotoxicity- and ER stress-induced β-cell death involves loss of Tfeb To determine if loss of β-cell Tfeb is unique to hypoxia, or also occurs under other diabetes-relevant conditions, we examined Tfeb levels in MIN6 β-cell exposed to different degrees of palmitic acid (PA)-induced lipotoxicity. As expected, PA caused a dose- and time-dependent MIN6 cell death (Figure 4.10a). Lipotoxic stress also resulted in loss of Tfeb protein that was paralleled by activation of pro-apoptotic caspase-3 (Figure 4.10b). Because PA induces ER stress-dependent β-cell death, we compared the PA-treated groups to cells treated with thapsigargin (Tg), which triggers ER stress by inhibiting ER Ca2+ uptake and depleting luminal Ca2+ stores (261). Like PA treatment, Tg-induced ER stress caused a significant reduction in Tfeb and induction of cleaved Caspase 3 (Figure. 4.10b). We next tested if ER-stressed MIN6 cells could be protected by augmenting Tfeb levels or activity. Notably, both overexpression of GFP-tagged Tfeb and activation of Tfeb with torin-1 reduced Tg-induced MIN6 cell death (Figure 4.10c). Taken together, these data indicate that loss of endogenous Tfeb contributes to ER stress-induced β-cell death, and that preventing or reversing Tfeb loss could help protect β-cells under a variety of diabetes-relevant conditions that cause prolonged ER stress. 98 Figure 4.10 – Tfeb promotes MIN6 β-cell survival and is reduced under lipotoxicity and ER stress. a) Representative images (left) and kinetic analysis of palmitate (PA)-induced MIN6 cell death quantified by the percentage of propidium iodide (PI) positive cells (right; n=3). b) Representative western blot (top) and densitometry quantification (bottom) of Tfeb and cleaved caspase 3 in MIN6 β-cells under PA-induced lipotoxicity and thapsigargin (Tg)-induced ER stress (n=3; **p < 0.01, ***p < 0.005 compared to BSA control by one-way ANOVA) c) Comparison of Tg-induced death in Tfeb:GFPpos MIN6 cells (green bar), Tfeb:GFPneg MIN6 cells (black bar) and Tfeb:GFPneg MIN6 cells treated with torin-1 (blue bar) (n=3; *p < 0.05 by one-way ANOVA). 99 4.3 Discussion To shed light on the impairment of autophagic flux observed in Chapter 3, we investigated the impact of hypoxia on β-cell lysosomes. We found that hypoxic islet cells had reduced expression of genes related to lysosomal homeostasis and accumulated dysfunctional lysosomes. Importantly, we also discovered that β-cells exposed to hypoxia, lipotoxicity and Tg-induced ER stress showed a reduction in Tfeb. Restoring Tfeb levels, either genetically or pharmacologically, improved lysosome- and autophagy-related gene expression and lysosomal function, and reduced β-cell apoptosis caused by hypoxia or ER stress. Impaired lysosomal function and autophagic turnover have been observed in cultured β-cells exposed to prolonged (gluco)lipotoxicity (146,195,248) or pro-inflammatory cytokines (147), and in β-cells from patients with T1D and T2D (148,149,252). Our findings show that similar impairments happen in hypoxic β-cells: mRNAs involved in lysosomal degradative function and pH regulation are reduced, while LysoTracker staining and cathepsin B activity assays established that lysosomes become functionally impaired. Altogether, this likely underlies the hypoxia-induced defects in β-cell autophagosome clearance. Although lysosomal physiology was negatively affected, we found that the amount of vesicular Lamp-1 immunostaining increased, indicating that the lysosomal population grew. This effect was more pronounced in our Atg5-deficient islet cells, which aligns with key roles for ATG5 in the turnover of lysosomal membranes by micro-autophagy (262) and in the selective turnover of damaged lysosomes by macro-autophagy (i.e. lysophagy) (263). It is therefore likely that hypoxia-induced disruption of 100 β-cell autophagy causes toxic accumulation of both damaged lysosomes and autophagosomes. A key finding of this study was that prolonged hypoxia, lipotoxicity, and ER stress cause a loss of β-cell TFEB. Decreases in TFEB mRNA and/or protein have been reported in diverse cellular pathologies, including cardiac glucolipotoxicity (264), pancreatitis (265), ethanol-induced liver injury (186), Thiel-Behnke corneal dystrophy (172), and in the diabetic kidney (266). To our knowledge it has not previously been reported in diabetic or stressed pancreatic β-cells. Nuclear TFEB is reduced in β-cells from human cadaveric donors with T2D (252), but it is not clear if it reflects an absolute loss of TFEB, impaired nuclear translocation, or both. While our results demonstrate that β-cell Tfeb mRNA and protein levels are reduced under several types of sustained in vitro stress, Tfeb phosphorylation and translocation remain to be investigated under the same conditions. Irrespective, we showed that lysosome/autophagy genes and lysosomal hydrolase activity are linked to β-cell Tfeb signaling and become impaired under conditions where Tfeb is reduced. Restoring Tfeb mitigated these impairments, which strongly indicates that disruption of Tfeb drives the lysosomal dysregulation in hypoxic β-cells. It is well-established that pharmacological inhibition of mTORC1 can activate TFEB and stimulate autophagy. Here, we found that the potent mTORC1/2 inhibitor torin-1 prevented hypoxia-induced loss of Tfeb in mouse β-cells. TFEB can promote its own expression by binding to CLEAR sites in its promoter (167). Moreover, TFEB stimulates the expression of MCOLN1, which facilitates lysosomal calcium release and CaN-dependent TFEB activation (104). Together, these mechanisms establish a positive feedback loop whereby activation of TFEB can 101 increase its own transcription. We speculate that this may be a mechanism by which torin-1 treatment restored Tfeb mRNA and protein in our mouse β-cells. Nutrient excess and diabetes are associated with sustained mTOR hyperactivity (267), and this has been suggested to promote β-cell autophagic failure under gluco- and lipo-toxic conditions (248). Hypoxia, in contrast, is expected to activate AMPK and suppress mTOR activity (219) so the robust effects of mTOR inhibition in hypoxic β-cells are perhaps unexpected. We did not specifically examine mTOR signaling, but our results indicate that its activity in hypoxic β-cells is sufficient to negatively affect Tfeb and lysosomal function. Using Tfeb knockout islet cells, we established that torin-1 treatment requires Tfeb to restore lysosomal cathepsin B activity under hypoxia, likely because it restores Tfeb-dependent expression of Ctsb and other genes that maintain the luminal environment necessary for normal hydrolase activity. Rapamycin has been reported to stimulate β-cell autophagy and improve their function and survival under conditions of glucolipotoxicity and during ER stress due to genetic defects in proinsulin folding (131,248,268). Surprisingly, rapamycin did not improve CTSB activity in our hypoxic β-cells. One potential explanation for this difference is that torin-1 inhibits mTORC1 more completely than rapamycin (269) and is a more potent TFEB activator in many cell types (157). Of note, rapamycin and torin-1 have both been reported to increase basal and stress-induced β-cell death (147,242,267,270), possibly due to inhibition of mTORC2 (270). It is therefore remarkable that we exclusively observed protective outcomes of torin-1 treatment under the conditions we tested. The reason for this difference is unclear, and it will be important 102 to further examine mTOR signaling, and how it affects Tfeb in β-cells under hypoxia and other stresses, including also in human islets. That β-cell Tfeb levels decrease in response to prolonged hypoxia, lipotoxicity, and ER calcium depletion suggests it might be linked to unmitigated ER stress. Others have shown that 16 hours of exposure to 25 mM glucose + 0.5 mM palmitate impaired LysoTracker Red staining, cathepsin B activity and autophagic clearance in the INS-1E β-cell line (195). These impairments were partially prevented by the chemical chaperone TUDCA, and therefore downstream of ER stress signaling (195). The same study reported that Tfeb mRNA expression and nuclear Tfeb translocation were increased after 8 hours of glucolipotoxic stress, but Tfeb was not examined at the 16-hour time point when lysosomes were disrupted. Overall, these prior findings align with our assessments of Tfeb levels in palmitate-stressed MIN6 β-cells (Figure 4.10). In 0.5 mM palmitate we saw a tendency for Tfeb to increase at 8 hours and decrease after 16 hours. Stronger lipotoxicity (1.5 mM palmitate) accelerated the loss of Tfeb and worsened MIN6 β-cell death. While we did not test rescue strategies under lipotoxicity, Tfeb overexpression and torin-1 treatment both reduced MIN6 death under Tg-induced ER stress. These findings have to be verified in primary β-cells, but collectively, our results are compatible with the idea that ER stress activates TFEB and autophagy, but unmitigated maladaptive ER stress may eventually contribute to β-cell autophagy failure by reducing TFEB. The molecular mechanisms by which β-cell Tfeb mRNA and protein are reduced are unclear and could depend on the specific type of stress. As discussed, mTORC1-dependent 103 phosphorylation might suppress TFEB autoregulation, and TFEB phosphorylation has also been linked to its proteasomal degradation (271). In response to ER stress the UPR shuts down global protein translation, but because TFEB and TFE3 are activated as part of the PERK-dependent UPR they are likely among the proteins that become selectively translated (161). Because PERK signaling activates TFEB, and TFEB conversely helps sustain the adaptive UPR (161) (Figure 4.9), impairment of one could possibly inhibit the other in a vicious cycle. Existing reports suggest other mechanisms by which cellular TFEB levels can be reduced. Intriguingly, a number of microRNAs (miRNAs) regulate TFEB under stress (272). For instance, miR-30b-5p has been shown to suppress TFEB-dependent gene expression and autophagy (273), and it is induced in hypoxic cardiomyocytes (274). In studies of atherosclerosis, ER stress increased the expression of miR-33 (275), which also suppresses TFEB and a lot of its target genes (276). Other miRNAs that may down-regulate TFEB include miRNA-128, miRNA-29, and miRNA-300-3p (reviewed in (272)). Although we have not come across studies linking these specific miRNAs to stressed or diabetic β-cells, roles for miRNAs are possible and could include indirect effects via targets such as mTOR. Finally, it is also noteworthy that the tumor suppressor Programmed Cell Death 4 (PDCD4) can suppress TFEB translation (277), and is induced by hypoxia in MIN6 β-cells (278). These processes all warrant consideration as possible mechanisms for β-cell TFEB loss. Chapter 3 demonstrated that autophagy is important for the viability of syngeneic islet grafts, suggesting that boosting β-cell autophagy could improve islet graft function and transplantation outcomes. In this regard, it will be important to determine if TFEB and lysosomal 104 clearance are compromised in transplanted β-cells. Our in vitro hypoxia experiments in this study, and studies of proinflammatory β-cell stress done by others (147), suggest they might be. Therefore, strategies to drive autophagy should also aim to preserve or restore lysosomal function to avoid the cytotoxic effects of accumulating autophagosomes and damaged lysosomes. With the Edmonton Protocol, rapamycin (also known as sirolimus) became a key immunosuppressant used in clinical islet transplantation (14). Although rapamycin stimulates autophagy and can protect stressed β-cells under some conditions, there is overwhelming evidence that rapamycin has islet toxic effects that may contribute to loss of adequate graft function in the long-term (reviewed in (279)). It has been suggested that the β-cell toxicity of rapamycin involves excessive autophagy activation and intracellular digestion (242). Our data indicate that rapamycin is ineffective at promoting lysosomal function (Figure 4.8b). It remains to be tested if rapamycin, like torin-1, activates TFEB in β-cells. Conceivably, rapamycin’s β-cell toxicity may be exacerbated by it stimulating autophagosome formation without adequate priming of lysosomal clearance. It is interesting that another major immunosuppressive drug used in islet transplantation, the CaN inhibitor FK506 (tacrolimus), has islet-toxic effects linked to suppression of TFEB signaling (280), and that similar TFEB impairment has been reported for the kidney toxicity of the CaN inhibitor and immunosuppressant, cyclosporin A (281). TFEB shows promise as a therapeutic target. Its ability to promote both autophagy initiation and clearance makes it a compelling candidate for β-cell protection in diabetes and islet transplantation. Our data from this study, and recent work by other groups, support this potential. 105 In a screen for CaN activators, Lim et al. discovered a small molecule, MSL-7, that potently stimulated TFEB, promoted autophagy and improved the metabolic profiles of ob/ob and HFD-fed mice (175). Further studies demonstrated that TFEB activation by MSL-7 stimulates β-cell clearance of toxic hIAPP aggregates (196). Directly related to existing diabetes treatments, the GLP1R agonist exendin 4 stimulates β-cell autophagy by activating TFEB in a calcium and CaN-dependent manner (176). Because of the complexities and potential toxicity of directly manipulating signaling nodes like mTOR and CaN, other possibilities are being explored for TFEB activation. Interesting examples include trehalose, which can stimulate TFEB by inducing a low level of lysosomal stress (282), and curcumin analog C1, a small molecule that binds TFEB at the N terminus and directly promotes its nuclear translocation (283). Testing if such mTOR-independent compounds can activate and/or restore TFEB and autophagy in stressed β-cells may identify new possibilities for preserving their function and survival. Clinical transplantation of donor- or stem cell-derived islets may benefit most readily from new β-cell protective drugs because grafts can be targeted locally using drug eluting microparticles (284), which minimizes systemic effects of the treatment. To summarize, this study demonstrated that hypoxia reduces TFEB levels and causes lysosomal failure in β-cells. TFEB loss also occurs under severe glucolipotoxicity and ER stress, correlating with increased β-cell apoptosis. Enhancing TFEB, either genetically or pharmacologically, improves lysosomal function and β-cell survival, suggesting that TFEB loss contributes to β-cell failure under chronic stress. Our results highlight the potential of restoring 106 or activating TFEB as strategies for β-cell protection in diabetes and islet transplantation. Future research is needed to understand the molecular mechanisms of TFEB signaling and dysregulation in stressed β-cells. 107 Chapter 5: β-Cell-specific Tfeb knockout has minor and sex-specific effects on glucose tolerance: Possible compensation by Tfe3. 5.1 Background In Chapter 4, we determined that lysosomes in hypoxic β-cells become impaired because of Tfeb downregulation and dysregulation of gene expression. Importantly, loss of β-cell Tfeb also happened under lipotoxicity and chemically induced ER stress. Here, we establish a new mouse model with conditional Tfeb deletion to directly test the impact of Tfeb deficiency on β-cell function and glucose homeostasis under basal and stressed conditions in vivo. Despite the increasing evidence for in vivo β-cell lysosomal and autophagic failure in diabetes, the underlying causes are unknown. One study has reported lower nuclear TFEB staining in β-cells of pancreas sections from human donors with T2D (252). However, little is known about the consequences of β-cell TFEB deficiency, and it remains to be determined if dysregulation of TFEB contributes to β-cell failure in diabetes pathogenesis. Loss of TFEB might affect both β-cell viability and physiology. For instance, lysosomal degradation of proinsulin and insulin secretory granules affects insulin secretion (129,130,246), and Pasquier. et al. recently made the surprising discovery that TFEB suppresses insulin gene transcription (260). While conducting the work in this chapter, the first report of a conditional β-cell Tfeb knockout mouse was published (40). In this paper, Park et al. found that Tfeb is important for activation of β-cell mitophagy in response to mitochondrial stress in vitro. When fed a HFD, their Tfeb knockout mice showed impaired diet-induced β-cell mitophagy, exacerbated β-cell failure and 108 diabetes (40). This paper provided detailed insights into Tfeb-dependent processes in the β-cell, but Tfeb was knocked out using the RIP2-Cre mouse model. As discussed previously, this is likely to have introduced off-target deletion, thus leaving it unclear what the impact of β-cell-specific Tfeb deficiency is in vivo. Transcription factor targeting of genes is sex-biased across human tissues (285), and recent studies indicate that TFEB-dependent signaling might show sex differences. For instance, a study found that Tfeb protein levels differ in muscle from male and female mice (286). In human cells, the androgen receptor has been reported to bind and function as a co-activator of TFEB (287,288). This suggests some degree of sex-dependent modulation of TFEB transcriptional activity. It is not known if β-cells show male-female differences in TFEB-dependent transcription or if there are sex differences in the outcomes of β-cell TFEB deficiency. In this chapter, we establish a novel mouse with Ins1Cre-driven recombination to evaluate the impact of β-cell-specific Tfeb knockout on glucose tolerance, including in models of STZ- and diet-induced β-cell stress. In the latter, we consider the potential for sex differences in Tfeb-dependent phenotypes. Motivated by results that indicate possible Tfe3-dependent compensation in our HFD-fed mice with β-cell Tfeb knockout, we conduct preliminary investigations of gene expression in islets with single and double β-cell Tfeb and Tfe3 deficiency. 109 5.2 Results 5.2.1. β-Cell-specific Tfeb knockout mice do not develop glucose intolerance As detailed in Chapter 2, we obtained Tfebflox/flox mice from Dr. Andrea Ballabio (TIGEM, Italy) (155) and bred these with Ins1Cre knock-in mice to establish a colony of β-cell-specific Tfeb knockout mice (TfebβKO) and littermate Tfeb wildtype controls (TfebβWT). The islets from our TfebβKO mice showed ~75% loss of Tfeb mRNA (Figure 5.1a), as well as a significant loss of Tfeb protein, as seen by islet western blot and immunostaining of dispersed β-cells (Figure 5.1b,c). Figure 5.1 – Confirmation of Tfeb knockout and C57BL/6 Nnt genotype a) qPCR confirmation of Tfeb mRNA loss in pancreatic islets from 12–30-week-old TfebβKO and TfebβWT littermates. Residual Tfeb mRNA in TfebβKO islets is likely from non-β-cells. Data represent mean ± SEM, ***p < 0.001 by paired Student’s t-test. (n=4; 3 female 3 and 1 male) b) Confirmation of Tfeb protein loss by Western blot in pancreatic islets from 30-week-old female TfebβKO and TfebβWT littermates. c) Confirmation of Tfeb protein loss by immunofluorescent staining in dispersed islet cells from 12–15-week-old TfebβKO and TfebβWT littermates. Data represent mean ± SEM. **p < 0.01, by paired Student’s t-test. (n=3; 2 female (red) & 1 male (blue)). d) Representative PCR genotyping of Nnt allele of a TfebβKO mouse and control mice with known C57BL/6N (6N) and C57BL/6J (6J) genetic background. Amplicon size: Nnt wildtype allele – 579 bp (6N); Nnt mutant allele – 743 bp (6J). –ve: no template control. 110 Ins1Cre and Tfebflox/flox mice are on the C57BL/6J and C57BL/6N backgrounds, respectively. This means that they carry wildtype (B6N) and mutant (B6J) nicotinamide nucleotide transhydrogenase (Nnt) genes, and this difference has been associated with differences in metabolic phenotypes and related gene expression (289). Our mice were bred such that our colony became stable on B6N (Figure 5.1d), and all animals used in the following were therefore on the B6N background. We tracked body weight gain, fasting blood glucose, and glucose tolerance in chow diet-fed TfebβWT and TfebβKO mice of both sexes between 8 and 27 weeks of age. Interestingly, we found that the deletion of β-cell Tfeb did not worsen glucose tolerance and even improved it in older TfebβKO mice (Figure 5.2a). As they aged, TfebβKO and TfebβWT littermates also showed similar weight gain and had comparable fasting glucose levels, with the possible exception of a slight improvement in older TfebβKO males (Figure 5.2b,c). Insulin sensitivity did not appear to differ between 29-week-old TfebβKO and TfebβWT littermates, but a larger sample size is needed to draw firm conclusions regarding insulin tolerance (Figure 5.2d). Our IPGTTs were done after a 6-hour fast, starting in the morning. Because Tfeb is activated under low nutrient conditions, we asked if the glucose tolerance phenotypes may differ after a longer overnight fast. Overnight fasting did not cause any appreciable changes to the glucose tolerance profiles of older female and male mice; TfebβKO mice still showed similar or slightly improved glucose tolerance (Figure 5.2e). Taken together, our data show that β-cell-specific deletion of Tfeb does not cause detrimental metabolic phenotypes in chow-fed mice. 111 112 We next examined the function of β-cells from female TfebβWT and TfebβKO mice ex vivo. Using the Magic Red assay, we compared lysosomal cathepsin B activity and found that it was reduced in TfebβKO β-cells (Figure 5.3a). In contrast, there were no major differences in the overall function of TfebβKO and TfebβWT islets; preliminary imaging experiments did not show obvious differences in glucose-stimulated cytosolic Ca2+ changes (Figure 5.3b), and although TfebβKO islets tended toward lower glucose- and KCl-stimulated insulin secretion, this was not significant (Figure 5.3c). This suggests that although Tfeb deletion impairs lysosomal hydrolase activity (and possibly other aspects of lysosomal physiology), Tfeb is not required for overall glucose-stimulated β-cell function. This is consistent with the observation that our TfebβKO mice did not have impaired glucose tolerance. Figure 5.2 – Body weight, IPGTT and ITT in chow-fed male and female TfebβWT & TfebβKO mice a) Intraperitoneal glucose tolerance tests (IPGTTs) of 17- and 27-week-old female (top) and male (bottom) TfebβKO and TfebβWT mice after 6 hours fasting. The rightmost panels summarize the IPGTT tests done between 8 and 27 weeks of age as the area under the curve (AUC). b) Fasting glucose of female (top) and male (bottom) TfebβKO and TfebβWT mice from 8-27 weeks of age. c) Body weight of female (top) and male (bottom) TfebβKO and TfebβWT mice from 8-27 weeks of age. d) Insulin tolerance test of 29-week-old female (top) and male (bottom) TfebβKO and TfebβWT mice. e) IPGTTs of 27–33-week-old female and male TfebβKO and TfebβWT mice after overnight fasting. Data are shown as mean ± SEM. *p < 0.05, **p < 0.01, ***p < 0.001 by repeated measures 2-way ANOVA and Sidak’s multiple comparison tests (Panels (a)-(c), n=6-10 females and n=5-8 males; Panel (d), n=2 females and n=3 males; Panel (e), n=5 females and n=4 males). 113 5.2.2. β-Cell Tfeb knockout sensitizes to MLD STZ-induced glucose intolerance Although we saw no major phenotypes related to β-cell function or glucose tolerance in chow-fed TfebβKO mice, this does not exclude a role for Tfeb in β-cell adaptation under conditions of stress. To explore this in vivo, we compared the glucose tolerance of male TfebβWT and TfebβKO mice after multiple low-dose (MLD) STZ injections, which induces inflammatory and ER stress in their β-cells (198,199). This experiment included age-matched Ins1Cre mice as additional controls. Before and after MLD STZ treatment, the body weights were comparable between all groups (Figure 5.4a). However, the TfebβKO mice showed an accelerated loss of Figure 5.3 – TfebβKO mice have reduced islet-cell CTSB activity but normal islet function a) Quantitative comparison of cathepsin B activity in islet cells from 16–18-week-old female TfebβWT and TfebβKO mice. Data represent mean ± SEM of 3 independent experiments analyzed by paired Student’s t-test, **p < 0.01. b) Preliminary Fura-2 fluorescence imaging of cytosolic Ca2+ responses in islets from 18-week-old female TfebβWT and TfebβKO mice. Traces represent recordings of 3 WT islets and 16 KO islets obtained from 4 animals; 1 TfebβWT and 3 TfebβKO). c) Static incubation assays quantifying insulin secretion in islets from 15-18-week-old female TfebβWT and TfebβKO mice. Data represent mean ± SEM of 3 independent experiments analyzed by two-way ANOVA and Sidak’s multiple comparison tests. 114 glucose control and impaired glucose tolerance compared to the control mice (Figure 5.4b,c). These data indicate that TFEB may have a critical role in β-cell adaptation under conditions related to strong ER and inflammatory stress in vivo (198,199). 5.2.3. β-Cell Tfeb knockout does not worsen glucose tolerance in HFD-fed mice We next investigated the role of β-cell Tfeb in the setting of T2D-relevant metabolic stress by feeding male and female TfebβKO and TfebβWT mice a high-fat diet (HFD) or control diet (CD) starting at 8 weeks of age (see Chapter 2 for diet details). Animals of both genotypes gained weight on HFD as expected, and no differences in weight gain or random and fasted blood Figure 5.4 – TfebβKO mice are sensitized to multiple low-dose STZ-induced hyperglycemia Male littermates of 13-17 weeks old TfebβKO and TfebβWT, and age-matched Ins1Cre mice were injected with multiple low dose Streptozotocin (STZ, 50 mg/kg) for 5 continuous days and monitored for blood glucose and body weight. Glucose tolerance was assessed by IPGTT at experimental endpoint. a) Body weight of TfebβKO and TfebβWT, and age-matched Ins1Cre mice post MLD-STZ administration. b) Random blood glucose measurements of TfebβKO, TfebβWT, and age-matched Ins1Cre mice within two weeks of the first STZ injection. c) IPGTTs performed at the experimental endpoint – i.e. 14 days after the first STZ injection or when the random blood glucose exceeded 20 mM in 2 consecutive tests. Data are shown as mean ± SEM; n=4 mice of each genotype, *p<0.05, **p<0.01 by repeated measures two-way ANOVA and Sidak’s multiple comparison test. 115 glucose levels were observed between genotypes (Figure 5.5a). Along with increased body weight, there was a progressive impairment of glucose tolerance in both sexes. However, β-cell Tfeb KO did not worsen the glucose intolerance in either male or female animals - in HFD-fed female mice, it even transiently improved glucose tolerance compared to littermate controls (Figure 5.5b). While this difference was modest in size, it was robust across several cohorts of mice. Interestingly, this was not seen in male TfebβKO mice, hinting at sex differences in the role of β-cell Tfeb during early adaptation to HFD-induced stress. There were no genotype-dependent differences in insulin tolerance in mice of either sex after 5 or 17 weeks of HFD-feeding (Figure 5.5c), suggesting the glucose tolerance differences seen in TfebβWT and TfebβKO females may be connected to β-cell function. 116 117 To evaluate the combined impacts of Tfeb deletion and diet-induced stress on β-cell function, we first compared ex vivo Ca2+ responses and insulin secretion in islets from TfebβWT and TfebβKO after 17 weeks on the HFD. When exposed to a stepwise ramp increase in extracellular glucose, islets from HFD-fed male and female mice responded with progressively larger cytosolic Ca2+ influx, starting at 5 mM glucose (Figure 5.6a). In the presence of 5 mM glucose, the average Ca2+ levels of both female and male TfebβKO islets were modestly higher than those of TfebβWT islets. However, the total Ca2+ entry at each glucose step – quantified as the area under the curve – did not show significant differences between the genotypes or the sexes (Figure 5.6b). Figure 5.5 – β-Cell Tfeb knockout does not worsen high-fat diet-induced glucose intolerance a) Body weight, random and fasting glucose of female and male TfebβKO and TfebβWT mice fed a high-fat diet (HFD) or control diet (CD) for 16 weeks, starting at 8 weeks of age. b) Intraperitoneal glucose tolerance tests of female and male TfebβKO and TfebβWT mice at 0, 4, 8 and 16 weeks on HFD or CD. c) Insulin tolerance test of female and male TfebβKO and TfebβWT mice after 5 or 17 weeks HFD or CD. Data are mean ± SEM. n=12-14 females on HFD and n=3-7 on CD; n=9 males on HFD and n=3 on CD, *p < 0.05, **p < 0.01, by repeated measures two-way ANOVA followed by Sidak’s multiple comparison tests. 118 Insulin secretion also did not differ between TfebβWT and TfebβKO islets in both sexes. Females showed a notable tendency toward better islet secretory function, with lower basal and higher stimulated insulin release, compared to males, but the trends in absolute insulin levels and stimulation indices did not reach statistical significance (Figures 5.6c,d). Given that Tfeb has been reported to facilitate β-cell mitophagy and sustain mitochondrial health under HFD-induced metabolic stress (40), we examined mitochondrial respiration in Figure 5.6 – Ca2+ signaling and insulin secretion in islets from HFD-fed TfebβWT & TfebβKO mice a) Cytosolic Ca2+ traces recorded in islets from TfebβWT and TfebβKO mice after 17 weeks on HFD. Left: average traces of 13 TfebβWT & 11 TfebβKO islets from 3 female mice of each genotype; 3-6 islets from each mouse. Right: average traces of 9 TfebβWT and 10 TfebβKO islets from 3 male mice of each genotype; 2-5 islets from each mouse. b) Cumulative Ca2+ entry quantified by the area under the curve at each step of the ramp in panel (a). c) Glucose- and KCl-stimulated insulin secretion in static incubations of batches of 15 islets. d) Stimulation index (stimulated secretion relative to basal secretion) calculated for each mouse in panel (c). Data in panels (b)-(d) represent mean ± SEM and were analyzed by two-way ANOVA and Sidak’s multiple comparison tests (n=5 female and n=4 male mice of each genotype). 119 whole islets from our HFD-fed TfebβWT and TfebβKO mice using Seahorse XFe24-based measurements of oxygen consumption rates. Using a combination of glucose stimulation and a Seahorse “Mito Stress Test” (Figure 5.7a), we quantified the basal, glucose-stimulated and ATP-coupled oxygen consumption, as well as mitochondrial proton leak and spare respiratory capacity (290) (Figure 5.7b). Although islets from our HFD-fed TfebβKO mice of both sexes showed trends toward reduced mitochondrial function, none of the respiratory measures were significantly impaired by Tfeb deficiency (Figure 5.7). Overall, this finding is consistent with our other ex vivo measures of Figure 5.7 – Mitochondrial respiration in islets from HFD-fed TfebβWT & TfebβKO mice. a) Mean Seahorse-based oxygen consumption rates (OCR) in islets from female and male TfebβKO and TfebβWT mice after 18 weeks on a high-fat diet. b) Quantification of basal, glucose-stimulated and ATP-linked respiration, as well as mitochondrial proton leak and spare respiratory capacity, based on the glucose stimulations and Mito Stress Tests in panel (a). Data represent mean ± SEM. Statistical comparisons were done using Student’s t-test (n=4 female and n=5 male mice of each genotype). 120 islet function, but it is in notable contrast to the respiratory dysfunction of HFD-fed Tfeb knockout islets reported by Park et al. (40). 5.2.4. Sex-dependent transcriptional changes and potential Tfe3-mediated compensation in HFD-fed TfebβKO mice Since we did not detect obvious phenotypic differences in the HFD-fed TfebβKO and TfebβWT animals or their ex vivo islets, we asked whether deletion of Tfeb affected the expression of known gene targets and if there may be compensatory changes to the functionally overlapping family member Tfe3. We started by comparing Tfe3 mRNA levels in islets from HFD-fed TfebβKO and TfebβWT mice and indeed detected transcriptional induction of Tfe3 following Tfeb knockout (Figure 5.8). We further compared the expression of select genes in the CLEAR network and observed several intriguing changes associated with lysosomal function and autophagy. While the levels of Ctsb, Atg5 and Atp6v1h were reduced in TfebβKO islets, as would be expected, other lysosomal genes increased, including Lamp1 and Mcoln1 (Figure 5.8). 121 Importantly, there were notable sex differences in these transcriptional changes. Overall, male knockout islets seemed more resistant to the loss of lysosomal genes, and Mcoln1 even showed opposite responses with induction in males and loss in female TfebβKO islets. Because Tfeb and Tfe3 have also been associated with mitochondrial function, including via PGC-1α (167,291,292), we further tested PGC-1α-regulated genes related to the mitochondrial electron transport chain. Also, there were apparent sex-dependent changes; Sdhb, Cox4l1 and Atp5e were all upregulated in females, while male islets only showed a smaller increase in Atp5e expression (Figure 5.8). 0123Islet mRNA (Rel WT)TfebTfe3Lamp1CtsdAtp6v1hMcoln1Atg5Map1lc3blysosomes autophagosomesSdhbCox4l1Atp5eMitochondria************************* ****** ***** ****** *********Female TfebβWTFemale TfebβKOMale TfebβWTMale TfebβKOFigure 5.8 – Tfe3 up-regulation and sex-specific effects on lysosomal and mitochondrial transcriptional adaptation to diet-induced stress in islets from HFD-fed TfebβKO mice. qPCR analysis of Tfeb, Tfe3 and select target genes related to lysosomal function, autophagy and mitochondrial oxidative phosphorylation in islets from female and male TfebβKO and TfebβWT mice after 18 weeks on HFD. Expression levels are normalized to TfebβWT of the same sex. Data represent mean ± SEM; n=6 females and n=5 males of each genotype, *p < 0.05, **p < 0.01, ***p < 0.001, ****p < 0.0001. Each gene was analyzed separately by one-way ANOVA followed by the Tukey multiple comparison test. 122 Next, we used immunofluorescence staining to determine how the transcriptional changes were related to Tfe3 and Lamp1 protein levels. In agreement with our gene expression data, Tfeb knockout increased Tfe3 and Lamp1 immunoreactivity in dispersed β-cells from male and female HFD-fed mice (Figure 5.9). At the same time, there was an increase in the ratio of nuclear-to-cytosolic Tfe3 immunoreactivity, indicating that Tfe3 nuclear translocation and transcriptional activity were enhanced in TfebβKO β-cells (Figure 5.9a,b). Interestingly, the changes to Tfe3 levels and translocation were more prominent in males than in females. More work is needed to determine the significance of these findings, but we believe that Tfe3 may be partially compensating for the loss of Tfeb and causally linked to some of the sex differences in gene expression that we observe following β-cell Tfeb KO. Figure 5.9 –Tfe3 and Lamp1 immunostaining in β-cells from HFD-fed TfebβKO & TfebβKO mice. a) Representative Tfe3 and Lamp1 immunostaining in β-cells from female and male TfebβWT and TfebβKO mice after 18 weeks on a high-fat diet. All β-cells were identified by insulin staining (not shown). b) Quantification of β-cell Tfe3 and Lamp1 immunostaining intensities. Tfe3 staining intensity was quantified as a whole-cell average and separately within the nucleus (identified by DAPI staining) and cytosol (non-nuclear area). Lamp1 staining intensity was quantified in the cytosol. Data represent mean ± SEM on a per-cell basis. *p<0.05, ****p<0.0001 by one-way ANOVA followed by the Tukey multiple comparison test (n=31 to 39 cells from 3 females and 3 males of each genotype). 123 To explore possible Tfe3 compensation in Tfeb-deficient β-cells, we designed adenoviral vectors for β-cell expression of Cre alone or in combination with Tfe3 shRNA. For identification of β-cells, these vectors, and a third shRNA[scramble] control vector, also express a TagBFP2 reporter under the rat Ins2 promoter (Figure 5.10a,b). By transducing isolated Tfebflox/flox islets with the vectors, we generated islets with normal Tfeb / Tfe3 expression, single β-cell Tfeb KO, and combined β-cell Tfeb KO / Tfe3 knockdown. In two preliminary tests, we transduced male islets with different MOIs and found that β-cell Tfeb KO increased islet Tfe3 mRNA levels. Moreover, short-term Tfeb KO suppressed the expression of Ctsb, Cstd and Atp6v1h, whereas the levels of Lamp1, Lamp2 and Mcoln1 increased (Figure 5.10c). The gene expression changes induced by in vitro Tfeb KO were very similar to our results in islets from male HFD-fed TfebβKO mice (Figure 5.8). Importantly, concurrent Tfe3 knockdown prevented the amplification of Lamp1, Lamp2 and Mcoln1 after Tfeb KO. The other transcripts Ctsb, Cstd and Atpv1h, were further reduced by combined Tfeb and Tfe3 deficiency (Figure 5.10c). In summary, we show that Tfe3 is upregulated in Tfeb knockout β-cells in vitro and under HFD-induced stress in vivo. Our data strongly support the possibility that Tfe3 may drive compensatory expression of a subset of genes for lysosomal biogenesis and function in Tfeb-deficient β-cells. Moreover, we see indications that the Tfeb/Tfe3-dependent regulation of these transcripts shows sexual dimorphisms in β-cells. Further work is warranted to generate a more complete picture of the Tfeb- and Tfe3-regulated transcriptional networks and their mutual roles for β-cell adaptation in diabetes. 124 Figure 5.10 – Tfe3 upregulation and partial compensatory gene expression in the absence of Tfeb a) Maps of the adenoviral vectors used for β-cell expression of Cre alone or in combination with shRNA against Tfe3, as well as a control vector expressing a scrambled shRNA. All vectors also express the fluorescent protein TagBFP2 under the rat Ins2 promoter to identify β-cells. b) Representative images from a confocal z-stack showing TagBFP2 fluorescence in a whole islet transduced with AdV.Ins2>TagBFP2:Cre vector. Scale bar = 30 µm. c) Preliminary qPCR analysis of Tfeb, Tfe3, and other select target genes related to lysosomal function in islets from ~15-week-old male Tfebflox/flox mice 4 days following transduction with the three adenoviral vectors shown in panel (a). (n=1 male mouse at each MOI). 125 5.3 Discussion In this study, we generated a mouse with Ins1Cre-driven Tfeb knockout and examined the importance of β-cell Tfeb for glucose tolerance, including under in vivo stress. We found that β-cell Tfeb KO worsened the loss of glucose tolerance in MLD-STZ-treated mice. However, in striking contrast to recent reports, we did not detect any genotype-specific glucose intolerance phenotype or worsened β-cell dysfunction when our TfebβKO animals were fed HFD for 17 weeks. Islets from our HFD-fed TfebβKO mice showed increased expression of Tfe3, as well as select genes associated with lysosomal, autophagic and mitochondrial function. Gene expression changes in islets with combined β-cell Tfeb KO and Tfe3 knockdown suggested that Tfe3 partially compensates for the transcriptional defects in Tfeb-deficient β-cells. Finally, we found that islets from HFD-fed mice showed sex differences in the gene expression changes resulting from Tfeb KO. Full knockout of Tfeb causes embryonic lethality in mice (293), but tissue-specific knockout of Tfeb has been widely used in animal models. In mice, conditional deletion of Tfeb caused metabolic defects in multiple tissues, including the liver and muscle, suggesting that Tfeb is directly involved in metabolic processes in addition to lysosomal functions (162,167,189). Considering this, we hypothesized that β-cell Tfeb knockout would lead to impaired β-cell function and glucose intolerance phenotypes. However, our observations did not support this - compared to their TfebβWT littermates, female TfebβKO mice showed a transient improvement in their glucose tolerance at the early stages of HFD feeding. While we were doing this work, 126 Park et al. published a paper reporting that Tfeb KO impaired β-cell mitochondrial homeostasis and caused markedly worsened glucose tolerance when the mice were fed a HFD (40). The reason for our conflicting results is unclear but may be related to differences in experimental models and design. First and foremost, they used the RIP2-Cre transgenic mouse, which raises the same concerns of confounding effects from off-target hypothalamic recombination we discussed in Chapter 3. The diverging phenotypes could also be related to the genetic background of the mice. Differences in glucose metabolism have been identified in C57BL/6 sub-strains (289). Mice on the B6/J background have a mutation in the nicotinamide nucleotide transhydrogenase (Nnt) gene, which is associated with mitochondrial redox homeostasis, and B6/J animals have lower insulin secretion and worse HFD-induced glucose intolerance (289). Nnt genotyping established that our Tfeb mouse colonies are stable on the B6/N background, i.e. Nnt wildtype. The genetic makeup of the Tfeb KO mice in Park et al. is unclear, but because RIP-Cre mice from Jax are on a B6/J background, they could be B6/J or mixed J/N, and thus more susceptible to HFD-induced β-cell stress and glucose intolerance than our mice. Other experimental factors may also lead to differences in the results. For instance, we performed IPGTTs in the afternoon after a 6-hour fast, whereas Park. et al. fasted their mice overnight. However, since overnight fasting did not alter the overall glucose tolerance phenotype of our TfebβKO mice (Figure 5.5) this is an unlikely explanation for the major differences between our Tfeb knockout mice and theirs. Our data suggest that the lack of β-cell dysfunction and glucose intolerance phenotypes in 127 our TfebβKO mice may involve transcriptional compensation by Tfe3. TFE3 and TFEB form homo- and hetero-dimers; they are both inhibited by mTORC1 phosphorylation, and both regulate genes in the CLEAR network (294). Although their cellular functions overlap, studies indicate TFE3 and TFEB are cooperative rather than fully redundant; overexpression of TFEB only partially rescued defects associated with TFE3 knockout, and the presence of both is required for maximal effects in regulating lysosomal biology and cellular stress response (113,160,161). Ex vivo, islets from our 17-weeks HFD-fed TfebβKO mice had increased expression of Tfe3 and a subset of shared gene targets, and we also saw some support for this at the protein level by immunostaining. Adenovirus-mediated β-cell Tfeb knockout also increased Tfe3 expression in short-term islet cultures without added stress factors. In our hands, Tfe3 up-regulation is, therefore, a robust feature of β-cell Tfeb knockout, and our shRNA knockdown experiments showed that Tfe3 compensates for some of the transcriptional defects associated with lack of Tfeb. Considering these results, it is surprising that islets from Park’s mice with β-cell Tfeb KO did not have increased Tfe3 mRNA compared to the Tfeb WT controls (40). Moreover, HFD-feeding stimulated islet Tfe3 expression in their Tfeb WT mice but not in their Tfeb KO mice (40). The differences between our results could be reconciled if Tfe3-dependent compensation only occurs under moderate stress and our Tfeb-deficient β-cells are less stressed than Park’s (e.g. due to their Nnt genotype). This has yet to be formally tested, but our data indirectly support the former. In contrast to what we saw in response to HFD feeding, our TfebβKO mice were sensitized to the more acute and severe β-cell stress induced by MLD-STZ 128 injections. In Chapter 4, we found that both Tfeb and Tfe3 expression were reduced in islets under chronic hypoxic stress. This indicates a lack of mutual compensation that might explain why hypoxic β-cells develop lysosomal dysfunction and impaired autophagic flux. Altogether, this suggests that careful evaluation of both Tfeb and Tfe3 is warranted in various diabetes-related models and stress conditions. Work is underway in our lab to investigate these questions using viruses for single and double β-cell Tfeb/Tfe3 deficiency in vitro and in vivo. Another important question we explored was possible sex differences in Tfeb-dependent β-cell stress responses. We observed that HFD-fed female TfebβKO mice showed a transient period with better glucose tolerance than their HFD-fed TfebβWT littermates, but this was not seen in males. Ex vivo, our HFD-fed male and female TfebβKO mice showed different changes in islet gene expression relative to their sex-matched TfebβWT littermates. Female TfebβKO islets specifically upregulated their expression of mitochondrial genes, whereas male TfebβKO islets tended to be more resistant to the loss of lysosomal genes than female TfebβKO islets. Of note, the Tfeb target Mcoln1 responded oppositely in male and female HFD-fed TfebβKO islets. The Mcoln1 gene encodes an ion channel that regulates lysosomal calcium-mediated activation of CaN, and thus, indirectly, the activity of both Tfeb and Tfe3 (103,104,161). This mechanism was found to be important for the activation of Tfeb-regulated β-cell mitophagy under mitochondrial stress, but this was only examined in male mice (40). Based on our data, it seems possible that suppression of Mcoln1 may limit Tfe3 activation and compensation of lysosome-related genes in our female TfebβKO β-cells, whereas high expression of Mcoln1 promotes it in 129 males. Moreover, sex hormones have been linked to the regulation of CaN expression (295). It is, therefore, an interesting question for further study how much males and female β-cells differ in their Tfeb- and Tfe3-dependent gene expression and whether the Mcoln1-CaN axis may be an important node for these sex differences. In summary, we generated what we believe is currently the most β-cell specific Tfeb knockout mouse. Our examinations of these mice indicate that β-cell Tfeb is dispensable for normal glucose tolerance under HFD-induced metabolic stress. This contradicts a recent report, and we speculate this may reflect critical differences in the Cre-driver and/or in Nnt genotype and β-cell stress levels. Our data further indicate that Tfe3 is upregulated and promotes the expression of a subset of shared Tfeb/Tfe3 gene targets when Tfeb is deleted in β-cells. More work is needed to determine the possible context-dependent nature of this relationship, to identify the scope of this transcriptional network, and to establish the consequences of this for β-cell stress responses in vitro and in vivo. Finally, we have provided the first comparison of male and female β-cell Tfeb knockout mice. We provided compelling evidence that sexually dimorphic transcriptional responses occur in the β-cells of our HFD-fed Tfeb knockout mice. The extent of these sex differences is an important question for future investigation, and overall, our data support the need for more investigation of biological sex as a factor in β-cell stress, adaptation and failure. 130 Chapter 6: Conclusion 6.1 Research summary The overall goal of this thesis was to determine the role of autophagy and lysosomal homeostasis in pancreatic β-cells under conditions of diabetes- and transplantation-relevant stress. In Chapter 3, we first used Ins1Cre knock-in mice to drive β-cell-specific Atg5 deletion, avoiding the potential for Ins2-driven off-target recombination that is a caveat of prior studies on β-cell autophagy deficiency (32,127,128,133,145,233). With this improved model, we established that basal β-cell autophagy is required for maintaining normal glucose tolerance. Using in vitro and in vivo models of short-term Atg5 deletion, we then addressed whether autophagy promotes or prevents β-cell survival under hypoxia and after islet transplantation. Prior studies suggest both possibilities but have not tested this directly (46,60). Our results showed that loss of autophagy exacerbates hypoxia-induced β-cell death and islet graft failure, demonstrating that autophagy is β-cell protective under these conditions. Furthermore, we found that prolonged hypoxia impairs autophagic flux, which Chapter 4 revealed is linked to compromised lysosomal homeostasis and a significant loss of the transcriptional master regulator TFEB. We further established that β-cell TFEB is reduced under ER stress, as well as lipotoxic conditions that are known to impair autophagic clearance (146,195,248). Importantly, we demonstrated that activating Tfeb can restore lysosomal activity and improve viability in mouse β-cells. In Chapter 5 we again used the Ins1Cre mouse line, this time to generate a mouse model with Tfeb deletion that is more β-cell-specific than what was used in a recent study (40). In contrast with this prior paper, we did not 131 see evidence for β-cell mitochondrial dysfunction and diabetes when we exposed our conditional Tfeb KO mice to HFD-induced metabolic stress. Ex vivo examination of islets from our mice and in vitro knockout experiments suggest this might be attributed to differences in compensatory Tfe3 expression, which we speculate may be linked to differences in the genetic background of our mice, and subsequent differences in the levels of β-cell stress. Further work is needed to test these predictions. Overall, this thesis enhances our understanding of autophagy's protective roles in stressed β-cells, identifies TFEB dysregulation as a possible cause of autophagy loss and β-cell failure in diabetes, and underscores the potential for improving β-cell stress adaptation by enhancing TFEB function. 6.2 Limitations and caveats When characterizing our β-cell-specific Atg5 and Tfeb knockout mice, we tracked the metabolic phenotypes of both male and female littermates. However, it is a limitation of this thesis that sex differences were not comprehensively examined in all studies. For example, we only investigated the role of Tfeb under MLD-STZ-induced stress in male mice, and our islet transplantation experiments only compared grafts of female islets transplanted into diabetic male recipients. We did this primarily because male mice tend to develop more robust and sustained STZ-induced hyperglycemia, and female littermates then were available to act as matched syngeneic islet donors. However, the performance of transplanted human donor islets (296) and the in vivo maturation of human embryonic stem cell-derived islet grafts (297) appear to be better in female 132 recipients, potentially due to differences in circulating sex hormones. Also, intrinsic sex differences in islet ER stress resilience (19) could conceivably affect MLD-STZ-induced damage, as well as the viability of transplanted islets. If we explore graft autophagy further, it will therefore be relevant to examine the role of recipient sex and to assess how islet donor sex affects graft survival and function in the same recipient environment. Moreover, the in vitro experiments in Chapters 3 and 4 were primarily done using islets from male mice. Given that our male and female TfebβKO mice exhibited differences in glucose tolerance and islet gene expression, it will be important to also investigate possible sex-differences in all in vitro aspects of Tfeb-regulated autophagy and stress adaptation. We used the most β-cell specific Cre-expressing mouse model available to drive gene deletion, but it is a limitation of our experimental design that we only used flox/flox littermates as controls for our Atg5βKO and TfebβKO mice. To control for the possible effects of losing insulin expression from one Ins1 allele and for β-cell expression of Cre, heterozygous breeding would ideally have been used to also generate Ins1Cre ‘wildtype’ littermates. Recent work has shown that NOD mice backcrossed with Ins1Cre mice are partially protected from developing T1D (298), so we recognize the lack of Cre expression in our controls as a potential caveat. Nonetheless, we are not aware of reports of altered glucose metabolism or β-cell function in chow- or HFD-fed Ins1Cre mice, and overall, we do not believe Ins1Cre has significantly affected our results. In our MLD-STZ experiment we included a group of age-matched Ins1Cre mice, and the outcome did not indicate that the worsened glucose tolerance in TfebβKO animals was 133 caused by the transgene. Moreover, Tfebflox/flox;Ins1Cre/wt and Tfebflox/flox mice only showed very minor differences in glucose tolerance irrespective of the experimental diet. While we cannot rule out that the presence of Ins1Cre contributed to the slightly improved glucose tolerance in older TfebβKO mice, our results do indicate that Ins1Cre does not have notable metabolic effects in our colonies. This also means that it is unlikely to have contributed significantly to the striking phenotype of our Atg5βKO mice. Our combined in vitro data provide clear evidence that 16-24 hours of hypoxic culture induces maladaptive ER stress, impairs the adaptive UPR, lowers β-cell Tfeb mRNA and protein levels, and impairs both lysosomal function and autophagic clearance. However, because our studies were limited to relatively late stages of hypoxic stress, they do not address the progression of these cellular events - including when and how Tfeb-regulated lysosomal autophagy and the UPR become dysfunctional. It should also be noted that hypoxia and circadian rhythms regulate each other (299), which could possibly contribute to the impairments at later time points. Moreover, we only examined the effects of hypoxic culture at 1% O2. While this level of O2 is pertinent for our understanding of the hypoxic stress faced by transplanted islets, it would also be relevant to determine the impact of more moderate hypoxia that islets may encounter during periods of high metabolic demand. We further acknowledge that the atmospheric oxygen levels of ~20% in our control cultures may be higher than what islet cells normally experience in the pancreas, which could possibly be as low as ~5% (210). Despite this limitation, our control conditions allow for valuable comparison with existing literature and our 134 in vitro experiments. Our syngeneic mouse islet transplantation experiments revealed that Atg5 helps sustain graft function in the first weeks following surgery. Unfortunately, our ability to examine the islet grafts was limited. We attempted to extract the surviving control islet grafts to compare the expression of Tfeb, as well as other lysosome and autophagy genes by qPCR. However, we were unable to locate several of the grafts under the kidney capsule and get clean enough samples for meaningful data. It therefore remains to be determined if transplanted β-cells also develop defects in Tfeb and autophagic flux, as our in vitro hypoxia and ER stress data strongly suggest may be the case. When assessing the impact of hypoxia, we found that CTSB activity is impaired. Our TfebβKO and torin-1 rescue experiments showed that Tfeb promotes CTSB expression and activity, and CTSB has previously been linked to lysosomal function and autophagic clearance in β-cells (195). This indicates that it is the lysosomal impairment that disrupts β-cell autophagy under hypoxia, but we acknowledge that this has not been directly demonstrated. Additionally, it is a limitation that autophagic flux was not assessed alongside torin-1-induced normalization of gene expression and CTSB activity. Further work, including also assessments of lysosomal pH, calcium signaling and ultrastructure, could help establish the extent and primary features of lysosomal dysfunction. Finally, our TfebβKO mouse studies leave some questions regarding the phenotypes of our mice. Isolated TfebβKO islets clearly had impaired expression of Tfeb target genes and reduced 135 lysosomal CSTB activity. However, because overall glucose tolerance and ex vivo islet function were normal, it should be investigated if autophagic flux is affected or how the cells otherwise compensate for the observed lysosomal impairments. We also lack clarity on the transient improvement in glucose tolerance observed in female mice after ~4 weeks on HFD. In this regard, it is a limitation that we only compared ex vivo islets after 16-18 weeks of HFD-feeding, and that we have not measured β-cell mass, islet insulin expression, insulin content and in vivo insulin secretion. We assayed the insulin content of ex vivo islets from HFD-fed TfebβWT and TfebβKO mice but found that the insulin extraction in some cases was incomplete and the protocol needed to be adjusted. The data were therefore inconclusive and not included in the thesis. Given the role of autophagy in insulin granule degradation and the recent finding that TFEB and TFE3 regulate insulin expression (260), further investigation into insulin homeostasis in our Tfeb models is warranted. 6.3 Future directions In addition to the question of potential sex-differences in Tfeb-dependent gene expression and glucose tolerance, this thesis has highlighted important areas for future research. Overall, our findings point to TFEB dysregulation as a driver of autophagy failure in stressed β-cells, possibly also in the progression of T1D and T2D where impaired β-cell autophagy is increasingly recognized as a likely pathogenic factor. However, the different phenotypes we observed in our MLD-STZ and HFD-stressed TfebβKO mice suggest that TFEB’s importance depends on the 136 type or severity of stress. As discussed in Chapter 5, we hypothesize that TFE3 can compensate for loss of TFEB function under moderate β-cell stress, but not when the insult is too severe or chronic - either because TFE3 only regulates a subset of TFEB gene targets or because TFE3 itself may become impaired, as indicated by loss of Tfe3 mRNA in hypoxic mouse islets. To address these questions, it will be necessary to carefully investigate both TFEB and TFE3 under conditions where β-cell autophagy is positively or negatively affected by stress. It will also be a priority to examine autophagic function and survival in mouse β-cells with single and double Tfeb/Tfe3 deficiency. In vitro, this can be done using Tfebflox/flox cells and adenovirus constructs described in Chapter 5. In vivo we would use AAV8-Ins1-mediated expression of Cre with and without simultaneous shRNA-mediated knockdown of Tfe3. We have bred Tfebflox/flox mice that also express the RFP-EGFP-LC3 autophagy reporter, which can be used to evaluate the effects on autophagic flux. In vitro, we can use multi-well high content confocal imaging to track β-cell autophagy and death, including the time-dependent effects of exposure to different levels of pro-inflammatory cytokines, ER stress, glucolipotoxicity and hypoxia. To test if Tfe3-dependent compensation helps sustain β-cell function and glucose homeostasis in TfebβKO mice, we would compare HFD-induced glucose intolerance in animals with AAV-induced single and double β-cell Tfeb/Tfe3 deficiency. To also get an idea of redundant versus selective transcriptional regulation by Tfeb and Tfe3, we intend to use RNA-Seq analyses to generate a comprehensive picture of the mRNA changes that follow their individual and combined loss. This will also allow more comprehensive insights into the overall β-cell state than the candidate gene approach used 137 in this thesis. Because bulk islet gene expression includes changes in non-β-cells, RNA-Seq will ideally be done using FACS-purified β-cells, or if necessary, single-cell RNA-Seq. We successfully used the mTORC1 inhibitor torin-1 and Tfeb overexpression to improve lysosomal function and β-cell survival under hypoxia and/or ER stress. As discussed, these findings support the idea that promoting TFEB activity may be a strategy to protect β-cells during diabetes development and after transplantation of donor- or stem cell-derived islets. As a first step toward this, we would test various compounds that have been reported to activate TFEB for their ability to induce TFEB:EGFP translocation in cultured β-cells. We would then test if those that promote TFEB translocation can stimulate autophagy and promote survival under stress. Also here, we would use a high-content imaging approach to determine the concentration- and time-dependent effects of the different compounds. Doing this side-by-side in TfebβWT and TfebβKO cells can tell us if the effects are Tfeb-dependent or not. When considering possible diabetes therapies, AAV-mediated gene delivery has potential as a means to achieve β-cell-targeted effects. To date, AAVs have mostly been used to express Cre for β-cell-specific gene deletion in rodent models, a powerful approach we also used in this thesis. Of note, however, it has also been demonstrated that β-cell-targeted expression of other transgenes by AAVs can have disease-modulating effects in mouse models of diabetes (201). Moving forward, it is an important goal for us to determine if AAV-Ins1-Tfeb delivery can improve β-cell function and prevent diabetes in mouse models of autoimmune T1D and obesity-related T2D. Even if β-cells do not exhibit a significant reduction in TFEB levels, as we 138 observed under in vitro stress, activating and overexpressing TFEB may stimulate autophagic clearance, and possibly also PGC-1α-regulated mitochondrial metabolism, to help sustain β-cell adaptation and function. To determine if TFEB and TFE3 are altered in the progression of diabetes, it will be important to also characterize their endogenous levels, phosphorylation and intracellular localization in control groups. Finally, it will be essential to investigate the roles and protective potential of TFEB and TFE3 in human islets and β-cells. We found that hypoxia reduces TFEB immunoreactivity in human β-cells. However, beyond this important observation, the questions addressed in this thesis have not been explored in human islets. To investigate this, we could use similar approaches and examine human β-cell function and survival under various stress conditions following TFEB/TFE3 knockdown or overexpression. Additionally, it would be important to examine whether TFEB and TFE3 levels or functions are impaired in islets from human donors with diabetes, compared to islets from healthy donors. For this purpose, new online resources and tools from the Alberta Diabetes Institute IsletCore could be valuable, as they provide access to linked multi-omics and functional phenotyping data from islets isolated from hundreds of human donors with and without diabetes (300). In the context of islet transplantation, it will be interesting to assess if human donor islets and stem cell-derived islets can be treated with TFEB activators or transduced with AAV-TFEB vectors to improve graft survival and function. Stem cell-derived islets also provide opportunities of using gene-editing to target TFEB and other autophagy pathways for improved transplantation outcomes and T1D treatment (301). Overall, 139 the mouse work presented in this thesis has reinforced the importance of β-cell autophagy, identified Tfeb dysregulation as a potential cause of autophagy failure in stressed β-cells, and highlighted Tfeb as a promising target for β-cell protection. Ultimately, the therapeutic relevance of TFEB in islet transplantation and diabetes treatment will depend on its function, and possible dysfunction, in human β-cells. 140 References 1. Sun, H., Saeedi, P., Karuranga, S., Pinkepank, M., Ogurtsova, K., Duncan, B. B., Stein, C., Basit, A., Chan, J. C. N., Mbanya, J. C., Pavkov, M. E., Ramachandaran, A., Wild, S. H., James, S., Herman, W. H. H., Zhang, P., Bommer, C., Kuo, S. H., Boyko, E. J. J., and Magliano, D. J. (2022) IDF Diabetes Atlas: Global, regional and country-level diabetes prevalence estimates for 2021 and projections for 2045. Diabetes Res Clin Pr 183, 109119 2. Xie, J., Wang, M., Long, Z., Ning, H., Li, J., Cao, Y., Liao, Y., Liu, G., Wang, F., and Pan, A. (2022) Global burden of type 2 diabetes in adolescents and young adults, 1990-2019: systematic analysis of the Global Burden of Disease Study 2019. BMJ 379, e072385 3. Tokarz, V. L., MacDonald, P. E., and Klip, A. (2018) The cell biology of systemic insulin function. The Journal of cell biology 217, 2273-2289 4. Forbes, J. M., and Cooper, M. E. (2013) Mechanisms of diabetic complications. Physiol Rev 93, 137-188 5. Chen, C., Cohrs, C. M., Stertmann, J., Bozsak, R., and Speier, S. (2017) Human beta cell mass and function in diabetes: Recent advances in knowledge and technologies to understand disease pathogenesis. Molecular metabolism 6, 943-957 6. Back, S. H., and Kaufman, R. J. (2012) Endoplasmic reticulum stress and type 2 diabetes. Annual review of biochemistry 81, 767-793 7. Bratanova-Tochkova, T. K., Cheng, H., Daniel, S., Gunawardana, S., Liu, Y. J., Mulvaney-Musa, J., Schermerhorn, T., Straub, S. G., Yajima, H., and Sharp, G. W. (2002) Triggering and augmentation mechanisms, granule pools, and biphasic insulin secretion. Diabetes 51 Suppl 1, S83-90 8. Rutter, G. A., Pullen, T. J., Hodson, D. J., and Martinez-Sanchez, A. (2015) Pancreatic beta-cell identity, glucose sensing and the control of insulin secretion. Biochem J 466, 203-218 9. Kalwat, M. A., and Cobb, M. H. (2017) Mechanisms of the amplifying pathway of insulin secretion in the beta cell. Pharmacol Ther 179, 17-30 10. Merrins, M. J., Corkey, B. E., Kibbey, R. G., and Prentki, M. (2022) Metabolic cycles and signals for insulin secretion. Cell metabolism 34, 947-968 11. Weir, G. C., Laybutt, D. R., Kaneto, H., Bonner-Weir, S., and Sharma, A. (2001) beta-cell adaptation and decompensation during the progression of diabetes. Diabetes 50, S154-S159 12. Eguchi, N., Vaziri, N. D., Dafoe, D. C., and Ichii, H. (2021) The Role of Oxidative Stress 141 in Pancreatic beta Cell Dysfunction in Diabetes. Int J Mol Sci 22, 1509 13. Cnop, M., Welsh, N., Jonas, J. C., Jorns, A., Lenzen, S., and Eizirik, D. L. (2005) Mechanisms of pancreatic beta-cell death in type 1 and type 2 diabetes: many differences, few similarities. Diabetes 54 Suppl 2, S97-107 14. Shapiro, A. M., Pokrywczynska, M., and Ricordi, C. (2017) Clinical pancreatic islet transplantation. Nature reviews. Endocrinology 13, 268-277 15. Kale, A., and Rogers, N. M. (2023) No Time to Die-How Islets Meet Their Demise in Transplantation. Cells 12, 796 16. Negi, S., Park, S. H., Jetha, A., Aikin, R., Tremblay, M., and Paraskevas, S. (2012) Evidence of endoplasmic reticulum stress mediating cell death in transplanted human islets. Cell Transplant 21, 889-900 17. Ramkumar, K. M., Sekar, T. V., Bhakkiyalakshmi, E., Foygel, K., Rajaguru, P., Berger, F., and Paulmurugan, R. (2013) The impact of oxidative stress on islet transplantation and monitoring the graft survival by non-invasive imaging. Curr Med Chem 20, 1127-1146 18. Gannon, M., Kulkarni, R. N., Tse, H. M., and Mauvais-Jarvis, F. (2018) Sex differences underlying pancreatic islet biology and its dysfunction. Molecular metabolism 15, 82-91 19. Brownrigg, G. P., Xia, Y. H., Chu, C. M. J., Wang, S., Chao, C., Zhang, J. A., Skovso, S., Panzhinskiy, E., Hu, X., Johnson, J. D., and Rideout, E. J. (2023) Sex differences in islet stress responses support female beta cell resilience. Mol Metab 69, 101678 20. Jo, S., Beetch, M., Gustafson, E., Wong, A., Oribamise, E., Chung, G., Vadrevu, S., Satin, L. S., Bernal-Mizrachi, E., and Alejandro, E. U. (2023) Sex Differences in Pancreatic beta-Cell Physiology and Glucose Homeostasis in C57BL/6J Mice. J Endocr Soc 7, bvad099 21. Mauvais-Jarvis, F. (2016) Role of Sex Steroids in beta Cell Function, Growth, and Survival. Trends in endocrinology and metabolism: TEM 27, 844-855 22. Drigo, R. A. e. a. (2019) Aging of human endocrine pancreatic cell types is heterogeneous and sexspecific. BioRxiv 23. Li, T., Jiao, W., Li, W., and Li, H. (2016) Sex effect on insulin secretion and mitochondrial function in pancreatic beta cells of elderly Wistar rats. Endocr Res 41, 167-179 24. Jacobo-Piqueras, N., Theiner, T., Geisler, S. M., and Tuluc, P. (2024) Molecular mechanism responsible for sex differences in electrical activity of mouse pancreatic beta cells. JCI Insight 9, e171609 142 25. Lytrivi, M., Castell, A. L., Poitout, V., and Cnop, M. (2020) Recent Insights Into Mechanisms of beta-Cell Lipo- and Glucolipotoxicity in Type 2 Diabetes. J Mol Biol 432, 1514-1534 26. Katsarou, A., Gudbjornsdottir, S., Rawshani, A., Dabelea, D., Bonifacio, E., Anderson, B. J., Jacobsen, L. M., Schatz, D. A., and Lernmark, A. (2017) Type 1 diabetes mellitus. Nature reviews. Disease primers 3, 17016 27. Dludla, P. V., Mabhida, S. E., Ziqubu, K., Nkambule, B. B., Mazibuko-Mbeje, S. E., Hanser, S., Basson, A. K., Pheiffer, C., and Kengne, A. P. (2023) Pancreatic beta-cell dysfunction in type 2 diabetes: Implications of inflammation and oxidative stress. World J Diabetes 14, 130-146 28. Jo, S., and Fang, S. (2021) Therapeutic Strategies for Diabetes: Immune Modulation in Pancreatic beta Cells. Front Endocrinol (Lausanne) 12, 716692 29. Kanak, M. A., Takita, M., Kunnathodi, F., Lawrence, M. C., Levy, M. F., and Naziruddin, B. (2014) Inflammatory response in islet transplantation. Int J Endocrinol 2014, 451035 30. Montane, J., Klimek-Abercrombie, A., Potter, K. J., Westwell-Roper, C., and Bruce Verchere, C. (2012) Metabolic stress, IAPP and islet amyloid. Diabetes, obesity & metabolism 14 Suppl 3, 68-77 31. Denroche, H. C., and Verchere, C. B. (2018) IAPP and type 1 diabetes: implications for immunity, metabolism and islet transplants. J Mol Endocrinol 60, R57-R75 32. Rivera, J. F., Costes, S., Gurlo, T., Glabe, C. G., and Butler, P. C. (2014) Autophagy defends pancreatic beta cells from human islet amyloid polypeptide-induced toxicity. The Journal of clinical investigation 124, 3489-3500 33. Hernandez, M. G., Aguilar, A. G., Burillo, J., Oca, R. G., Manca, M. A., Novials, A., Alcarraz-Vizan, G., Guillen, C., and Benito, M. (2018) Pancreatic beta cells overexpressing hIAPP impaired mitophagy and unbalanced mitochondrial dynamics. Cell Death Dis 9, 481 34. Montane, J., Cadavez, L., and Novials, A. (2014) Stress and the inflammatory process: a major cause of pancreatic cell death in type 2 diabetes. Diabetes Metab Syndr Obes 7, 25-34 35. Sato, Y., Endo, H., Okuyama, H., Takeda, T., Iwahashi, H., Imagawa, A., Yamagata, K., Shimomura, I., and Inoue, M. (2011) Cellular hypoxia of pancreatic beta-cells due to high levels of oxygen consumption for insulin secretion in vitro. The Journal of biological chemistry 286, 12524-12532 36. Gerber, P. A., and Rutter, G. A. (2017) The Role of Oxidative Stress and Hypoxia in 143 Pancreatic Beta-Cell Dysfunction in Diabetes Mellitus. Antioxid Redox Signal 26, 501-518 37. Hetz, C., Zhang, K., and Kaufman, R. J. (2020) Mechanisms, regulation and functions of the unfolded protein response. Nature reviews. Molecular cell biology 21, 421-438 38. Pearson, G. L., Gingerich, M. A., Walker, E. M., Biden, T. J., and Soleimanpour, S. A. (2021) A Selective Look at Autophagy in Pancreatic beta-Cells. Diabetes 70, 1229-1241 39. Stancill, J. S., Broniowska, K. A., Oleson, B. J., Naatz, A., and Corbett, J. A. (2019) Pancreatic beta-cells detoxify H(2)O(2) through the peroxiredoxin/thioredoxin antioxidant system. The Journal of biological chemistry 294, 4843-4853 40. Park, K., Lim, H., Kim, J., Hwang, Y., Lee, Y. S., Bae, S. H., Kim, H., Kim, H., Kang, S. W., Kim, J. Y., and Lee, M. S. (2022) Lysosomal Ca(2+)-mediated TFEB activation modulates mitophagy and functional adaptation of pancreatic beta-cells to metabolic stress. Nature communications 13, 1300 41. Komatsu, H., Kandeel, F., and Mullen, Y. (2018) Impact of Oxygen on Pancreatic Islet Survival. Pancreas 47, 533-543 42. Olsson, R., Olerud, J., Pettersson, U., and Carlsson, P. O. (2011) Increased numbers of low-oxygenated pancreatic islets after intraportal islet transplantation. Diabetes 60, 2350-2353 43. Sato, Y., Inoue, M., Yoshizawa, T., and Yamagata, K. (2014) Moderate hypoxia induces beta-cell dysfunction with HIF-1-independent gene expression changes. PloS one 9, e114868 44. Tsuyama, T., Sato, Y., Yoshizawa, T., Matsuoka, T., and Yamagata, K. (2023) Hypoxia causes pancreatic beta-cell dysfunction and impairs insulin secretion by activating the transcriptional repressor BHLHE40. EMBO Rep 24, e56227 45. Bensellam, M., Maxwell, E. L., Chan, J. Y., Luzuriaga, J., West, P. K., Jonas, J. C., Gunton, J. E., and Laybutt, D. R. (2016) Hypoxia reduces ER-to-Golgi protein trafficking and increases cell death by inhibiting the adaptive unfolded protein response in mouse beta cells. Diabetologia 59, 1492-1502 46. Zheng, X., Zheng, X., Wang, X., Ma, Z., Gupta Sunkari, V., Botusan, I., Takeda, T., Bjorklund, A., Inoue, M., Catrina, S. B., Brismar, K., Poellinger, L., and Pereira, T. S. (2012) Acute hypoxia induces apoptosis of pancreatic beta-cell by activation of the unfolded protein response and upregulation of CHOP. Cell death & disease 3, e322 47. Iwawaki, T., Akai, R., Kohno, K., and Miura, M. (2004) A transgenic mouse model for monitoring endoplasmic reticulum stress. Nature medicine 10, 98-102 48. Walter, P., and Ron, D. (2011) The unfolded protein response: from stress pathway to 144 homeostatic regulation. Science 334, 1081-1086 49. Pobre, K. F. R., Poet, G. J., and Hendershot, L. M. (2019) The endoplasmic reticulum (ER) chaperone BiP is a master regulator of ER functions: Getting by with a little help from ERdj friends. The Journal of biological chemistry 294, 2098-2108 50. Sharma, R. B., Landa-Galvan, H. V., and Alonso, L. C. (2021) Living Dangerously: Protective and Harmful ER Stress Responses in Pancreatic beta-Cells. Diabetes 70, 2431-2443 51. Gupta, S., McGrath, B., and Cavener, D. R. (2010) PERK (EIF2AK3) regulates proinsulin trafficking and quality control in the secretory pathway. Diabetes 59, 1937-1947 52. Harding, H. P., Zyryanova, A. F., and Ron, D. (2012) Uncoupling proteostasis and development in vitro with a small molecule inhibitor of the pancreatic endoplasmic reticulum kinase, PERK. The Journal of biological chemistry 287, 44338-44344 53. Scheuner, D., Vander Mierde, D., Song, B., Flamez, D., Creemers, J. W., Tsukamoto, K., Ribick, M., Schuit, F. C., and Kaufman, R. J. (2005) Control of mRNA translation preserves endoplasmic reticulum function in beta cells and maintains glucose homeostasis. Nat Med 11, 757-764 54. Chan, J. Y., Luzuriaga, J., Maxwell, E. L., West, P. K., Bensellam, M., and Laybutt, D. R. (2015) The balance between adaptive and apoptotic unfolded protein responses regulates beta-cell death under ER stress conditions through XBP1, CHOP and JNK. Molecular and cellular endocrinology 413, 189-201 55. Allagnat, F., Christulia, F., Ortis, F., Pirot, P., Lortz, S., Lenzen, S., Eizirik, D. L., and Cardozo, A. K. (2010) Sustained production of spliced X-box binding protein 1 (XBP1) induces pancreatic beta cell dysfunction and apoptosis. Diabetologia 53, 1120-1130 56. Usui, M., Yamaguchi, S., Tanji, Y., Tominaga, R., Ishigaki, Y., Fukumoto, M., Katagiri, H., Mori, K., Oka, Y., and Ishihara, H. (2012) Atf6alpha-null mice are glucose intolerant due to pancreatic beta-cell failure on a high-fat diet but partially resistant to diet-induced insulin resistance. Metabolism 61, 1118-1128 57. Harding, H. P., Zeng, H., Zhang, Y., Jungries, R., Chung, P., Plesken, H., Sabatini, D. D., and Ron, D. (2001) Diabetes mellitus and exocrine pancreatic dysfunction in perk-/- mice reveals a role for translational control in secretory cell survival. Molecular cell 7, 1153-1163 58. Gao, Y., Sartori, D. J., Li, C., Yu, Q. C., Kushner, J. A., Simon, M. C., and Diehl, J. A. (2012) PERK is required in the adult pancreas and is essential for maintenance of glucose homeostasis. Molecular and cellular biology 32, 5129-5139 145 59. Xu, G., Chen, J., Jing, G., Grayson, T. B., and Shalev, A. (2016) miR-204 Targets PERK and Regulates UPR Signaling and beta-Cell Apoptosis. Mol Endocrinol 30, 917-924 60. Song, S., Tan, J., Miao, Y., Sun, Z., and Zhang, Q. (2018) Intermittent-Hypoxia-Induced Autophagy Activation Through the ER-Stress-Related PERK/eIF2alpha/ATF4 Pathway is a Protective Response to Pancreatic beta-Cell Apoptosis. Cell Physiol Biochem 51, 2955-2971 61. Jiang, D., and Wan, F. (2018) Exendin-4 protects INS-1 cells against palmitate-induced apoptosis through the IRE1alpha-Xbp1 signaling pathway. Exp Ther Med 16, 1029-1035 62. Yong, J., Parekh, V. S., Reilly, S. M., Nayak, J., Chen, Z., Lebeaupin, C., Jang, I., Zhang, J., Prakash, T. P., Sun, H., Murray, S., Guo, S., Ayala, J. E., Satin, L. S., Saltiel, A. R., and Kaufman, R. J. (2021) Chop/Ddit3 depletion in beta cells alleviates ER stress and corrects hepatic steatosis in mice. Sci Transl Med 13, eaba9796 63. Song, B., Scheuner, D., Ron, D., Pennathur, S., and Kaufman, R. J. (2008) Chop deletion reduces oxidative stress, improves beta cell function, and promotes cell survival in multiple mouse models of diabetes. The Journal of clinical investigation 118, 3378-3389 64. Klionsky, D. J. e. a. (2021) Guidelines for the use and interpretation of assays for monitoring autophagy (4th edition)(1). Autophagy 17, 1-382 65. Kocaturk, N. M., and Gozuacik, D. (2018) Crosstalk Between Mammalian Autophagy and the Ubiquitin-Proteasome System. Frontiers in cell and developmental biology 6, 128 66. Kroemer, G., Marino, G., and Levine, B. (2010) Autophagy and the integrated stress response. Mol Cell 40, 280-293 67. Klionsky, D. J., Petroni, G., Amaravadi, R. K., Baehrecke, E. H., Ballabio, A., Boya, P., Bravo-San Pedro, J. M., Cadwell, K., Cecconi, F., Choi, A. M. K., Choi, M. E., Chu, C. T., Codogno, P., Colombo, M. I., Cuervo, A. M., Deretic, V., Dikic, I., Elazar, Z., Eskelinen, E. L., Fimia, G. M., Gewirtz, D. A., Green, D. R., Hansen, M., Jaattela, M., Johansen, T., Juhasz, G., Karantza, V., Kraft, C., Kroemer, G., Ktistakis, N. T., Kumar, S., Lopez-Otin, C., Macleod, K. F., Madeo, F., Martinez, J., Melendez, A., Mizushima, N., Munz, C., Penninger, J. M., Perera, R. M., Piacentini, M., Reggiori, F., Rubinsztein, D. C., Ryan, K. M., Sadoshima, J., Santambrogio, L., Scorrano, L., Simon, H. U., Simon, A. K., Simonsen, A., Stolz, A., Tavernarakis, N., Tooze, S. A., Yoshimori, T., Yuan, J., Yue, Z., Zhong, Q., Galluzzi, L., and Pietrocola, F. (2021) Autophagy in major human diseases. The EMBO journal 40, e108863 68. (2016) The Nobel Prize in Physiology or Medicine 2016., NobelPrize.org 69. Uttenweiler, A., and Mayer, A. (2008) Microautophagy in the yeast Saccharomyces cerevisiae. Methods Mol Biol 445, 245-259 146 70. Schafer, J. A., Schessner, J. P., Bircham, P. W., Tsuji, T., Funaya, C., Pajonk, O., Schaeff, K., Ruffini, G., Papagiannidis, D., Knop, M., Fujimoto, T., and Schuck, S. (2020) ESCRT machinery mediates selective microautophagy of endoplasmic reticulum in yeast. The EMBO journal 39, e102586 71. Kaushik, S., and Cuervo, A. M. (2012) Chaperone-mediated autophagy: a unique way to enter the lysosome world. Trends Cell Biol 22, 407-417 72. Salvador, N., Aguado, C., Horst, M., and Knecht, E. (2000) Import of a cytosolic protein into lysosomes by chaperone-mediated autophagy depends on its folding state. The Journal of biological chemistry 275, 27447-27456 73. Galluzzi, L., Baehrecke, E. H., Ballabio, A., Boya, P., Bravo-San Pedro, J. M., Cecconi, F., Choi, A. M., Chu, C. T., Codogno, P., Colombo, M. I., Cuervo, A. M., Debnath, J., Deretic, V., Dikic, I., Eskelinen, E. L., Fimia, G. M., Fulda, S., Gewirtz, D. A., Green, D. R., Hansen, M., Harper, J. W., Jaattela, M., Johansen, T., Juhasz, G., Kimmelman, A. C., Kraft, C., Ktistakis, N. T., Kumar, S., Levine, B., Lopez-Otin, C., Madeo, F., Martens, S., Martinez, J., Melendez, A., Mizushima, N., Munz, C., Murphy, L. O., Penninger, J. M., Piacentini, M., Reggiori, F., Rubinsztein, D. C., Ryan, K. M., Santambrogio, L., Scorrano, L., Simon, A. K., Simon, H. U., Simonsen, A., Tavernarakis, N., Tooze, S. A., Yoshimori, T., Yuan, J., Yue, Z., Zhong, Q., and Kroemer, G. (2017) Molecular definitions of autophagy and related processes. The EMBO journal 36, 1811-1836 74. Johansen, T., and Lamark, T. (2020) Selective Autophagy: ATG8 Family Proteins, LIR Motifs and Cargo Receptors. J Mol Biol 432, 80-103 75. Onishi, M., Yamano, K., Sato, M., Matsuda, N., and Okamoto, K. (2021) Molecular mechanisms and physiological functions of mitophagy. The EMBO journal 40, e104705 76. Narendra, D., Tanaka, A., Suen, D. F., and Youle, R. J. (2008) Parkin is recruited selectively to impaired mitochondria and promotes their autophagy. The Journal of cell biology 183, 795-803 77. Lazarou, M., Sliter, D. A., Kane, L. A., Sarraf, S. A., Wang, C., Burman, J. L., Sideris, D. P., Fogel, A. I., and Youle, R. J. (2015) The ubiquitin kinase PINK1 recruits autophagy receptors to induce mitophagy. Nature 524, 309-314 78. Soleimanpour, S. A., Gupta, A., Bakay, M., Ferrari, A. M., Groff, D. N., Fadista, J., Spruce, L. A., Kushner, J. A., Groop, L., Seeholzer, S. H., Kaufman, B. A., Hakonarson, H., and Stoffers, D. A. (2014) The diabetes susceptibility gene Clec16a regulates mitophagy. Cell 157, 1577-1590 79. Nakatogawa, H. (2020) Mechanisms governing autophagosome biogenesis. Nature 147 reviews. Molecular cell biology 21, 439-458 80. Axe, E. L., Walker, S. A., Manifava, M., Chandra, P., Roderick, H. L., Habermann, A., Griffiths, G., and Ktistakis, N. T. (2008) Autophagosome formation from membrane compartments enriched in phosphatidylinositol 3-phosphate and dynamically connected to the endoplasmic reticulum. The Journal of cell biology 182, 685-701 81. Katsuragi, Y., Ichimura, Y., and Komatsu, M. (2015) p62/SQSTM1 functions as a signaling hub and an autophagy adaptor. FEBS J 282, 4672-4678 82. Zhen, Y., Spangenberg, H., Munson, M. J., Brech, A., Schink, K. O., Tan, K. W., Sorensen, V., Wenzel, E. M., Radulovic, M., Engedal, N., Simonsen, A., Raiborg, C., and Stenmark, H. (2020) ESCRT-mediated phagophore sealing during mitophagy. Autophagy 16, 826-841 83. Mizushima, N. (2020) The ATG conjugation systems in autophagy. Curr Opin Cell Biol 63, 1-10 84. Turco, E., Fracchiolla, D., and Martens, S. (2020) Recruitment and Activation of the ULK1/Atg1 Kinase Complex in Selective Autophagy. J Mol Biol 432, 123-134 85. Kijanska, M., Dohnal, I., Reiter, W., Kaspar, S., Stoffel, I., Ammerer, G., Kraft, C., and Peter, M. (2010) Activation of Atg1 kinase in autophagy by regulated phosphorylation. Autophagy 6, 1168-1178 86. Karanasios, E., Walker, S. A., Okkenhaug, H., Manifava, M., Hummel, E., Zimmermann, H., Ahmed, Q., Domart, M. C., Collinson, L., and Ktistakis, N. T. (2016) Autophagy initiation by ULK complex assembly on ER tubulovesicular regions marked by ATG9 vesicles. Nature communications 7, 12420 87. Kotani, T., Kirisako, H., Koizumi, M., Ohsumi, Y., and Nakatogawa, H. (2018) The Atg2-Atg18 complex tethers pre-autophagosomal membranes to the endoplasmic reticulum for autophagosome formation. Proc Natl Acad Sci U S A 115, 10363-10368 88. Menon, M. B., and Dhamija, S. (2018) Beclin 1 Phosphorylation - at the Center of Autophagy Regulation. Frontiers in cell and developmental biology 6, 137 89. Walczak, M., and Martens, S. (2013) Dissecting the role of the Atg12-Atg5-Atg16 complex during autophagosome formation. Autophagy 9, 424-425 90. Lorincz, P., and Juhasz, G. (2020) Autophagosome-Lysosome Fusion. J Mol Biol 432, 2462-2482 91. Szatmari, Z., and Sass, M. (2014) The autophagic roles of Rab small GTPases and their upstream regulators: a review. Autophagy 10, 1154-1166 148 92. Wickner, W. (2010) Membrane fusion: five lipids, four SNAREs, three chaperones, two nucleotides, and a Rab, all dancing in a ring on yeast vacuoles. Annual review of cell and developmental biology 26, 115-136 93. Stoka, V., Turk, V., and Turk, B. (2016) Lysosomal cathepsins and their regulation in aging and neurodegeneration. Ageing Res Rev 32, 22-37 94. Schwake, M., Schroder, B., and Saftig, P. (2013) Lysosomal membrane proteins and their central role in physiology. Traffic 14, 739-748 95. Huynh, K. K., Eskelinen, E. L., Scott, C. C., Malevanets, A., Saftig, P., and Grinstein, S. (2007) LAMP proteins are required for fusion of lysosomes with phagosomes. The EMBO journal 26, 313-324 96. Andrejewski, N., Punnonen, E. L., Guhde, G., Tanaka, Y., Lullmann-Rauch, R., Hartmann, D., von Figura, K., and Saftig, P. (1999) Normal lysosomal morphology and function in LAMP-1-deficient mice. The Journal of biological chemistry 274, 12692-12701 97. Eskelinen, E. L., Schmidt, C. K., Neu, S., Willenborg, M., Fuertes, G., Salvador, N., Tanaka, Y., Lullmann-Rauch, R., Hartmann, D., Heeren, J., von Figura, K., Knecht, E., and Saftig, P. (2004) Disturbed cholesterol traffic but normal proteolytic function in LAMP-1/LAMP-2 double-deficient fibroblasts. Molecular biology of the cell 15, 3132-3145 98. Bandyopadhyay, U., Kaushik, S., Varticovski, L., and Cuervo, A. M. (2008) The chaperone-mediated autophagy receptor organizes in dynamic protein complexes at the lysosomal membrane. Molecular and cellular biology 28, 5747-5763 99. Xiong, J., and Zhu, M. X. (2016) Regulation of lysosomal ion homeostasis by channels and transporters. Sci China Life Sci 59, 777-791 100. Forgac, M. (2007) Vacuolar ATPases: rotary proton pumps in physiology and pathophysiology. Nature reviews. Molecular cell biology 8, 917-929 101. Mauvezin, C., Nagy, P., Juhasz, G., and Neufeld, T. P. (2015) Autophagosome-lysosome fusion is independent of V-ATPase-mediated acidification. Nature communications 6, 7007 102. Medina, D. L. (2021) Lysosomal calcium and autophagy. Int Rev Cell Mol Biol 362, 141-170 103. Scotto Rosato, A., Montefusco, S., Soldati, C., Di Paola, S., Capuozzo, A., Monfregola, J., Polishchuk, E., Amabile, A., Grimm, C., Lombardo, A., De Matteis, M. A., Ballabio, A., and Medina, D. L. (2019) TRPML1 links lysosomal calcium to autophagosome biogenesis through the activation of the CaMKKbeta/VPS34 pathway. Nature communications 10, 5630 149 104. Medina, D. L., Di Paola, S., Peluso, I., Armani, A., De Stefani, D., Venditti, R., Montefusco, S., Scotto-Rosato, A., Prezioso, C., Forrester, A., Settembre, C., Wang, W., Gao, Q., Xu, H., Sandri, M., Rizzuto, R., De Matteis, M. A., and Ballabio, A. (2015) Lysosomal calcium signalling regulates autophagy through calcineurin and TFEB. Nature cell biology 17, 288-299 105. Tian, X., Gala, U., Zhang, Y., Shang, W., Nagarkar Jaiswal, S., di Ronza, A., Jaiswal, M., Yamamoto, S., Sandoval, H., Duraine, L., Sardiello, M., Sillitoe, R. V., Venkatachalam, K., Fan, H., Bellen, H. J., and Tong, C. (2015) A voltage-gated calcium channel regulates lysosomal fusion with endosomes and autophagosomes and is required for neuronal homeostasis. PLoS Biol 13, e1002103 106. Zhu, F., Miao, Y., Cheng, M., Ye, X., Chen, A., Zheng, G., and Tian, X. (2022) The CACNA1A Mutant Disrupts Lysosome Calcium Homeostasis in Cerebellar Neurons and the Resulting Endo-Lysosomal Fusion Defect Can be Improved by Calcium Modulation. Neurochem Res 47, 249-263 107. Kim, S. W., Kim, M. K., Hong, S., Choi, A., Choi, J. H., Muallem, S., So, I., Yang, D., and Kim, H. J. (2022) The intracellular Ca(2+) channel TRPML3 is a PI3P effector that regulates autophagosome biogenesis. Proc Natl Acad Sci U S A 119, e2200085119 108. Rabanal-Ruiz, Y., Otten, E. G., and Korolchuk, V. I. (2017) mTORC1 as the main gateway to autophagy. Essays Biochem 61, 565-584 109. Kim, J., Kundu, M., Viollet, B., and Guan, K. L. (2011) AMPK and mTOR regulate autophagy through direct phosphorylation of Ulk1. Nature cell biology 13, 132-141 110. Hosokawa, N., Hara, T., Kaizuka, T., Kishi, C., Takamura, A., Miura, Y., Iemura, S., Natsume, T., Takehana, K., Yamada, N., Guan, J. L., Oshiro, N., and Mizushima, N. (2009) Nutrient-dependent mTORC1 association with the ULK1-Atg13-FIP200 complex required for autophagy. Molecular biology of the cell 20, 1981-1991 111. Dunlop, E. A., Hunt, D. K., Acosta-Jaquez, H. A., Fingar, D. C., and Tee, A. R. (2011) ULK1 inhibits mTORC1 signaling, promotes multisite Raptor phosphorylation and hinders substrate binding. Autophagy 7, 737-747 112. Martina, J. A., Chen, Y., Gucek, M., and Puertollano, R. (2012) MTORC1 functions as a transcriptional regulator of autophagy by preventing nuclear transport of TFEB. Autophagy 8, 903-914 113. Martina, J. A., Diab, H. I., Lishu, L., Jeong, A. L., Patange, S., Raben, N., and Puertollano, R. (2014) The nutrient-responsive transcription factor TFE3 promotes autophagy, lysosomal biogenesis, and clearance of cellular debris. Science signaling 7, ra9 150 114. Napolitano, G., Esposito, A., Choi, H., Matarese, M., Benedetti, V., Di Malta, C., Monfregola, J., Medina, D. L., Lippincott-Schwartz, J., and Ballabio, A. (2018) mTOR-dependent phosphorylation controls TFEB nuclear export. Nature communications 9, 3312 115. Palmieri, M., Impey, S., Kang, H., di Ronza, A., Pelz, C., Sardiello, M., and Ballabio, A. (2011) Characterization of the CLEAR network reveals an integrated control of cellular clearance pathways. Human molecular genetics 20, 3852-3866 116. Settembre, C., and Medina, D. L. (2015) TFEB and the CLEAR network. Methods in cell biology 126, 45-62 117. Tang, H. W., Weng, J. H., Lee, W. X., Hu, Y., Gu, L., Cho, S., Lee, G., Binari, R., Li, C., Cheng, M. E., Kim, A. R., Xu, J., Shen, Z., Xu, C., Asara, J. M., Blenis, J., and Perrimon, N. (2021) mTORC1-chaperonin CCT signaling regulates m(6)A RNA methylation to suppress autophagy. Proc Natl Acad Sci U S A 118, e2021945118 118. Herzig, S., and Shaw, R. J. (2018) AMPK: guardian of metabolism and mitochondrial homeostasis. Nature reviews. Molecular cell biology 19, 121-135 119. Margariti, A., Li, H., Chen, T., Martin, D., Vizcay-Barrena, G., Alam, S., Karamariti, E., Xiao, Q., Zampetaki, A., Zhang, Z., Wang, W., Jiang, Z., Gao, C., Ma, B., Chen, Y. G., Cockerill, G., Hu, Y., Xu, Q., and Zeng, L. (2013) XBP1 mRNA splicing triggers an autophagic response in endothelial cells through BECLIN-1 transcriptional activation. The Journal of biological chemistry 288, 859-872 120. Wei, Y., Pattingre, S., Sinha, S., Bassik, M., and Levine, B. (2008) JNK1-mediated phosphorylation of Bcl-2 regulates starvation-induced autophagy. Molecular cell 30, 678-688 121. Zalckvar, E., Berissi, H., Eisenstein, M., and Kimchi, A. (2009) Phosphorylation of Beclin 1 by DAP-kinase promotes autophagy by weakening its interactions with Bcl-2 and Bcl-XL. Autophagy 5, 720-722 122. B'Chir, W., Maurin, A. C., Carraro, V., Averous, J., Jousse, C., Muranishi, Y., Parry, L., Stepien, G., Fafournoux, P., and Bruhat, A. (2013) The eIF2alpha/ATF4 pathway is essential for stress-induced autophagy gene expression. Nucleic Acids Res 41, 7683-7699 123. Marciniak, S. J., Yun, C. Y., Oyadomari, S., Novoa, I., Zhang, Y., Jungreis, R., Nagata, K., Harding, H. P., and Ron, D. (2004) CHOP induces death by promoting protein synthesis and oxidation in the stressed endoplasmic reticulum. Genes & development 18, 3066-3077 124. Grotemeier, A., Alers, S., Pfisterer, S. G., Paasch, F., Daubrawa, M., Dieterle, A., Viollet, B., Wesselborg, S., Proikas-Cezanne, T., and Stork, B. (2010) AMPK-independent induction of autophagy by cytosolic Ca2+ increase. Cell Signal 22, 914-925 151 125. Ma, Q. (2013) Role of nrf2 in oxidative stress and toxicity. Annu Rev Pharmacol Toxicol 53, 401-426 126. Ren, L., Yang, H., Cui, Y., Xu, S., Sun, F., Tian, N., Hua, J., and Peng, S. (2017) Autophagy is essential for the differentiation of porcine PSCs into insulin-producing cells. Biochemical and biophysical research communications 488, 471-476 127. Jung, H. S., Chung, K. W., Won Kim, J., Kim, J., Komatsu, M., Tanaka, K., Nguyen, Y. H., Kang, T. M., Yoon, K. H., Kim, J. W., Jeong, Y. T., Han, M. S., Lee, M. K., Kim, K. W., Shin, J., and Lee, M. S. (2008) Loss of autophagy diminishes pancreatic beta cell mass and function with resultant hyperglycemia. Cell Metab 8, 318-324 128. Ebato, C., Uchida, T., Arakawa, M., Komatsu, M., Ueno, T., Komiya, K., Azuma, K., Hirose, T., Tanaka, K., Kominami, E., Kawamori, R., Fujitani, Y., and Watada, H. (2008) Autophagy is important in islet homeostasis and compensatory increase of beta cell mass in response to high-fat diet. Cell metabolism 8, 325-332 129. Riahi, Y., Wikstrom, J. D., Bachar-Wikstrom, E., Polin, N., Zucker, H., Lee, M. S., Quan, W., Haataja, L., Liu, M., Arvan, P., Cerasi, E., and Leibowitz, G. (2016) Autophagy is a major regulator of beta cell insulin homeostasis. Diabetologia 59, 1480-1491 130. Yamamoto, S., Kuramoto, K., Wang, N., Situ, X., Priyadarshini, M., Zhang, W., Cordoba-Chacon, J., Layden, B. T., and He, C. (2018) Autophagy Differentially Regulates Insulin Production and Insulin Sensitivity. Cell reports 23, 3286-3299 131. Bachar-Wikstrom, E., Wikstrom, J. D., Ariav, Y., Tirosh, B., Kaiser, N., Cerasi, E., and Leibowitz, G. (2013) Stimulation of autophagy improves endoplasmic reticulum stress-induced diabetes. Diabetes 62, 1227-1237 132. Bartolome, A., Guillen, C., and Benito, M. (2012) Autophagy plays a protective role in endoplasmic reticulum stress-mediated pancreatic beta cell death. Autophagy 8, 1757-1768 133. Quan, W., Hur, K. Y., Lim, Y., Oh, S. H., Lee, J. C., Kim, K. H., Kim, G. H., Kim, S. W., Kim, H. L., Lee, M. K., Kim, K. W., Kim, J., Komatsu, M., and Lee, M. S. (2012) Autophagy deficiency in beta cells leads to compromised unfolded protein response and progression from obesity to diabetes in mice. Diabetologia 55, 392-403 134. Supale, S., Li, N., Brun, T., and Maechler, P. (2012) Mitochondrial dysfunction in pancreatic beta cells. Trends in endocrinology and metabolism: TEM 23, 477-487 135. Twig, G., Elorza, A., Molina, A. J., Mohamed, H., Wikstrom, J. D., Walzer, G., Stiles, L., Haigh, S. E., Katz, S., Las, G., Alroy, J., Wu, M., Py, B. F., Yuan, J., Deeney, J. T., Corkey, B. E., and Shirihai, O. S. (2008) Fission and selective fusion govern mitochondrial segregation 152 and elimination by autophagy. EMBO J 27, 433-446 136. Sidarala, V., Pearson, G. L., Parekh, V. S., Thompson, B., Christen, L., Gingerich, M. A., Zhu, J., Stromer, T., Ren, J., Reck, E. C., Chai, B., Corbett, J. A., Mandrup-Poulsen, T., Satin, L. S., and Soleimanpour, S. A. (2020) Mitophagy protects beta cells from inflammatory damage in diabetes. JCI Insight 5, e141138 137. Choi, S. E., Lee, S. M., Lee, Y. J., Li, L. J., Lee, S. J., Lee, J. H., Kim, Y., Jun, H. S., Lee, K. W., and Kang, Y. (2009) Protective role of autophagy in palmitate-induced INS-1 beta-cell death. Endocrinology 150, 126-134 138. Martino, L., Masini, M., Novelli, M., Beffy, P., Bugliani, M., Marselli, L., Masiello, P., Marchetti, P., and De Tata, V. (2012) Palmitate activates autophagy in INS-1E beta-cells and in isolated rat and human pancreatic islets. PloS one 7, e36188 139. Wu, J., Kong, F., Pan, Q., Du, Y., Ye, J., Zheng, F., Li, H., and Zhou, J. (2017) Autophagy protects against cholesterol-induced apoptosis in pancreatic beta-cells. Biochemical and biophysical research communications 482, 678-685 140. Sheng, Q., Xiao, X., Prasadan, K., Chen, C., Ming, Y., Fusco, J., Gangopadhyay, N. N., Ricks, D., and Gittes, G. K. (2017) Autophagy protects pancreatic beta cell mass and function in the setting of a high-fat and high-glucose diet. Scientific reports 7, 16348 141. Chu, K. Y., O'Reilly, L., Mellet, N., Meikle, P. J., Bartley, C., and Biden, T. J. (2019) Oleate disrupts cAMP signaling, contributing to potent stimulation of pancreatic beta-cell autophagy. The Journal of biological chemistry 294, 1218-1229 142. Chu, K. Y., O'Reilly, L., Ramm, G., and Biden, T. J. (2015) High-fat diet increases autophagic flux in pancreatic beta cells in vivo and ex vivo in mice. Diabetologia 58, 2074-2078 143. Rivera, J. F., Gurlo, T., Daval, M., Huang, C. J., Matveyenko, A. V., Butler, P. C., and Costes, S. (2011) Human-IAPP disrupts the autophagy/lysosomal pathway in pancreatic beta-cells: protective role of p62-positive cytoplasmic inclusions. Cell Death Differ 18, 415-426 144. Shigihara, N., Fukunaka, A., Hara, A., Komiya, K., Honda, A., Uchida, T., Abe, H., Toyofuku, Y., Tamaki, M., Ogihara, T., Miyatsuka, T., Hiddinga, H. J., Sakagashira, S., Koike, M., Uchiyama, Y., Yoshimori, T., Eberhardt, N. L., Fujitani, Y., and Watada, H. (2014) Human IAPP-induced pancreatic beta cell toxicity and its regulation by autophagy. The Journal of clinical investigation 124, 3634-3644 145. Kim, J., Cheon, H., Jeong, Y. T., Quan, W., Kim, K. H., Cho, J. M., Lim, Y. M., Oh, S. H., Jin, S. M., Kim, J. H., Lee, M. K., Kim, S., Komatsu, M., Kang, S. W., and Lee, M. S. (2014) 153 Amyloidogenic peptide oligomer accumulation in autophagy-deficient beta cells induces diabetes. J Clin Invest 124, 3311-3324 146. Las, G., Serada, S. B., Wikstrom, J. D., Twig, G., and Shirihai, O. S. (2011) Fatty acids suppress autophagic turnover in beta-cells. The Journal of biological chemistry 286, 42534-42544 147. Lambelet, M., Terra, L. F., Fukaya, M., Meyerovich, K., Labriola, L., Cardozo, A. K., and Allagnat, F. (2018) Dysfunctional autophagy following exposure to pro-inflammatory cytokines contributes to pancreatic beta-cell apoptosis. Cell death & disease 9, 96 148. Muralidharan, C., Conteh, A. M., Marasco, M. R., Crowder, J. J., Kuipers, J., de Boer, P., and Linnemann, A. K. (2021) Pancreatic beta cell autophagy is impaired in type 1 diabetes. Diabetologia 64, 865-877 149. Masini, M., Bugliani, M., Lupi, R., del Guerra, S., Boggi, U., Filipponi, F., Marselli, L., Masiello, P., and Marchetti, P. (2009) Autophagy in human type 2 diabetes pancreatic beta cells. Diabetologia 52, 1083-1086 150. Liu, H., Javaheri, A., Godar, R. J., Murphy, J., Ma, X., Rohatgi, N., Mahadevan, J., Hyrc, K., Saftig, P., Marshall, C., McDaniel, M. L., Remedi, M. S., Razani, B., Urano, F., and Diwan, A. (2017) Intermittent fasting preserves beta-cell mass in obesity-induced diabetes via the autophagy-lysosome pathway. Autophagy 13, 1952-1968 151. Fujimoto, K., Hanson, P. T., Tran, H., Ford, E. L., Han, Z., Johnson, J. D., Schmidt, R. E., Green, K. G., Wice, B. M., and Polonsky, K. S. (2009) Autophagy regulates pancreatic beta cell death in response to Pdx1 deficiency and nutrient deprivation. The Journal of biological chemistry 284, 27664-27673 152. Goginashvili, A., Zhang, Z., Erbs, E., Spiegelhalter, C., Kessler, P., Mihlan, M., Pasquier, A., Krupina, K., Schieber, N., Cinque, L., Morvan, J., Sumara, I., Schwab, Y., Settembre, C., and Ricci, R. (2015) Insulin granules. Insulin secretory granules control autophagy in pancreatic beta cells. Science 347, 878-882 153. Napolitano, G., and Ballabio, A. (2016) TFEB at a glance. J Cell Sci 129, 2475-2481 154. Martina, J. A., and Puertollano, R. (2017) TFEB and TFE3: The art of multi-tasking under stress conditions. Transcription 8, 48-54 155.Settembre, C., Zoncu, R., Medina, D. L., Vetrini, F., Erdin, S., Erdin, S., Huynh, T., Ferron, M., Karsenty, G., Vellard, M. C., Facchinetti, V., Sabatini, D. M., and Ballabio, A. (2012) A lysosome-to-nucleus signalling mechanism senses and regulates the lysosome via mTOR and TFEB. EMBO J 31, 1095-1108 154 156. Puertollano, R., Ferguson, S. M., Brugarolas, J., and Ballabio, A. (2018) The complex relationship between TFEB transcription factor phosphorylation and subcellular localization. The EMBO journal 37, e98804 157. Roczniak-Ferguson, A., Petit, C. S., Froehlich, F., Qian, S., Ky, J., Angarola, B., Walther, T. C., and Ferguson, S. M. (2012) The transcription factor TFEB links mTORC1 signaling to transcriptional control of lysosome homeostasis. Science signaling 5, ra42 158. Steingrimsson, E., Copeland, N. G., and Jenkins, N. A. (2004) Melanocytes and the microphthalmia transcription factor network. Annu Rev Genet 38, 365-411 159. Hemesath, T. J., Steingrimsson, E., McGill, G., Hansen, M. J., Vaught, J., Hodgkinson, C. A., Arnheiter, H., Copeland, N. G., Jenkins, N. A., and Fisher, D. E. (1994) microphthalmia, a critical factor in melanocyte development, defines a discrete transcription factor family. Genes & development 8, 2770-2780 160. Pastore, N., Brady, O. A., Diab, H. I., Martina, J. A., Sun, L., Huynh, T., Lim, J. A., Zare, H., Raben, N., Ballabio, A., and Puertollano, R. (2016) TFEB and TFE3 cooperate in the regulation of the innate immune response in activated macrophages. Autophagy 12, 1240-1258 161. Martina, J. A., Diab, H. I., Brady, O. A., and Puertollano, R. (2016) TFEB and TFE3 are novel components of the integrated stress response. The EMBO journal 35, 479-495 162. Pastore, N., Vainshtein, A., Klisch, T. J., Armani, A., Huynh, T., Herz, N. J., Polishchuk, E. V., Sandri, M., and Ballabio, A. (2017) TFE3 regulates whole-body energy metabolism in cooperation with TFEB. EMBO Mol Med 9, 605-621 163. Zhao, G. Q., Zhao, Q., Zhou, X., Mattei, M. G., and de Crombrugghe, B. (1993) TFEC, a basic helix-loop-helix protein, forms heterodimers with TFE3 and inhibits TFE3-dependent transcription activation. Molecular and cellular biology 13, 4505-4512 164. Sardiello, M., Palmieri, M., di Ronza, A., Medina, D. L., Valenza, M., Gennarino, V. A., Di Malta, C., Donaudy, F., Embrione, V., Polishchuk, R. S., Banfi, S., Parenti, G., Cattaneo, E., and Ballabio, A. (2009) A gene network regulating lysosomal biogenesis and function. Science 325, 473-477 165.Medina, D. L., Fraldi, A., Bouche, V., Annunziata, F., Mansueto, G., Spampanato, C., Puri, C., Pignata, A., Martina, J. A., Sardiello, M., Palmieri, M., Polishchuk, R., Puertollano, R., and Ballabio, A. (2011) Transcriptional activation of lysosomal exocytosis promotes cellular clearance. Dev Cell 21, 421-430 166. Settembre, C., and Ballabio, A. (2011) TFEB regulates autophagy: an integrated coordination of cellular degradation and recycling processes. Autophagy 7, 1379-1381 155 167. Settembre, C., De Cegli, R., Mansueto, G., Saha, P. K., Vetrini, F., Visvikis, O., Huynh, T., Carissimo, A., Palmer, D., Klisch, T. J., Wollenberg, A. C., Di Bernardo, D., Chan, L., Irazoqui, J. E., and Ballabio, A. (2013) TFEB controls cellular lipid metabolism through a starvation-induced autoregulatory loop. Nature cell biology 15, 647-658 168. Palmieri, M., Pal, R., Nelvagal, H. R., Lotfi, P., Stinnett, G. R., Seymour, M. L., Chaudhury, A., Bajaj, L., Bondar, V. V., Bremner, L., Saleem, U., Tse, D. Y., Sanagasetti, D., Wu, S. M., Neilson, J. R., Pereira, F. A., Pautler, R. G., Rodney, G. G., Cooper, J. D., and Sardiello, M. (2017) mTORC1-independent TFEB activation via Akt inhibition promotes cellular clearance in neurodegenerative storage diseases. Nature communications 8, 14338 169. Pi, H., Li, M., Tian, L., Yang, Z., Yu, Z., and Zhou, Z. (2017) Enhancing lysosomal biogenesis and autophagic flux by activating the transcription factor EB protects against cadmium-induced neurotoxicity. Scientific reports 7, 43466 170. Pena-Llopis, S., Vega-Rubin-de-Celis, S., Schwartz, J. C., Wolff, N. C., Tran, T. A., Zou, L., Xie, X. J., Corey, D. R., and Brugarolas, J. (2011) Regulation of TFEB and V-ATPases by mTORC1. The EMBO journal 30, 3242-3258 171. Ploper, D., and De Robertis, E. M. (2015) The MITF family of transcription factors: Role in endolysosomal biogenesis, Wnt signaling, and oncogenesis. Pharmacological research 99, 36-43 172. Wang, L., Zhao, C., Zheng, T., Zhang, Y., Liu, H., Wang, X., Tang, X., Zhao, B., and Liu, P. (2022) Torin 1 alleviates impairment of TFEB-mediated lysosomal biogenesis and autophagy in TGFBI (p.G623_H626del)-linked Thiel-Behnke corneal dystrophy. Autophagy 18, 765-782 173. Alesi, N., Khabibullin, D., Rosenthal, D. M., Akl, E. W., Cory, P. M., Alchoueiry, M., Salem, S., Daou, M., Gibbons, W. F., Chen, J. A., Zhang, L., Filippakis, H., Graciotti, L., Miceli, C., Monfregola, J., Vilardo, C., Morroni, M., Di Malta, C., Napolitano, G., Ballabio, A., and Henske, E. P. (2024) TFEB drives mTORC1 hyperactivation and kidney disease in Tuberous Sclerosis Complex. Nature communications 15, 406 174. Malek, M., Wawrzyniak, A. M., Ebner, M., Puchkov, D., and Haucke, V. (2022) Inositol triphosphate-triggered calcium release from the endoplasmic reticulum induces lysosome biogenesis via TFEB/TFE3. The Journal of biological chemistry 298, 101740 175. Lim, H., Lim, Y. M., Kim, K. H., Jeon, Y. E., Park, K., Kim, J., Hwang, H. Y., Lee, D. J., Pagire, H., Kwon, H. J., Ahn, J. H., and Lee, M. S. (2018) A novel autophagy enhancer as a therapeutic agent against metabolic syndrome and diabetes. Nature communications 9, 1438 176. Zummo, F. P., Krishnanda, S. I., Georgiou, M., O'Harte, F. P., Parthsarathy, V., Cullen, K. 156 S., Honkanen-Scott, M., Shaw, J. A., Lovat, P. E., and Arden, C. (2022) Exendin-4 stimulates autophagy in pancreatic beta-cells via the RAPGEF/EPAC-Ca(2+)-PPP3/calcineurin-TFEB axis. Autophagy 18, 799-815 177. Scott, I., Webster, B. R., Chan, C. K., Okonkwo, J. U., Han, K., and Sack, M. N. (2014) GCN5-like protein 1 (GCN5L1) controls mitochondrial content through coordinated regulation of mitochondrial biogenesis and mitophagy. The Journal of biological chemistry 289, 2864-2872 178. Yuan, L., Yuan, Y., Liu, F., Li, L., Liu, J., Chen, Y., Cheng, J., and Lu, Y. (2021) PGC-1alpha alleviates mitochondrial dysfunction via TFEB-mediated autophagy in cisplatin-induced acute kidney injury. Aging (Albany NY) 13, 8421-8439 179. Chauhan, S., Goodwin, J. G., Chauhan, S., Manyam, G., Wang, J., Kamat, A. M., and Boyd, D. D. (2013) ZKSCAN3 is a master transcriptional repressor of autophagy. Molecular cell 50, 16-28 180. Cinque, L., De Leonibus, C., Iavazzo, M., Krahmer, N., Intartaglia, D., Salierno, F. G., De Cegli, R., Di Malta, C., Svelto, M., Lanzara, C., Maddaluno, M., Wanderlingh, L. G., Huebner, A. K., Cesana, M., Bonn, F., Polishchuk, E., Hubner, C. A., Conte, I., Dikic, I., Mann, M., Ballabio, A., Sacco, F., Grumati, P., and Settembre, C. (2020) MiT/TFE factors control ER-phagy via transcriptional regulation of FAM134B. The EMBO journal 39, e105696 181. Martina, J. A., and Puertollano, R. (2018) Protein phosphatase 2A stimulates activation of TFEB and TFE3 transcription factors in response to oxidative stress. The Journal of biological chemistry 293, 12525-12534 182. Ferron, M., Settembre, C., Shimazu, J., Lacombe, J., Kato, S., Rawlings, D. J., Ballabio, A., and Karsenty, G. (2013) A RANKL-PKCbeta-TFEB signaling cascade is necessary for lysosomal biogenesis in osteoclasts. Genes & development 27, 955-969 183. Yan, S. (2022) Role of TFEB in Autophagy and the Pathogenesis of Liver Diseases. Biomolecules 12, 672 184. Lin, C. W., Zhang, H., Li, M., Xiong, X., Chen, X., Chen, X., Dong, X. C., and Yin, X. M. (2013) Pharmacological promotion of autophagy alleviates steatosis and injury in alcoholic and non-alcoholic fatty liver conditions in mice. J Hepatol 58, 993-999 185. Thomes, P. G., Trambly, C. S., Fox, H. S., Tuma, D. J., and Donohue, T. M., Jr. (2015) Acute and Chronic Ethanol Administration Differentially Modulate Hepatic Autophagy and Transcription Factor EB. Alcohol Clin Exp Res 39, 2354-2363 186. Chao, X., Wang, S., Zhao, K., Li, Y., Williams, J. A., Li, T., Chavan, H., Krishnamurthy, 157 P., He, X. C., Li, L., Ballabio, A., Ni, H. M., and Ding, W. X. (2018) Impaired TFEB-Mediated Lysosome Biogenesis and Autophagy Promote Chronic Ethanol-Induced Liver Injury and Steatosis in Mice. Gastroenterology 155, 865-879 e812 187. Pastore, N., Blomenkamp, K., Annunziata, F., Piccolo, P., Mithbaokar, P., Maria Sepe, R., Vetrini, F., Palmer, D., Ng, P., Polishchuk, E., Iacobacci, S., Polishchuk, R., Teckman, J., Ballabio, A., and Brunetti-Pierri, N. (2013) Gene transfer of master autophagy regulator TFEB results in clearance of toxic protein and correction of hepatic disease in alpha-1-anti-trypsin deficiency. EMBO molecular medicine 5, 397-412 188. Jung, S. H., Lee, W., Park, S. H., Lee, K. Y., Choi, Y. J., Choi, S., Kang, D., Kim, S., Chang, T. S., Hong, S. S., and Lee, B. H. (2020) Diclofenac impairs autophagic flux via oxidative stress and lysosomal dysfunction: Implications for hepatotoxicity. Redox Biol 37, 101751 189. Mansueto, G., Armani, A., Viscomi, C., D'Orsi, L., De Cegli, R., Polishchuk, E. V., Lamperti, C., Di Meo, I., Romanello, V., Marchet, S., Saha, P. K., Zong, H., Blaauw, B., Solagna, F., Tezze, C., Grumati, P., Bonaldo, P., Pessin, J. E., Zeviani, M., Sandri, M., and Ballabio, A. (2017) Transcription Factor EB Controls Metabolic Flexibility during Exercise. Cell metabolism 25, 182-196 190. Erlich, A. T., Brownlee, D. M., Beyfuss, K., and Hood, D. A. (2018) Exercise induces TFEB expression and activity in skeletal muscle in a PGC-1alpha-dependent manner. Am J Physiol Cell Physiol 314, C62-C72 191.Tsunemi, T., Ashe, T. D., Morrison, B. E., Soriano, K. R., Au, J., Roque, R. A., Lazarowski, E. R., Damian, V. A., Masliah, E., and La Spada, A. R. (2012) PGC-1alpha rescues Huntington's disease proteotoxicity by preventing oxidative stress and promoting TFEB function. Science translational medicine 4, 142ra197 192. Zhang, J., Johnson, J. L., He, J., Napolitano, G., Ramadass, M., Rocca, C., Kiosses, W. B., Bucci, C., Xin, Q., Gavathiotis, E., Cuervo, A. M., Cherqui, S., and Catz, S. D. (2017) Cystinosin, the small GTPase Rab11, and the Rab7 effector RILP regulate intracellular trafficking of the chaperone-mediated autophagy receptor LAMP2A. The Journal of biological chemistry 292, 10328-10346 193. Israeli, T., Riahi, Y., Garzon, P., Louzada, R. A., Werneck-de-Castro, J. P., Blandino-Rosano, M., Yeroslaviz-Stolper, R., Kadosh, L., Tornovsky-Babeay, S., Hacker, G., Israeli, N., Agmon, O., Tirosh, B., Cerasi, E., Bernal-Mizrachi, E., and Leibowitz, G. (2022) Nutrient Sensor mTORC1 Regulates Insulin Secretion by Modulating beta-Cell Autophagy. Diabetes 71, 453-469 194. Pasquier, A., Pastore, N., D'Orsi, L., Colonna, R., Esposito, A., Maffia, V., De Cegli, R., 158 Mutarelli, M., Ambrosio, S., Tufano, G., Grimaldi, A., Cesana, M., Cacchiarelli, D., Delalleau, N., Napolitano, G., and Ballabio, A. (2023) TFEB and TFE3 control glucose homeostasis by regulating insulin gene expression. EMBO J 42, e113928 195. Zummo, F. P., Cullen, K. S., Honkanen-Scott, M., Shaw, J. A. M., Lovat, P. E., and Arden, C. (2017) Glucagon-Like Peptide 1 Protects Pancreatic beta-Cells From Death by Increasing Autophagic Flux and Restoring Lysosomal Function. Diabetes 66, 1272-1285 196. Kim, J., Park, K., Kim, M. J., Lim, H., Kim, K. H., Kim, S. W., Lee, E. S., Kim, H. H., Kim, S. J., Hur, K. Y., Kim, J. H., Ahn, J. H., Yoon, K. H., Kim, J. W., and Lee, M. S. (2021) An autophagy enhancer ameliorates diabetes of human IAPP-transgenic mice through clearance of amyloidogenic oligomer. Nature communications 12, 183 197. Hara, T., Nakamura, K., Matsui, M., Yamamoto, A., Nakahara, Y., Suzuki-Migishima, R., Yokoyama, M., Mishima, K., Saito, I., Okano, H., and Mizushima, N. (2006) Suppression of basal autophagy in neural cells causes neurodegenerative disease in mice. Nature 441, 885-889 198. Like, A. A., and Rossini, A. A. (1976) Streptozotocin-induced pancreatic insulitis: new model of diabetes mellitus. Science 193, 415-417 199. Ahn, C., An, B. S., and Jeung, E. B. (2015) Streptozotocin induces endoplasmic reticulum stress and apoptosis via disruption of calcium homeostasis in mouse pancreas. Molecular and cellular endocrinology 412, 302-308 200. Courtade, J. A., Wang, E. Y., Yen, P., Dai, D. L., Soukhatcheva, G., Orban, P. C., and Verchere, C. B. (2017) Loss of prohormone convertase 2 promotes beta cell dysfunction in a rodent transplant model expressing human pro-islet amyloid polypeptide. Diabetologia 60, 453-463 201. Montane, J., Bischoff, L., Soukhatcheva, G., Dai, D. L., Hardenberg, G., Levings, M. K., Orban, P. C., Kieffer, T. J., Tan, R., and Verchere, C. B. (2011) Prevention of murine autoimmune diabetes by CCL22-mediated Treg recruitment to the pancreatic islets. The Journal of clinical investigation 121, 3024-3028 202. Sabatini, P. V., Speckmann, T., Nian, C., Glavas, M. M., Wong, C. K., Yoon, J. S., Kin, T., Shapiro, A. M. J., Gibson, W. T., Verchere, C. B., and Lynn, F. C. (2018) Neuronal PAS Domain Protein 4 Suppression of Oxygen Sensing Optimizes Metabolism during Excitation of Neuroendocrine Cells. Cell Rep 22, 163-174 203. Wang, X., Hao, J., Metzger, D. L., Mui, A., Ao, Z., Verchere, C. B., Chen, L., Ou, D., and Warnock, G. L. (2009) Local expression of B7-H4 by recombinant adenovirus transduction in mouse islets prolongs allograft survival. Transplantation 87, 482-490 159 204. Aharoni-Simon, M., Shumiatcher, R., Yeung, A., Shih, A. Z., Dolinsky, V. W., Doucette, C. A., and Luciani, D. S. (2016) Bcl-2 Regulates Reactive Oxygen Species Signaling and a Redox-Sensitive Mitochondrial Proton Leak in Mouse Pancreatic beta-Cells. Endocrinology 157, 2270-2281 205. Luciani, D. S., White, S. A., Widenmaier, S. B., Saran, V. V., Taghizadeh, F., Hu, X., Allard, M. F., and Johnson, J. D. (2013) Bcl-2 and Bcl-xL Suppress Glucose Signaling in Pancreatic beta-Cells. Diabetes 62, 170-182 206. Hammar, E., Parnaud, G., Bosco, D., Perriraz, N., Maedler, K., Donath, M., Rouiller, D. G., and Halban, P. A. (2004) Extracellular matrix protects pancreatic beta-cells against apoptosis: role of short- and long-term signaling pathways. Diabetes 53, 2034-2041 207. Gamble, A., Pepper, A. R., Bruni, A., and Shapiro, A. M. J. (2018) The journey of islet cell transplantation and future development. Islets 10, 80-94 208. Hogrebe, N. J., Ishahak, M., and Millman, J. R. (2023) Developments in stem cell-derived islet replacement therapy for treating type 1 diabetes. Cell Stem Cell 30, 530-548 209. Lau, J., Henriksnas, J., Svensson, J., and Carlsson, P. O. (2009) Oxygenation of islets and its role in transplantation. Curr Opin Organ Transplant 14, 688-693 210. Carlsson, P. O., Liss, P., Andersson, A., and Jansson, L. (1998) Measurements of oxygen tension in native and transplanted rat pancreatic islets. Diabetes 47, 1027-1032 211. Bensellam, M., Duvillie, B., Rybachuk, G., Laybutt, D. R., Magnan, C., Guiot, Y., Pouyssegur, J., and Jonas, J. C. (2012) Glucose-induced O(2) consumption activates hypoxia inducible factors 1 and 2 in rat insulin-secreting pancreatic beta-cells. PloS one 7, e29807 212. Hogan, M. F., and Hull, R. L. (2017) The islet endothelial cell: a novel contributor to beta cell secretory dysfunction in diabetes. Diabetologia 60, 952-959 213. Okajima, Y., Matsuzaka, T., Miyazaki, S., Motomura, K., Ohno, H., Sharma, R., Shimura, T., Istiqamah, N., Han, S. I., Mizunoe, Y., Osaki, Y., Iwasaki, H., Yatoh, S., Suzuki, H., Sone, H., Miyamoto, T., Aita, Y., Takeuchi, Y., Sekiya, M., Yahagi, N., Nakagawa, Y., Tomita, T., and Shimano, H. (2022) Morphological and functional adaptation of pancreatic islet blood vessels to insulin resistance is impaired in diabetic db/db mice. Biochim Biophys Acta Mol Basis Dis 1868, 166339 214. Lavallard, V., Cottet-Dumoulin, D., Wassmer, C. H., Rouget, C., Parnaud, G., Brioudes, E., Lebreton, F., Bellofatto, K., Berishvili, E., Berney, T., and Bosco, D. (2020) NLRP3 Inflammasome is Activated in Rat Pancreatic Islets by Transplantation and Hypoxia. Scientific reports 10, 7011 160 215. Forrester, S. J., Kikuchi, D. S., Hernandes, M. S., Xu, Q., and Griendling, K. K. (2018) Reactive Oxygen Species in Metabolic and Inflammatory Signaling. Circulation research 122, 877-902 216. Semenza, G. L. (2009) Regulation of oxygen homeostasis by hypoxia-inducible factor 1. Physiology (Bethesda) 24, 97-106 217. Garcia-Contreras, M., Tamayo-Garcia, A., Pappan, K. L., Michelotti, G. A., Stabler, C. L., Ricordi, C., and Buchwald, P. (2017) Metabolomics Study of the Effects of Inflammation, Hypoxia, and High Glucose on Isolated Human Pancreatic Islets. J Proteome Res 16, 2294-2306 218. Samanta, D., and Semenza, G. L. (2017) Maintenance of redox homeostasis by hypoxia-inducible factors. Redox Biol 13, 331-335 219. Chun, Y., and Kim, J. (2021) AMPK-mTOR Signaling and Cellular Adaptations in Hypoxia. Int J Mol Sci 22, 9765 220. Bellot, G., Garcia-Medina, R., Gounon, P., Chiche, J., Roux, D., Pouyssegur, J., and Mazure, N. M. (2009) Hypoxia-induced autophagy is mediated through hypoxia-inducible factor induction of BNIP3 and BNIP3L via their BH3 domains. Molecular and cellular biology 29, 2570-2581 221. Mazure, N. M., and Pouyssegur, J. (2010) Hypoxia-induced autophagy: cell death or cell survival? Curr Opin Cell Biol 22, 177-180 222. Li, L., Wang, Z. V., Hill, J. A., and Lin, F. (2014) New autophagy reporter mice reveal dynamics of proximal tubular autophagy. J Am Soc Nephrol 25, 305-315 223. Mauthe, M., Orhon, I., Rocchi, C., Zhou, X., Luhr, M., Hijlkema, K. J., Coppes, R. P., Engedal, N., Mari, M., and Reggiori, F. (2018) Chloroquine inhibits autophagic flux by decreasing autophagosome-lysosome fusion. Autophagy 14, 1435-1455 224. Kuma, A., Hatano, M., Matsui, M., Yamamoto, A., Nakaya, H., Yoshimori, T., Ohsumi, Y., Tokuhisa, T., and Mizushima, N. (2004) The role of autophagy during the early neonatal starvation period. Nature 432, 1032-1036 225. Devaskar, S. U., Singh, B. S., Carnaghi, L. R., Rajakumar, P. A., and Giddings, S. J. (1993) Insulin II gene expression in rat central nervous system. Regul Pept 48, 55-63 226. Magnuson, M. A., and Osipovich, A. B. (2013) Pancreas-specific Cre driver lines and considerations for their prudent use. Cell metabolism 18, 9-20 227.Coupe, B., Ishii, Y., Dietrich, M. O., Komatsu, M., Horvath, T. L., and Bouret, S. G. (2012) 161 Loss of autophagy in pro-opiomelanocortin neurons perturbs axon growth and causes metabolic dysregulation. Cell Metab 15, 247-255 228. Kaushik, S., Arias, E., Kwon, H., Lopez, N. M., Athonvarangkul, D., Sahu, S., Schwartz, G. J., Pessin, J. E., and Singh, R. (2012) Loss of autophagy in hypothalamic POMC neurons impairs lipolysis. EMBO Rep 13, 258-265 229. Meng, Q., and Cai, D. (2011) Defective hypothalamic autophagy directs the central pathogenesis of obesity via the IkappaB kinase beta (IKKbeta)/NF-kappaB pathway. J Biol Chem 286, 32324-32332 230. Thorens, B., Tarussio, D., Maestro, M. A., Rovira, M., Heikkila, E., and Ferrer, J. (2015) Ins1(Cre) knock-in mice for beta cell-specific gene recombination. Diabetologia 58, 558-565 231. McCall, M., Toso, C., Emamaullee, J., Pawlick, R., Edgar, R., Davis, J., Maciver, A., Kin, T., Arch, R., and Shapiro, A. M. (2011) The caspase inhibitor IDN-6556 (PF3491390) improves marginal mass engraftment after islet transplantation in mice. Surgery 150, 48-55 232. Komatsu, M., Waguri, S., Ueno, T., Iwata, J., Murata, S., Tanida, I., Ezaki, J., Mizushima, N., Ohsumi, Y., Uchiyama, Y., Kominami, E., Tanaka, K., and Chiba, T. (2005) Impairment of starvation-induced and constitutive autophagy in Atg7-deficient mice. J Cell Biol 169, 425-434 233. Pei, X., Wang, H., Xu, P., Liang, K., and Yuan, L. (2022) The core autophagy protein ATG5 controls the polarity of the Golgi apparatus and insulin secretion of pancreatic beta cells. Biochem Biophys Res Commun 629, 26-33 234. Herrera, P. L. (2000) Adult insulin- and glucagon-producing cells differentiate from two independent cell lineages. Development 127, 2317-2322 235. Postic, C., Shiota, M., Niswender, K. D., Jetton, T. L., Chen, Y., Moates, J. M., Shelton, K. D., Lindner, J., Cherrington, A. D., and Magnuson, M. A. (1999) Dual roles for glucokinase in glucose homeostasis as determined by liver and pancreatic beta cell-specific gene knock-outs using Cre recombinase. J Biol Chem 274, 305-315 236.Mehran, A. E., Templeman, N. M., Brigidi, G. S., Lim, G. E., Chu, K. Y., Hu, X., Botezelli, J. D., Asadi, A., Hoffman, B. G., Kieffer, T. J., Bamji, S. X., Clee, S. M., and Johnson, J. D. (2012) Hyperinsulinemia drives diet-induced obesity independently of brain insulin production. Cell metabolism 16, 723-737 237. Chipurupalli, S., Ganesan, R., Martini, G., Mele, L., Reggio, A., Esposito, M., Kannan, E., Namasivayam, V., Grumati, P., Desiderio, V., and Robinson, N. (2022) Cancer cells adapt FAM134B/BiP mediated ER-phagy to survive hypoxic stress. Cell death & disease 13, 357 162 238. Fang, Y., Tan, J., and Zhang, Q. (2015) Signaling pathways and mechanisms of hypoxia-induced autophagy in the animal cells. Cell biology international 39, 891-898 239. Liang, R., Liu, N., Cao, J., Liu, T., Sun, P., Cai, X., Zhang, L., Liu, Y., Zou, J., Wang, L., Ding, X., Zhang, B., Shen, Z., Yoshida, S., Dou, J., and Wang, S. (2022) HIF-1alpha/FOXO1 axis regulated autophagy is protective for beta cell survival under hypoxia in human islets. Biochim Biophys Acta Mol Basis Dis 1868, 166356 240. Kim, D. S., Song, L., Wang, J., Wu, H., Gou, W., Cui, W., Kim, J. S., and Wang, H. (2018) Carbon Monoxide Inhibits Islet Apoptosis via Induction of Autophagy. Antioxid Redox Signal 28, 1309-1322 241. Tanemura, M., Saga, A., Kawamoto, K., Machida, T., Deguchi, T., Nishida, T., Sawa, Y., Doki, Y., Mori, M., and Ito, T. (2009) Rapamycin induces autophagy in islets: relevance in islet transplantation. Transplant Proc 41, 334-338 242. Tanemura, M., Ohmura, Y., Deguchi, T., Machida, T., Tsukamoto, R., Wada, H., Kobayashi, S., Marubashi, S., Eguchi, H., Ito, T., Nagano, H., Mori, M., and Doki, Y. (2012) Rapamycin causes upregulation of autophagy and impairs islets function both in vitro and in vivo. Am J Transplant 12, 102-114 243. Marasco, M. R., and Linnemann, A. K. (2018) beta-Cell Autophagy in Diabetes Pathogenesis. Endocrinology 159, 2127-2141 244. Button, R. W., Roberts, S. L., Willis, T. L., Hanemann, C. O., and Luo, S. (2017) Accumulation of autophagosomes confers cytotoxicity. The Journal of biological chemistry 292, 13599-13614 245. Szenci, G., Csizmadia, T., and Juhasz, G. (2023) The role of crinophagy in quality control of the regulated secretory pathway. J Cell Sci 136, jcs260741 246. Pasquier, A., Vivot, K., Erbs, E., Spiegelhalter, C., Zhang, Z., Aubert, V., Liu, Z., Senkara, M., Maillard, E., Pinget, M., Kerr-Conte, J., Pattou, F., Marciniak, G., Ganzhorn, A., Ronchi, P., Schieber, N. L., Schwab, Y., Saftig, P., Goginashvili, A., and Ricci, R. (2019) Lysosomal degradation of newly formed insulin granules contributes to beta cell failure in diabetes. Nature communications 10, 3312 247. Meszaros, G., Pasquier, A., Vivot, K., Goginashvili, A., and Ricci, R. (2018) Lysosomes in nutrient signalling: A focus on pancreatic beta-cells. Diabetes, obesity & metabolism 20 Suppl 2, 104-115 248. Mir, S. U., George, N. M., Zahoor, L., Harms, R., Guinn, Z., and Sarvetnick, N. E. (2015) Inhibition of autophagic turnover in beta-cells by fatty acids and glucose leads to apoptotic cell 163 death. The Journal of biological chemistry 290, 6071-6085 249. Trudeau, K. M., Colby, A. H., Zeng, J., Las, G., Feng, J. H., Grinstaff, M. W., and Shirihai, O. S. (2016) Lysosome acidification by photoactivated nanoparticles restores autophagy under lipotoxicity. The Journal of cell biology 214, 25-34 250. Jung, M., Lee, J., Seo, H. Y., Lim, J. S., and Kim, E. K. (2015) Cathepsin inhibition-induced lysosomal dysfunction enhances pancreatic beta-cell apoptosis in high glucose. PLoS One 10, e0116972 251. Platt, F. M., d'Azzo, A., Davidson, B. L., Neufeld, E. F., and Tifft, C. J. (2018) Lysosomal storage diseases. Nature reviews. Disease primers 4, 27 252. Ji, J., Petropavlovskaia, M., Khatchadourian, A., Patapas, J., Makhlin, J., Rosenberg, L., and Maysinger, D. (2019) Type 2 diabetes is associated with suppression of autophagy and lipid accumulation in beta-cells. Journal of cellular and molecular medicine 23, 2890-2900 253. Luzio, J. P., Hackmann, Y., Dieckmann, N. M., and Griffiths, G. M. (2014) The biogenesis of lysosomes and lysosome-related organelles. Cold Spring Harb Perspect Biol 6, a016840 254. Trivedi, P. C., Bartlett, J. J., Perez, L. J., Brunt, K. R., Legare, J. F., Hassan, A., Kienesberger, P. C., and Pulinilkunnil, T. (2016) Glucolipotoxicity diminishes cardiomyocyte TFEB and inhibits lysosomal autophagy during obesity and diabetes. Biochimica et biophysica acta 1861, 1893-1910 255. Munoz-Sanchez, J., and Chanez-Cardenas, M. E. (2019) The use of cobalt chloride as a chemical hypoxia model. J Appl Toxicol 39, 556-570 256. Nishida, Y., Arakawa, S., Fujitani, K., Yamaguchi, H., Mizuta, T., Kanaseki, T., Komatsu, M., Otsu, K., Tsujimoto, Y., and Shimizu, S. (2009) Discovery of Atg5/Atg7-independent alternative macroautophagy. Nature 461, 654-658 257. Rodriguez-Muela, N., Hernandez-Pinto, A. M., Serrano-Puebla, A., Garcia-Ledo, L., Latorre, S. H., de la Rosa, E. J., and Boya, P. (2015) Lysosomal membrane permeabilization and autophagy blockade contribute to photoreceptor cell death in a mouse model of retinitis pigmentosa. Cell Death Differ 22, 476-487 258. Lin, X. X., Sen, I., Janssens, G. E., Zhou, X., Fonslow, B. R., Edgar, D., Stroustrup, N., Swoboda, P., Yates, J. R., 3rd, Ruvkun, G., and Riedel, C. G. (2018) DAF-16/FOXO and HLH-30/TFEB function as combinatorial transcription factors to promote stress resistance and longevity. Nature communications 9, 4400 259. Carey, K. L., Paulus, G. L. C., Wang, L., Balce, D. R., Luo, J. W., Bergman, P., Ferder, I. 164 C., Kong, L., Renaud, N., Singh, S., Kost-Alimova, M., Nyfeler, B., Lassen, K. G., Virgin, H. W., and Xavier, R. J. (2020) TFEB Transcriptional Responses Reveal Negative Feedback by BHLHE40 and BHLHE41. Cell Rep 33, 108371 260. Pasquier, A., Pastore, N., D'Orsi, L., Colonna, R., Esposito, A., Maffia, V., De Cegli, R., Mutarelli, M., Ambrosio, S., Tufano, G., Grimaldi, A., Cesana, M., Cacchiarelli, D., Delalleau, N., Napolitano, G., and Ballabio, A. (2023) TFEB and TFE3 control glucose homeostasis by regulating insulin gene expression. The EMBO journal, e113928 261. Cunha, D. A., Hekerman, P., Ladriere, L., Bazarra-Castro, A., Ortis, F., Wakeham, M. C., Moore, F., Rasschaert, J., Cardozo, A. K., Bellomo, E., Overbergh, L., Mathieu, C., Lupi, R., Hai, T., Herchuelz, A., Marchetti, P., Rutter, G. A., Eizirik, D. L., and Cnop, M. (2008) Initiation and execution of lipotoxic ER stress in pancreatic beta-cells. J Cell Sci 121, 2308-2318 262. Lee, C., Lamech, L., Johns, E., and Overholtzer, M. (2020) Selective Lysosome Membrane Turnover Is Induced by Nutrient Starvation. Dev Cell 55, 289-297 e284 263. Hung, Y. H., Chen, L. M., Yang, J. Y., and Yang, W. Y. (2013) Spatiotemporally controlled induction of autophagy-mediated lysosome turnover. Nature communications 4, 2111 264. Trivedi, P. C., Bartlett, J. J., Mercer, A., Slade, L., Surette, M., Ballabio, A., Flibotte, S., Hussein, B., Rodrigues, B., Kienesberger, P. C., and Pulinilkunnil, T. (2020) Loss of function of transcription factor EB remodels lipid metabolism and cell death pathways in the cardiomyocyte. Biochim Biophys Acta Mol Basis Dis 1866, 165832 265. Wang, S., Ni, H. M., Chao, X., Wang, H., Bridges, B., Kumer, S., Schmitt, T., Mareninova, O., Gukovskaya, A., De Lisle, R. C., Ballabio, A., Pacher, P., and Ding, W. X. (2019) Impaired TFEB-mediated lysosomal biogenesis promotes the development of pancreatitis in mice and is associated with human pancreatitis. Autophagy 15, 1954-1969 266. Yang, C., Chen, X. C., Li, Z. H., Wu, H. L., Jing, K. P., Huang, X. R., Ye, L., Wei, B., Lan, H. Y., and Liu, H. F. (2021) SMAD3 promotes autophagy dysregulation by triggering lysosome depletion in tubular epithelial cells in diabetic nephropathy. Autophagy 17, 2325-2344 267. Ardestani, A., Lupse, B., Kido, Y., Leibowitz, G., and Maedler, K. (2018) mTORC1 Signaling: A Double-Edged Sword in Diabetic beta Cells. Cell Metab 27, 314-331 268. Bugliani, M., Mossuto, S., Grano, F., Suleiman, M., Marselli, L., Boggi, U., De Simone, P., Eizirik, D. L., Cnop, M., Marchetti, P., and De Tata, V. (2019) Modulation of Autophagy Influences the Function and Survival of Human Pancreatic Beta Cells Under Endoplasmic Reticulum Stress Conditions and in Type 2 Diabetes. Front Endocrinol (Lausanne) 10, 52 165 269. Thoreen, C. C., Kang, S. A., Chang, J. W., Liu, Q., Zhang, J., Gao, Y., Reichling, L. J., Sim, T., Sabatini, D. M., and Gray, N. S. (2009) An ATP-competitive mammalian target of rapamycin inhibitor reveals rapamycin-resistant functions of mTORC1. The Journal of biological chemistry 284, 8023-8032 270. Barlow, A. D., Xie, J., Moore, C. E., Campbell, S. C., Shaw, J. A., Nicholson, M. L., and Herbert, T. P. (2012) Rapamycin toxicity in MIN6 cells and rat and human islets is mediated by the inhibition of mTOR complex 2 (mTORC2). Diabetologia 55, 1355-1365 271. Sha, Y., Rao, L., Settembre, C., Ballabio, A., and Eissa, N. T. (2017) STUB1 regulates TFEB-induced autophagy-lysosome pathway. EMBO J 36, 2544-2552 272. Cora, D., Bussolino, F., and Doronzo, G. (2021) TFEB Signalling-Related MicroRNAs and Autophagy. Biomolecules 11, 985 273. Guo, H., Pu, M., Tai, Y., Chen, Y., Lu, H., Qiao, J., Wang, G., Chen, J., Qi, X., Huang, R., Tao, Z., and Ren, J. (2021) Nuclear miR-30b-5p suppresses TFEB-mediated lysosomal biogenesis and autophagy. Cell death and differentiation 28, 320-336 274. Zhang, L., and Jia, X. (2019) Down-regulation of miR-30b-5p protects cardiomyocytes against hypoxia-induced injury by targeting Aven. Cell Mol Biol Lett 24, 61 275. Sun, Y., Zhang, D., Liu, X., Li, X., Liu, F., Yu, Y., Jia, S., Zhou, Y., and Zhao, Y. (2018) Endoplasmic Reticulum Stress Affects Lipid Metabolism in Atherosclerosis Via CHOP Activation and Over-Expression of miR-33. Cell Physiol Biochem 48, 1995-2010 276. Ouimet, M., Ediriweera, H., Afonso, M. S., Ramkhelawon, B., Singaravelu, R., Liao, X., Bandler, R. C., Rahman, K., Fisher, E. A., Rayner, K. J., Pezacki, J. P., Tabas, I., and Moore, K. J. (2017) microRNA-33 Regulates Macrophage Autophagy in Atherosclerosis. Arterioscler Thromb Vasc Biol 37, 1058-1067 277. Cao, B., Chen, X., Li, Y., Zhou, T., Chen, N., Guo, Y., Zhao, M., Guo, C., Shi, Y., Wang, Q., Du, X., Zhang, L., and Li, Y. (2024) PDCD4 triggers alpha-synuclein accumulation and motor deficits via co-suppressing TFE3 and TFEB translation in a model of Parkinson's disease. NPJ Parkinsons Dis 10, 146 278. Kumar, S., Marriott, C. E., Alhasawi, N. F., Bone, A. J., and Macfarlane, W. M. (2017) The role of tumour suppressor PDCD4 in beta cell death in hypoxia. PLoS One 12, e0181235 279. Berney, T., and Secchi, A. (2009) Rapamycin in islet transplantation: friend or foe? Transpl Int 22, 153-161 280. Park, K., Sonn, S. K., Seo, S., Kim, J., Hur, K. Y., Oh, G. T., and Lee, M. S. (2023) 166 Impaired TFEB activation and mitophagy as a cause of PPP3/calcineurin inhibitor-induced pancreatic beta-cell dysfunction. Autophagy 19, 1444-1458 281. Li, Z. H., An, N., Huang, X. J., Yang, C., Wu, H. L., Chen, X. C., Pan, Q. J., and Liu, H. F. (2021) Cyclosporine A blocks autophagic flux in tubular epithelial cells by impairing TFEB-mediated lysosomal function. J Cell Mol Med 25, 5729-5743 282. Jeong, S. J., Stitham, J., Evans, T. D., Zhang, X., Rodriguez-Velez, A., Yeh, Y. S., Tao, J., Takabatake, K., Epelman, S., Lodhi, I. J., Schilling, J. D., DeBosch, B. J., Diwan, A., and Razani, B. (2021) Trehalose causes low-grade lysosomal stress to activate TFEB and the autophagy-lysosome biogenesis response. Autophagy 17, 3740-3752 283. Sun, Y., Wang, X., Chen, B., Huang, M., Ma, P., Xiong, L., Huang, J., Chen, J., Huang, S., and Liu, Y. (2022) TFEB-Mediated Lysosomal Restoration Alleviates High Glucose-Induced Cataracts Via Attenuating Oxidative Stress. Invest Ophthalmol Vis Sci 63, 26 284. Kuppan, P., Kelly, S., Polishevska, K., Hojanepesov, O., Seeberger, K., Korbutt, G. S., and Pepper, A. R. (2020) Co-localized immune protection using dexamethasone-eluting micelles in a murine islet allograft model. Am J Transplant 20, 714-725 285. Lopes-Ramos, C. M., Chen, C. Y., Kuijjer, M. L., Paulson, J. N., Sonawane, A. R., Fagny, M., Platig, J., Glass, K., Quackenbush, J., and DeMeo, D. L. (2020) Sex Differences in Gene Expression and Regulatory Networks across 29 Human Tissues. Cell reports 31, 107795 286. Triolo, M., Oliveira, A. N., Kumari, R., and Hood, D. A. (2022) The influence of age, sex, and exercise on autophagy, mitophagy, and lysosome biogenesis in skeletal muscle. Skelet Muscle 12, 13 287. Cortes, C. J., Miranda, H. C., Frankowski, H., Batlevi, Y., Young, J. E., Le, A., Ivanov, N., Sopher, B. L., Carromeu, C., Muotri, A. R., Garden, G. A., and La Spada, A. R. (2014) Polyglutamine-expanded androgen receptor interferes with TFEB to elicit autophagy defects in SBMA. Nat Neurosci 17, 1180-1189 288. Blessing, A. M., Rajapakshe, K., Reddy Bollu, L., Shi, Y., White, M. A., Pham, A. H., Lin, C., Jonsson, P., Cortes, C. J., Cheung, E., La Spada, A. R., Bast, R. C., Jr., Merchant, F. A., Coarfa, C., and Frigo, D. E. (2017) Transcriptional regulation of core autophagy and lysosomal genes by the androgen receptor promotes prostate cancer progression. Autophagy 13, 506-521 289. Nemoto, S., Kubota, T., and Ohno, H. (2022) Metabolic differences and differentially expressed genes between C57BL/6J and C57BL/6N mice substrains. PloS one 17, e0271651 290. Brand, M. D., and Nicholls, D. G. (2011) Assessing mitochondrial dysfunction in cells. Biochem J 435, 297-312 167 291. Salma, N., Song, J. S., Arany, Z., and Fisher, D. E. (2015) Transcription Factor Tfe3 Directly Regulates Pgc-1alpha in Muscle. J Cell Physiol 230, 2330-2336 292. Evans, T. D., Zhang, X., Jeong, S. J., He, A., Song, E., Bhattacharya, S., Holloway, K. B., Lodhi, I. J., and Razani, B. (2019) TFEB drives PGC-1alpha expression in adipocytes to protect against diet-induced metabolic dysfunction. Science signaling 12, eaau2281 293. Steingrimsson, E., Tessarollo, L., Reid, S. W., Jenkins, N. A., and Copeland, N. G. (1998) The bHLH-Zip transcription factor Tfeb is essential for placental vascularization. Development 125, 4607-4616 294. Steingrimsson, E., Tessarollo, L., Pathak, B., Hou, L., Arnheiter, H., Copeland, N. G., and Jenkins, N. A. (2002) Mitf and Tfe3, two members of the Mitf-Tfe family of bHLH-Zip transcription factors, have important but functionally redundant roles in osteoclast development. Proc Natl Acad Sci U S A 99, 4477-4482 295. Shang, D., Wang, L., Klionsky, D. J., Cheng, H., and Zhou, R. (2021) Sex differences in autophagy-mediated diseases: toward precision medicine. Autophagy 17, 1065-1076 296. Lemos, J. R. N., Baidal, D. A., Poggioli, R., Fuenmayor, V., Chavez, C., Alvarez, A., Linetsky, E., Mauvais-Jarvis, F., Ricordi, C., and Alejandro, R. (2022) Prolonged Islet Allograft Function is Associated With Female Sex in Patients After Islet Transplantation. J Clin Endocrinol Metab 107, e973-e979 297. Saber, N., Bruin, J. E., O'Dwyer, S., Schuster, H., Rezania, A., and Kieffer, T. J. (2018) Sex Differences in Maturation of Human Embryonic Stem Cell-Derived beta Cells in Mice. Endocrinology 159, 1827-1841 298. Skovso, S., Overby, P., Memar-Zadeh, J., Lee, J. T. C., Yang, J. C. C., Shanina, I., Sidarala, V., Levi-D'Ancona, E., Zhu, J., Soleimanpour, S. A., Horwitz, M. S., and Johnson, J. D. (2022) beta-Cell Cre Expression and Reduced Ins1 Gene Dosage Protect Mice From Type 1 Diabetes. Endocrinology 163, bqac144 299. O'Connell, E. J., Martinez, C. A., Liang, Y. G., Cistulli, P. A., and Cook, K. M. (2020) Out of breath, out of time: interactions between HIF and circadian rhythms. Am J Physiol Cell Physiol 319, C533-C540 300. Ewald, J. D., Lu, Y., Ellis, C. E., Worton, J., Kolic, J., Sasaki, S., Zhang, D., Dos Santos, T., Spigelman, A. F., Bautista, A., Dai, X. Q., Lyon, J. G., Smith, N. P., Wong, J. M., Rajesh, V., Sun, H., Sharp, S. A., Rogalski, J. C., Moravcova, R., Cen, H. H., Manning Fox, J. E., Consortium, H.-D., Atlas, E., Bruin, J. E., Mulvihill, E. E., Verchere, C. B., Foster, L. J., Gloyn, A. L., Johnson, J. D., Pepper, A. R., Lynn, F. C., Xia, J., and MacDonald, P. E. (2024) HumanIslets: An integrated platform for human islet data access and analysis. bioRxiv 168 301. Leite, N. C., Pelayo, G. C., and Melton, D. A. (2022) Genetic manipulation of stress pathways can protect stem-cell-derived islets from apoptosis in vitro. Stem Cell Reports 17, 766-774
Thesis/Dissertation
eng
Cell and Developmental Biology
Vancouver : University of British Columbia Library
Attribution-NonCommercial-NoDerivatives 4.0 International
Graduate
Luciani, Dan S.
Roles of autophagy and transcription factor EB (TFEB) in β-cell viability and function
Text