Functional characterization of stomatin-like protein 2
Panagiotis Mitsopoulos, M.Sc.
Department of Microbiology and Immunology
McGill University, Montréal, QC, Canada
August 2017
A thesis submitted to McGill University in partial fulfillment of the requirements of the degree of Doctor of Philosophy
© Panagiotis Mitsopoulos 2017
To my parents, Chris and Despina
You provided me the opportunity to pursue anything
Abstract
Stomatin-like protein (SLP)-2 is a widely expressed and highly conserved protein identified in a proteome analysis of detergent-insoluble, glycolipid-enriched microdomains of human T cells activated through the antigen receptor. SLP-2 partitions mostly in mitochondria but a low abundance pool is present at the plasma membrane. SLP-2 is upregulated upon T cell activation, and its deletion in T cell-specific SLP-2 knockout mice results in a post-transcriptional defect in
IL-2 production and decreased T cell responses. Mechanistically, SLP-2 binds the mitochondrial phospholipid cardiolipin and interacts with prohibitin (PHB) proteins, correlating with impaired cardiolipin compartmentalization and respiratory function in SLP-2-deficient T cells. In our current model of SLP-2 function, SLP-2 recruits PHBs to cardiolipin to form cardiolipin- enriched microdomains in the mitochondrial inner membrane. We hypothesize that these specialized microdomains may provide an environment necessary for the optimal assembly and function of multichain complexes, including respiratory supercomplexes and translation machinery.
Respiratory chain complexes naturally associate to form supercomplexes for optimal electron transfer and function, and cardiolipin is known to be important in this process. We found that in the absence of SLP-2, T cells exhibited defective assembly of complex I-III2 and I-
III2-IV1-3 supercomplexes but not of individual respiratory complexes. This correlated with increased respiration uncoupled from ATP production, a greater reliance on glycolysis, and significantly delayed T cell proliferation when glycolysis was inhibited, together demonstrating the importance of SLP-2 in mitochondrial respiratory function.
Mitochondrial translation occurs in close proximity to the mitochondrial inner membrane and is essential for generating core polypeptides of respiratory chain complexes. Given the
3 regulatory role of SLP-2 in processes closely associated with the mitochondrial inner membrane, we examined the impact of Slp-2 deletion on mitochondrial translation. 35S-Methionine/cysteine pulse labeling of SLP-2-deficient T cells showed a significant impairment in the production of several mitochondrial DNA-encoded polypeptides following T cell activation, including Cytb,
COXI, COXII, COXIII, and ATP6. This defect was post-transcriptional but Slp-2 deletion did not affect mitochondrial ribosome assembly.
Lastly, naturally occurring and synthetic compounds exist that bind PHBs and can have various beneficial effects on cell function. However, studies on these PHB ligands in the context of immunity are limited and the effect of SLP-2:PHB interaction on the activities of these compounds has never been examined. We found that the synthetic PHB ligands FL3 and Mel6 modulated T cell function. Specifically, FL3 elicited increased IL-2 production in T cells stimulated with superantigen but had no effect on α-CD3/CD28-induced T cell activation.
Conversely, Mel6 impaired IL-2 production when T cells were stimulated with superantigen but significantly increased α-CD3/CD28-induced T cell activation. Interestingly, experiments using
T cell-specific SLP-2 knockout mice and co-immunoprecipitation studies indicate that these effects are independent of SLP-2:PHB interaction.
Taken together, these results demonstrate the regulatory function of SLP-2 in multiple essential processes associated with the mitochondrial inner membrane, and support a model of functional membrane microdomain organization orchestrated by SLP-2.
4 Résumé
Stomatin-like protein (SLP)-2 est une protéine largement exprimée et hautement conservée qui a
été identifiée lors d’une analyse protéomique des microdomaines insoluble enrichis en glycolipides des cellules T humaines activées par le récepteur pour l'antigène. SLP-2 se retrouve principalement dans les mitochondries et de façon moins abondante dans la membrane plasmique. L’expression de SLP-2 est augmentée suite à l'activation des lymphocytes T et l’abolition de son expression dans les lymphocytes T de souris entraîne un défaut post- transcriptionnel de la production d'IL-2 et une diminution des réponses des lymphocytes T. SLP-
2 lie la cardiolipine des phospholipides mitochondriaux et interagit avec les prohibitines (PHBs), ce qui est en correlation avec une altération de la compartimentation de la cardiolipine et de la fonction respiratoire dans les cellules T déficientes en SLP-2. Selon le modèle actuel, SLP-2 recruterait des PHBs à la cardiolipine pour former des microdomaines enrichis en cardiolipine dans la membrane interne mitochondriale. Nous émettons l'hypothèse que ces microdomaines spécialisés fournissent un environnement nécessaire à l'assemblage et la fonction optimale des complexes à plusieurs chaînes, y compris les supercomplexes respiratoires et ceux de la machinerie de traduction.
Les complexes de la chaîne respiratoire s'associent naturellement pour former des supercomplexes qui sont nécessaires pour un transfert d'électrons optimal, et la cardiolipine est connue pour être importante dans ce processus. Nous avons constaté qu'en l'absence de SLP-2, les lymphocytes T présentaient un défaut dans l’assemblage des complexes I-III2 et I-III2-IV1-3 dans les supercomplexes, mais pas les complexes respiratoires individuels. Ce défaut est associé
à une augmentation de la respiration découplée de la production d'ATP, à une dépendance accrue
à la glycolyse et un retard significatif de la prolifération des lymphocytes T lorsque la glycolyse
5 est inhibée, le tout démontrant l'importance de SLP-2 dans la fonction respiratoire mitochondriale.
La traduction mitochondriale se produit à proximité immédiate de la membrane interne mitochondriale et est essentielle pour générer les polypeptides des complexes de la chaîne respiratoire. Compte tenu du rôle réglulateur de SLP-2 dans les processus étroitement associés à la membrane interne mitochondriale, nous avons examiné l'impact de la délétion de SLP-2 sur la traduction mitochondriale. Des études de marquage bref radioactif utilsant de la 35S- méthionine/cystéine sur des lymphocytes T déficients en SLP-2 ont montré une altération significative de la production de plusieurs polypeptides codés par l'ADN mitochondriale après activation des lymphocytes T, y compris Cytb, COXI, COXII, COXIII et ATP6. Cette altération
était post-transcriptionnel mais la suppression de SLP-2 n'a pas affecté l'assemblage du ribosome mitochondrial.
Finalement, il existe des composés naturels et synthétiques qui lient les PHBs et qui peuvent avoir divers effets bénéfiques sur la fonction cellulaire. Cependant, les études de ces ligands des PHBs sont limitées dans le contexte de l'immunité et l'effet de l'interaction de SLP-2:
PHB sur les activités de ces composés n'a jamais été examiné. Nous avons constaté que les ligands PHB synthétiques FL3 et Mel6 modulent la fonction des cellules T. Plus précisément,
FL3 a provoqué une augmentation de la production d'IL-2 dans les cellules T stimulées par un superantigène mais n'a eu aucun effet sur l'activation des lymphocytes T induite par le CD3 et le
CD28. À l'inverse, Mel6 a diminué la production d'IL-2 lorsque les cellules T ont été stimulées avec un superantigène mais a augmenté de façon significative l'activation des lymphocytes T induite par le CD3 et le CD28. Il est intéressant de noter que les expériences utilisant des souris décientes dans l’expression de SLP-2 spécifiquement dans les lymphocytes T et des études de
6 co-immunoprécipitation indiquent que ces effets sont indépendants de l'interaction entre SLP-2 et les PHBs.
L’ensemble de ces résultats démontre la fonction régulatrice du SLP-2 dans de multiples processus essentiels associés à la membrane interne de la mitochondrie et appuie un modèle d'organisation membranaire de microdomaines fonctionnels orchestrée par SLP-2.
7 Acknowledgements
First and foremost, I sincerely thank Dr. Joaquin Madrenas who has been an excellent mentor throughout the course of my studies. Next, I thank Dr. Sylvie Fournier for her support and for operating as my co-supervisor in the latter stages of my degree. I thank my advisory committee members Dr. Heidi McBride and Dr. Amit Bar-Or for helpful discussions and their guidance on my project. It was a pleasure to work alongside my many colleagues in the lab over the years. I thank Camille Stegen, Adam Peres, Dr. Zhigang Li, Dr. Andrey Gimenez, Joao Ormonde,
Andreea Damian, Dr. Benoit Levast, Ben Li, Cynthia Tang, Jelani Clarke, and Michelle Wang for helpful discussions and creating an exciting work environment. I thank Frank Chang, Calvin
Liang, Nikita Ber, and Orsi Lapohos, undergraduate students that I was fortunate to work with on this project. I also thank Dr. Darah Christie, Holly Lemmon, Teresa Fernandez, Luan Chau, and
Thu Chau, colleagues that together created a stimulating scientific environment and enjoyable atmosphere when I first joined the lab. I thank Dr. Stanley Dunn and Dr. Ewa Cairns for their support early in my studies, and I thank everyone in the Department of Microbiology and
Immunology at McGill University for providing an excellent learning environment during my studies.
Finally, I thank the most important people in my life: my father Chris, my mother
Despina, my brother Ahileas, and my sister Angela⎯I would not be me without you; Amanda, who brings me true joy and happiness; Camillo, Ashley, and my niece and nephews, the source of so many smiles; and Pouya, Sean, Holly, and Teresa for being such wonderful friends.
8 Table of Contents
ABSTRACT ...... 3 RÉSUMÉ ...... 5 ACKNOWLEDGEMENTS ...... 8 LIST OF TABLES ...... 11 LIST OF FIGURES ...... 12 LIST OF ABBREVIATIONS ...... 14 ORIGINAL CONTRIBUTIONS TO KNOWLEDGE ...... 18 CONTRIBUTIONS OF AUTHORS ...... 21 CHAPTER 1: INTRODUCTION ...... 24 SPFH PROTEINS AND MEMBRANE COMPARTMENTALIZATION ...... 24 The SPFH protein superfamily ...... 24 The stomatin family of proteins ...... 26 Stomatin-like protein 2 ...... 28 Prohibitins ...... 31 Prohibitin ligands ...... 33 T CELLS ...... 35 T cell bioenergetics ...... 36 MITOCHONDRIAL FUNCTION ...... 38 Respiratory chain supercomplexes ...... 39 Mitochondrial translation ...... 41 RATIONALE AND SPECIFIC AIMS ...... 45 FIGURE LEGENDS ...... 48 PREFACE TO CHAPTER 2 ...... 52 CHAPTER 2: IDENTIFICATION OF MULTIMOLECULAR COMPLEXES AND SUPERCOMPLEXES IN COMPARTMENT-SELECTIVE MEMBRANE MICRODOMAINS ...... 53 ABSTRACT ...... 54 INTRODUCTION AND RATIONALE ...... 55 DETERGENT-INSOLUBLE GLYCOSPHINGOLIPID (DIG) MICRODOMAIN ISOLATION ...... 58 PLASMA MEMBRANE ISOLATION ...... 64 CARDIOLIPIN-ENRICHED MITOCHONDRIAL MEMBRANE MICRODOMAIN ISOLATION ...... 69 MITOCHONDRIAL SUPERCOMPLEX IDENTIFICATION BY BLUE-NATIVE GEL ELECTROPHORESIS ...... 72 DISCUSSION ...... 79 SUMMARY ...... 81 ACKNOWLEDGEMENT ...... 81 Table 2.1. Preparation of gradient gels for blue-native gel electrophoresis...... 82 FIGURE LEGEND ...... 83 PREFACE TO CHAPTER 3 ...... 85 CHAPTER 3: STOMATIN-LIKE PROTEIN 2 IS REQUIRED FOR IN VIVO MITOCHONDRIAL RESPIRATORY CHAIN SUPERCOMPLEX FORMATION AND OPTIMAL CELL FUNCTION ...... 86 ABSTRACT ...... 87 INTRODUCTION ...... 88
9 RESULTS ...... 90 DISCUSSION ...... 97 MATERIALS AND METHODS ...... 102 ACKNOWLEDGMENTS ...... 107 FIGURE LEGENDS ...... 108 PREFACE TO CHAPTER 4 ...... 118 CHAPTER 4: STOMATIN-LIKE PROTEIN 2 DEFICIENCY RESULTS IN IMPAIRED MITOCHONDRIAL TRANSLATION ...... 119 ABSTRACT ...... 120 INTRODUCTION ...... 121 RESULTS ...... 123 DISCUSSION ...... 127 MATERIALS AND METHODS ...... 130 ACKNOWLEDGMENTS ...... 134 FIGURE LEGENDS ...... 135 PREFACE TO CHAPTER 5 ...... 143 CHAPTER 5: MODULATION OF T CELL ACTIVATION BY THE PROHIBITIN LIGAND DERIVATIVES FL3 AND MEL6 ...... 144 ABSTRACT ...... 145 INTRODUCTION ...... 146 RESULTS ...... 148 DISCUSSION ...... 151 MATERIALS AND METHODS ...... 154 ACKNOWLEDGMENTS ...... 156 FIGURE LEGENDS ...... 157 CHAPTER 6: DISCUSSION ...... 164 APPENDIX ...... 170 SUPPLEMENTARY MATERIALS AND METHODS ...... 170 SUPPLEMENTARY FIGURE LEGENDS ...... 172 REFERENCES ...... 175
10 List of Tables
Chapter 2
Table 2.1. Preparation of gradient gels for blue-native gel electrophoresis...... 82
11 List of Figures
Chapter 1
Figure 1.1. SLP-2 secondary structure ...... 49 Figure 1.2. Structures of PHB ligands and derivatives ...... 50 Figure 1.3. Model of SLP-2 function in the context of T cell activation ...... 51
Chapter 2
Figure 2.1. Procedure for two-dimensional blue-native gel electrophoresis (2D-BN-PAGE) .... 84
Chapter 3
Figure 3.1. SLP-2-deficient T cells have an impaired mitochondrial respiration profile characterized by increased uncoupled respiration and reliance on glycolysis ...... 112 Figure 3.2. Reduced RCS detection and cell growth in Slp-2-/- MEFs ...... 113 Figure 3.3. SLP-2-deficient T cells have defective RCS expression ...... 114 Figure 3.4. Formation of individual respiratory chain complexes is not altered by Slp-2 deletion ...... 115 Figure 3.5. Deletion of Slp-2 results in delayed T cell proliferation when glycolysis is restricted ...... 116 Figure 3.6. Deletion of Slp-2 results in defective post-transcriptional IL-2 production when glycolysis is restricted ...... 117
Chapter 4
Figure 4.1. Mitochondrial translation is impaired in SLP-2-deficient T cells upon stimulation 138 Figure 4.2. Mitochondrial DNA stability and PGC-1α expression are not affected by deletion of Slp-2 in T cells ...... 139 Figure 4.3. SLP-2 deficiency does not impair mitochondrial transcription in T cells ...... 140 Figure 4.4. SLP-2 migrates in sucrose-density gradients similarly to mitoribosome complexes but does not affect mitoribosome assembly ...... 141 Figure 4.5. IL-2 production is decreased in SLP-2 T-KO T cells following activation ...... 142
Chapter 5
Figure 5.1. Activation of Jurkat T cells is increased by FL3 and decreased by Mel6 in response to superantigen ...... 159 Figure 5.2. Mel6 activates T cells differentially depending on the stimulus ...... 160 Figure 5.3. Effects of FL3 or Mel6 on T cell signaling following activation ...... 161
12 Figure 5.4. Effects of FL3 or Mel6 on IL-2 responses following activation of SLP-2-deficient T cells ...... 162 Figure 5.5. SLP-2:PHB1/2 and PHB1:PHB2 interactions are not affected by FL3 or Mel6 ..... 163
Appendix
Supplementary Figure 1. SLP-2-deficiency results in increased spare respiratory capacity in activated T cells ...... 173 Supplementary Figure 2. Slp-2 is systemically deleted in viable Slp-2-/- mice ...... 174
13 List of Abbreviations
AAA ATPases associated with diverse cellular activities
ASIC Acid-sensing ion channel
ATP Adenosine triphosphate
B2M β2 microglobulin
BN-PAGE Blue-native polyacrylamide gel electrophoresis
Co-IP Co-immunoprecipitation
CD Cluster of differentiation cDNA Complimentary deoxyribonucleic acid
CFSE Carboxyfluorescein diacetate succinimidyl ester
CTLA-4 Cytotoxic T lymphocyte-associated antigen 4
Cre-ERT2 Cre recombinase-estrogen receptor T2
DCFDA 5-(and-6)-chloromethyl-2',7'-dichlorodihydrofluorescein diacetate,
acetyl ester
DDM n-Dodecyl β-D-maltoside
DEG/ENaC Degenerin/epithelial sodium channel
DIG Detergent-insoluble glycosphingolipid
DI-GEM Detergent-insoluble glycolipid-enriched microdomains
DNA Deoxyribonucleic acid
ECAR Extracellular acidification rate
EDTA Ethylenediaminetetraacetic acid
ELISA Enzyme-linked immunosorbent assay
14 ERK Extracellular signal-regulated kinase
FAD Flavin adenine dinucleotide
FCCP Carbonyl cyanide 4-(trifluoromethoxy)phenylhydrazone
GAPDH Glyceraldehyde 3-phosphate dehydrogenase
GFP Green fluorescent protein
GTPase Guanosine triphosphate hydrolase
HIG Hypoxia-induced gene
HPRT Hypoxanthine guanine phosphoribosyltransferase
IFN Interferon
KO Knockout
IκB Inhibitor of κB
IL Interleukin
LAT Linker for activation of T cells
LSU Ribosomal large subunit
MEF Mouse embryonic fibroblast
MFI Mean fluorescence intensity
MHC Major histocompatibility complex mRNA Messenger ribonucleic acid
MRP Mitochondrial ribosomal protein mtDNA Mitochondrial deoxyribonucleic acid mTOR Mechanistic target of rapamycin
NAD Nicotinamide adenine dinucleotide
NBT Nitro blue tetrazolium
15 NF-AT Nuclear factor of activated T cells
NF-κB Nuclear factor-κB
PAA Polymethacrylic acid
PAGE Polyacrylamide gel electrophoresis
PBS Phosphate-buffered saline
PCR Polymerase chain reaction
PGC-1α Proliferator-activated receptor γ coactivator-1α
PHB Prohibitin
PLC Phospholipase C
PMCB Plasma membrane coating buffer
PMSF Phenylmethyl sulfonyl fluoride pSMAC Peripheral supramolecular activation cluster
PVDF Polyvinylidene fluoride qRT-PCR Quantitative reverse-transcriptase polymerase chain reaction
OCR Oxygen consumption rate
OPA1 Optic atrophy 1
OXPHOS Oxidative phosphorylation
RCS Respiratory chain supercomplexes
ROS Reactive oxygen species
RPL19 Ribosomal protein L19 rRNA Ribosomal ribonucleic acid
SBG Serva Blue G
SCAF1 Supercomplex assembly factor 1
16 SDS Sodium dodecyl sulfate
SEE Staphylococcal enterotoxin E
SLP Stomatin-like protein
SPFH Stomatin/prohibitin/flotillin/HflK/C
SSU Ribosomal small subunit
TCA Tricarboxylic acid
TCR T cell receptor
TEMED Tetramethylethylenediamine
TGF Transforming growth factor
TFH T follicular helper
T-KO T cell-specific knockout
TIM Translocase of the inner mitochondrial membrane
TH T helper
TLR Toll-like receptor
TMRM Tetramethylrhodamine methyl ester
TOM Translocase of the outer mitochondrial membrane tRNA Transfer ribonucleic acid
WCL Whole-cell lysate
WT Wild type
Zap70 Zeta chain-associated protein kinase 70
17 Original Contributions to Knowledge
Chapter 3
Mitsopoulos, P., Chang, Y.H., Wai, T., König, T., Dunn, S.D., Langer, T., Madrenas, J.
Stomatin-like protein 2 is required for in vivo mitochondrial respiratory chain supercomplex formation and optimal cell function. Mol Cell Biol. 2015; 35: 1838-47.
• SLP-2 deficiency results in increased uncoupled respiration in activated T cells,
particularly under conditions of limited glycolytic activity.
• SLP-2 deficiency results in an increased reliance on glycolysis in activated T cells.
• SLP-2 is specifically required for the formation of RCS (e.g., (I-III2)1-2, I-III2-IV1-3, I-III2,
and III2-IV arrangements) in mouse T cells and MEFs, but does not influence the
assembly of individual respiratory chain complexes.
• Under conditions of restricted glycolytic activity (i.e., when T cells are forced to rely on
OXPHOS for energy production), Slp-2 deletion results in delayed T cell proliferation
that cannot be rescued by recombinant IL-2 protein supplementation to overcome the IL-
2 defect in SLP-2-deficient T cells.
Chapter 4
Mitsopoulos, P., Lapohos, O., Weraarpachai, W., Antonicka, H., Chang, Y.H., Madrenas, J.
Stomatin-like protein 2 deficiency results in impaired mitochondrial translation. PLoS One.
2017; 12: e0179967.
18 • SLP-2 is required for efficient translation of most or all mitochondria-encoded peptides in
activated T cells, indicating that SLP-2 is a general regulator of mitochondrial translation.
• SLP-2 co-migrates with mitoribosomes in sucrose-density centrifugation experiments and
may interact indirectly with mitoribosomes, but its deletion does not affect mitoribosome
assembly.
Chapter 5
Mitsopoulos, P., Lapohos, O., Désaubry, L., Madrenas, J. Modulation of T cell activation by the prohibitin ligand derivatives FL3 and Mel6. 2017; Manuscript in preparation.
• The PHB ligand FL3 modulates T cell activation since it consistently elicits increased IL-
2 production when T cells are stimulated with SEE but not α-CD3/CD28 antibodies.
• The PHB ligand derivative Mel6 modulates T cell activation since it decreases IL-2
levels upon SEE stimulation but elicits an increase in α-CD3/CD28-induced T cell
activation.
• The mechanism(s) underlying the effects of FL3 and Mel6 on the modulation of T cell
activation do not involve SLP-2, and neither FL3 or Mel6 interfere with SLP-2:PHB1/2
or PHB1:PHB2 interaction.
Appendix
• SLP-2 deficiency results in significantly increased spare respiratory capacity in activated
T cells.
19 • The generation of Slp-2-/- mice was achieved by a non-conventional method involving the
breeding of adult, tamoxifen-inducible, Slp-2-floxed, β-actin Cre-ERT2 mice following
induction of Cre recombinase activity. These mice are viable, contain no SLP-2 at the
DNA or protein levels, and exhibit no gross phenotypic abnormalities.
20 Contributions of Authors
Chapter 1: Introduction
Written by Mitsopoulos P.
Chapter 2
Mitsopoulos, P., Madrenas, J. Identification of multimolecular complexes and supercomplexes in compartment-selective membrane microdomains. Methods Cell Biol. 2013; 117: 411-31.
Mitsopoulos, P. wrote the manuscript. Madrenas, J. contributed to manuscript preparation and edited final manuscript.
Chapter 3
Mitsopoulos, P., Chang, Y.H., Wai, T., König, T., Dunn, S.D., Langer, T., Madrenas, J.
Stomatin-like protein 2 is required for in vivo mitochondrial respiratory chain supercomplex formation and optimal cell function. Mol Cell Biol. 2015; 35: 1838-47.
Mitsopoulos, P. contributed to experimental design, data interpretation, performed experiments for Figures 3.1, 3.3, 3.4, and 3.6, and wrote the manuscript. Chang, Y.H. performed experiments for Figure 3.5. Wai, T. performed experiments for Figure 3.2 and contributed to experimental design. König, T. performed experiments for Figure 3.2. Dunn, S.D. and Langer, T. contributed to experimental design and edited the manuscript. Madrenas, J. is the principal investigator, and contributed to experimental design, data interpretation and manuscript preparation.
21
Chapter 4
Mitsopoulos, P., Lapohos, O., Weraarpachai, W., Antonicka, H., Chang, Y.H., Madrenas, J.
Stomatin-like protein 2 deficiency results in impaired mitochondrial translation. PLoS One.
2017; 12: e0179967.
Mitsopoulos, P. contributed to experimental design, data interpretation, performed or contributed to all experiments, and wrote the manuscript. Lapohos, O. contributed to experiments for Figures
4.2B and 4.3. Weraarpachai, W. contributed to experiments for Figure 4.1. Antonicka, H. contributed to experiments for Figure 4.1 and edited the manuscript. Chang, Y.H. performed experiments for Figure 4.5. Madrenas, J. is the principal investigator, and contributed to experimental design, data interpretation and manuscript preparation.
Chapter 5
Mitsopoulos, P., Lapohos, O., Désaubry, L., Madrenas, J. Modulation of T cell activation by the prohibitin ligand derivatives FL3 and Mel6. 2017; Manuscript in preparation.
Mitsopoulos, P. contributed to experimental design, data interpretation, performed all experiments, and wrote the manuscript. Lapohos, O., contributed to experiments for Figure 5.4.
Désaubry, L. synthesized the FL3 and Mel6 compounds. Madrenas, J. is the principal investigator, and contributed to experimental design, data interpretation and manuscript preparation.
22 Chapter 6: Discussion
Written by Mitsopoulos, P.
Appendix
Mitsopoulos, P. contributed to experimental design, data interpretation, and performed all experiments. Madrenas, J. is the principal investigator and contributed to experimental design and data interpretation.
23 Chapter 1: Introduction
SPFH Proteins and Membrane Compartmentalization
Biological membranes are composed of a multitude of lipid, protein, and sugar species, and it is estimated that one-third of the genome encodes membrane proteins [1]. Membranes assemble via the hydrophobic properties of lipids, and interactions between membrane components can spontaneously lead to domains of specific composition [1]. An important characteristic of biological membranes is their asymmetry, a feature that is actively maintained [2, 3].
Importantly, many membrane-associated processes including cell signaling, vesicle fusion and budding, cellular adhesion, and cell division require transient clustering or separation of specific membrane components [4-6].
The organization of biological membranes is due in part to certain proteins that act as scaffolds for lipids and proteins. For instance, in mitochondrial membranes it has been proposed that certain members of the SPFH protein superfamily, including the integral membrane proteins prohibitins (PHBs), have a role as scaffolds in membrane organization and may contribute to the recruitment of specific proteins to functional sites [7]. There are many proteins in the SPFH superfamily in which there is little known about their function. The aim of this work is to characterize the function of a specific SPFH protein, namely stomatin-like protein (SLP)-2, and its role in membrane-associated processes.
The SPFH protein superfamily
The SPFH protein superfamily consists of Stomatins, Prohibitins, Flotillins, and the bacterial
HflK/C proteins due to their shared, highly conserved SPFH protein domain [8]. These proteins are associated with membranes of various organelles, including mitochondria, Golgi, early
24 endosomes, endoplasmic reticulum, and plasma membrane [9]. Though the function of the SPFH domain remains unclear, it is proposed that the membrane association of SPFH superfamily proteins may occur due to the presence of this domain [8, 10, 11]. For example, the SPFH domain of podocin and its Caenorhabditis elegans homologue MEC-2 were proposed to facilitate lipid association by binding to cholesterol [12]. It is thought that the SPFH protein superfamily plays a role in the organization of membrane domains by directly binding to lipids and other proteins to facilitate membrane compartmentalization. Indeed, many SPFH family members, including members of the stomatin family, are enriched in detergent-insoluble glycolipid-enriched microdomains (DI-GEMs) [9].
PHBs are mitochondrial proteins that are involved in an array of cellular activities, and are found in other cellular compartments including the nucleus and plasma membrane [13, 14].
These proteins function in the mitochondria as chaperone proteins [15], and are involved in the compartmentalization of mitochondrial membranes [7, 16] and mitochondrial biogenesis [15].
PHBs have also been implicated in other cellular processes including the organization of mitochondrial DNA nucleoids [17], cell proliferation [18], and the regulation of lifespan [19,
20]. PHBs will be reviewed in more detail below.
Flotillins are ubiquitously expressed proteins found in intracellular membranes and at the plasma membrane [21-23]. The two homologous members of the flotillin protein family, namely flotillin-1 and flotillin-2, form homo- and hetero-oligomeric complexes [24], and their expression patterns influence one other [25]. These proteins do not span the membrane but rather associate with the membrane by interacting with other membrane proteins [26]. Flotillins are proposed to organize lipid rafts and act as scaffolding proteins within them [21]. Originally these proteins were discovered in axon regeneration of goldfish retinal ganglion cells [27]. Since then,
25 flotillins have been associated with several other cellular processes attributable to their function in lipid rafts, including endocytosis [28, 29], insulin signaling [30], cell proliferation [31], membrane protein recruitment [32], T cell activation [33], neuronal differentiation [34], and tumor progression [35].
The bacterial proteins HflK and HflC (together, HflK/C) have been identified as modulators of FtsH, an ATP-dependent protease anchored to the cytoplasmic membrane of bacteria [36]. Together, FtsH and HflK/C form a high-molecular weight complex at the cytoplasmic membrane known as the FtsH holoenzyme [36, 37]. In this way, HflK/C acts as a negative regulator of membrane substrates for FtsH [36, 38]. However, HflK/C does not appear to be essential for overall bacterial function [36, 39]. Taken together, the research conducted thus far on the function of SPFH proteins points to an involvement in the formation or regulation of
DI-GEMs that have importance in diverse cellular activities.
The stomatin family of proteins
The stomatin family is a highly conserved family of proteins with homologues present widely across evolution, including in prokaryotes, archaea, fungi, nematodes, insects, plants, amphibians, birds, rodents, primates, and others [40]. It consists of five members: stomatin, SLP-
1, SLP-2, SLP-3, and podocin [41-44]. Stomatin, the first member of this family to be identified, is phosphorylated and palmitoylated [45, 46], can form oligomers, and associate with lipid rafts
[47, 48]. Stomatin was originally found to be deficient in erythrocytes of patients with stomatocytosis [43], a hemolytic anemia caused by increased membrane permeability to sodium and potassium [49]. This suggested a role for this protein in the regulation of cation channels and it was thought to be the cause of stomatocytosis [50, 51]. Thus the protein was named stomatin,
26 referencing the distinct mouth shape of erythrocytes in these patients [43]. Although it was later shown following the generation of stomatin knockout mice that this protein was not responsible for the erythrocyte phenotype in stomatocytosis patients [52], further research pointed to a role for stomatin in ion channel regulation. There is evidence that stomatin homologues in
Caenorhabditis elegans are involved in the regulation of the degenerin/epithelial sodium channel
DEG/ENaC family, including the acid-sensing ion channels (ASIC) [53, 54], a finding corroborated in human and mouse systems [55]. Moreover, the stomatin homologue unc-1 in C. elegans was identified to control sensitivity to diethyl ether [56], a phenotype later observed in stomatin knockout mice [57]. The mechanism of ion channel regulation by stomatin is not yet understood.
SLP-1 is the least-studied mammalian member of the stomatin family. In addition to its
N-terminal stomatin domain, it contains a sterol carrier protein-2 domain at its C-terminus that is unique among mammalian stomatin family members [58]. Human SLP-1 is predominantly expressed in the nervous system [42], and was shown to localize to late endosomes and can affect stomatin recruitment to these compartments [59]. SLP-1 also localizes in vesicles and at the plasma membrane in neurites of dorsal root ganglia neurons, and was found to specifically inhibit the ASIC1a subunit in these cells [60]. Furthermore, SLP-1 was found to bind and inhibit the degradation of the F-box protein Fbw7-γ, a specificity factor for the Skp1-Cul1-F-box protein ubiquitin ligase complex that targets proteins required for cellular proliferation [61].
SLP-3 is not widely expressed and is mainly studied in the dorsal root ganglia. It was first identified in olfactory sensory neurons [62] and has 77% amino acid sequence similarity with
MEC-2 in C. elegans [63]. It was found that SLP-3 knockout mice have decreased mechanosensitivity in neurons of the dorsal root ganglia resulting in reduced tactile acuity, and
27 that SLP-3 can inhibit the ASIC proteins similar to stomatin [64]. Furthermore, an N-terminal hydrophobic region of SLP-3 is required for its localization in highly mobile vesicles and is important for interaction with ASIC proteins [65], and the interaction of SLP-3 with ASIC proteins modulates nociceptor mechanosensitivity [66]. Finally, SLP-3 binds cholesterol to facilitate force transfer [67] and inhibition of SLP-3 oligomerization can modulate touch hypersensitivity [68].
Podocin is expressed in the kidneys and its mutation is the most common cause of heriditary nephrotic syndrome [69]. Podocin was first identified due to its mutation in patients with familial idiopathic nephrotic syndrome [70]. This protein forms oligomers and acts as a scaffolding protein that organizes lipid-protein domains involving nephrin and the actin cytoskeleton [71]. Podocin is required for the formation of the slit diaphragm, a specialized intercellular junction that connects neighboring podocyte foot processes within glomeruli [72].
Furthermore, like other stomatin family members podocin has been studied in mechanotransduction and was found to associate with membrane cholesterol and directly bind the TRP ion channel TRPV-6 in the slit diaphragm [12, 73]. In general, the stomatin family members have been identified in DI-GEMs and are thought to organize functional domains in the membrane that may be important for complex receptor assemblies and cellular processes.
Stomatin-like protein 2
The human Slp-2 gene is located on chromosome 9p13 and contains 1071 nucleotides encoding a
357 amino acid protein [44, 74]. It was first identified in mature human erythrocytes where it was associated with high molecular weight complexes, but was subsequently found to be widely expressed in many tissues including heart, brain, kidney, lung, placenta, pancreas, skeletal
28 muscle, and liver [44, 74]. SLP-2 shares sequence homology with other stomatin family members in the predicted C-terminal and proximal β-sheet domains but is unique in that it lacks a putative hydrophobic transmembrane domain and palmitoylation sites [10, 44, 75], leading to the suggestion that it diverged from the other stomatin family members early in evolution [40,
44]. Human and mouse SLP-2 are highly conserved and share several putative myristoylation sites thought to facilitate membrane association [10, 75, 76]. SLP-2 also contains an N-terminal mitochondrial targeting sequence [10, 77, 78], a feature unique among stomatin family members.
A model of the SLP-2 secondary structure is depicted in Figure 1.1. Taken together, these features make SLP-2 the most unique member of the stomatin family and a prime candidate for further study.
The subcellular localization of SLP-2 is primarily the mitochondria. This was consistent with the presence of a mitochondrial targeting sequence shown in a study using fusion proteins of green fluorescent protein (GFP) with either full length SLP-2, SLP-2 residues 1-50 (which contain the mitochondrial targeting sequence), or SLP-2 lacking residues 1-50; the two former fusion proteins were localized to the mitochondria whereas the latter was not [77]. Accordingly, numerous studies have identified SLP-2 as part of the mitochondrial proteome in several species including humans and mice [79-84]. The mitochondrial targeting sequence is cleaved following entry into the organelle, and SLP-2 becomes tightly associated with the mitochondrial inner membrane on the side of the intermembrane space [10, 77, 85]. There, SLP-2 interacts with
PHB1 and PHB2 [10, 77, 86], and binds to the mitochondrial phospholipid cardiolipin in a dose- dependent manner [86]. There is also evidence that, similar to its family members, SLP-2 is able to associate with itself either directly by homo-oligomerizing or indirectly as part of multiprotein complexes [78]. Knockdown of SLP-2 in cell lines resulted in decreased mitochondrial
29 membrane potential [10], and decreased levels of PHB1, PHB2, and certain respiratory chain complex I and IV subunits [77].
There exists at least one secondary, albeit quantitatively much smaller pool of SLP-2 at the plasma membrane that has been detected in erythrocytes [44] and T cells [76, 78]. In T cells, it was found that SLP-2 expression is upregulated following activation in humans and mice [76,
87], and the modulation of SLP-2 levels affects the quality of effector T cell responses [76].
SLP-2 was found to interact with components of the TCR signalosome during T cell activation
[76] and its levels were enhanced in the pSMAC where signaling TCR microclusters are located
[78]. Furthermore, the mitochondrial and plasma membrane pools of SLP-2 coalesce at the immunological synapse during T cell activation in a polymerized actin-dependent manner [76,
78].
More information was gathered with the generation of the T cell-specific SLP-2 knockout mouse. Deletion of SLP-2 in T cells resulted in a post-transcriptional defect in IL-2 production in response to TCR ligation and reduced CD4+ T cell responses [87]. Furthermore, SLP-2 deficiency resulted in a host of mitochondrial changes including impaired cardiolipin compartmentalization in mitochondrial membranes, decreased levels of certain complex I and II subunits, and decreased activities of complexes I and II+III [87]. These changes were in line with
SLP-2 overexpression studies in human T cells showing increases in T cell responses, mitochondrial biogenesis, cardiolipin content, respiratory complex I and II activities, and intracellular ATP levels [87] suggesting a more optimal environment for the assembly of respiratory chain components [86]. Taken together, these data indicate that SLP-2 likely facilitates functional membrane compartmentalization in mitochondrial membranes and at the plasma membrane [78]. Additional evidence supporting the notion that SLP-2 is an important
30 regulator of membrane microdomain organization came from the study of macrophages with impaired intracellular cholesterol trafficking. The expression of SLP-2 was significantly increased under these conditions and SLP-2 knockdown led to abnormal raft composition and decreased responsiveness of macrophages to multiple TLR ligands [88].
SLP-2 has been implicated in other cellular and mitochondrial processes. There is evidence that SLP-2 is involved in the pro-survival response of mitochondrial hyperfusion induced by stress in mammalian cells [10, 89]. This is corroborated by findings for the SLP-2 ortholog in C. elegans that is required for mitochondrial hyperfusion during reoxygenation following anoxia [90]. Another recent study found that the organization of mitochondrial membrane scaffolds by SLP-2 is important for anchoring a large protease complex at the mitochondrial inner membrane composed of the rhomboid protease PARL and the i-AAA protease YME1L [91]. PARL is able to regulate respiratory chain complex I activity and mitophagy [92, 93], and the mitochondrial inner membrane proteases YME1L and OMA1 balance mitochondrial fusion and fission by mediating the processing of optic atrophy 1 (OPA1), a dynamin-like GTPase [94]. Through this association, SLP-2 is able to regulate the proteolytic activities of PARL and OMA1 peptidase [91]. Lastly, SLP-2 has been identified as an oncogenic-related protein and is upregulated in a variety of cancers correlating with increased mortality [95-97]. SLP-2 has been shown to be involved in a variety of cellular processes and additional studies are required to precisely characterize its function.
Prohibitins
Prohibitin 1 (PHB1; B-cell receptor associated protein-32, BAP-32) and prohibitin 2 (PHB2;
BAP-37) are highly conserved and ubiquitously expressed members of the PHB family of
31 proteins [13]. PHBs are composed of an N-terminal alpha helix with hydrophobic regions that anchor the protein in the membrane, an evolutionarily conserved SPFH domain important for association with membrane microdomains and for protein-protein interactions, and a C-terminal coiled-coil domain also involved in protein-protein interactions, including that between PHB1 and PHB2 [13, 98, 99]. PHB1 and PHB2 are 53% homologous in amino acid sequence [100], and are dependent on co-expression for stability since knockdown of either PHB1 or PHB2 results in the loss of the other [15, 101, 102]. Furthermore, PHB1 and PHB2 knockout mice are both lethal during embryonic development [103, 104], underscoring their functional significance.
PHB1 and PHB2 are mainly expressed in mitochondria where they heterodimerize, and
12-16 of these heterodimers associate to form large ring-like structures of approximately 1 MDa at the inner membrane [99, 105]. Homodimers have not been detected [99, 102]. PHBs are associated with mitochondrial AAA (m-AAA) protease and there is evidence that the PHB complex functions as a chaperone to prevent the degradation of functional proteins [101].
Moreover, PHBs have been shown to be important for the maintenance of mitochondrial cristae structure through its involvement in the cleavage of OPA1 [18, 106]. PHBs have also been suggested to facilitate the compartmentalization of the mitochondrial inner membrane [7], and may be important for organizing cardiolipin in these membranes [7, 16].
PHBs are not exclusive to mitochondria as there have been reports of their presence in other subcellular locations including the nucleus and plasma membrane. In the nucleus PHBs regulate transcriptional activation and the cell cycle, whereas at the plasma membrane PHBs function as transmembrane adaptors that activate downstream signaling [14]. Interestingly, unlike in mitochondria heterodimerization has not been reported for the function of PHB1 and
PHB2 in these subcellular compartments [107]. Furthermore, PHBs have been studied for their
32 role in protection against apoptosis. This occurs through upregulation of phospho-ERK, which leads to increases in anti-apoptotic factors that prevent cytochrome c release and subsequent caspase 3 activation [98].
Studies are now focusing on the function of PHBs in immunity. It has been shown that the cellular expression of PHB1 and PHB2 is significantly increased following T cell activation
[86, 108], including a considerable proportion at the plasma membrane [109]. Elevated PHB levels were also measured in peripheral T lymphocytes in patients with B-lymphoproliferative diseases [110]. Furthermore, it is suggested that PHBs at the surface of activated T cells are involved in the T cell receptor-mediated signaling cascade [109]. Indeed, there is evidence that cell surface PHBs may be involved in inhibition of ERK1/2 phosphorylation following T cell receptor engagement through interaction with the lectin Siglec-9 [111]. The function of PHBs in the regulation of T cell activation requires further research and targeting these proteins may provide novel mechanistic insights into this process.
Prohibitin ligands
Several compounds have been identified to interact with and specifically bind to PHB1 and/or
PHB2 to influence their function and affect cellular processes. These compounds have been termed PHB ligands. Perhaps the most actively studied PHB ligands are the phytochemicals collectively termed flavaglines. Flavaglines are a group of naturally occurring and synthetic compounds with a cyclopenta[b]benzofuran skeleton [112]. Rocaglamide was the first of these compounds identified from the medicinal plant Aglaia elliptifolia in 1982 [113]. Since then, more than 100 flavaglines have been identified possessing various pharmacological effects including anticancer, neuroprotective, and cardioprotective properties [112, 114]. Although these
33 flavaglines were known to elicit a wide range of effects, the mechanism of action was not well defined. Recently, it was determined using affinity chromatography-coupled mass spectrometry that PHB1 and PHB2 are direct targets of flavaglines [115]. In addition to the naturally occurring flavaglines, which include the rocaglamide-related compound rocaglaol, there is significant interest in synthetically modifying these compounds to achieve more potent effects. One promising synthetic PHB ligand is the rocaglaol derivative FL3 (Figure 1.1A-B). FL3 has specifically been shown to bind PHB1 and PHB2 [115], to protect against experimental colitis
[116], and to possess anticancer [117] and cardioprotective [118] effects. In the context of immunity, it was reported that rocaglamide and certain derivatives exhibited immunosuppressive activity in T cells by inhibiting cytokine gene expression by blocking NF-AT activity [119].
However, the effect of FL3 on T cell function has not yet been tested.
Another group of PHB ligands is the fully synthetic small molecule melanogenin and derivatives (Figure 1.1C-D). Melanogenin was shown to promote the induction of pigmentation in melanocytes by directly binding PHB1, as confirmed by affinity purification [120]. This effect was achieved by melanogenin interfering with the interaction between PHB1 and a transcription factor, allowing it to proceed with induction of tyrosinase expression, the rate-limiting enzyme in melanin biosynthesis [120]. Although no further investigations on melanogenin have been reported, this compound represents an additional category of PHB ligand whose function in other contexts, including immunity, may be of interest.
The study of PHB ligands on immunity is of significant interest yet research in this context is currently limited. T cells represent an ideal model to study the effects of these compounds on adaptive immunity since PHBs have already been implicated in T cell activation.
Furthermore, it is becoming increasingly clear that cellular bioenergetics and mitochondrial
34 function are among the processes essential for T cell function, and that membrane compartmentalization, perhaps driven in part by certain proteins in the SPFH superfamily (e.g., stomatins and PHBs), may be key to these mechanisms. As such, the following sections will focus on cellular bioenergetics and mitochondrial function in the context of T cells.
T Cells
T cells are a subset of cells involved in adaptive immune responses that function to directly lyse infected cells or to activate other immune cells to clear invading pathogens. Our knowledge of basic T cell function and T cell activation has been well reviewed [121]. T cells express the T cell receptor for antigen (TCR) on the cell surface, and this receptor recognizes the unique structure of specific combinations of foreign antigenic peptide bound to a major histocompatibility complex (MHC) molecule displayed on antigen-presenting cells. The variance in the ligand-binding region of the TCR between each T cell provides a diverse T cell repertoire able to recognize and respond to innumerable foreign antigens deriving from invading pathogens. For a T cell to become activated and differentiate into an effector cell, three conditions must be met: the TCR and co-receptor (i.e., CD4 or CD8) must recognize the specific peptide:MHC complex; the co-stimulatory molecules on the surface of the antigen-presenting cell (e.g., B7.1 and B7.2) and T cell (e.g., CD28) must interact; and cytokines (e.g., IL-6, IL-12,
TGF-β) that control differentiation into the appropriate effector cell must be produced by the antigen-presenting cell to act on the T cell. CD28-dependent co-stimulation of the activated T cell will induce expression of IL-2, the cytokine that drives T cell proliferation and differentiation, and the necessary component for a high-affinity IL-2 receptor. Following
+ + activation, T cells differentiate into CD8 cytotoxic T cells, CD4 T helper cells (e.g., TH1, TH2,
35 TH17, or TFH), or T regulatory cells, and after clearance of the pathogen a small proportion further differentiate into memory T cells.
T cell bioenergetics
The bioenergetics of T cells underlies their activation, differentiation and function. Naive T cells exhibit a quiescent phenotype since they do not require the induction of metabolic pathways needed for the generation of DNA, lipids, and proteins [122]. Thus, naive T cells mainly use the available nutrients to maximize ATP production through oxidative phosphorylation (OXPHOS) and engage fatty acid oxidation [123, 124]. Glycolysis, an alternative ATP-generating bioenergetic program, is not relied upon as a major source of energy by T cells in this state [124,
125].
Naive T cells proliferate and differentiate upon antigen recognition, and these processes require the engagement of the appropriate metabolic programs. Notably, early signaling events including activation of the mechanistic target of rapamycin (mTOR) complex leads to multiple cellular processes including autophagy, the synthesis of lipids and proteins, and a significant increase in the uptake of nutrients such as glucose [122]. Glucose is imported by the transporter
Glut1 and is mainly used for glycolysis [126]. During conventional glycolysis, one molecule of glucose is catabolized into two molecules of pyruvate by a series of cytosolic enzymatic reactions that do not require oxygen, yielding two reduced nicotinamide adenine dinucleotide
(NADH) and two net ATP molecules [122, 127]. However, activated T cells mainly utilize glucose to engage in aerobic glycolysis [128], also termed the Warburg effect from observations in cancer biology [129]. Aerobic glycolysis differs from conventional glycolysis in that, despite the presence of oxygen, pyruvate is converted to lactate by the enzyme lactate dehydrogenase
36 instead of being imported into the mitochondria to enter the tricarboxylic acid (TCA) cycle, a set of reactions that generates reducing equivalents to fuel OXPHOS [130]. It has been suggested that this step is necessary for the maintenance of the cellular NAD+ pool required for subsequent rounds of glycolysis [127]. In addition to providing a means for rapid ATP generation, it is speculated that shifting the metabolic program towards aerobic glycolysis upon T cell activation increases the availability of glycolytic precursors required for biosynthetic reactions [130, 131].
There is also evidence that glycolysis is required for the effector function of activated T cells, and this is suggested to be a result of engagement of the cellular GAPDH pool in the glycolytic program leaving it unable to bind adenylate-uridylate rich elements in the 3ʹ untranslated region of IFN-γ mRNA [128]. However, glycolysis is not required for the survival or proliferation of activated T cells [128].
The substantial increase in aerobic glycolysis measured following T cell activation does not replace the requirement for energy production by OXPHOS. Indeed, OXPHOS is increased significantly in activated T cells [123, 126, 132], is sufficient to support T cell activation [132], and is essential for T cell proliferation [128, 132]. Furthermore, memory T cells have been found to possess increased mitochondrial spare respiratory capacity (i.e., the amount of a cell's maximal respiration that is not being utilized), which contributes to a rapid re-engagement of key metabolic reactions when necessary [133]. Mitochondria provide other functions required for T cell activation, namely acting as a sink for cytosolic calcium (Ca2+) upon TCR engagement [134,
135] and generating reactive oxygen species [132, 136]. The regulation of mitochondrial function in activated T cells is poorly understood and studies focused on elucidating these mechanisms will be necessary for advancements in immune therapies. The following sections
37 will focus on several key mitochondrial functions that are necessary for optimal cell function, including that of T cells.
Mitochondrial Function
The primary function of mitochondria is energy production by oxidative phosphorylation. This process is conducted by the generation of a proton gradient across the mitochondrial inner membrane that provides the proton-motive force required to drive ATP synthesis. There are five protein complexes associated with the inner membrane that are responsible for this process, collectively termed the respiratory chain: NADH:ubiquinone reductase (Complex I); succinate:coenzyme Q reductase (Complex II); coenzyme Q:cytochrome c oxidoreductase
(Complex III); cytochrome c oxidase (Complex IV); and ATP synthase (Complex V) [137-139].
Two electron carrier molecules are also involved, namely coenzyme Q (also termed ubiquinone) embedded in the lipid bilayer and cytochrome c localized on the external surface of the inner membrane [137, 140]. Together, these form an electron transport chain with Complex I being the entry point for electrons donated by NADH [138]. Ten protons are translocated to the intermembrane space per molecule of NADH by the proton-pumping Complexes I, III, and IV
[138]. Complex II is an additional entry point for electrons to enter the chain. The electron transport chain culminates with the reduction of molecular oxygen, the ultimate electron acceptor, by Complex IV to form water, and the proton-motive force generated by the proton gradient across the inner membrane drives ATP synthesis by Complex V [138, 139]. The respiratory chain is a proficient energy-producing machine capable of producing up to 36 molecules of ATP per molecule of glucose, making it the primary source of cellular ATP in many systems [139].
38
Respiratory chain supercomplexes
Recently, it has been shown that mitochondrial respiratory complexes have supramolecular interactions in the mitochondrial inner membrane to form respiratory chain supercomplexes
(RCS) [141-144]. Superassembly of the respiratory chain is a feature observed in a diverse array of species including yeast and other fungi, plants, vertebrates and invertebrates [137]. Though non-superassembled respiratory complexes are fully functional, RCS assemble to facilitate more efficient electron transfer during oxidative phosphorylation, to allow for the utilization of different electron transport pathways and substrates, to stabilize Complex I and other complexes, and to limit the production of reactive oxygen species generated from electron transport [142,
145-149]. The existence of RCS has been confirmed by electron microscopy and single particle image processing [150, 151]. RCS occur mainly between complexes I, III, and IV (termed the respirasome) with various stoichiometries (e.g., I-III2-IV1-4), but can also occur between
Complexes I and III (e.g., (I-III2)1-2) or Complexes III and IV (e.g., III2-IV) [137]. The majority of Complex I is associated with supercomplexes to maintain its stability [149, 152]. Complex III mainly forms dimers, which can be either free or associated with supercomplexes [137].
Complex V can form dimers and oligomers [144, 153], and Complex II is thought to remain fluid
[142, 145] but has been detected in Complex I-II-III-IV-containing supercomplexes [154].
Coenzyme Q and cytochrome c have also been shown to associate with RCS [154], and recent evidence indicates that there are two distinct pools of coenzyme Q that are dedicated to reducing equivalents from NADH or FADH2 [147]. The increasingly prevalent model is one in which there exists plasticity between the formation and dissociation of RCS depending on metabolic
39 demands and other factors, as opposed to the fluid or solid models in which respiratory chain complexes exhibit no supramolecular interactions or are statically superassembled, respectively.
The dynamic nature of RCS organization and dissociation has been observed in a variety of conditions and disease states. Starvation of mice reduced the amount of I-III2-IV RCS in liver mitochondria [147] and several studies have shown that RCS distribution can change with ageing: Complex I-containing RCS were decreased in aged rat heart mitochondria [155], and a similar effect was observed with ageing rat brain mitochondria, particularly involving I-III2 RCS
[156]. Furthermore, RCS formation was decreased in pathological conditions including heart failure [157] and Barth syndrome [158]. Interestingly, the formation of RCS is not necessary for bioenergetic function. Non-superassembled respiratory chain complexes are able to respire and certain common laboratory mouse strains (i.e., C57Bl/6J and Balb/cJ) that harbour a mutation in
SCAF1, rendering them unable to form Complex III-IV-containing RCS, have no serious bioenergetic deficiencies [147, 159]. However, the evidence indicates that RCS formation is an important feature of the respiratory chain and more research is required to establish the mechanisms governing RCS formation and function.
The molecular machinery involved in the formation, maintenance, and regulation of RCS is not well characterized. However, it is clear that the composition of the mitochondrial inner membrane is a vital factor as exemplified by the phospholipid cardiolipin. Cardiolipin is mainly localized to mitochondrial membranes [160, 161] and is unique among phospholipids in that it has a dimeric structure composed of diphosphatidylglycerol resulting in a specific conical structure [162]. Cardiolipin is essential for the organization of functional protein-protein and protein-lipid microdomains where superassembly of respiratory complexes occurs [137, 163].
Barth syndrome, a human genetic disease characterized by cardiomyopathy, skeletal myopathy,
40 and neutropenia, is caused by a mutation in the tafazzin gene that impairs cardiolipin remodeling and destabilizes RCS [158], a feature of cardiolipin present in other systems [164, 165].
Furthermore, cardiolipin has been shown to physically bind to Complexes I, III, IV, and V [166-
168], and to be required for the activities of these complexes [169-171]. It has also been suggested that cardiolipin may fill the spaces between complexes organized into RCS [140].
Thus, cardiolipin is an important factor for proper RCS formation and function.
Several proteins have also been identified to be important for RCS formation. Two related Saccharomyces cerevisiae proteins, Rcf-1 and Rcf-2, and a mammalian homolog hypoxia-induced gene (HIG)2A, are necessary for assembly of mature Complex IV and affect
Complex III-IV RCS formation [172-174]. Depletion of C11orf83, a protein that binds cardiolipin and phosphatidic acid and is required for Complex III assembly and stabilization, resulted in decreased amounts of III-IV- but not I-III-IV-containing supercomplexes [175].
Lastly, supercomplex assembly factor I (SCAFI) has been shown to be required for the superassembly of Complexes III and IV, but does not affect the individual assembly or function of either complex [147, 159]. The identification of additional factors required for the assembly and maintenance of RCS will be necessary to define the mechanisms governing this process and may be important for the modulation of mitochondrial function.
Mitochondrial translation
The mitochondrial proteome is encoded almost entirely in the nucleus, but these organelles maintain their own set of DNA and possess the requisite machinery to transcribe and translate mitochondria-encoded genes. Mitochondrial (mt)DNA is a maternally inherited, circular, double- stranded molecule [176] consisting of 16,569 base pairs in humans [177] and 16,295 base pairs
41 in mice [178]. It is polyplasmic, with each mitochondrion typically containing many copies, and is continuously replicating independent of the cell cycle and also in non-dividing cells [179,
180]. Genes are located on both the heavy (H) and light (L) strands and the mitochondrial genome lacks introns and has only one major non-coding region, termed the displacement (D)- loop [177, 178]. MtDNA is organized into nucleoids, protein-DNA complexes containing proteins involved in mtDNA maintenance, replication, and transcription [180]. In total, mtDNA encodes 37 genes of which 13 code for essential polypeptide subunits of respiratory chain complexes I, III, IV, and V, while the remaining two rRNA and 22 tRNA genes are necessary for mitochondrial translation [181, 182]. Furthermore, mtDNA has an approximate 10-fold greater mutation rate than nuclear DNA likely due to factors including limited DNA repair, lack of histones, and proximity to reactive oxygen species [183, 184].
Transcription of mtDNA originates from three promoters, two on the H strand and one on the L strand, to generate polycistronic molecules [180]. These units are further processed such that the tRNAs are excised and the remaining rRNAs and mRNAs are polyadenylated [180], resulting in the generation of nine monocistronic and two dicistronic mRNAs that can be translated by the mitoribosome [177]. These mitochondrial mRNAs differ from their nuclear counterparts since they contain few 5ʹ untranslated nucleotides [185]. Furthermore, they differ from both prokaryotic and nuclear mRNAs in that they are uncapped and contain a poly(A) tail that immediately follows or is part of the stop codon [177]. Upon completion of mRNA processing, mitochondrial translation is initiated and becomes coordinated with the expression, import, and membrane integration of nuclear-encoded mitochondrial proteins, essential processes for the assembly of respiratory chain complexes.
42 The mitochondrial proteome contains an estimated 1,200 proteins of which most originate in the nucleus [180, 186]. These nuclear-encoded mitochondrial proteins are synthesized on cytosolic ribosomes and translocated as pre-proteins to the mitochondria [187].
The import of these pre-proteins occurs either post-translationally or in a co-translational manner with cytosolic ribosomes bound to the cytosolic side of the outer mitochondrial membrane [188-
190]. Cytosolic chaperones keep the pre-proteins unfolded to prevent aggregation and enable entrance into the translocation channels located in both the outer and inner mitochondrial membranes [191]. These include translocase of the outer mitochondrial membrane (TOM complex) and translocase of the inner mitochondrial membrane (TIM)23 complex [190, 192].
Pre-proteins will follow one of several routes to reach their destination based on their targeting signals. Many involved in mitochondrial translation or respiratory chain complex assembly are directed to their destination by processes involving the TIM23 complex. This complex can mediate translocation of pre-proteins to the inter membrane space, pre-proteins can enter the matrix through the TIM23 channel, and inner membrane pre-proteins are either inserted directly or first transported to the matrix before membrane integration in processes involving TIM23
[190]. Once the imported pre-proteins reach their destination, targeting sequences are proteolytically cleaved and the proteins can fold into their mature forms [190, 192].
The mitochondrial translation system differs substantially from its eukaryotic cytosolic counterpart, but is instead more similar to the prokaryotic translation system [182]. Human mitochondrial ribosomes (mitoribosomes) are composed of two rRNAs (12S and 16S) and approximately 81 mitochondrial ribosomal proteins (MRPs) that are organized into a 28S small subunit (SSU) and a 39S large subunit (LSU) [182]. The functional monosome is comprised of the SSU and LSU and sediments as a 55S particle [193]. Its density is lower than the 80S or 70S
43 monosomes of eukaryotic or bacterial cells, respectively [193], since mammalian mitoribosomes have a substantially greater protein-to-RNA ratio as a result of less rRNA domains and increased protein mass [194]. In addition to rRNA and MRPs, the basic mitochondrial translation machinery is comprised of tRNAs, initiation, elongation, and termination factors, mitochondrial aminoacyl-tRNA synthetases, and methionyl-tRNA transformylase [182, 194]. Furthermore, the functional core of the mitoribosome has been evolutionarily conserved and thus contains the tRNA aminoacyl (A)-, peptidyl (P)- and exit (E)-sites [195]. Mitochondrial protein synthesis occurs in three phases: initiation, elongation, and termination, and the details of these phases are described in several reviews [196, 197].
Mitochondrial translation occurs in close proximity to the mitochondrial inner membrane
[198, 199]. Indeed, biochemical studies have shown that nearly half of mammalian mitoribosomes interact with the mitochondrial inner membrane and it is suggested that this association is mediated in part by mitoribosome interaction with integral or membrane-bound proteins [200]. Few such proteins have been identified, but these may include OXA1L, a polytopic membrane insertase of the mitochondrial inner membrane [201] that has been cross- linked to LSU components [202], and LetM1, a mitochondrial inner membrane protein with a large matrix domain interacts with the mitoribosomal protein MRPL36 [203]. Furthermore, it is possible that only membrane-bound mitoribosomes are translationally active [204]. This idea is supported by a study showing that yeast mitoribosomes become anchored to the inner membrane during translation [205]. This close association with the membrane may be functionally advantageous since it is thought that the mitochondrial gene products are inserted into the inner membrane as they are being synthesized on the mitoribosome [182]. This co-translational activity may be necessary since all 13 mitochondrially translated proteins are extremely
44 hydrophobic core proteins of the respiratory chain and must be inserted into the membrane to prevent aggregation and precipitation in the matrix [181, 188, 189].
It is hypothesized that mitoribosome assembly occurs in two mitochondrial subcompartments: initiated in the nucleoids, sites of mtDNA maintenance, replication, and transcription, and completed in the RNA granules, where post-transcriptional RNA processing and maturation occur [206, 207]. These compartments contain mitoribosome assembly factors,
MRPs, and proteins involved in RNA stability [206, 208]. It has been reported that several
GTPases may be involved in mitoribosome assembly, including Mtg1 and Mtg2, mitochondrial inner membrane-associated proteins that interact with the LSU [209], and C4orf14, which is implicated in SSU assembly [210]. Additionally, the helicase DDX28 and the Fas-activated serine threonine kinase family member FASTKD2 have been shown to be required for LSU assembly [208, 211, 212], whereas C7orf30 mediates LSU stability [213, 214]. The identification of other factors mediating mitoribosome assembly or stability will help to clarify the mechanisms involved in this process.
Rationale and Specific Aims
Initial studies in our laboratory were focused on identifying novel players in the regulation of T cell activation through the antigen receptor. This was initiated by performing a proteome analysis of DI-GEMs of activated T cells. In addition to proteins known to be involved in T cell activation, a novel protein, SLP-2, was identified whose levels were enhanced in this cellular compartment [76]. Members of the stomatin family of proteins, to which SLP-2 belongs, have been implicated in the formation and organization of membrane microdomains [48, 75].
Subsequent work showed that SLP-2, a mainly mitochondrial protein that is widely expressed,
45 had at least one other subcellular pool at the plasma membrane and modulated T cell activation
[76, 78, 86]. To further characterize the function of SLP-2 in T cell responses, our group generated T cell-specific SLP-2 knockout (SLP-2 T-KO) mice. Using these mice, it was found that CD4+ T cell responses and the activities of certain mitochondrial respiratory chain complexes were impaired [87]. Mechanistic studies showed that SLP-2 binds the mitochondrial phospholipid cardiolipin in a dose-dependent manner [86]. Furthermore, the data suggested that
SLP-2 recruits PHBs to cardiolipin in the mitochondrial inner membrane to form functional membrane microdomains where the respiratory complexes associate [87]. A model of mitochondrial SLP-2 function in the context of T cell activation is depicted in Figure 1.3. Since
SLP-2 is involved in the formation of specialized cardiolipin-enriched membrane microdomains in the mitochondrial inner membrane, we hypothesized that SLP-2 would be required for optimal respiratory chain formation and mitochondrial function.
It is known that cardiolipin is essential for the function of the respiratory chain [137,
163], and that respiratory chain complexes naturally associate to form supercomplexes for optimal electron transfer and function [137, 140]. Based on our model of SLP-2 function, my first specific aim is to characterize the role of SLP-2 in the organization and function of mitochondrial respiratory chain complexes and supercomplexes. To do this, T cells from
SLP-2 T-KO mice will be examined for bioenergetic performance, formation of respiratory chain complexes and supercomplexes, and T cell function.
Given the importance of SLP-2 in the formation of specialized membrane microdomains in the mitochondrial inner membrane, my second specific aim is to determine whether SLP-2 is required for proper mitochondrial translation, a critical cellular process that occurs in close association to the mitochondrial inner membrane [198-200]. To test this, the level of de novo
46 synthesis of mitochondria-encoded peptides and the extent of mitoribosome assembly will be measured in SLP-2-deficient T cells.
Finally, there are extra-mitochondrial pools of SLP-2 and PHBs and it is known that
PHBs are involved in a host of cellular processes [14, 98]. Furthermore, naturally occurring and synthetic compounds exist that bind PHBs and can have various beneficial effects on cell function [114]. However, it is not known what role the association of SLP-2 with PHBs plays in these processes. My third specific aim is to determine the effect of the synthetic PHB ligand derivatives FL3 and Mel6 on T cell function and to assess whether SLP-2 is involved in the mechanism(s) underlying these effects. To do this, I will test the effect of FL3 and Mel6 on T cell activation in control or SLP-2-deficient T cells, and determine whether the binding of these compounds to PHBs affects SLP-2:PHB interaction.
Taken together, this work will be important for elucidating the function of SLP-2 in the regulation of key cellular processes occurring in close association with cellular membranes and membrane proteins.
47 Figure Legends
Figure 1.1. SLP-2 secondary structure. SLP-2 is a 37 kDa protein that contains an N-terminal mitochondrial targeting sequence, several putative myristoylation sites, a highly conserved SPFH domain common to members of its protein superfamily, and an α-helix-rich domain.
Figure 1.2. Structures of PHB ligands and derivatives. (A) Rocaglaol, (B) FL3, (C)
Melanogenin, (D) Mel6. Adapted from [117] and [120].
Figure 1.3. Model of SLP-2 function in the context of T cell activation. Side view and top view of the mitochondrial inner membrane depicting SLP-2 binding to cardiolipin (CL) and interacting with PHB1 and PHB2 in regions where the respiratory complexes localize. During T cell activation, SLP-2 protein levels are increased in these regions, leading to the recruitment of additional CL and PHBs to form microdomains for optimal respiratory chain function. Complex
I (CI; NADH dehydrogenase); Complex II (CII; succinate dehydrogenase); Complex III (CIII; bc1 complex); Complex IV (CIV; cytochrome c oxidase); Complex V (CV; ATP synthase).
Figure is adapted from Christie, D. 2012 [215].
48 Figure 1.1
49 Figure 1.2
50 Figure 1.3
51 Preface to Chapter 2
Membrane microdomains are very dynamic and heterogeneous membrane structures, and their composition and function are widely varied depending on their subcellular location. Originally, we identified SLP-2 in a proteome analysis of detergent-insoluble microdomains from activated
T cells stimulated through the antigen receptor. This chapter focuses on the methodology of membrane microdomain isolation, specifically of detergent-insoluble glycosphingolipid microdomains, plasma membranes, and cardiolipin-enriched mitochondrial membrane microdomains. Moreover, the process of mitochondrial supercomplex identification by blue- native gel electrophoresis is detailed, which will be an important concept in the subsequent chapter.
52 Chapter 2: Identification of Multimolecular Complexes and Supercomplexes in Compartment-Selective Membrane Microdomains
Panagiotis Mitsopoulos and Joaquín Madrenas
From
The Department of Microbiology and Immunology, McGill University, Montreal, QC, Canada
H3A 2B4
This chapter contains the following published manuscript:
Mitsopoulos P, Madrenas J. Identification of multimolecular complexes and supercomplexes in compartment-selective membrane microdomains. Methods Cell Biol. 2013; 117: 411-431.
Copyright © 2013. Elsevier Inc.
53 Abstract
Cellular membranes contain specialized microdomains that play important roles in a wide range of cellular processes. These microdomains can be found in the plasma membrane and other membranes within the cell. Initially labeled lipid rafts and defined as being resistant to extraction by nonionic detergents and enriched in cholesterol and glycosphingolipids, we now understand that these membrane microdomains are very dynamic and heterogeneous membrane structures whose composition and function can vary widely depending on their cellular location. Indeed, though they are classically associated with the plasma membrane and have been shown to facilitate a wide variety of processes, including signal transduction and membrane trafficking, specialized membrane microdomains have also been identified in other membranes including those in the mitochondria. These mitochondrial membrane microdomains are enriched in cardiolipin, the signature phospholipid of the mitochondria, and may have important implications in metabolism by facilitating optimal assembly and function of the mitochondrial respiratory chain. Furthermore, isolation of multimolecular complexes while retaining their supramolecular interactions has been critical to the study of mitochondrial respiratory supercomplexes. Here, we discuss methods to isolate various membrane microdomains, including detergent-insoluble glycosphingolipid microdomains, mitochondrial cardiolipin-enriched microdomains, and blue- native gel electrophoresis of mitochondrial membranes.
54 Introduction and rationale
The fluid mosaic model of biological membranes by Singer and Nicolson (1972) described the membrane as a primarily lipid matrix with proteins distributed randomly throughout [216].
However, shortly thereafter it was proposed that membranes contain “clusters of lipids”, these clusters being “quasicrystalline” regions surrounded by more freely dispersed liquid crystalline lipid molecules. This concept of membrane lipid microdomains was refined to claim that these clusters would contain “lipids in a more ordered state”. More importantly, the organization of lipids into membrane microdomains provided an additional conceptual framework for membrane heterogeneity across cellular subcompartments and thus the possibility of structural and functional significance for these microdomains [217].
Since its original proposal, the concept of membrane microdomains has been better characterized at the molecular level. Under the term of lipid rafts, membrane microdomains were described as being resistant to extraction by nonionic detergents and enriched in cholesterol and glycosphingolipids [218]. The term detergent-insoluble microdomains is more encompassing of its heterogeneity in composition than lipid rafts, as some of these microdomains may be selectively enriched for other phosphiolipids. For simplicity, we will be using both indistinguishably in this paper. Lipid raft structures were thought to be stably held together by lipid-lipid interactions [218]. It has also become clear that lipid rafts are heterogeneous, dynamic sterol- and sphingolipid-enriched nanoscale microdomains that compartmentalize cellular processes [218], including membrane trafficking, signal transduction, and cell polarization [219].
These microdomains are ordered assemblies of specific proteins in which the meta-stable resting state can be activated to coalesce by specific lipid-lipid, lipid-protein, and protein-protein interactions [220-222].
55 The compartmentalization of the plasma membrane into detergent insoluble microdomains is a very dynamic process. For example, in T cells, the formation of an immunological synapse correlates with clustering of glycolipid-enriched microdomains. These microdomains are enriched with signaling molecules critical for the initiation and sustainability of T cell antigen receptor mediated signaling such as lck, linker for activation of T cells (LAT), and protein kinase C theta, and their biological integrity is essential for proper T cell antigen receptor mediated signaling [223-226]. Lipid rafts move into the immunological synapse in response to CD3 and CD28 ligation, and formation of these microdomains is important for normal signaling leading to T cell activation. Lipid rafts also contain regulators of signaling such as CTLA-4, a negative regulator of T cell activation that partitions within lipid rafts and relocates to the immunological synapse [227].
In addition to the plasma membrane, detergent-insoluble microdomains have been identified in other cellular membranes including mitochondrial membranes [87, 228, 229]. In mitochondria, cardiolipin is the major phospholipid and evidence suggests that there exist microdomains selectively enriched in cardiolipin [230]. Cardiolipin is a phospholipid that has been shown to be important for optimal function of the mitochondrial respiratory chain. Thus, it is possible that the respiratory chain operates within cardiolipin-enriched microdomains in the mitochondrial inner membrane. Isolation of these microdomains will be important for the biochemical characterization of respiratory chain complexes and supercomplexes, and to determine their biological relevance.
In this paper, we will concentrate on the characterization of multimolecular complexes and supercomplexes in detergent-insoluble microdomains from mitochondrial membranes, specifically cardiolipin-enriched microdomains. These molecular structures are a focus of
56 intense study, specifically understanding how they are assembled and how they ensure optimal cellular respiration. There is an increasing amount of experimental evidence suggesting that mitochondrial respiratory complexes are not simply randomly dispersed within the mitochondrial inner membrane but rather undergo ordered supramolecular associations. These mitochondrial respiratory supercomplexes were first identified by Schagger and von Jagow when they performed blue-native gel electrophoresis on digitonin-solubilized yeast and mammalian mitochondria [143, 144]. Since then, supercomplexes have been found in many mammalian tissues, as well as other organisms including fish, fungi, and bacteria [142]. These supercomplexes mainly consist of the standard respiratory complexes I, III, and IV in various stoichiometries (e.g., I-III2-IV1-4), though there may be some cases in which complex II is involved [154]. Arrangement of respiratory complexes into supercomplexes has been suggested to aid in substrate channeling and may be important for optimal activity of the respiratory chain.
Furthermore, there is evidence that cardiolipin may be necessary for supercomplex assembly
[165]. The reader is referred to several excellent reviews on mitochondrial respiratory supercomplexes for further details [140, 142].
Here, we will describe methodology for isolating various membrane microdomains, including detergent-insoluble glycosphingolipid microdomains from plasma membranes for biochemical characterization, and methods for isolating cardiolipin-enriched mitochondrial membranes and native protein complexes including the mitochondrial respiratory chain supercomplexes. These techniques will provide the tools necessary to effectively study multimolecular complexes and oligomeric structures in their intrinsic membrane location and explore the biological functions associated with these specialized membrane microdomains.
57 Detergent-insoluble glycosphingolipid (DIG) microdomain isolation
The following protocol is designed for the isolation of detergent-insoluble glycosphingolipid microdomains from cellular membranes, in particular T lymphocytes. This protocol is modified from methods described previously [224, 231-234].
A) Materials i) Reagents
Mes buffer, pH 6.5
NaCl
Triton X-100
EDTA
Na3VO4 (prepared fresh)
PMSF (phenylmethyl sulfonyl fluoride)
Aprotinin
Sucrose
Distilled deionized water (ddH2O)
ii) Equipment
Multi-Purpose Rotator (Scientific Industries Inc.)
Dounce homogenizer with loose-fitting plunger (Wheaton)
Pasteur pipettes
Ultracentrifuge (e.g., Optima XL-90 with SW40 rotor, Beckman)
58 iii) Buffers and Sucrose
• DIG Lysis buffer: (25 mM Mes pH 6.5, 150 mM NaCl, 0.5% Triton X-100, 1 mM EDTA, 1
mM fresh Na3VO4, 1 mM PMSF, 10 µg/mL aprotinin in ice-cold ddH2O)
For 30 mL:
750 µL of 1 M Mes Buffer, pH 6.5
1500 µL of 3 M NaCl
1500 µL of 10% Triton X-100
60 µL of 0.5 M EDTA, pH 8.0
300 µL of fresh 0.1 M Na3VO4
300 µL of 100 mM PMSF
30 µL of 10 mg/mL Aprotinin
25.56 mL of ddH2O
For 50 mL:
1250 µL of 1 M Mes Buffer, pH 6.5
2500 µL of 3 M NaCl
2500 µL of 10% Triton X-100
100 µL of 0.5 M EDTA, pH 8.0
500 µL of fresh 0.1 M Na3VO4
500 µL of 100 mM PMSF
50 µL of 10 mg/mL Aprotinin
42.6 mL of ddH2O
• MBS buffer: (25 mM Mes pH 6.5, 150 mM NaCl)
59 For 300 mL:
7.5 mL of 1 M Mes Buffer, pH 6.5
15 mL of 3 M NaCl
277.5 mL of ddH2O
• 85% (w/v) sucrose: (64 g sucrose + 40 mL MBS buffer)
• 35% (w/v) sucrose: Dilute 85% sucrose in MBS with 1 mM EDTA and 1 mM Na3VO4
4 samples (21 mL)
8.65 mL of 85% sucrose
210 µL of 0.1 M Na3VO4
42 µL of 0.5 M EDTA
12.098 mL of MBS
6 samples (31 mL)
12.77 mL of 85% sucrose
310 µL of 0.1 M Na3VO4
62 µL of 0.5 M EDTA
17.858 mL of MBS
• 5% (w/v) sucrose: Dilute 85% sucrose in MBS with 1 mM EDTA and 1 mM Na3VO4
60
4 samples (21 mL)
1.24 mL of 85% sucrose
210 µL of 0.1 M Na3VO4
42 µL of 0.5 M EDTA
19.508 mL of MBS
6 samples (31 mL)
1.83 mL of 85% sucrose
310 µL of 0.1 M Na3VO4
62 µL of 0.5 M EDTA
28.798 mL of MBS
Note: Mix sucrose mixtures at 4°C, rotating for 2 min.
• 2× Sample buffer
o Dilute 4× sample buffer (refer to "Plasma membrane isolation" section) with DIG
lysis buffer (1:1)
§ e.g., 100 µL of 4× sample buffer + 100 µL of DIG lysis buffer
B) Method i) Isolation of DIG fractions
• Prepare 40×106 to 100×106 cells per group. For fewer cells, use less lysis buffer to keep the
cell concentration high.
• Stimulate cells as appropriate.
• The following steps must be performed on ice:
o Prepare the buffers and sucrose mixtures (see Materials: Buffers and Sucrose).
61
o Gently resuspend cell pellet in DIG lysis buffer. Lyse for 30 min.
o Homogenize each lysate in a pre-chilled Dounce homogenizer (Wheaton; 7 mL glass
tube with glass plunger) by applying 10 gentle strokes with the loose-fitting plunger.
o Mix homogenized lysate with an equal volume of 85% (w/v) sucrose, then rotate at
4°C for 2 min.
o Place this mixture at the bottom of a 14 × 95 mm Ultra-Clear ultracentrifuge tube
(Beckman). Slowly overlay this mixture with 5 mL of 35% sucrose mixture. Next,
carefully overlay the 35% layer with 5 mL of 5% sucrose mixture. The overlays can
be made in a variety of ways (e.g., Pasteur pipette, pipette-aid set to “slow”, or a
gradient-former apparatus), provided it is performed carefully and consistently.
o Centrifuge at 200,000 × g for 16 – 18 h at 4°C in a pre-chilled SW40 rotor.
o Place tubes on ice, discard the upper 3 – 4 mL, then extract the opaque band and the
35/5% interface (1 mL total), and label “DIG”. Discard the next 5 mL, then extract
the bottom 2 mL from the tube and label “Soluble”. Alternatively the entire tube may
be divided into 12 × 1 mL fractions. The extraction can be performed with a 1 mL
pipetman or by making a side puncture with a butterfly needle (this requires use of
polyallomer soft tubes).
• Aliquot and add 4× sample buffer (at a 1:4 ratio), boil at 95°C for 5 min, and western blot
directly, or immunoprecipitate the fractions and then western blot. Alternatively, DIGs may
be stored at -70°C. DIGs (but not the soluble fraction) can be pelleted by diluting the fraction
with an equal volume of MBS and then centrifuging at 20,000 × g for 10 min at 4°C.
Additionally, protein can be precipitated and then assayed.
62
ii) Immunoprecipitation of DIG Fractions
• Pre-coat protein A/G beads with 1 µg antibody in 500 µL PBS for 2 h at 4°C.
• Split the 1 mL DIG fractions into two portions of 500 µL in microfuge tubes.
• Add 500 µL MBS to each tube, then vortex.
• Spin at 20,000 × g for 10 – 30 min at 4°C.
• Remove and discard 950 µL of the supernatant from each tube with a pipette without
disturbing the pellet.
• Add 900 µL of DIG lysis buffer to one tube, vortex, then transfer mixture to the other tube
and vortex.
• Centrifuge the beads coated with antibody (i.e., maximum speed for 3 seconds). Discard the
supernatant and wash the antibody-coated beads once with 500 µL of DIG lysis buffer.
Centrifuge again and discard the supernatant.
• Add the DIG fraction to the antibody-coated beads, then rotate gently at 4°C for 2 h or
overnight.
• Wash three times with 1 mL of DIG lysis buffer, centrifuging at maximum speed for 3
seconds each time.
• Add 2× sample buffer, boil at 95°C for 5 minutes, then proceed with western blotting.
• Note: Soluble fractions should be immunoprecipitated directly.
63
Plasma membrane isolation
This protocol is designed to obtain highly purified plasma membranes for biochemical studies, and also produces an internal membrane fraction. This protocol has been adapted from Chaney and Jacobson 1983 [235] and Cells: A Laboratory Manual [236].
A) Materials i) Reagents
Mes buffer
NaCl
Sorbitol
Polymethacrylic acid (PAA)
Imidazole
Aprotinin
Leupeptin
PMSF
Na3VO4
Ludox
Histodenz
Sodium dodecyl sulfate (SDS)
β-Mercaptoethanol
Tris
Glycerol
Bromophenol blue
64
ddH2O
ii) Buffers and Solutions
• PMCB (plasma membrane coating buffer)
o 20 mM MES
o 150 mM NaCl
o 280 mM sorbitol (182.19 g/mol)
o pH range: 5.0 – 5.5
• PMCB pH 6-7
o Before adjusting the pH of the PMCB above, take a 50 mL aliquot (it should be
pH 7).
• PAA polymethacrylic acid (require 5 mL per sample)
o 1 mg/mL PAA in PMCB (pH 6-7)
§ First prepare 50 mg/mL stock in water (dissolves slowly), then dilute
accordingly.
• Lysis buffer (prepare 50 mL for 2 samples)
o 2.5 mM imidazole, pH 7.0
o 10 µg/mL aprotinin
o 10 µg/mL leupeptin
o PMSF and Na3VO4 can be added as well
• Ludox
o Make 5% v/v mixture of ludox in PMCB (5 mL per sample).
• Histodenz (Nicodenz)
65
o Measure 10 g powder, add 5.5 mL lysis buffer.
o Vortex repeatedly, takes approximately 30 – 60 min to dissolve.
o Take 7 mL of this and top up to 10 mL with lysis buffer.
o This solution can be frozen as a stock, but ensure it completely dissolves upon
thawing.
• 4× Sample buffer (prepare 50 mL, can be stored at room temperature)
o 8% SDS
o 8% β-Mercaptoethanol
o 250 mM Tris, pH 6.8
o 40% glycerol (v/v)
o 2% Bromophenol blue
o ddH2O
• PBS (phosphate-buffered saline), pH 7.4
o 8.0 g of NaCl
o 0.2 g of KCl
o 1.44 g of Na2HPO4
o 0.24 g of KH2PO4
o up to 1L ddH2O
iii) Equipment
• Optima XL-90 ultracentrifuge with SW40 rotor (Beckman), or similar
66
o This protocol is designed for an 11 mL tube. Depending on the rotor used,
calculate the volume of lysate and other Histodenz layers accordingly so that they
fit in the tube. Smaller rotors (e.g., SW60, Beckman) may also be used.
B) Method i) Silica coating
The purpose of this method is to encapsulate the cells in ludox, crosslink with PAA to form a cast around the membrane, and then lyse the cells using a Dounce homogenizer.
• Count the cells of interest (use between 5 – 50 × 106 cells).
• Wash cells with PBS.
• Resuspend cells in 1 mL of PMCB.
• Add 5 mL of 5% ludox in a 50 mL conical tube.
• Aspirate the cells into a large syringe.
• Attach a 20 gauge needle, then add cells from syringe one drop at a time to the ludox
solution while manually swirling the tube.
• Add PMCB to a final volume of 20 mL.
• Centrifuge at 900 × g for 3 min, then aspirate the supernatant. Appearance of a “fluffy”
pellet generally means that the cells are already being lysed or dead. This is not desired.
• Wash twice with 20 mL PMCB.
• Resuspend in 1 mL of PMCB.
• Add the silica-coated cells drop-wise as described previously to 5 mL of the 1 mg/mL
PAA solution in a fresh 50 mL tube while swirling manually.
• Dilute by topping up to 20 mL PMCB.
67
• Centrifuge at 900 × g for 3 min, then discard the supernatant.
• Resuspend in 2 mL of cold lysis buffer, put on ice for 30 min and agitate occasionally.
• Lyse cells in pre-chilled Dounce homogenizer using the loose-fit pestle (ten strokes) and
then the tight-fit pestle (three strokes).
• Check that cells have lysed using a light microscope.
ii) Fractionation
Now the plasma membrane is very dense and can be isolated from internal membranes and nuclear debris.
• Centrifuge silica membranes (900 × g for 3 min).
• Remove fluid phase and transfer it to microfuge tubes. This contains internal membranes
and the cytosolic fraction.
o Spin the fluid phase at 20,000 × g for 10 min at 4°C.
o The pellet is internal membranes and the new fluid phase is cytosol.
o Add a small amount of lysis buffer to the internal membranes, plus 4× sample
buffer, then boil and use for western blotting.
o Take the cytosol phase and add 4× sample buffer, boil and use for Western
blotting.
• Resuspend the silica membranes in 11 mL of lysis buffer.
• Layer this onto 2 mL of 70% Histodenz in the ultracentrifuge tube.
• Place in an ultracentrifuge rotor (i.e., SW40, Beckman) and spin at 28,000 × g for 30 min
in an Optima XL-90 ultracentrifuge (Beckman) or similar.
• Carefully remove and discard all fluid phase.
68
• Resuspend the pellet in 1 mL of lysis buffer.
• Transfer into microfuge tube, keep on ice.
• Wash three times with 1 mL of lysis buffer (spin in table-top microcentrifuge at
maximum speed (e.g., 20,000 × g) for 5 seconds).
• The pellet contains purified plasma membrane.
• To solubilize proteins for western blotting:
o Add small amount of lysis buffer (i.e., 500 µL) containing 1.0% SDS.
o Sonicate (several short bursts, keep on ice as much as possible).
o Boil for 5 min.
o Pellet on table-top centrifuge at 20,000 × g for 10 min at 4°C.
o The supernatant now contains solubilized proteins from the plasma membranes.
Transfer the supernatant to a new tube, add 4× sample buffer, then boil and use
for western blotting.
o Discard the silica pellet.
Cardiolipin-enriched mitochondrial membrane microdomain isolation
The following protocol is designed to isolate cardiolipin-enriched microdomains from mitochondrial membranes. This protocol has been adapted from the method introduced by Ciarlo et al. [229].
A) Materials i) Reagents
69
HEPES
NaCl
Triton X-100
Aprotinin
Tris-HCl, pH 8.8
SDS
EDTA
Tris, pH 7.5
Sucrose ddH2O
ii) Buffers
• Isolation buffer
o 250 mM sucrose
o 10 mM Tris, pH 7.5
o 1 mM EDTA
• Extraction buffer
o 25 mM HEPES, pH 7.5
o 150 mM NaCl
o 1% Triton X-100
o 10 µg/mL aprotinin
• Solubilization buffer
o 50 mM Tris-HCl, pH 8.8
70
o 1% SDS
o 5 mM EDTA
B) Method i) Mitochondria Isolation
• Harvest cells of interest and isolate intact mitochondria from the cells. The Qproteome
Mitochondria Isolation kit (Qiagen) is recommended. Perform all steps on ice.
• Alternatively, mitochondria can be isolated from cells by differential centrifugation
following homogenization using a Dounce homogenizer. Here, we outline mitochondria
isolation from primary activated T cells. This protocol has been adapted from Ogilvie et
al. [237] and may vary depending on cell type.
o All steps must be performed on ice or at 4°C.
o Harvest cells of interest and wash twice with ice-cold PBS (refer to "Plasma
membrane isolation" section), centrifuging each time at 380 × g for 5 min at 4°C.
o Resuspend cell pellet in 5 mL of ice-cold isolation buffer and transfer to a pre-
chilled Dounce homogenizer.
o Homogenize cells on ice by applying 20 strokes using the tight-fit pestle of the
Dounce homogenizer. The number of strokes should be optimized for each cell
type.
o Transfer homogenate into 4 microfuge tubes and centrifuge twice at 600 × g for
10 min at 4°C.
o Collect the supernatants and transfer to new microfuge tubes.
71
o Centrifuge at 10,000 × g for 20 min at 4°C. The pellet contains mitochondria. The
supernatant (i.e., cytosol) can be collected for quality control analysis (e.g.,
measurement of cytochrome C protein levels).
ii) Cardiolipin-enriched Microdomain Isolation
• Lyse mitochondrial pellet by resuspending in 1 mL of extraction buffer, then incubate for
20 minutes on ice.
• Collect lysates and centrifuge at 20,000 × g in a table-top microcentrifuge for 2 min at
4°C.
• Collect the supernatants containing Triton X-100-soluble material. Centrifuge the pellets
a second time (20,000 × g for 30 sec at 4°C) to remove the remaining soluble material.
• Solubilize pellets in 100 µL solubilization buffer.
• Shear DNA by passage through a 22-gauge needle.
• Analyze both Triton X-100-soluble and -insoluble material by western blot. Load the
fraction samples by volume.
Mitochondrial supercomplex identification by blue-native gel electrophoresis
Blue-native gel electrophoresis was first applied to digitonin-solubilized mitochondrial membranes by Schagger et al. [144] leading to the identification of mitochondrial respiratory supercomplexes. Blue-native refers to the fact that the separated protein complexes are not denatured but rather retain enzymatic activity while their electrophoretic separation relies on binding of the dye Serva Blue G to the protein. Treatment of samples with the mild detergent digitonin allows for many protein-protein interactions to remain intact, allowing for analysis of
72
supramolecular associations of the respiratory complexes and other proteins. Blue-native samples are run on a first-dimension non-denaturing polyacrylamide gel to separate protein complexes based on size. Analysis of the components of each multi-protein complex from the first-dimension gel can be achieved by cutting the appropriate gel slice and running it on a denaturing second-dimension polyacrylamide gel followed by western blotting. Silver staining may also be performed on first- or second-dimension gels to visualize multiprotein complexes or their components, respectively. This technique can be used to gather a considerable amount of information on multiprotein complexes and mitochondrial respiratory supercomplexes in particular. For this reason, blue-native gel electrophoresis is now one of the most commonly used techniques for studying mitochondrial respiratory supercomplexes. The following protocol for the isolation of mitochondrial respiratory supercomplexes by blue-native gel electrophoresis has been adapted from Schagger and von Jagow, 1991 [143], and Sasarman et al., 2008 [238].
A) Materials i) Reagents
Aminocaproic acid
Bis-tris
Tricine
Serva Blue G
Acrylamide/bisacrylamide
EDTA n-Dodecyl β-D-maltoside (DDM)
Ammonium persulfate
73
Glycerol
TEMED ddH2O
ii) Equipment
Gradient former
Peristaltic pump
Gel electrophoresis apparatus
iii) Buffers and Solutions
• 3× Gel buffer (1.5 M aminocaproic acid, 150 mM Bis-tris, pH 7.0)
o 19.68 g aminocaproic acid
o 3.14 g Bis-tris
o up to 100 mL ddH2O
• Cathode buffer (15 mM Bis-tris, 50 mM Tricine, pH 7.0)
o 3.14 g Bis-tris
o 8.96 g Tricine
o up to 1 L ddH2O
• Blue cathode buffer
o 100 mL Cathode buffer
o 0.02 g Serva Blue G
• Anode buffer (50 mM Bis-tris, pH 7.0)
o 20.93 g Bis-tris
74
o up to 2 L ddH2O
• Acrylamide/bisacrylamide mix (48% acrylamide, 1.5% bisacrylamide (99.5T,C))
o 24.0 g acrylamide
o 0.75 g bisacrylamide
o up to 50 mL ddH2O
• Membrane buffer
o 0.5 mL 3× Gel buffer
o 0.5 mL 2 M aminocaproic acid
o 4 µL 500 mM EDTA
• SBG sample buffer (0.75 M aminocaproic acid, 5% Serva Blue G)
o 3.75 mL 2 M aminocaproic acid
o 0.5 g Serva Blue G
o up to 10 mL ddH2O
• 2 M Aminocaproic acid
o 13.12 g aminocaproic acid
o 50 mL ddH2O
• 10% n-Dodecyl β-D-maltoside (0.1 g/mL)
• 10% Ammonium persulfate (0.1 g/mL)
• Serva Blue G causes a negative charge shift of the proteins and is able to remain tightly
bound to the protein [143].
• Aminocaproic acid supports the solubilizing properties of neutral detergents and allows
for omission of any salt that would impair electrophoresis [143].
75
• Bistris is a base with a pK in the slightly acidic range (pK 6.5-6.8) and is thus able to
stabilize pH 7.5 in the gel [143].
• Tricine was found to empirically have the optimum pK (8.15) and the appropriate relative
mobility at pH 7.5 to allow protein separation within polyacrylamide gradient gels [143].
B) Method i) Sample Preparation
• Prepare samples for blue-native gel electrophoresis. Use isolated mitochondria from cells
or tissues (refer to "Mitochondria isolation" section). Alternatively, isolation of
mitochondria from cells in suspension may not be necessary due to the high
concentration of mitochondrial respiratory complexes relative to other cellular proteins.
• Wash cells/mitochondria once with PBS (refer to "Plasma membrane isolation" section),
then resuspend in ice-cold PBS.
• Measure the protein concentration: take an aliquot of cell suspension, sonicate or use
detergent (e.g., CHAPS), and measure protein concentration (e.g., using BCA Protein
Assay kit (Pierce)).
• Add more PBS, then centrifuge the cells.
• Resuspend cells in PBS to a final concentration of 5 mg/mL according to the protein
concentration measured above. Place cells in microfuge tube.
• Add an equal volume of digitonin (4 mg/mL) and incubate on ice for 5 minutes (final
concentration of cells is 2.5 mg/mL, final concentration of digitonin is 2 mg/mL,
digitonin:protein ratio is 0.8).
• Following 5 minute incubation, add PBS to a final volume of 1.5 mL.
76
• Centrifuge at 10,000 × g for 10 min.
• Resuspend pellet (i.e., membranes) in Membrane buffer (half the volume of PBS used to
bring the sample to 5 mg/mL previously).
• Add 10% DDM to a final concentration of 1% (e.g., 1/10 volume of Membrane buffer
added in the previous step), then incubate on ice for 15 min.
• Centrifuge at 20,000 × g for 20 min at 4°C.
• Place supernatant in a new microfuge tube and measure the protein concentration (e.g.,
using BCA Protein Assay kit (Pierce)).
• Add SBG sample buffer (e.g., half the volume of 10% DDM added previously).
• Apply the samples to a blue-native gel for electrophoresis. Alternatively, samples may be
stored at -20°C.
ii) Performing Blue-Native Gel Electrophoresis
• Prepare 6% and 15% resolving gel mixture according to Table 1.
• Fill the tubing and 1 cm of cassette height with ddH2O.
• Fill the front reservoir of the gradient mixer with 2.8 mL of 6% gel mixture and the rear
reservoir with 2.3 mL of 15% gel mixture, then place the conical insert into the rear
reservoir.
• Fill the gel cassette by means of underlaying the gel under water using a peristaltic pump
at speed 9.
• After polymerization, wash gel surface with 1× Gel buffer.
• Prepare 2.5 mL of 4% stacking gel mixture according to Table 1, then insert comb and
fill to the top with stacking gel mixture.
77
• After polymerization, remove comb and wash wells with 1× GB.
• Fill the wells with Blue cathode buffer, and underlay the samples into the wells (10 - 30
µg protein per well).
• Place gel cassettes into the apparatus and fill upper and lower reservoir with blue cathode
buffer and anode buffer, respectively.
• Run for 15 min at 40 V, then at 80 V until the dye reaches 2/3 of the gel. Replace the
blue cathode buffer with cathode buffer and continue the electrophoresis until the dye
front reaches the end of the gel.
• Remove the gel from the plates and proceed with either i) western blotting; ii) 2D gel
electrophoresis; or iii) silver staining (Figure 2.1).
iii) Analysis by Western Blotting
• For western blotting, place gels in Towbin semi-dry transfer buffer for 20 min on a
rocker.
• Place a polyvinylidene fluoride (PVDF) membrane in methanol for 1 min, rinse three
times with ddH2O, then place in Towbin semi-dry transfer buffer for 20 min on a shaker.
• Prepare stack on a semi-dry transfer unit as follows: thick blot paper, thin blot paper,
PVDF membrane, gel, thin blot paper, thick blot paper. Ensure there are no bubbles
between any of the layers to avoid poor transfer. Transfer for 42 min, 16 V, 0.26 limit (or
0.52 limit for two gels).
• Proceed with immunoblotting using specific antibodies. Note: some antibodies are able to
detect proteins in their native conformation. An alternative is to denature the gel prior to
membrane transfer by placing it in SDS-containing 1× sample buffer (refer to "Plasma
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membrane isolation" section) on a shaker for 30 min, boil briefly in a microwave (e.g., 10
sec), and place on a shaker for an additional 15 min. This may allow binding by
antibodies that require the protein to be denatured.
iv) 2D Blue-Native Polyacrylamide Gel Electrophoresis (2D BN-PAGE)
• For 2D gel electrophoresis, cut the gel slice of interest from the 1D blue-native gel.
• Place the gel slice in denaturing SDS-containing 1× sample buffer on a shaker for 30 min,
boil briefly in a microwave (e.g., 10 sec), and place on a shaker for an additional 15 min.
• Prepare SDS-containing 10% polyacrylamide gels using a comb designed to hold a gel
slice and a separate well to load the molecular weight ladder.
• Proceed with polyacrylamide gel electrophoresis as normal, then transfer to a PVDF
membrane and blot with the antibody of interest.
v) Silver Staining
• For silver staining of 1D or 2D blue-native gels, the Silver Stain Plus kit (Bio-Rad) is
recommended.
Discussion
Identification of multimolecular complexes within the biochemical microenvironment in which they function is a first step towards characterizing their functional role and targeting them for therapeutic purposes. The biophysical environment in which assembly of the complexes takes place may in itself not only define critical stages for their formation but also focus the search for molecular interactions. We have discussed here several methods to isolate specialized
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membrane microdomains from various membranes of the cell, including the plasma membrane and mitochondrial membranes, and the analysis of multimolecular complexes and supercomplexes in cardiolipin-enriched microdomains in the mitochondrial inner membrane.
A few aspects regarding the outlined protocols deserve further consideration. As reported by others, in our hands, mitochondrial respiratory supercomplex isolation is best achieved using the detergent digitonin since it is sufficiently mild to preserve the supramolecular interactions of multichain protein complexes [154]. Some supercomplexes (and all individual complexes) can be extracted using other detergents, namely Triton X-100, NP-40 (Igepal CA-
630), Tween-20, and DDM [154]. As a control to measure the total amount of individual complexes, we recommend treating the samples of interest with DDM since all individual complexes are detectable by western blotting following this treatment with no detectable proportion associated with supercomplexes. Meanwhile, the detergents Brij-96V, cholate,
Empigen BB, perfluoro octanoic acid, and CHAPS are largely unable to extract any individual complexes or supercomplexes [154].
It is also important to keep in mind that the detection of an array of molecules within the fraction of detergent-insoluble microdomains of a given compartment does not necessarily correlate with colocalization of these molecules within the same microdomains. This is due to the heterogenity of lipid rafts, determined by many factors including ectodomain interactions between surface receptors [239, 240].
Finally, we acknowledge that there is an ongoing debate about the biological significance of the biochemical preparation of detergent-insoluble microdomains, and the multi-molecular complexes that are isolated within these compartments. There is still controversy on whether these domains do actually exist as discrete entities in vivo or rather result from the biochemical
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manipulation of the cells and their membranes. This debate is apparent when discussing the existence and biological significance of mitochondrial respiratory supercomplexes. Assessment of the different sides of this argument goes beyond the scope of this methodological paper.
Summary
It is becoming increasingly apparent that there are specialized membrane microdomains in many biological membranes, including the plasma and mitochondrial membranes, and that these microdomains are heterogeneous with varying compositions. The support for the existence of these microdomains in a natural setting and their formation, independently of biochemical manipulation or as an experimental artifact, is still under debate. However, through the use of different approaches, including those outlined in this paper, it is becoming increasingly clear that these specialized membrane microdomains are biologically relevant in distinct ways depending on their location and composition. It is hoped that the techniques detailed here will be helpful to explore novel models to test and verify the functional correlates that will support their biological significance.
Acknowledgement
We thank Dr. Eric Shoubridge (McGill University, Montréal, QC) for helpful comments and discussions in the development of protocols for mitochondrial isolation and 2D-BN-PAGE.
Work at the Madrenas laboratory is funded by the CIHR. J.M. holds a Tier I Canada Research
Chair in Human Immunology.
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Table 2.1. Preparation of gradient gels for blue-native gel electrophoresis.
Volumes listed are sufficient for 1 or 2 gels (i.e., 2.5 or 5.0 mL, respectively).
4% Stacking gel 6% Resolving gel 15% Resolving gel
2.5 mL 5.0 mL 2.5 mL 5.0 mL 2.5 mL 5.0 mL ddH2O 1.45 mL 2.87 mL 2.72 mL 5.44 mL 0.84 mL 1.68 mL
3× Gel buffer 0.82 mL 1.64 mL 1.65 mL 3.3 mL 1.65 mL 3.3 mL
Acrylamide/bisacrylamide 0.2 mL 0.4 mL 0.6 mL 1.2 mL 1.5 mL 3.0 mL
Glycerol - - - - 1 mL 2 mL
Ammonium persulfate 30 µL 60 µL 30 µL 60 µL 5 µL 10 µL
TEMED 3 µL 6 µL 2 µL 4 µL 2 µL 4 µL
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Figure legend
Figure 2.1. Procedure for two-dimensional blue-native gel electrophoresis (2D-BN-PAGE).
(A) Non-denatured samples are loaded into the wells of a non-denaturing 4 – 16% gradient polyacrylamide gel. Following electrophoresis, a lane of interest is cut from the gel and this gel slice is placed in denaturing sample buffer prior to its placement atop a denaturing polyacrylamide gel (e.g., 10%). Following second-dimension PAGE, the gel can be silver- stained to visualize the proteins that comprise the first-dimension multiprotein complex bands.
Alternatively, the proteins from the gel can be transferred to a PVDF membrane for western blot detection of specific proteins (not shown). (B) Silver stain of 2D-BN-PAGE of isolated mitochondria from adult C57BL/6 mouse brain. A molecular weight ladder is included on the far right-hand portion of the gel.
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Figure 2.1
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Preface to Chapter 3
Previous data generated from our laboratory led us to a model of SLP-2 function in the mitochondria such that SLP-2 recruits PHBs to cardiolipin-enriched microdomains at the mitochondrial inner membrane, forming functional microdomains where the respiratory complexes operate. Recently, respiratory chain complexes have been shown to possess supramolecular interactions to form respiratory chain supercomplexes (RCS) that allow for more efficient electron transfer and function. Additionally, it is known that cardiolipin is required for
RCS function. Since we have previously identified SLP-2 to be important for the formation of cardiolipin-enriched microdomains and mitochondrial respiratory chain function, we wanted to utilize specific techniques including blue-native gel electrophoresis to study the role of SLP-2 in the formation and function of RCS.
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Chapter 3: Stomatin-like Protein 2 is Required for In vivo Mitochondrial
Respiratory Chain Supercomplex Formation and Optimal Cell Function
Panagiotis Mitsopoulosa, Yu-Han Changa, Timothy Waib, Tim Königb, Stanley D. Dunnc,
Thomas Langerb, Joaquín Madrenasa
From
aMicrobiome and Disease Tolerance Centre, Department of Microbiology and Immunology,
McGill University, Montreal, Quebec, Canada; bInstitute for Genetics, Cologne Excellence
Cluster on Cellular Stress Responses in Aging Associated Diseases, Center for Molecular
Medicine, University of Cologne, Cologne, Germany; and cDepartment of Biochemistry,
University of Western Ontario, London, Ontario, Canada
This chapter contains the following published manuscript:
Mitsopoulos P, Chang YH, Wai T, Konig T, Dunn SD, Langer T, Madrenas J. Stomatin-like protein 2 is required for in vivo mitochondrial respiratory chain supercomplex formation and optimal cell function. Mol Cell Biol. 2015; 35: 1838-1847.
Copyright © 2015. American Society for Microbiology
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Abstract
Stomatin-like protein (SLP)-2 is a mainly mitochondrial protein that is widely expressed and is highly conserved across evolution. We have previously shown that SLP-2 binds the mitochondrial lipid cardiolipin and interacts with prohibitin-1 and -2 to form specialized membrane microdomains in the mitochondrial inner membrane, which are associated with optimal mitochondrial respiration. To determine how SLP-2 functions, we performed bioenergetic analysis of primary T cells from T cell-selective Slp-2 knockout mice under conditions that forced energy production to come almost exclusively from oxidative phosphorylation. These cells had a phenotype characterized by increased uncoupled mitochondrial respiration and decreased mitochondrial membrane potential. Since formation of mitochondrial respiratory chain supercomplexes (RCS) may correlate with more efficient electron transfer during oxidative phosphorylation, we hypothesized that the defect in mitochondrial respiration in SLP-2-deficient T cells was due to deficient RCS formation. We found that in the absence of SLP-2, T cells had decreased levels and activities of complex I-III2 and I-III2-IV1-3 RCS, but no defects in assembly of individual respiratory complexes. Impaired
RCS formation in SLP-2-deficient T cells correlated with significantly delayed T cell proliferation in response to activation under conditions of limiting glycolysis. Altogether, our findings identify SLP-2 as a key regulator of the formation of RCS in vivo, and show that these supercomplexes are required for optimal cell function.
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Introduction
Stomatin-like protein (SLP)-2 is a mainly mitochondrial protein that is widely expressed and is highly conserved across evolution [40, 78, 79, 84]. We have previously shown that SLP-2 binds the mitochondrial phospholipid cardiolipin and interacts with prohibitin (PHB)-1 and -2, which are proposed to form specialized cardiolipin-enriched microdomains in the mitochondrial inner membrane important for optimal respiratory function [86, 87]. Indeed, we showed that deletion of Slp-2 results in decreased cardiolipin microdomains and increased mitochondrial respiration uncoupled from ATP synthase activity, a defect overcome by an increased reliance on glycolysis
[87].
Recently, it has been shown that mitochondrial respiratory complexes are not randomly dispersed throughout the mitochondrial inner membrane, but instead have supramolecular interactions allowing them to form respiratory chain supercomplexes (RCS) [141-144]. RCS are commonly isolated from mitochondrial membranes using mild detergents, usually digitonin, followed by blue-native polyacrylamide gel electrophoresis (BN-PAGE), and their existence has been confirmed by electron microscopy and single particle image processing [150, 151]. RCS occur mainly between complexes I, III, and IV (NADH-coenzyme Q reductase, ubiquinol- cytochrome c reductase, and cytochrome c oxidase, respectively) with various stoichiometries, whereas complex V (ATP synthase) can form dimers and oligomers [144, 153], and complex II
(succinate-CoQ reductase) is thought to remain fluid [142, 145]. Coenzyme-Q, and cytochrome c have also been shown to associate with RCS [154], and recent evidence indicates that there are two distinct pools of coenzyme-Q that are dedicated to reducing equivalents from NADH or
FADH2 [147]. RCS assemble to facilitate more efficient electron transfer and to allow for the utilization of different electron transport pathways and substrates during oxidative
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phosphorylation, to stabilize complex I and other complexes, and to limit the production of reactive oxygen species generated from electron transport during oxidative phosphorylation
[142, 145-149]. Altogether, these effects support the importance of RCS for optimal mitochondrial function.
The molecular machinery involved in the formation, maintenance, and regulation of RCS is not well characterized. The defects in RCS assembly/stability observed in some human genetic diseases have provided clues regarding the requirements of RCS formation and maintenance. For example, Barth syndrome, characterized by cardiomyopathy, skeletal myopathy, and neutropenia, is caused by a mutation in the tafazzin gene that impairs cardiolipin remodeling and destabilizes RCS [158], a feature of cardiolipin present in other systems [164, 165]. Cardiolipin has been shown to physically bind to complexes I, III, IV, and V [167, 168], and to be required for the activities of these complexes [169, 171]. It has also been suggested that cardiolipin may fill the spaces between complexes organized into RCS. Thus, cardiolipin is an important factor for proper RCS formation.
Several proteins have also been identified to be important for RCS formation. Two related Saccharomyces cerevisiae proteins, Rcf-1 and Rcf-2, and a mammalian homolog hypoxia-induced gene (HIG)2A, are necessary for assembly of mature complex IV and affect complex III-IV RCS formation [172-174]. Furthermore, supercomplex assembly factor I
(SCAFI) has been shown to act as a RCS chaperone by specifically allowing assembly of complex III-IV supercomplexes [147, 159]. The identification of additional factors required to assemble and maintain RCS will be necessary to define the mechanisms governing this process and may be important for the modulation of mitochondrial function.
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Given our previous findings that SLP-2 forms cardiolipin-enriched microdomains in the mitochondrial inner membrane, and that deletion of Slp-2 results in decreased levels and activities of certain mitochondrial respiratory chain components and increased uncoupled respiration [87], we sought to examine whether SLP-2 was required for the formation of RCS.
Here, we show that the efficiency of mitochondrial respiration in primary T cells lacking SLP-2 expression is impaired when the cell must rely almost exclusively on oxidative phosphorylation for energy production, and that this effect is associated with decreased amounts of (I-III2)1-2 and
I-III2-IV1-3 RCS. Functionally, this defect results in decreased T cell proliferation when glycolysis, but not oxidative phosphorylation, is restricted. These data identify SLP-2 as a key regulator of mitochondrial respiratory function by linking formation of cardiolipin-enriched microdomains with optimal assembly of RCS, and point to a functional role of RCS in T cells in vivo.
Results
SLP-2-deficient T cells have increased uncoupled respiration and decreased mitochondrial membrane potential, both exacerbated under conditions of limiting glycolysis.
We have previously generated T cell-specific Slp-2 knockout (SLP-2 T-K/O) mice and have shown that deletion of Slp-2 in T cells in vivo negatively impacts the function of the mitochondrial respiratory chain, resulting in decreased levels of some complex I and II subunits as well as decreased activities of complexes I and II + III [87]. In addition, using extracellular flux technology to assess cellular respiration, we have shown that when glucose is available,
SLP-2-deficient T cells from these mice exhibit increased mitochondrial respiration uncoupled from ATP synthase activity. In these T cells, inefficient mitochondrial respiration was
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compensated by an increased reliance on glycolysis to preserve function [87]. To test the metabolic function of SLP-2 T-K/O T cells under conditions of almost exclusive reliance upon oxidative phosphorylation, we incubated primary wild-type (WT) and SLP-2 T-K/O T cells in media containing galactose but lacking glucose, thereby severely limiting the glycolytic rate since this sugar enters the glycolytic pathway at a much slower rate than glucose as it is a much poorer substrate for hexokinase [241]. We found that although the basal cellular and mitochondrial OCRs were unchanged between WT and SLP-2-deficient T cells (Figure 3.1A-B), there was a trend towards decreased percentage of mitochondrial respiration coupled to ATP synthase activity in SLP-2-deficient T cells cultured in both glucose- and galactose-containing media relative to WT T cells (Figure 3.1C). Though this did not reach statistical significance, analysis of uncoupled mitochondrial respiration rates revealed a significant increase in this parameter in SLP-2-deficient T cells that was significantly greater when glycolysis was limiting
(Figure 3.1D). This increase in uncoupled mitochondrial respiration is comparable to the increase observed in hearts of a mouse model of diabetes and obesity [242]. We also confirmed cells were respiring normally using FCCP as a positive control (data not shown). Although total cellular
ATP levels were not changed (Figure 3.1E), SLP-2-deficient T cells exhibited a decrease in mitochondrial membrane potential when cultured in galactose-containing media (Figure 3.1F), pointing to impaired function of the mitochondrial respiratory chain. This correlated with an increased reliance by SLP-2 T-K/O T cells on glycolysis when cultured in both glucose- or galactose-containing media, as noted by the significantly greater basal extracellular acidification rates (Figure 3.1G). Finally, SLP-2-deficient T cells exhibited increased levels of reactive oxygen species (ROS) (Figure 3.1H), a characteristic of mitochondrial respiratory chain
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impairment. Taken together, these data show an impaired mitochondrial respiratory profile in the absence of SLP-2 that was worsened by experimentally limiting the cellular glycolytic rate.
SLP-2 deficiency results in defective RCS formation in cell lines.
The makeup of RCS can vary between tissues and species, but RCS are commonly thought to occur as I-III2-IV1-4, or as I-III2. In the current study, we have estimated RCS stoichiometries based on previous literature [81, 144, 243] and approximate molecular weights. The previous data showing impaired mitochondrial respiratory function in the absence of SLP-2, as well as our findings that SLP-2 is important for the formation of cardiolipin-enriched membrane microdomains necessary for optimal mitochondrial respiratory function [86, 87] suggested that there might be impaired RCS formation in the absence of SLP-2. To test this, we first generated
Slp-2-knockout mouse embryonic fibroblasts (Slp-2-/- MEFs) in vitro and performed BN-PAGE of WT and Slp-2-/- MEF mitochondrial lysates. Immunoblot analyses following BN-PAGE showed a dramatic reduction of respiratory complex I (NDUFA9 and NDUFB6), III (UQCRC2), and IV (MTCO1) subunits in RCS, specifically of (I-III2)1-2, I-III2-IV3, I-III2-IV2, I-III2-IV, I-III2, and III2-IV arrangements (Figure 3.2A). The significant reductions in the levels of RCS correlated with a decreased growth rate of Slp-2-/- MEFs compared to WT controls when cultured in either glucose- or galactose-containing media (Figure 3.2B). Interestingly, Slp-2-/- MEFs failed to proliferate when glycolysis was limiting (Figure 3.2B, lower panel), consistent with our previous findings that SLP-2-deficient cells had impaired mitochondrial respiratory function.
Together, these data showed defective RCS formation in MEFs in the absence of SLP-2, which correlated with decreased cell cycling.
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Primary SLP-2-deficient T cells have defective RCS formation in vivo.
Next we assessed RCS formation in vivo from mitochondria of primary T cells lacking SLP-2.
To do this, we immunoblotted 1D BN-PAGE gels of digitonin-solubilized mitochondrial lysates from WT or SLP-2-deficient T cells with antibodies against subunits of respiratory complex I
(NDUFA9 and NDUFS3), III (UQCRC2 and Core 1), IV (MTCO1), and V (ATP5A) (Figure
3.3A). We found decreased or undetectable amounts of RCS containing complex I and III subunits, specifically of I-III2-IV3, I-III2-IV2, I-III2-IV, and I-III2 arrangements (Figure 3.3A).
Immunodetection of PHB1 and SLP-2 revealed that these proteins exhibit electrophoretic mobilities conspicuously overlapping with those of larger RCS (Figure 3.3A). The co-migration of PHBs and SLP-2 observed here is in line with previous reports from our group and others that show that SLP-2 associates with PHBs [77, 86].
To further document the decreased complex I association with RCS and its functional significance, we performed a complex I in-gel activity assay on a non-denatured first dimension gel to visualize enzymatically active complex I (Figure 3.3B). Complex I activity, as indicated by the development of purple formazan, was decreased in high-molecular weight regions corresponding to RCS, including I-III2-IV2 and I-III2 (Figure 3.3B).
To corroborate our findings obtained using first dimension BN-PAGE, we cut non- denatured 1D BN-PAGE gels of the same aforementioned WT and SLP-2 T-K/O T cell mitochondrial lysates in half and loaded high- (Figure 3.3C) and low-molecular weight (Figure
3.3D) portions onto second dimension SDS polyacrylamide gels, then immunoblotted them with antibodies directed against subunits of complexes I, II, III, IV, and V. Consistent with the 1D
BN-PAGE results, we observed decreased levels of complex I, III, and IV subunits associated with RCS in mitochondria of SLP-2-deficient T cells relative to WT controls, including I-III2-
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IV3, I-III2-IV2, I-III2-IV, and I-III2 arrangements (Figure 3.3C). The migration of MTCO1 with
RCS, faintly detectable by 1D BN-PAGE analysis (Figure 3.3A), can be better visualized by 2D
BN/SDS-PAGE analysis (Figure 3.3C). Complex V has been reported to associate with RCS under certain circumstances [154], but has also been commonly found to associate into dimers and multimers [144, 153]. We found that this technique was sufficiently sensitive to show migration of complex V at various regions of high-molecular weight that correspond to the migration patterns of RCS, suggesting that these may be multimers of complex V or perhaps associations with certain RCS (Figure 3.3C). However, the multimers of complex V were decreased or absent upon Slp-2 deletion (Figure 3.3C). The levels of non-RCS-associated complexes did not appear different between WT and SLP-2 T-K/O T cells, as can be seen in figure 3.3C (complexes I, III2, and V) and figure 3.3D (complexes II and IV). Furthermore, 2D
BN/SDS-PAGE analysis revealed that PHBs and SLP-2 are associated in high-molecular weight protein complexes, as observed in both first- (Figure 3.3A) and second-dimension gels (Figure
3.3C). Taken together, these results show that lack of SLP-2 expression is associated with deficient RCS formation in vivo.
Formation of individual respiratory complexes is not affected by SLP-2 deficiency. To determine whether the defect in supercomplex formation in SLP-2-deficient cells was a result of defective formation of individual respiratory complexes, we solubilized mitochondrial membranes with the detergent dodecyl maltoside, which has been shown to disrupt RCS formation while maintaining individual respiratory complex interactions [154]. Immunoblotting membranes from first dimension BN-PAGE gels showed no differences in the levels of individual complex I, II, III,
IV, or V in SLP-2 T-K/O T cells compared to WT controls (Figure 3.4A). Subsequent
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immunoblotting for PHB1 and SLP-2 (Figures 3.4B and C, respectively) showed that these proteins maintain their migration pattern at high molecular weight when membranes are solubilized with dodecyl maltoside compared to digitonin (Figure 3.3A). These results indicated that the defect observed in the formation of RCS in the absence of SLP-2 was not a result of a deficiency in individual respiratory complex formation.
SLP-2 deficiency results in delayed T cell proliferation under conditions of limited glycolysis.
We next studied the functional consequences of impaired formation of RCS in SLP-2-deficient T cells. This system provided us with the opportunity to assess the activation and proliferation of
WT and SLP-2 T-K/O T cells in response to α-CD3/CD28 stimulation under conditions of either unlimited or limited glycolysis. First we assessed early activation events by determining the upregulation of CD25 and the down-regulation of CD62L by flow cytometry. CD25 was significantly upregulated following activation of WT and SLP-2 T-K/O T cells (Figure 3.5A, left panel). Such upregulation was significantly decreased in T cells activated in conditions of restricted glycolysis (i.e., galactose-containing media) or restricted oxidative phosphorylation
(i.e., oligomycin-containing media), yet neither depended upon SLP-2. CD62L expression was decreased in all activation conditions and this down-regulation was also unaffected by SLP-2 deficiency (Figure 3.5A, middle panel). The viability of T cells was unaffected in all activation conditions except when T cells were restricted in their ability to use both glycolysis and oxidative phosphorylation, under which almost all T cells died (Figure 3.5A, right panel). These results indicated that deficiency of SLP-2 in T cells did not affect early activation events.
Next, we studied the effect of SLP-2 deficiency on T cell cycling. We found that SLP-2- deficient T cells proliferated similarly to WT T cells under conditions of maximal activation with
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α-CD3/CD28 and unlimited supply of glucose (Figure 3.5B). However, under conditions of limited glycolysis, SLP-2-deficient T cells displayed a significantly delayed profile of proliferation (Figure 3.5B). On a population level and after three days of stimulation, only 18% of SLP-2-deficient T cells had gone into high cycling compared to 37% of WT T cells whereas
47% of SLP-2-deficient T cells remained in low cycling compared to only 24% of WT T cells
(Figure 3.5C). By day 3 after stimulation, SLP-2-deficient T cells were on average delayed by at least one-cycle (Figure 3.5D), and this could not be attributed to decreased cell viability (Figure
3.5E). Furthermore, we found that SLP-2-deficient T cells cultured in glucose-restricted conditions were delayed in entry to the cell cycle 37 h following stimulation, as determined by the delayed increase in Ki67 expression (Figure 3.5F). Non-stimulated cells cultured in glucose, and stimulated cells cultured with neither glucose nor galactose exhibited poor viability after 24 hours and failed to proliferate after up to 5 days (data not shown).
To examine the cause of the delayed proliferation of SLP-2-deficient T cells, we determined the interleukin (IL)-2 response of these T cells to α-CD3/CD28 stimulation in conditions of unlimiting and limiting glycolysis. As shown in figure 3.6A, SLP-2-deficient T cells had decreased production of IL-2. Such a defect was apparent both in cultures containing glucose as well as in cultures containing galactose but not glucose. Furthermore, as we had previously shown, the defective IL-2 response in SLP-2-deficient T cells was a result of a post- transcriptional mechanism as the production of IL-2 mRNA was similar between WT and SLP-
2-deficient T cells cultured in glucose- (Figure 3.6B) or galactose-containing media (Figure
3.6C). To further elucidate the mechanism responsible for the impairment in IL-2 production, we performed intracellular IL-2 staining of α-CD3/CD28-stimulated WT and SLP-2 T-K/O T cells treated with brefeldin A at the time of stimulation to prevent cytokine secretion. We found that
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the percentage of IL-2+ SLP-2 T-K/O T cells was reduced relative to WT T cells when cultured in glucose- (16.2 vs 22.5%) or galactose-containing media (18.1 vs 28.3%). These data indicate that the post-transcriptional impairment of IL-2 production in SLP-2 T-K/O T cells is likely due to a defect in protein translation and not secretion. Although the IL-2 levels were decreased, we found no difference in IκB phosphorylation (Figure 3.6D) or mitochondrial calcium release
(Figure 3.6E) following stimulation, suggesting normal T cell signaling. To determine whether the lower IL-2 levels were responsible for the delayed proliferation of glucose-restricted SLP-2- deficient T cells, we supplemented the cultures with 50 ng/mL of mouse recombinant (mr)IL-2 at the time of stimulation. However, this did not restore the delay in proliferation of glucose- restricted SLP-2-deficient T cells (Figure 3.6F), indicating an alternate mechanism. Altogether, these results documented a functional defect associated with the impaired formation of RCS in the absence of SLP-2.
Discussion
The molecular machinery that supports the optimal assembly and function of respiratory complexes in the mitochondrial inner membrane is not fully defined. Recent evidence suggests that different proteins may be involved in the assembly of individual components of the respiratory chain into RCS, i.e., assemblies of respiratory chain complexes with defined stoichiometries. In the context of this model, our findings identify SLP-2 as a critical player in the formation of these RCS, specifically (I-III2)1-2, I-III2-IV3, I-III2-IV2, I-III2-IV, I-III2, and III2-
IV arrangements, and potentially complex V-containing dimers and multimers. The precise mechanism by which SLP-2 affects RCS formation, whether it is primarily important for RCS assembly or stability or rather for regulating the expression of individual respiratory complex
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subunits, is currently unknown and is beyond the scope of this study. However, our data indicate that the mechanism responsible for defective RCS formation in the absence of SLP-2 does not relate to improper formation of individual respiratory complexes. Using T cells from a genetically engineered mouse strain in which the Slp-2 gene is specifically deleted in T cell progenitors, we have shown here that the defect in RCS formation is linked with an impaired respiratory profile, more apparent in conditions of limited glycolysis, and this translates into a defective functional T cell response.
There is evidence to suggest that the lipid composition of the mitochondrial inner membrane is key for proper RCS formation, particularly since the interactions between complexes occur within the lipid bilayer of the membrane while the matrix portions do not strongly interact [145]. The mitochondrial signature phospholipid cardiolipin is a critical component of this environment and has been shown to be essential for proper RCS formation
[142, 145, 165], with defective cardiolipin remodeling and RCS instability associated with Barth syndrome [141, 158]. We have previously shown that SLP-2 binds cardiolipin and interacts with
PHB1 and PHB2, and that in SLP-2-deficient T cells, the levels of cardiolipin and PHBs in mitochondrial detergent-insoluble membranes are significantly decreased [86, 87]. As a consequence, deficiency in SLP-2 expression would impair the proper compartmentalization of the mitochondrial membrane and result in defective RCS formation. This would be consistent with the evidence that SLP-2 can dimerize [78] and may form homooligomeric complexes similar to PHBs [102]. Indeed, SLP-2 was found to migrate in a high-molecular weight complex by BN-PAGE in rat liver, potentially as a homooligomeric complex [81], and a recent report has claimed that a plant homologue of Slp-2 in Arabidopsis thaliana may also be linked to RCS organization in plants [244]. These reports corroborate our findings and, together with our data
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showing impaired RCS formation both in Slp-2-/- MEFs and SLP-2-deficient T cells, point to a critical role of SLP-2 in RCS formation, one that is conserved across different tissues and species.
The abnormal compartmentalization of cardiolipin in the absence of SLP-2 may explain the defective RCS formation, particularly of (I-III2)1-2 and I-III2-IV1-3 arrangements. The formation of cardiolipin-enriched membrane microdomains facilitated by SLP-2 [86, 87] may provide a platform for optimal RCS assembly and/or stability in the inner membrane of the mitochondria. Formation of these microdomains likely involves PHBs in addition to SLP-2, and
PHBs have been previously proposed to be involved in the assembly of respiratory complexes
[245]. PHBs comigrate with RCS by 2D BN/SDS-PAGE in this study and in [154], while SLP-2 also migrated with RCS, though at a slightly slower rate than PHBs. The interaction between
SLP-2 and PHBs [77, 86, 87] may have structural and functional implications in the regulation of mitochondrial function [230], though PHB oligomerization is not affected by Slp-2 deletion in our model. As we previously reported, our data shows that the levels of PHBs present in the detergent (i.e., triton X-100, digitonin, or dodecyl maltoside)-soluble fraction of mitochondrial lysates in the absence of SLP-2 are not different from control levels [87]. Furthermore, SLP-2 does not seem to directly interact with RCS because experiments with plant mitochondria using complex I mutants lacking proper RCS associations showed that the migration pattern of a plant
Slp-2 homolog is largely unchanged [244]. It has also recently been reported that the Cox7a2l gene in several strains of mice, including C57BL/6J, carries a mutation that prevents the formation of complex IV-containing RCS. The mouse strains used in this study have a
C57BL/6N Tac background and our data demonstrates that these mice retain the ability to form complex IV-containing RCS.
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The defective formation of RCS was associated with an impaired respiratory profile. This profile is characterized by increased uncoupled respiration, decreased mitochondrial membrane potential, and an increased reliance on glycolysis. In addition, we observed a slight but significant increase in ROS produced upon stimulation of SLP-2-deficient T cells. Reactive oxygen species are produced by the respiratory chain under normal circumstances [246] and defective RCS formation, particularly of I-III-containing RCS, leads to increased ROS production [145]. This supports our findings of defective (I-III2)1-2 and I-III2-IV1-3 RCS formation in the absence of SLP-2.
It has recently been reported that either glycolysis or oxidative phosphorylation alone is sufficient to sustain normal T cell proliferation in response to activation [128]. SLP-2 plays a role in T cell activation [76] and its deficiency is associated with impaired T cell responses [87].
We show here that proliferation of SLP-2 T-K/O T cells is not significantly affected compared to that of WT T cells when oxidative phosphorylation is restricted if glycolysis is not limiting, but is significantly delayed when glycolysis is restricted. Furthermore, this delay in proliferation is due in part to a delayed entry into the cell cycle. These findings would be expected in light of the bioenergetics profile of these T cells and suggest that the metabolic output of the respiratory chain in SLP-2 T-K/O T cells is insufficient to meet the demands required for optimal T cell proliferation. Such a defect may be secondary to impaired RCS assembly and/or stability, and results in delayed proliferation. However, this delayed cycling cannot be rescued by restoration of IL-2 levels, indicating the involvement of other factors.
Chang and colleagues have recently reported that the expression of several cytokines, including IL-2 and interferon (IFN)-γ, is decreased when glycolysis is restricted [128]. This decrease in cytokine expression is mediated by a post-transcriptional mechanism involving
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increased ability for glyceraldehyde-3-phosphate dehydrogenase (GAPDH) to bind to AU-rich elements within the 3ʹ un-translated region of these respective mRNAs, effectively reducing their translation in response to decreased glycolysis [128]. We had already shown that SLP-2-deficient
T cells have a post-transcriptional defect in IL-2 production in response to stimulation through the antigen receptor [87]. However, contrary to the observation by Chang et al. with limited glycolysis in WT cells, the defect observed in SLP-2-deficient T cells is selective for IL-2 and does not occur for other cytokines such as IFN-γ or IL-17. Furthermore, the post-transcriptional
IL-2 defect in SLP-2-deficient T cells is present to the same extent under conditions of limiting or unlimiting glycolysis, and is likely a result of impaired protein translation and not secretion.
Altogether, these results suggest that the mechanism responsible for the post-transcriptional defect in IL-2 production in SLP-2 T-K/O T cells is independent of GAPDH, and may involve a novel pathway linked to impaired mitochondrial respiration.
In summary, our data show that SLP-2 is required for optimal RCS formation in the mitochondrial inner membrane. In the absence of SLP-2 there is significant loss of (I-III2)1-2 and
I-III2-IV1-3 RCS, which may be due to a defect in assembly or lack of stability of these supercomplexes. More importantly, abnormal RCS expression correlates with impaired mitochondrial respiration and results in decreased effector cell function. Further studies are required to determine the mechanism whereby SLP-2 functions to allow for optimal RCS formation: whether it is a direct function of SLP-2, or rather if it is indirect, perhaps through the generation of specialized cardiolipin-enriched microdomains in the mitochondrial inner membrane. Together, these findings provide evidence that RCS are a relevant functional entity and identify SLP-2 as a key player in the assembly and/or stability of these supercomplexes.
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Materials and Methods
Mice. T cell-specific Slp-2 knockout mice in the C57BL/6N Tac background were generated as described [87]. Briefly, Slp-2lox/wt mice were crossed with CD4-Cre mice (Taconic Farms,
Hudson, NY) to generate Slp-2lox/lox/Cre+ (SLP-2 T-K/O) or Slp-2lox/lox/Cre- (WT) mice. Breeding colonies were derived from the same Slp-2lox/lox breeders and kept in parallel. Mice were maintained in the animal facility at McGill University with approval of the Comparative
Medicine and Animal Resources Centre in accordance with the Canadian Council on Animal
Care Guidelines.
Mouse Embryonic Fibroblasts. Primary WT and Slp-2-/- mouse embryonic fibroblasts (MEFs) were isolated from E13.5 Slp-2lox/lox conditional embryos [87] and immortalized using a plasmid expressing SV40 large T antigen [18]. Immortalized WT and Slp-2-/- MEFs were then generated in vitro as previously described [94]. Slp-2-/- clones were confirmed by PCR and immunoblot analyses.
Cell Culture. T cells were isolated from WT and SLP-2 T-K/O mouse spleens by negative selection using the EasySep™ Mouse T cell Isolation kit (Stemcell Technologies, Vancouver,
Canada). Cells were stimulated with 5 µg/mL α-CD3ε and 2 µg/mL α-CD28 plate-bound antibodies (eBioscience, San Diego, CA), and cultured in 25 mM glucose- or galactose- containing medium with or without the addition of 1 nM oligomycin. Glucose- and galactose- containing media were prepared from glucose-free RPMI supplemented with 25 mM glucose or galactose, 10% dialyzed fetal bovine serum, 1 mM sodium pyruvate, 2 mM L-glutamine, 0.1%
β-mercaptoethanol, and penicillin/streptomycin. Immortalized WT and Slp-2-/- MEFs were
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cultured in glucose-free DMEM+glutamax (Gibco, Carlsbad, CA) supplemented with 25 mM glucose or galactose, 10% dialyzed fetal calf serum (Gibco), 1 mM sodium pyruvate, and non- essential amino acids. Galactose-containing media contained little or no glucose. For cell growth experiments, WT and Slp-2-/- MEFs were seeded at 50,000 cells/plate and harvested every 24 hours to assess viable cell number by trypan-blue exclusion. Cells were incubated at 37°C in a humidified atmosphere of 5% CO2.
Bioenergetic Analysis. Oxygen consumption and extracellular acidification rates (OCR/ECAR) were measured using a Seahorse XF-24 analyzer (Seahorse Bioscience, North Billerica, MA) as described previously [87]. Briefly, following stimulation for 48 hours, WT and SLP-2 T-K/O T cells were suspended in Seahorse XF assay media supplemented with 2 mM L-glutamine, 2 mM sodium pyruvate, and 25 mM glucose or galactose and seeded onto XF24 V7 24-well cell culture plates coated with 50 µg/mL poly-D-lysine at 500,000 cells/well. OCR and ECAR (a measure of lactic acid formed during glycolysis) were measured prior to and following the sequential additions of 10 µM oligomycin, 100 nM rotenone and 1 µM antimycin A. Basal OCR refers to the respiration rate measured prior to the addition of mitochondrial drugs. Mitochondrial OCR was calculated by subtracting the OCR in the presence of rotenone/antimycin A from the basal
OCR. Uncoupled OCR refers to oligomycin-insensitive mitochondrial respiration. The percentage of coupled OCR was calculated relative to the total mitochondrial OCR.
Blue-native gel electrophoresis. Blue-native gel electrophoresis was performed based on the method introduced by Schagger and von Jagow with some modifications [143, 238, 245, 247].
Briefly, mitochondria of WT and SLP-2 T-K/O T cells stimulated for 48 hours were isolated
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using a dounce homogenizer and differential centrifugation as described [247]. Mitochondria were lysed using digitonin (4 g/g protein) or n-dodecyl β-D-maltoside (1.6 g/g protein) for 5 min and 20 µg protein was loaded per lane in Novex 3 – 12% Bis-Tris gels (Life Technologies,
Grand Island, NY) and run on ice. Crude mitochondria (100 µg) from WT and Slp-2-/- MEFs were solubilized in 6 g/g digitonin and electrophoresed in 3 – 9% gels. First dimension gels were either denatured with sodium dodecyl sulfate (SDS)-containing buffer for 2D BN/SDS-PAGE, transferred to PVDF membranes for immunoblotting, stained with silver (Silver Stain Plus kit,
Bio-Rad, Hercules, CA), or assayed for in-gel complex I activity (0.1 M Tris-HCl, 0.14 mM
NADH, and 1.0 mg/mL nitro blue tetrazolium (NBT) with gentle agitation [248]). For 2D
BN/SDS-PAGE, 1D gel slices were cut in half and high- and low-molecular weight portions of
WT and SLP-2 T-K/O slices were loaded onto the same 10% 2D SDS polyacrylamide gel. 2D gels were either transferred to PVDF membranes for immunoblotting or stained with silver.
Immunoblotting. PVDF membranes containing protein transferred from 1D BN-PAGE or 2D
BN/SDS-PAGE gels were blotted with antibodies against subunits of mouse complex I
(NDUFA9, NDUFB6, NDUFS3), complex II (SDHA), complex III (UQCRC2, Core 1), complex
IV (MTCO1), and complex V (ATP5A), or Total OXPHOS Antibody Cocktail (Mitosciences,
Eugene, OR), α-PHB1 (Santa Cruz Biotechnology, Santa Cruz, CA), and α-SLP-2 (Protein Tech
Group Inc., Chicago, IL). For phospho-ΙκΒ detection, WT or SLP-2 T-K/O T cells were stimulated with Mouse T Activator Dynabeads (Life Technologies) for the indicated times, then washed with ice-cold PBS containing 400 µM sodium orthovanadate and resuspended in lysis buffer containing 1% Triton X-100 with phosphatase and protease inhibitors for 30 min on ice, as described [87]. Lysates were separated on a 10% SDS-PAGE gel and transferred to a PVDF
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membrane, then immunoblotted with α-phospho-ΙκΒ (Ser32), α-ΙκΒ (Cell Signaling, Danvers,
MA) , α-SLP-2, and α-GAPDH (Chemicon International, Temecula, CA) antibodies.
Enzyme-linked Immunosorbent Assay. WT and SLP-2 T-K/O T cells were stimulated for 24 hours in glucose- or galactose-containing media and IL-2 levels were quantified using the Mouse
IL-2 ELISA Ready-SET-Go!® kit (eBioscience).
Flow Cytometry. To assess T cell proliferation, WT and SLP-2 T-K/O T cells were loaded with 1
µM carboxyfluorescein diacetate succinimidyl ester (CFSE), then stimulated (1 µg/mL α-
CD3ε, 2 µg/mL α-CD28, plate-bound) in glucose- or galactose-containing media with or without oligomycin for 0, 2, 3, 4, and 5 days. To measure CD25 and CD62L expression, cells were blocked with 10 µg/mL Mouse Fc Block (BD Pharmingen, Franklin Lakes, NJ), then stained with 2 µg/mL of mouse BV421 α-CD62L (BD Horizon), APC α-CD25, and APC eFluor 780 α-
CD3 antibodies (eBioscience). For IL-2 supplementation studies, 50 ng/mL mouse recombinant
IL-2 (eBiosciences) was added to the culture media at the start of the experiment. For intracellular IL-2 determination, T cells were treated with BD GolgiPlug (BD Biosciences) containing brefeldin A at the time of α-CD3/CD28 stimulation and fixed 24 h later using BD
Cytofix/Cytoperm (BD Biosciences), then stained with APC α-IL-2 (eBiosciences). Viability was assessed using the Zombie Aqua Fixable Viability kit (BioLegend, San Diego, CA). To assess mitochondrial membrane potential, stimulated WT and SLP-2 T-K/O T cells were stained in non-quench mode with 20 µM tetramethylrhodamine methyl ester (TMRM) for 1 h with no washing [249]. Reactive oxygen species levels were assessed by measuring fluorescence of cells stained with 5-(and-6)-chloromethyl-2',7'-dichlorodihydrofluorescein diacetate, acetyl ester (Life
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Technologies). Cell cycle entry was measured using APC-conjugated anti-Ki67 antibody
(BioLegend). For calcium release assay, cells were stained with Fluo-3-AM (4 µg/mL) and Fura
Red (10 µg/mL; Invitrogen) for 30 minutes. Following baseline measurement, ionomycin (2
µg/mL), FCCP (50 nM), or EDTA (5 mM) were added and fluorescence was measured for 5 min. Flow cytometry was performed using an LSRFortessa flow cytometer (BD), and data were analyzed with FlowJo software (Treestar Inc., Ashland, OR).
Quantitative reverse-transcriptase PCR. WT and SLP-2 T-K/O T cells were cultured for 4, 12, and 24 hours with or without α-CD3/CD28 stimulation in glucose- or galactose-containing media. Cellular RNA was isolated using the RNeasy® Plus Mini Kit (Qiagen, Mississauga, ON,
Canada), and quantified using NanoDrop (Thermo Scientific, Wilmington, DE). cDNA was synthesized from 220.08 ng of total RNA using the Omniscript® Reverse Transcription Kit
(Qiagen), and transcript levels were quantified on the CFX96™ Real-Time System (Bio-Rad) using iTaq™ Universal SYBR® Green Supermix (Bio-Rad). Primer sequences were as follows.
IL-2: forward 5’-CCTGGAGCAGCTGTTGATGG-3’ and reverse 5’-
CAGAACATGCCGCAGAGGTC-3’; RPL19: forward 5’-AACTCCCGTCAGCAGATCAG-3’ and reverse 5’-CATTGGCAGTACCCTTCCTC-3’; HPRT: forward 5’-
AGTCCCAGCGTCGTGATTAG-3’ and reverse 5’-CAGAGGGCCACAATGTGATG-3’.
Relative IL-2 transcript levels were determined by comparing IL-2 Ct values of WT or SLP-2 T-
K/O T cells stimulated for 4, 12, and 24 h in glucose- or galactose-containing media to that of non-stimulated T cells, then normalized to RPL19 and HPRT expression.
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ATP Quantification. WT or SLP-2 T-K/O T cells were stimulated with plate-bound α-
CD3/CD28 for 2 days in complete media, then incubated for 1 h in glucose- or galactose- containing media prior to measurement of ATP using the ATP Determination Kit (Molecular
Probes, Eugene, OR), as described [87].
Acknowledgments
We thank Dr. Julie St-Pierre, Dr. Heidi McBride, and Dr. Eric Shoubridge (McGill University) for helpful comments regarding bioenergetics and blue-native gel electrophoresis, Dr. Russell
Jones (McGill University) for access to the Seahorse XF analyzer, and members of the Madrenas laboratory for comments and criticisms. This work was funded by grants from the Canadian
Institutes of Health Research to J.M. and European Research Council to T.L. T.W. holds a HFSP long term fellowship. J.M. holds a Tier I Canada Research Chair in Human Immunology
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Figure Legends
Figure 3.1. SLP-2-deficient T cells have an impaired mitochondrial respiration profile characterized by increased uncoupled respiration and reliance on glycolysis. T cells from
WT (filled symbols/bars) or SLP-2 T-K/O (empty symbols/bars) mice were stimulated for 48 h followed by 1 h incubation in glucose (circles)- or galactose (triangles)-containing media.
Following measurement of basal oxygen consumption rate (OCR) (A), the mitochondrial drugs oligomycin, rotenone and antimycin A were sequentially added to determine total mitochondrial
OCR (B), as well as mitochondrial OCR coupled (C) or uncoupled (D) to ATP synthase activity.
(E) ATP levels were quantified in whole-cell lysates of WT or SLP-2 T-K/O T cells treated as previously indicated (n = 3). (F) Mitochondrial membrane potential was measured by flow cytometry in T cells stimulated for 48 h prior to 24 h incubation in glucose- or galactose-only conditions using TMRM in non-quench mode (FCCP: positive control) (n = 4). (G) Basal extracellular acidification rate (ECAR), a measure of glycolytic rate, was measured in T cells concurrently with OCR from (A). (H) Reactive oxygen species levels were determined in cells treated as in (F) by measuring DCFDA geometric mean fluorescence intensity (Geo. MFI) (n =
4). (A-D,G) Dot plots depict the mean OCR or ECAR per 106 cells of six mice per genotype performed in at least duplicate and analyzed in pairs. (F,H) Each dot represents one mouse and the mean was calculated from 4 mice per genotype and analyzed in pairs. *: p < 0.05 and **: p <
0.01 relative to WT control in glucose- or galactose-containing media, as indicated.
Figure 3.2. Reduced RCS detection and cell growth in Slp-2-/- MEFs. (A) BN-PAGE immunoblot analysis of mitochondria isolated from WT and Slp-2-/- immortalized fibroblasts. To visualize individual complexes and RCS, membranes were blotted with antibodies directed
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against complex IV (MTCO1) and complex I (NDUFA9) (left panel) or complex III (UQCRC2) and complex I (NDUFB6) (right panel). Membranes were subsequently blotted for complex II
(SDHA) and complex V (ATP5A). (B) Viable cell counts of WT (black squares) and Slp-2-/-
(white triangles) fibroblasts cultured in 25 mM glucose- or galactose-containing media were measured every 24 hours for 5 days by trypan-blue exclusion from an initial seeding of 50,000 cells. Data indicate mean ± S.D. of three independent experiments.
Figure 3.3. SLP-2-deficient T cells have defective RCS expression. Digitonin-solubilized mitochondrial lysates from WT or SLP-2 T-K/O T cells were separated using BN-PAGE under non-denaturing conditions. (A) Membranes of 1D gels were immunoblotted a maximum of two times with antibodies against subunits of each respiratory complex as follows: NDUFA9,
NDUFS3; UQCRC2, Complex III core 1; MTCO1 (complex IV subunit I); ATP synthase subunit alpha (ATP5A); PHB1; SLP-2. The stoichiometries of each RCS band are indicated and arrows point to the most apparent defects. (B) In-gel complex I activity was performed on a 1D
BN-PAGE gel. Purple colour indicates areas of complex I activity. Arrowheads indicate RCS containing decreased levels of complex I subunits (A) or activity (B; filled) or complex III subunits (A; open) when Slp-2 is deleted. First dimension gel slices were cut and loaded onto second dimension denaturing gels to compare high- (C) and low-molecular weight complexes
(D). Membranes were immunoblotted with antibodies against respiratory chain subunits (CI: complex I subunit NDUFB8; CII: complex II subunit 30 kDa; CIII: complex III subunit core 2;
CIV: complex IV subunit I, CV: ATP synthase subunit alpha), then PHB1 and SLP-2. Red boxes indicate RCS containing decreased levels of complex I, III, IV, and V subunits. Data are representative of three (A, C-D) or two (B) independent experiments.
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Figure 3.4. Formation of individual respiratory chain complexes is not altered by Slp-2 deletion. Dodecyl maltoside-solubilized mitochondrial lysates from WT or SLP-2 T-K/O T cells were separated using BN-PAGE under non-denaturing conditions. Membranes were immunoblotted with (A) a cocktail of antibodies against respiratory chain subunits (CI: complex
I subunit NDUFB8; CII: complex II subunit 30 kDa; CIII: complex III subunit core 2; CIV: complex IV subunit I, CV: ATP synthase subunit alpha), then (B) PHB1 and (C) SLP-2.
Figure 3.5. Deletion of Slp-2 results in delayed T cell proliferation when glycolysis is restricted. WT (black bars) and SLP-2 T-K/O T cells (white bars) were resting or stimulated in glucose- or galactose-containing media with or without oligomycin for the indicated times. (A)
Expression of CD25 and CD62L were measured after 0 and 24 hours by flow cytometry (n = 3) and data are represented as fold changes in geometric MFI relative to resting T cells at 0 h.
Viability was concurrently measured and was normalized to the glucose-only condition. (B) T cell proliferation was assessed by measurement of CFSE dilution on days 0, 2, 3, 4, and 5 post- stimulation by flow cytometry. Plots depict representative WT and SLP-2 T-K/O T cell distributions (n = 3). (C) CFSE distribution of WT and SLP-2 T-K/O T cells on day 3 grown in galactose-containing media is expanded from B, and three proliferative subsets (hi, mid, and lo) are indicated. (D) The number of proliferation cycles of T cells stimulated in galactose- containing media was calculated by taking log1/2 of the relative CFSE geometric MFI normalized to day 0. (E) Cell viability of CFSE-stained cells is measured at day 3 by flow cytometry. (F)
Stimulated cells cultured in glucose- or galactose-containing media were analyzed for cellular
Ki67 expression by flow cytometry. Data are reported as mean +/- SEM. *: p < 0.05 and ***: p <
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0.001. N.D.: not detected. N.S.: not significant. a: p < 0.001 relative to WT T cells in glucose- containing media; b: p < 0.001 relative to SLP-2 T-K/O T cells in glucose-containing media.
Figure 3.6. Deletion of Slp-2 results in defective post-transcriptional IL-2 production when glycolysis is restricted. WT and SLP-2 T-K/O T cells were stimulated in glucose- or galactose- containing media for the indicated times. (A) IL-2 protein levels were measured in supernatants by ELISA at 24 hours following stimulation of WT (black bars) or SLP-2 T-K/O T cells (white bars) in glucose- or galactose-containing media. IL-2 mRNA levels were assessed using quantitative reverse-transcriptase PCR at 4, 12 and 24 hours post-stimulation in (B) glucose or
(C) galactose-containing media (WT: closed circles; SLP-2 T-K/O: open circles). (D) WT or
SLP-2 T-K/O T cells were stimulated with α-CD3/CD28-coated beads for the indicated times and whole-cell lysates were blotted with α-phospho-IκB, α-IκB, α-SLP-2, and α-GAPDH antibodies. Numbers on left axis indicate molecular weight in kDa. (E) Calcium release was determined by flow cytometry in WT (left panel) or SLP-2 T-K/O (right panel) T cells by measuring the Fluo-3-AM:Fura Red ratio. Samples were analyzed for baseline fluorescence, then analyzed immediately following addition of 2 µg/mL ionomycin, 50 nM FCCP (positive control), or 5 mM EDTA (negative control). (F) Proliferation profiles of stimulated CFSE- stained cells supplemented with or without mrIL-2 (50 ng/mL) at t = 0 h. Pie charts indicate the percentage of cells in low (white), medium (gray), and high (black) proliferative states after 72 h.
Data are reported as mean +/- SEM. ***: p < 0.001.
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Figure 3.1
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Figure 3.2
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Figure 3.3
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Figure 3.4
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Figure 3.5
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Figure 3.6
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Preface to Chapter 4
Having established that SLP-2 regulates the formation of mitochondrial RCS likely by forming specialized membrane microdomains, we wanted to investigate whether this protein was important for another process that occurs in close association with the mitochondrial inner membrane, namely mitochondrial translation. Mitochondrial translation occurs independently of cytosolic translation and is an important cellular process since it is necessary for the translation of 13 core polypeptides of respiratory chain complexes and the ATP synthase. Mutation or defective translation of these genes can lead to mitochondrial disease. Furthermore, there is evidence that functional mitoribosomes become anchored to the inner membrane. This chapter focuses on the role of SLP-2 on mitochondrial translation.
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Chapter 4: Stomatin-like Protein 2 Deficiency Results in Impaired
Mitochondrial Translation
Panagiotis Mitsopoulos1, Orsolya Lapohos1, Woranontee Weraarpachai2,3, Hana Antonicka2, Yu-
Han Chang1, Joaquín Madrenas1*
From
1 Microbiome and Disease Tolerance Centre, Department of Microbiology and Immunology,
McGill University, Montreal, Quebec, Canada, and Los Angeles Biomedical Research Institute at Harbor-UCLA Medical Center, Torrance, California, United States of America. 2 Department of Human Genetics and Montreal Neurological Institute, McGill University, Montreal, Quebec,
Canada. 3 Department of Biochemistry, Faculty of Medicine, Chiang Mai University, Chiang
Mai, Thailand
This chapter contains the following published manuscript:
Mitsopoulos P, Lapohos O, Weraarpachai W, Antonicka H, Chang YH, Madrenas J. Stomatin- like protein 2 deficiency results in impaired mitochondrial translation. PLoS One. 2017; 12: e0179967.
Copyright © 2017. Public Library of Science
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Abstract
Mitochondria translate the RNAs for 13 core polypeptides of respiratory chain and ATP synthase complexes that are essential for the assembly and function of these complexes. This process occurs in close proximity to the mitochondrial inner membrane. However, the mechanisms and molecular machinery involved in mitochondrial translation are not fully understood, and defects in this process can result in severe diseases. Stomatin-like protein (SLP)-2 is a mainly mitochondrial protein that forms cardiolipin- and prohibitin-enriched microdomains in the mitochondrial inner membrane that are important for the formation of respiratory supercomplexes and their function. Given this regulatory role of SLP-2 in processes closely associated with the mitochondrial inner membrane, we hypothesized that the function of SLP-2 would have an impact on mitochondrial translation. 35S-Methionine/cysteine pulse labeling of resting or activated T cells from T cell-specific Slp-2 knockout mice showed a significant impairment in the production of several mitochondrial DNA-encoded polypeptides following T cell activation, including Cytb, COXI, COXII, COXIII, and ATP6. Measurement of mitochondrial DNA stability and mitochondrial transcription revealed that this impairment was at the post-transcriptional level. Examination of mitochondrial ribosome assembly showed that
SLP-2 migrated in sucrose-density gradients similarly to the large ribosomal subunit but that its deletion at the genetic level did not affect mitochondrial ribosome assembly. Functionally, the impairment in mitochondrial translation correlated with decreased interleukin-2 production in activated T cells. Altogether, these data show that SLP-2 acts as a general regulator of mitochondrial translation.
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Introduction
Mitochondria are essential for the function of most mammalian cells. These organelles are vital for a host of cellular processes and are typically the chief source of cellular energy production due to their ability to perform oxidative phosphorylation (OXPHOS) [250]. Though most genes encoding mitochondrial proteins are located in the nucleus, mitochondrial (mt) DNA encodes 37 genes of which 13 code for essential polypeptide subunits of respiratory chain complexes I, III,
IV, and V, while the remaining two ribosomal (r) RNA and 22 transfer (t) RNA genes are necessary for mitochondrial translation [182, 251]. Defects in the processes leading to expression of mitochondria-encoded polypeptides, including mtDNA maintenance, transcription, and translation, can result in improper assembly and function of mitochondrial respiratory chain complexes, a feature common to a heterogeneous set of severe, often fatal, mitochondrial diseases [252].
Mitochondrial genes are transcribed and translated in the matrix by processes involving unique molecular machinery that is separate and distinct from their nuclear and cytoplasmic counterparts. With regards to translation, mitochondrial ribosomes (mitoribosomes) are made of components generated from both the nuclear and mitochondrial genomes [253]. The assembly of these components is hypothesized to occur in two mitochondrial subcompartments in two steps: early in nucleoids, which are centers of mtDNA maintenance, replication and transcription; and later in RNA granules, locations where post-transcriptional RNA processing and maturation occur [206, 207]. Furthermore, it is known that mitochondrial translation occurs in close proximity to the mitochondrial inner membrane [198, 199]. Indeed, a biochemical study showed that nearly half of mammalian mitoribosomes interact with the mitochondrial inner membrane
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[200], suggesting that this process is mediated in part by mitoribosome interaction with integral or membrane-bound proteins.
Recent evidence in yeast mitochondria indicates that the mitoribosome binds to membrane-associated proteins [254] and becomes anchored to the inner membrane during the translation of nascent polypeptides [205]. A similar process may occur during mammalian mitochondrial translation such that the 13 extremely hydrophobic core proteins of the respiratory chain are available for immediate membrane insertion during a coordinated process with nuclear- encoded proteins in which assembly of the respiratory chain complexes occurs. In this context, the local microenvironment of the mitochondrial inner membrane including its lipid and protein composition may be critical for the regulation of this process.
We have previously shown that stomatin-like protein (SLP)-2, a mainly mitochondrial protein of the SPFH family of proteins, binds to cardiolipin and functions to form specialized membrane microdomains involving cardiolipin and prohibitins in the mitochondrial inner membrane that are important for the activities of certain respiratory chain complexes [86, 87], and the formation of respiratory chain supercomplexes [255]. Given this regulatory role of SLP-
2, it is possible that its function may regulate other processes that depend on compartmentalization of the mitochondrial inner membrane, including translation. Here, we show that SLP-2 is required for de novo mitochondrial protein synthesis upon T cell activation in mice, and that this function is not mediated at the level of mtDNA stability or mitochondrial transcription. Furthermore, we show that SLP-2 migrates in sucrose-density gradients similarly to mitoribosome complexes but that its deletion at the genetic level does not alter mitoribosome assembly. Finally, the impairment in mitochondrial translation correlates with decreased T cell
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activation. These data identify SLP-2 as a key regulator of mitochondrial translation during T cell activation, a function that may extend to other tissues or cell-types.
Results
De novo synthesis of mitochondria-encoded polypeptides is dependent on SLP-2 in activated T cells. To investigate whether SLP-2 functions to directly or indirectly regulate mitochondrial translation, we measured de novo mitochondrial protein synthesis in resting or stimulated T cells from wild type (WT) or T cell-specific Slp-2 knockout (SLP-2 T-KO) mice by detection of 35S- methionine/cysteine incorporation into nascent mitochondrial polypeptides (Fig 4.1A). We found that in WT T cells, the de novo level of every mitochondria-encoded polypeptide was lower during resting conditions than upon activation (48 hours post-stimulation), with 11 of 13 polypeptides being significantly less abundant (Fig 4.1B). Under resting conditions, deletion of
Slp-2 had no effect on mitochondrial translation compared to WT control cells. However, upon stimulation, SLP-2-deficient T cells failed to upregulate the translation of many mitochondria- encoded polypeptides to the same extent as stimulated WT T cells, with de novo COXI, Cytb,
COXII, COXIII, and ATP6 protein levels being significantly lower than in activated WT T cells.
The finding that multiple mitochondria-encoded polypeptides failed to be upregulated in SLP-2- deficient T cells following T cell activation suggested a general defect in mitochondrial translation in the absence of SLP-2.
The impairment of mitochondria-encoded polypeptide synthesis in SLP-2-deficient T cells occurs post-transcriptionally. General defects in the translation of mitochondria-encoded genes may be secondary to a variety of factors, including poor maintenance of mtDNA, a defect in
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mitochondrial transcription, or improper assembly of the mitoribosome [251]. To narrow down these possibilities, we first assessed mtDNA stability by measuring its abundance in WT or SLP-
2-deficient T cells under resting conditions or during T cell activation by quantitative PCR.
Given our previous finding that overexpression of SLP-2 in human T cells increased mtDNA content [76, 86], it was possible that a lack of SLP-2 in mouse T cells would have a negative impact on mtDNA maintenance. We found that both WT and SLP-2-deficient T cells had significantly increased mtDNA content upon T cell activation after 24 and 40 hours when compared to their respective unstimulated controls (Fig 4.2A). There were no differences in mtDNA levels between WT and SLP-2-deficient T cells at any time studied, indicating that the mitochondrial translation defect in SLP-2-deficient T cells was not related to mtDNA maintenance.
Peroxisome proliferator-activated receptor γ coactivator-1α (PGC-1α) is a master regulator of mitochondrial biogenesis and metabolism [256, 257]. PGC-1α expression is increased in situations of cellular energy stress and can be regulated by various post-translational modifications including phosphorylation of multiple residues [256, 257]. We have previously reported that SLP-2 overexpression increased PGC-1α gene and protein expression in human T cells, and that SLP-2 was upregulated prior to PGC-1α upregulation in control human peripheral blood mononuclear cells following stimulation [86]. To determine whether the disparity in mitochondrial translation between activated WT and SLP-2 T-KO T cells was due to impaired mitochondrial biogenesis in SLP-2-deficient T cells, we measured the gene and protein expression of PGC-1α under resting conditions and upon T cell activation. We found that while
PGC-1α mRNA levels decreased following T cell activation (Fig 4.2B), total PGC-1α protein levels increased under similar conditions, as did post-translationally-modified PGC-1α (Fig
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4.2C) that may reflect active phosphorylated forms of the protein [258]. SLP-2 and PHB1 protein levels were also substantially increased following T cell activation, corroborating previous findings [86]. However, SLP-2 deficiency did not affect PGC-1α abundance at the gene or protein levels, indicating that mitochondrial biogenesis was not a contributing factor to the defective mitochondrial translation observed in SLP-2-deficient T cells upon activation.
Since there were no apparent defects in mtDNA stability or PGC-1α expression in resting or activated SLP-2-deficient T cells, we next examined whether mitochondrial transcription was occurring normally under these conditions. To test this, we performed quantitative reverse- transcriptase (qRT)-PCR to measure transcript levels of the genes corresponding to four of the most significantly decreased polypeptides, ND2, CYTB, COX2, and ATP6, as shown in Fig 4.1.
Additionally, it should be noted that one gene from each respiratory chain complex is represented in this selection. We found that the transcript levels were equally abundant for each gene both prior to and following T cell activation, and that deletion of Slp-2 did not affect the abundance of these transcripts compared to WT T cells over time (Figs 4.3A-D). Furthermore, the levels of 12S rRNA, an indicator of mitoribosome abundance, were not significantly affected by Slp-2 deletion (Fig 4.3E). These results indicated that the mitochondrial translation defect we observed in activated SLP-2-deficient T cells was post-transcriptional.
SLP-2 migrates in sucrose-density gradients similarly to mitoribosome complexes but does not affect their assembly. The assembly and stability of the mitoribosome is critical for mitochondrial translation to occur. Mammalian mitoribosomes are composed of a 28S small and
39S large subunit to form a 55S monosome [194]. The small subunit is composed of 12S rRNA and approximately 30 mitoribosomal proteins (MRPs), whereas the large subunit is composed of
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16S rRNA and approximately 53 MRPs [251]. To assess mitoribosome assembly in the absence of SLP-2, we separated mitochondrial lysates of resting or stimulated WT or SLP-2 T-KO T cells based on size by sucrose density sedimentation, then collected and resolved 14 equal fractions by
SDS-PAGE. Immunoblots of protein components of the small (e.g., MRPS15, MRPS22) or large
(e.g., MRPL14, MRPL44) mitoribosomal subunits revealed that under resting conditions, the amounts of the small and large mitoribosomal subunits, as well as amounts of fully assembled monosomes were not different in SLP-2-deficient T cells compared to WT T cells (Fig 4.4A).
Stimulated SLP-2 T-KO T cells exhibited a similar lack of differences in mitoribosome assembly as compared to WT T cells under the same conditions (Fig 4.4B). Interestingly, although some
SLP-2 remained in less dense fractions at the top of the gradient, this protein mainly sedimented across fractions that contained mitoribosomal subunits in both resting and stimulated conditions.
In fact, upon stimulation the sedimentation profile of SLP-2 shifted into denser fractions and was more abundant in those containing 55S monosomes compared to resting cells. The overlap in these sedimentation profiles indicates that we cannot exclude the possibility of a direct or indirect interaction between mitoribosomes and SLP-2. Immunoblotting of PHB2 revealed that its sedimentation profile did not match that of the mitoribosomes indicating that they are not directly associated. Altogether, these data document an important post-transcriptional function for SLP-2 in the production of mitochondria-encoded polypeptides.
The impaired synthesis of mitochondria-encoded polypeptides in the absence of SLP-2 correlates with decreased T cell activation. To determine whether the defect in mitochondrial translation occurring in activated SLP-2-deficient T cells was functionally relevant, we measured interleukin (IL)-2 production in WT and SLP-2 T-KO T cells stimulated through the antigen
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receptor. T cell activation is a vital process in the adaptive immune response that is required for
T cell differentiation and proliferation [121]. In accordance with our previous findings [87, 255], we found that T cells lacking SLP-2 had significantly decreased IL-2 production compared to their WT counterparts (Fig 4.5). These data indicate that the defect in mitochondrial translation occurring in the absence of SLP-2 has significant consequences on the function of T cells.
Discussion
The mechanisms and molecular machinery required for the translation of mitochondria-encoded genes are not fully defined. Our findings identify SLP-2 as a vital protein for efficient mitochondrial translation during the metabolically demanding conditions accompanying T cell activation. Indeed, our data indicate that SLP-2 is important as a general regulator of mitochondrial translation since the expression of multiple polypeptides, namely Cytb, COXI,
COXII, COXIII, and ATP6, is significantly affected by its absence. This defect in mitochondrial translation correlates with a functional impairment in T cell activation. Though the mechanism underlying this function of SLP-2 remains uncertain, the data presented here narrow the focus to a post-transcriptional role.
Deletion of Slp-2 did not affect mitoribosome assembly, although we found that SLP-2 migrated in sucrose-density gradients similarly to the large mitoribosomal subunit and to a lesser extent the assembled monosome. This finding is corroborated by a study showing SLP-2 co- migration with the 39S mitoribosome complex by complexome profiling [259]. More importantly, compared to resting conditions our data showed that SLP-2 migration increased in fractions containing the monosome during T cell activation, correlating with impaired translation efficiency of mitochondria-encoded polypeptides in SLP-2-deficient cells. Furthermore, deletion
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of Slp-2 did not affect mitoribosome abundance. These findings indicate that SLP-2 is either directly or indirectly involved in the function of the mitoribosome and affects its efficiency, but by itself it is not essential for its assembly since the mitoribosome migration profiles were unchanged upon Slp-2 deletion.
Mitochondrial translation is closely associated with the mitochondrial inner membrane, but its functional role in this process is poorly understood. Since SLP-2 forms specialized prohibitin- and cardiolipin-enriched microdomains in the mitochondrial inner membrane that are associated with the activities of certain respiratory complexes and the formation of respiratory chain supercomplexes, it is likely that SLP-2 may regulate mitochondrial translation by controlling the membrane composition at sites where translation occurs. We examined the migration profile of PHB2 with respect to mitoribosome complexes and found that its sedimentation profile on the sucrose gradient did not match that of mitoribosomes. However, a small amount of PHB2 was present in the same fractions as the small mitoribosomal subunit, in agreement with previous studies reporting that prohibitins contributed to mitochondrial translation [260] and co-migrated with the 28S subunit [259]. Taken together, the sucrose gradient sedimentation pattern suggests a weak or indirect association of prohibitins with the mitoribosome. Interestingly however, though the absence of SLP-2 impaired mitochondrial translation in activated T cells, both mitoribosome assembly and the migration of PHB2 were unchanged relative to activated WT T cells. This implies that SLP-2 is not by itself responsible for monosome assembly or the potential association of prohibitins with the mitoribosome.
Finally, we found that although T cells lacking SLP-2 were able to produce each mitochondria-encoded polypeptide, efficient mitochondrial translation was dependent on SLP-2 only following T cell activation. Initially this seemed counter-intuitive since OXPHOS is the
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predominant form of energy production in resting T cells and is supplanted by aerobic glycolysis following T cell activation [261]. However, not only is OXPHOS still engaged in activated T cells [255], it is significantly increased [262] and is required for the expression of early activation markers and for optimal T cell proliferation [128, 255]. Therefore, it is possible that resting T cells lacking SLP-2 had the opportunity to produce sufficient amounts of mitochondria- encoded polypeptides since they were in a steady state with relatively low metabolic demands.
However, activated T cells undergo rapid growth and become metabolically very active to support their proliferation and to exert their immune function [261]. We see this reflected in the significantly increased abundance of each mitochondria-encoded polypeptide in WT T cells during the transition from resting to activated states. However, SLP-2-deficient T cells did not have the capacity to increase mitochondrial translation, highlighting the importance of this protein. Furthermore, this impairment in mitochondrial translation may be a contributing factor to the defective functional responses of SLP-2-deficient T cells observed here and previously, which include decreased IL-2 production and delayed proliferation [255].
In summary, we have identified SLP-2 as a key protein for general mitochondrial translation in activated T cells. Although the mechanism behind this function requires further study, we have shown that SLP-2 migrates in sucrose-density gradients similarly to mitoribosome complexes and hypothesize that it may exert its effect through the compartmentalization of the mitochondrial inner membrane at sites where translation occurs.
Finally, we propose that this function may extend to other tissues or cell types, particularly those with great metabolic demands.
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Materials and Methods
Mice. We generated T cell-specific Slp-2 knockout mice in the C57BL/6N Tac background, as described previously [87]. Briefly, Slp-2lox/wt mice were crossed with CD4-Cre mice (Taconic
Farms, Hudson, NY) to generate Slp-2lox/lox/Cre+ (SLP-2 T-KO) or Slp-2lox/lox/Cre- (WT) mice.
Mice were maintained in the animal facility at McGill University with approval of the
Comparative Medicine and Animal Resources Centre in accordance with the Canadian Council on Animal Care Guidelines.
Cell Culture. T cells were isolated from spleens of WT or SLP-2 T-KO mice. Briefly, spleens were homogenized using physical disruption in cell culture media to obtain a single-cell suspension that was subsequently passed through a 40 µm mesh filter. Red blood cell lysis buffer
(144 mM ammonium chloride, 17 mM Tris, pH 7.2) was added for 2 min on ice, then cells were washed three times with cell culture media. T cells were isolated from these splenocytes by negative selection using the EasySep™ Mouse T cell Isolation kit (Stemcell Technologies,
Vancouver, Canada). Cells were resting or stimulated with 5 µg/mL anti-CD3ε and 2 µg/mL anti-CD28 plate-bound antibodies (eBioscience, San Diego, CA), and cultured in RPMI supplemented with 10% fetal bovine serum, 1 mM sodium pyruvate, 2 mM L-glutamine, 10 mM
HEPES, 0.1% β-mercaptoethanol, and penicillin/streptomycin, unless otherwise indicated. Cells were incubated at 37°C in a humidified atmosphere of 5% CO2.
Mitochondrial translation. Mitochondrial translation was performed as described [263]. Briefly,
T cells from WT or SLP-2 T-KO mice were resting or stimulated with anti-CD3/CD28 for 48 h, then washed with PBS and incubated for 30 minutes in labeling media (i.e., methionine/cysteine- free RPMI (Sigma-Aldrich, St. Louis, MO), 10% dialyzed fetal bovine serum (Gibco, Carlsbad,
CA), 1 mM sodium pyruvate, 2 mM L-glutamine, 10 mM HEPES, and penicillin/streptomycin).
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Emetine, a cytoplasmic translation inhibitor, was added to a final concentration of 100 µg/mL.
Following incubation for 5 minutes, cells were pulse-labeled with 200 µCi/mL of 35S- methionine/cysteine mix (Perkin Elmer, Waltham, MA) and incubated for 1 hour, then media was replaced with R10 + 10% FBS media for 10 minutes. After three washes, cells were resuspended in PBS and frozen at -80°C. Protein levels were measured using the Micro BCA kit
(Thermo Fisher Scientific, Waltham, MA). Samples were suspended in gel loading buffer (93 mM Tris-HCl, 7.5% glycerol, 1% SDS, 0.25 mg/mL bromophenol blue, 3% β-mercaptoethanol, pH 6.7), sonicated at 60% for 8 seconds, then 50 µg of each sample was loaded onto a 12 - 20% gradient polyacrylamide gel and electrophoresed at 8 mA for 16 hours. Gels were dried at 60°C under vacuum for 1 hour using an SGD2000 Slab Gel Dryer (Thermo Fisher Scientific), then exposed to a phosphoimager cassette for 3 days. Labeled translation products were detected by direct autoradiography using a Storm 840 Gel and Blot Imaging System and densitometric analysis was performed using ImageQuant TL software (GE Lifesciences, Buckinghamshire,
England).
Mitoribosome assembly. All sample processing was performed on ice unless otherwise indicated.
Mitochondria were isolated as described [247], with minor modification. Briefly, resting or anti-
CD3/CD28-stimulated WT or SLP-2 T-KO T cells were suspended in isolation buffer (320 mM sucrose, 10 mM Tris-HCl, EDTA-free complete protease inhibitor cocktail (Roche, Basel,
Switzerland), pH 7.4) and mitochondria were isolated using 50 strokes of a tight-fitting Dounce homogenizer. Cellular debris were pelleted (600 × g, 10 min, 4°C) and discarded. The mitochondria-containing supernatant fraction was then centrifuged (10,000 × g, 10 min, 4°C), then mitochondria were washed with isolation buffer and resuspended at 10 mg/mL in lysis buffer (1% Triton X-100, 260 mM sucrose, 100 mM KCl, 20 mM MgCl2, 10 mM Tris-HCl, 5
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mM β-mercaptoethanol, EDTA-free complete protease inhibitor cocktail (Roche), pH 7.5) for 20 minutes. Following centrifugation at 9300 × g for 45 min at 4°C, 272 µg of mitochondrial lysates were layered onto cold, 1 mL, 10 - 30% discontinuous sucrose gradients, then centrifuged at
100,000 × g for 2 h at 4°C using an SW 60 Ti rotor on an Optima LE-80K ultracentrifuge
(Beckman Coulter, Indianapolis, IN). Sucrose gradient buffer consisted of lysis buffer lacking
Triton X-100. Fourteen fractions of equal volume were collected starting from the top and total protein content was precipitated by addition of 20% trichloroacetic acid. Following three washes with acetone, precipitated proteins were dissolved at room temperature in sample buffer (2%
SDS, 2% β-mercaptoethanol, 62.5 mM Tris, pH 6.8, 10% glycerol, 0.5% bromophenol blue), then boiled and resolved by SDS-PAGE. Proteins were transferred to PVDF membranes and then blotted using anti-MRPL14 (Sigma-Aldrich), anti-MRPL44, anti-MRPS15, anti-MRPS22, anti-
SLP-2 (Proteintech Group, Rosemont, IL), and anti-PHB2 (Santa Cruz Biotechnology, Santa
Cruz, CA) antibodies. Cells from 4 mice per group were pooled to achieve sufficient protein content for detection in each experiment.
Quantitative PCR. For quantification of mitochondrial mRNA, total cellular RNA was isolated from resting or anti-CD3/CD28-stimulated WT or SLP-2 T-KO T cells at the indicated times using the RNeasy® Plus Mini kit (Qiagen, Mississauga, ON, Canada). RNA was quantified using
NanoDrop (Thermo Scientific, Wilmington, DE), and cDNA was synthesized from 88.3 ng of total RNA using the Omniscript® Reverse Transcription Kit (Qiagen). Transcript levels were quantified on the CFX96™ Real-Time System using iTaq™ Universal SYBR® Green Supermix
(Bio-Rad, Hercules, CA). Primers: ND2 (Fwd.: 5ʹ-CACCCTTGCCATCATCTAC-3ʹ, Rev.: 5ʹ-
CTGAATTCCAGGCCTACTC-3ʹ); Cytb (Fwd.: 5ʹ-GTAGACAAAGCCACCTTGAC -3ʹ, Rev.:
5ʹ-CCTGTTGGGTTGTTTGATCC-3ʹ); COX2 (Fwd.: 5ʹ-CCGAGTCGTTCTGCCAATAG-3ʹ,
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Rev.: 5ʹ-GATTTAGTCGGCCTGGGATG-3ʹ); ATP6 (Fwd.: 5ʹ-
ATTAGCCCACCAACAGCTAC-3ʹ, Rev.: 5ʹ-GGCTTACTAGGAGGGTGAATAC-3ʹ); 12S rRNA (Fwd.: 5ʹ- CTGCTCGCCAGAACACTACG -3ʹ, Rev.: 5ʹ-
TGAGCAAGAGGTGGTGAGGT -3ʹ).
For mtDNA quantification, WT or SLP-2 T-KO T cells were stimulated for the indicated times with Mouse T Activator Dynabeads (Life Technologies, Grand Island, NY), then washed with PBS and DNA was isolated using the GeneJET Genomic DNA Purification kit and normalized using NanoDrop (Thermo Scientific). The mtDNA primers used (Fwd.: 5ʹ-
CTAGAAACCCCGAAACCAAA-3ʹ, Rev.: 5ʹ-CCAGCTATCACCAAGCTCGT-3ʹ) were shown to provide accurate quantification of mtDNA in mouse tissues without co-amplification of nuclear mitochondrial insertion sequences, a confounding factor of accurate mtDNA quantification [264].
Immunoblotting. For PGC-1α detection, WT or SLP-2 T-KO T cells were stimulated with Mouse
T Activator Dynabeads (Life Technologies) for the indicated times, then washed with ice-cold
PBS containing 400 µM sodium orthovanadate and resuspended in lysis buffer containing 1%
Triton X-100 with phosphatase and protease inhibitors for 45 min on ice, as described [87].
Lysates were resolved by SDS PAGE and transferred to PVDF membranes, then immunoblotted with antibodies against PGC-1α, PHB1 (Santa Cruz Biotechnology), SLP-2 (Proteintech Group), and β-actin (Cell Signaling Technology, Danvers, MA). Densitometric analysis was performed using FluorChem software (Alpha Innotech, San Jose, CA).
Enzyme-linked immunosorbent assay (ELISA). T cells were isolated from spleens of WT or SLP-
2 T-KO mice and stimulated with 5 µg/mL anti-CD3ε and 2 µg/mL anti-CD28 for 18 h.
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Secretion of IL-2 was measured by ELISA using the mouse IL-2 ELISA kit (eBioscience) according to the manufacturer's instructions.
Acknowledgments
We thank the members of the Madrenas laboratory and Drs. Eric A. Shoubridge, Heidi McBride and Amit Bar-Or for helpful comments and criticisms.
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Figure Legends
Figure 4.1. Mitochondrial translation is impaired in SLP-2-deficient T cells upon stimulation. Resting or anti-CD3/CD28-stimulated T cells isolated from WT or SLP-2 T-KO mice were cultured with 35S-methionine/cysteine for 1 h in the presence of the cytoplasmic translation inhibitor, emetine. Cell lysates were resolved by SDS-PAGE and 35S- methionine/cysteine incorporation into nascent mitochondria-encoded polypeptides was measured by direct autoradiography. (A) Representative 35S-methionine/cysteine incorporation data are shown. Bands corresponding to mitochondrial polypeptides are denoted. (B)
Densitometric analysis of 35S-methionine/cysteine incorporation from multiple experiments was performed with all values relative to the WT Stimulated group, which was assigned a value of 1
(represented by dotted line). Values below dotted line indicate decreased densitometry relative to
WT Stimulated group. N = 4 (resting) or 7 (stimulated) mice per group. Statistical significance was calculated using Student's t test. *: p < 0.05; **: p < 0.01; ***: p < 0.001 compared to WT
Stimulated group.
Figure 4.2. Mitochondrial DNA stability and PGC-1α expression are not affected by deletion of Slp-2 in T cells. WT and SLP-2 T-KO T cells were resting or stimulated with anti-
CD3/CD28 for the indicated times, then total cellular DNA (A), RNA (B) or protein (C) content was isolated. (A) Levels of mtDNA were measured by qPCR and normalized to Rag1 DNA content. Values are relative to the 0 h WT condition. N = 3 mice per group. Statistical significance was calculated using two-way ANOVA. ***: p < 0.001 compared to respective 0 h
WT or T-KO control. (B) PGC-1α gene expression was measured by qRT-PCR and normalized to RPL19, B2M, and HPRT gene expression. Values are relative to the 0 h WT condition. N = 4
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mice per group. Statistical significance was calculated using two-way ANOVA. ***: p < 0.001 compared to respective 0 h WT or T-KO control. N.D.: Not detected. (C) Cell lysates were resolved by SDS-PAGE, transferred to membranes and immunoblotted with antibodies against
PGC-1α, SLP-2, PHB1, and β-actin. Left panel: representative blots are depicted; numbers indicate molecular weight in kDa. The upper band in the anti-PGC-1α blot shows post- translational modification of this protein that may reflect active phosphorylated PGC-1α. Right panel: densitometry was calculated relative to the 0 h WT condition for PGC-1α, SLP-2, and
PHB1 (N = 3). Statistical significance was calculated using Student's t test. *: p < 0.05 compared to 0 h WT control. N.D.: Not detected.
Figure 4.3. SLP-2 deficiency does not impair mitochondrial transcription in T cells. Total
RNA was isolated from resting (i.e., 0 h) or anti-CD3/CD28-stimulated WT (closed circles) or
SLP-2 T-KO (open circles) T cells after 4, 8, 24, or 40 h, then qRT-PCR was performed using primers for (A) Nd2, (B) Cytb, (C) Cox2, (D) Atp6, and (E) 12S rRNA. Normalization of gene expression was performed using Rpl19, B2m, and Hprt. N = 4 mice per group.
Figure 4.4. SLP-2 migrates in sucrose-density gradients similarly to mitoribosome complexes but does not affect mitoribosome assembly. Mitochondrial lysates from (A) resting or (B) anti-CD3/CD28-stimulated WT or SLP-2 T-KO T cells were layered onto 10 - 30% discontinuous sucrose gradients. Following sedimentation, 14 equal fractions were collected with fraction 1 starting from the top (10% sucrose) to fraction 14 at the bottom (30% sucrose) of the column. The total protein content of each fraction was resolved by SDS-PAGE then transferred to membranes that were blotted against anti-MRPS15, anti-MRPS22, anti-MRPL44, anti-
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MRPL14, anti-SLP-2, and anti-PHB2 antibodies. Fractions corresponding to the small (i.e., 28S) and large (i.e., 39S) mitoribosomal subunits, as well as the assembled mitoribosome (i.e., 55S) are indicated. Fraction numbers are indicated for each set of blots. Data are representative of two
(A) or three (B) independent experiments.
Figure 4.5. IL-2 production is decreased in SLP-2 T-KO T cells following activation.
Splenic T cells were isolated from WT or SLP-2 T-KO mice and stimulated with anti-
CD3/CD28. Secretion of IL-2 was measured after 18 h by ELISA. N = 5 mice per group.
Statistical significance was calculated using Student's t test. ***: p < 0.001 compared to WT group.
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Figure 4.1
138
Figure 4.2
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Figure 4.3
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Figure 4.4
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Figure 4.5
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Preface to Chapter 5
Numerous compounds, both naturally occurring and synthetic, have been found to induce a wide variety of beneficial effects including anticancer, cardioprotective, and neuroprotective properties. Interestingly, it has recently been shown that these compounds bind to PHB1 and/or
PHB2 to exert their effects, and are now collectively termed PHB ligands. It is well established that SLP-2 interacts with PHB1 and PHB2 but the role of SLP-2 in PHB ligand function has never been studied. Having already established a T cell-specific SLP-2 knockout mouse line, this provided us the unique opportunity to study the role of SLP-2 in relation to the effects of the synthetic PHB ligand derivatives FL3 and Mel6 in the context of T cell function.
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Chapter 5: Modulation of T Cell Activation by the Prohibitin Ligand
Derivatives FL3 and Mel6
Panagiotis Mitsopoulosa, Orsolya Lapohosa, Laurent Désaubry, Joaquín Madrenasa
From
aMicrobiome and Disease Tolerance Centre, Department of Microbiology and Immunology,
McGill University, Montreal, QC, Canada, and Los Angeles Biomedical Research Institute at
Harbor-UCLA Medical Center, Torrance, CA 90277, USA; bTherapeutic Innovation Laboratory,
UMR7200, CNRS/Université de Strasbourg, Illkirch, France.
Manuscript in preparation.
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Abstract
Naturally occurring and synthetic compounds have been identified to bind prohibitin (PHB)1 and/or PHB2 and exert a wide range of beneficial effects. PHBs have been implicated in the response of T cells during activation, but studies on PHB ligands in this context are limited.
Here, we examined the effects of FL3 and Mel6, synthetic derivatives of the PHB ligands rocaglaol and melanogenin, respectively, on T cell activation. We found that both FL3 and Mel6 modulated T cell activation but the effects were dependent on the route of stimulation.
Specifically, FL3 elicited increased IL-2 production in human and mouse T cells stimulated with
Staphylococcal enterotoxin E (SEE) but had no effect during α-CD3/CD28-induced T cell activation. Conversely, Mel6 impaired IL-2 production in these systems when T cells were stimulated with SEE but significantly increased T cell activation when cells were stimulated with
α-CD3/CD28. The mechanism(s) underlying these responses remains unclear, but our data suggest that the level of ERK1/2 phosphorylation may be a factor. However, the known interaction between PHB1/2 and stomatin-like protein (SLP)-2 was found to be unimportant for the action of FL3 or Mel6 based on T cell activation experiments using T cell-specific SLP-2 knockout mice and co-immunoprecipitation studies. The study of PHB ligands on T cell activation will be important for determining the function of PHBs in this process and may lead to novel therapeutic strategies.
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Introduction
Prohibitins (PHBs) are highly conserved and ubiquitously expressed proteins [13]. The two eukaryotic members of this family, PHB1 and PHB2, are highly homologous and contain an
SPFH domain shared by stomatins, flotillins, and the bacterial HflK/C proteins [13, 99, 100]. In mitochondria PHB1 and PHB2 hetero-dimerize, and these dimers are thought to associate to form large ring-like structures at the inner membrane that are important for respiratory chain function, protein metabolism, and maintenance of mitochondrial structure [7, 16, 99, 105]. In addition to mitochondria, PHBs are found in small quantities in other subcellular compartments including the nucleus and plasma membrane [14, 265], and have been associated with a diverse array of cellular functions. PHBs act as transcription factors in the nucleus to regulate the cell cycle, senescence, and tumor suppression [266], and PHB1 is a key factor in cell survival since it can suppress apoptosis by the intrinsic pathway through the Ras-Raf-MEK-Erk pathway [98,
267].
Recently, certain mitochondrial proteins have been implicated in the regulation of T cell activation, including PHBs and stomatin-like protein (SLP)-2. Cellular levels of PHB1 and
PHB2 are significantly increased upon activation through the antigen receptor [108], including at the cell surface [109], suggesting that these proteins may be important for T cell receptor- mediated signaling. Similar to PHBs, SLP-2 is a highly conserved, ubiquitously expressed protein that is upregulated in response to T cell activation [44, 74, 76, 87]. Deletion of Slp-2 results in impaired IL-2 production and CD4+ T cell responses in vivo [87]. We have shown that
SLP-2 interacts with PHBs and binds the mitochondrial phospholipid cardiolipin to form functional microdomains in the mitochondrial inner membrane that are associated with the respiratory chain activity, assembly of respiratory chain supercomplexes, and mitochondrial
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translation [86, 87, 255]. The functional interaction between PHB1, PHB2, and SLP-2 in the context of T cell activation requires further study.
Certain small molecules, both naturally occurring in plants and synthetically generated, have been shown to bind to PHBs and have wide-ranging effects. Several categories of such compounds, termed "PHB ligands," have been described including flavaglines and melanogenins. Flavaglines are a family of cyclopenta[b]benzofuran compounds isolated from
Aglaia species of medicinal plants [112]. Rocaglamide was the first of these compounds identified from A. elliptifolia in 1982 [113]. Since then, a large number of naturally occurring and synthetic flavaglines have been described, and their structures have been documented [117,
268]. Flavaglines have been studied mostly for their anticancer properties but also exhibit neuroprotective and cardioprotective effects [112, 114]. Flavaglines have been shown to bind
PHB1 and PHB2 [115], and this interaction has been attributed to many of the effects exhibited by these compounds. However, the impact of flavaglines on immune processes is not well characterized. It was shown in one study that rocaglamide and several derivatives decreased T cell activation and cytokine production, an effect that was associated with decreased NF-AT activation [119]. Furthermore, rocaglamide derivatives have been shown to inhibit NF-κB activation in Jurkat T cells [269]. It is not known whether these immunosuppressive properties are conserved among flavaglines.
The flavagline FL3 is a synthetic derivative of rocaglaol, itself a derivative of rocaglamide [117]. Similar to other flavaglines, FL3 has been shown to bind PHB1 and PHB2 by affinity purification [115], to protect against experimental colitis [116], and to exhibit anticancer
[117] and cardioprotective [118] properties. However, the immune properties of this compound have not been investigated.
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The fully synthetic small molecule melanogenin was identified in a screen for modulators of pigmentation in melanocytes. This compound was found to promote the induction of melanin through its binding of PHB1, an interaction confirmed by affinity purification [120]. The effect of melanogenin or its derivatives on T cell function has not been studied. To examine the effects of PHB ligands on T cell activation, we stimulated Jurkat or primary human T cells treated concurrently with FL3 or the melanogenin derivative Mel6. We found that FL3 increased T cell activation in response to Staphylococcal enterotoxin E (SEE) stimulation in both human and mouse T cells. In contrast, Mel6 exhibited immune-modulatory activity in human and mouse T cells depending on the stimulus, with SEE stimulation decreasing and α-CD3/CD28 increasing
IL-2 production. Lastly, we found that mouse T cells responded similarly to human T cells when activated in the presence of FL3 or Mel6, and that these compounds had no effect on SLP-
2:PHB1/2 or PHB1:PHB2 interactions. Evaluated together, it is possible that the opposing effects of FL3 and Mel6 on T cell function may reflect a differential mode of action of these compounds on PHBs.
Results
The PHB ligand derivatives FL3 and Mel6 modulate T cell activation.
The effects of the rocaglaol derivative FL3 and the melanogenin derivative Mel6 on T cell function have not been examined. In order to test the effects of these compounds on T cell activation, we first assessed the viability of Jurkat T cells to increasing concentrations of each compound for 24 hours to establish effective treatment concentrations. We found that 50 nM
FL3 and 10 µM Mel6 were optimal concentrations that did not significantly affect cell viability
(Fig. 5.1A). To investigate the effect of FL3 and Mel6 on T cell function, we next activated
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Jurkat T cells in the presence or absence of each of these compounds for 24 hours and measured
IL-2 production (Fig. 5.1B). We chose to stimulate these cells with SEE in the presence of LG2 antigen-presenting cells since Jurkat T cells elicit poor IL-2 responses to α-CD3/CD28 stimulation. Superantigens activate T cells by directly binding both the T cell receptor on T cells and the major histocompatibility complex class II on antigen-presenting cells in a way that bypasses conventional antigen processing and presentation [270]. We found that IL-2 production was significantly increased in the presence of FL3 and significantly decreased in the presence of
Mel6. To corroborate these findings using a more physiologically relevant system, we measured
IL-2 production from FL3- or Mel6-treated primary human peripheral T cells stimulated for 24 hours with either SEE in the presence of LG2 antigen-presenting cells (Fig. 5.2A), or α-
CD3/CD28-coated beads (Fig. 5.2B) to induce a conventional route of T cell activation. For direct comparison, these experiments were performed concurrently with SEE-stimulated Jurkat T cells (Fig. 5.2C). From these experiments, we found that T cells activated with SEE in the presence of FL3 elicited a consistent, significantly increased IL-2 response relative to control cells. However, FL3 had not effect on IL-2 production in response to α-CD3/CD28 stimulation in primary T cells. Perhaps more interesting was the fact that Mel6-treated primary T cells exhibited decreased IL-2 levels in response to SEE stimulation in a similar fashion to Jurkat T cells, but conversely produced significantly greater amounts of IL-2 when stimulated with α-
CD3/CD28. These data indicate that FL3 and Mel6 can differentially modulate the activation of
T cells depending on the stimulation conditions.
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The impact of FL3 or Mel6 on T cell receptor signaling.
FL3 has been shown to bind PHB1 and PHB2 by affinity purification [115], and the structural similarity of Mel6 to the PHB1-binding melanogenin [120] suggests it too may bind PHB1. To investigate the mechanism underlying the modulatory nature of FL3 and Mel6 with regards to T cell activation, we investigated early T cell signaling in the presence of each of these compounds. PBMCs or primary human T cells were pre-treated for 1 hour with FL3 or Mel6, then stimulated for 5 minutes by the addition of SEE or α-CD3/CD28-coated beads, respectively.
Immunoblots of these cell lysates indicated that there were no significant changes in levels of phosphorylated ERK1/2, PLCγ1, or Zap70 (Fig. 5.3). These data suggest that the modulatory nature of these compounds on the activation of T cells does not manifest at the level of early signaling events.
The effect of SLP-2 on PHB ligand function.
We have previously shown that SLP-2 interacts with PHB1 and PHB2 and functions to form cardiolipin-enriched microdomains in the mitochondrial inner membrane [86, 87]. These functional membrane microdomains appear to be important for mitochondrial and cellular function, since deletion of SLP-2 results in impaired respiratory chain supercomplex formation
[255], defective mitochondrial translation (Mitsopoulos et al. 2017, in preparation), and suboptimal T cell activation and function [87, 255]. Due to this association between PHBs and
SLP-2, we decided to investigate whether SLP-2 was important in mediating the modulatory effects of FL3 or Mel6 on T cell activation. To test this, splenocytes or T cells isolated from WT or SLP-2 T-KO mice were treated with FL3 or Mel6 and stimulated with SEE or α-CD3/CD28- coated beads for 24 hours (Fig. 5.4). First, we found that the IL-2 expression induced in mouse
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WT T cells in the presence of FL3 or Mel6 followed a similar pattern to that observed in human
T cells. Specifically, FL3 elicited a trend towards increased IL-2 levels in SEE-stimulated splenocytes but had no effect in α-CD3/CD28-stimulated T cells. Likewise, Mel6 significantly increased IL-2 expression of T cells in response to α-CD3/CD28 stimulation, although unlike human T cells, mouse T cells were not affected by Mel6 treatment when stimulated with SEE.
Next, we found that SLP-2 deletion resulted in significantly decreased IL-2 production in T cells in response to SEE (Fig. 5.4A) or α-CD3/CD28 (Fig. 5.4B) stimulation, corroborating our previous findings [87, 255]. However, the proportion of IL-2 production between WT and SLP-2
T-KO T cells was consistent when cells were treated with FL3 or Mel6 using either stimulus, suggesting that SLP-2 is not involved in the mechanism by which IL-2 expression is regulated by these compounds.
To further investigate the role of SLP-2 in the activities of FL3 and Mel6, we tested whether the interaction between SLP-2 and PHBs was altered in the presence of these compounds. Previously, we and others have shown that PHB1 and PHB2 interact with SLP-2
[10, 77, 86]. To test this, Jurkat T cells were treated for 2 h with FL3 or Mel6 and lysates were co-immunoprecipitated with α-SLP-2 (Fig. 5.6A), α-PHB1 (Fig. 5.6B), or α-PHB-2 (Fig. 5.6C).
We found that neither FL3 nor Mel6 affected the interaction between SLP-2 and PHB1 or PHB2, or the interaction between PHB1 and PHB2. These data indicate that SLP-2 is not important for the modulatory effects of FL3 or Mel6 on T cell activation.
Discussion
PHB ligands have been shown to exhibit a wide range of protective effects but our knowledge of these compounds on immunity is limited. Our findings identify the PHB ligand derivatives FL3
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and Mel6 as modulators of T cell activation. Indeed, these compounds differentially affect T cell activation depending on the stimulation conditions. Specifically, FL3 consistently elicited increased IL-2 production when T cells were stimulated with SEE but not α-CD3/CD28 antibodies. In contrast, Mel6 treatment resulted in decreased IL-2 levels upon SEE stimulation but elicited an increase in α-CD3/CD28-induced T cell activation. The mechanism underlying these effects requires further study but our data indicates that it does not involve SLP-2:PHB1/2 interaction.
The increase in T cell activation occurring with FL3 treatment in superantigen-stimulated cells suggests that this compound may enhance T cell responses. This finding is unexpected based on the structural similarity of FL3 with other flavaglines, since it was found in a previous study that rocaglamide and several derivatives decreased IL-2 expression of human T cells in response to α-CD3/CD28 stimulation [119]. These seemingly opposing findings may be explained by the stimulation conditions since α-CD3/CD28 stimulation in the presence of FL3 produced no significant effect on IL-2 production in any of our systems, a difference likely attributable to the structural differences between these compounds. The increased IL-2 production observed with FL3 treatment in our study occurred only in T cells stimulated with superantigen. There is evidence that PHBs may be involved in the ERK cascade since ERK signaling was inhibited by antibodies against PHB1 or PHB2 during T cell activation [109].
Indeed, our data indicates that ERK1/2 phosphorylation in PBMC may have been increased under these conditions suggesting the increase in IL-2 production was a result of enhanced TCR- mediated signaling.
Equally as interesting was the effect of Mel6 on T cell activation. In contrast to FL3,
Mel6 impaired IL-2 production in T cells stimulated with superantigen but increased IL-2 levels
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when stimulated with α-CD3/CD28. These are novel findings since neither Mel6 nor melanogenin have been tested on T cell activation. However, the mechanism remains unclear.
Our data indicate ERK1/2 phosphorylation was decreased in Mel6-treated PBMC stimulated with SEE, suggesting an impairment in TCR-mediated signaling, but ERK1/2 phosphorylation was unchanged in Mel6-treated T cells stimulated with α-CD3/CD28. This dependence on the route of T cell activation in relation to the action of PHB ligand derivatives is supported in the literature. Specifically, it has been shown that IL-2 production induced by α-CD3/CD28 antibodies in T cells were inhibited by the Salmonella Typhi virulence polysaccharide Vi by interacting with membrane PHBs and suppressing early activation events including ERK phosphorylation [271]. However, the authors show that activation of T cells by PMA/ionomycin does not affect IL-2 expression or ERK phosphorylation in this system. Additional mechanistic studies are required to understand the intricacies of PHBs and PHB ligands in the activation of T cells.
The IL-2 data presented here corroborates our previous studies showing that deletion of
Slp-2 results in significantly decreased IL-2 production following T cell activation [87, 255].
This defect occurs post-transcriptionally [87], but one remaining question was whether or not IL-
2 production was occurring at the maximal rate in activated SLP-2-deficient T cells. The finding that Mel6-treated SLP-2-deficient T cells stimulated with α-CD3/CD28 were able to significantly increase their production of IL-2 compared to control cells shows that SLP-2 T-KO
T cells are able to increase their IL-2-producing capacity. However, the inhibition of IL-2 production caused by Slp-2 deletion still occurs in the presence of Mel6 since Mel6-treated SLP-
2 T-KO T cells stimulated with α-CD3/CD28 produce significantly less IL-2 than WT T cells under the same conditions. These findings suggest that the mechanism responsible for limiting
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IL-2 production in the absence of SLP-2 is separate and distinct from that which causes enhanced IL-2 production when Mel6 is present.
Materials and Methods
Cell culture. Primary human peripheral blood mononuclear cells (PBMCs) were isolated from blood of healthy volunteers on Ficoll-Hypaque gradients. Volunteers provided informed consent in compliance with the Research Ethics Office at McGill University. Primary human T cells were isolated from PBMC preparations by negative selection using the EasySep Human T cell
Isolation kit (Stemcell Technologies, Vancouver, Canada). Primary mouse splenocytes were isolated from WT or SLP-2 T-KO mice, and primary mouse T cells were isolated from those splenocyte preparations by negative selection using the EasySep™ Mouse T cell Isolation kit
(Stemcell Technologies, Vancouver, Canada). Jurkat T cells (E6.1) were purchased from the
American Type Culture Collection (Manassas, VA), and LG2 B lymphoblastoid cells were kindly provided by Dr. Eric Long (NIAID, NIH, Rockville, MD). Human cells were cultured in
R10 media (i.e., RPMI supplemented with 10% fetal bovine serum, 1 mM sodium pyruvate, 2 mM L-glutamine, 10 mM HEPES, and penicillin/streptomycin). Mouse cells were cultured in
R10 media supplemented with 0.1% β-mercaptoethanol. Cells were incubated at 37°C in a humidified atmosphere of 5% CO2. Cells were resting or stimulated for the indicated times with
0.5 or 10 ng/mL Staphylococcal enterotoxin E in the presence of LG2 antigen-presenting cells
(1:1 ratio), or with human or mouse α-CD3/CD28-coated T-Activator Dynabeads (Life
Technologies, Grand Island, NY) at a 1:1 cell-to-bead ratio.
Mice. We previously generated T cell-specific Slp-2 knockout mice in the C57BL/6N Tac background [87]. Briefly, Slp-2lox/wt mice were crossed with CD4-Cre mice (Taconic Farms,
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Hudson, NY) to generate Slp-2lox/lox/Cre+ (SLP-2 T-KO) or Slp-2lox/lox/Cre- (WT) mice. Mice were maintained in the animal facility at McGill University with approval of the Comparative
Medicine and Animal Resources Centre in accordance with the Canadian Council on Animal
Care Guidelines.
Enzyme-linked Immunosorbent Assay. Levels of secreted human or mouse IL-2 in supernatants were measured using the appropriate human or mouse ELISA Ready-SET-Go!® kits
(eBioscience, San Diego, CA) according to the manufacturer's instructions.
Flow Cytometry. Viability was assessed using the Zombie Aqua™ Fixable Viability kit
(BioLegend, San Diego, CA). Flow cytometry was performed using an LSRFortessa flow cytometer (BD), and data were analyzed with FlowJo software (Treestar Inc., Ashland, OR).
Co-immunoprecipitation. Protein A agarose beads were coated with α-SLP-2 (Proteintech
Group, Rosemont, IL), α-PHB1, or α-PHB2 (Santa Cruz Biotechnology, Santa Cruz, CA) antibodies rotating overnight at 4°C. Unbound antibody was removed by washing and cell lysates were incubated overnight at 4°C on a rotator. Following immunoprecipitation, unbound proteins were washed and antibody-bound protein complexes were eluted by addition of sample buffer containing β-mercaptoethanol and boiling for 5 minutes. Immunoprecipitated samples were resolved by SDS-PAGE and subsequently immunoblotted using α-SLP-2, α-PHB1, or α-
PHB2 antibodies.
Immunoblotting. Following stimulation, cells were resuspended in lysis buffer containing 1%
Triton X-100 with phosphatase and protease inhibitors for 45 min on ice, as described [87].
Lysates were resolved by SDS PAGE and transferred to PVDF membranes, then immunoblotted with antibodies against phospho-ERK1/2, ERK1/2, phospho-PLCγ1, PLCγ1, phospho-p38, p38
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(Cell Signaling Technology, Danvers, MA), PHB1 (Santa Cruz Biotechnology), SLP-2
(Proteintech Group), and GAPDH (Chemicon International, Temecula, CA).
Acknowledgments
We thank the members of the Madrenas laboratory for helpful comments and criticisms. This research was supported by the Canadian Institutes of Health Research. J.M. holds a Tier I
Canada Research Chair in Human Immunology.
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Figure Legends
Figure 5.1. Activation of Jurkat T cells is increased by FL3 and decreased by Mel6 in response to superantigen. (A) Jurkat T cells were treated with control media or with increasing concentrations of FL3 (left panel) or Mel6 (right panel) and viability was measured after 24 h by flow cytometry. (B) Jurkat T cells and LG2 antigen-presenting cells were seeded at a 1:1 ratio and treated with control media, 50 nM FL3, or 10 µM Mel6 in the presence or absence of SEE
(0.5, or 10 ng/mL). Levels of secreted IL-2 were measured after 24 hours by ELISA. Data are representative of three independent experiments. ***: p < 0.001.
Figure 5.2. Mel6 activates T cells differentially depending on the stimulus. Primary human T cells were incubated for 24 h with control media, 50 nM FL3, or 10 µM Mel6 (A) in the presence of LG2 antigen-presenting cells with or without 10 ng/mL SEE, or (B) in the presence or absence of α-CD3/CD28-coated beads. (C) Jurkat T cells were concurrently incubated under the same conditions described for (A). Levels of secreted IL-2 were measured after 24 hours by ELISA.
Data are representative of three independent experiments.
Figure 5.3. Effects of FL3 or Mel6 on T cell signaling following activation. Primary human
PBMC (A) or T cells (B) were pre-treated for 1 h with control media, 50 nM FL3, or 10 µM
Mel6, then incubated for 5 min with or without the addition of 100 ng/mL SEE or α-CD3/CD28- coated beads, respectively. Cell lysates were resolved by SDS-PAGE and immunoblotting was performed with antibodies against the indicated proteins. Numbers indicate molecular weight in kDa.
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Figure 5.4. Effects of FL3 or Mel6 on IL-2 responses following activation of SLP-2-deficient
T cells. (A) Splenocytes from WT or SLP-2 T-KO mice were incubated with or without 10 ng/mL SEE in the presence of control media, 50 nM FL3, or 10 µM Mel6. Levels of secreted IL-
2 were measured by ELISA after 24 h. (B) Purified T cells from spleens of WT or SLP-2 T-KO mice were incubated with or without α-CD3/CD28-coated beads for 24 h in the presence of control media, 50 nM FL3, or 10 µM Mel6. Levels of secreted IL-2 were measured by ELISA after 24 h. Data are representative of three independent experiments.
Figure 5.5. SLP-2:PHB1/2 and PHB1:PHB2 interactions are not affected by FL3 or Mel6.
Jurkat T cells were treated with control media, 50 nM FL3, or 10 µM Mel6 for 2 h, then whole- cell lysates were prepared. A fraction of each whole-cell lysate was collected and remaining lysates were co-immunoprecipitated using (A) α-SLP-2, (B) α-PHB1, or (C) α-PHB2 antibodies.
Co-immunoprecipitations (Co-IP) and whole-cell lysates (WCL) were resolved by SDS-PAGE and immunoblotted with antibodies against the indicated proteins.
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Figure 5.1
159
Figure 5.2
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Figure 5.3
161
Figure 5.4
162
Figure 5.5
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Chapter 6: Discussion
The mitochondrial inner membrane is considered to be the most protein-rich membrane in the cell and is the site of several critical cellular processes [272]. Previous data generated by our group indicate that SLP-2 recruits PHBs to cardiolipin to form specialized microdomains in the mitochondrial inner membrane [78, 86, 87]. Here, we show that SLP-2 is necessary for respiratory chain supercomplex formation and mitochondrial translation, and compromise of these functions in SLP-2-deficient T cells is associated with impaired T cell function. The data generated here support our model of SLP-2 function in which SLP-2 regulates the formation of functional membrane microdomains necessary for key cellular processes.
The identification of novel factors that regulate the assembly and function of RCS is an area of burgeoning interest. Although various proteins have been identified to influence RCS, only one factor, SCAF1, has been identified thus far that exclusively affects the formation of
RCS, specifically those containing CIV [147, 159]. This is in contrast to other proteins that influence RCS assembly due to their importance in the formation of individual respiratory complexes [172-174]. Using Slp-2-deleted mouse T cells and MEFs, we have identified SLP-2 to be a critical player in the formation of RCS. Importantly, SLP-2 is required for the proper assembly of a large number of RCS arrangements, specifically (I-III2)1-2, I-III2-IV1-3, I-III2, and
III2-IV, and does not influence the assembly of individual respiratory chain complexes. Our findings are corroborated by a separate study linking the plant homologue of Slp-2 to RCS organization in Arabidopsis thaliana [244]. The mechanism by which SLP-2 regulates RCS formation remains unclear. We have found SLP-2 to migrate in a high-molecular weight complex by BN-PAGE as has been reported previously [81]. The composition of these high- molecular weight complexes is not known but they do migrate similarly to RCS, making it
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difficult to rule out indirect association between SLP-2 and RCS. In fact, our group has shown that SLP-2 binds cardiolipin [86], the signature mitochondrial phospholipid that is known to be an integral component of RCS formation and function since defective cardiolipin remodeling results in RCS instability and Barth syndrome [158]. It is possible that in the absence of SLP-2, cardiolipin-enriched microdomains become less stable and cause RCS dissociation. Whether
SLP-2 is associated with RCS is unclear but it is perhaps more likely that the formation of cardiolipin-enriched microdomains containing PHBs and SLP-2 are important sites of RCS formation in the mitochondrial inner membrane.
During our bioenergetic analyses of SLP-2-deficient T cells, we made a peculiar observation in regards to the spare respiratory capacity of these cells. Spare respiratory capacity is defined as the difference between the maximal and basal respiration rates [133].
Methodologically, we measured oxygen consumption rate after the addition of the protonophore
FCCP, which induces maximal respiratory activity by continuously disrupting the proton gradient across the mitochondrial inner membrane [133, 273]. This method reveals the maximal respiratory capacity, and by extension, the spare respiratory capacity of a cell. Specifically, we found that upon addition of FCCP, activated T cells lacking SLP-2 and thus lacking the ability to fully form functional RCS had a significantly greater respiration rate compared to WT T cells with normal RCS assembly (Supplementary Figure 1). This appeared to be a discrepant result since our body of work to that point indicated that SLP-2 was required for optimal mitochondrial respiration under physiological conditions. Since FCCP only elicits its desired effect in a narrow concentration range and its effect may be influenced by membrane composition [273], we performed an FCCP dose response on WT and SLP-2-deficient T cells. We found that the difference in spare respiratory capacity was maintained regardless of the FCCP concentration,
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and that the peak of FCCP-induced respiration was the same regardless of SLP-2 deficiency.
Recently, it was reported that in cells lacking functional SCAF1, a protein required for assembly of complex IV-containing RCS, total cellular respiration driven by glucose, pyruvate, and glutamine was significantly increased, whereas CIV respiration in intact cells was not affected by the presence of functional SCAF1 [147, 159]. This suggests that cells lacking the ability to form functional supercomplexes cannot regulate their respiration and would have increased oxygen consumption when the respiratory chain is induced to function at a high level. This is in line with the increased FCCP-induced respiration observed in SLP-2-deficient T cells, and coupled with our finding of increased respiration uncoupled from ATP production in these cells it suggests that functional RCS (and by extension, SLP-2) are required for efficient, ATP-generating mitochondrial respiration.
In addition to defective RCS assembly, SLP-2 deficiency also resulted in impaired mitochondrial translation during T cell activation. The fact that the translation of most or all mitochondria-encoded peptides were affected by the absence of SLP-2 points to a role for this protein as a general regulator of mitochondrial translation as opposed to being important for the translation of specific peptides. Although we showed that SLP-2 did not affect mitoribosome assembly, it is possible that the efficiency of the mitoribosome was impaired in SLP-2-deficient
T cells due to the defective formation of functional cardiolipin-enriched microdomains normally orchestrated by SLP-2. Indeed, the mitoribosome becomes anchored to the inner membrane
[205] and the location of translation is of great importance since it is thought that the newly synthesized, hydrophobic peptides are inserted into the membrane co-translationally [188, 189].
Furthermore, our data suggest that SLP-2 may indirectly interact with the mitoribosome, perhaps through the formation of these specialized membrane microdomains to create an ideal
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environment for mitoribosome function. Additional studies will be required to elucidate the mechanism of SLP-2 function on mitochondrial translation.
The defective mitochondrial function associated with SLP-2 deficiency in T cells translated into impaired T cell function. Specifically, SLP-2-deficient T cells exhibited decreased
T cell activation, and significant delays in cell cycle entry and proliferation under conditions of limiting glycolysis, effects that could not be overcome by supplementation of recombinant IL-2.
Recent advances in T cell bioenergetics have revealed that although glycolysis is upregulated upon T cell activation, it does not replace the requirement for energy production by OXPHOS.
Indeed, OXPHOS is increased significantly in activated T cells [123, 126, 132], is sufficient to support T cell activation [132], and is essential for T cell proliferation [128, 132]. Our data are in line with these findings and illustrate the functional consequences of defective mitochondrial processes associated with Slp-2 deletion.
Although it is known that SLP-2 interacts with PHBs [10, 77, 86], our findings show that
SLP-2 is not important for the function of the PHB ligand derivatives FL3 and Mel6 in modulating the activation of human and mouse T cells. Interestingly, during SEE stimulation
FL3 was able to induce IL-2 production in SLP-2-deficient T cells equal to that of WT T cells in the absence of FL3. Even more striking, during α-CD3/CD28 stimulation Mel6 was able to induce IL-2 production in SLP-2-deficient T cells to a level that far exceeded the amount generated in WT T cells lacking Mel6 treatment. These findings clearly demonstrate that SLP-2- deficient T cells possess the capacity to overcome the post-transcriptional defect inherent in these cells [87]. However, the mechanism of enhanced IL-2 production by FL3 and Mel6 as described is separate and distinct from that of the impairment in IL-2 production caused by the absence of SLP-2. This is illustrated by the fact that FL3- or Mel6-treated SLP-2-deficient T
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cells maintained an impaired IL-2 phenotype relative to similarly treated WT T cells. These findings highlight the complexities of T cell activation and will be important in developing a complete understanding of the regulation of this process.
We have previously found that conventional systemic SLP-2 knockout mice are embryonic lethal (C.D. Lemke and J. Madrenas, unpublished observation [87]). This observation was corroborated by a second report from an independent group that the loss of SLP-2 function is lethal in SLP-2 transgenic mice (Heppenstall, Seifert and Lewin, unpublished result [63]). This finding was not particularly surprising considering transgenic deletion of other mitochondrial proteins in mice results in lethality, including that of the SPFH family proteins PHB1 and PHB2
[103, 104]. However, our attempts to confirm this lethality using an alternate, non-conventional method of systemic SLP-2 deletion were unsuccessful. Specifically, Slp-2-floxed, β-actin Cre-
ERT2 mice were generated and Cre recombination was induced by a regimen of tamoxifen administration in adult mice. Deletion of Slp-2 in these mice is dependent on the bioavailability of the tamoxifen metabolite 4-hydroxytamoxifen [274] and results in variable amounts of deletion ranging from 47 to 87% depending on the tissue. After tamoxifen administration these mice were allowed to breed. Surprisingly, breeding was successful and some pups were found to have complete, systemic deletion of SLP-2 at both the gene and protein levels (Supplementary
Figure 2). Moreover, these SLP-2-/- mice exhibited no gross phenotypic abnormalities (i.e., normal lifespan, breeding capabilities, mobility, etc.) and were maintained independent of the
Cre-ERT2 transgene. It is likely that a certain proportion of the gametes of these mice underwent successful Cre recombination to delete the Slp-2 gene and gave rise to SLP-2-/- pups.
The reason for the discrepancy in the lethality of SLP-2 knockout mice is not known and requires further study. Sequencing of the regions upstream and downstream of the Slp-2 gene in
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both our conventional and Cre-lox transgenic mouse strains identified no genetic anomalies. One explanation pertains to the possible expression of the neomycin cassette that was inserted (in the opposite direction of Slp-2 transcription) by homologous recombination in place of the Slp-2 gene during the generation of our conventional SLP-2 knockout mice. It has been reported that under certain circumstances the neomycin cassette can be expressed and cause unexpected embryonic lethality in transgenic mice [275]. It is evident that further investigation is required to clarify the issue of embryonic lethality in SLP-2 knockout mice.
Taken together, the work presented here has demonstrated that SLP-2 is an essential protein for optimal mitochondrial function, and its absence translates to impaired T cell function.
Specifically, SLP-2 is required for proper RCS formation and translation of mitochondria- encoded peptides, and its deletion results in respiration increasingly uncoupled from ATP production, an increased reliance on glycolysis, and defective T cell effector responses including impaired IL-2 production and proliferation. These findings are in line with a model of SLP-2 function in which this protein orchestrates the organization of specialized, PHB- and cardiolipin- enriched membrane microdomains in the mitochondrial inner membrane important for key mitochondrial processes.
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Appendix
Supplementary Materials and Methods
Bioenergetic Analysis. Oxygen consumption rate (OCR) was measured using a Seahorse XF-24 analyzer (Seahorse Bioscience, North Billerica, MA). Following α-CD3/CD28 stimulation for 48 hours, WT and SLP-2 T-K/O T cells were suspended in Seahorse XF assay media supplemented with 2 mM L-glutamine, sodium pyruvate, and 25 mM glucose or galactose and seeded onto
XF24 V7 24-well cell culture plates coated with 50 µg/mL poly-D-lysine at 500,000 cells/well. T cells were placed at 37°C in a CO2-free atmosphere for 1 h prior to analysis. OCR was measured prior to and following the addition of FCCP (carbonyl cyanide 4-
(trifluoromethoxy)phenylhydrazone). Spare respiratory capacity was calculated as the maximal
OCR (i.e., following addition of FCCP) subtracted by the basal OCR.
Mice. Slp-2-floxed, β-actin Cre-ERT2 mice were generated by first generating the SLP-2lox/wt mouse on a C57BL/6 background [87]. These Slp-2-floxed mice were crossed with C57BL/6 mice transgenic for Cre-ERT2 under the control of the chicken β-actin promoter/enhancer and cytomegalovirus immediate-early enhancer (The Jackson Laboratory, Bar Harbor, ME). Slp-2- floxed mice lacking Cre-ERT2 were used as controls. Cre recombinase activity was induced in adult mice by administration of tamoxifen (3 mg/40 g b.w. in corn oil, i.p., once daily for up to
10 days). Breeding of tamoxifen-administered, adult Slp-2-floxed, β-actin Cre-ERT2 mice generated pups including those with a Slp-2-/- genotype.
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Polymerase chain reaction. Total genomic DNA was isolated using the Qiagen DNeasy Blood and Tissue kit (Qiagen, Mississauga, Canada) according to the manufacturer's instructions.
Polymerase chain reaction was performed to detect a portion of the Slp-2 gene using primers corresponding to genomic sequences upstream of and within the Slp-2 gene: Fwd: 5ʹ-
ACTTCCACCCTTCAGTCCAGGTCG-3ʹ; Rev: 5ʹ-ACTTGGATTCTGTGAAAGCAGACAC-
3ʹ; or to detect full-length Slp-2 using primers corresponding to genomic sequences flanking the gene: Fwd: 5ʹ-GCGGAGTGGACCAGCTGAAGGA-3ʹ Rev: 5ʹ-CAATGGGCATCTGAAT
GGGTCTC-3ʹ; The former produces a 406 bp product if the Slp-2 gene is present or no product if the Slp-2 gene is absent, and the latter produces a 534 bp product if the Slp-2 gene is absent
(i.e., this product is an amplification of the Slp-2-flanking regions surrounding the loxP site that remains following Cre recombination). Samples were resolved by DNA electrophoresis in a 1% agarose gel and visualized by ethidium bromide staining.
Immunoblotting. Freshly isolated tissues were homogenized in lysis buffer containing 1% Triton
X-100, 150 mM NaCl, 10 mM Tris (pH 7.6), 4 mM EDTA, 1 mM sodium orthovanadate, 10
µg/mL aprotinin, 10 µg/mL leupeptin, and 25 µM p-nitrophenyl-pʹ-guanidinobenzoate for 30 minutes on ice. Following centrifugation (10,700 × g, 10 min, 4°C), lysates were mixed with sample buffer containing β-mercaptoethanol, boiled for 5 minutes, then resolved by SDS PAGE and transferred to PVDF membranes. Immunoblotting was performed with antibodies against
SLP-2 (Proteintech Group), and GAPDH (Chemicon International, Temecula, CA).
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Supplementary Figure Legends
Supplementary Figure 1. SLP-2-deficiency results in increased spare respiratory capacity in activated T cells. T cells from WT (filled symbols) or SLP-2 T-KO (empty symbols) mice were stimulated for 48 h with α-CD3/CD28 and assessed for spare respiratory capacity.
Following measurement of basal oxygen consumption rate (OCR), increasing concentrations of
FCCP were added to induce maximal respiration. Spare respiratory capacity was calculated by subtracting basal OCR from maximal OCR. Data are reported as mean ± SEM of n = 3 mice per group. *: p < 0.05 and **: p < 0.01 relative to WT control.
Supplementary Figure 2. Slp-2 is systemically deleted in viable Slp-2-/- mice. Genomic DNA was isolated from Slp-2+/+, Slp-2+/-, and Slp-2-/- mice and PCR was performed using primers corresponding to (A) genomic sequences upstream of and within the Slp-2 gene, or (B) genomic sequences flanking the Slp-2 gene. In A, the amplification of a 406 bp PCR product indicates the presence of the Slp-2 gene on at least one allele, while the absence of a PCR product indicates that Slp-2 is deleted on both alleles. In B, the presence of a 534 bp PCR product indicates deletion of the Slp-2 gene on at least one allele. Based on the data, the genotypes of the mice are as follows: M1: Slp-2+/+, M2: Slp-2-/-, M3: Slp-2+/-. NTC: no template control. Ctrl: control (i.e.,
534 bp deletion product confirmed by DNA sequencing). Numbers indicate size in bp. (C)
Immunoblotting of whole tissue lysates from adult mouse littermates was performed to detect
SLP-2 protein levels in Slp-2+/+, Slp-2+/-, and Slp-2-/- mice. Mouse genotype is indicated for each lane. All tissues were isolated from one mouse per genotype. Loading control: GAPDH.
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Supplementary Figure 1
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Supplementary Figure 2
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