Mechanisms of Caspase-3 Regulation in the Execution of Cell Death
DISSERTATION
Presented in Partial Fulfillment of the Requirements for the Degree Doctor of Philosophy in the Graduate School of The Ohio State University
By
Yadira Malavez
Graduate Program in Molecular, Cellular and Developmental Biology
The Ohio State University
2012
Andrea I. Doseff, Adviser
Dissertation Committee:
Dr. Clay B. Marsh
Dr. Harold A. Fisk
Dr. Tsonwin Hai
Copyright by
Yadira Malavez
2012
Abstract
Apoptosis is an evolutionarily conserved mechanism necessary for the homeostasis in multicellular organisms. The cysteine protease caspase-3 has a key role in apoptosis for its central role in the execution of the apoptotic cascade. The molecular mechanisms that regulate caspase-3 activation are not completely understood.
Previously, our laboratory demonstrated that caspase-3 is phosphorylated by PKCδ in human monocytes. However, the role of caspase-3 phosphorylation during apoptosis has not been elucidated. In this investigation, it was observed that PKCδ is necessary for caspase-3 phosphorylation. An interaction motif in caspase-3 was identified to be necessary for PKCδ interaction. Five PKCδ phosphorylated sites in caspase-3 were mapped utilizing mass spectroscopy. Phosphorylation of specific sites promoted caspase-
3 autocatalytic cleavage and apoptosis, in vitro and in vivo. Caspase-3 phosphorylation acts in a positive feedback mechanism to amplify the apoptotic cascade. These results suggest a novel regulatory mechanism to control caspase-3 apoptotic activity and execution of cell death.
Furthermore, the apoptotic and immunological heterogeneity in CD14+CD16+ and
CD14+CD16- monocytes was analyzed. It was observed that CD14+CD16+ cells were more susceptible to undergo spontaneous apoptosis, in part due to upregulation of the activity of the caspases. CD14+CD16+ monocytes release more TNF-α compared to the ii
CD14+CD16- counterparts. Furthermore, the expression of the members of the PKC family was characterized in monocyte subsets. These results suggested that elevated expression of PKCε may play a role in the pro-inflammatory role of CD14+CD16+ monocytes. The current investigation highlights the importance of caspase-3 and the members of the PKC family as important regulators of cell death and survival pathways.
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Dedication
This document is dedicated to my mother,
Mayra Acevedo Ponce,
for her love and support.
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Acknowledgements
I would like to thank Dr. Andrea I. Doseff for giving me the opportunity to be part of her laboratory and for her guidance and advice throughout these years. I would like to express my sincere appreciation to Drs. Clay B. Marsh, Harold A. Fisk and Tsonwin Hai, for accepting being in my committee and for their guidance. Next, I would like to thank the members of the Doseff laboratory, especially Drs. Oliver Voss, Martha E. Gonzalez-
Mejia, Arti Singh, and Mr. Daniel Arango. It was a pleasure to work with each of them and I am grateful for all the memories that I will take with me throughout my life.
Special thanks to Hassan Kamran and Justin Tossey, for their great help in the cloning of the phospho-mutants, and Maria Belen Federico for her help with the development of the cell free system protocol. I am deeply grateful with Dr. Wei Huang and Dr. Prabakaran
Nagarajan for their incredible help providing the PKCβ-/- and guidance for the isolation of mouse embryonic fibroblast. I also would like to thank Dr. Nancy Lill and Nurettin
Sever for allowing me to utilize their equipment necessary for the transfection of monocytes. Thanks to Lizanel Feliciano, Fabiola Jara, Greetchen Diaz, and Dr. Rebecca
Tirado-Corbala, for your friendship and support all these years. Thanks to my family for always believing in me and giving me their love and support. Special thanks to my husband, Esbal Jimenez, for his support and faith in me. Lastly, thanks God for giving me strength to achieve this goal.
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Vita
November, 6th 1979…………………………………San Juan, PR.
1997 – 2002………………………………………... B.S., Microbiology. University of Puerto Rico, Humacao, PR.
2002 – 2005………………………………………... M.S., Food Microbiology. University of Puerto Rico, Mayaguez, PR.
2006 – Present……………………………………… Graduate Research Associate Working towards Ph.D., Molecular Cellular and Developmental Biology. The Ohio State University, Columbus, OH.
Publications
Malavez Y., Voss O.H., Gonzalez-Mejia M.E. and Doseff A.I. PKCδ phosphorylation of caspase-3 regulates cell death and survival through a complex regulatory network (In preparation, 2011).
Malavez Y., Voss O.H., Gonzalez-Mejia M.E. and Doseff A.I. Characterization of cell fate and immune response of CD14+CD16- and CD14+CD16+ monocytes (In preparation, 2011).
Malavez Y., Gonzalez-Mejia M.E. and Doseff A.I. 2008. Protein Kinase C Delta (PRKCD). PKC. Atlas of Genetics and Cytogenetics in Oncology and Haematology. URL: http://AtlasGeneticsOncology.org/Genes/
Fields of Study
Major Field: Molecular, Cellular and Developmental Biology
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Table of Contents
Abstract……………………………………………………….………………….... ii
Dedication…………………………………………………………….……………. iv
Acknowledgments……………………………………………….…………………. v
Vita……………………………………………………………………..…………... vi
List of Figures………………………………………………………………………. xii
List of Tables…………………………………………………..…….……………... xiv
Abbreviations………………………….………………………….………………… xv
Chapter 1…………………………………..……………………….………..…….. 1
1.1 Introduction………….………………………………………………….….…… 1
1.2 Apoptosis…………………………………………..………….………….…...... 2
1.3 Caspase family…………………………………………..…….………………... 4
1.4 Apoptotic pathways ….………………………………………….….………….. 8
1.4.1 Caspase substrates ………………………………..……….……...………. 11
1.5 Regulation of caspase activity….……………..…………...……..……….……. 13
1.5.1 Inhibitors of apoptosis (IAP)……………………………………...………. 14
1.5.2 Regulation of caspase activity by phosphorylation....………………..…… 15
1.6 Caspase-3…………………………………………………..…………..……….. 17
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1.6.1 Caspase-3 structure and activation……………………………………….. 19
1.6.2 Caspase-3 functions……………..……………………..…………..…….. 22
1.6.3 Potential role of caspase-3 in human diseases…………...……………….. 23
1.6.4 Regulation of caspase-3 activity……………………….…………..…….. 25
1.6.4.1 Ubiquitination……………………………………………..……… 26
1.6.4.2 Nitrosylation……………………..………………………....…….. 27
1.6.4.3 Heat shock proteins …………………..………….…….…….…... 27
1.7 Protein kinase C delta (PKCδ) .…………….…..…………....…….…….….…. 29
1.7.1 The protein kinase C (PKC) family……………………….……..…..…… 29
1.7.2 PKCδ structure…..…………………………..……………..….…..……… 30
1.7.3 Role of PKCδ in apoptosis ..……….……………………..…..………..… 30
1.7.4 Substrates of PKCδ during apoptosis....…………………..….……..…… 32
1.8. Regulation of cell death in the immune response……….…………....………... 34
1.8.1 Innate and adaptive immune response.……………………………….….. 34
1.8.2 Monocytes and Macrophages ………………………..……..…….……... 35
1.8.3 The role of monocytes in the immune response...... 37
1.8.3.1 Toll-like receptors…………………………..……..……..…...….. 37
1.8.4 Monocyte subsets……….………………………………..….…………... 40
1.9 Summary………………………………………………………..……..………... 42
Chapter 2 Materials and methods………………………………….……….…… 58
2.1 Reagents and chemicals…………………………………………….…………... 58
2.2 Cloning and mutagenesis………………………………………….……………. 60
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2.3 Tissue culture …………………………………………..…………….….……... 66
2.4 Isolation of mouse embryonic fibroblasts (MEF)………………….……….…... 68
2.5 Extract preparation and immunoblotting……………………………..……….... 69
2.6 Protein quantitation and Western blot analysis………………………….…….... 69
2.7 Immunoprecipitation and in vitro kinase assays………………….……...... 71
2.8 Protein expression and purification………………………….……….……….... 72
2.9 Activation of caspase-3 by recombinant caspase-9…………….……….…….... 74
2.10 Activation of caspase-3 by MCF-7 cell extracts …………………..…………. 74
2.11 Caspase-3 activity assays ……………………………………………….…….. 75
2.12 In vitro kinase assays with recombinant caspase-3 ……………….…….…….. 76
2.13 Monocyte subset isolation ……………………….………………..…….…….. 78
2.14 Inhibition of caspase activity …………………………………….………..….. 79
2.15 Electroporation of monocyte subsets…………………………………….……. 80
2.16 IgG clustering ……………………………………………………….………... 80
2.17 Detection of TNF-α by ELISA …………………………………...………….. 81
2.18 Detection of PKC isoforms in monocyte subsets ………………….……….... 82
2.19 Flow cytometry ……..…………………………………………………..…..... 82
2.22 Statistical analysis……………………………………………………….….…. 83
Chapter 3 PKCδ phosphorylation of caspase-3 regulates the execution of 84 apoptosis……………………………………………………………………………. 3.1 Abstract……………………………………………………………………...….. 84
3.2 Introduction………………………………………………………………..…..... 85
3.3 Results…………………………………………………………………..……..... 87
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3.3.1 PKCδ is necessary for caspase-3 phosphorylation during cell death..… 87
3.3.2 Identification of caspase-3 domains phosphorylated and involved in 88 the interaction with PKCδ…………….………………..……….…………… 3.3.3 Identification of PKCδ interaction motif in caspase-3……………….... 89
3.3.4 Identification of PKCδ phosphorylation sites in caspase-3……..…...... 91
3.3.5 Ser12 is important for caspase-3 phosphorylation..………….……….... 94
3.3.6 The phosphorylation of Ser36 is necessary for the PKCδ dependent 95 phosphorylation of caspase-3………………………………….…………….. 3.3.7 Caspase-3 phosphorylation modulates its protease activity…….…...... 96
3.3.8 Role of Ser12 and Ser36 Phosphorylation of in the autocatalytic 97 cleavage of caspase-3….…………………………………....……………….. 3.3.9 Role of caspase-3-Ser36 phosphorylation in apoptosis…..…….……… 100
3.3.10 Phosphorylation of Ser36 is important for caspase-3 cleavage during 102 apoptosis……………………….…………………………………………….. 3.4 Discussion.…………………………………………………………………...... 103
Chapter 4 Characterization of cell fate and immune response of CD14+CD16- and 123 CD14+CD16+ monocytes ………………………….………………..…………….. 4.1 Abstract…………………………………………………………………….…… 123
4.2 Introduction………………………………………….…………………....…….. 124
4.3 Results…………………………………………………………..……………..... 129
4.3.1 Optimization of CD14+CD16- and CD14+CD16+ isolation conditions….... 129
4.3.2 Classical and non-classical monocyte subsets have differences in 130 spontaneous apoptosis...... 4.3.3 CD14+CD16- and CD14+CD16+ apoptosis is dependent on caspase-3...…. 133
4.3.4 Caspase cascade in monocyte subsets..……...…………………….…...... 132
4.3.5 LPS-induced TNF-α production is elevated in non-classical monocytes… 134
4.3.6 Expression of PKC-isoforms in different monocyte subsets…..………..... 135
4.3.7 The expression of PKCε is necessary for LPS-induced TNF-α release in 135 CD14+CD16+ monocytes.……………………….……….…….……..………… x
4.4 Discussion………………………………………………….……….…………... 136
Chapter 5 Conclusions and future direction……………………….…...... 152
References…..…………………..……………………………………………...... 163
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List of Figures
Fig. 1.1 Conservation of the apoptotic pathway in Caenorhabditis elegans and 43 Homo sapiens………………………………………………………………………. Fig. 1.2 Comparison of the domain structure of human caspases………………….. 44
Fig. 1.3 Initial processing site and substrate specificity of caspases………………. 45
Fig. 1.4 Apoptotic pathways……………………………………………………….. 46
Fig. 1.5 Comparison of the crystal structure of the active effector caspases………. 47
Fig.1.6 Schematic representation of procaspase-7 activation……………………… 48
Fig. 1.7 Mechanism of caspase-3 activation……………………………………….. 49
Fig.1.8 PKC family………………………………………………………………… 50
Fig.1.9 PKCδ structural domains and phosphorylation sites………………………. 51
Fig.1.10 Model of PKCδ activation………………………………………………… 52
Fig. 1.11 Hematopoiesis…………………………………………………………….. 53
Fig. 1.12 TLR4 Pathway…………………………………………………………… 54
Fig. 3.1 PKCδ is necessary for caspase-3 phosphorylation during cell death……… 110
Fig. 3.2 Identification of caspase-3 domains phosphorylated and involved in the 111 association with PKCδ……………………………………………………………… Fig. 3.3 Identification of PKCδ interaction motif in caspase-3…………………….. 112
Fig. 3.4 Identification of PKCδ phosphorylation sites in caspase-3………………... 113
Fig. 3.5 PKCδ phosphorylates caspase-3 Ser12 and Ser36 in vitro………………….. 114
Fig. 3.6 Caspase-3-Ser12 is an important phosphorylation site in HeLa cells………. 115
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Fig. 3.7 The phosphorylation of caspase-3-Ser36 is necessary for the PKCδ………. 116 dependent phosphorylation of caspase-3……………………………………………
Fig. 3.8 Caspase-3 phosphorylation modulates its protease activity in vitro………. 117
Fig. 3.9 Phosphorylation of caspase-3-Ser12 and caspase-3-Ser36 regulates the 118 second autocatalytic cleavage………………………………………………………. Fig. 3.10 Phosphorylation of caspase-3-Ser36 regulates apoptosis…………………. 119
Fig. 3.11 Phosphorylation of caspase-3-Ser36 is important for caspase-3 cleavage 120 during apoptosis……………………………………………………………………. Fig. 3.12. Model……………………………………………………………………. 121
Fig. 4.1 Monocyte subsets purification scheme……………………………………. 143
Fig. 4.2. Classical and non-classical monocyte subsets have differences in 144 spontaneous apoptosis……………………………………………………………… Fig. 4.3 CD14+CD16- and CD14+CD16+ monocytes require caspase-3 to undergo 145 apoptosis…………………………………………………………………………… Fig. 4.4 Apoptosis in monocyte subsets involves the activation of caspase-8 and 146 caspase-9…………………………………………………………………………… Fig.4.5 TNF-α production in monocyte subsets……………………………………. 147
Fig. 4.6 Expression of PKC isoforms in monocyte subsets………………………… 148
Fig. 4.7 The expression of PKCε is necessary for TNF-α release in CD14+CD16+ 149 monocytes………………………………………………………………………….. Fig. 4.8. Summary of the results for the characterization of cell death and immune 150 response in monocyte subsets…….………………………………………………… Fig. 5.1 Model…………………………………………………………………….. 162
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List of Tables
Table 1.1 Caspase family functions and classifications…………………………….. 55
Table 1.2 Caspase inhibitors and their inhibitory 56 constants…………………………………………………………………………..... Table 1.3 TLR receptors and ligands……………………………………………..... 57
Table 2.1 List of antibodies…………………………………………………….….. 59
Table 2.2 List of primers……………………………………………………………. 62
Table 2.3 List of clones in pQE31 vector……………………………………...…… 63
Table 2.4 Clones in pENTR/D-TOPO vector………………..……………...……... 64
Table 2.5 Clones in pcDNA-4/His Max vector (Xpress)……………..……...…….. 65
Table 2.6 List of siRNA oligonucleotides.…………………………………...…….. 80
Table 3.1. Table 3.1. Catalytic parameters of recombinant caspase-3-WT and 122 phospho-mutant proteins……………………………………………………………. Table 4.1 Optimized monocyte subset isolation conditions……………...………… 151
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List of Abbreviations
Abbreviation Description AAP Amyloid protein AFC 7-amino-4-trifluoromethylcoumarin Ala Alanine AMC Amino-4-methylcoumarin Apaf-1 Activating factor 1 AraC Arachidonic acid ARP2 and ARP3 Actin-related protein 2/3 complex Asp Aspartic acid B cells B lymphocytes Bcl B cell lymphoma BIRs Baculovirus IAP repeats BSA Bovine serum albumin C. elegans Caenorhabditis elegans c-Abl Abelson tyrosine kinase CAD Caspase-activated DNAse CARD Caspase recruitment domain Casp-3 Caspase-3 CCR Chemokine receptors CCR Chemokine receptors CD14 LPS receptor CD14+CD16- Clasical-monocytes CD14+CD16+ Non-clasical monocytes CD16 (FcγIII receptor CDK1 Cyclin-dependent kinase ced Cell death abnormal cFlipL Cellular FLICE- inhibitory protein, long isoform CHO Aldehyde group
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List of Abbreviations, continued
CL Cleaved Cys Cysteine DAPI 4-6-diamidino-2-phenylindole DC Dendritic cells DD Death domain DDD Three Asp residues DED Death effector domain DISC Death inducing signaling complex DMSO Dimethyl sulfoxide DNA-PK DNA-dependent protein kinase DR6 Death receptor 6 DYRK1A Dual-specificity tyrosine-(Y)-phosphorylation-regulated kinase A1 E1 Ub-activating enzyme E2 Ub-conjugating enzyme E3 Ub ligase EGF Epidermal growth factor egl-1 Egg-laying abnormal ERK1 and ERK2 Extracellular signal-regulated kinases FADD Fas-associated DD FasL Fas ligand FL Full-length FMK Fluoromethylketone FMLP N-formyl-methionine-leucine-phenylalanine Gly Glycine GM-CSF Granulocyte macrophage colony stimulating factor Grasp65 Golgi-stacking protein H2B Histone 2B HAT Histone acetyl transferase HCK Hemopoietic cell kinase HDAC2 Histone deacetylase-2 HRP Horseradish peroxidase HSCs Pluoripotent hematopietic stem cells Hsp27 Small heat shock protein 27 IAP Inhibitor of Apoptosis ICAD Inhibitor of caspase-activated DNAse xvi
List of Abbreviations, continued
ICE Interleukin -1b-converting enzyme IFN-β Interferon β IKKβ IκB kinase β IL-1 Interleukin-1 IL-1β Interleukin-1b IPTG 1-thio-β-D-galactopyranoside IRAK IL-1 receptor associated kinase IκB inhibitor of the nuclear factor κB, NF-κB LBP LPS-binding protein LPS Lipopolysaccharides LYN Tyrosine-protein kinase Lyn MAP Mitogen-activated protein M-CSF Macrophage colony-stimulating factor MEF Mouse embryonic fibroblasts MHC-II Major hystocompatibility complex-II MKK2/3 Mitogen-activated protein kinase-activated protein kinases MOMP Mitochondria outer membrane depolarization Mst1 Mammalian Sterile 20-like kinase 1 NF-κB Nuclear factor-κB NK cells Natural killer NKCC-1 Na-K-2Cl co-transporter-1 PAC-1 Procaspase activating compound-1 PAK2 p21-activated protein kinase-2 PAMPs Pathogens-associated molecular patterns PARG Poly-(ADP-ribose) glycohydrolase PARP-1 DNA repair proteins, like poly (ADP-ribose) polymerase-1 PB1 PKCδ binding motif 1 PB2 PKCδ binding motif 2 PBM1 PKCδ binding mutant 1 PBM2 PKCδ binding mutant 2 PBMC Peripheral blood mononuclear cells PBS Sterile phosphate buffered saline PCD Program cell death PD Parkinson’s disease
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List of Abbreviations, continued
PDB Protein Data Bank PDK1 phosphoinositide-dependent kinase 1 PKA Protein kinase alpha PKB Protein kinase beta PKC Protein kinase C PKCα PKC alpha PKCβI PKC beta I PKCβII PKC beta II PKCγ PKC gamma PKCδ PKC delta PKCε PKC epsilon PKCζ PKC zeta PKCη PKC eta PKCθ PKC theta PKCι/λ PKC iota/lambda PMA Phorbol 12-myristate 13-acetate PP1 Protein phosphatase 1 PSB Positive selection buffer PT Permeability transition pore rcaspase-3 Recombinant caspase-3 ROCK-1 Rho-associated kinase-1 ROS Reactive oxygen species rPKCδ Recombinant PKCδ RSK2 Ribosomal S6 kinase 2 S100A9 Protein S100-A9 Scd5 Stearoyl-CoA desaturase 5 Ser Serine sHsp Small heat shock protein SMC Smooth muscle cell accumulation Src Tyrosine kinase sarcoma T cells T lymphocytes TEMED N,N,N’,N’-Tetra-methyl-ethylene-diamine Thr Threonine TIR Toll/IL-1 receptor
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List of Abbreviations, continued
TLRs Toll-like receptors TNF Tumor necrosis factor TPA 12-O-tetradecanoylphorbol-13-acetate TRAIL TNF-related apoptosis inducing ligand Ub Ubiquitin V Variable regions Xpress pcDNA-4/His Max vector zIEALal Benyzloxycarbonyl-Ile-Glu(OtBu)-Ala-Leucinal
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Chapter 1
1.1 Introduction
A balance between cell death and cell survival is established to control cell numbers in multicellular organisms. The auto-demolition is controlled by program cell death (PCD) or apoptosis. Apoptosis is an evolutionarily conserved mechanism that plays an important role in embryonic development, as a defense mechanism during immune reactions and for the physiological removal of cells. Deregulation of apoptosis contributes to human conditions such as neurodegenerative diseases, ischemic damage, autoimmune disorders, and cancer. The cysteine protease caspase-3 is an essential regulator of the apoptosis pathway for its central role in the execution of cell death. The molecular mechanisms that regulate caspase-3 activation are still incompletely understood. This work identifies novel aspects in the mechanisms responsible for the regulation of caspase-3 apoptotic activity and presents their contribution in the regulation of cell death and survival. A better understanding of the molecular mechanisms that modulate apoptosis is necessary to elucidate the pathways that regulate biological systems and may contribute to the development of new therapies for diseases with dysregulated cell fate.
1
1.2 Apoptosis
In animals, millions of cells are eliminated daily though a program cell death or apoptosis (1-3). The term apoptosis is of Greek origin, that means “dropping off or falling off” in analogy to leaves falling of a tree or petals falling off a flower (4). This term describes the morphological processing observed in cells undergoing cell death.
The characteristic morphological changes that occur during apoptosis include cell shrinkage, loss of contact with neighbor cells, chromatin condensation, nuclear fragmentation, plasma membrane blebbing or budding, and cell fragmentation into compact membrane enclosed structures, called apoptotic bodies, which contain cytosol fragmented DNA and organelles (1, 4-6). Scavenger cells such as monocytes and macrophages recognize the phosphatidylserine receptor exposed in the plasma membrane of the apoptotic bodies and remove them through the process of phagocytosis (7). The engulfed apoptotic bodies are processed within the phagosome. The fusion of the lysosome with the phagosome allows the digestion of the apoptotic bodies by lysosomal hydrolases promoting an efficient re-utilization of the cell components, in the absence of inflammation (8). This process has important roles during embryonic development, differentiation, regulation and function of cells of the immune system, homeostasis maintenance and for the removal of damaged cells (9-13).
The cellular morphological changes that occur during apoptosis are accompanied by biochemical changes including the activation of the evolutionary conserved cysteine- aspartate-proteases, named caspases. The first caspase was initially identified in the genetic model organism, Caenorhabditis elegans (C. elegans), described as a molecule
2 involved in the regulation of cell death during development, named “cell death abnormal”
(ced)-3 (8, 14). Subsequent studies performed by Sydney Brenner, Robert Horvitz and
John E. Sulston received in 2002 the Nobel Prize Physiology and Medicine for the elucidation of the organ development and cell death pathway in C. elegans. They identified four genes that function sequentially to control the onset of apoptosis (Fig.1.1).
The activation of the effector ced-3 is in part mediated by the adaptor protein ced-4. In the absence of apoptosis, ced-4 is inhibited by interaction with the anti-apoptotic protein ced-9. During apoptosis egg-laying abnormal (egl-1) inhibits ced-4, allowing ced-3 activation for the execution of apoptosis. Orthologs of these proteins were later identified in higher eukaryotes. In humans this pathway is regulated by the family of caspases and other regulatory proteins (Section 1.3). Ced-9 is the ortholog of the mammalian B cell lymphoma (Bcl)-2 and Bcl-XL, while egl-1 is a BH3-only member of the Bcl2 family
(explained in Section 1.4). Ced-4 and ced-3 are the orthologues of the mammalian activating factor 1 (Apaf-1) and caspase-9 (8, 15). Since then, 14 members of the caspase family have been identified, of which 13 are found in humans; with the exception of caspase-11 only found in mice (16, 17).
The first human caspase identified, interleukin-1-converting enzyme (ICE), or caspase-1, was found to be involved in the maturation of the pro-inflammatory cytokine, interleukin-1 (IL-1β) (18-22). The overexpression of caspase-1 in mammalian cells was sufficient to induce cell death, demonstrating that have an analogous role in cell death as ced-3 (23, 24). Later, caspase-3 expression was shown to be necessary for mammalian cell apoptosis, supporting the role of the caspases in cell death (25). Recent studies have
3 revealed additional roles for caspases in proliferation, differentiation and inflammation
(9-13, 26, 27). Caspase-1 was shown to have an important role during inflammatory reactions (19). Based on these findings caspases were identified as inflammatory or apoptotic and great progress have been made to define the complex networks that regulate their function.
1.3 Caspase family
Caspases are cysteine proteases that are synthesized as inactive zymogens composed of three domains; an N-terminal domain, named prodomain, an intermediate large subunit (~ 20 kDa), and a C-terminal domain (~ 10 kDa) (Fig. 1.2). The size of the prodomain can vary among caspases, being long in initiator caspases (~ 200 amino acids) and short in executioner caspases (20 - 30 amino acids). The intermediate domain has the conserved QACXG (X is Arg, Gln or Gly) pentapeptide that contains the catalytic cysteine (Cys) (6). This pentapeptide is a signature of all caspases important for the identification of members of the family.
All caspases share a number of distinct features and can be classified based on their function, substrate specification, and phylogenetic relationships (28). Caspase-1, 4,
5, 11, 12, and, 13 are involved in inflammation and cytokine maturation; caspase-14 is involved in keratinocyte differentiation and maintenance of skin barrier and caspase-2, 3,
4
6, 7, 8, 9, and 10 are involved in apoptosis (29, 30). The activation of the apoptosis cascade is a process tightly regulated to activate this signaling cascade when needed.
Additional classification of the caspases has been made based on the mechanisms of caspase activation, categorizing them as initiators or executioners (Table 1.1) (31-33).
The prodomain of initiator caspases contain important interaction regions, such as the caspase recruitment domain (CARD) found in caspase-1, 2, 4, 5, 9, 11, and 12 and the death effector domain (DED) found in caspase-8 and -10. These domains mediate the interaction with accessory molecules that contain the same motif (Fig. 1.2) (34, 35).
Initiator caspases, like caspase-8 and -9, are activated by apoptotic stimuli and do not require a proteolytic cleavage by other caspases for their activation. In contrast, executioner caspases, such as caspase-3, -6, -7 and -14 require a proteolytic cleavage by an activator to initiate their activation (25, 36-39).
Structural information is available for some active caspases, including caspase-1,
-2, -3, -6, -7, -8, and -9. Analysis of the caspase-3 crystal structure revealed that the active enzyme is a heterodimer composed of two large (~ 20 kDa, α) and two small (~ 10 kDa, β) subunits aligned in a head to tail configuration, αββα. The heterodimer is composed of a core of 6 anti-parallel β strands in each monomer forming a continuous
12-stranded β-sheets in the enzyme active site flanked by α-helices (6, 16). The caspase active site is formed by flexible loops (L1-L4, L2’located in the adjacent monomer) composed of the S4-S3-S2-S1 specificity sub-sites. The specificity sub-sites bind the residues P4-P3-P2-P1 located in the target proteins. The loop L1 and a portion of the L2,
5 that contains the catalytic Cys, are part of the large subunit, while the L3 and L4 loops are part of the small subunit (33, 36). The size and topology of the L1 and L3 loops is conserved among all caspases, while the L2 and L4 have great variation. The S1 sub-site is contains the L1, L2, and L3 loops and is characterized for a narrow and deep structure.
The narrow structure of the S1 binding pocket limits the P1 residue to an Asp. This structural characteristic provides unique substrate specificity to all caspases for the cleavage of susbtrates that contain a tetrapeptide (P4-P3-P2-P1) that has an Asp at the position P1 (Fig. 1.3).
The specificity of each caspase is given by the size of the caspase S4 sub-site, which defines the preference for the binding of residues at the position P4 (40). Previous studies identified the substrate specificity of caspases by cleavage of specific peptides linked to amino-4-methylcoumarin (AMC) (40). Measurements of fluorescence over time allowed the classification of the caspases into three groups based on the preference for the P4 binding site (Fig. 1.3). Group I (caspase-1, -4, and -5) have the widest S4- binding site which favors large hydrophobic amino acids and cleave the (W/L)EHD tetrapeptide motif. Group II, composed of effector caspases (caspase-2, -3, and -7), has a narrow pocket with high hydrogen bond interactions that favors an Asp at the P4 position and cleave the DEXD tetrapeptide motif (X is any amino acid). Group III, composed mostly of initiator caspases (caspase-6, -8, and -9), has an intermediate size in the S4 pocket preferring small hydrophobic amino acids, such as Val and Leu. Group III cleaves (I/V/L)EXD tetrapeptide motifs (21, 28, 40).
6
Further analysis revealed that initiator caspases had a preference for the cleavage of tetrapeptides contained in their own structure, which allow them to perform self- activation (Fig.1.3) (41, 42). In contrast, the executioner caspases contained sequences optimal for the cleavage by an initiator caspase (Group III) (43). This indicated that the effector caspases required a cleavage by an initiator caspase for their activation.
Based on the substrate specificity of the caspases synthetic inhibitors were designed (44, 45). These inhibitors are highly specific in their substrate recognition and are widely used to investigate the regulation of caspases. The caspase inhibitors are membrane permeable peptides linked to either fluoromethylketone (FMK) or aldehyde group (CHO). These inhibitors bind to the active Cys-residue of a caspase in a reversible or irreversible manner. Reversible inhibitors contain a specific tetra-peptide-sequences followed by an CHO (45), whereas irreversible inhibitors contain a FMK group (44). A tri-peptide-inhibitor was developed to be able to bind and inhibit almost all caspases (e.g.
VAD-FMK). The VAD-FMK inhibits caspase-3 with a Ki of 43 nM, while it restrains caspase-8 and caspase-9 activation with a Ki of 2.5 and 3.9 nM, respectively (Table 1.2).
The use of caspase inhibitors provided important information about the mechanism of caspase activation and functions (40, 46-48). DEVD-FMK, is a potent caspase-3 inhibitor Ki of 0.32 nM (Table 1.2), but can also inhibit caspase-7, at least in vitro, with a lower affinity (Ki 0.92 nM) (Table 1.2) (40). These studies provided invaluable information about the structural regulatory mechanisms that provide the specificity of the members of the caspase family and allowed the elucidation of the apoptotic pathways.
7
1.4 Apoptotic pathways
The apoptotic cascade can be activated through two main pathways, termed the extrinsic and intrinsic pathway (Fig. 1.4). The former is activated by apoptotic stimuli like chemotherapeutic agents, cytokines, ROS, growth factor deprivation, anoxia, and
DNA damage all of which induce loss of mitochondrial transmembrane potential (ΔΨm).
This process is characterized by the formation of the permeability transition pore (PT), and release of cytochrome c from the inner membrane of the mitochondria to the cytoplasm (49, 50). Once in the cytoplasm, cytochrome c binds Apaf-1 and in the presence of ATP/dATP recruits procaspase-9 to the apoptosome heptameric-complex (51,
52). Apaf-1 contains a CARD domain that mediates the interaction with caspase-9. The oligomerization of procaspase-9 induces an autocatalytic cleavage leading to the dimerization and subsequent activation of the executioner caspases (53, 54). Active caspase-9 is responsible of the first cleavage required for the activation of the effectors caspase-3 and -7. These effector caspases cleave numerous cellular targets that direct the controlled demolition of the cell (discussed in Section 1.4.1). In addition, caspase-3 directs a positive feedback loop by cleaving caspase-9 at Asp330, enhancing caspase-9 apoptotic activity by 8-fold (55).
The activation of the intrinsic apoptotic pathway is regulated at multiple stages to prevent improper activation of the cell death machinery. Among the proteins involved in this pathway, the Bcl-2 family has been recognized as important regulators of the mitochondrial-dependent apoptosis (56). Bcl-2 family members share one to three Bcl-2
8 homology domains named BH1-3 and are divided in two groups of pro and anti-apoptotic proteins involved in the regulation of the permeability of the outer mitochondria membrane. The anti-apoptotic members of the Bcl-2-family, including Bcl-2, Bcl-xL,
Bcl-w, Mcl-1, and A1, prevent the mitochondria outer membrane depolarization
(MOMP) and the subsequent release of cytochrome c. Whereas the pro-apoptotic members of the Bcl family (Bax, Bak and Bock) and the BH3-only proteins (Bid, Bad,
Bik, Bim, Bmf, Hrk, Noxa, and Puma) promote mitochondria membrane permeabilization (30). The BH3-only proteins detect cellular stress or damage and induce Bax-Bak oligomerization. Currently, there are two models that that explains how the BH3-only proteins promote MOMP. The sensitizer model proposes that BH3-only proteins, such as Hrk, Noxa, Bad, Bik, and Bmf, promote cell death indirectly by binding the pro-survival Bcl-2 proteins liberating Bax and Bak. The second model proposes that some BH3-only proteins, such as Bid, Bim, and Puma, directly bind to Bax and Bak promoting a conformational change that allows their oligomerization (57). The MOMP opens a channel that allows the release of pro-apoptotic proteins from the mitochondria intermembrane space, such as cytochrome c, SMAC/DIABLO, and OMI/HTRA2 to the cytoplasm (Fig. 1.4).
In addition, other regulators of the intrinsic apoptotic pathway are the Inhibitor of
Apoptosis (IAP), first identified in baculoviruses for their involvement in suppression of host cell death response to viral infection (58). This family is composed of several members, including c-IAP1, c-IAP2, XIAP (X-linked AIP), and ML-IAP (melanoma
IAP), that inhibit cell death by direct binding to initiator and effector caspases. Most 9
IAPs contain a conserved baculoviral IAP repeat (59), which contains ~ 80 amino acids folded around a zinc atom. The BIR domains are the protein interaction domain that mediates the inhibition of caspases (60). During apoptosis, the activity of the IAPs is inhibited by SMAC/DIABLO, after its release from the mitochondria. This protein shares an IAP-binding tetra-peptide motif preventing the inhibition of caspases by the
IAP family. Binding of SMAC/DIABLO to XIAP allows the activation of caspase-9 by
Apaf-1 and cytochrome c, and the subsequent activation of effector caspases to execute cell death.
The activation of the extrinsic pathway is mediated by death receptors located at the cell membrane (61). The family of death receptors is composed of 6 members, such as Fas (CD95 or Apo-1), TNF-R1, TRAIL-R1 (DR4) TRAIL-R2 (DR5 or Apo-2), death receptor 3 (DR3, Apo-3, Tramp or Wsl-1 or Lard), and death receptor 6 (DR6). Death receptors are transmembrane proteins containing an N-terminal extracellular domain and a C-terminal intracellular tail. The extracellular domain contains five cysteine-rich repeats and the cytoplasmic region contains a death domain (DD-domain). The DD- domain allows the initiation of the death signal upon binding of the cognate ligand (62).
Binding of the ligand causes oligomerization of the receptor through the DD-domain promoting the recruitment of adapter proteins to the complex, such as the Fas-associated
DD (FADD). The adapter proteins contain death effector domain (DED) that mediates the recruitment of initiator caspases containing a DED (caspase-8 or caspase-10) or the cellular FLICE- inhibitory protein, long isoform (cFlipL). The interaction of these proteins allows the formation of the death inducing signaling complex (DISC) (63). The 10
DISC promotes the dimerization of caspase-8 by increasing the concentration of inactive monomers, promoting its activation (64-67). Activation of caspase-8 by the CD95-DISC involves the activation of the effector caspases, caspase-3, caspase-6 and caspase-7 (68).
Cells harboring the capacity to activate the extrinsic pathway through this signaling cascade are called type I cells.
Type II cells produce less active caspase-8 at the DISC level and involve the additional activation of the mitochondrial cell death pathway upon treatment with Fas ligand (FasL) (69, 70). In these cells caspase-8 cleaves Bid, a Bcl-2 family member, and the cleaved Bid translocates and accumulates in the mitochondria. Cleaved Bid induces
Bax-Bak oligomerization causing MOMP and cell death (71, 72). The involvement of the intrinsic pathway in cells treated with Fas L amplifies the apoptotic cascade. Since type II cells rely on the activation of the mitochondrial pathway it can be inhibited by overexpression of members of the Bcl-2 family, such as Bcl-2 and Bcl-XL (73, 74).
Thus, activation of the apoptotic program leads to a controlled proteolysis of cell components that can either activate or inactivate the target protein.
1.4.1 Caspase substrates
During apoptosis, nearly 400 proteins are cleaved by proteases, representing ~ 2% of human proteome (75). The cleavage of cytoplasmic and nuclear proteins during cell death cause the morphological characteristics associated with apoptotic cells. The characteristic apoptotic cellular condensation occurs after the caspase-dependent cleavage of cytoskeleton components, such as β-catenin, E-cadherin, actin, vimentin, GAS2, α-
11 adducin, SLK, gelsolin, and cytoketatin-18 (26). Membrane blebbing has been associated with the cleavage of α-fodrin, major component of the cortical cytoskeleton
(76). Under physiological conditions, α-fodrin is cleaved by calpain for the process of cytoskeletal remodeling, producing an intermediate cleaved fragment of 150 kDa. This intermediate polypeptide is specifically cleaved during apoptosis by caspase-3 producing a 120 kDa fragment (77). Membrane blebbing has also been associated with cleavage of
Rho-associated kinase-1 (ROCK-1). Cleavage of ROCK-1 promotes its activity and subsequent phosphorylation of myosin light chain (78) (Fig. 1.4).
DNA condensation is also controlled by the caspase-3-dependent cleavage of the
Mammalian Sterile 20-like kinase 1 (Mst1). Cleavage of Mst1 promotes its activation and subsequent phosphorylation of histone 2B (H2B) followed by DNA condensation
(79). Nuclear fragmentation is associated with the caspase-mediated cleavage of nuclear lamins (lamin A, B and C) and ROCK-1 mediated contraction of actin bundles. Actin surrounds the nuclear lamina, the cleavage of lamins and the contraction of actin bundles causes the collapse of the nucleus dispersing the nuclear material into the cytoplasm (74).
Lamins A and C are cleaved by caspase-6, while caspase-3 cleaves lamin B (80). DNA fragmentation is caused by the endonuclease-mediated chromatin cleavage in the inter- nucleosomal spaces by CAD (caspase-activated DNAse). In healthy cells, CAD activity is inhibited by ICAD (inhibitor of caspase-activated DNAse) (81). However, during apoptosis, caspase-3 cleavage of ICAD releases CAD inhibition allowing DNA hydrolysis (82, 83).
12
The fragmentation of cellular organelles, such as Golgi apparatus is an important event during apoptosis, facilitating the packing of these organelles into apoptotic bodies.
This process is associated with caspase-mediated cleavage of the Golgi-reassembly stacking protein 65 (Grasp65), golgin-160, p155, cytoplasmic dynein intermediate chains
(CD-IC), p150, syntaxin-5, and giantin (84-87).
As part of the apoptotic cell disassembly caspases, specifically caspase-3, cleave and inhibit essential proteins ranging from anti-apoptotic molecules, such as Bcl-2 and
DNA repair proteins like poly (ADP-ribose) polymerase-1 (PARP-1), poly-(ADP-ribose) glycohydrolase (PARG) and DNA-dependent protein kinase (DNA-PK), inhibiting the
DNA repair machinery (88-90). Caspases also cleave the cell cycle regulator retinoblastoma (Rb) (91). Despite the progress made in the identification of proteins cleaved during apoptosis, further studies are needed to identify the proteases responsible for these events.
1.5 Regulation of caspase activity
Caspases cleave multiple targets with diverse biological function to direct the controlled demolition of the cells. Lack of apoptosis is associated with cancer and developmental malformations. In contrast, excessive apoptosis is associated with several diseases, such as neurodegenerative disorders, ischemic injury, and AIDS (4, 39, 92). To control the activation of the apoptotic cascade cells have developed strategies to regulate the activity of caspases though the activity of IAPs and post translational modifications.
13
1.5.1 Inhibitors of apoptosis
The members of the IAP family have 1-3 baculovirus IAP repeats (BIRs) and a caspase recruitment domain (55). XIAP, IAP-1 and IAP-2 contain three BIR domains and the third domain (BIR3) inhibits caspase-9 activity, by blocking the homodimerization required for its activation (93, 94). The XIAP inhibition of caspase-9 is released by Smac/DIABLO allowing the initiation of the apoptotic cascade (95).
XIAP BIR domain also inhibits caspase-3 and -7 by binding their active site in a reverse orientation blocking the entry of substrates (93, 96). Through this mechanism
XIAP prevents the second auto-catalytic cleavage and activation of these caspases.
Activation of the extrinsic apoptosis by treatment with UV or TRAIL activates caspase-3 causing the cleavage and inactivation of XIAP in melanoma cells (97). Silencing of
XIAP promotes caspase-3 cleavage and activation after co-treatment with TRAIL and
UVB; whereas overexpression of a non-cleavable XIAP mutant prevented caspase-3 activity (97). These studies suggest that caspase-3 activation and inactivation is a dynamic process and the dysregulation of this balance is associated with malignancy
(98). Elevated expression of XIAP was found in several breast cancer cell lines (98).
Treatment of ovarian cancer epithelial cells with the chemotherapeutic drug cisplatin reduced the expression of XIAP and induces the cleavage of caspase-3 and -9. Thus, inhibition of XIAP promotes the execution of apoptosis (97, 99, 100).
cIAP1 has also been shown to associate with caspase-3 and caspase-7 during different steps of their processing (101). cIAP1 binds to active caspase-7 tetramer at the
14
AKPD motif found at the N-terminus of the large subunit, while it binds to the SGIS motif in the caspase-3-prop17. The AKPD motif is important for cIAP1 ubiquitination of caspase-7. The SGIS was found to be necessary for caspase-3 ubiquitination in vitro, but not in cells. The SGIS (aa 29-32) motif seems to be important for caspase-3 activation, since its replacement with the caspase-7 homologous sequence, AKPD, prevents TRAIL induced caspase-3 cleavage (101). Further studies will be necessary to determine how cIAP1 regulates caspase-3 activity and execution of apoptosis.
Collectively, these studies support the role of the members of the IAP family as important regulators of caspases that inhibit apoptosis by blocking the activation of initiator or executioner caspases.
1.5.2 Regulation of caspase activity by phosphorylation
Phosphorylation is an important regulatory mechanism to control the activity of caspases. (102-109). Caspase-9 was the first member of the family shown to be regulated by phosphorylation (105). Caspase-9 phosphorylation by the pro-survival kinase
AKT/PKB (protein kinase beta) at Ser196 inhibits its activity. After that finding other studies have identified additional phosphorylable sites in caspase-9. Caspase-9-Thr125 is the best characterized phosphorylation site shown to be targeted of the extracellular signal-regulated kinases (ERK1 and ERK2) MAPK, cyclin-dependent kinase (CDK1), and the dual-specificity tyrosine-(Y)-phosphorylation-regulated kinase A1 (DYRK1A).
ERK1/2 phosphorylates caspase-9-Thr125 in cells treated with 12-O- tetradecanoylphorbol-13-acetate (TPA) or epidermal growth factor (EGF) inhibiting its
15 activity (102, 103, 110, 111). Inhibition of caspase-9 by CDK1 phosphorylation of
Thr125 was shown to prevent apoptosis during the mitosis phase of the cell cycle. The phosphorylation of this site represses caspase-9 activation and elevated levels of phosphorylated Thr125 were found in gastric carcinomas, suggesting that it could be a marker for malignancy (102). In addition, DYRK1A phosphorylation of caspase-9 was shown to protect retina cells from apoptosis during neuronal cell development (111).
Additional phosphorylation sites have also been characterized. Caspase-9-Ser144 has been shown to be a target of Protein kinase C zeta (PKCζ) in response to hyperosmotic stress
(104). Also, casein kinase 2 (CK2) phosphorylates Ser348 repressing caspase-9 activity in mice (106). In addition, Abelson tyrosine kinase (c-Abl) phosphorylation of Tyr153 was reported to enhance caspase-9 activity in response to DNA damage, but the mechanism has not been elucidated (107)
Several studies have also reported caspase-8 regulation through phosphorylation.
The tyrosine kinase sarcoma (Src) phosphorylates caspase-8-Tyr380 repressing Fas- induced apoptosis in cells treated with EGF (112). Caspase-8-Thr263 is phosphorylated by the ribosomal S6 kinase 2 (RSK2) and overexpression of RSK2 was found to reduce apoptosis and delays caspase-8 activation in cells treated with FasL. Caspase-8 was also shown to be phosphorylated by p38-MAPK and CDK1 (113). Caspase-8-Ser387 phosphorylation by CDK1/Cyclin B1 prevents-Fas mediated apoptosis during mitosis and
CDK1 silencing allows caspase-8 activation and apoptosis. Caspase-8-Ser387 phosphorylation was elevated in cancer cell lines and breast cancer tissue, suggesting that caspase-8-Ser387 phosphorylation may contribute to tumorigenesis (114). 16
The activity of effector caspases is also regulated through phosphorylation.
Caspase-7 was recently shown to be phosphorylated by p21-activated protein kinase-2
(PAK2) repressing its activation in vitro (115). Caspase-3 was shown to be phosphorylated by p38-MAPK at Ser150 preventing Fas-induced apoptosis in human neutrophils (113). We have previously shown that caspase-3 is phosphorylated by protein kinase C delta (PKCδ) (116). Preliminary studies suggest that the PKCδ dependent phosphorylation of caspase-3 promotes its activity in human monocytes (116).
However, the molecular mechanisms of how PKCδ regulates caspase-3 activity have not been elucidated. Altogether, these results suggest an important role of caspase phosphorylation controlling life and death decisions.
1.6 Caspase-3
The caspase-3 gene is located in chromosome 4 (4q34/8) and was identified as the homolog of CED-3 and caspase-1 in C. elegans and humans, respectively (117). Two independent groups identified this enzyme to be responsible for the cleavage of PARP and characterized it as a pro-apoptotic protease. It was named Yama (after the Hindu god of the death) and Apopain and was later re-named caspase-3 (25, 118). Evidence for the critical role of caspase-3 during apoptosis was provided by the usage of malignant cells that lack caspase-3 expression (119, 120). Cell lines that lack caspase-3 expression are resistant to apoptosis through the intrinsic and extrinsic pathways. KMH2, a malignant tumor cell line of nodular lymphocyte predominance Hodgkin’s disease (NLPHD), lacks caspase-3 expression and showed resistance to apoptosis induced with FasL agonist
17
(121). MCF-7 breast cancer cells lack caspase-3 expression due to a 47-base pair deletion in exon-3, which makes them resistant to the intrinsic pathway triggered by the chemotherapeutic agents etoposide and doxorubicin (122). These cells are sensitive to apoptosis induced by staurosporine or TNF-α, but lack some of the apoptotic morphological features such as shrinkage, membrane blebbing and DNA fragmentation.
Reconstitution of caspase-3 expression in MCF-7 cells followed by treatment with TNF-α restored the apoptotic morphological changes (120). These results suggest that another caspase can substitute for caspase-3 during TNF-α or staurosporine induced apoptosis, but indicate that caspase-3 is important for the execution of some key events in the demolition phase of apoptosis like DNA fragmentation, cell shrinkage and membrane blebbing (122).
Caspase-3-/- mouse were developed to assess caspase-3 importance during development and apoptosis. The C57BL/6 caspase-3-/- mouse were significantly smaller than caspase-3 wild-type animals and died 1-3 weeks after birth (92). These mice had supernumerary enlarged deformed brains caused by the lack of apoptosis necessary during brain development (92) and the lack of apoptosis correlated with reduced caspase-
7 expression. However, the 129S1/SvImJ caspase-3-/- mice strain lack developmental defects due to an elevated expression of caspase-7 in the brain (123). Attempting to identify the role of caspase-3 and caspase-7 in apoptosis a double knockout casp-3-/-
/casp-7-/- (DKO) was developed. These mice died rapidly after birth with defects in cardiac development. Isolated mouse embryonic fibroblasts (MEF) were highly resistant to mitochondrial and death receptor induced apoptosis (124). These studies provided 18 evidence of the compensatory role of members of the caspase family and differences in the activity of mice strains. The usage of cell lines such as MCF-7 and KMH2 that lack caspase-3 expression is valuable to understanding the role of caspase-3 in cell death.
1.6.1 Caspase-3 structure and activation
The structure of the mature active executioner caspases, including caspase-3, -6 and -7 have been determined by X-ray crystallography (Fig. 1.5). The active caspase-3, -
6, and -7 share similar structures, composed of two heterodimer, each one containing a large p20 subunit and a small subunit p10. The three effector caspases contain similar arrangements of the loop bundle structures L1-L4 and antiparallel conformation of the monomers (Fig. 1.5). Caspase-3 has 54% sequence homology with caspase-7 and shares a preference for the cleavage of the synthetic substrate DEVD-AFC (AFC, 7-amino-4- trifluoromethylcoumarin) substrate with a Km of 9.7 and 11 μM, respectively (125-127).
Despite this fact, caspase-3 cleaves a wider array of proteins during apoptosis and is largely responsible for the demolition phase of apoptosis (75). Caspase-6 shares a sequence homology of 41% with caspase-3 and a limited amount of substrates have been identified, including lamin A and C (88, 128). Caspase-6 has a different substrate specificity than caspase-3 and -7, showing a preference for the cleavage of the substrate
VEID-AFC substrate with a Km of 30 μM, while it cleaves the DEVD-AFC with a Km of 180 μM (129). Previously, pro-caspase-6 was shown to be cleaved by caspase-3, but it can also be activated in the absence of caspase-3 activity by caspase-8 (52, 130).
19
The crystal structure of inactive caspase-3 (pro-caspase-3) remains unresolved, due to lack of electron density in the prodomain. Thus models of caspase-3 activation have been done based on the structure of the inactive caspase-7 (pro-caspase-7), (127,
131). Pro-caspase-7 substrate binding groove is composed of four flexible loops L1-L4 and one loop from the adjacent monomer, L2’ (Fig.1.6). The L1 and L4 loop are the sides of the active site; the L3 loop comprises the bottom, and the L2 loop contains the catalytic Cys183. Analysis of pro-caspase-7 crystal structure revealed that a covalent bond between the L2 and L2’ loops locks the enzyme in an inactive conformation. The covalent bond causes the catalytic cysteine to be rotated away from the S1 binding site maintaining the L2’ in the central cavity causing the disruption of the active site (127).
The caspase-6 crystal structure was recently generated and revealed that it contains a longer L2 loop than other effector caspases (~ 10 amino acids) (132). The longer L2 loop was described to play a role in caspase-6 auto-activation. However, caspase-3 and -7 have shorter L2 loops and this difference has been suggested to prevent the self- activation of caspase-3 and -7 (Fig. 1.5 and 1.6) (132).
During apoptosis caspase-3 is cleaved by an initiator caspase at Asp175, which separates the large (p17: 17 kDa) from the small (p12: 12kDa) subunit (Fig.1.7) (133).
This event is followed by an autocatalytic cleavage at Asp28 that releases the prodomain
(28 aa) (39, 133). An alternative cleavage at Asp9 has also been reported in vitro (134).
Mutations of the three cleavage sites prevent caspase-3 cleavage of the fluorogenic substrate DEVD-AFC, but do not affect the binding to the substrate (135). The two subunits of the p17 and p12 domains align forming the active caspase-3 (133). 20
Activation of caspase-3 increases its protease activity by ~ 100-fold allowing the controlled dismantling of the cell during apoptosis (136).
The activation of caspase-3 has been shown to be structurally regulated by a sequence of three Asp residues (DDD) (179-181), located in the p12 subunit, named
“safety catch” (Fig.1.7). The safety catch sequence has been proposed to maintain the pro-caspase-3 in a dormant state preventing its activation by upstream caspases and the autocatalytic cleavage in the absence of apoptosis. Mutations of the Asp residues to Ala caspase-3-D179-181A, or exposure to acidic buffers (pH 5.25 - 5.75) promotes the first cleavage by initiator caspases and activation of caspase-3. The fact that lower pH can trigger this cleavage is interesting as acidification has been reported in dying cells (135).
Structural and mutational studies have demonstrated that the amino acid side chains in the dimer interface are important for the caspase-3 active site formation, activation and specificity (137). Mutation of residues in the dimer interface
(Glu167Ala/Asp169Ala) prevented caspase-3 auto-activation and cleavage at Asp175. In addition, the double mutant Glu173Ala/Asp175Ala showed changes in substrate specificity
(138). In contrast, mutation of caspase-3- Val266Glu in the dimer interface affected the conformation of the active side loops and resulted in the pseudoactivation of caspase-3 in the absence of cleavage, increasing the activity of caspase-3 by ~ 60-fold. Transient expression of HEK-293 cells with caspase-3-Val266Glu in HEK-293 cells increased apoptosis by 30% in the absence of a proteolytic cleavage (139).
21
These studies suggest that caspase-3 exists inactive in the cell due to structural characteristics that prevent its activation in the absence of apoptotic stimuli. This structural regulation may serve as a way to restrain its activity in healthy cells since the activation of caspase-3 is the point of no returns that when unleashed will cause the controlled demolition of the cell.
1.6.2 Caspase-3 functions
Caspase-3 plays an important role in the execution of apoptosis activated through the extrinsic and intrinsic apoptosis pathway. Diverse studies have identified caspase-3 as the major protease responsible for the cleavage of structural cytoplasmic proteins, regulatory proteins, DNA-repair proteins, cell cycle regulatory proteins, and proteins in the apoptotic cascade (26). The analysis of cellular targets of caspase-3 and caspase-7 revealed redundancy in the cleavage of a few substrates, including PARP and ROCK-1
(77, 122, 125).
In addition of the role of caspase-3 in apoptosis, in recent years it became apparent that caspase-3 is also involved in non-apoptotic processes such as cellular differentiation but the mechanisms that specify this role have not yet been investigated.
Activation of caspase-3 has been reported during erythropoiesis (140). Similar observations have been reported during osteogenic differentiation, as caspase-3-/- mice show reduced osteoclast activity caused by reduced proliferation and differentiation of bone marrow stromal cells (141). During myoblast differentiation, myotube and myofiber formation was significantly decreased in caspase-3-/- primary cells (142). 22
Interestingly, caspase-3 activation was shown to be important during monocyte to macrophage differentiation with IL-32. However, the differentiation did not require the activity of the initiator caspases caspase-1, -8 and -9 (143). In addition, treatment with the macrophage colony-stimulating factor (M-CSF) induced monocyte to macrophage differentiation requiring some activity of caspase-3 and -9 (144). However, our group showed that M-CSF treatment activated AKT and induced survival but found no difference in caspase-3 activity (145). Thus, caspase-3 is an important regulator of cell death and survival and a better characterization of the mechanisms that regulate caspase-3 activity would be of use for the development of new strategies to modulate cell fate.
1.6.3 Potential role of caspase-3 in human diseases
Caspase-3 has been implicated in human pathologies, such as cancer, diabetes,
Alzheimer, and Parkinson disease, among others. However, its specific role has not been elucidated. Early studies identified caspase-3 as an attractive target for the treatment of cancer due to the resistance to apoptosis observed in radiotherapy and chemotherapy treated tumor cells. Caspase-3 was shown to be activated in cancer cells treated with chemotherapeutic drugs; including doxorubicin, etoposide, staurosporine, and cysplatin
(120, 146, 147). Reconstitution of caspase-3 expression in MCF-7 breast cancer cell line sensitizes them to radiotherapy (148). In addition, a screen of peptides identified the small molecule called procaspase activating compound-1 (PAC-1) as an activator of caspase-3 in vitro. PAC-1 was shown to activate caspase-3 by binding to the safety catch tripeptide inducing apoptosis in lung cancer cell lines (135). In addition, PAC-1
23 significantly delayed the growth of subcutaneous lung tumors in a mouse xenograft model. Diverse types of cancer are characterized to have elevated levels of procaspase-3
(~ 2 - 20-fold), including non-small cell carcinoma, melanoma, leukemia, and colon cancer. PAC-1 directly activates caspase-3 at low concentrations, which make it effective for the treatment of tumor cells that harbor elevated caspase-3 expression, showing low toxicity to the non-cancerous tissue. Later studies developed a PAC-1 derivate, S-PAC-1, which also activates caspase-3 and induces partial regression in dogs with leukemia (149). Thus, an assessment of the levels of pro-caspase-3 expression could be utilized to assess the appropriate therapy to target caspase-3 for the treatment of cancer.
Caspase-3 has been shown to be important role in the development of diabetes.
Caspase-3-/- mice are protected from diabetes and show β cell apoptotic resistance after treatment with the glucose analog streptozotocin (150, 151). Furthermore, the activity of caspase-3 is important for diabetic retinopathy, one of the most common complications in diabetes. Diabetic retinopathy causes alterations in blood flow, death of retinal pericytes
(perivascular contractile cells), basement membrane thickening, increases in vascular permeability, capillary occlusion, and retina degeneration, causing vision loss and even blindness (152). Retina capillary cells from diabetic mice express active caspase-3 and its activity has been associated with the development of diabetic retinopathy (153). Some studies have found that the pericyte apoptosis that occurs during diabetic retinopathy is mediated by caspase-3 and could provide a therapeutic perspective for the treatment of this condition (154). 24
Chronic neurodegenerative diseases, such as Alzheimer and Parkinson’s disease, are characterized by progressive and irreversible loss of synaptic function. The role of caspase-3 in Alzheimer’s disease is not completely understood, but some studies have implicated caspase-3 in disease development (155-157). Abnormal levels of active caspase-3 were detected in the brains of Alzheimer’s patients. Caspase-3 is involved in the proteolysis of the amyloid protein (AAP), and formation of amyloid αβ peptide, which is associated with the development of Alzheimer’s disease (155, 156). A recent study reported that caspase-3 activity correlated with reduction of spine density in pyramidal neurons. Inhibition of caspase-3 in the Tg2576-APPswe Alzheimer’s mouse model restores postsynaptic density, spine size and rescued memory (157). In
Parkinson’s disease (PD) patients, the increase in active caspase-3 in the substantia nigra correlated with increase in apoptosis of dopaminergic neurons (158, 159).
As a whole, these studies suggest that caspase-3 activity is important for diverse biological processes and may play a role in the development or pathology of human malignancies. However, the precise mechanisms that regulate the activation of caspase-3 remain unclear and require further experimental analysis.
1.6.4 Regulation of caspase-3 activity
Caspase-3 has been shown to be regulated through diverse mechanisms to control the execution of cell death. As described in previous sections, caspase-3 activation is inhibited by members of the IAP family (Section 1.5.1). In addition, caspase-3 activity has been shown to be modulated by phosphorylation (Section 1.5.2). Additional studies 25 have found that caspase-3 can be regulated through ubiquitination and by interaction with heat shock proteins.
1.6.4.1 Ubiquitination
Caspase-3 stability and activity have been shown to be regulated by ubiquitination. Ubiquitination of a target protein involves the attachment of ubiquitin
(Ub) protein (8 kDa) to a substrate in a complex series of reactions substrates are signal for proteasome proteolysis (60). The reaction is catalyzed by an Ub-activating enzyme
(E1), an Ub-conjugating enzyme (E2), and an Ub ligase (E3) (60, 160). The overexpression of the E3 Ub-ligase SAG/ROC-SCFB-TrCP reduces endogenous caspase-3, while silencing of these proteins increases caspase-3 expression levels in HEK293 cells
(161). Further analysis identified that ubiquitination may affect the stability of the cleaved caspase-3 subunits (162). The polyubiquitination of the p17 and p12 subunit of caspase-3 targets them for proteasomal degradation. Treatment with proteasome inhibitors like lactacystin and zIEALal (benyzloxycarbonyl-Ile-Glu(OtBu)-Ala-Leucinal) prevented the degradation of these domains. Hence, active caspase-3 stability is potentially regulated by polyubiquitination. The polyubiquitination of caspase-3 has been detected in vitro utilizing recombinant caspase-3; however, it has not been detected in cells undergoing apoptosis (101). Thus, polyubiquitination is a potential mechanism to control the stability of the active caspase-3 that may play a role controlling its activity during the execution of apoptosis, but more information is needed to assess its specific role in cell death. 26
1.6.4.2 Nitrosylation
The activity of caspase-3 is also regulated by the S-nitrosylation. Caspase-3 is nitrosylated in human cell lines and is de-nitrosylated in cells treated with FasL (163).
Caspase-3 nitrosylation inhibits caspase-3 activity by binding the catalytic cysteine,
Cys163. However, through de-nitrosylation caspase-3 activity is restored. Human lymphocytic cell lines have increased levels of S-nitrosylated caspase-3 in the mitochondria, suggesting that this process may play a role in the intrinsic apoptotic pathway (164). Thus, caspase-3 nitrosylation and de-nitrosylation is a dynamic process that prevents caspase-3 activation in the absence of apoptosis and re-activates its activity when needed.
1.6.4.3 Heat shock proteins
Heat shock proteins are chaperones synthesized by cells in response to heat, physiological, physical or chemical stress, assisting protein folding, preventing protein aggregation and reducing regulatory protein complex disruption (165). Heat shock proteins are classified as large or small according to their molecular weight. Large heat shock proteins have an ATP-dependent chaperone activity and include Hsp100, 90, 70, and 60 (166, 167). The small HSPs (sHsps) group is composed of 11 family members that have molecular weights between 15-40 kDa (166, 168). Their chaperone activity plays important roles in protein folding, prevention of protein aggregation, control of regulatory proteins, and movement of protein into cellular compartments (165). sHsp are characterized by having a conserved crystallin domain. sHsps have anti-apoptotic 27 functions against a wide range of cellular stresses (169-174). Also, α-crystallin, has been shown to protect cells from staurosporine, TNF-α, UV light, hydrogen peroxide, and etoposide treatments (171-174). α-crystallin consists of two similar subunits, αA- and
αB-crystallin, that form heteroaggregates (175). αB-crystallin inhibits the autocatalytic maturation of procaspase-3 and cytochrome c release during TNF-α-induced apoptosis of breast cancer cells (169, 170). Hsp27, another member of the family, plays an important role in the regulation of monocyte spontaneous apoptosis (176). Hsp27 is constitutively expressed in monocytes, whereas αB-crystallin is not (176). The expression of Hsp27 increases during monocyte to macrophage differentiation to repress apoptosis. Hsp27 was found to bind caspase-3 prodomain preventing caspase-3 dependent apoptosis in human monocytes through the inhibition of caspase-3 autocatalytic cleavage. Thus,
Hsp27 expression represses the apoptotic fate of monocytes.
Hsp27 is phosphorylated by p38 MAPK, PKCδ and mitogen-activated protein kinase-activated protein kinases (MKK2/3), during heat and oxidative stress on Ser15,
Ser78 and Ser82 (177-179). Phosphorylation of Hsp27 has been linked with loss of chaperone activity and inhibition of its anti-apoptotic function (180, 181). However, the phosphorylated form of Hsp27 can block the Fas-dependent pathway by preventing the
DAXX-Ask complex formation and subsequently inhibiting caspase-8 activation (182).
As in caspase-3, the role of Hsp27 phosphorylation is not well understood and further studies are needed to elucidate its role in the balance of cell death and survival.
28
1.7 Protein Kinase C delta (PKCδ)
1.7.1 The protein kinase C (PKC) family
The protein kinase C (PKC) family of Ser/Thr kinases consists of 11 isoforms further divided into subfamilies based on structural motifs and cofactor requirements
(Fig. 1.8) (183). The classical PKCs (α, βI, βII and γ) require 1,2-diacylglycerol, phosphatidylserine (PS), and Ca2+. They possess two cysteine rich-motifs involved in the interaction with phorbol ester and DAG, and a conserved region 2 (C2) for the binding to
Ca2+. The novel PKCs (δ, ε, θ, and η) require DAG for their activation and lack the C2 region that binds Ca2+, thus they are calcium independent. Novel PKCs have a C2-like region at the N-terminal domain that plays a role in protein-protein interaction. The atypical PKCs (ζ, τ and ι/λ) lack DAG, phorbol ester and Ca2+ binding regions, and are
DAG and Ca2+ independent. The differences in cofactor requirement between PKC isoforms confer specificity. Previous studies have demonstrated the involvement of
PKCs in monocyte differentiation, immunity and apoptosis (184-188). The cPKCs,
PKCα and PKCβI have been shown to regulate monocyte differentiation into macrophages and dendritic cells, respectively (184, 188). PKCζ and PKCε have important roles in the release of pro-inflammatory cytokines and activation of the nuclear factor-κB (NF-κB) signaling pathway, explained in Section 1.9.4.1 (186, 187). PKCδ plays an important role in regulation of apoptosis during multiple stages of the apoptotic cascade (116).
29
1.7.2 PKCδ structure
PKCδ, a novel PKC member, is a 78 kDa protein composed of a regulatory and a catalytic domain (Fig. 1.9) (189). The N-terminal regulatory domain contains an auto- inhibitory region, named pseudosubstrate, and four conserved domains, the C1 and C2- like in the regulatory region and the C3 and C4 located in the catalytic region. PKCδ also contains five variable regions (V); the V1 and V2 regions are located in the PKCδ regulatory domain, the V3 hinge region, separates the catalytic and regulatory domains, and the V4 and V5 regions are located in the catalytic domain (183, 190) (Fig. 1.9). The
C1 motif contains DAG/PMA (Phorbol 12-myristate 13-acetate) binding sequences that allow the interaction with a hydrophilic cleft located at a hydrophobic surface of this domain. The binding to the hydrophobic cleft forms a contiguous hydrophobic surface that promotes PKC binding to membranes. The C2-like region located at the N terminal domain has the same core residues of the C2 domain, but it lacks the essential calcium coordinating acidic residues that allows classical PKCs to bind Ca2+ (191). The C3 and
C4 are required for ATP/substrate binding and catalytic activity of the enzyme, respectively. The pseudosubstrate domain, located between the C1 and C2 motifs, maintains PKCδ in an inactive conformation by blocking the access to the substrate binding pocket (183, 192, 193).
1.7.3 Role PKCδ in apoptosis
PKCδ exists in the cell in an immature inactive conformation that requires post- translational modifications to achieve catalytic maturity before activation by DAG/PMA 30
(Fig. 1.9). Multiple studies have analyzed the activation of PKCδ during apoptosis that identified sequential steps that are involved in the catalytic maturation of this kinase. In the immature PKCδ, the pseudosubstrate domain interacts with the substrate-binding region of the catalytic site locking the kinase in an inactive state (194). During apoptosis, phosphorylation of the activation loop allows the catalytic maturation of PKCδ (195).
PKCδ catalytic maturation involves the auto-phosphorylation of Ser662 in the C3 domain and phosphorylation of Ser505 and Ser643 of the V5 region (Fig. 1.9). PKD1 has been shown to phosphorylate PKCδ-Ser505, while the Ser643 can be phosphorylated by PKCζ
(196). Next, PKCδ binds the cellular, nuclear or mitochondrial membrane through the
C1/C2 domains causing a conformational change that release the pseudosubstrate (197).
Subsequent auto-phosphorylation of Ser299, Ser302 and Ser304 are markers of PKCδ maturation.
Caspase-3 participates in the catalytic maturation of this kinase by cleaving the
Asp327 located in the hinge V3 region, separating the regulatory domain from the catalytic domain (Fig. 1.10) (198). Treatment with TNF-α, Fas, etoposide, mitomycin, cytosine arabinoside, etoposide, UV, and ionizing radiation induce the cleavage of PKCδ in various cell types (199-205). The cleaved catalytic fragment of PKCδ translocates to the nucleus utilizing its NLS (nuclear localization signal) to trigger apoptosis (199, 200).
Moreover, it has been suggested that the PKCδ catalytic fragment may serve to amplify downstream events in the apoptotic pathway (206). Overexpression of the catalytic fragment in the absence of an apoptotic stimulus was sufficient to induce apoptosis in a variety of cell types (207). Moreover, in glioma cells, treatment with etoposide in the
31 presence of the PKCδ inhibitor, rottlerin, inhibited PKCδ activation and showed decrease in the cleaved PKCδ (208, 209). PKCδ associates with caspase-3 and phosphorylates it promoting apoptosis in human primary monocytes (210). These results suggest an essential team-work of these molecules in the induction of apoptosis (201). Thus, PKCδ not only participates in later events of apoptosis by acting downstream of the caspases, but also participates in the regulation of early stages of apoptosis.
In light of the central role of PKCδ in apoptosis, PKCδ-/- mice were found to be protected against γ-irradiation induced apoptosis (211). Furthermore, vein segments from the PKCδ-/- isografted to carotid arteries of recipient mice develop a severe arteriosclerosis characterized by high monocyte and macrophage infiltration and smooth muscle cell (SMC) accumulation. The isolated SMC from PKCδ-/- mice are resistant to apoptosis induced by multiple stimuli and showed reduced caspase-3 activation, PARP cleavage and cytochrome c release, compared to the wild-type (212). Altogether, these studies support the role of PKCδ as an important pro-apoptotic kinase and the collaboration between PKCδ and caspase-3 is important for the execution of cell death.
Thus, elucidation of the role of PKCδ phosphorylation of caspase-3 in cell death is essential to understand the pathways that control cell fate.
1.7.4 Substrates of PKCδ during apoptosis
PKCδ phosphorylates numerous targets with diverse functions which contribute to the execution of the apoptotic pathway (213). PKCδ phosphorylates nuclear proteins contributing to key steps of the apoptosis pathway. DNA-PK that plays an important 32 role in the repair of double strand breaks has been shown to be a target of PKCδ (214).
The activated PKCδ directly binds and phosphorylates DNA-PK causing its disassociation from DNA and inhibition of DNA repair in response to DNA damage.
PKCδ was shown to interact with and phosphorylate the tumor protein 53 (p53), an important regulator of the cell cycle events, inducing apoptosis in response to DNA damage (215). Furthermore, PKCδ phosphorylates Rad9, a key factor involved in checkpoint regulation of the DNA damage response (216).
PKCδ also participates in the disassembly of the nuclear lamina, one of the hallmarks of apoptosis (217). During arachidonic acid (AraC)-induced apoptosis PKCδ translocates to the nucleus and phosphorylates lamin B. Phosphorylation was shown to occur concomitantly with the caspase-3 dependent cleavage of lamin B, which contributes to the characteristic nuclear appearance of apoptotic cells. In addition, PKCδ phosphorylates the small heat shock protein 27 (Hsp27) during apigenin-induced apoptosis (218). The phosphorylation of Ser15 and Ser82 inhibits the cytoprotective activity of Hsp27 promoting apoptosis in leukemia cells.
Although the majority of studies indicate that PKCδ is a pro-apoptotic kinase, there are some studies that have described non-apoptotic functions associated with the phosphorylation of cellular targets. PKCδ have been shown to phosphorylate various transcription factors, including p300, Sp1 and NF-κB. PKCδ phosphorylation of p300 inhibits its histone acetyl transferase (HAT) activity preventing nucleosome histone acetylation and gene expression (219). In addition, PKCδ phosphorylates Sp1 regulating it’s binding to histone deacetylase (HDAC)-2. PKCδ phosphorylates NF-κB in response
33 to TNF-α (220). Treatment of osteosarcoma cells with TNF-α induces PKCδ phosphorylation of NF-κB and its kinase activity was necessary for NF-κB nuclear translocation (220). PKCδ phosphorylates gamma actin regulating its interaction with the airway epithelial Na-K-2CI cotransporter (NKCC1) (221). NKCC1 activity is associated with cytoskeletal anchoring that affects cell shape and assembly of transport proteins.
Collectively, these results show the important role of PKCδ in the regulation of apoptosis and suggest that it may have additional roles in the absence of cell death.
1.9 Regulation of cell death in the innate immune response
1.9.1 Innate and adaptive immune response
The immune system provides protection against infectious diseases through self and non-self discrimination through cell-surface recognition (222, 223). Cells of the immune system have the potential to activate an inflammatory or apoptotic response.
The cells of the immune system originate in the blood marrow from the pluoripotent hematopietic stem cells (HSCs) (Fig. 1.11). The HSCs give rise to two progenitor cells, the lymphoid and myeloid stem cells (222-224). The lymphoid progenitors differentiate into B lymphocytes (B cells), T lymphocytes (T cells) and natural killers (NK cells). The myeloid progenitors give rise to megakaryocytes, erythroblast, basophil, eosinophil, and granulocyte/monocyte progenitors. These cells are the line of defense against pathogens through the acquired and innate immune response (225-227).
34
1.9.2 Monocytes and macrophages
Monocytes are essential components of the innate immune system. They constitute ~ 5 - 10% of the total peripheral blood mononuclear cells (PBMC). They circulate in the blood stream for 48 - 72 hours and in the absence of any stimuli, undergo spontaneous apoptosis in a caspase-3 dependent manner (116, 228, 229). We previously showed that caspase-3 is phosphorylated by PKCδ during monocyte apoptosis (116). The in vitro phosphorylation of caspase-3 promoted its activity suggesting that this may be a regulatory process to regulate monocyte cell death. However, monocyte spontaneous apoptosis can be prevented by exposure to differentiation factors, inflammatory signals, or by malignant transformation (145, 229, 230).
Differentiation factors such as macrophage colony stimulating factor (M-CSF) and granulocyte macrophage colony stimulating factor (GM-CSF) prevent monocyte apoptosis by promoting their differentiation into macrophages or dendritic cells. These cells have longer life spans that can range from months to years (223). Macrophages have an important role in tissue homeostasis in the clearance of senescent or apoptotic cells through tissue remodeling and repair after inflammation (231). Macrophages have a high phagocytosis capability and present antigens through the major hystocompatibility complex-II (MHC-II) to activate the adaptive immune response through the activation of
T cells. These phagocytic cells are highly heterogeneous with diverse specialized functions adopted in the anatomical location where they reside. In contrast, dendritic cells (DC) have reduced proteolytic capacity but they perform phagocytosis of pathogens and secrete cytokines to activate T cells and initiate their response (232). Thus,
35 monocytes can differentiate into macrophages or DC under different stimuli. M-CSF treatment induces monocyte to macrophage differentiation inhibiting cell death (145).
M-CSF was shown to activate the survival kinase Akt/PKB, which phosphorylates and inhibits caspase-9 and the apoptotic signaling cascade (105, 233). In addition, as mentioned in Section 1.8, the members of the PKC family have been shown to play a role in the monocyte to macrophage differentiation. Treatment with PMA induces monocyte to macrophage differentiation in a PKCα dependent manner (188). Alternatively, GM-
CSF and IL-4 treatment induced monocytes to differentiate into DC, in a PKCβ dependent manner (188, 234). Treatment of monocytes with the Gӧ6976 or the PKCβI specific inhibitor prevent PKCβI activation and differentiation into DC (188).
In addition, monocytes escape their apoptotic fate by exposure to inflammatory stimuli. Exposure to the Gram-negative lipopolysaccharides (LPS) and secretion of cytokines such as TNF-α and IL-1β have been suggested to inhibit monocyte cell death
(229, 230). Circulating monocytes become activated in the presence of pathogens, and the activated monocytes produce pro-inflammatory cytokines that cause inflammation
(235). Inflammation has three phases: initiation, maintenance and resolution (227, 235,
236). The pathogen is recognized by a monocyte or macrophage causing their activation.
The inflammatory response initiates and is maintained through the release of pro- inflammatory cytokines, chemokines and recruitment of monocytes, macrophages and other inflammatory cells to the site of infection (237). TNF-α is one of the most abundant cytokines released by monocytes and macrophages stimulated with LPS (238).
The secreted TNF-α recruits additional macrophages to sites of inflammation, and
36 promotes the release of additional pro-inflammatory cytokines, such as interferon β (IFN-
β). Through the phagocytic activity of monocytes and macrophages, bacteria and apoptotic cells are removed leading to the resolution of inflammation (236). Therefore, it is accepted that monocytes exhibit prolonged life span by inflammatory stimuli, leading to the accumulation of monocytes and macrophages in the site of infection. Altogether, monocyte survival and apoptosis maintains homeostasis by allowing their endurance in the presence of infection and restricting their survival when their activity is not needed.
1.9.4. The role of monocytes in the innate immune response
1.9.4.1 Toll-like receptors
Monocytes become activated by recognition of pathogens through cell surface receptors. Among these receptors, the Toll-like receptors are important mediators of the innate immune system. These receptors are evolutionary conserved and were first identified in Drosophila as an essential receptor for the dorsoventral specification (239).
Mutation of Toll receptors caused the development of fungal infections suggesting an important role of these receptors in the innate immune system (240). Mammalian homologs of the Toll receptors were identified and designated Toll-like receptors (TLRs).
Thirteen TLRs have been identified in mammals that recognize diverse pathogens- associated molecular patterns (PAMPs), including bacterial components such as peptidoglycan and lipoteichoic acid in Gram-positive bacteria (TLR1/2) and lipopolysaccharide and flagellin proteins (TLR4), RNA (TLR3, 7 and 8) and DNA
37
(TLR9, 10) nucleic acids (241-243) (Table 1.3) (244). Activation of the TLRs triggers signaling pathways leading to the production of proinflammatory cytokines as well as the maturation of antigen presenting cells of the innate immune system. The TLRs function as important sensors of the innate immune system that identify and activate the innate immune response to eliminate the invading pathogens.
The TLR4 recognizes LPS in the bacteria cell wall of Gram-negative bacteria and is the best characterized member of the TLR family. The activation of TLR4 involves the activity of the LPS-binding protein (LBP) and CD14 receptor (Fig. 1.12). In the circulation, LBP recognizes LPS and transfers it to the CD14 receptor (245). The CD14 presents LPS to the extracellular portion of the TLR4 receptor. TLR4 exists in a complex with MD-2 protein. The interaction between LPS and the TLR4-MD-2 causes oligomerization of the receptor and recruitment of adaptor proteins through interaction with the Toll/IL-1 receptor (TIR) domain, which received its name for the high similarity to the interleukin-1 (IL-1) receptor family (243).
The TLR4 signaling has been divided into the myeloid differentiation primary response gene-88 (MyD88)-dependent and MyD88 independent pathways. MyD88 is an important adaptor protein that binds TLR4 through the TIR domain. Mouse macrophages from the MyD88-/- mice are unable to activate TLR4 in response to LPS suggesting a key role of MyD88 in the production of proinflammatory cytokines (246, 247). The MyD88 independent pathway involves the transcription of Type I IFNs and IFN-inducible genes that play a role in the host defense against viral infections, as well as activation of antigen
38 presenting cells (248, 249). This pathway promotes the release of cytokines involved in the activation of T cells, B cells and NK cells (250).
During the activation of the TLR4 in a MyD88-dependent pathway, MyD88 recruits and activates the IL-1 receptor associated kinase (IRAK) (Fig. 1.12). IRAK activation promotes the phosphorylation and activation of the IκB kinase β (IKKβ) (251).
IKKβ binds to two members of the IKK family, IKKα and IKKγ forming the IKK complex. Once IKKβ is active it promotes the phosphorylation of IκB (inhibitor of the nuclear factor κB, NF-κB) and targets it for degradation. Degradation of IκB allows the activation and nuclear translocation of NF-κB for the transcription of pro-inflammatory cytokines, such as TNF-α, IL-1β and IL-6 (252). In addition, IgG complexes bind to Fc receptor family members FcγRI (CD64), FcγRII (CD32) and FcγRIII (CD16) and stimulate the release of TNF-α, IL-1β and IL-6 by activation of NF-κB (253). The signaling cascade that regulates the release of pro-inflammatory cytokines by binding to
IgG in monocytes has not been elucidated.
PKCs have been shown to regulate cytokine secretion in monocytes and macrophages. Stimulation of monocytes and macrophages with LPS promotes PKCζ activity inducing NF-κB activity, however the mechanisms have not been elucidated
(254). Recent studies have suggested that PKCε plays a role in the TLR4 pathway, but the role in this pathway is not well understood. PKCε was recently shown to be recruited to the TLR4 in a MyD88 dependent manner (255) (Fig. 1.12). Macrophages from the
PKCε-/- mice show reduced activation NF-κB after LPS stimulation and release low amount of TNF-α, IL-1β, PGE2, and nitric oxide compared to wild-type mice (186). NF-
39
κB activation was suppressed by inhibition of PKCε in human monocytes treated with
LPS or the bacterial chemoattractant N-formyl-methionine-leucine-phenylalanine
(FMLP) preventing the release of pro-inflammatory cytokines (256, 257). Altogether, the
TLR pathway mediates the recognition of pathogens and the release of pro-inflammatory cytokines through a complex regulatory network. Monocytes and macrophages act as sensors of for the detection of pathogens in the body triggering the apoptosis cascade when is needed.
1.9.3 Monocyte subsets
Monocytes are a heterogeneous population that exhibit differences in the expression of the cell surface antigens CD14 (receptor for the bacterial lipopolysaccharide, LPS) and the low affinity receptor for immunoglobulin G, CD16
(FcγIII receptor) (258). Classical monocytes express CD14 and lack the expression of
CD16 (CD14+CD16-), the non-classical monocytes express CD14 and CD16
(CD14+CD16+). The classical monocytes comprise 95% and the non-classical represent approximately 5% of the total monocytes. The non-classical monocytes can be further subdivided based on the expression level of CD14, into the CD14 high expression
(CD14++CD16+) and into CD14 low expression cells (CD14dimCD16+) (259). These two subpopulations represent 4.7% and 0.8%, respectively of the total monocytes.
Transcriptional analysis of CD14+CD16- and CD14+CD16+ gene expression showed 83% homology. Seventeen percent of CD14+CD16+ cells were found to be > 2-fold upregulated or downregulated compared to CD14+CD16- cells (260). These cells differ
40 in the expression of cytokine, adhesion molecules and chemokine receptors (CCR).
Among these genes, CD14+CD16+ cells were found to have higher expression of genes involved in FCγ receptor mediated phagocytosis, such as actin-related protein 2/3 complex (ARP2 and ARP3), tyrosine-protein kinase Lyn (LYN) and hemopoietic cell kinase (HCK). In contrast, the CD14+CD16- subset has an upregulation of genes involved in antimicrobial function, such as cathepsin G (CTSG), lysozyme C (LYZ), myeloperoxidase (MPO), and protein S100-A9 (S100A9) (261). The CD14+CD16+ cells have been associated with a more mature phenotype for the expression of the major histocompatibility complex receptors, such as HLA-DR, -DP and -DQ, normally expressed in macrophages (261, 262).
The function of the CD14+CD16+ monocytes remains unclear, but recent findings showed this subpopulation is expanded constituting ~ 20 - 30% monocytes in diseases like rheumatoid arthritis, asthma, tuberculosis, AIDS, sepsis, and cancer (263-265).
Patients with acute and chronic infections express high levels of CD14+CD16+ cells and treatment with antibiotics reduced this monocyte subpopulation correlating with clinical improvement (265). Based on these studies the CD16+ monocytes have been described to be pro-inflammatory. Elevated levels of CD16+ cells have been correlated with high expression of the pro-inflammatory cytokine TNF-α in the blood and low production of the anti-inflammatory cytokine IL-10 (266). More studies are needed to characterize the role of monocyte subsets to determine their role in inflammatory diseases.
41
Summary
This thesis focuses on the characterization of the mechanisms that regulate activation and execution of cell death. We observed that caspase-3 is phosphorylated during apoptosis by PKCδ. The domains of caspase-3 that bind and are phosphorylated by
PKCδ were identified. We discovered a novel PKCδ binding motif necessary for caspase-3 phosphorylation and binding. We identified and characterized the sites in caspase-3 phosphorylated by PKCδ and assessed their role in the proteolytic activity of caspase-3. We present in this work how the phosphorylation of caspase-3 has the potential to promote or inhibit its protease activity. This study presents a novel mechanism that could be of use for the development of therapies for the treatment of cancer of neurological disorders. In addition, we analyzed differences in apoptosis and
PKC expression in monocyte subsets and characterize their role in monocyte functions.
We identified differences in the release of TNF-α and PKCε expression between monocyte subsets. We show that PKCε expression plays an important role in the secretion of pro-inflammatory cytokines in monocytes exposed to LPS. A better understanding of the differences between monocyte subsets is needed for the development of new therapies for the treatment of inflammatory diseases. Our findings present a model for the regulation of cell death and survival pathways through the regulation of caspase-3 and members of the PKC family.
42
Caenorhabditis elegans Homo sapiens
Apoptotic stimuli Apoptotic stimuli
Egl-1 Bid, Bim
Ced-9 BCL-2
Ced-4 Apaf-1
Ced-3 Caspase-9
Caspase-3, -7
Apoptosis Apoptosis
Fig. 1.1 Conservation of the apoptotic pathway in Caenorhabditis elegans and Homo sapiens. In C. elegans, apoptosis-inducing stimuli promote the activation of the ced-3 inactive enzyme causing cell death. In mammals, cell death-inducing stimuli mediate first the activation of the initiator caspases, like caspase-9. Once activated, caspase-9 cleaves and activates the executioners, caspase-3 or 7, to cause apoptosis.
43
Fig. 1.2 Comparison of the domain structure of human caspases. All caspases are composed of 3 domains a prodomain, a middle domain and carboxyl-terminal domain. The QACXG motif identifies all members of the caspase family. The CARD or DED-domains are found in the prodomain of certain caspases.
44
QACXG
Name Initial processing site Substrate specificity P4 P3 P2 P1 S4 S3 S2 S1
Caspase-1 W F K D W E H D Caspase-2 D Q Q D D E D D Caspase-3 I E T D D E V D Caspase-4 W V R D W/L E H D Caspase-5 W V R D W/L E H D Caspase-6 T E V D V E H D Caspase-7 I Q A D D E V D Caspase-8 V E T D L E T D Caspase-9 P E P D L E H D Caspase-10 I E A D A E V D Caspase-12 A T A D * * * D Caspase-14 * * * * W E M D Ced-3 D S V D D E T D
Fig. 1.3 Initial processing site and substrate specificity of caspases. The first step in the activation of a caspase involves the cleavage at processing site at the carboxyl-terminal domain. Once caspases are activated they recognize 4 specific amino acid sequence in their cellular target protein. Initiator caspases have substrate specificity that allows their own autocatalytic cleavage. *, not specified. Modified figure from Degterev et al., 2003 (6).
45
Fig. 1.4 Apoptotic pathways (95).
46
Fig. 1.5 Comparison of the crystal structure of the active effector caspases. The structure of an inhibitor-bound caspase-3 (Protein Data Bank, PDB :2H5J), -6 (PDB : 3P45), and -7 (PDB: E3DR), which is representative of other caspase structures, are shown. The four surface loops (L1–L4 ) are shown for one of the monomers. Each caspase monomer is composed of two chains colored. The first monomer is colored in pink and yellow and the second monomer is colored in blue and green. Structures were generated in Pymol Molecular Graphic System.
47
Fig 1.6 Schematic representation of procaspase-7 activation. The active loops are in an inactive conformation unfavorable for catalysis. The interdomain loop is in a closed conformation and by cleavage at Asp198 rearranges the active site loops and produces a free N-terminus in the small subunit (L2’). These changes capacitates caspase-7 active site for substrate binding, which further induces drastic conformational changes (127).
48
Initiator caspase
QACRG DDD 28 175 pro p17 p12 Inactive caspase-3 L1 L2 L3 L4
28 pro p17 p12 1st Cleavage
28 2nd Autocatalytic pro p17 p12 cleavage
p12 p17 Active p17 Caspase-3 p12
pro Prodomain, 28 aa Cleavage site
p17 Large domain, 17 kDa QACRG Conserved motif (161-165)
p12 Small domain, 12 kDa DDD Safety catch (179-181) L1- L4 loops
Fig. 1.7 Mechanism of caspase-3 activation. Caspase-3 is synthesized as an inactive zymogen consisting of a prodomain (Pro), intermediate domain (p17) and a carboxyl-terminal domain (p12). Caspase-3 contains an active cysteine (C) closed by a conserved QACRG sequence and contains a safety catch sequence (DDD). Caspase-3 activation requires a two step cleavage. Active caspase-3 enzyme is a homodimer composed of two p17- and two p12-domains. The configuration of the active site requires 4 protruding loops (L1-L4). Upon activation, caspase-3 cleaves specific targets after aspartic acid residues to induce apoptosis.
49
Hinge Regulatory domain Catalytic domain H2N COOH
Classical PKCs C1 C2 C3 C4 α, βI, βII and γ
C2 C1 C3 C4 Novel PKCs like δ, ε, θ, η
Atypical PKCs C3 C4 ζ , ι/λ
C1 domain: phorbol ester C1 C3 C3 domain binding region
Lipid binding region C4 C4 domain
Phosphatidic acid C2 C2 domain: Ca2+ binding motif binding region
C2 C2 like domain like
Fig 1.8 PKC family. The members of the PKC family are divided in three subfamilies that include the classical, novel and atypical PKCs. The structural differences are shown (194).
50
C1 domain: phorbol ester C1 C4 C4 domain binding region
C2 C2 like domain Lipid binding region
C3 C3 domain
Fig 1.9 PKCδ structural domains and phosphorylation sites. Phosphorylation sites are represented by the single letter amino acid code. S and Y represent serine and tyrosine amino acids, respectively (194).
51
Fig 1.10 Model of PKCδ activation. Sequential activation and subcellular localization of PKCδ are illustrated in the model.
52
Hematopoietic stem cell (HSC)
Myeloid progenitor Lymphoid progenitor
Eosinophil Mega Erythroblast Basophil Granulocyte/ T cell B cell NK cell progenitor karyocyte progenitor monocyte progenitor
Neutrophil Monoblast
Thrombocytes Basophil
Monocyte
Erythrocytes Eosonophil
CD14+CD16- CD14+CD16+
Dendritic Macrophages cells
Fig. 1.11 Hematopoiesis. Modified from Dingli and Pacheco, 2010 (224).
53
LPS
LBP TLR4 MD2
MyD88
Caspase-9 IRAK
Caspase-3 IKK complex P Cell Death IκB NFκB
TNF-α IL-6, IL-1β NFκB
Fig. 1.12 TLR4 pathway. Simplified modified from Lu et al., 2008 (267).
54
Name Function Subfamily Caspase-1 Initiator Inflammation Caspase-4 Initiator Inflammation Caspase-5 Initiator Inflammation Caspase-12 Initiator Inflammation Caspase-13 Initiator Inflammation Caspase-2 Initiator Apoptosis Caspase-8 Initiator Apoptosis Caspase-9 Initiator Apoptosis Caspase-10 Initiator Apoptosis Caspase-3 Executioner Apoptosis Caspase-6 Executioner Apoptosis Caspase-7 Executioner Apoptosis Caspase-14 Executioner Differentiation
Table 1.1 Caspase family functions and classifications. Modified figure from Degterev et al., 2003 (6).
55
Name z-DEVD-FMK Ac-YVAD-CHO z-VAD-FMK XIAP c-IAP c-IAP2 Caspase-1 17 .76 2.5 >10 -4 n.d. n.d. Caspase-2 1710 >104 2400 n.d. n.d. n.d. Caspase-3 0.32 >104 43 0.7 108 35 Caspase-4 132 362 130 n.d. n.d. n.d. Caspase-5 205 163 5.3 n.d. n.d. n.d. Caspase-6 31 >104 98 >10 -4 n.d. n.d. Caspase-7 0.92 >104 39 20 .42 0.29 Caspase-8 1.6 352 2.5 >10 -4 n.d. n.d. Caspase-9 0.6 970 3.9 1 n.d. n.d. Caspase-10 12 408 n.d. >10 -4 n.d. n.d.
Table 1.2 Caspase inhibitors and their inhibitory constants. n.d.- Not determined Ki values. Values are expressed in nM. Modified figure from Degterev et al., 2003 (6).
56
Toll-Like Receptor Ligand
TLR1 + TLR2 Bacterial lipoproteins TLR2 + TLR6 Bacterial lipoproteins, lipoteichoic acid, yeast cell wall mannans TLR2 GPI anchors (parasites), bacterial porins, peptidoglycan, HMGBI TLR3 dsRNA TLR4 LPS, HSPs, HMGBI, some viral proteins TLR5 Bacterial flagellin TLR7 ssRNA (virus) TLR8 ssRNA (virus) TLR9 CpG-containing DNA (virus and bacteria) TLR10 Unknown TLR11 Unknown TLR12 Toxoplasma profilin TLR13 Unknown TLR14 Unknown
Table 1.3. TLR receptors and ligands (244).
57
Chapter 2
Materials and Methods
2.1 Reagents and chemicals
Bovine serum albumin (BSA), MG-132 proteasome inhibitor, etoposide, - mercaptoethanol, Tween 20, phenylmethylsulfonyl fluoride (PMSF), imidazole, chymostatin, pepstatin, leupeptin, antipain, ATP, bromophenol blue (BPB), cytochalasin
B, NP-40, sodium glycerophosphate, sodium pyrophosphate, sodium orthovanadate, sodium fluoride, Triton-X-100, paraformaldehyde, N,N,N’,N’-Tetra-methyl-ethylene- diamine (TEMED), LPS, Histopaque-1077, and 4-6-diamidino-2-phenylindole (DAPI) were obtained from Sigma (St. Louis, MO). Restriction enzymes, including BamHI,
EcoRI and XhoI were obtained from New England Biolabs (Ipswich, MA).
The caspase inhibitors, IETD-FMK, LEHD-FMK and DEVD-FMK, the substrate
IETD-AFC, and the Annexin V staining buffer, were obtained from BD Biosciences (San
Jose, CA). The caspase substrates, DEVD-AFC and LEHD-AFC were obtained from MP
Biomedicals (Solon, OH). Histone 2B (H2B) was obtained from Roche (Indianapolis,
IN). Reagents obtained from Research Products International corporation (RPI, Prospect,
IL) were: agarose, K2HPO4, KCl, Na2HPO4, Na2H2PO4, EDTA, NaCl, KH2PO4,sucrose,
DTT, MgCl2, MnCl2, SDS, Tris, urea, PIPES, ampicillin and kanamycin.
Monocyte isolation products, including CD16 and CD14 magnetic microbeads,
LS-column, LD-columns were obtained from Miltenyi Biotec (Auburn, CA). Additional 58 reagents, including yeast extract, tryptone and glycerol were received from EMD
(Gibbstown, NJ). HEPES, Coomassie brilliant blue R-250, methanol and glacial acetic acid, were purchased from Fisher Scientific (Pittsburgh, PA).
The antibodies used in this work are listed in Table 2.1.
Table 2.1. List of antibodies
Catalog Antibody Clone Type number Company Location anti-active caspase-3 Asp175 Polyclonal 9661S Cell Signaling Danvers, MA
anti-active-caspase-3- * Polyclonal 559341 Cell Signaling Danvers, MA
anti-FITCalexa- 633 - polyclonal A21070 Invitrogen Carlsbad, CA
anti-bid FL-195 Polyclonal SC-11423 Santa Cruz Santa Cruz, CA
anti-caspase-3 19 Monoclonal 610323 BD Biosciences San Jose, CA
anti-CD14-APC * Polyclonal 555399 BD Biosciences San Jose, CA
anti-CD16-PE * Monoclonal 130-091- Miltenyi Biotec Auburn, CA
anti-CD16-PE * Monoclonal 130245-091 - Miltenyi Biotec Auburn, CA
Anti-cleaved PARP F21- Monoclonal 558576245 BD Biosciences San Jose, CA
anti-GAPDH FL852-335 Polyclonal SC-25778 Santa Cruz Santa Cruz, CA
anti-IgG * Monoclonal SC-2025 Santa Cruz Santa Cruz, CA
anti-IgG * Polyclonal SC-2027 Santa Cruz Santa Cruz, CA
anti-IgG-PE G18- Polyclonal 555787 BD Biosciences San Jose, CA
anti-mouse-HRP 145* Monoclonal NA931 Amersham Arlington Heights,
anti-PKC alpha (PKCα) C-20 Polyclonal SC-208 Santa Cruz Santa Cruz,IL CA
anti-PKC beta I (PKCβI) C16 Polyclonal SC-209 Santa Cruz Santa Cruz, CA
anti-PKC beta II C18 Polyclonal SC-210 Santa Cruz Santa Cruz, CA
anti-PKC(PKCβ deltaII )(PKCδ) C-20 Polyclonal SC-937 Santa Cruz Santa Cruz, CA
59
Table 2.1 List of antibodies, continued anti-PKC epsilon (PKCε) C-15 Polyclonal SC-214 Santa Cruz Santa Cruz, CA
anti-PKC eta (PKCη) C15 Polyclonal SC-215 Santa Cruz Santa Cruz, CA anti-PKC gamma (PKCγ) C19 Polyclonal SC-211 Santa Cruz Santa Cruz, CA
anti-PKC iota/lambda H-76 Polyclonal SC-11399 Santa Cruz Santa Cruz, CA
anti-PKC(PKCι/λ) theta (PKCθ) C-18 Polyclonal SC-212 Santa Cruz Santa Cruz, CA
anti-PKC zeta (PKCζ) C20 Polyclonal SC-215 Santa Cruz Santa Cruz, CA
anti-PKCδ C20 Polyclonal SC-937 Santa Cruz Santa Cruz, CA
anti-rabbit-HRP * Polyclonal NA934 Amersham Arlington Heights,
anti-Xpress * Monoclonal R91025 Invitrogen Carlsbad,IL CA
anti-α-fodrin * Monoclonal MAB1622 Millipore Charlottesville, VA
anti-β-tubulin AA2 Monoclonal 05-661 Millipore Charlottesville, VA
2.2 Cloning and mutagenesis
The caspase-3-FL in pQE31 vector (AB # 468, Table 2.3.1) was used as a template for PCR to create point mutations in the identified phosphorylation sites utilizing the Quick Change Site-directed Mutagenesis Kit (Cat.-No.: 200515, Stratagene,
Cedar Creek, TX). The primers utilized are listed in Table 2.2. The mutations were confirmed by sequencing (Macrogen, Korea) utilizing the primer PAO-37 (Table 2.2).
The clones were subsequently subcloned into pENTR/D-TOPO vector (Table 2.3.2)
(Cat.-No.: K2400-20, Invitrogen) utilizing the primers PAO-120 and PAO-188 (Table
2.2) to introduce BamHI/EcoRI restriction sites. The clones were screened and analyzed by sequencing. Subsequently the mutant caspase-3 was subcloned into the mammalian expression vector pcDNA-4/His Max vector (Xpress) and used for expression of Xpress- tagged proteins (Table 2.3.3). 60
Caspase-3 potential PKCδ binding sites, named caspase-3-PBM1 and 2 (Fig. 3.3), were mutated and replaced, respectively. Caspase-3-PBM1-FL (AB-831)was created by replacing the four amino acid SGIS (aa 29 - 32) in caspase-3-FL (AB-419) by site directed mutagenesis (Stratagene, Cedar Creek, TX) using the primer set PAO-470:PAO-
471 (Table 2.2). PBM2 was generated by deletion of sequences QACRGTEL (aa 161-
168) in caspase-3-FL (AB-419) first two XhoI sites flanking the motif using the primer set: PAO-405:PAO-406 and PAO-407:PAO-408 (Table 2.2). Next, the DNA was digested with XhoI and a fragment of 490 bp was re-ligated using T4-Ligase (Invitrogen,
Carlsbad, CA) creating the caspase-3-PBM2-FL (AB-848). Colonies were selected and the DNA was purified and screened by restriction digestion with BamHI/EcoRI enzymes.
To generate caspase-3-PBM1-prop17 (AB-836), the caspase-3-PBM1-FL (AB-831) was used as a template to amplify by PCR utilizing the primer set PAO-120:PAO-188 (Table
2.2). The DNA from the caspase-3-PBM2-FL (AB-848) was utilized as a template to amplify caspase-3-PBM2-prop17 (AB-807) utilizing the primer set PAO-120:PAO-396
(Table 2.2).
61
Table 2.2 Primer List
Number Purpose Primer Sequence (5'-3') PAO-37 Sequencing Forward CGGATAACAATTTCACACAG PAO-69 S36A Forward GA ATA TCC CTG GAC AAC GCG TAT AAA ATG GAT TAT C PAO-70 S36A Reverse GAT AAT CCA TTT TAT ACG CGT TGT CCA GGG ATA TTC PAO-71 S36D Forward GAATATCCCTGGACAACGATTATAAAATGGATTATC PAO-72 S36D Reverse GATAATCCATTTTATAATCGTTGTCCAGGGATATTC PAO-77 T77A Forward AGCAAACCTCAGGGAAGCTTTCAGAAACTTGAAAT PAO-78 T77A Reverse ATTTCAAGTTTCTGAAAGCTTCCCTGAGGTTTGCT PAO-79 T77D Forward GCAAACCTCAGGGAAGACTTCAGAAACTTGAAAT PAO-80 T77D Reverse ATTTCAAGTTTCTGAAGTCTTCCCTGAGGTTTGC To add PAO-120 Forward CACCGGATCCATGGAGAACACTGAAAAC BamH1 PAO-129 S12D Forward TCA GTG GAT TCA AAA GAC ATT AAA AAT TTG GAA PAO-130 S12D Reverse TTC CAA ATT TTT AAT GTC TTT TGA ATC CAC TGA PAO-131 S32A Forward ATG GAC TCT GGA ATA GAC CTG GAC AAC AGT TAT PAO-132 S32A Reverse ATA ACT GTT GTC CAG GGC TAT TCC AGA GTC CAT PAO-135 S58A Forward AAG AAT TTT CAT AAG GCC ACT GGA ATG ACA TCT PAO-136 S58A Reverse AGA TGT CAT TCC AGT GGC CTT ATG AAA ATT CTT PAO-137 S58D Forward AAG AAT TTT CAT AAG GAC ACT GGA ATG ACA TCT PAO-138 S58D Reverse AGA TGT CAT TCC AGT GTC CTT ATG AAA ATT CTT PAO-139 T59A Forward AAT TTT CAT AAG AGC GCT GGA ATG ACA TCT CGG PAO-140 T59A Reverse CCG AGA TGTCAT TCC AGC GCT CTT ATG AAA ATT PAO-141 T59D Forward AAT TTT CAT AAG AGC GAT GGA ATG ACA TCT CGG PAO-142 T59D Reverse CCG AGA TGTCAT TCC ATC GCT CTT ATG AAA ATT PAO-143 T67A Forward ACA TCT CGG TCT GGT GCA GAT GTC GAT GCA GCA PAO-144 T67A Reverse TGC TGC ATC GAC ATC TGC ACC AGA CCG AGA TGT PAO-145 T59D Forward ACA TCT CGG TCT GGT GAT GAT GTC GAT GCA GCA PAO-146 T59D Reverse TGC TGC ATC GAC ATC ATC ACC AGA CCG AGA TGT PAO-151 S12G Forward TCA GTG GAT TCA AAA GGC ATT AAA AAT TTG GAA PAO-152 S12G Reverse TTC CAA ATT TTT AAT GCC TTT TGA ATC CAC TGA Add EcoR1 PAO-188 Reverse GAA TTC TTA GTC TGT CTC AAT GCC ACA GTC CAG TTC in casp-3-
prop17 Add EcoR1 PAO-189 Reverse GAA TTC TTA GTG ATA AAA ATA GAG TTC in casp-3-
prop17
62
Table 2.3 Clones in pQE31 vector
Clone ID Description Restriction sites
AB-468 Caspase-3-FL-WT-PQE31 BamHI 1/Sal1 BamHI 1/Sal1 AB-488 Casp-3-FL-T59A-PQE31 BamHI 1/Sal1 AB-504 Casp-3-FL-S12G-PQE31 BamHI 1/Sal1 AB-528 Casp-3-FL-S36A-PQE31 BamHI 1/Sal1 AB-534 Casp-3-FL-T67A-PQE31 BamHI 1/Sal1 AB-538 Casp-3-FL-T67D-PQE31 BamHI 1/Sal1 AB-540 Casp-3-FL-S12G/S36A-PQE31 BamHI 1/Sal1 AB-541 Casp-3-FL-S12G/T59A-PQE31 BamHI 1/Sal1 AB-542 Casp-3-FL-S36D-PQE31 BamHI 1/Sal1 AB-543 Casp-3-FL-S12D-PQE31 BamHI 1/Sal1 AB-545 Casp-3-FL-T77D-PQE31 BamHI 1/Sal1 AB-546 Casp-3-FL-T59D-PQE31 BamHI 1/Sal1 AB-560 Casp-3-FL-S12G/ S36A/T59A/ T67A/T77A (M5-A/G)-PQE31 BamHI 1/Sal1 AB-561 Casp-3-FL-T77A-PQE31 BamHI 1/Sal1 AB-563 Casp-3-FL-S12D/S36D-PQE31 BamHI 1/Sal1 AB-564 Casp-3-FL-S12D/T59D-PQE31
63
Table 2.4 Clones in pENTR/D-TOPO vector
Restriction Clone ID Description sites used BamH1/EcoR1 AB-536 Casp-3-FL-S36A, T59A-pENTR/D-TOPO BamH1/EcoR1 AB-586 Casp-3-FL-S12D-pENTR/D-TOPO Casp-3-prop17-S12G/ S36A/ T59A/ T67A/ T77A (M5A)-pENTR/D- BamH1/EcoR1 AB-596 TOPO BamH1/EcoR1 AB-660 Casp-3-prop17-S12D/ S36D-pENTR/D-TOPO BamH1/EcoR1 AB-661 Casp-3-prop17-S12G/ S36A -pENTR/D-TOPO BamH1/EcoR1 AB-562 Casp-3-FL-S36D, T59D-pENTR/D-TOPO BamH1/EcoR1 AB-666 Casp-3-FL-S36D-pENTR/D-TOPO BamH1/EcoR1 AB-667 Casp-3-FL-S36A-pENTR/D-TOPO BamH1/EcoR1 AB-668 Casp-3-FL-S12D/ S36D-pENTR/D-TOPO BamH1/EcoR1 AB-669 Casp-3-FL-S12G/ S36A -pENTR/D-TOPO BamH1/EcoR1 AB-745 Casp-3-FL-T59A-pENTR/D-TOPO BamH1/EcoR1 AB-746 Casp-3-FL-T67A-pENTR/D-TOPO BamH1/EcoR1 AB-747 Casp-3-FL-T77A-pENTR/D-TOPO BamH1/EcoR1 AB-748 Casp-3-FL-S12G/ S36A / T59A-pENTR/D-TOPO BamH1/EcoR1 AB-749 Casp-3-FL-S12G/ S36A/ T59A/ T67A/ T77A (M5A)-pENTR/D-TOPO Casp-3-FL-S12G/ S32A/ S36A/ S58A/ T59A/ T67A/ T77A (M7A)- BamH1/EcoR1 AB-750 pENTR/D-TOPO BamH1/EcoR1 AB-751 Casp-3-FL-S12D/ S36D/ T59D/ T67D/ T77D (M5D)-pENTR/D-TOPO BamH1/EcoR1 AB-753 Casp-3-FL-S12G/ S36A / T67A/ T77A (M4A)-pENTR/D-TOPO BamH1/EcoR1 AB-755 Casp-3-prop17-S36A-pENTR/D-TOPO BamH1/EcoR1 AB-756 Casp-3-prop17-S36D-pENTR/D-TOPO BamH1/EcoR1 AB-757 Casp-3-prop17-S12G/ S36A , T59A-pENTR/D-TOPO BamH1/EcoR1 AB-758 Casp-3-prop17-S12D/ S36D/ T59D-pENTR/D-TOPO BamH1/EcoR1 AB-759 Casp-3-prop17-S12D/ S36D/ T59D/ T67D/ T77D (M5D)-pENTR/D-TOPO AB-760 Casp-3-prop17-S12G/ S32A/ S36A/ S58A/ T59A/ T67A/ T77A (M7A)- BamH1/EcoR1 pENTR/D-TOPO
64
Table 2.4 Clones in pENTR/D-TOPO vector vector, continued
BamH1/EcoR1 AB-761 Casp-3-prop17-S12G-pENTR/D-TOPO BamH1/EcoR1 AB-762 Casp-3-prop17-S12D-pENTR/D-TOPO BamH1/EcoR1 AB-766 Casp-3-FL-S10A-pENTR/D-TOPO BamH1/EcoR1 AB-767 Casp-3-prop17-S10A-pENTR/D-TOPO BamH1/EcoR1 AB-585 Casp-3-FL-S12G-pENTR/D-TOPO BamH1/EcoR1 AB-763 Casp-3-prop17-T59A-pENTR/D-TOPO BamH1/EcoR1 AB-764 Casp-3-prop17-T67A-pENTR/D-TOPO BamH1/EcoR1 AB-765 Casp-3-prop17-T77A-pENTR/D-TOPO Casp-3-FL-S12G/ S32A/ S36A/ S58A/ T59A/ T67A/ T77A (M7A)- BamH1/EcoR1 AB-750 pENTR/D-TOPO BamH1/EcoR1 AB-808 Casp-3-FL-S32A-pENTR/D-TOPO BamH1/EcoR1 AB-809 Casp-3-FL-S32D-pENTR/D-TOPO
Table 2.5 Clones in pcDNA-4/His Max vector (Xpress)
Clone Restriction Description ID sites used BamH1/EcoR1 AB-419 Caspase-3-FL-WT-pcDNA 4/His Max Xpress BamH1/EcoR1 AB-421 Caspase-3-prop17-pcDNA 4/His Max Xpress BamH1/EcoR1 AB-422 Caspase-3-p17-pcDNA 4/His Max Xpress BamH1/EcoR1 AB-423 Caspase-3-p12-pcDNA 4/His Max Xpress BamH1/EcoR1 AB-744 Casp-3-FL-S36A-pcDNA 4/His Max Xpress BamH1/EcoR1 AB-752 Casp-3-FL-S12D/ S36D/ T59D/ T67D/ T77D (M5D)-pcDNA 4/His Max Xpress BamH1/EcoR1 AB-754 Casp-3-FL-S12G/ S36A / T67A/ T77A (M4A)-pcDNA 4/His Max Xpress BamH1/EcoR1 AB-580 Casp-3-FL-S12G-pcDNA 4/His Max Xpress BamH1/EcoR1 AB-581 Casp-3-FL-S12D-pcDNA 4/His Max Xpress
65
Table 2.5 Clones in pcDNA-4/His Max vector (Xpress), continued
BamH1/EcoR1 AB-582 Casp-3-prop17-S12G-pcDNA 4/His Max Xpress BamH1/EcoR1 AB-583 Casp-3-prop17-S12D-pcDNA 4/His Max Xpress Casp-3-prop17-S12G/ S36A/ T59A/ T67A/ T77A (M5A)-pcDNA 4/His Max BamH1/EcoR1 AB-602 Xpress Casp-3-prop17-S12G/ S32A/ S36A/ S58A/ T59A/ T67A/ T77A (M7A)- BamH1/EcoR1 AB-604 pcDNA 4/His Max Xpress BamH1/EcoR1 AB-662 Casp-3-prop17-S12D/ S36D-pcDNA 4/His Max Xpress BamH1/EcoR1 AB-663 Casp-3-prop17-S36D-pcDNA 4/His Max Xpress BamH1/EcoR1 AB-664 Casp-3-prop17-S36A-pcDNA 4/His Max Xpress BamH1/EcoR1 AB-665 Casp-3-prop17-S12G/ S36A -pcDNA 4/His Max Xpress BamH1/EcoR1 AB-670 Casp-3-FL-S36D-pcDNA 4/His Max Xpress BamH1/EcoR1 AB-671 Casp-3-FL-S12D/ S36D-pcDNA 4/His Max Xpress BamH1/EcoR1 AB-672 Casp-3-FL-S12G/ S36A -pcDNA 4/His Max Xpress Casp-3-prop17-S12D/ S36D/ T59D/ T67D/ T77D (M5D)-pcDNA 4/His Max BamH1/EcoR1 AB-713 Xpress BamH1/EcoR1 AB-714 Casp-3-prop17-S12D/ S36D/ T59D-pcDNA 4/His Max Xpress BamH1/EcoR1 AB-776 Casp-3-FL-S12G/ S36A/ T59A/ T67A/ T77A (M5A)-pcDNA 4/His Max Xpress BamH1/EcoR1 AB-770 Casp-3-prop17-S10A-pcDNA 4/His Max Xpress BamH1/EcoR1 AB-777 Casp-3-prop17-T59A-pcDNA 4/His Max Xpress BamH1/EcoR1 AB-778 Casp-3-prop17-T67A-pcDNA 4/His Max Xpress BamH1/EcoR1 AB-779 Casp-3-prop17-T77A-pcDNA 4/His Max Xpress BamH1/EcoR1 AB-807 Casp-3-prop17-PBM2-pcDNA 4/His Max Xpress BamH1/EcoR1 AB-831 Casp-3-FL-PBM1-pcDNA 4/His Max Xpress BamH1/EcoR1 AB-836 Casp-3-prop17-PBM1-pcDNA 4/His Max Xpress BamH1/EcoR1 AB-848 Casp-3-FL-PBM2-pcDNA 4/His Max Xpress
2.3 Tissue culture
Hela and MCF-7 cells were obtained from the American Type Culture collection
(ATCC) and were cultured in DMEM (Cat No.: 11995, Invitrogen) at 37°C in 5% CO2. 66
The growth media was supplemented with 5% of heat inactivated fetal bovine serum
(FBS, Cat.No.: S11150, Atlanta Biologicals, Lawrenceville, GA) and 1% penicillin/streptomycin (Cat-No.: 15070063, Invitrogen). Mouse embryonic fibroblasts were cultured in DMEM (Cat.-No.: 11995-065, Invitrogen). Primary human monocytes were cultured in RPMI 1640 (Cat-No.: 11875119, Invitrogen) at 37°C in 5% CO2, in the absence of growth factors or antibiotics.
HeLa cells were transiently transfected with indicated clones (chapter 3). Cells were seeded at a confluency of 40% in 10 cm dishes containing 10 ml of DMEM (Cat.-
No.: 11995, Invitrogen), supplemented with 5% FBS and 1% penicillin/streptomycin
(Cat-No.: 15070063, Invitrogen). A transfection mix containing 8 µg of DNA and 20 µl of Lipofectamine 2000 (Cat-No.: 11668-019, Invitrogen), was incubated with 0.8 ml of
DMEM for 30 min, at room temperature. The cells were rinsed with PBS and 4.2 ml of
DMEM were added. After incubation, the DNA/Lipofectamine mix was added drop- wise to the cells. The cells were incubated for 5 h and after that 5 ml of DMEM 5% FBS were added. Cells were incubated for additional 24 h. HeLa cells transfected with caspase-3 p17 and p12 domains were treated with 20 μM MG132 for 4 h after transfection (Cat.No: BML-PI102-0005, Enzo Life Sciences, Farmingdale, NY) proteasome inhibitor to avoid the degradation (162).
MCF7 cells were transiently transfected with the Xpress-tagged caspase-3-WT and phospho-mutants. The cells were seed into 35 mm plates at 40% confluency in
DMEM, supplemented with 5% FBS and 1% penicillin/streptomycin (40% confluence).
Twenty four hours later, a transfection mix containing 2.5 µg of DNA and 0.75 µl of the
67
Xfect polymer (Cat-No.: 631317 , Clontech), was incubated with 0.2 ml of Xfect buffer for 10 min, at room temperature. The cells were rinsed with PBS and 1 ml of DMEM was added. After incubation, the DNA/Xfect mix was added drop-wise to the cells. The cells were incubated for 4 h. The media was replaced with 1 ml of DMEM supplemented with 10% FBS. The cells were incubated for 24 h before treatment.
MCF7 cells transfected with caspase-3-WT or mutants were treated with 200 µM etoposide (Cat-No.: E1383, Sigma) for various amounts of times. The effect of specific mutations in apoptosis was analyzed by Western blot, DEVD-AFC and flow cytometry analysis, explained in subsequent sections of the materials and methods.
2.4 Isolation of mouse embryonic fibroblasts
Mouse embryonic fibroblast (MEF) from WT and PKCδ-/- C57BL/6 mice were kindly provided by Dr. Mary Reyland laboratory, Denver, Colorado. The C57BL/6 and
PKCβ-/- mice utilized for MEF isolation were kindly provided by Drs. Wei Huang and
Kamal Mehta (The Ohio State University, Columbus, OH). The protocol for MEF isolation was modified from the MEF isolation protocol kindly provided Dr. Prabakaran
Nagarajan (Columbus, OH). MEF from WT and PKCβ-/- C57BL/6 mice were obtained from embryos between 12.5-14.5 days of gestation were extracted and rinsed in PBS.
The skull and liver were dissected (268). A 21 G syringe was placed inside a tube containing 2 ml of DMEM culture media (Cat.-No.: 11995-065, Invitrogen) supplemented with 20% FBS and 1% penicillin/streptomycin (Cat-No.: 15070063,
Invitrogen). Each embryo was placed inside the syringe and was minced by six cycles of
68 aspiration and ejection. Cells were plated into a 10 cm culture dish containing 8 ml of
DMEM supplemented with 20% FBS, and were allowed to adhere for 48 h. Cells were split in a 1:1 ratio in DMEM supplemented with 5% FBS every 3-4 days until they reached passage 5. At passage 5 the MEF were utilized for experiments and were viable until passage 10.
2.5 Extract preparation and immunoblotting
Cells were collected by centrifugation and rinsed once with PBS and the cell pellet was snap-frozen in liquid nitrogen. MCF-7 and monocyte cell extracts (were prepared by lysis of 2 x 106 cells in 20 µl of Nonidet P-40 (NP-40) buffer (10 mM Tris - pH 7.5, 0.5% NP-40, 5 mM EDTA – pH 8, 10 mM sodium glycerophosphate, 5 mM sodium pyrophosphate, 1 mM sodium orthovanadate, 1 mM DTT, 0.1 mM PMSF and 2
µg/ml protease inhibitors). HeLa cell extract were prepared by lysis of 2 x 106 cells in 20
µl of Tween 20 buffer (50 mM Hepes - pH 7.5, 150 mM NaCl, 0.1% Tween 20, 1 mM
EDTA, 2.5 mM EGTA, 10% glycerol, 50 mM NaFl, 10 mM sodium glycerophosphate, 5 mM sodium pyrophosphate, 1 mM sodium orthovanadate, 1 mM DTT, 0.1 mM PMSF and 2 µg/ml protease inhibitors). Cells were lysed by continuous vortex at 4°C. The lysates were centrifuged at 13,200 rpm for 10 min at 4 °C. The supernatant were aliquoted and kept at – 80°C for future use.
2.6 Protein quantitation and Western blot analysis
The protein concentration was determined using the colorimetric Bradford assay from BioRad. The Bradford reagent was diluted 1:5 in water and 199 µl of the solution 69 were incubated with 1 µl of bovine serum albumin (BSA) standard or samples, for 5 min.
BSA was utilized as a standard control in a range of 0.5 to 8 μg. The absorbance at 595 nm was measured in the Victor X3 micro plate reader (Perkin Elmer, Waltham,
Massachusetts) and the protein concentration was determined utilizing the Work Out software (Perkin Elmer, version 2.5, 2008).
Samples analyzed by Western blot were boiled for 5 min at 95°C in the presence of 5 x Laemmli buffer (250 mM Tris, pH 6.8, 10% SDS, 50% glycerol, 0.5% bromophenol blue (BPB), and 1.78 M β-mercaptoethanol). The proteins were resolved by a one dimension polyacrylamide gel electrophoresis. The gels were composed of a
5% stacking gel (acrylamide/bisacrylamide ratio 29:1, 0.13 M Tris-Cl, pH 6.8, 0.1%
SDS, 0.1 ammonium persulfate (APS), and 0.775 mg/ml TEMED) and 15% resolving gel
(acrylamide/bisacrylamide ratio 29:1, 0.39 M Tris-Cl, pH 8.8, 0.1% SDS, 0.1 APS, and
0.31 mg/ml TEMED). All gels were run in a BioRad mini-protean II system in running buffer (25 mM Tris and 190 mM Glycine). The samples and protein marker (SeeBlue
Plus 2, Cat-No.: LC5925, Invitrogen) were resolved by running at 100 V for 10 min, followed by 150 V for 90 min. The gels were either stained with Coomassie staining solution (0.25% Coomassie brilliant blue R-250, 50% methanol, and 10% glacial acetic acid) or transferred to nitrocellulose membranes (0.20 µm, Whatman, USA) for 70 min on transfer buffer (25 mM Tris, pH 8.3, 192 mM glycine, 20% methanol). The membranes were blocked in a solution containing 5% powder milk in TBS buffer (10 mM Tris-Cl, pH 7.6, 0.15 M NaCl, and 0.5% Tween 20) for 0.5 h at room temperature.
Membranes were incubated overnight at 4°C with the primary antibodies used in a
70
1:1000 to 1:4000 dilutions in a solution containing TBS buffer and either 0.25% BSA or
5%powder milk. After incubation the membranes were rinsed 3 times with 10 ml of TBS buffer. The secondary antibodies were diluted in a 1:4000 ratio in TBS buffer containing either 0.25% BSA or 5% powder milk. Secondary antibodies linked to horseradish peroxidase (HRP) were incubated with the membranes for 1 h at room temperature.
Membranes were rinsed 3 times with 10 ml of TBS buffer. The proteins were visualized utilizing the enhanced chemiluminescence solution (ECL, Cat-No.: RPN2106,
Amersham).
2.7 Immunoprecipitations and in vitro kinase assays
Immunoprecipitation reactions were performed utilizing cell extracts at a concentration of 1 µg/µl were prepared in a Tween 20 buffer 50 mM Hepes - pH 7.5, 150 mM NaCl, 0.1% Tween 20, 1 mM EDTA, 2.5 mM EGTA, 10% glycerol, 50 mM sodium fluoride, 10 mM sodium glycerophosphate, 5 mM sodium pyrophosphate, 1 mM sodium orthovanadate, 1 mM DTT, 0.1 mM PMSF and 2 µg/ml protease inhibitors). Anti-Xpress
(Cat-No.: R91025, Invitrogen) or anti-PKCδ (Cat-No.: SC-937, Santa Cruz
Biotechnology, Santa Cruz CA) or IgG isotype control antibodies were utilized (Table
2.1). The immunoprecipitation reactions were incubated for 14 h at 4°C with constant rotation. The reactions were incubated with 30 µl of protein G agarose [50% slurry] (Cat-
No.: 15920-010, Invitrogen) for 1 h at 4°C with constant rotation. The beads were rinsed
4 times with 0.7 ml of Tween 20 buffer and resuspended in 20 μl of 2X Laemmli buffer boiled at 95ºC for 5 min and were resolved by SDS-PAGE.
71
Alternatively, the IPs were utilized for subsequent kinase assay reactions. The
IPs were rinsed twice with 700 μl of kinase buffer (25 mM HEPES, pH 7.4, 10 mM
MnCl2, 1 mM MgCl2), and resuspended in 20 µl of kinase buffer supplemented with 500 nM ATP and 10 µCi of [γ-32P] ATP (Perkin Elmer), and a mixture of phosphatidylserine/ diacylglycerol were added at a final concentration of 200 µg/ml and 20 μg/ml, respectively. For the immunoprecipitation and kinase reaction in MEF, 5 µg of H2B were added as exogenous substrate (Fig. 3.1). In all cases the phosphorylation reaction were carried out at 37°C for 1 h. The reactions were stopped by addition of 5X Laemmli buffer, boiled for 5 min at 95ºC, resolved by SDS-PAGE, and the samples were analyzed by Western blot and autoradiography.
2.8 Protein expression and purification
To obtain caspase-3 pure proteins, plasmids containing 6xHis-tag caspase-3
(Table 2.3.1) were transformed into M15 Escherichia coli (Cat.-No.: 34210, Qiagen,
Valencia, CA). M15 cells contain a pREP4 plasmid that encodes a Lac-repressor to regulate protein expression. For protein expression, starting cultures of 20 ml were grown for 14 h at 37ºC in Terrific Broth media [2% glucose, 2.4% yeast extract, 1.2% tryptone, 4% glycerol, 0.17 M KH2PO4, 0.72 M K2HPO4], containing 100 μg/ml ampicillin and 30 μg/ml kanamycin as selection markers. The overnight cultures were diluted in Terrific Broth to an optical density (OD600) of 0.1. Cultures were grown until they reached an optical density of OD600 of 0.5, for approximately 2 h at 37ºC. Protein expression was induced by incubation with 1 mM isopropyl 1-thio-β-D- galactopyranoside (IPTG, Gold Biotechnology, St. Louis, MO) for 30 min at 20ºC. The 72 cultures were harvested by centrifugation at 6,000 rpm, for 5 min at 4ºC. Bacterial pellets were resuspended in 10 ml of sonication buffer (50 mM sodium phosphate, 150 mM sodium chloride, 1% Tween 20, 5 mM β-mercaptoethanol, 0.1 mM PMSF, and 2
µg/ml CLAP) and lysed by sonication (10 seconds per cycle, repeated 25 times, output: 8, dutycycle: 80, Branson Sonifier 450, ½ inch disrupter horn, VWR Scientific, West
Chester, PA). The lysates were then centrifuged at 12,000 rpm for 10 min at 4ºC, and the supernatants were treated with 10 µg/ml RNAse (Invitrogen, Carlsbad, CA) and 5 µg/ml
DNAse (Invitrogen, Carlsbad, CA), and were filtered with miracloth (Calbiochem, San
Diego, CA). The lysates were incubated with 125 μl of nickel-nitrilotriacetic acid- agarose beads (Cat-No.: 30210, Qiagen) containing 50% slurry for 90 min with constant agitation at 4ºC. The supernatant containing Ni+ beads were loaded into a column. The beads were rinsed twice with 5 ml sonication buffer followed by two rinses with 5 ml washing buffer (50 mM HEPES pH 7.4, 300 mM NaCl, 10% glycerol, 1% Tween 20, 1 mM PMSF, and 2 µg/ml CLAP). The proteins were eluted with a discontinuous imidazole gradient of 30 mM, 50 mM, and 100 mM imidazole in washing buffer. The proteins were eluted by constant addition of 1 ml of each buffer, from a total volume of
10 ml. Elutions were analyzed by SDS-PAGE and stained with Coomassie Brilliant Blue
R-250 solution (0.25% (w/v) Coomassie brilliant blue R-250, 50% (v/v) methanol and
10% (v/v) glacial acetic acid). The elutions containing the proteins of interest were dialyzed two times for 3 h at 4°C, with constant stirring, in caspase-3 reaction buffer (50 mM HEPES, pH 7.4, 50 mM NaCl, 10% sucrose, 0.1 mM PMSF, and 1 mM DTT) in a
1:1000 dilution. Dialyzed proteins were snap-frozen in liquid nitrogen and stored at -
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80°C for future use. The proteins were stable for 7 cycles of freeze and thaw as determined by the DEVD-AFC activity assay.
2.9 Activation of caspase-3 by recombinant caspase-9
For activation assays, 50 ng recombinant purified his-tagged caspase-3-WT or phospho-mutants were incubated with 40 units of recombinant caspase-9 (Cat-No.:
SE173, MP Biological, Solon, OH) at 37°C in a caspase-3 reaction buffer (50 mM
HEPES, pH 7.4, 50 mM NaCl, 10% sucrose, 0.1 mM PMSF, and 1 mM DTT) in a total volume of 5 μl. The samples were collected over time and snap-frozen in liquid nitrogen and stored at -70°C for future use. The activity of caspase-3 was measured through the cleavage of the tetra-peptide DEVD-AFC (activity assay) or by Western blot analysis.
The DEVD assay was performed with 1 μl of the reaction. For the Western blot analysis,
2.5 μl of the reaction were run on the gel and the immunoblot was performed using anti- caspase-3 antibody (Cat-No.: 610323, BD Biosciences) (Table 2.1).
2.10 Activation of recombinant caspase-3 by MCF7 S100 cell extracts
To obtain cell free lysates referred as S100 lysates, MCF-7 cells were lysed in the
KPM buffer (50 mM Pipes-pH 7.0, 50 mM KCl, 100 µM PMSF, 5 mM EGTA, 1 mM
DTT, 2mM, 2 µg/ml CLAP, 10 µg/ml cytochalasin B) by 5 rounds of freezing and thawing (5 times) as previously described. This protocol is modified from Cullen et al.,
2008 (269). Briefly, ten million cells were lysed in 500 μl of KPM buffer. The lysates were centrifuged at 55,000 rpm for 1 h at 4°C, supernatants comprising cytosolic 74 fractions, were collected, aliquoted and snap-frozen in liquid nitrogen and kept at -70 for future use.
For the activation of caspase-3 using cell free systems, 40 ng of the recombinant caspase protein were incubated with 75 μg of S100 lysates and were supplemented with
50 µM cytochrome c, and 1 μM dATP in KPM buffer; to reach a total volume of 5 µl.
The reactions were incubated at 37°C for 0, 1, 3, and 4 h. The samples were collected over time and were snap-frozen in liquid nitrogen and stored at -70°C for future use.
Caspase-3 activity and processing were determined by the DEVD-AFC activity assay
(Section 2.13) or immunoblots.
2.11 Caspase Activity assays
Caspase activities were generated as previously described (229). Briefly, caspase-
3 and caspase-9 activities were determined by incubation of 10 µg of cell extracts or 10 ng of recombinant caspase-3 with 20 μM of 7-amino-4-trifluoromethyl coumarin assay
(AFC) linked to the tetra-peptide DEVD-AFC (caspase-3, Cat-No.: 03AFC13810, MP
Biomedicals, Solon, OH), or linked to LEHD (caspase-9, Cat-No.: 0219944501, MP
Biomedicals, Solon, OH) in a cyto-buffer (10% glycerol, 50 mM Pipes, pH 7.0, 1 mM
EDTA, 1 mM DTT). For the detection of caspase-8 activity, the lysates were incubated with the IETD-AFC (Cat-No.: 556552, BD Biosciences, in a cyto-buffer (20% glycerol,
100 mM Hepes, pH 7.5, 1 mM EDTA, 1 mM DTT) at 37°C for 30 min. Release of free
AFC was measured using the Cytofluor 4000 fluorometer (Perceptive Co, Framingham,
75
MA), (filters: excitation, 400 nm; emission, 505 nm). The enzymatic activity of the caspase is proportional to the fluorescence released as the consequence of the cleavage of the AFC. To calculate the enzymatic activity, a graph of fluorescence vs. time (min) was generated. The slope of the graph was calculated and divided by the amount of protein utilized in the assay. Utilizing known concentrations of AFC, ranging from 0 – 80 µM) a standard curve was generated. The activity of caspase-3 was expressed as nM
AFC/min/amount of protein.
2.12 In vitro kinase assays with recombinant caspase-3
The recombinant caspase-3 proteins were phosphorylated in vitro by human recombinant PKCδ (Cat.-No.: P2293, Invitrogen). One hundred ng of recombinant caspase-3 were added to 20 µl of kinase reaction mixture containing 25 mM HEPES, pH
32 5.2, 10 mM MnCl2, 1 mM MgCl2, 500 nM ATP and 4 Ci of [γ- P] ATP (Perkin Elmer) in the presence of 2.5 ng of PKCδ. The specific activity of PKCδ is 1603 nmole of phosphate transferred to a peptide substrate per minute per mg of total protein at 30°C. A mixture of phosphatidylserine (PS) (Cat-No.: 770035, Avanti Polar Inc. Alabaster, AL) and diacylglycerol (DAG) (Cat-No.: 870520P, Avanti Polar Inc.) were added to the reactions as cofactors at a final concentration of 200 µg/ml and 20 µg/ml, respectively.
The phosphorylation reaction was carried out at 37°C for 0.5 h. Reactions were stopped by the addition of 5 µl of 5X Laemmli buffer [250 mM Tris, pH 6.8, 10% SDS, 50% glycerol, 0.5% bromophenol blue (BPB) and 1.78 M β-mercaptoethanol]. The samples
76 were boiled at 95ºC for 5 min and resolved by Western blot. The membrane was exposed to a film for the detection of the phosphorylated proteins by autoradiography.
A serial dilution of known concentration of [γ-32P] ATP, ranging from 0.5 pCi to
25 was spotted on a filter paper and standards were visualized by autoradiography. The density (Intensity/mm2) of the band obtained for each standard was analyzed utilizing the
Quantity One software (Version 4.6.6, Bio-Rad) and graphed the density (Intensity/mm2) vs. the calculated counts per minute (CPM) values of each of the [γ-32P] ATP standards.
The density of the signal was used to calculate the corresponding cpm-values
(CPMSample). To calculate the total amount of ATP incorporated CPMSample-values were multiplied by the total amount of ATP (pmol) that was used for the kinase reaction
(ATPtotal = the sum of radioactive and non-radioactive ATP). These values were divided by the total amount of CPMs that were supplied during the kinase reaction (CPMtotal).
The specific activity was determined by division of the total amount of incorporated
ATP by the total amount of protein (μg) and kinase reaction time (min), described in the following formula:
CPM sample * ATPtotal CPM Specific _ activity total ng protein * min
For the detection of caspase-3 by chemiluminescence, the membrane was blocked with 5% milk in TBS containing 0.05% Tween 20. The membrane was incubated with anti-caspase-3 antibodies (1:2000) overnight at 4ºC (Cat.-No.: 610323, BD Biosciences,
San Jose, CA) (Table 2.1). Membrane was rinsed with TBS buffer three times and was
77 incubated with horseradish peroxidase conjugated anti-mouse secondary antibody
(1:4000, GE, Piscataway, NJ) (Table 2.1).
2.13 Monocyte subset isolation
Human blood from normal patients was obtained from the American Red Cross.
The blood was diluted 1:1 with sterile phosphate buffered saline (PBS) solution and was subjected to a Histopaque-1077 gradient (Sigma), at 2,000 rpm for 20 min at 18 °C. The peripheral mononuclear layer (PBMC) was collected and rinsed twice in cold RPMI 1640
(Invitrogen). Hypolysis of red blood cells was performed in 2 ml of endotoxin free water, for 50 sec at room temperature. The reaction was stopped by addition of 2X PBS.
The cells were rinsed in RPMI, centrifuged at 1,050 rpm and a fraction stained with
Trypan blue was utilized for counting in a hemocytometer. Cell pellets were resuspended at a concentration of 1 x 106 cells in 180 µl of positive selection buffer (PSB, 137 mM
NaCl, 2.7 mM KCl, 10 mM Na2HPO4, 2 mM KH2PO4, 2 mM EDTA, 0.5% BSA). Cells were incubated with FcR blocking reagent in a ration of 100 μl of reagent per 108 cells and non-monocyte depletion cocktail (Cat-No.: 120-000-249, Miltenyi Biotec), at a ratio of 100 µl beads/1 x108 cells. The cell/bead suspension was loaded onto a LD columns
(Cat-No.: 130-042-402), non-labeled cells eluted in the flow-through and were incubated
CD16 magnetic microbeads (Cat-No.: 120-000-249, Miltenyi Biotec) at a ratio of 80 µl beads/1 x 108 cells for 15 min at 4ºC. The cell/bead suspension was loaded onto a LS column (Cat-No.: 130-042-401) and CD16+ and CD16- labeled cells were isolated by positive and negative selection. The flow through consisted of CD16- cells, whereas
78
CD16+ remained in the column due to the magnetic field. The cells are eluted by removal from the magnets and addition of 5 ml of PSB. CD16+ and CD16- were resuspended in PSB and were incubated with CD14-magnetic microbeads at a concentration of 16 µl beads/1 x 107 cells (Cat-No.: 130-050-201, Miltenyi Biotec). The
CD14+ positive cells were isolated by positive selection in the LS-column. The flow- through contained non-monocytic cells. CD14+ cells remained bound to the column and were eluted in 5 ml of PBS by removal of the column from the magnetic field. The isolated monocytes were kept at a concentration of 2 x 106 cells/ml. The purity of the
CD14+CD16+ and CD14+CD16- monocyte sub-populations were assessed by flow cytometry, described in Section 2.21. Monocytes were seeded for experiments at a density of 0.5 x 106 cells/ml were cultured for various length of time in RPMI 1640.
2.14 Inhibition of caspase activity
Purified monocytes were incubated at a concentration of 0.5 x 106 cells/ml for 8 h in the presence of DEVD-FMK (Cat-No.: 550378), IETD-FMK (Cat-No.: 550380), and
LEHD-FMK (Cat-No.: 550381) (BD Biosciences, San Jose, CA), caspase-3, caspase-8, or caspase-9: inhibitors of respectively. All inhibitors were dissolved in dimethyl sulfoxide (DMSO, Sigma), and used final concentrations of 1 to 25 M. The activity of caspase-3, caspase-8, and caspase-9 was determined by incubating 10 µg of cell extract with the substrates: DEVD-AFC, IETD-AFC, and LEHD-AFC, respectively, as described in Section 2.13.
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2.15 Electroporation of monocyte subsets
One million cells were resuspended in 20 μl of the electroporation buffer, provided by the manufacturer (Cat-No.: V4X P-3032, Amaxa, Cologne, Germany). Six pmol of PKCε or siRNA-control were added to the solution and transferred to the nucleocuvette strips (Table 2.4). The EA-100 program was used for the electroporation of human primary monocytes in the Amaxa 4D-Nucleofector X-Unit, according to the manufacturer’s specifications. After transfection the cells were resuspended in 100 μl of
RPMI containing 10% FBS and were transferred to culture tubes and incubated for 24 h before stimulation with LPS.
Table 2.6 List of siRNA oligonucleotides
siRNA Company Primer Sequence (5'-3') Scramble Dharmacon Sense UUCUCCGAACGUGUCACGUtt Scramble Dharmacon Antisense ACGUGACACGUUCGGAGAAtt PKCε Dharmacon Sense AAGCCCCUAAAGACAAUGAAGtt PKCε Dharmacon Antisense CUUCAUUGUCUUUAGGGGCUUtt
2.16 IgG clustering assay
The IgG clustering assay was perform as previously described (270). utilized to mimic an FCRγ mediated immune response and stimulate the production of pro- inflammatory cytokines. A 96-well plate was coated with chrome pure human IgG (Cat-
No.: 009-000-003, Jackson ImmunoResearch Laboratories, West Grove, PA), at a concentration of 10 µg/ml diluted in PBS and incubated for 16 h at 4°C. Next day, the plates were rinsed three times with 300 μl of PBS. Monocytes were plated in wells at a
80 density of 200,000 cells/200 µl of RPMI 1640, for 16 h at 37°C. The supernatant was collected for TNF-α detection by enzyme-linked immunosorbent assay (ELISA).
2.17 Detection of TNF-α by ELISA
The release of TNF-α in monocyte subpopulations was measured by a sandwich
ELISA following the manufacturer instructions (Cat-No.: DY210 R&D, Minneapolis,
MN). A 96-well plate was coated with the capture antibody (4 µg/ml) diluted in PBS and was incubated for 16 h at room temperature. Next day, the plates were rinsed three times with washing buffer (PBS pH 7.4, 0.05% Tween 20) then blocked with 300 µl of reagent diluent (PBS pH 7.4, BSA 5%, 0.05% Tween 20) for 1 h at room temperature. After incubation, the plates were rinsed three times with washing buffer and coated with TNF-α standard, at concentrations ranging from 1000 pg/ml to 15 pg/ml, or with 100 µl of the sample, for 2 h at room temperature. The plates were rinsed three times with washing buffer then coated with 100 µl of detection antibody (200 ng/ml) and incubated for 2 h, at room temperature. Then, the wells were rinsed three times with washing buffer and were incubated for 20 min, in the dark, with 0.1 ml of streptavidin-HRP (dilution 1:200). After incubation, the plate was rinsed and incubated with 0.1 ml of the substrate solution for 20 min. Reactions were stopped by the addition of 50 µl of H2SO4 solution (2 N) and the color intensity was measured with the Victor X3 micro plate reader (Perkin Elmer) at a wavelength of 450 nm.
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2.18 Detection of PKC isoforms in monocyte subsets
The isolated monocytes were lysed utilizing the NP-40 buffer, as described previously. The detection of PKC isoforms were performed utilizing 5 µg of protein. For
50 µg of protein was utilized for PKC gamma and PKC lambda/iota were. The proteins were loaded on an 8% acrylamide gel and were run for 2 h at 100 V. The proteins were transferred to a Protran Nitrocellulose Blotting Membranes (0.20 µm, Whatman, USA) for 70 min at 100 V on transfer buffer (25 mM Tris, pH 8.3, 192 mM glycine, 20%
Methanol). The membranes were blocked with 5% milk in TBS containing 0.05%
Tween 20. The membrane was incubated with antibodies for PKC proteins at a dilution of 1:2000. Recombinant PKC proteins (Pan Vera, Madison, WI) were used as a positive control, including PKCα (P2345), PKCβI (P2346), PKCβII (P2347) PKCγ (P2350), PKCδ
(P2293), PKCε (P2349), PKCθ (P2995), PKCη (P2635), PKCζ (P2351), and PKCι/λ
(PV3230). Additional immunoblots were carried out with anti-caspase-3 (BD
Biosciences), anti-active caspase-3 (Cell Signaling, Danvers, MA) and anti-β-Tubulin
(Millipore, Charlottesville, VA) antibodies. Membranes were rinsed with TBS buffer three times and were incubated with horseradish peroxidase conjugated anti-mouse secondary antibody (1:4000, GE, Piscataway, NJ) (Table 2.1).
2.19 Flow cytometry analysis
CD14+CD16- and CD14+CD16+ cells at a final concentration of 0.5 x 105, were resuspended in 100 µl of Annexin V staining buffer [0.01 M Hepes/NaOH pH 7.4,0.14 M
NaCl, 2.5 mM CaCl2] (BD Biosciences) containing 10 ng/ml of the following antibodies:
82
FITC-conjugated anti-CD14-APC (BD Biosciences), anti-CD16-PE (Miltenyi Biotec), anti-cleaved PARP-FITC (BD Biosciences), anti-active caspase-3-FITC (Cell Signaling) or IgG isotype control; and were incubated for 15 min at room temperature (Table 2.1).
Cells were rinsed once with 0.5 ml of Annexin V buffer and centrifuged at 1,200 rpm for
5 min and were resuspended in 0.5 ml of Annexin V staining buffer containing 1% paraformaldehyde. The isolated cell purity was determined by flow cytometry analysis utilizing the BD Biosciences FACSCalibur flow cytometer and the FCS Express V3 software.
2.20 Statistical analysis
The sample deviations were calculated and expressed as standard error of the mean (SEM). The calculated standard error was divided by the square root of the sample size (n). The statistical significance was determined by student t test using the Graph Pad
Prism, version 5. A variance with P ≤ 0.05, were considered statistically significant.
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Chapter 3
PKCδ phosphorylation of caspase-3 regulates the execution of apoptosis
3.1 Abstract
Caspase-3 is an important mediator of cell death and its activation is central in the apoptotic cascade. Its activity is tightly regulated through positive and negative signals that modulate its activation for the execution of apoptosis when needed. We have previously shown that caspase-3 is phosphorylated by PKCδ. However, the role of caspase-3 phosphorylation during apoptosis is not completely understood. In this study we showed that PKCδ is necessary for caspase-3 phosphorylation, in vivo. We showed the domains of caspase-3 that bind to and are phosphorylated by PKCδ. We further identified an interaction motif in caspase-3 necessary for PKCδ binding and phosphorylation. In addition, we mapped the sites in caspase-3 phosphorylated by PKCδ.
Phosphorylation of Ser12 and Ser36 promoted caspase-3 activity by promoting the kinetics of the autocatalytic cleavage. We identified Ser36 as an important amino acid necessary for caspase-3 phosphorylation and execution of apoptosis. These results support a model by which apoptosis is regulated by the PKCδ dependent phosphorylation of caspase-3 and the phosphorylation of specific sites has the potential to shift the balance between cell death and survival.
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3.2 Introduction
Apoptosis is an evolutionarily conserved mechanism important for development, immunity, and tissue homeostasis through the physiological removal of cells (271). The family of cysteine proteases, termed caspases, is responsible for the execution of apoptosis in higher eukaryotes. Caspases exist in the cell as inactive proenzymes that require a proteolytic cleavage for their activation. Apoptosis can be activated through the intrinsic or extrinsic pathway, depending the origin of the death stimulus (Fig. 1.5) (6).
Activation of the intrinsic pathway by UV radiation, chemotherapeutic drugs and oxidative stress, causes the release of cytochrome c from the mitochondria (27, 30). In the cytosol, cytochrome c binds the apoptotic factor Apaf-1 and induces its oligomerization, recruiting and causing the dimerization and autocatalytic activation of the initiator caspase, caspase-9 (Fig. 1.5) (41, 54, 57). The active caspase-9 cleaves and activates effectors caspases, such as caspase-3.
The proenzyme or full-length caspase-3 (caspase-3-FL) is composed of 3 domains; the prodomain or N-terminal, the p17 or intermediate and the p12 or C-terminal domain (Fig. 1.8). The activation of caspase-3 involves a two-step proteolytic processing. The first cleavage at caspase-3-D175 is mediated by an initiator caspase, like caspase-9, which separates the prop17 (composed of the prodomain and p17 domain) and p12 domains (25, 37). A second autocatalytic cleavage separates the prodomain and the p17 domain (39). Two p17 and two p12 domains rearrange forming a heterodimer that forms the active enzyme (133). The active caspase-3 cleaves structural and regulatory proteins that cause DNA fragmentation, membrane blebbing, and formation of apoptotic
85 bodies, some of the typical morphological changes associated with apoptosis. During the demolition phase of apoptosis caspase-3 cleaves the amino acid sequence DEXD (X is any amino acid) found in its cellular targets including α-fodrin, poly (ADP-ribose) polymerase-1 (PARP-1), and PKCδ (6, 77, 125). The serine/threonine kinase, PKCδ, is proteolytically cleaved by caspase-3 at the hinge region releasing the 40 kDa catalytically active fragment (Fig. 1.10). The release of PKCδ cleaved (PKCδ-CL) fragment has been shown to be important for later stages in the apoptotic cascade (Fig. 1.10) (272).
We have previously shown that PKCδ associates with and phosphorylates caspase-3 (116). Currently, the role of PKCδ phosphorylation of caspase-3 during apoptosis has not been completely elucidated. PKCδ is a pro-apoptotic kinase responsible for the phosphorylation of membrane, cytoplasmic and nuclear proteins during apoptosis (213). Previously, classical PKCs were shown to require an interaction motif for the binding and phosphorylation of p53, which is not required by PKCδ (273).
However, while several PKCδ substrates are known, including γ-actin, p300, Sp1, DNA- dependent protein kinase (DNA-PK), and Hsp-27 (Gonzalez-Mejia and Doseff unpublished data), how the association and therefore specificity is defined remains unclear (214, 215, 221, 274). Other kinases, such as protein kinase alpha (PKA), phosphoinositide-dependent kinase 1 (PDK1), mitogen-activated protein (MAP) kinase, and cyclin dependent kinase (CDK), interact with their substrates through “recognition motifs” located in short amino acid sequences adjacent to the phosphorylation sites (275).
Linear motifs can be about 3 - 8 amino acids in length and are characterized as areas of transient interaction involved in signaling networks (276). These motifs have been
86 suggested to provide specificity and stability to the kinase-substrate interaction important for efficient phosphorylation and transduction of the signaling cascade (277, 278). To gain understanding on what defines PKCδ substrate specificity we analyzed the primary structure of PKCδ-interacting substrates to assess whether they share a common motif that mediates the interaction. Characterization of the substrate interaction motif necessary for PKCδ binding would provide valuable information for the development of new therapies that could selectively inhibit this kinase (279-284).
In this study, we showed that PKCδ is necessary for caspase-3 phosphorylation, in vivo. We identified the domains of caspase-3 involved in the association with PKCδ and discovered an interaction motif responsible for their association. In addition, 5 sites in caspase-3 were identified as targets of PKCδ, in vitro. We found phosphorylation of specific sites promote caspase-3 cleavage and apoptosis. These results provide a novel regulatory mechanism controlling caspase-3 apoptotic activity and execution of cell death.
3.3 Results
3.3.1 PKCδ is necessary for caspase-3 phosphorylation during cell death
Previously we demonstrated that caspase-3 is phosphorylated by PKCδ in human monocytes and the increase in caspase-3 activity during monocyte cell death correlates with the increase in PKCδ activity (116). However; the role of PKCδ phosphorylation of caspase-3 during apoptosis is not understood. We utilized mouse embryonic fibroblast
(MEF) from WT, PKCδ-/- and PKCβ-/- knockout mice to investigate whether PKCδ is
87 necessary for caspase-3 phosphorylation during apoptosis. Immunoblot analysis showed similar levels of caspase-3 in the WT, PKCδ-/- and PKCβ-/- MEF (Fig. 3.1 A). PKCδ-/-
MEF lack of PKCδ expression, but express normal levels of PKCβI, whereas MEF from
-/- PKCβ mice lack PKCβI expression but have normal levels of PKCδ (Fig. 3.1 A). MEF were treated with 10 μM doxorubicin for 3 h to induce apoptosis or left untreated.
Lysates were immunoprecipitated with anti-caspase-3 antibodies or IgG isotype control, and subjected to in vitro kinase assays. H2B was utilized as an exogenous substrate, as we showed previously that it is highly phosphorylated by the caspase-3 associated kinase
(116). Caspase-3 associated phosphorylation increased during apoptosis in WT and
PKCβ-/- MEF (Fig 3.1 B, lane 3 and 9). However, no increase in the caspase-3 associated phosphorylation was observed in PKCδ-/- MEF (Fig. 3.1 B, lane 6). These results suggest that PKCδ is necessary for caspase-3 phosphorylation during cell death.
3.3.2 Identification of caspase-3 domains phosphorylated and involved in the interaction with PKCδ
To identify the domains of caspase-3 that are phosphorylated by PKCδ, HeLa cells were transiently transfected with the N-terminal Xpress-tag pcDNA4-HisMax
(Xpress vector) plasmids encoding human caspase-3 full-length (FL) or different domains generated during caspase-3 proteolytic processing through apoptosis (Table 2.3.3).
Immunoprecipitation of the Xpress-tagged caspase-3 proteins and in vitro kinase assays were performed. Consistent with our previous results, caspase-3-FL, consisting of the prodomain, p17 and p12 domains, was found to be phosphorylated (Fig. 3.2 A, lane 2)
88
(116). The phosphorylation of caspase-3-p17 domain showed ~ 2-fold increase compared with caspase-3-FL, while prop17 showed an even higher phosphorylation of ~
5-fold. (Fig. 3.2 A, lanes 2-4). In contrast, no phosphorylation was observed in the p12 or C-terminus domain, even though all domains were expressed at similar levels (Fig 3.2
A, lanes 5-9).
Next, we identified the domains of caspase-3 involved in the association with
PKCδ. Lysates from cells transiently expressing full length or different domains of caspase-3 were immunoprecipitated with anti-PKCδ antibodies or IgG isotype control.
We observed that PKCδ associates with caspase-3-FL, as previously reported (Fig 3.2 B, lane 2) (116). The prop17 and p17 domains showed an increase in the association with
PKCδ of ~ 5 and ~ 2 fold, respectively, compared to caspase-3-FL (Fig 3.2 B, lanes 3 and
4 vs. 2). However, the p12 domain showed no interaction with PKCδ, but the lack of association is not due to its expression, as all domains were similarly expressed (Fig 3.2
B, lanes 5-9). Collectively, these results demonstrate that the prop17 and p17 domains associate with and are phosphorylated by PKCδ, and suggest that the p12 domain is not necessary for the association.
3.3.3 Identification of PKCδ interaction motif in caspase-3
Interaction motifs have been identified for several substrates of protein kinases.
However, how PKCδ associates with its substrates is not yet known (275, 277, 278).
Protein linear motifs have been shown to be important regions for protein-protein interaction (285). Utilizing the Dilimot program (http://dilimot.russelllab.org) we 89 analyzed the amino acid sequence of substrates that have been shown to directly interact with PKCδ, including Hsp27, γ-actin, p53, p300, DNA-PK, and SP1 (285). This program has identified the motif of interaction of MDM2/p53 and stearoyl-CoA desaturase 5
(Scd5)/protein phosphatase 1 (PP1) (285, 286). Two linear motifs were identified as potential PKCδ binding motif, referred as PB1 and PB2, in caspase-3. The two predicted binding motifs were located in the p17 caspase-3 domain (Fig. 3.3 A). PB1 corresponds to the amino acids S29GXS32 (X corresponding to any amino acid) just adjacent to the boundary between the prodomain and the p17 domain (Fig 3.3 A). PB2 corresponds to amino acids Q161AXXXXXL168, contains the conserved QACXG sequence characteristic of all members of the caspase family (Fig. 1.2). Comparative studies showed that PB1 is found in all substrates that were showed to directly interact with PKCδ, except p53 which had an SXXS motif instead. PB2 was found in all proteins except γ-actin (Fig. 3.3 A).
The conservation of these sequences in PKCδ associating substrates analyzed suggested that they could correspond to important motifs for the kinase:substrate interaction (Fig.
3.3 A, grey). To assess the role of the predicted motifs, site directed mutagenesis of the
PB1 was performed to replace the SGIS amino acids with 4 alanines (4A: aa 32 - 36) in caspase-3 FL and prop17, referred as PKCδ binding mutant 1 (PBM1) (Fig. 3.3). The
QACRGTEL (aa 161-168) motif was deleted in both caspase-3-FL and caspase-3- prop17, referred to as PKCδ binding mutant 2 (PBM2) (Fig. 3.3 B). To assess the effect of the interaction motif in PKCδ binding and phosphorylation, HeLa cells were transiently transfected with caspase-3-WT-prop17, caspase-3-PBM1-prop17 or caspase-
3-PBM2-prop17 (Fig 3.3 C). Immunoprecipitation of the Xpress-tagged caspase-3
90 proteins and in vitro kinase assays were performed. Our results show that the phosphorylation was reduced by 75% in the caspase-3-PBM1-prop17 and no reduction in phosphorylation was observed in the caspase-3-PBM2-prop17, compared to caspase-3-
WT-prop17 (Fig. 3.3 C, lanes 3 and 4 vs. 2). PKCδ binding was also reduced by 75% in caspase-3-PBM1-prop17, while it showed equal binding to caspase-3-PBM2-prop17 as with the caspase-3-WT-prop17 (Fig. 3.3 C, lanes 3 vs. 3 and 4). In addition, similar experiments were done using full-length caspase-3 proteins. The phosphorylation of caspase-3 by PKCδ was completely inhibited in caspase3-PBM1-FL (Fig. 3.3 D, lane 3).
In addition, the PKCδ association and phosphorylation of caspase-3 was abolished in caspase-3 PBM1-FL, but no reduction in PKCδ binding or phosphorylation was observed in the caspase-3-PBM2-FL. These results show that the SGIS motif is necessary for
PKCδ binding and phosphorylation of caspase-3. Furthermore, the conservation of this motif suggests that it could play an important role in the PKCδ association and phosphorylation of additional substrates.
3.3.4 Identification of PKCδ phosphorylation sites in caspase-3
To identify the PKCδ phosphorylation sites in caspase-3 we cloned the caspase-3 prop17 domain into the pQE31 bacterial expression vector, as it was the domain that showed the highest phosphorylation by PKCδ. Bacterial recombinant capase-3-prop17 was affinity purified and subjected to in vitro kinase assays using recombinant PKCδ
(rPKCδ) (Table 2.3). We found that caspase-3-prop17 was phosphorylated by rPKCδ,
91 but not by the heat inactivated rPKCδ (Fig. 3.4 A, upper panel, (+) and (-) respectively).
Thus, these results demonstrate that caspase-3-prop17 is a direct substrate of PKCδ.
To map the PKCδ phosphorylation sites on caspase-3, the caspase-3-prop17 was phosphorylated in vitro by rPKCδ or heat-inactivated rPKCδ in the presence of non- radioactive ATP, and was analyzed by mass spectrometry (Section 2.1) (287). A peptide coverage of more than 90% of the prop17-caspase-3 domain using trypsin was obtained and the generated peptides were verified using the Mascot software by searching against the non-redundant human proteome (288). The mass spectrometry data identified 5 phosphorylation sites located in the human caspase-3 prop17 domain (Fig. 3.4 B). Ser12 was located in the prodomain and Ser36, Thr59, Thr67, Thr77 were located in the p17 domain. Evolutionary conservation analysis showed that Ser12, Ser36 and Thr59 are conserved in vertebrates, Thr67 was conserved from humans to fruit fly and Thr77 was conserved in mammals and fruit fly (Fig 3.4 B). The conservation of the caspase-3 phosphorylable amino acids suggests an important role of these sites in the regulation of caspase-3.
To assess the role of phosphorylation on caspase-3, site directed mutagenesis was performed to replace the five amino acids with alanine or glycine to create the phospho- dead mutant, reffered as caspase-3-M5-A/G-prop17 [S12G/S36A/T59A/T67A/T77A] (Table
2.3). Purified recombinant caspase-3-WT-prop17 and caspase-3-prop17-M5-A/G were subjected to in vitro kinase assays, using rPKCδ or heat-inactivated rPKCδ (-). Mutation of the 5 identified phosphorylation sites inhibited the PKCδ-dependent phosphorylation compared to caspase-3-WT-prop17 (Fig. 3.4 C, lane 3 vs. 2). However, no
92 phosphorylation of caspase-3-WT-prop17 was observed in reactions containing the heat inactivated rPKCδ (Fig. 3.4 C, lane 1). In addition, to evaluate the effect of caspase-3 phosphorylable sites in vivo, caspase-3-M5-A/G-prop17 was cloned into the mammalian expression vector Xpress, thus obtaining amino-terminal tagged Xpress caspase-3-M5-
A/G-prop17 (for details see Table 2.3). Lysates from HeLa cells transiently expressing the Xpress-tagged caspase-3-WT-prop17 or caspase-3-M5-A/G-prop17 were immunoprecipitated with anti-Xpress or IgG isotype control antibodies and subsequently subjected to in vitro kinase assays. Mutation of the 5 sites target of PKCδ prevents caspase-3-prop17 phosphorylation in vivo (Fig. 3.4 D, lane 3 vs. 2). Immunoblot with anti-Xpress antibodies showed equal immunoprecipitation of both caspase-3 proteins.
These results suggest that the phosphorylated amino acids identified by mass spectroscopy analysis are important for caspase-3 phosphorylation.
Next, the contribution of each independent phosphorylable site was evaluated.
For this purpose, the identified phosphorylation sites in caspase-3 were individually mutated to either glycine or alanine to create single phospho-null mutants [S12G, S36A,
T59A, T67A, and T77A] and were expressed in bacteria. The purified recombinant caspase-3-WT and phospho-null mutants showed a purity of ~ 90%, as determined by
Coomassie staining (Fig. 3.5 A). The recombinant proteins were incubated with rPKCδ and subjected to in vitro kinase assays. We found that mutations in Ser12 and Ser36
(caspase-3-S12G and caspase-3-S36A) abolished caspase-3 phosphorylation, while negligible reductions in phosphorylation were observed in caspase-3-T59A, caspase-3-
93
T67A, and caspase-3-T77A (Fig. 3.5 B). Hence, these results suggest that Ser12 and Ser36 are the main sites phosphorylated by PKCδ.
3.3.5 Ser12 is important for caspase-3 phosphorylation
To determine the role of Ser12 in caspase-3 phosphorylation, a phospho-mimetic mutant in which Ser12 was replaced by an Asp was created by site directed mutagenesis in the caspase-3-FL and also in the caspase-3-prop17, referred as caspase-3-S12D. Both the null and phospho-mimetic mutants, caspase-3-S12G and caspase-3-S12D, were cloned into mammalian Xpress vector. Lysates from HeLa cells transiently expressing caspase-
3-WT-FL or mutants were immunoprecipitated with anti-Xpress antibodies or IgG isotype control and were subjected to in vitro kinase assays. Caspase-3-S12G-FL and caspase-3-S12D-FL showed a reduction in phosphorylation of 90%, compared to caspase-
3-WT-FL (Fig. 3.6 A, lanes 3-4 vs. 2). However, no differences in the binding of PKCδ were observed between caspase-3-WT-FL, caspase-3-S12G-FL and caspase-3-S12D-FL.
Similar experiments were conducted with the caspase-3-S12G-prop17 and caspase-3-
S12D-prop17. In this case, a ~ 80% reduction in phosphorylation of caspase-3 was observed (Fig. 3.6 B, lanes 3 and 4 vs. 2), and no differences in the association of caspase-3 with PKCδ were observed. These results suggest that Ser12 has an important role in caspase-3 phosphorylation.
94
3.3.6 The phosphorylation of Ser36 is necessary for the PKCδ dependent
phosphorylation of caspase-3
Next, we assessed the role of Ser36 in caspase-3 phosphorylation. First, the phospho-mimetic mutant S36D was created by site directed mutagenesis in the caspase-3-
FL and caspase-3-prop17 and was cloned into the Xpress vector. The phospho-null and phospho-mimetic caspase-3-S36A and caspase-3-S36D, respectively, were used for transient transfection into HeLa cells (Table 2.3). The proteins were immunoprecipitated with anti-Xpress antibodies and subjected to in vitro kinase assays. Mutation of S36A prevented caspase-3-FL phosphorylation (Fig. 3.7 A, lane 3). In contrast, caspase-3-
S36D-FL showed a ~ 2 fold increase in phosphorylation, with no change in PKCδ association, compared to caspase-3-WT-FL (Fig. 3.7 A, lane 4 vs. 2). Similar experiments showed that mutation of S36A inhibited caspase-3-prop17 phosphorylation, while no effect in the phosphorylation was observed in the caspase-3-S36D-prop17 (Fig.
3.7 B). No differences in the association with PKCδ were observed in caspase-3-WT- prop17, caspase-3-S36A-prop17 or caspase-3-S36D-prop17. These results indicate that
Ser36 is an important site for regulation of caspase-3 phosphorylation.
Next we determined whether the inhibition in phosphorylation observed in the caspase-3-S36A mutant was specific. For this purpose we mutated Ser32 to alanine (S32A), a site located in the p17 domain that lacks evolutionarily conservation (Fig. 3.3, Table
2.4). HeLa cells were transiently transfected with the Xpress tagged caspase-3-WT, caspase-3-S32A or caspase-3-S36A. The lysates were immunoprecipitated with anti-
Xpress antibodies and subjected to in vitro kinase assays. Mutation of S32A did not 95 prevent caspase-3-FL phosphorylation, compared to caspase-3-WT-FL (Fig. 3.7 C, lane 3 vs. 2). In contrast, caspase-3-S36A-FL showed no phosphorylation but showed equal binding to PKCδ as the caspase-3-WT-FL and caspase-3-S32A-FL (Fig. 3.7 C). These results suggest that the phosphorylation of Ser36 is specific and by mutating this amino acid we prevent caspase-3 phosphorylation.
3.3.7 Caspase-3 phosphorylation modulates its protease activity
Next we evaluated the effect the newly identified phosphorylable amino acids in caspase-3 protease activity. For this purpose, recombinant caspase-3-WT, caspase-3- phospho-null (S12G, S36A, T59A, T67A, and T77A) and phospho-mimetic (S12D, S36D,
T59D, T67D, and T77D) proteins were incubated with recombinant caspase-9 for 0, 0.5, 1, or 2 h at 37ºC, to promote the first cleavage and the subsequent second autocatalytic cleavage necessary for caspase-3 activation. The mixtures were used to determine the activity of caspase-3 by the DEVD-AFC assay. The enzymatic activity of caspase-3-WT increases over time and shows a peak in activity of 40 fold by 1 h and 80 fold by 2 h (Fig.
3.8, A). All caspase-3-phospho-null recombinant proteins showed similar kinetics of activation as caspase-3-WT (Fig. 3.8 A). However, changes in the activity of caspase-3 were observed in the caspase-3 phospho-mimetic proteins (Fig. 3.8 B). Caspase-3-S12D and caspase-3-S36D showed significantly higher activity than caspase-3-WT. After 1 h, caspase-3-S12D and caspase-3-S36D had an increase in activity of 100 fold and by 2 h showed an increase of 120 fold (Fig. 3.8 B, 1 h, P <0.005, 2 h, P <0.05). In contrast,
T59D, T67D and T77D showed reduced activities. Caspase-3-T77D showed 50% lower 96 activity than caspase-3-WT over time (1 h and 2 h, P < 0.05). In contrast, the caspase-3 mutations T59D and T67D inhibited caspase-3 activity (P <0.005). These results suggest that caspase-3 contains phosphorylation sites that positively and negatively modulate its activity.
To evaluate whether the simultaneous phosphorylation of Ser12 and Ser36 promotes caspase-3 activity; both amino acids were mutated (S12G/S36A and S12D/S36D).
Similar experiments were conducted to activate the recombinant caspase-3-WT, caspase-
3-S12G/S36A and caspase-3-S12D/S36D by incubation with caspase-9. We found that the phospho-mimetic mutant caspase-3-S12D/S36D did not promote higher activity compared to the caspase-3-WT or caspase-3-S12G/S36A. There was no statistical difference in the activities of these proteins (P > 0.05). These results suggest that the simultaneous phosphorylation of Ser12 and Ser36 does not promote caspase-3 activity.
3.3.8 Role of Ser12 and Ser36 phosphorylation in the autocatalytic cleavage of
caspase-3
The increase in caspase-3 activity observed in the recombinant caspase-3-S12D and caspase-3-S36D phospho-mimetic proteins suggested that the phosphorylation may affect caspase-3 cleavage. To assess whether the phosphorylation of these sites affect caspase-3 cleavage and activation, caspase-3-WT, caspase-3-S12G, caspase-3-S12D, caspase-3-S36A, and caspase-3-S36D were incubated with recombinant caspase-9 for different periods of time. Immunoblots show that caspase-3-WT has a polypeptide at 36 kDa corresponding to the full-length unprocessed zymogen (Fig. 3.9 A, lane 1, Casp-3- 97
FL); which was processed to a 22 kDa polypeptide corresponding to the prop17 domain by ~ 0.5 h (Fig. 3.9 A, lane 2, prop17), and was further processed after 1 h into a 17 kDa corresponding to the p17 large domain of caspase-3 (Fig. 3.9 A, lane 3). At this time, some of the precursor caspase-3 was still observed, but by 2 h of incubation caspase-3-
WT was completely processed (Fig. 3.9 A, lane 4). Similar kinetics of processing were observed in the caspase-3-S12G and caspase-S36A phospho-null proteins. In both cases, the prop17 polypeptide was found at ~ 1 h and limited amount of the p17 domain was observed, and after 2 h the inactive full length was fully cleaved (Fig. 3.8 A, lanes 3-4, 7-
8 and 15-16). In contrast, caspase-3-S12D and caspase-3-S36D showed an increase of 4 fold in the accumulation of the p17 by 1 h compared to caspase-3-WT or the phospho- null mutant proteins (Fig. 3.8 A, lanes 3, 11 and 19).
Next, the effect of phosphorylation in the processing of caspase-3 was evaluated in a cell free system. This system was previously shown to activate the apoptotic signaling cascade, reconstituting apoptotic features (269). The activation of caspase-3 is mediated by endogenous caspase-9, cytochrome c and dATP as part of the intrinsic pathway (Section 1.4). MCF-7 cytosolic extracts lacking endogenous caspase-3 were supplemented with purified recombinant caspase-WT or phosphorylation mutants. The first product of processing, prop17 polypeptide, was observed in caspase-3-WT at ~ 3 h
(Fig. 3.9 B, lane 3), and by 4 h the p17, product of the second autocatalytic cleavage, was observed and was accompanied by the disappearance of the full length zymogen (Fig. 3.9
B, lane 4). Similar kinetics of processing were observed in caspase-3-S12G and caspase-
S36A mutants (Fig. 3.9 B). In contrast, both phospho-mimetic mutants showed faster
98 kinetics of processing. The p17 domain, product of caspase-3 autocatalytic cleavage, was cleaved by 3 h in caspase-3-S12D and caspase-3-S36D, at a time when caspase-3-WT and phospho-null mutants only showed the prop17 polypeptide (Fig. 3.9 lanes 11 and 19 vs.
3, 7 and 15. It was noticed that by 3 h, the p17 polypeptide of the caspase-3-S36D was 2 fold higher than in caspase-S12D (3.9 B, lanes 11 vs. 19). After incubation for 4 h the caspase-3-S12D and caspase-3-S36D showed complete cleavage of the precursor protein
(Fig. 3.9 B, lanes 12 and 20). Altogether, these results suggest that phosphorylation of
Ser12 and Ser36 increase the activity of caspase-3 by increasing the kinetics of the autocatalytic cleavage.
Utilizing the same cell free reactions described, we evaluated whether the faster processing kinetics of caspase-3 phospho-mimetic had an effect on the cleavage of the endogenous α-fodrin, a substrate of caspase-3. Under physiological conditions, α-fodrin is cleaved (CL) by calpain, producing a 150 kDa polypeptide (77). During apoptosis, the
150 kDa α-fodrin polypeptide is specifically cleaved by caspase-3, producing a 120 kDa polypeptide. Incubation of caspase-3-WT, caspase-3-S12G and caspase-S36A with the
MCF-7 S-100 cleaved α-fodrin into the 120 kDa polypeptide by 4 h (Fig. 3.8 B lower panel, lanes 4, 8 and 16). However, the caspase-3-S12D and caspase-3-S36D cleaved α- fodrin by 3 h (Fig. 3.8 B lower panel, lanes 11 and 19). These results indicate that phosphorylation of Ser12 and Ser36 promotes the activation of caspase-3 resulting in faster cleavage of substrates during apoptosis.
The increase in α-fodrin cleavage observed in caspase-3-S12D and caspase-3-S36D made us explore whether the phosphorylation was affecting the affinity of caspase-3 for
99 its substrates. For this purpose, we incubated recombinant caspase-3-WT or phospho- mutants, with recombinant caspase-9 for 2 h at 37 ºC and evaluated the ability of the active caspase-3 to cleave the DEVD-AFC substrate. Plots of initial velocity vs. substrate concentration were utilized to determine the parameters Km and kcat for each protein. As shown Table 3.1, the Km of caspase-3-WT, caspase-3-phospho-null and phospho-mimetic mutants were comparable, showing values from 3.57-7.78 μM. These parameters are similar to the recombinant caspase-3-WT values published previously, where it has been reported that caspase-3-WT Km range is between 3 -11 μM (63, 129,
289). The catalytic efficiency (kcat) was also comparable among the proteins analyzed.
The kcat ranged between 1.08 – 1.46 s-1. The catalytic efficiency (kcat/Km) of the proteins was also similar, ranging from 1.61 x 105 – 3.61 x 105. This data shows that the phosphorylation of caspase-3 does not affect the affinity or the catalytic efficiency of the cleavage of substrates.
Taken together, these results suggest an important role of caspase-3-Ser12 and caspase-3-Ser36 as regulators of caspase-3 activation through the modulation of the autocatalytic cleavage and potentially provide a mechanism in which PKCδ regulates caspase-3 processing during apoptosis.
3.3.9 Role of caspase-3-Ser36 phosphorylation in apoptosis
To determine the role of caspase-3 phosphorylation in cell death, we evaluated characteristic phenotypic changes that occur in apoptotic cells. MCF7 cells were transiently transfected with Xpress-tagged caspase-3-WT, caspase-3-S12G, caspase-3- 100
S12D, caspase-3-S36A or caspase-3-S36D and subsequently treated with 200 μM etoposide or diluent-control DMSO for 12 h. The percentage of apoptotic cells with active caspase-
3 and DNA fragmentation was increased from ten percent in the control to 25% after treatment of caspase-3-WT, caspase-3-S12G and caspase-3-S12D MCF-7 cells (Fig. 3.10
A). In contrast, 15% of cells transfected with caspase-3-S36A were apoptotic after treatment, representing 50% fewer apoptotic cells compared to caspase-3-WT. Whereas
45% of caspase-3-S36D-cells were apoptotic (Fig. 3.10 A, caspase-3-WT vs. caspase-3-
S36A, or caspase-3-S36D, time 12 h, P < 0.05) (Fig. 3.10 A).
Next, similar experiments were conducted to investigate the role of caspase-3 phosphorylation in the cleavage of the substrate PARP. The percentage of PARP cleavage was increased by about 20% compared to DMSO treated samples in cells transiently transfected with caspase-3-WT, caspase-3-S12G and caspase-3-S12D (Fig. 3.10
B). However, cells transfected with caspase-3-S36A showed a decrease of ~ 2 fold in cleavage of PARP, compared to caspase-3-WT (Fig. 3.10 B). Cells transfected with caspase-3-S36D showed an increase of 0.5 fold of cells with cleaved PARP, compared to caspase-3-WT (Fig. 3.10 B, caspase-3-WT vs. caspase-3-S36A, time 12 h, P < 0.05).
These data suggests that in cells treated with etoposide caspase-3-Ser12 phosphorylation does not affect apoptosis or the cleavage of the caspase-3 substrate PARP. However, the phosphorylation of caspase-3-Ser12 could play a role in apoptosis induced by different stimuli. The phosphorylation of Ser36 is important for caspase-3 execution of the cell death program and affects the typical apoptotic morphological events.
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3.3.10 Phosphorylation of Ser36 is important for caspase-3 cleavage during
apoptosis
To determine the importance of the caspase-3-Ser36 phosphorylation in caspase-3 activation during cell death, MCF7 cells were transiently transfected with Xpress tagged caspase-3-WT, caspase-3-S36A or caspase-3-S36D and subsequently treated with 200 μM etoposide or DMSO for various times to induce apoptosis (Fig. 3.11 A). Caspase-3-WT was cleaved gradually over time, showing the cleaved active form by 6 h of treatment, at the same time where α-fodrin was cleaved. The cleavage of caspase-3-S36A and α-fodrin were detected by 9 h of treatment showing slower kinetics of activation and cleavage of the substrate. In contrast, caspase-3-S36D was active and cleaved α-fodrin by 3 h.
During apoptosis PKCδ is activated by phosphorylation and by the caspase-3 dependent cleavage producing the 40 kDa catalytically active fragment (Fig. 1.9). The cleavage of PKCδ was analyzed in the transiently transfected MCF-7 cells. PKCδ was cleaved by caspase-3-WT by 3 h of etoposide treatment, while it was detected by 9 h in caspase-3-S36A (Fig. 3.11). In contrast, the cleavage of PKCδ was detected after 1 h in caspase-3-S36D.
Furthermore, the activity of caspase-3 was evaluated using the DEVD-AFC assay
(Fig. 3.11 B). Cells expressing caspase-3-WT showed a gradual time dependent increase in caspase-3 activity during the etoposide treatment. Cells expressing caspase-3-S36A showed lower activity than caspase-3-WT throughout all the times evaluated. In contrast, cells expressing caspase-3-S36D showed an increase in the activity by 1 and 3 h of etoposide treatment, reaching a maximum by 3 h (Fig. 3.11 B). Altogether, these results 102
suggest that the phosphorylation of Ser36 is essential for caspase-3 activation and
cleavage of caspase-3 substrates during cell death.
3.4 Discussion
Caspases are a family of cysteine proteases that are activated through proteolytic
cleavage to execute the apoptotic program. Activation of apoptosis through the extrinsic
or intrinsic pathway causes the oligomerization and autocatalytic cleavage of the
activator caspases; caspase-8 and caspase-9. These caspases cleave and activate effector
caspases, such as caspase-3 (6). The activation of the members of the caspase family is
tightly regulated through positive and negative signals that restrain the apoptotic activity
in healthy cells and activate a controlled cellular demolition when needed. Previously we
showed that caspase-3 is phosphorylated in human monocytes by PKCδ (116). In this
study we further investigated PKCδ phosphorylation of caspase-3 and characterized the
role of the phosphorylation in cell death. We observed that caspase-3 associated
phosphorylation increases in WT and PKCβ-/- MEF during doxorubicin-induced
apoptosis (Fig 3.1), whereas PKCδ-/- MEF showed no increase in the caspase-3-
associated phosphorylation. These results suggest that PKCδ is necessary for caspase-3
phosphorylation.
Next, we mapped the domains of caspase-3 that mediate PKCδ binding and
phosphorylation. We observed that PKCδ interacts and phosphorylates caspase-3-FL in
HeLa cells (Fig 3.2). However, PKCδ showed an increase in the phosphorylation and
binding to the caspase-3-p17 and caspase-3-prop17 of 2 and 5 fold, respectively. 103
However, no phosphorylation or binding to PKCδ was observed in the p12 domain.
These results suggest that caspase-3 phosphorylation and interaction with PKCδ could be structurally regulated in the inactive proenzyme. We could speculate that caspase-3 cleavage by an activator caspase during apoptosis could release the p12 domain allowing
PKCδ interaction and phosphorylation of the caspase-3 p17 and prop17 domains. Further studies are required to assess the role of p12 domain in caspase-3 phosphorylation.
The recognition of small sequences by some protein kinases have been shown to mediate specificity and are important for the efficient substrate phosphorylation (275,
277, 278). ERK1/2 recognizes a specific interaction motif located in caspase-9- prodomain important for the phosphorylation of caspase-9-T125 (290). The MAPK family has well conserved interaction motifs. However, no information is available about an interaction motif required for PKCδ phosphorylation (290, 291). The increase in PKCδ association with the caspase-3-p17 and -prop17 domains suggested the presence of an interaction motif in caspase-3 that could mediate PKCδ binding. Protein linear motifs have been characterized to be sites for protein-protein interaction (285). The analysis of the amino acid sequence of several substrates shown to directly interact with PKCδ revealed common linear motifs. The motifs SGXS (PB1) and QAXXXXXL (PB2) were found in the majority of PKCδ substrates analyzed. PB1 was identified as an important motif for PKCδ interaction and phosphorylation (Fig. 3.3). The conservation of this motif in all substrates analyzed suggests that this motif could be important for PKCδ phosphorylation of target proteins.
104
Next we analyzed the phosphorylated caspase-3 by mass spectrometry analysis and identified 5 sites phosphorylated by PKCδ (Fig 3.4). Sequence analysis showed that all phosphorylation sites are conserved in mammals and this conservation could suggest an important role in the regulation of caspase-3. Previously, a conserved sequence of
DDD (179-181), named “safety catch”, was found to be an important for the inhibition of caspase-3 auto-activation in the absence of apoptosis. We showed that the identified sites are important for PKCδ phosphorylation of caspase-3. Mutation of all sites to a non-phophorylatable alanine or glycine prevented PKCδ phosphorylation of caspase-3
(Fig 3.4 C, D). The mutation of each site to alanine identified the caspase-3-Ser12 and caspase-3-Ser36 as the main PKCδ phosphorylation sites (Fig. 3.4E). Furthermore, the mutation of caspase-3-S12G, caspase-3-S12D or caspase-3-S36A inhibited caspase-3 phosphorylation in HeLa cells (Fig 5). In contrast, mutation of caspase-3-S36D increased caspase-3 phosphorylation (Fig. 5A). The phosphorylation of caspase-3-Ser36 could be a regulatory step that controls the phosphorylation of additional sites. Caspase-3-Ser12 and caspase-3-Ser36 are located in caspase-3-prodomain and p17 domain adjacent to the identified PKCδ interaction motif, in agreement with previous studies that identified binding motifs in close proximity of the phosphorylation sites. Furthermore, caspase-3-
Ser12 and caspase-3-Ser36 are not conserved in other effector caspases, like caspase-6 and
-7. These results are in agreement with in silico analysis of the sequences of these caspases that showed that they lack the identified PKCδ binding motif. Caspase-3 shows high homology with caspase-7 (54%). However, the prodomain and the initial portion of the intermediate domain are highly divergent in these proteins (125-127).
105
We previously showed that phosphorylation of recombinant caspase-3 by PKCδ and subsequent activation results in an increase in activity of caspase-3 of 50% and 30% after 1 and 2 h of incubation with caspase-9, compared to the unphosphorylated caspase-
3. Our results support our previous findings and identify Ser12 and Ser36 as important phosphorylation sites that promote the activity of caspase-3 (Fig. 3.12). In this study, we showed that phosphorylation of caspase-3-Ser12 and caspase-3-Ser36 promoted caspase-3 activity by ~ 50% and ~ 30% after 1 and 2 h of incubation with caspase-9, respectively
(Fig. 3.7 B). In contrast, the phosphorylation of Thr59, Thr67 and Thr77 inhibited caspase-
3 activity in vitro, (Fig. 3.12). Mutation of these sites to alanine did not affect the protease activity of these proteins (Fig. 3.7 A), suggesting that the effect on the activity of caspase-3 is caused by the effect of the phospho mimetic amino acid and not just by the mutation of the sites. These results suggest that PKCδ has the capacity to activate and repress caspase-3 activity and further studies should address the relevance of the phosphorylation of Thr59, Thr67 and Thr77 in apoptosis. Caspase-7 was recently shown to be phosphorylated by PAK2 in breast cancer cells treated with staurosporine inhibiting its activation (115).
Previously we showed that PKCδ phosphorylation of caspase-3 was shown to be important for monocyte apoptosis (116). Since the caspase-3-Ser12 and caspase-Ser36 were shown to be important for PKCδ phosphorylation of caspase-3 and the caspase-3-
S12D and caspase-3-S36D phospho-mimetic mutants were shown to increase caspase-3 in vitro, we focused in these sites to evaluate the role of the phosphorylation in caspase-3 activation and in the execution of apoptosis. Caspase-9 or cell-free system activation of
106 caspase-3-S12D or caspase-3-S36D revealed faster kinetics of the second autocatalytic cleavage (Fig. 3.8) without affecting the affinity of caspase-3 for the DEVD-AFC. The increase in the Km observed in the caspase-3-S12D is within the range observed in the literature; however, it showed similar catalytic efficiency as caspase-3-WT, phospho-null and phospho-mimetic mutants. However, it could reflect a subtle difference in affinity of caspase-3 for the cleavage of substrates when is phosphorylated at Ser12 or Ser36. These data suggest a novel role of the PKCδ phosphorylation of caspase-3 as a regulator of caspase-3 autocatalytic cleavage and activation (Fig. 3.12). Previously, p38 was shown to phosphorylate caspase-3-Ser150 in human neutrophils (113). However, the phosphorylation of caspase-3-Ser150 inhibits caspase-3 protease activity in human neutrophils. We have shown here that caspase-3 associated phosphorylation requires
PKCδ and mutation of caspase-3-S12G and caspase-3-S36A inhibited caspase-3 phosphorylation (Fig 3.5 and 3.6). Caspase-3 phosphorylation may have diverse roles in different cells and further studies should address the role of caspase-3-Ser150 phosphorylation in caspase-3 cleavage.
Furthermore, the phosphorylation of caspase-3-Ser36 was shown to be important for the execution of apoptosis in MCF-7 cells treated with etoposide (Fig. 3.10 B).
Caspase-S36A had a significant reduction in DNA fragmentation and PARP cleavage after etoposide treatment, which are hallmarks of apoptosis (81). Caspase-3-S36D showed significantly higher apoptosis compared to the caspase-3-WT (Fig. 3.10 B). In contrast the phosphorylation of caspase-3-Ser12 did not affect apoptosis or cleavage of
PARP after etoposide treatment. However, the in vitro activation of caspase-3-Ser12
107 suggested that this site was important for caspase-3 activation and replacement of this site reduced caspase-3 phosphorylation (Fig. 3.5-3.8). We hypothesize that Ser12 phosphorylation could play a role in apoptosis induced by a different stimuli. Further experiments should asses the role of caspase-3-Ser12 in cell death.
A time course of etoposide treatment was performed to evaluate the role of caspase-3-Ser36 phosphorylation in apoptosis (Fig 3.10). Cells transfected with caspase-
3-S36A treated with etoposide had lower activity and showed a delay in caspase-3, PKCδ and α-fodrin cleavage, compared to caspase-3-WT. In contrast, cells transfected with caspase-3-S36D showed fast activation, and faster cleavage of caspase-3, α-fodrin and
PKCδ. The release of PKCδ catalytic domain through caspase-3 cleavage has been associated with an amplification of the apoptotic cascade (292-294). The PKCδ dependent phosphorylation of caspase-3 at Ser36 may act in a positive feedback loop to promote caspase-3 activation and cleavage and further activation of PKCδ in the demolition phase of apoptosis. These results show for the first time the requirement of caspase-3 phosphorylation for the execution of apoptosis. These results may explain how caspase-3 is the main effector caspase in the apoptotic cascade, regardless of the caspase-
7 shared affinity for the cleavage of the DEVD-containing substrates. Caspase-7 phosphorylation by PAK2 was shown to inhibit its activity in MCF-7 cells treated with staurosporine, suggesting a different role of caspase-7 in the regulation of chemotherapy induced cell death (115). Previously, the phosphorylation of caspase-9 by c-Abl was shown to be required for the activation of apoptosis, suggesting that the phosphorylation is an important mechanism to regulate the activation of the apoptotic cascade (107).
108
Modulation of caspase-3 phosphorylation may provide a regulatory mechanism that has the potential to control cell death and survival.
Dysregulation of apoptosis has been linked to cancer and inflammatory diseases, such as rheumatoid arthritis and atherosclerosis (295-297). The elucidation of mechanisms that regulate caspase-3 activity is important for the development of effective therapies for diseases that involve the dysregulation of the apoptotic cascade. Cancer treatments target proteins upstream in the apoptotic pathways that induce the activation of caspase-3 (297, 298). However, the development of new strategies that promote caspase-
3 phosphorylation and activation could lead to more effective therapies. The results provided by this study suggest a novel regulatory mechanism to control caspase-3 apoptotic activity and execution of cell death that could be used for the development of new treatments for cancer and other diseases with dysregulated cell fate.
109
Fig. 3.1 PKCδ is necessary for caspase-3 phosphorylation during cell death. A, Caspase-3 (casp-3), -/- -/- PKCδ and PKCβI, expression was analyzed in C57BL/6 MEF WT, PKCδ and PKCβ from mice by immunoblot with anti-casp-3, anti-PKCδ, anti-PKCβI,and β-tubulin antibodies. B, Lysates from MEF treated with 10 μM doxorubicin for 3 h were immunoprecipitated (IP) with anti-casp-3 antibodies or an IgG isotype control, followed by in vitro kinase assays with [γ-32P] ATP and H2B, as an exogenous substrate. Kinase reaction products were resolved by SDS-PAGE and transferred to membranes. Phosphorylated H2B was visualized by autoradiography and the same membrane was immunoblotted with anti-casp-3 antibodies. The results shown are representative of three independent experiments.
110
Fig. 3.2 Identification of caspase-3 domains phosphorylated and involved in the association with PKCδ. A, Lysates from HeLa cells expressing caspase-3 (casp-3) domains (Input) were immunoprecipitated (IP) with anti-Xpress antibodies or IgG isotype control and subjected to in vitro kinase assays in the presence of [γ-32P] ATP. Kinase reaction products were resolved by SDS-PAGE and transferred to membranes. The phosphorylated proteins were visualized by autoradiography. Aliquots of the lysates used in the IPs were run on the same membrane (Input) and were immunoblotted with anti- Xpress antibodies. B, Lysates were IP with anti-PKCδ or IgG isotype control antibodies. The IPs were resolved by SDS-PAGE, transferred to membranes. Aliquots of the lysates used in the IPs were run on the same membrane (input) and immunoblotted with anti-Xpress and anti-PKCδ antibodies. Interaction assays were performed by Dr. Voss (B). The results shown are representative of three independent experiments.
111
Fig. 3.3 Identification of PKCδ interaction motif in caspase-3. A, Schematic representation of PKCδ binding motif in human casp-3-prop17 (Swiss-Prot: P42574), DNA-PK (GenBank: AAA79184.1), γ-Actin (GenBank: CAA27723), Hsp27 (GenBank: BAB17232.1), p53 (GenBank: BAC16799.1), p300 (Swiss- Prot: Q09472.2), and Sp1 (NP_612482.2.1). Conserved amino acids (aa) are highlighted in grey and the regions are indicated. The predicted PKCδ binding (PB) motifs are highlighted in black. B, Schematic representation of caspase-3 clones generated. C-D, Lysates from HeLa cells expressing different caspase-3 (casp-3) constructs were immunoprecipitated (IP) with anti-Xpress antibodies or an IgG isotype control and were subjected to in vitro kinase assays with [γ-32P] ATP. Kinase reaction products were resolved by SDS- PAGE and transferred to membranes. Casp-3 phosphorylation of was visualized by autoradiography and the membranes were immunoblotted with anti-Xpress and anti-PKCδ antibodies. Transiently transfected cells expressed C, casp-3-WT-prop17, casp-3-PBM1-prop17 or casp-3-PBM2-prop17; D, casp-3-WT-FL, casp-3-PBM1-FL or casp-3-PBM2-FL. C-D, interaction assays were performed by Dr. Voss. The results shown are representative of three independent experiments.
112
Fig. 3.4 Identification of PKCδ phosphorylation sites in caspase-3. A, Human purified recombinant casp-3-WT-prop17 was subjected to in vitro kinase assays with [γ-32P] ATP with active or heat inactivated rPKCδ. Phosphorylated kinase products were resolved by SDS-PAGE, transferred to membranes, visualized by autoradiography. The same membranes were immunoblotted with anti-casp-3 and anti-PKCδ antibodies. B, Phosphorylated casp-3-WT-prop17 was analyzed by liquid chromatography mass spectrometry (LC-MS) and the identified sites are represented in the diagram and highlighted in grey. The sequence of human caspase-3 (Swiss-Prot: P42574) was compared to caspase-3 orthologs; including chimpazee (Swiss-Prot: P70677), mouse (Swiss-Prot: P55213), rat (Swiss-Prot: Q5IS54), pig (Swiss-Prot: Q95ND5), chicken (GenBank: AAC32602), and fruit fly (GenBank: AAF55329). C, In vitro kinase assays with [γ-32P] ATP and active rPKCδ performed with rcasp-3-WT-prop17 and rcasp-3-M5-A/G-prop17. D, Immunoprecipitation and in vitro kinase assays of Xpress tagged caspase-3-WT-prop17 or casp-3-M5-A/G- prop17. C-D, Kinase reaction products were resolved by SDS-PAGE and transferred to membranes. The phosphorylation of casp-3 was detected by autoradiography and the membranes were immunoblotted with anti-casp-3 (C) or Xpress antibodies (D). The results shown are representative of three independent experiments.
113
Fig. 3.5 PKCδ phosphorylates caspase-3 Ser12 and Ser36 in vitro. A, Coomassie staining of the 6xHis-tag purified human caspase-3 recombinant proteins. B, Recombinant proteins were phosphorylated in vitro with rPKCδ. Kinase reaction products were resolved by SDS-PAGE and transferred to membranes. Phosphorylation was detected by autoradiography and membranes were immunoblotted with anti-casp-3 antibodies. The results shown are representative of three independent experiments.
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Fig. 3.6 Caspase-3-Ser12 is an important phosphorylation site in HeLa cells. Lysates from HeLa cells expressing different caspase-3 (casp-3) constructs were immunoprecipitated (IP) with an anti-Xpress antibodies or IgG isotype control and subjected to in vitro kinase assays, with [γ-32P] ATP. Kinase reaction products were resolved by SDS-PAGE and transferred membranes. Phosphorylated proteins were visualized by autoradiography and membranes were immunoblotted with anti-Xpress and anti-PKCδ antibodies. Transiently transfected cells expressed: A, casp-3-WT-FL, casp-3-S12G-FL or casp-3-S12D-FL; B, casp-3-WT-prop17, casp-3-S12G-prop17 or casp-3-S12D-prop17. The results shown are representative of three independent experiments.
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Fig. 3.7 The phosphorylation of caspase-3-Ser36 is necessary for the PKCδ dependent phosphorylation of caspase-3. Lysates from HeLa cells expressing different caspase-3 (casp-3) constructs were immunoprecipitated (IP) with an anti-Xpress antibodies or IgG isotype control and subjected to in vitro kinase assays with [γ-32P] ATP. Kinase reaction products were resolved by SDS-PAGE and transferred to membranes and the phosphorylated proteins were visualized by autoradiography. Membranes were immunoblotted with anti-Xpress and anti-PKCδ antibodies. Transiently transfected cells expressed: A, casp-3-WT-FL, casp-3-S36A-FL or casp-3-S36D-FL; B, casp-3-WT-prop17, casp-3-S36A- prop17 or casp-3-S36D-prop17; C, casp-3-WT-prop17, casp-3-S32A-prop17 or casp-3-S36A-prop17. The results shown are representative of three independent experiments.
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Fig. 3.8 Caspase-3 phosphorylation modulates its protease activity in vitro. A-C, 6xHis-tag recombinant caspase-3 proteins were activated in vitro by incubation with recombinant caspase-9 for 0, 0.5, 1, and 2 h at 37˚C, and the activity was monitored by DEVD-AFC assays. The recombinant proteins analyzed were A, casp-3-WT-FL ( ), casp-3-S12G-FL ( ), casp-3-S36A-FL ( ), casp-3-T59A-FL ( ), casp- 3-T67A-FL ( ), and casp-3-T77A-FL( ); B, casp-3-WT-FL ( ), casp-3-S12D-FL ( ), casp-3-S36D-FL ( ),casp-3-T59D-FL ( ), casp-3-T67D-FL ( ), and casp-3-T77D-FL( ); and C, casp-3-WT-FL ( ), casp-3- S12G/S36A-FL ( ) and casp-3-S12D/S36D-FL ( ). Results are expressed as means ± SEM (n = 3). ** P < 0.005 and * P < 0.05.
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Fig. 3.9 Phosphorylation of caspase-3-Ser12 and caspase-3-Ser36 regulates the second autocatalytic cleavage. 6xHis-tag full length recombinant casp-3-WT, casp-3-S12G, casp-3-S12D, casp-3-S36A, and casp- 3-S36D were activated in vitro by incubation with A, recombinant caspase-9 for 0, 0.5, 1, and 2 h at 37˚C; or B, MCF-7 S-100 cell free system for 0, 1, 3, or 4 h at 37 ºC. A-B, Proteins were resolved by SDS-PAGE and immunoblotted with anti-caspase-3 antibodies. B, The cleavage of α-fodrin was analyzed by immunoblot with anti-α-fodrin antibodies. C, Plot of initial velocity (V0) vs. substrate concentration for recombinant casp-3-WT-FL ( ), casp-3-S12G-FL ( ), casp-3-S12D-FL ( ), casp-3-S36A-FL ( ), and casp-3- S36D-FL ( ). The solid lines represent fits to the Michaelis-Menten equation. The parameters for the fits are shown in Table 3.1. Results are expressed as means ± SEM (n = 3). ** P < 0.005 and * P < 0.05.
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Fig. 3.10 Phosphorylation of caspase-3-Ser36 regulates apoptosis. MCF-7 transiently expressing casp-3- WT, casp-3-S12G, casp-3-S12D casp-3-S36A, or casp-3-S36D were treated with 200 μM etoposide (200 µM) or diluent-control DMSO (Ctrl) for 12 h. A, Cells were stained with DAPI, anti-cleaved-active casp-3- FITC and anti-Xpress-Alexa 633 antibodies. B, Quantification of the percentage of apoptotic cells determined by triple staining. C, MCF-7 transiently expressing casp-3 clones were treated with 200 μM etoposide or diluent control were stained with DAPI, anti-cleaved-PARP-FITC and anti-Xpress-Alexa 633 antibodies. D, Quantification of the percentage of cleaved PARP determined by triple staining. Results are expressed as means ± SEM (n = 3). * P < 0.05.
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Fig. 3.11 Phosphorylation of caspase-3-Ser36 is important for caspase-3 cleavage during apoptosis. MCF-7 transiently expressing casp-3-WT, casp-3-S36A-FL or casp-3-S36D-FL were treated with 200 µM etoposide or diluent-control DMSO (Ctrl) for 12 h. A, Lysates were resolved by SDS-PAGE and immunoblotted with anti-Xpress, anti-active-casp-3, anti-α-fodrin, anti-PKCδ, and anti-β-Tubulin antibodies. B, Casp-3 activity was determined by the DEVD-AFC assay for the lysates obtained in A. Results are expressed as means ± SEM (n = 3).
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Fig. 3.12 Model
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Table 3.1 Catalytic parameters
Casp-3-WT S12G S12D S36A S36D Km (μM) 3.57 ± 0.23 4.27 ± 0.44 7.78 ± 0.47 4.06 ± 0.34 4.04 ± 0.44 Kcat (s-1) 1.13 ± 0.02 1.14 ± 0.03 1.25 ± 0.02 1.08 ± 0.02 1.46 ± 0.04 Kcat/Km (M-1 s-1) 3.17x105 2.69x105 1.61x105 2.67x105 3.61x105
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Chapter 4
Characterization of cell fate and immune response of CD14+CD16- and
CD14+CD16+ monocytes
4.1 Abstract
Monocytes are essential components of the innate immune system that circulate for a short period of time and undergo spontaneous apoptosis. These cells are a heterogeneous population that can be subdivided in two main groups: classical monocytes (CD14+CD16-) and non-classical monocytes (CD14+CD16+). We developed a purification scheme and identified differences in spontaneous apoptosis between monocyte subsets. Our results show that CD14+CD16+ cells are more susceptible to undergo spontaneous apoptosis and have significantly higher activity of caspase-3, -8 and
9. Utilizing different pharmacological caspase inhibitors we observed that apoptosis of monocyte subsets is dependent on caspase-3, -8 and -9, suggesting that it occurs though the activation of the intrinsic and extrinsic apoptotic pathways. Analysis of cytokine production showed that CD14+CD16+ express significantly higher TNF-α over time upon
LPS treatment. Our results show that the elevated production of TNF-α observed in
CD14+CD16+ is in part caused by high expression of PKCε. Taken together, these findings show that upregulation of the activity of apoptotic caspases regulates
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CD14+CD16+ cell fate, while under inflammatory conditions the elevated PKCε expression promotes the release of proinflammatory cytokines.
4.2 Introduction
Monocytes are essential components of the innate immune system responsible of the recognition and clearance of pathogens and apoptotic cells (228). Monocytes have a key role in the initiation and resolution of inflammation through the release of pro- inflammatory cytokines and reactive oxygen species (ROS), phagocytosis, and activation of the acquired immune response (235, 299, 300). Recent studies have discovered that blood monocytes are a heterogeneous population that differ in the expression of the cell surface antigens CD14 (receptor for the bacterial lipopolysacharide, LPS) and the low affinity receptor for immunoglobulin G, CD16 (FcγIII receptor) (258). Classical monocytes express CD14 and lack the expression of CD16 (CD14+CD16-); however non- classical monocytes express the CD14 receptor and co-express CD16 (CD14+CD16+)
(258). The CD14+CD16- are the predominant population in healthy individuals and the
CD14+CD16+ account for 5 - 10% of all monocytes (301). The role of CD14+CD16+ in the innate immunity remains unknown.
Monocytes circulate in the bloodstream for 24 - 72 h and undergo apoptosis in the absence of any stimuli (229, 302). Limited information is available about differences in apoptosis in monocyte subpopulations. However, recent studies have suggested that
CD14+CD16+ monocytes are more susceptible to glucocorticoid treatment and oxidative stress than classical monocytes (303, 304). We have previously shown that caspase activation is crucial for apoptosis of classical monocytes (229). Caspases are aspartate-
124 specific cysteine proteases that have a key role in the execution of cell death. The activation of apoptosis could occur through two main pathways, the extrinsic and intrinsic pathways that activate the initiator caspases, caspase-8 and caspase-9, respectively (26). The extrinsic apoptotic pathway involves the activation of the death receptors of the tumor necrosis factor (TNF) receptor superfamily, such as Fas or TNF- related apoptosis inducing ligand (TRAIL) (62). Activation of this signaling pathway results in the activation of the initiator caspase, caspase-8 and the subsequent cleavage of the effector caspase-3. In contrast, the intrinsic apoptotic pathway is activated by permeabilization of the mitochondrial membrane releasing cytochrome c, formation of the apoptosome resulting in the activation of caspase-9. The active caspase-9 cleaves and activates caspase-3 triggering the apoptotic cascade (51, 52). Cross-talk between the two apoptotic pathways has been reported through the caspase-8 dependent cleavage of Bid.
The cleaved Bid translocates and promotes mitochondrial permeabilization and activation of the intrinsic apoptotic pathway (71, 72). The activation of the extrinsic and intrinsic apoptotic pathway converges in the activation of caspase-3. Caspase-3 cleaves nuclear and cytoplasmic proteins that cause the controlled demolition of the cell (26). Caspase-3 can also cleave the effector caspase-6 during apoptosis as part of a positive feedback mechanism to cleave and activate more caspase-8 (26).
Monocyte spontaneous cell death has been shown to be regulated through the extrinsic and intrinsic apoptotic pathways. Monocytes express Fas and the Fas L during spontaneous apoptosis. Treatment with the anti-Fas antibody blocks the interaction between Fas and Fas ligand and the subsequent activation of the extrinsic apoptotic
125 cascade (305). In addition, treatment with the caspase-9 inhibitor LEHD-FMK or the caspase-3 inhibitor DEVD-FMK, prevents monocyte spontaneous apoptosis highlighting the important role of the members of the caspase family in this process (145, 229).
Monocytes escape their apoptotic fate by exposure to inflammatory stimuli, differentiation factors or by malignant transformation, through the inhibition of the caspases (145, 229). During an inflammatory response, exposure to lipopolysaccharides
(LPS) and cytokines such as TNF-α and IL-1β have been shown to inhibit monocyte cell death (229, 230). Diverse studies have shown that CD14+CD16+ monocytes are expanded in inflammatory diseases, such as rheumatoid arthritis, asthma, tuberculosis,
AIDS and sepsis (263-265). Patients with acute and chronic infections express high levels of CD14+CD16+ cells and the treatment with antibiotics has been correlated with a reduction in this monocyte population and clinical improvement on patient’s health (265).
Because of the expansion during inflammatory disease and reduced expression of the anti-inflammatory cytokine IL-10, the CD14+CD16+ monocytes have been associated with a pro-inflammatory phenotype (263-265, 306). More studies are needed to characterize the role of monocyte subsets to be able to develop new therapies for the treatment of inflammatory diseases.
Stimulation of monocytes with lipopolysaccharide (LPS) occurs through the Toll- like receptor 4 (TLR4) (307). During infection, the lipopolysaccharide binding protein
(LBP) recognizes and facilitates the binding of LPS to the CD14 receptor, and then the
CD14 transfers LPS to the TLR4/MD-2 complex (245, 267). LPS recognition induces
TLR4 oligomerization which allows the binding between LPS and MD-2 (243)
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(Fig.1.14). MyD88 binds to the LPS-stimulated TLR4 and recruits the IL-1 receptor associated kinase-1 (IRAK1) that promotes the phosphorylation and activation of the IκB kinase β (IKKβ) (251). IKKβ, part of the IKK complex, phosphorylates the inhibitor of
NF-κB (IκB) causing its degradation and the release of the transcription factor NF-κB
(246, 247, 251). NF-κB translocates to the nucleus and activates the transcription of pro- inflammatory cytokines, including TNF-α (308, 309). Increase in TNF-α production has been implicated in the pathobiology of inflammatory diseases, including rheumatoid arthritis, Crohn’s disease and multiple sclerosis. Suppression of TNF-α expression has been proposed as a strategy to slow down the progression of inflammation (310-312).
The members of the protein kinase C (PKC) family are important regulators of diverse cellular processes such as proliferation, survival, immunity, and cell death. The
PKC family is composed of 11 isoenzymes classified by structural motifs and cofactor requirements (Fig. 1.9). Classical PKCs (α, βI, βII, , and γ) require calcium, 1,2- diacylglycerol (DAG) and phosphatidylserine (PS); novel PKCs (δ, ε, η, and θ) require
DAG for their activation, and atypical PKCs (ζ and λ/ι) activation is calcium and DAG independent (313). The members of the PKC family have been shown to be involved in monocyte differentiation, apoptosis and release of pro-inflammatory cytokines. PKCα and PKCβI have been shown to regulate monocyte differentiation into macrophages and dendritic cells, respectively (188). PKCδ is an important regulator of monocyte apoptosis through the phosphorylation of caspase-3 (116). Phosphorylation promotes the activity of caspase-3 and the cleavage of nuclear and cytoplasmic substrates including PKCδ
(194, 314). PKCδ knockout mice thymocytes are resistant to apoptotic stimuli
127 demonstrating its importance for the execution of apoptosis (315, 316). PKCζ knockout mice have defective NF-κB signaling (187). In addition, PKCε knockout mice have impaired innate immunity and fail to clear bacterial infections (186). Macrophages from the PKCε-/- mice release low levels of TNF-α after stimulation with LPS. Recent studies have linked PKCε to the TLR4 pathway, but its function in this signaling cascade is not well understood (255). Stimulation with LPS recruits PKCε to the TLR4 complex in a
MyD88 dependent manner. Activation of PKCε was involved in the transcriptional activation of NF-κB (255). The expression of PKCs in CD14+CD16+ monocytes has not been analyzed and based on the relevant role of these proteins a better understanding of the expression of the members of the PKC family should provide clues to identify the functional role of monocyte subsets in the innate immune response.
In this study we identified differences in spontaneous apoptosis between different monocytes subsets. We found that CD14+CD16+ cells are more susceptible to undergo spontaneous apoptosis in part caused by upregulation of the activity of the members of the caspase family. We observed that CD14+CD16+ monocytes release higher levels of
TNF-α than classical monocytes, when stimulated with LPS. Expression analysis of the members of the PKC family revealed that CD14+CD16+ have higher expression of PKCε compared to the CD14+CD16- monocytes. Our results implicate PKCε as a critical component of the LPS signaling cascade and may play a role in the pro-inflammatory phenotype associated with CD14+CD16+ monocytes.
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4.3 Results
4.3.1 Optimization of CD14+CD16- and CD14+CD16+ isolation conditions
To study the signaling pathways involved in apoptosis in different monocyte subsets, we developed a purification scheme to isolate CD14+CD16- and CD14+CD16+.
The peripheral blood mononuclear cells (PBMC) were isolated from human buffy coats of normal subjects by a Ficoll gradient to remove plasma, red blood cells and granulocytes. Remaining red blood cells were removed by hypolysis (Fig. 4.1). Next, natural killer (NK), eosinophil and neutrophils were removed by binding to the CD15 and
CD56 receptor, while monocytes and additional lymphocytes eluted in the flow-through
(negative selection). Cells were incubated with anti-CD16 antibodies conjugated to magnetic beads. Cells bound to the column were CD16+ and the flow through contained
CD16- cells. Both CD16+ and CD16- were incubated with CD14-microbeads for the isolation of monocytes. The amount of CD14 and CD16 microbeads per number of cells
(bead/cell ratio) indicated by the manufacturer was 100/100, which corresponded to 100
μl of beads/108 cells. Purification of CD14+CD16- and CD14+CD16+ under these conditions provided population that showed 84% and 73% purity, respectively, as determined by flow cytometry analysis (Table 4.1). Different ratios were tested to improve the purity of the isolated populations. The 80/100 bead to cell ratio provided populations of CD14+CD16- and CD14+CD16+ cells that had 95% and 85% purity, respectively. The relative yield of the purifications was ~ 10 x 106 for CD14+CD16- and
~ 4 x 106 for CD14+CD16+ from per buffy coat (Table 4.1). This purification scheme
129 was utilized for the purification of monocyte subsets utilized in the experiments described in this chapter.
4.3.2 Classical and non-classical monocyte subsets have differences in spontaneous
apoptosis
In healthy individuals the number of CD14+CD16+ cells represents around 5% of the total population of monocytes, but this percentage has been shown to increase in patients with inflammatory diseases (263-265). The reason for the differences in the numbers of monocyte subsets observed is not known. To assess whether monocyte subsets have differences in cell death that play a role in the ratio of monocytes found in healthy individuals, we examined the life span of monocyte subsets measuring the percentage of apoptosis. After 4 h of incubation about 10% of CD14+CD16- and 30% of
CD14+CD16+ were apoptotic, as indicated by Annexin V/7AAD staining (Fig. 4.2). The amount of apoptotic cells increased to 30% for CD14+CD16- and 90% in CD14+CD16+ cells after 8 h. These results show a ~ 3 fold increase in the amount of CD14+CD16+ apoptotic cells after 4 and 8 h (Fig. 4.2). Analysis of the percentage of cells stained only with Annexin V (early apoptosis) and cells stained with Annexin V/7AAD (late apoptosis) show that by 8 h of incubation 25% of the CD14+CD16- cells were in early apoptosis and only 5% were in late apoptosis. In contrast, at 8 h 40% of CD14+CD16- cells were in early apoptosis and 50% were in late apoptosis.
The percentage of active caspase-3 cells increased over time for both monocyte subsets, supporting our observations that these cells are undergoing apoptosis (Fig. 4.2
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B). After incubation for 4 h the percentage of active caspase-3 in CD14+CD16- was 8% and by 8 h increased to 30%, which represents a 10 fold increase, compared to time 0
(Fig. 4.2 A, white bars). In CD14+CD16+ the percentage of active caspase-3 by 4 and 8 h was 35 and 55% apoptosis, respectively, that corresponds to a 2 and 3 fold increase compared to time 0 (Fig. 4.2 B, black bars). The amount of caspase-3-positive cells was
2.5 times higher in CD14+CD16+ after incubation for 4 h, compared to CD14+CD16-. At
8 h incubation, CD14+CD16+ showed 1.5 more caspase-3-positive cells compared to
CD14+CD16- monocytes.
In addition, analysis of the activity of caspase-3 in these samples show that after 4 and 8 h, the CD14+CD16- exhibited 4 and 9 fold increase in the activity of caspase-3, as compared to time 0 (Fig. 4.2 C, white bars). CD14+CD16+ showed 70% and 40% higher activity of caspase-3 by 4 and 8 h, respectively, compared to CD14+CD16- (Fig. 4.2 C, black bars). Supporting these observations, CD14+CD16- showed the second cleavage of caspase-3 (p17) after 8 h, at the same time when PKCδ cleavage was detected. In contrast, in CD14+CD16+ the active caspase-3 p17 polypeptide was detected by 4 h. The cleavage of PKCδ was also detected at 4 h in CD14+CD16+ cells. At 8 h, the amount of the active caspase-3 (second cleavage) and cleaved PKCδ were 4 and 3 times stronger than in CD14+CD16-, respectively. Altogether, these results suggest that CD14+CD16- and CD14+CD16+ have differences in apoptosis.
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4.3.3 CD14+CD16- and CD14+CD16+ apoptosis is dependent on caspase-3
To assess whether caspase-3 activity is necessary for CD14+CD16- and
CD14+CD16+ apoptosis we utilized a pharmacological inhibitor of caspase-3. Treatment with 1μM DEVD-FMK significantly reduced caspase-3 activity by 3 and 5 fold on
CD14+CD16- and CD14+CD16+, respectively, compared to cells undergoing apoptosis treated with the diluent-control DMSO (-) (Fig. 4.3 A, CD14+CD16- vs. CD14+CD16+, 1
μM, caspase-3 P < 0.001). Treatment with 25 μM DEVD-FMK completely inhibited caspase-3 activity in monocyte subsets (Fig. 4.3 A, CD14+CD16- vs. CD14+CD16+, 25
μM, caspase-3 P < 0.001). The cleavage of Bid was analyzed as marker of apoptosis.
After incubation for 8 h, CD14+CD16+ showed more cleavage of Bid than CD14+CD16-
(Fig. 4.3). Inhibition of caspase-3 by treatment with 1 and 25 μM DEVD-FMK gradually prevented the cleavage of Bid in CD14+CD16- and CD14+CD16+. These results suggest that monocyte subsets undergo apoptosis in a caspase-3-dependent manner.
4.3.4 Caspase cascade in monocyte subsets
The activity of initiator caspases was analyzed to assess their role in the differences in spontaneous apoptosis observed in CD14+CD16- and CD14+CD16+. The activity of caspase-8 was increased by 6 fold in CD14+CD16- and 10 fold in
CD14+CD16+ compared to freshly isolated monocytes. The activity of caspase-8 was significantly higher in CD14+CD16+ compared to the CD14+CD16- after 8 h (Fig. 4.4 A,
CD14+CD16- vs. CD14+CD16+, time 8 h, caspase-8 P < 0.005). Analysis of caspase-9
132 activity showed a 5 fold increase in CD14+CD16- compared to 7 fold increase in
CD14+CD16+ 8 h. The activity of caspase-9 was ~ 3 fold higher in CD14+CD16+ compared to classical monocytes (Fig. 4.4 B, CD14+CD16- vs. CD14+CD16+, time 8 h, caspase-9 P < 0.05). These results show that both caspase-8 and caspase-9 are activated during spontaneous apoptosis and suggests differences in the temporal regulation of caspase activation in different monocyte subsets could be involved in the increase in apoptosis observed in CD14+CD16+.
To define the caspase cascade involved in monocyte apoptosis, monocytes undergoing spontaneous apoptosis were treated with caspase-8 and caspase-9 pharmacological inhibitors. CD14+CD16- and CD14+CD16+ showed a reduction in caspase-8 activity of 2 and 3 fold after treatment with 1 μM IETD-FMK, compared to
DMSO treated samples, respectively (Fig. 4.4 C, grey bars). The inhibition of caspase-8 activity significantly reduced caspase-3 activity in CD14+CD16- by 2.5 fold and by 3 fold in CD14+CD16+, compared to controls (Fig. 4.4 A, dashed bars) (Fig. 4.4 A,
CD14+CD16- and CD14+CD16+, DMSO vs. IETD-FMK, 8 h, P < 0.0001).
Inhibition of caspase-9 by treatment of LEHD-FMK decreases caspase-9 activity by about 2 and 4 fold in CD14+CD16- and CD14+CD16+, respectively (Fig. 4.4 D, grey bars). However, LEHD-FMK treatment of monocyte subsets inhibited caspase-3 activity by ~ 3 and 3.5 fold in CD14+CD16- and CD14+CD16+, respectively (Fig. 4.4 D, dashed bars). The inhibition in the activity of caspase-9 and caspase-3 were statistically significant for both monocyte subset (Fig. 4.4 D, CD14+CD16- and CD14+CD16+, DMSO vs. LEHD-FMK, 8 h, caspase-9 activity P < 0.005 and P < 0.05, respectively; and
133 caspase-3 P < 0.005). These results suggest that the activity of caspase-3 and apoptosis of CD14+CD16- and CD14+CD16+ depends in the activation of caspase-8 and caspase-9 suggesting that the spontaneous apoptosis of monocyte subsets occur by activating the intrinsic and extrinsic pathway.
4.3.5 LPS-induced TNF-α production is elevated in non-classical monocytes
Increased production of TNF-α has been found in inflammatory diseases (317). In addition, increased numbers of CD14+CD16+ have been reported during inflammation
(263-265). To evaluate the contribution of each monocyte subset to LPS-induced TNF-α release, CD14+CD16+ and CD14+CD16- were stimulated with 10 ng/ml LPS, for different times or PBS as diluent control (Fig. 4.5 A). LPS-stimulation resulted in a time dependent increase in TNF-α production for both monocyte subsets. The levels of TNF-α were significantly higher over time in CD14+CD16+ compared to CD14+CD16- cells.
After incubation for 4 and 8 h, CD14+CD16+ released 3.5 and 5 times more TNF-α than
CD14+CD16- (Fig. 4.5 A, CD14+CD16- vs. CD14+CD16+, 4 h P < 0.05, 8 h P < 0.005).
Next, the release of TNF-α was evaluated in human monocytes upon clustering the FcγR receptor (Fig. 4.5 B). The CD14+CD16+ cells release a significantly higher amount of TNF-α compared to CD14+CD16- (Fig. 4.5 A, CD14+CD16- vs. CD14+CD16+,
P < 0.05). These results suggest that CD14+CD16+ have a more pro-inflammatory response to LPS and by FcγR clustering compared to classical monocytes.
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4.3.6 Expression of PKC-isoforms in different monocyte subsets
The members of the PKC family are important modulators of monocyte cell fate and immunological response (116, 186). To determine the contribution of PKCs in the differences observed in apoptosis and immune regulators in monocyte subsets the expression of PKC isoenzymes was investigated in CD14+CD16- and CD14+CD16+.
Monocyte subsets expressed similar levels of classical PKCs (PKCα, PKCβI and PKCβII).
No significant differences in the expression of the atypical PKCζ were observed in monocyte subsets, whereas PKCγ and PKCι/λ were not detected. In the case of novel
PKCs, CD14+CD16- and CD14+CD16+ express similar level of PKCδ and very low but similar expression of PKCη and PKCθ were observed (Fig. 4.6). In contrast, the expression of PKCε was significantly higher in CD14+CD16+ (Fig. 4.6 A and B,
CD14+CD16- vs. CD14+CD16+ n=7, P < 0.005). The elevated expression of PKCε may help explain some of the differences observed between monocyte subsets.
4.3.7 The expression of PKCε is necessary for elevated LPS-induced TNF-α release
in CD14+CD16+ monocytes
To determine whether differences in expression of PKCε play a role in the immune response mediated by monocytes, LPS-induced TNF production was evaluated.
The expression of PKCε was reduced in 50% in CD14+CD16- and CD14+CD16+ transfected with siRNA-PKCε (Fig. 4.7 A). CD14+CD16- and CD14+CD16+ transfected with siRNA-control released ~ 6000 and ~ 12,000 pg, respectively, compared to samples
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treated with PBS control. The release of TNF-α in monocyte subsets was significantly
reduced by silencing PKCε expression. CD14+CD16- showed a reduction of ~ 30% in
TNF-α expression by silencing PKCε expression (Fig. 4.7 B, CD14+CD16-, control vs.
PKCε siRNA treated with LPS P < 0.05). However, silencing PKCε expression in
CD14+CD16+ reduced TNF-α expression by ~ 50% (Fig. 4.7 B, CD14+CD16+, control vs.
PKCε siRNA treated with LPS P < 0.05). Interestingly, the levels of TNF-α observed in
CD14+CD16+ after silencing PKCε expression were similar as the classical monocytes
transfected with siRNA-control. These demonstrate that the elevated production of TNF-
α observed in CD14+CD16+ cells stimulated by LPS is in part caused by high expression
of PKCε.
4.4 Discussion
Monocytes play an important role in the innate immune response and recent studies have characterized these cells as heterogeneous populations (116, 228, 229).
Previous studies have shown that CD14+CD16- and CD14+CD16+ have differences in abundance in the blood; however in inflammatory diseases the ratios of monocyte subsets are altered (264, 301). The differences in the abundance of monocyte subsets may suggest a differential response to apoptosis; however monocyte subset apoptosis is not well understood.
Previous studies evaluated changes in cell fate and immune response of classical monocytes. At the time there was no knowledge about the heterogeneous monocyte subsets and the techniques utilized to purify them did not allow the separation of these
136 cells. Recent technology has taken advantage of the differences in the expression of the
CD16 receptor to isolate the main monocyte populations, the CD14+CD16- and
CD14+CD16+. Optimization of the isolation techniques allow the isolation of monocyte subsets with higher purity and lower amount of contaminant cells that allow the characterization of CD14+CD16- and CD14+CD16+ to understand their biological roles in cell death and immune response. Changes in the bead/cell ratio allowed us to isolate
CD14+CD16- and CD14+CD16+ with purities of 95% and 85%, which is comparable to the purities reported by previous studies (Fig. 1.1) (260).
Previously, CD14+CD16+ monocytes were reported to be more susceptible to glucocorticoids, suggesting that treatments with anti-inflammatory drug may deplete the non-classical monocytes as part of their mechanism of action in an anti-inflammatory therapy. In addition, a recent article reported that CD14+CD16+ cells are more susceptible to oxidative stress induced apoptosis (304). This study shows that
CD14+CD16+ exhibit higher susceptibility to undergo spontaneous apoptosis compared to classical monocytes (Fig. 4.2). Both monocyte subsets undergo apoptosis in a caspase- dependent manner (Fig. 4.3 A). We observed significantly higher activity of caspase-3, -
8, and -9 in CD14+CD16+ non-classical monocytes (Fig. 4.3 and 4.4). Consistently, the increase in caspase activities was accompanied by higher levels of cleaved caspase substrates, including PKCδ and Bid (Fig. 4.3). These results suggest that CD14+CD16+ cells have different life span than CD14+CD16-. The elevated apoptosis observed in the
CD14+CD16+ monocytes may act as a regulatory mechanism to suppress their abundance in healthy patients.
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It has been reported that oxidative stress induces Fas-mediated apoptosis in classical monocytes (318). In addition, classical monocytes undergo apoptosis after phagocytosis of pathogens in a Fas-receptor mediated pathway, which activates caspase-8 before changes in mitochondrial membrane potential (319). In T and B lymphocytes it has been shown that induction of apoptosis by etoposide treatment causes activation of caspase-8 in a caspase-3 and caspase-6 dependent manner, in the absence of the DISC formation (320). Our studies support a type II Fas mediated apoptosis during apoptosis of both monocyte subsets. The cleavage of Bid has been shown to mediate the cross-talk between the extrinsic and intrinsic pathway (71, 72). These studies support our findings suggesting that human monocyte subsets undergo spontaneous apoptosis activating the intrinsic and extrinsic pathway.
We have previously shown that classical monocytes undergo apoptosis in a caspase-3 dependent manner (229). However, the role of caspase-3 in CD14+CD16+ apoptosis is not understood. The inhibition of caspase-3 with the DEVD-FMK prevented the cleavage of the caspase-8 substrate Bid in both monocyte subsets treated with DEVD-
FMK (Fig.4.3). Utilizing pharmacological inhibitors we observed that caspase-3 activity is regulated by caspase-8 and caspase-9 in both monocyte subsets (Fig. 4.4). These results demonstrate the involvement of the extrinsic and intrinsic apoptotic pathways in the activation of caspase-3 for the execution of apoptosis.
The CD14+CD16+ monocytes release low levels of the anti-inflammatory cytokine
IL-10 and are expanded during inflammatory diseases (264, 266, 301). Analysis of the release of pro-inflammatory cytokines, such as TNF-α, IL-1β and IL-6, have been
138 conflicting since some reports showed reduced or no differences in the expression of these cytokines, while others showed increase in expression of TNF-α, concomitant with an increase in density of CD14+CD16+ cells (266, 321). However, many studies have been done analyzing whole blood making it difficult to assess differences in cytokine release between monocyte subsets (265, 266, 321, 322). Our results showed that isolated
CD14+CD16+ stimulated with LPS produced significantly higher TNF-α than
CD14+CD16- (Fig. 4.5 A). In addition, we showed that CD14+CD16+ cells release significantly higher TNF-α as a result of Fcγ receptor clustering compared to
CD14+CD16- cells (Fig. 4.5 B). Hence, these results may suggest that upon different stimuli CD14+CD16+ produce higher amounts of TNF-α supporting their role as the pro- inflammatory subset.
In an inflammatory environment, exposure to LPS and secretion of cytokines such as TNF-α and IL-1β have been suggested to inhibit monocyte cell death; however, in the absence of any stimuli these cells undergo spontaneous apoptosis (230). The higher susceptibility of CD14+CD16+ monocytes to undergo apoptosis may control the accumulation of the pro-inflammatory monocytes in the absence of inflammation and could play an important role in the resolution phase of the inflammatory response. The
CD14+CD16+ monocytes have been shown to be expanded in inflammatory diseases, such as Crohn’s disease, rheumatoid arthritis, asthma, sarcoidosis, pancreatitis, and sepsis, among others (264, 301). Interestingly, patients with acute and chronic infections express high levels of CD14+CD16+ cells and antibiotics treatment correlates with a reduction in these cells and clinical improvement on patient’s health (265). The increase
139 in numbers of non-classical monocytes during inflammation may suggest an inhibition of the apoptotic cascade in CD14+CD16+ cells in inflammatory diseases. CD14+CD16+ cells showed significantly inhibition of caspase-3, -8 and -9 when treated their respective inhibitors (Fig. 4.3 and 4.4). Since monocyte differentiation and stimulation have been shown to inhibit monocyte cell death, it is tempting to hypothesize that the accumulation of CD14+CD16+ in inflammatory diseases may occur by inhibition of the apoptotic machinery allowing their survival. Further studies are needed to determine how the inflammatory environment changes the susceptibility of CD14+CD16+ monocytes to undergo spontaneous apoptosis.
Diverse studies have analyzed the expression of the members of the PKC family in classical monocytes; however, this is the first report to document differences in the expression of PKCs between CD14+CD16- and CD14+CD16+ monocytes. CD14+CD16+ had higher expression of PKCε, compared to classical monocytes (Fig. 4.7). The high expression of PKCε observed in CD14+CD16+ may suggest an important role in the expression of pro-inflammatory cytokines, since PKCε-/- mice are highly sensitive to
Gram-negative and Gram-positive bacteria (186). Also, macrophages from PKCε-/- mice have reduced expression of TNF-α and IL-1β in response to LPS and IFNγ due to a reduction in NF-κB activation. In addition, PKCε was shown to be necessary for NF-κB activation in of monocytes stimulated with the bacterial chemoattractant N-formyl- methionine-leucine-phenylalanine (FMLP) (256). Stimulation of monocytes with LPS activates the TLR4 pathway and activates a signaling cascade that mediates the transcription and release of pro-inflammatory cytokines through the activation of NF-κB.
140
More evidence linked PKCε with the TLR4 pathway but the role of this kinase in the signaling cascade was unknown. A study identified that PKCε recruited to the TLR4 in a
MyD88 dependent manner, linking it to the initial steps of the activation of TLR4 pathway (255). This study shows that silencing expression of PKCε reduced LPS- induced TNF-α release by 35% and 50% in CD14+CD16+ and CD14+CD16-, respectively
(Fig. 4.7). The expression stimulated with LPS was at comparable levels as the TNF-α expression levels observed in CD14+CD16- monocytes. These results suggest that the elevated PKCε expression may mediate the high production of TNF-α and pro- inflammatory phenotype associated with non-classical monocytes.
In summary, our results show differences in spontaneous apoptosis and immunological response of monocyte subpopulations (Fig. 4.8). CD14+CD16+ monocytes are highly susceptible to apoptosis due to the upregulation in the activity of members of the caspase family. These cells release significantly higher amounts of TNF-
α in response to LPS in part due to the elevated expression of PKCε. These results support a model in which in healthy individuals the accumulation of the pro-inflammatory
CD14+CD16+ monocytes is regulated through apoptosis to maintain homeostasis.
Exposure to pathogens and activation by FcγR-binding may prevent cell death through inhibition of the members of the caspase family and allow the accumulation of
CD14+CD16+, observed during inflammatory diseases. Elevated expression of PKCε mediates activation of the TLR4 signaling pathway that mediates high release of TNF-α and may promote faster recruitment of leukocytes to the site of infection. However, further evaluation CD14+CD16+ dynamics in the presence or absence of infection may
141 provide new mechanisms to modulate the density and immunological response of
CD14+CD16+ in inflammatory diseases.
142
Fig. 4.1 Monocyte subsets purification scheme. CD14+CD16- and CD14+CD16+ cells were isolated from the PBMC of healthy donors through a Ficoll density gradient and magnetic cell sorting. Cells were stained with anti-CD14-APC and anti-CD16-PE antibodies. Purity of monocyte subpopulations was assessed by flow cytometry.
143
Fig. 4.2. Classical and non-classical monocyte subsets have differences in spontaneous apoptosis. CD14+CD16- and CD14+CD16+ cells undergoing spontaneous apoptosis cultured for 4 and 8 h. A, Percentage of apoptotic cells was determined by flow cytometry of cells labeled with Annexin V and 7AAD. B, Percentage of cleaved caspase-3 determined by flow cytometry of cells labeled with active caspase-3-FITC antibody. C, Activity of caspase-3 was determined by DEVD-AFC assay. D, Presence of full length and cleaved active caspase-3 and PKCδ were analyzed by immunoblot. Same membranes were re-blotted with β-tubulin antibodies. Results are expressed as an average ± SEM (n=3). ** P < 0.005 and * P < 0.05.
144
Fig. 4.3 CD14+CD16- and CD14+CD16+ monocytes require caspase-3 to undergo apoptosis. A, The activity of caspase-3 was measured by the DEVD-AFC assay in freshly isolated monocytes (NT), cells treated for 8 h with 1, 25 μM DEVD-FMK or diluent-control (DMSO). B, Cell lysates were resolved by SDS-PAGE and were immunoblotted with, anti-Bid and anti-β-Tubulin antibodies. Results are expressed as an average ± SEM (n=3). *** P < 0.0001.
145
Fig. 4.4 The activation of caspase-3 during monocyte apoptosis is dependent on caspase-8 and caspase-9. Caspase activity was determined in freshly isolated monocytes (NT) or after incubation for 8 h. A, Caspase-8 activity was measured by the IETD-AFC assay. B, Caspase-9 activity was measured by the LEHD-AFC assay. C-D, The activity of caspase-3, caspase-8 and caspase-9 was determined by the DEVD-AFC, IETD-AFC and LEHD-AFC assay, respectively, in freshly isolated monocytes (NT), cells treated for 8 h with (C) 1 μM IETD-FMK, (D) 1 μM LEHD-FMK or diluent-control (DMSO). Results are expressed as mean ± SEM (n=3). *** P < 0.0001, ** P < 0.005 and * P < 0.05.
146
Fig. 4.5. TNF-α production in monocyte subsets. A, Monocyte subsets were stimulated with 10 ng/ml for 2, 4, and 8 h. B, Monocyte subsets were incubated with 10 μg/ml immobilized human with IgG for 16 h to cluster FCRγ receptor. TNF-α was measured by ELISA. Results are expressed as an average ± SEM (n=3). ** P < 0.005 and * P < 0.05.
147
Fig. 4.6 Expression of PKC isoforms in monocyte subsets. A, Cell lysates from 2 donors and recombinant PKCs (control) were resolved by SDS-PAGE and immunoblotted anti-PKC and β-Tubulin antibodies. B, Relative expression of PKCs were normalized by β-tubulin. Results are expressed as ± SEM (n=7). ** P < 0.005.
148
Fig. 4.7 The expression of PKCε is necessary for high TNF-α release in CD14+CD16+ monocytes. A, PKCε expression in cells electroporated with siRNA-PKCε or siRNA-control. B, Monocyte subsets were stimulated with 10 ng/ml LPS for 8 h. Supernatants were collected and the secreted TNF-α was measured by ELISA. * P < 0.05.
149
Fig. 4.8. Summary of the results for the characterization of cell death and immune response in monocyte subsets.
150
Bead/cell ratio Cell concentration (x 106) Purity (%) (%)
CD14+CD16- CD14+CD16+ CD14+CD16- CD14+CD16+ 100/100 85 9 84 73 100/80 80 8 87 77 80/80 108 11 88 70 80/100 75 ± 7 4 ± 1 95 85
Table 4.1 Optimized monocyte subset isolation conditions. CD14+CD16- and CD14+CD16+ cells were isolated from the PBMC of healthy donors through a Ficoll density gradient and magnetic cell sorting. Ratio of magnetic bead to cells was modified and the cell purity and concentration were determined. Cells were stained with anti-CD14-APC and anti-CD16-PE antibodies. Purity of monocyte subpopulations was assessed by flow cytometry.
151
Chapter 5
Conclusions and Further Directions
The members of the caspase family are important regulators of apoptosis (6).
Several studies have revealed that the balance between cell death and survival is controlled by complex regulatory networks that maintain cell homeostasis (323).
Dysregulation in this balance is associated with pathologies including cancer, inflammatory diseases and sepsis (4, 39, 92, 324). Caspase-3 has a central role in the apoptotic cascade since it is activated by the extrinsic and intrinsic apoptotic pathways and is the main effector protease responsible for the cleavage of structural and regulatory proteins during the demolition phase of apoptosis (26). The activity of caspase-3 is positively and negatively regulated to maintain homeostasis and allow its activation when needed. Diverse studies have shown that caspase-3 is regulated by protein-protein interaction and post-translational modifications, including phosphorylation (93, 97, 113,
116, 176). We have shown here that caspase-3 is phosphorylated during apoptosis and the expression of PKCδ is necessary for the phosphorylation (Fig. 3.1). Elucidation of the role of PKCδ phosphorylation of caspase-3 in cell death is essential to understand the pathways that control cell fate. For this purpose, we mapped the domains of caspase-3 that are phosphorylated and interact with PKCδ (Fig. 3.2 and 3.3). We identified the caspase-3 prop17 and p17 domains as the main subunits phosphorylated in involved in the association with PKCδ. Our results showed that the C-terminus domain of caspase-3
152 is not phosphorylated nor associates with PKCδ. However, our results suggest that the p12 may have a repressive role in the phosphorylation of caspase-3, since the caspase-3 full-length containing the p12 domain exhibits reduced phosphorylation compared to all other domains that lack the p12 C-terminal domain (Fig. 3.2). Preliminary experiments suggest that the removal of the p12 domain enhances PKCδ binding and phosphorylation of caspase-3-prop17. Our studies suggest that the cleavage of caspase-3 releases the p12 domain enhancing the phosphorylation of the prop17 and p17 domains. Further studies should address the role of the p12 domain in the regulation of caspase-3 phosphorylation and binding to PKCδ.
We further identified a motif in caspase-3 necessary for PKCδ binding and phosphorylation of caspase-3 (Fig. 3.3, 5.1). This motif is located in an unstructured region in the p17 domain at the boundary with the prodomain. Caspase-3-prodomain and initial portion of the p17 domain are predicted be unstructured in nature, based on the procaspase-7 crystal structure (127). Interestingly, linear motifs or unstructured regions have also been suggested to be enriched in sites that can be phosphorylated (325). For example, v-cyclin-CDK6 phosphorylates two serine residues in an unstructured loop of
Bcl-2 inhibiting its anti-apoptotic activity (326). In this study we identified the SGIS motif located in close proximity to the two main sites in caspase-3 phosphorylated by
PKCδ (Fig. 3.4). The SGIS is located 17 amino acids upstream of Ser12 and three amino acids downstream of Ser36. This interaction motif was found to be necessary for caspase-
3 phosphorylation. Replacement of the SGIS motif inhibited caspase-3-FL phosphorylation and binding to PKCδ. Interestingly, we observed a significant reduction
153 in PKCδ binding and phosphorylation by mutation of the SGIS in caspase-3-prop17, but it was not completely inhibited. These results suggest that the binding of PKCδ to the prop17 could be stabilized by residues located in the prodomain. This hypothesis is supported by the increase in PKCδ binding and phosphorylation of caspase-3 in the prop17 domain in comparison to the p17 domain (Fig. 3.2).
The conservation of the SGXS motif in PKCδ substrates suggest that this sequence provides specificity for the efficient phosphorylation of substrates. Preliminary experiments suggest that the SGXS motif is also necessary for PKCδ phosphorylation and binding to Hsp27. Previously, classical PKCs were shown to require a 27 amino acid motif for the binding and phosphorylation of p53, but it was not required for the binding of PKCδ (273). Our results show that p53 only contains an SXXS motif instead of the
SGXS motif found in other PKCδ substrates. In addition, we could not identify the
SGXS motif in the sequences of other effector caspases, including caspase-6 and caspase-
7, suggesting that they are not targets of PKCδ. Further studies are required to evaluate whether the SGXS motif is necessary for PKCδ binding and phosphorylation of substrates. Previous studies identified the SGIS motif as the binding site of cIAP1 in the caspase-3-prop17 (Fig. 5.1). The effect of the binding of PKCδ in the interaction between cIAP and caspase-3 is unknown. We could hypothesize that during apoptosis, the binding of PKCδ to caspase-3 may release the inhibitor and allow the execution of cell death. Further studies should address how the interaction between caspase-2 and cIAP is modulated by the binding to PKCδ (101). The identification of PKCδ binding motif and the role of the binding during apoptosis could potentially be used for the
154 development of therapies to target this kinase in diseases such as cancer and arthrosclerosis, associated with PKCδ activity (189).
The phosphorylation of caspase-3 by PKCδ was further investigated and we identified five residues by mass spectrometry (Fig. 5.1). We observed that mutation of these sites prevented the phosphorylation of the caspase-3-prop17 (Fig. 3.4). The identified sites showed great evolutionary conservation suggesting that they may play an important role in the regulation of caspase-3. The sites Ser12, Ser36 and Thr59 were conserved in vertebrates; Thr67 was conserved from humans to fruit fly; and Thr77 was conserved in mammals and fruit fly. Mutation of the individual sites revealed different contributions to caspase-3 phosphorylation (Fig. 3.5). We identified Ser12 and Ser36 as the main sites in caspase-3 phosphorylated by PKCδ (Fig. 3.5-3.7).
We analyzed the effect of the phosphorylation of these sites in the activity proteolytic activity of caspase-3 and showed that the phosphorylation could promote or repress caspase-3 activity (Fig. 3.8). Phosphorylation of Thr59, Thr67 and Thr77 repressed caspase-3 activity, while the phosphorylation of Ser12 and Ser36 promoted caspase-3 activity in vitro (Fig. 5.1). Since the mutation of these sites to alanine did not affect the proteolytic activity of these proteins we consider that the effect in the activity of caspase-
3 is caused by the effect of the phospho mimetic amino acid and not caused just by the mutation of the sites. Interestingly, the mutation of Thr59, Thr67 and Thr77 had less contribution to caspase-3 phosphorylation by PKCδ. These results suggest that PKCδ has the capacity to activate and repress caspase-3 activity. However, further studies should address the relevance of the phosphorylation of Thr59, Thr67 and Thr77 in apoptosis.
155
While another study showed that the phosphorylation of caspase-3 by p38 inhibits neutrophil apoptosis, our results suggest PKCδ responsible for caspase-3 phosphorylation, and we identified Ser12 and Ser36 as the main phosphorylation sites in vivo and in vitro. We have shown that PKCδ is necessary for caspase-3 phosphorylation in doxorubicin-induced apoptosis in MEF (Fig 3.1). In addition, we demonstrated that null-mutations caspase-3-S12G and caspase-3-S36A inhibit caspase-3 phosphorylation on the full length and prop17 caspase-3 in HeLa cells (Fig. 3.6-3.7). Thus, the phosphorylation of Ser150 may be specific mechanism for the repression of caspase-3 in neutrophils.
For the first time we show that phosphorylation of caspase-3 promotes its activation and apoptosis (Fig. 3.8-3.11). Other caspases have been shown to be negatively regulated by phosphorylation (102-106, 110, 111, 113, 115, 327), and so far only the phosphorylation of caspase-9 at Tyr153 was shown to be necessary for the execution of cell death (107). We have shown here that Ser12 and Ser36 have an important role in the activation of caspase-3 (Fig. 3.8). These observations are in agreement with our previous observations that showed that phosphorylation of recombinant caspase-3 by PKCδ followed by activation with caspase-9 results in an increase in activity that is about two times compared to the non-phosphorylated caspase-3
(116). We have shown here phosphorylation of Ser12 and Ser36 promotes the kinetics of the autocatalytic cleavage without affecting the affinity of caspase-3 for the synthetic substrate DEVD-AFC (Fig. 3.9, 5.1). We further evaluated the role of these sites in apoptosis and showed that Ser36 is necessary for caspase-3 execution of cell death.
156
Inhibition of Ser36 phosphorylation repressed caspase-3 activity, prevented the cleavage of target proteins and inhibited apoptosis in MCF-7 cells treated with etoposide (Fig.
3.10-3.11). In contrast, the phosphorylation of Ser36 promoted apoptosis suggesting that this site is a key regulator of caspase-3 activity. Ser12 phosphorylation did not reflect an effect in cells treated with etoposide. However, we cannot disregard the potential role of the phosphorylation of Ser12 under different stimulus. Further studies should assess the role of Ser12 phosphorylation in cells induced to undergo apoptosis through stimulus that activate the intrinsic and extrinsic pathway.
We have described a mechanism by which caspase-3 activation is regulated by
PKCδ phosphorylation. The phosphorylation of caspase-3 by PKCδ enhances the apoptotic signaling cascade in cells treated with etoposide. As previously shown, we observed a positive feedback loop in the activation of caspase-3 and cleavage of PKCδ in cells undergoing apoptosis (Fig. 3.11 A, 5.1). Caspase-3-S36D showed faster kinetics of
PKCδ cleavage, suggesting that the positive feedback activation between caspase-3 and
PKCδ occurs and may play a role in the increase in apoptosis (Fig. 3.11 A). In contrast with these results, caspase-7 was recently shown to be negatively regulated by PAK2 phosphorylation during chemotherapeutic-induced apoptosis (115). The difference in the role of the phosphorylation in the activities of these effector caspases could explain why caspase-3 is the main effector caspase during apoptosis.
The interaction between caspase-3 with Hsp27 inhibits caspase-3 second autocatalytic cleavage and activation (176). Since our studies reflect that the phosphorylation of Ser12 and Ser36 promotes the kinetics of the second autocatalytic
157 cleavage it would be important to assess how the phosphorylation of these sites affect the interaction between caspase-3 and Hsp27 (Fig. 5.1). In addition, we have recently shown that PKCδ phosphorylates Hsp27 at Ser15 and Ser82 in apigenin-induced apoptosis of leukemia cells (218). Unpublished work in our lab revealed that the phosphorylation of
Hsp27 by PKCδ prevents its binding to caspase-3 (Gonzalez-Mejia and Doseff). Thus,
PKCδ regulates important steps in the apoptotic cascade by phosphorylating Hsp27 and caspase-3. Further work should evaluate how the PKCδ dependent phosphorylation affects caspase-3 and Hsp27 binding.
An understanding of the mechanisms that regulate caspase-3 activation is an important step towards effective therapies for a variety of diseases related to dysregulation of the apoptotic pathway. In the case of cancer, multiple therapies have been developed to target upstream events in the apoptotic cascade (298). Therapies that target the members of the Bcl-2 family activate a signaling cascade that results in the activation of caspase-3 (328). The development of new therapies that target the effector caspases would be of great interest. Recently, a synthetic peptide was designed to directly target caspase-3 in tumor cells (149, 329). The synthetic peptide S-PAC-1 activates caspase-3 and has been shown to be effective for the treatment of leukemia (39).
Thus, the development of therapies that regulate caspase-3 phosphorylation could be effective for the treatment of diseases with dysregulated apoptosis.
We have previously shown that caspase-3 is important for monocyte apoptosis
(116). At the time were that study was done, little information was available about the monocyte heterogeneity. Recent studies have characterized the monocyte subpopulations
158 and have shown that the main subsets, CD14+CD16- and CD14+CD16+ are differentially represented in the blood of healthy individuals (258). CD14+CD16- represents 90% of the total monocytes, while the CD14+CD16+ represents 5% (301). A recent article reported that CD14+CD16+ cells are more susceptible to oxidative stress induced apoptosis, which suggested that the abundance of these cells could be regulated through apoptosis (304). We analyzed whether these monocyte subsets have differences in apoptosis that could explain differences in the representation of these cells in the blood.
We observed significantly higher spontaneous apoptosis in CD14+CD16+ compared to
CD14+CD16- (Fig. 4.2). The activity of caspase-3, -8 and -9 was significantly higher in these cells suggesting that upregulation of the activity of these proteases play a role in the higher susceptibility of these cells to undergo spontaneous apoptosis (Fig. 4.3).
We previously analyzed the requirement of caspase-3 for monocyte apoptosis.
However, at that time the techniques utilized for the purification of monocytes did not separate the two subsets and due to the lower abundance of CD14+CD16+ it is unknown whether these monocytes undergo apoptosis through the same pathway as the
CD14+CD16- monocytes. Our studies revealed that CD14+CD16+ also require caspase-3 to undergo spontaneous apoptosis (Fig. 4.3). We showed that inhibition of caspase-3 prevents monocyte apoptosis and the cleavage of Bid, a protein cleaved during monocyte apoptosis. The role of the phosphorylation of caspase-3 in monocyte subsets has not been evaluated. Further studies should assess whether the phosphorylation of caspase-3 plays a role in the differences in apoptosis observed between monocyte subsets.
159
The CD14+CD16+ monocytes has been shown to expand during inflammatory diseases and could reach about 30% of the total monocytes (223, 265). These cells release low levels of the anti-inflammatory cytokine IL-10, suggesting that they may have a proinflammatory function (264, 266, 301). Interestingly, exposure to LPS and secretion of cytokines such as TNF-α and IL-1β have been shown to inhibit monocyte cell death; however, in the absence of any stimuli these cells undergo apoptosis (Fig. 1.5)
(230). The higher susceptibility of CD14+CD16+ monocytes to undergo apoptosis may control their accumulation in healthy individuals. We hypothesize that during infection
CD14+CD16+ cells should proliferate allowing an increase of their population for the production of proinflammatory cytokines to combat infections.
The expansion of the CD14+CD16+ cells have been correlated with increase in levels of proinflammatory cytokines such as TNF-α in the blood (266). CD14+CD16+ have been previously shown to have differences in the susceptibility to glucocorticoids
(303). The higher susceptibility of CD14+CD16+ suggests that treatments with anti- inflammatory drugs may deplete the non-classical monocytes as part of their mechanism of action in an anti-inflammatory therapy. The increase in cytokine production associated with the expansion of CD14+CD16+ has been analyzed in whole blood which complicates the assessment of the contribution of each monocyte (266). We purified
CD14+CD16- and CD14+CD16+ to assess of the contribution of each monocyte subset in the production of TNF-α after stimulation with LPS. We observed that CD14+CD16+ have 3 fold higher expression of TNF-α after stimulation with LPS, compared to
CD14+CD16- (Fig. 4.5). We also observed a higher TNF-α release in CD14+CD16+ by
160 clustering the IgG receptor, compared to CD14+CD16- monocytes. These results support the role of CD14+CD16+ as a proinflammatory subset.
The PKC family has been shown to have important roles in monocyte differentiation, immunological response and apoptosis (116, 186-188). We analyzed the expression of PKCs that could explain the heterogeneity observed in CD14+CD16- and
CD14+CD16+ cells. Our studies only revealed that CD14+CD16+ had 3 fold higher expression of PKCε (Fig. 4.7). Analysis of the roles of PKCε in monocytes suggested a potential role in their immunological response. PKCε knockout mice have impaired innate immunity and fail to clear bacterial infections (186). Recent studies have linked
PKCε to the TLR4 pathway, which is activated by stimulation with LPS (255). We show here that silencing of PKCε in CD14+CD16+ repressed TNF-α release to similar levels to
CD14+CD16- (Fig. 4.7). These results suggest that the high expression of PKCε in
CD14+CD16+ may sustain their proinflammatory phenotype. The role of PKCε in the
IgG clustering-mediated TNF-α production has not been elucidated. The signaling pathway that mediates the production of pro-inflammatory cytokines by monocyte clustering of IgG is not understood (Fig. 5.1). Further studies should assess the role of
PKCε in CD14+CD16+ in the FcγR pathway and in the release of additional proinflammatory cytokines (Fig. 5.1).
Altogether, this work presents a novel mechanism to control cell fate through the
PKCδ dependent phosphorylation of caspase-3. It also characterizes differences in apoptosis and immunological response in monocyte subsets. These results present a model for the regulation of cell death and survival pathways that involves a complex
161 regulatory network regulated by caspase-3 and the members of the PKC family. This knowledge could be used for the development of new strategies for the treatment of pathologies associated with dysregulation of apoptosis and proinflammatory diseases.
Fig. 5.1 Model
162
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