CD56+ T-cells in Relation to Cytomegalovirus in Healthy Subjects and Kidney Transplant Patients
Institute of Infection and Global Health
Department of Clinical Infection, Microbiology and
Immunology
Thesis submitted in accordance with the requirements of the
University of Liverpool for the degree of Doctor in Philosophy by
Mazen Mohammed Almehmadi
December 2014
- 1 - Abstract
Human T cells expressing CD56 are capable of tumour cell lysis following activation with interleukin-2 but their role in viral immunity has been less well studied. The work described in this thesis aimed to investigate CD56+ T-cells in relation to cytomegalovirus infection in healthy subjects and kidney transplant patients (KTPs). Proportions of CD56+ T cells were found to be highly significantly increased in healthy cytomegalovirus-seropositive (CMV+) compared to cytomegalovirus-seronegative (CMV-) subjects (8.38% ± 0.33 versus 3.29%± 0.33; P < 0.0001). In donor CMV-/recipient CMV- (D-/R-)- KTPs levels of CD56+ T cells were 1.9% ±0.35 versus 5.42% ±1.01 in D+/R- patients and 5.11% ±0.69 in R+ patients (P 0.0247 and < 0.0001 respectively). CD56+ T cells in both healthy CMV+ subjects and KTPs expressed markers of effector memory-
RA T-cells (TEMRA) while in healthy CMV- subjects and D-/R- KTPs the phenotype was predominantly that of naïve T-cells. Other surface markers, CD8, CD4, CD58, CD57, CD94 and NKG2C were expressed by a significantly higher proportion of CD56+ T-cells in healthy CMV+ than CMV- subjects. Functional studies showed levels of pro-inflammatory cytokines IFN-γ and TNF-α, as well as granzyme B and CD107a were significantly higher in CD56+ T-cells from CMV+ than CMV- subjects following stimulation with CMV antigens. This also resulted in higher levels of proliferation in CD56+ T-cells from CMV+ than CMV- subjects. Using class I HLA pentamers, it was found that CD56+ T-cells from CMV+ subjects contained similar proportions of antigen-specific CD8+ T cells to CD56- T cells in healthy donors of several different HLA types. A comprehensive gene expression profile by microarrays of CMV-stimulated sorted CD56+ T-cells was conducted which showed that expression of 106 genes involved in innate and adaptive immune responses was significantly changed, almost all being increased. Some of these changes were validated via RT-PCR and flow cytometry, the latter indicating higher ISG-15 protein expression in response to CMV stimulation in CMV+ than CMV- subjects. Overall, these differences may reflect the expansion and enhanced functional activity of CMV-specific CD56+ memory T cells. In view of the link between CD56 expression and T-cell cytotoxic function, this strongly implicates CD56+ T cells as being an important component of the cytotoxic T-cell response to CMV.
- 2 - Dedication
I dedicate this work to my mother (Salha) and my father (Janakiram et al.) who pray for me constantly and inspire me. A dedication also goes to my wife (Khadija) and my sons (Aser) and (Moath) who were always inspiring me with their smiles throughout my PhD. Also, special thanks to Taif University and the Saudi Cultural Buereau for their support throughout my scholarship years.
Acknowledgment
All Thanks to Allah for his countless blessing. I would like to express my sincere appreciation and thanks to my supervisor Dr. Steve Christmas, you have been a remarkable mentor. I would like to thank you for your assisting thoughout my PhD. I would also like to thank my second supervisor Dr Brian Flangan for his invaluable assistance to the study, and thanks to the Department of Women’s and Children’s Health of Alder Hey Children’s Hospital. Sincere gratitude to Sally Heyworth, Mr Abdul Hammad and all renal transplant recipients from Royal Liverpool and Broadgreen University Hospital Transplant Unit for their participation in this study. I am also indebted to Ms Caroline Broughton, Dr Shamsher Ahmed, Dr Suliman Alomar, Dr Waleed Mahallawi, Dr Ali Alejenef, Dr Debbie Howarth and Dr Raju Gautam for their support provided at all levels during this thesis. I am also grateful to all members of the staff of the Department of Clinical Infection, Microbiology and Immunology in IGH, Liverpool University for being cooperative during this course. A special thanks to Dr. Lucille Rainbow, Centre for Genomic Research, IIB for her vital contribution. Thanks also to Ayman Mubarak, Abdullah Aljurayyan, Mustafa Halawi, Wael Alturaiki, and Zaid Al-bayati for their invaluable friendship. Finally, a special thanks to my family. Words cannot express how grateful I am to my mother (Salha), father (Janakiram et al.), wife (Khadija), sons (Aser and Moath), brothers and sisters for all of the help and support provided. Your prayers for me were what sustained me thus far.
- 3 - List of Publications
Almehmadi M, Flanagan BF, Khan N, Alomar S, Christmas SE. Increased numbers and functional activity of CD56⁺ T cells in healthy cytomegalovirus positive subjects. Immunology. 2014 Jun;142(Varnum et al.):258-68.
Al Omar S, Flanagan BF, Almehmadi M, Christmas SE. The effects of IL-17 upon human natural killer cells. Cytokine. 2013 Apr;62(1) 123-30.
Manuscript under preparation
Almehmadi M, Hammad A, Heyworth S, Christmas SE. CD56+ T cells are increased in kidney transplant patients following cytomegalovirus infection.
- 4 - Table of Contents
Abstract 2
Dedication 3
Acknowledgements 3
List of publications 4
Table of contents 5
List of figures 13
List of tables 18
Abbreviations complete list 19
1. Introduction 22
1. Summary of the immune system 23
1.1 Antigen presentation 24
1.2 T-lymphocytes 25
1.2.1 CD4 + T-cells 25
1.2.2 CD8+ T-cells 28
1.2.3 γδ-T-cells 28
1.2.4 Natural Killer cells (NK cells) 29
1.3 B-Lymphocytes 29
1.4 CD56+ T cells (CD56+CD3+ cells) 29
1.4.1 History of CD56+ T-cells 29
1.4.2 CD56+ T-cell functional capability combines that of T-cells 34
and NK cells
1.4.3 CD56 neural adhesion molecule (NCAM) 34
- 5 - 1.4.4 CD56+ T cells functions in infections 34
1.5 Human Cytomegalovirus (HCMV) 40
1.5.1 Cytomegalovirus Transmission, Latency and Pathogenicity 42
1.5.1.1 Transmission 42
1.5.1.2 Latency 44
1.5.1.3 HCMV Pathogenicity 45
1.5.1.3.a Solid Organ Transplantation (SOT) 45
1.5.2 Immune response to HCMV 49
1.5.2.1 Innate Immunity 49
1.5.2.2 Natural Killer cells (NK) 49
1.5.2.2.a Missing self-hypothesis (Reduction in MHC-I expression) 52
1.5.2.2.b Self-cells expression of self-induced Ligands 52
1.5.2.3 Adaptive Immune response 53
1.5.2.3.1 Humoral Response to CMV 53
1.5.2.3.2 T-cell-mediated immune responses 54
1.5.2.3.2.a γδT-cells in response to CMV 54
1.5.2.3.2.b CD8+T-cells in response to CMV 54
1.5.2.3.2.c CD4+ T cells in response to CMV 57
1.5.3 CMV immune-evasion 59
1.5.3.1 CMV genes homologue to the host genes 60
1.5.3.2 CMV evading detection by Cytotoxic T cells 61
1.6 Kidney Transplant Rejection 61
1.6.1 T cells and B cells interface and generation of antibodies 63
1.6.2 T-cell role response in graft rejection and graft tolerance 64
1.6.2.1 The allorecognition process 65
- 6 - 1.6.2.1.a Direct allorecognition pathway 66
1.6.2.1.b Indirect allorecognition pathway 67
1.6.2.1.c Semi-direct allorecognition pathway 68
1.6.3 NK cell allorecognition 69
1.6.4 Development of Alloresponses 70
1.6.4.1 The alloresponse of CD4+ T-cells 71
1.6.4.2 The alloresponse of CD8+ T -cells 71
1.6.5 CD56+ T-cells immunological role to Graft 72
1.6.6 The role of CMV in allograft rejection 73
1.7 Aims and objectives 74
2. Materials and Methods 76
2.1 CMV IgG ELISA 76
2.1.1 CMV IgG ELISA assay Principle 76
2.1.2 Reagents and materials supplied and preparation 76
2.1.3 ELISA assay procedure and measurement 77
2.2 CMV lysate preparation 78
2.2.1 Fibroblast Cell Culture 78
2.2.2 CMV lysate preparation protocol 79
2.3 Peripheral blood 80
2.3.1 PBMC Donors 80
2.3.2 PBMC Isolation 81
2.3.3 Counting of PBMCs 82
2.4 Flow cytometry (FACSCalibur™ BD Biosciences) 82
2.4.1 Surface staining for phenotypic analysis 83
- 7 - 2.4.2 Functional analysis of CD56+ T-cells 85
2.4.2.1 Intracellular staining of CD56+ T-cells 85
2.4.2.2 Degranulation assay of CD56+ T-cells; detection by LAMP-1 87
(CD107a) expression
2.4.2.2.1 MHC negative targets 87
2.4.2.3.2 PBMC stimulation by K562 cells and CMV lysate 88
2.4.2.4 Proliferation assay of CD56+ T-cells 89
2.4.2.4.1 Carboxy-Fluorescein Succinimidyl Ester (CFSE) assay 89
2.4.2.4.2 PBMC CFSE labelling 90
2.4.2.4.3 PBMC stimulation by CMV lysate and SEB 90
2.4.2.5 CD56+ T-cell CMV specificity detection 92
2.4.2.5.1 HLA tissue type 93
2.4.2.5.2 CD56+ T-cells Pro5 Pentamer staining 96
2.4.2.6 Identifying CD56+ T-cell gene expression profile in response to 96
CMV
2.4.2.6.1 CD56+ T-cell sorting 97
2.4.2.6.2 RNA extraction protocol 98
2.4.2.6.3 DNase-treatment of RNA 99
2.4.2.6.4 One-Colour Microarray-Based Gene Expression 100
2.4.2.6.5 Data Analysis of microarrays using GeneSpring GX9.0 101
2.4.2.6.6 Validation of Microarrays results 102
2.4.2.6.7 Synthesis of cDNA (Reverse Transcription) 103
2.4.2.6.8 RT-PCR gene expression analysis 103
2.4.2.6.9 Protein expression analysis 105
2.4.2.6.9a CD38 surface staining of CD56+ T-cells 106
- 8 - 2.4.2.6.9b ISG-15 intracellular staining of CD56+ T-cells 106
3. Frequency and phenotype of CD56+ T-cells in CMV 110
seropositive and seronegative healthy subjects
3.1 CMV sero-status by applying IgG ELISA assay 110
3.2 Frequency of CD56+ T-cells in CMV seropositive and 112
seronegative healthy subjects
3.2.1 Introduction 112
3.2.2 Aims 112
3.2.3 Results 112
3.2.3.1 CD56+ T-cells relative numbers in healthy CMV seropositive 113
and seronegative subjects
3.2.3.2 Relative numbers of CD56-CD3+ cells in healthy CMV 116
seropositive and seronegative subjects
3.2.3.3 Relative numbers of NK cells (CD56+CD3-cells) in healthy 118
CMV seropositive and seronegative subjects
3.2.3.4 CD56+ T-cell levels compared in different age groups 119
3.2.3.5 CD56+ T-cells compared according to gender 120
3.3 Characterisation of the phenotype of CD56+ T-cells in CMV 121
seropositive and seronegative healthy subjects
3.3.1 Introduction 121
3.3.2 Aims of this study 127
3.3.2 Results 127
3.3.3.1 Expression of CD4 and CD8 by CD56+ T-cells 127
3.3.3.2 CD56+ T-cells; naive and memory markers 131
- 9 - 3.3.3.3 CD56+ T-cell expression of Lymphocyte function-associated 136
antigens
3.3.3.4 CD56+ T-cell expression of CD57, CD161, CD94/NKG2C and 138
CD16
3.3.3.5 CD56+ T-cells expression of KIRs CD158 a/h, CD158e and 141
CD158f
3.3.3.6 CD56+ T-cell expression of CD25 and CD69 143
3.3.3.7 CD56+ T-cell expression of TCRVα7.2 and γδ-TCR 145
3.3.3.8 CD56+ T-cell expression of exhaustion markers 148
3.3.4 Discussion 150
4. Functional responses in CD56+ T cells from CMV 160
seropositive and CMV seronegative healthy subjects
4.1 Cytokine responses of CD56+ T-cells 160
4.1.1 Introduction 160
4.1.2 Aims 161
4.1.3 Results 161
4.1.3.1 IFN-γ expression following stimulation with CMV lysate 162
4.1.3.2 TNF-α expression following stimulation with CMV lysate 165
4.1.3.3 IL-2 expression following stimulation with CMV lysate 167
4.1.3.4 Perforin content following stimulation with CMV lysate 170
4.1.3.5 Granzyme A expression following stimulation with CMV 172
lysate
4.1.3.6 Granzyme B expression following stimulation with CMV lysate 176
4.2 Degranulation assay of CD56+ T-cells; detection by LAMP-1 179
- 10 - (CD107a) expression
4.2.1 Aim 179
4.2.2 Cell surface exposure of CD107a in response to stimulation 179
4.3 Proliferative responses of CD56+ T cells to stimulation 185
4.3.1 Aims 185
4.3.2 Proliferative responses of CD56+ T cells to CMV antigens 185
4.4 Antigen specificity of CD8+CD56+ T cells in CMV+ subjects 188
4.4.1 Aims 188
4.4.2 CD56+ T-cells Pro5 pentamer staining 188
4.5 Discussion 190
5. Frequency and phenotype of CD56+ T-cells in CMV 198
seropositive and seronegative Renal Transplant Patients
5.1 Introduction 198
5.2 Aim 199
5.3 Results 199
5.3.1 Frequency of CD56+ T-cells in CMV seropositive and 199
seronegative renal transplant patients
5.3.2 CD56+ T-cell levels during CMV viraemia 200
5.3.3 Relative numbers of CD56-CD3+ cells in KTPs 202
5.3.4 Relative numbers of CD56+CD3- NK cells in KTPs 204
5.4 Characterisation of the phenotype of CD56+ T-cells in KTPs 204
5.4.1 Aims 205
5.4.2 Results 205
5.4.2.1 Expression of CD4 and CD8 by CD56+ T-cells in KTPs 205
- 11 - 5.4.2.2 CD56+ T-cells; naive and memory markers in KTPs 206
5.4.2.3 CD56+ T-cells; CD11a, NKG2C and CD161 in KTPs 208
5.4.2.4 CD56+ T-cells; CMV HLA Pentamer labelling 209
5.5 Discussion 211
6. Study to identify CD56+ T-cell gene expression profile in 219
response to CMV
6.1 Introduction 219
6.2 Aim 220
6.3 Results 220
6.3.1 One-Colour Microarray-Based Gene Expression 220
6.3.2 Validation of microarrays results 237
6.3.3 RT-PCR gene expression analysis 237
6.3.4 ISG-15 and CD38 protein expression analysis by FACs 243
6.3.4.1 ISG-15 protein expression by CD56+ T-cells 244
6.3.4.2 CD38 protein expression by CD56+ T-cells 246
6.4 Discussion 247
7. General conclusion 255
8. Biblography 261
Appendix 294
- 12 - List of Figures
1.1 Naïve CD4+ T-cells differentiate in response to different cytokines 27
1.2 CD56+ T-cells levels between healthy and malaria patients 36
1.3 CD56+ T-cell response to SEB. 37
1.4 Hepatic CD56+/- T-cell and NK cell cytokine response 39
1.5 HCMV 3-D structure 41
1.6 HCMV invasion of the host cell 43
1.7 MCMV infections in three different mice strains 51
1.8 CD4 + and CD8+ T cell responses to HCMV 56
1.9 Three allorecognition pathways 70
2.1 ELISA assay principle. 78
2.2 PBMC separation 82
2.3 Gating strategies 85
2.4 Proliferation gating strategies 91
2.5 Tissue typing gating strategy 94
2.6 Microarrays workflow flow-charts 101
3.1 CD56+ T-cell levels in representative CMV + and CMV- subjects 115
3.2 Percentages of CD56+ T-cells in CMV+ and CMV- subjects 116
3.3 Percentages of CD56- T-cells in CMV+ and CMV- subjects 117
3.4 Frequencies of CD56+CD3-NK cells in CMV+ and CMV- subjects 118
3.5 CD56+ T-cells compared by age group 119
3.6 CD56+ T-cell levels according to gender 120
3.7 Levels of CD56+ T-cells in males and females with the same CMV 121
- 13 - sero-status
3.8 Expression of CD4 and CD8 by CD56+ T-cells. 128
3.9 Expression of CD4 and CD8 by CD56+ T-cells in CMV+ and CMV- 129
subjects
3.10 Expression of CD4 vs CD8 by CD56+ T-cells 130
3.11 Memory and naive markers expressed by CD56+ T-cells 132
3.12 Comparison of memory and naive markers expressed by CD56+ T- 134
cells
3.13 LFA-1& 3 expression by CD56+ T-cells 136
3.14 LFA-1& 3 expression by CD56+ T-cells between CMV+/- 137
3.15 CD57, CD161, CD94/NKG2C and CD16 expression by CD56+ T- 138
cells
3.16 CD57, CD161, CD94/NKG2C and CD16 expression by CD56+ T- 140
cells in CMV+ and CMV- subjects
3.17 KIR expression by CD56+ T-cells. 142
3.18 KIR expression by CD56+ T-cells in CMV+ and CMV- subjects 143
3.19 CD69 and CD25 expression by CD56+T-cells 144
3.20 CD69 and CD25 expression by CD56+T-cells in CMV+ and CMV- 145
subjects
3.21 TCRVα7.2 and γδ-TCR expression by CD56+ T-cells. 146
3.22 TCRVα7.2 and γδ-TCR expression by CD56+ T-cells in CMV+ and 147
CMV- subjects
3.23 PD-1, CTLA-4 and KLRG-1 expression by CD56+ T-cells in CMV+ 148
and CMV- subjects
3.24 PD-1, CTLA-4 and KLRG-1 expression by CD56+ T-cells in CMV+ 149
- 14 - and CMV- subjects
3.25 Phenotypic differences between CD56+ T cells from CMV 156
seropositive and CMV seronegative healthy subjects
3.26 Phenotypic differences between CD56+ T cells from CMV 157
seropositive and CMV seronegative healthy subjects
4.1 SEB as a positive control for CD56+T-cells 162
4.2 Levels of IFN-γ expression 164
4.3 Levels of IFN-γ expression in CMV+ and CMV- subjects 165
4.4 Levels of TNF-α expression 166
4.5 Levels of TNF-α expression in CMV+ and CMV- subjects 167
4.6 Levels of IL-2 expression 168
4.7 Levels of IL-2 expression in CMV + and CMV- subjects 169
4.8 Levels of perforin 171
4.9 Levels of perforin in CMV+ and CMV- subjects 172
4.10 Levels of Granzyme A expression 174
4.11 Levels of Granzyme A expression in CMV + and CMV- subjects 175
4.12 Levels of Granzyme B expression 177
4.13 Levels of Granzyme B expression in CMV + and CMV- subjects 178
4.14 Levels of CD107a expression after PBMCs pulsed with K562 181
4.15 Levels of CD107a expression after PBMCs pulsed with K562 in 182
CMV + and CMV- subjects
4.16 Levels of CD107a expression after PBMCs stimulated with CMV. 183
4.17 Levels of CD107a expression after PBMCs stimulated with CMV in 184
CMV + and CMV- subjects.
4.18 Representative plots of CFSE labelled PBMCs 186
- 15 - 4.19 The percentage of proliferated cells labelled by CFSE 187
4.20 Representative plots of Pro5 CMV pentamer-labelling 189
4.21 Comparison of the percentage of pentamer labelled cells in CD56- 190
and CD56+ T cells.
5.1 Scatter plots comparing the percentage of CD56+ T-cells between 200
different KTP groups.
5.2 KTPs studied who had an episode of viraemia 201
5.3 D+R- patients studied who had an episode of viraemia 202
5.4 Scatter plots comparing the percentage of CD56- T-cells between 203
different KTP groups.
5.5 Frequency of CD56+CD3-NK cells in KTPs 204
5.6 The percentage of CD8+ and CD4+ CD56+ T-cells compared 206
between KTP groups
5.7 Memory and naive markers expressed by CD56+ T-cells in KTPs. 208
5.8 CD11a, NKG2C and CD161 expression on CD56+ T-cells from 209
KTPs
5.9 Representative density plots of HLA-pentamer-labelling 210
5.10 Comparision of HLA-pentamer-labelled cells 211
5.11 Levels of CD56+ T-cells in healthy subjects and KTPs 213
6.1 Microarray chip 221
6.2 Gene normalization 222
6.3 Significantly expressed genes using the filtration method. 223
6.4 Hierarchical clustering of the genes. 229
6.5 Genes normally regulated by IFNs and up-regulated in CD56+ T cells 232
following CMV stimulation.
- 16 - 6.6 Genes upregulated according to function by CD56+ T-cells due to an 233
anti-CMV response.
6.7 Genes upregulated according to immune and defence response by 234
CD56+ T-cells due to an anti-CMV response.
6.8 Genes upregulated according to stress by CD56+ T-cells due to an 236
anti-CMV response.
6.9 Ubiquitination cycle 238
6.10 Amplification plot showing curves by RT-PCR. 239
6.11 Comparison between means of gene expression in RT-PCR and 243
Microarrays
6.12 Levels of ISG-15 protein expression. 244
6.13 Levels of ISG-15 protein expression by density plot 245
6.14 CD38 expression 247
- 17 - List of Tables
1.1 Direct and indirect clinical effects caused by HCMV activation 48
in SOT
2.1 List of andtibodies against surface markers and their suitable 84
fluorochromes that have been used to phenotype CD56+ T-cells
in healthy subjects and patients
2.2 List of intracellular mAb that have been used in these 87
experiments. Different types of surface mAb were used with IC
mAb
2.3 Reagents that were used in proliferation assays of CD56+ T- 90
cells
2.4 Reagents used in pentamer labelling. 95
2.5 Reagents used in RT-PCR analysis of gene expression 104
2.6 RT-PCR thermal conditions 105
3.1 IgG ELISA detection 111
3.2 Phenotypic characteristics of memory T-cells 123
3.3 Markers applied to phenotypic analysis of CD56+ T-cells in 124
CMV seropositive and seronegative donors.
4.1 Cytokine and cytotoxic mediator detection. 192
6.1 Detailed information about genes whose expression 224
significantly changed
6.2 Significantly up- or down-regulated genes expressed by CD56+ 231
T-cells in response to CMV categorised according to fold
- 18 - change (FC).
6.3 RT-PCR results for IFN-γ 240
6.4 RT-PCR results for ISG-15 241
6.5 RT-PCR results of USP-18 242
Abbreviations complete list
AIDS Acquired immune deficiency syndrome
APC Antigen presenting cells
BCR B-cell receptor
BMT Bone marrow transplant
CCR Chemokine receptor
CD Cluster of differentiation
CFSE Carboxy-Fluorescein Succinimidyl Ester
CIK Cytokine-induced killer cells
CMV Cytomegalo virus
CNS Central nervous system
CPE Cytopathic effect
CTL Cytotoxic T-cells
CTLA CytotoxicT-lymphocyte associated protein
CXC Chemokine motif
D+R- Donor + recipient -
DC Dendritic cells
D-R- Donor - recipient -
- 19 - E1A Human adenovirus type 6 E1A 13S gene
EBV Epstein Barr virus
ER Endoplasmic reticulum
ERCC External RNA Control Consortium
FACS Flurescence Activated Cell Sorter/ing
FC Fold change
FceRI Fcε receptor 1
GVT Graft-versus tumour
HEV Hepatitis E virus
HHV Human herpesvirus
HIV Human immune-deficiency virus
HRP Horseradish peroxidase
IE Immediate early
IFN Interferon
IgE Immunoglobulin E
IL Interleukin
ISG Interferon stimulated genes iTreg Induced regulatory T-cells
KLRG killer lectin-like receptor G
KTP Kidney transplant patient
LAK Lymphokine-activated killer cells
LIR Leucocyte immunoglobulin like receptor
LPS Lipopolysaccharide
- 20 - P815 Cell line tumour cells
PAMPs Pathogen associated molecular patterns
PD Programmed cell death
TNF Tumour necrosis factor
TNFR tumor necrosis factor receptor super gene family
UL83 unique long 82
αβ T cell Alpha beta T-cells
γδ T cell Gamma delta T-cells
- 21 -
Chapter 1
Introduction
- 22 - 1. The immune system
The mammalian immune system is composed of two main components, the innate and the adaptive immune system, which work with each other to protect the body from harmful pathogens. The immune system has to adapt rapidly to detect and neutralize different types of pathogens. Furthermore, the immune system has methods to discriminate between self and non-self cells; these methods are essential to avoid targeting host cells. However in some abnormal states self-cells can be targeted by immune effector mechanisms, leading to the development of autoimmune diseases. If the activity of the immune system is modulated and reduced, this may lead to an invasion by pathogens, even those that were not able to form a real threat previously (Abbas and Lichtman 2009).
The innate immune system is relatively non-specific; it includes humoral barriers, secretory molecules, cells and anatomical barriers. Furthermore, anatomical barriers involve mechanical factors such as the epithelial surfaces that are normally impervious to pathogen infiltration (Kimbrell and Beutler 2001).
Chemical factors also are part of these barriers for example fatty acids in the sweat that prevent the growth and colonization of bacteria. Also, biological factors are part of these barriers and assist in preventing infection such as the normal flora that competes with pathogenic bacteria for nutrients inside the gastrointestinal tract of the body. Humoral factors also have essential roles in defending against infection when anatomical barriers are penetrated by the pathogens. Humoral components are effective in initiating inflammation that is characterised by recruitment of phagocytic cells to the site of pathogen invasion.
This system involves for example complement components, the coagulation system, and lysozyme. Cellular components are important in the inflammatory
- 23 - response to pathogens. Cells involved include neutrophils, macrophages, NK cells and eosinophils. Both macrophages and neutrophils are essential parts of the phagocytic system that engulf the pathogens and neutralize them (Abbas and
Lichtman 2009).
The adaptive immune system is the more specific type of host immunity. This system is antigen specific and requires antigen presentation. Lymphocytes are a crucial part of the adaptive immune system, of which there are several types. The major subsets are B-cells that mediate humoral immune responses and T-cells that regulate cell mediated responses, and both of these cell types are derived from the bone marrow as a result of haematopoietic stem cell division (Kimbrell and
Beutler 2001).
1.1 Antigen presentation
Human leucocyte antigens (HLA) that are also named MHC have an essential role in adaptive immune system. HLA is expressed on cell surfaces and their main function is to facilitate communication of the body cells with leucocytes. HLA is divided into two types, HLA-I and HLA-II (Penn 2005).
HLA-I is expressed on all nucleated body cells, and this class’s main function is to express peptides to CD8+ T-cells. This peptide is either self-antigen expressed by healthy cells or foreign antigen that is expressed by abnormal cells under intracellular stress. Endogenous proteins are digested into peptides in the cytosol via proteolytic proteasomes, then transported by transporter proteins into the ER
- 24 - where peptides are bound to newly synthesized HLA-I due to affinity and then transported to the cell surface (Van Hateren, 2010).
HLA-II, however, is expressed mainly by APC. The latter cells are macrophages, dendritic cells (DC) and B-cells. Extracellular pathogens are engulfed by a process called phagocytosis or endocytosis by APC, then extracellular pathogens are digested by proteases in the endosome. In the ER α, β and invariant chains are synthesized and assembled, then transported via the Golgi body to endosomes and here invariant chain is digested. Moreover, peptides from extracellular pathogens are loaded into HLA-II molecule and transported into the cell surface for CD4+ T- cell recognition (Penn 2005).
1.2 T-lymphocytes
T-lymphocytes have several different subsets. These subsets share the expression of TCR or T-cell receptor. TCR activates in response to the expression of antigen on HLA-I or II by antigen presenting cells (APC).
1.2.1 CD4 + T-cells
Activated CD4+ T-cells were identified by Mosmann and Coffman to have 2 different subsets Th1 and Th2 in 1986. Moreover, currently about four different types have been described, Th1, Th2, Th17 and iTreg cells (Zhu and Paul 2008).
T-helper cells have essential functions in modulating the adaptive immune system, these roles are exerted by their ability to produce cytokines and
- 25 - chemokines to activate and recruit other types of cells (figure 1.1) (Zhu and Paul
2008).
A role for Th1 cells is in resistance to intracellular pathogens; the cytokines mainly produced by Th1 cells are IFN-γ and IL-2 and lymphotoxin-α (LTα). In addition, LTα is considered a marker for multiple sclerosis disease progression
(Selmaj et al. 1991). IFN-γ released by Th1 is essential for increasing macrophage antimicrobial activity (Suzuki et al. 1988).
The other main type of T-helper cells are Th2 cells, which are important in mediating host defence against extracellular pathogens. Th2 cells are potent cytokine producers of amphiregulin, IL-4, IL-5, IL-9, IL-10, IL-13 and IL-25
(Zhu and Paul 2008). The release of IL-4 by Th2 cells is the major mediator of
IgE class switching in B-cells. Furthermore, IgE targets FceRI receptors on basophils and mast cells, when cross linked with a multivalent ligand leading to the release of active mediators like histamine and serotonin and cytokines like IL-
4, IL-13 and TNF-α (Zhu and Paul 2008).
- 26 -
Figure 1.1 Naïve CD4+ T-cells differentiate. In response to different cytokines
aanaive CD4+T-cells differentate into specific types of CD4+ T-cells (Zhu and
Paul 2008).
Th17 cells are one of the helper cells subsets; this subtype releases IL-17A and F,
IL-21 and IL-22. Additionally, IL-17A and F target the IL-17RA receptor, and they have the ability to recruit and activate neutrophils in response to extracellular pathogens. IL-21 plays a role in stimulating Th17 differentiation, similar to the role of IFN-γ to Th1 and IL-4 to Th2 (Korn et al. 2007, Nurieva et al. 2007, Zhou et al. 2007).
T regulatory cells or classically known as suppressor T cells, are a subset of T- cells that preserve tolerance to self-antigens and regulate immune response homeostasis (Zhu and Paul 2008, Sakaguchi 2004, Joffre et al. 2008). T-regs are
CD4+CD25+ cells and develop and emigrate from the thymus. Increase in percentage of T-reg cells was found to help prevent autoimmune diseases and
- 27 - allograft rejection (Joffre et al. 2008). Several studies have investigated the mechanism of T-reg function and they are thought to mediate their function through direct contact with target cells (Shevach 2006). Moreover, T-regs are able to release the cytokines TGF-β, IL-10 and IL-35 (Zhu and Paul 2008).
1.2.2 CD8+ T-cells
CD8+ T-cells are a subset of lymphocytes also known as T-killer cells, cytolytic
T-cells, Killer T-cells or Cytotoxic T lymphocytes (CTL). This subset acquired these several names due to their function in killing cancer cells; pathogen infected cells and impaired cells due to any effect. CTL have the ability to recognise peptides presented on the extracellular part of MHC-I molecules. These peptides are generated in the intracellular region of the cells via the degradation of cytosolic proteins in the proteasome. Consequently, CD8+ T-cells eliminate cells that express their specific antigen peptides presented on MHC-I.
1.2.3 γδ-T-cells
This subset of T-cells expresses TCR that is distinct from other types of T-cells.
γδ-cells TCR is made of one γ chain and one δ chain. Additionally, the frequency of γδ-cells is much less than αβT-cells, but this type of T-cells is found more in gut mucosal and is more abundant in intraepithelial lymphocytes.
- 28 - 1.2.4 Natural Killer cells (NK cells)
NK cells have a critical role in the rapid immune response of the innate immune system. NK cells, a type of lymphocyte, are cytotoxic against foreign pathogen- infected cells that have down regulated class I MHC. NK cells express inhibitory receptors that detect self-class I MHC and prevent NK cells from initiating the killing process against self-cells that are not infected. The NK cell activation process is regulated by activating and inhibitory receptors and, also, by MHC-I associated proteins on the target cells (Iversen et al. 2005); (Lanier 2005).
1.3 B-cells (B Lymphocytes)
The other type of effector immune response is the humoral immune response; this type is modulated by B-cells. As well as being responsible for antibody production, B-cells also perform an essential role in antigen presentation. B-cells have a B-cell receptor (BCR) which is a membrane-bound immunoglobulin that also helps to distinguish B-cells from T-cells. Once B-cells become activated they differentiate into plasma cells or memory B-cells.
1.4 CD56+ T cells (CD56+CD3+ cells)
1.4.1 History
Several efforts have been attempted in recent decades for the development of effective immunotherapy against tumours by using cytotoxic antitumor cells (Lu and Negrin 1994). LAK cells (Lymphokine-activated killer cells) are described as
- 29 - the primary prototype of CIK cells (Cytokine-induced killer cells) (Linn and Hui
2010). Rosenberg and colleagues first generated LAK cells in 1980. LAK cells are described as lymphoid cells that highly proliferate several folds after being induced by IL-2 that is essential for cell growth, proliferation and differentiation.
LAK cells show cytotoxic capabilities different from NK cells and CTL
(cytotoxic T lymphocytes) (Grimm et al. 1983), still they share some properties with NK cells such as the surface marker CD56 and the lack of MHC restriction
(Ramsdell and Golub 1987). Leu19 is the previous name for CD56 (Linn and Hui
2010). Moreover, LAK cells are not MHC restricted and they are capable of specifically lysing cancerous cells, even those resistant to killing by NK cells.
Also, LAK cells do not target self-cells and are specific for tumour cells (Fagan and Eddleston 1987). These cells are cytotoxic against autologous and allogeneic tumour cells types (Grimm et al. 1983). The molecularcloning of IL-2 in 1983 by
Taniguchi and colleagues subsequently allowedintravenous administration of IL-2 and LAK cells in bulk amounts reducing the number and size of secondary tumours in animal tumour models and preliminary human studies (Fagan and
Eddleston 1987), though IL-2 administration can lead to toxicities such as hypotension and vascular leakage (Jennings et al. 2013). LAK cells derived from peripheral blood cells of tumour patients were introduced with IL-2 by intravenous administration into tumour patients with hepatic or pulmonary metastases and reduced the size of the tumour more than 50% in less than three weeks. Each tumour patient received LAK cells obtained from his/ her peripheral blood cells (Rosenberg et al. 1985).
Moreover, NK/LAK cell immunotherapy illustrates potent immune responses toward MHC-I negative tumours such as neuroblastoma and by the use of MD-
- 30 - DCs (monocyte–derived dendritic cells) removes the requirement for the use of potentially toxic IL-2 (Valteau-Couanet et al. 2002).
Even though LAK cells have shown some benefits toward tumour treatment, the use of these cells was held back by shortage of enough cells for cancer immunotherapy (Rosenberg et al. 1986). Furthermore, LAK cells are CD3- and hence are unable to generate an antigen-specific response (Ramsdell and Golub
1987).
In 1986, tumour-infiltrating lymphocytes (TILs) show about 100 times more effective therapeutic outcomes than LAK cells in metastatic tumours (Rosenberg et al. 1986). TILs are lymphocytes that migrated from the blood stream toward the tumour site then infiltrated the tumour; their major effector cell phenotype is
MHC-restricted CD3+CD56-CD8+ (Lu and Negrin 1994); (Yoo et al. 1990).
These cells are efficient in treatment of mice with tumours resistant toward
LAK/IL-2 adoptive immunotherapy (Rosenberg et al. 1986). Preparation of these cells is different from LAK cells. They are harvested from tumours and then incubated with IL-2 in culture medium (Rosenberg et al. 1986). However, as with
LAK cells these cells are also held back by the complexity of generating enough cells to use them in cancer immunotherapy. Moreover, functional modifications occur during the harvesting process from the human tissue (Lu and Negrin 1994).
TILs have been reported to have a beneficial outcome in extending survival in cancers such as skin cancer (Haanen et al. 2006) and ovarian cancer (Ma and Gu
1991) (Sharma et al. 2007). LAK cells are limited by the inability to generate high numbers that is further hampered after applying chemotherapy to the cancer patients. On other hand, TILs are more effective than LAK cells, although those
TILs cells harvested from metastatic and advanced tumours were reported to have
- 31 - less cytotoxicity and proliferation ability than those harvested from primary tumours (Yoo et al. 1990); (Lu and Negrin 1994).
In 1991, a new study reported an advanced procedure to generate high numbers of effector cytotoxic cells against human B lymphoma cell lines. This study was conducted in SCID (severe combined immunodeficiency) mice that lack both T and B-lymphocytes. Additionally, human lymphoma cells were injected into these mice to grow and allow examination of the interaction between human lymphocytes and these lymphoma tissues (Schmidt-Wolf et al. 1991). CIK cells
(cytokine induced killer cells) were generated through ex-vivo culturing of peripheral blood mononuclear cells (PBMCs) with the periodic introduction of
IFN-γ, anti-CD3(OKT-3) molecular antibody and interleukin 2 (IL-2) over a 21-
28 day period (Schmidt-Wolf et al. 1991). The purpose of using these specific cytokines as well as anti-CD3 mAb was to induce T cells to proliferate several folds. Moreover, IFN-γ was used to induce the cytotoxic capability of these cells by the induction of IL-2 receptors on these cells before adding IL-2. Additionally,
IFN-γ induces monocytes to produce IL-12 and up-regulate LFA-3 (CD58) which are essential for CD56+CD3+ cell expansion (Lopez et al. 2001). After that, adding IL-2 to these cells will result in activation of more cells even those that have not been activated by using IL-2 alone in LAK cell cultures. Also, integrating IL-2 genes into CIK cells has boosted IL-2 secretion when cultured with DC, that in turn improved CIK antitumour activity towards a pancreatic cancer cell line (Nagaraj et al. 2004). Moreover, the addition of IL-1α can enhance the cytotoxicity of the cells (Schmidt-Wolf et al. 1991). As a result of using these cytokines to generate these cells they are then named Cytokine-Induced killer cells or CD56+ T-cells. In this study CIK cells showed greater cytotoxicity
- 32 - against B-lymphoma cells than LAK cells, and only slight cytotoxicity against mouse bone marrow stem cells and human haematopoietic engraftment (Schmidt-
Wolf et al. 1991). Moreover, the use of CIK cells against tumours does not require the administration of IL-2 into the host (Linn and Hui 2010). The higher capability of CIK cells than LAK cells can be due to the higher number of generated cells (Lu and Negrin 1994). CIK cell can also be generated from cord blood (Introna et al. 2006).
CIK cells are phenotypically different from LAK cells and are CD3+CD56+; furthermore the levels of these cells in normal human peripheral blood are between 1% and 5%. However, during ex vivo expansion they are generated from
CD3+CD56- cells and not generated from CD3-CD56+ cells in an ex vivo environment. CD56 acquisition is correlated with increased cytotoxicity of these cells. CD56 was acquired by lymphocytes in an ex vivo environment that already expressed the phenotype CD4+CD8+, CD4-CD8- and CD4-CD8+ after the cells were cultured with IL-2 and anti-CD3 mAb. However, CD4+CD8- cells did not acquire CD56 under these conditions (Lu and Negrin 1994). Among the expanded population of CIK cells the majority of the cells are CD8+ cells, moreover,
CD3+CD56+ cells comprise almost 90% whereas the rest are CD3+CD56- cells and CD3-CD56+ cells (Sangiolo 2011); (Niam et al. 2011); (Hontscha et al.
2011).
CIK cells share some properties with NK cells such as the bulky granular morphology, killing targets in non-MHC restricted manner and the physical expression of CD56, and lectin-like glycoprotein NKG2A, CD94, NKG2C (Saikh et al. 2003) and NKG2D. However, CIK cells lack the NK cell markers NKp46,
NKp44 and have only low expression of NKp30 (Pievani et al. 2011).
- 33 - 1.4.2 CD56+ T-cell functional capability combines that of T-cells and NK cells
On the surface of CD56+ T-cells there are high levels of expression of TCR/CD3 receptors. Studies involving stimulation of TCR/CD3 receptor using P815 tumour cells loaded with anti-CD3 monoclonal antibody led to the production of the cytokines IFN-γ and TNF-α. In addition, CD56+ T-cells retained antigen specificity when PBMCs from CMV seropositive donors were cultured for 21 days after which the proliferated cells showed potent specificity when stained using CMV peptide loaded HLA-A*0201 tetramers. Furthermore, activated
CD56+ T-cells that retained their specificity expressed the NK cell activation markers NKG2D, NKp30 and DNAM-1 that indicated the capacity of these cells to exert TCR specificity and NK cell killing capacity (Pievani et al. 2011).
1.4.3 CD56 / neural adhesion molecule (NCAM)
CD56 is a membrane glycoprotein and generally known to be a marker of NK cells and central nervous system (CNS) cells, however many studies showed the expression of this marker on T-cells and other types of cell. Moreover, this cluster of differentiation is not well studied. Some studies suggested a possible role in establishing effector to target cell adhesion and conjugation functions (Vergelli et al. 1996). CD56 is part of the Ig super-family and is a 140kDA isoform, which has six sites for N-linked glycosylation modified by polysialic acid (Pittet et al. 2000).
CD8+ T-cell cytolytic effector activity has been found to correlate with increase in CD56 surface expression. Moreover, CD8+ T-cells with high levels of CD56
- 34 - expression were found to have more IFN-γ and granzyme-B, also, direct cytolysis by CD8+ T-cells is conducted through CD8+CD56+ cells only (Pittet et al. 2000).
As well as being expressed on neuronal tumours, CD56 has been found on several other tumours such as myeloma and myeloid leukaemia and anti-CD56 has been developed for purposes of antibody based applied immunotherapy of cancer
(Jensen and Berthold 2007).
1.4.4 CD56+ T-cell functions in infectious disease
CD56+ T-cells have been studied for many years in relation to tumours; however, they have not been studied thoroughly with regards to microbial infection. Some studies have shown that CD56+T-cells increase during infection. Parasitaemia in malaria induced CD56+ T-cells to expand in patients more than healthy subjects
(Watanabe et al. 2003) (Figure 1.2).
- 35 -
Figure 1.2 CD56+ T-cells levels between healthy and malaria patients. Two
colour scatter plots displaying the difference between the levels of CD56+ T-cells
in healthy and two malaria patients infected with two different parasites. CD56+ T
cells have expanded more in the patient infected with P. vivax than P. falciparum
(Watanabe et al. 2003).
These cells were abundant in the liver of those patients infected with malaria
(Watanabe et al. 2003). During influenza viral infection considerable numbers of
CD8+ T-cells develop CD56 which was noted in mice (Assarsson et al. 2000);
(Kambayashi et al. 2000), moreover, CD8+ T-cells develop NK cell associated molecules that may be involved in regulation of MHC-I-dependent T-cell effector functions (Kambayashi et al. 2000); (Slifka et al. 2000).
CD4+CD56+ T-cells expand in response to Hepatitis B virus antigens restricted to
MHC-II; also CD4+CD56+ T cells migrated to the nasopharynx in response to herpes simplex virus infection, consequently, these cells correlate with the adaptive immune response mechanism toward infection (Barnaba et al. 1994);
- 36 - (Taddesse-Heath et al. 2003). Moreover, two days incubation with the superantigen staphylococcal enterotoxin B (SEB) of mononuclear cells purified from NK cells resulted in 3 fold expansion of CD56+ T-cells (Figure 1.3) (Saikh et al. 2003).
Figure 1.3 CD56+ T-cell responses to SEB. Two color scatter plots showing
CD56+ T cells after stimulation by SEB for two days at 37°C/ 5% CO2. The mononuclear cell population was depleted of NK cells, and a clear expansion of
CD56+CD3+ cells from 5.8% (A) to 16% (B) is seen (Saikh et al. 2003).
CD4+CD56+ T-cells are abundant in the liver of mice, they produce IL-4 and furthermore can support Th2 cell development (Emoto et al. 1999). These cells shifted their IL-4 production to IFN-γ after infection by M. bovis BCG (Emoto et al. 1999). Increase of IFN-γ expression by CD56+ T-cells was also detected when
PBMCs were cultured and stimulated with an EBV-transformed lymphoblastoid cell line (Fu et al. 2012). CD56+ T-cells also increased in HIV-positive Chinese offspring born from HIV infected mothers, moreover, the phenotype characteristics of the cells in HIV positive children showed down-regulation of
- 37 - other NK cell markers: CD16, NKp80, NKp44, NKp30 and CD244 (Paiardini et al. 2005). Moreover, up-regulation of NKG2D, KIR3DL1and CD45RO was also observed on CD56+ T cells (Fu et al. 2012). CD56+ T-cells have also been found to enhance DC function; this was noticed when DCs from HIV positive children were incubated with CD56+ T-cells which induced the expression of CD83, CD86 and CD80 on DCs more than co-culturing DCs obtained from HIV negative controls with CD56+ T-cells. Clearly, the previous finding show that CD56+ T- cells are capable of inducing the maturation of DCs in an HIV positive environment, which also has indirect effects on other immune cells regulated by
DCs (Fu et al. 2012).
In normal adult liver, proportions of CD56+ T-cells were found to range from
13.7% to 27.2% of total lymphocytes stained by anti-CD3 and anti-CD56 conjugated antibodies. Moreover, after purifying these cells into three groups:
CD56+CD3+, CD56+CD3- and CD56-CD3+ cells; only CD56+CD3- cells were capable of direct lysis of K562. Furthermore, inducing MNC obtained from normal adult liver by using PMA/ionomycin showed that both CD3+CD56- and
CD3+CD56+ cells produced IFN-γ, TNF-α, IL-2 and IL-4. However,
CD56+CD3- cells produced only IFN-γ and TNF-α (Figure 1.4) (Doherty et al.
1999).
- 38 -
Figure 1.4 Hepatic CD56+/- T-cell and NK cell cytokine response. MNC
obtained from the liver and stimulated with PMA/ionomycin, MNC were
stained by three conjugated antibodies CD3, CD56 and the third colour for
detection of intracellular cytokine; the latter is displayed in the histograms. The
shaded histograms represent the un-stimulated negative control. NT cells are
CD56+ T cells and the histograms show that these cells produced Th1 cytokines
IFN-γ, TNF-α, IL-2 and the Th2 cytokine IL-4 but no IL-5 (Doherty et al.
1999).
In contrast to the previous findings that showed increase percentages of CD56+ T- cells in response to infection, CD56+ T-cells numbers have been found to decrease in hepatitis E virus (HEV) infected patients compared to healthy subjects. However, even though their numbers are reduced, the cells express
- 39 - activation marker CD69 more strongly than healthy controls; this could be explained as these cells are migrating toward the liver which is the site of inflammation and also the clearance site of apoptotic cells (Srivastava et al. 2008).
To clarify this, HEV seropositive patients were treated by IFN-α and it was found that NK cells in these patients started to express both CD69 and annexin-V, the latter being an apoptosis marker. Therefore, both NK and CD56+ T-cells migrated to the site of inflammation and their activation was due to activation-induced death of both NK and CD56+ T-cells which led to reduction in their number
(Srivastava et al. 2008). Moreover, in HCV seropositive patients CD56+ T-cell levels were found to be non-significantly lower in patients than healthy subjects which was apparently due to continuous activation and apoptosis leading to a net reduction in total number of CD56+ T-cells (Lucas et al. 2003).
1.5 Human Cytomegalovirus (HCMV)
HCMV is the largest virus among human herpes-viruses family, which has a genome size of 235 kb encoding 165 genes. HCMV is a β-human herpes virus type 5, the envelope of this virus consists of a lipid bilayer that include several significant viral glycoproteins. Moreover, the envelope surrounds a virion composed of a linear double stranded DNA inside an icosahedral nucleo-capsid that surrounded by a proteinaceous matrix named the tegument (Figure 1.5)
(Crough and Khanna 2009); (Chen et al. 1999); (Mocarski 2007).
- 40 -
Figure 1.5 HCMV 3-D structures. This figure illustrating different components
of the virus (Crough and Khanna 2009).
The virion diameter of HCMV ranges between 200–300 nm. The tegument section holds several large proteins, minor proteins and RNA of cellular and viral source (Crough and Khanna 2009); (Mocarski 2007); (Varnum et al. 2004). The large proteins such as that encoded by the UL48 gene and UL99-encoded pp28, also, the phospho-protein 65 (pp65) also named unique long 83 (UL83) are present abundantly and form the lower matrix. Moreover, the virion trans- activator pp71 also known as unique long 82 (UL83) is present plentifully and forms the upper matrix protein. Furthermore, the large matrix phospho-protein, which is the herpes-virus, core virion maturation protein pp150 is encoded by the
UL32 gene (Crough and Khanna 2009); (Varnum et al. 2004).
The proteins of the tegument have several functions that assist in virus activities.
These functions generally consist of assembly and disassembly of virions during virus invasion of the target cell. Also, other functions of the tegument proteins are to modify the target cell response and to control interaction of virus infection and
- 41 - invasion (Crough and Khanna 2009); (Mocarski 2007). The target cell or the host cell provides a lipid bi-layer envelope enclosing the tegument, which is derived from endoplasmic reticulum and Golgi body. Moreover, the tegument has more than 20 glycoproteins encoded by the virus. These glycoproteins gB, gH, gL, gM, gN and gO are for cell attachment and penetration (Crough and Khanna 2009,
Varnum et al. 2004). After cell invasion by the virus, synthesis of viral proteins is commenced in three phases, first is the immediate early (IE) which begins from zero time up to 2 hours after successful invasion, second is the delayed early between 2 hours after the invasion to 24 hours and the late phase at 24 hours after successful infection (Figure 1.6) (Crough and Khanna 2009); (Stinski 1978).
1.5.1 Cytomegalovirus Transmission, Latency and Pathogenicity
1.5.1.1 Transmission
Human cytomegalovirus sero-positivity prevalence ranges between 30%-90% in developed countries, this also increases in developing countries and old people
(Crough and Khanna 2009); (Staras et al. 2006). In healthy individuals HCMV mostly remains non-pathogenic in the latent state. However, in immune- compromised patients, the fetus or in neonates HCMV can be reactivated and give rise to pathological sequelae. In addition, HCMV transmission occurs by several methods from infected individuals. HCMV can be transmitted by direct contact between body secretions or fluids such as saliva and blood, also, through organ transplantation. Moreover, it can be transmitted from the mother to baby by breast-feeding or during pregnancy via placental transfer (Crough and Khanna
2009, Sia and Patel 2000).
- 42 -
Figure 1.6 HCMV invasion of the host cell. It commences via the endocytic pathway or by direct fusion; attachment to the host cell occurs by viral glycoproteins that bind to the host surface receptors. After that, nucleocapsids are released into the host cytoplasm as a result of the fusion between cellular membrane and virus envelope. Then, nucleocapsids migrate to the nucleus where virus DNA is released and the expression of virus genes starts, followed by viral replication, maturation and viral synthesis, and encapsulation of the viral DNA to form the capsids. Moreover, a second envelopment occurs in the
ER-Golgi intermediate compartment and another two stages of envelopment before release of new viruses by exocytosis from the plasma membrane
(Crough and Khanna 2009).
- 43 - 1.5.1.2 HCMV latency and reactivation
HCMV latency mechanisms have been studied and examined in infected host cells via both in vitro and ex vivo experimental models. Some investigations suggested that HCMV proceeds to latency in the absence of active virus replication (Cheung et al. 2006); (Goodrum et al. 2002). UL138 viral sequences were the first viral sequences discovered to be very significant for HCMV latency, this was supported through studies on CD14+ monocytes and CD34+ progenitor cells in HCMV sero-positive individuals (Crough and Khanna 2009); (Goodrum et al. 2007). The mechanisms of latency may start upon virus access to the body and proceed to the latent phase without viral gene expression. Furthermore, the virus expresses genes necessary for effective latency upon entry and may not express genes required for infection. Also, disturbance of active HCMV infection after virus entry leads to advancement into the latency phase (Crough and Khanna
2009); (Cheung et al. 2006); (Goodrum et al. 2002); (Kondo et al. 1994); (Reeves et al. 2005). Additionally, granulocyte-macrophage progenitor (GMP) infection with HCMV induces virus latency-associated transcripts encoded in the MIE promoter region of the genome of HCMV, and also detected in seropositive healthy individuals, though this type of infection was not successful for in vitro infection (Crough and Khanna 2009); (Kondo et al. 1994); (Kondo and Mocarski
1995); (Kondo et al. 1996); (Landini et al. 2000); (White et al. 2000). Transcripts of the UL111.5A region that encodes a variant IL-10 homologue were detected in latently infected GMP cells, in naturally infected bone marrow cells and in granulocyte colony stimulating factor (G-CSF) mobilized peripheral blood samples collected from allograft donors (Crough and Khanna 2009); (Jenkins et al. 2004). However, the mechanism of change from the latency phase to
- 44 - reactivation is still unclear, nevertheless, HCMV reactivates during situations of stress, infection, immunosuppression and inflammation (Crough and Khanna
2009); (Kutza et al. 1998); (Mutimer et al. 1997); (Prosch et al. 2000). Moreover, several other factors can facilitate a change of HCMV status from latency towards the reactivation phase such as TNF-α, catecholamines, norepinephrine and epinephrine. Moreover, TNF-α has a significant role in the reactivation process of
HCMV, as the HCMV IE enhancer/promoter region is found to be induced by
TNF-α. Correspondingly, TNF-α binds to TNF receptor on CMV-infected cells, leading to protein kinase C and NF-kB activation (Crough and Khanna 2009);
(Prosch et al. 1995); (Stein et al. 1993). Cyclic AMP (cAMP) induces the stimulation of IE enhancer/promoter leading to HCMV reactivation where cAMP concentration rises due to stress, epinephrine, norepinephrine and pro- inflammatory prostaglandins (Crough and Khanna 2009); (Varnum et al. 2004);
(Kline et al. 1998).
1.5.1.3 HCMV Pathogenicity
CMV is a potent immunogen that activates the immune system rapidly. HCMV can cause serious health problems in individuals with non-developed immune systems or who are immunocompromised. However, CMV hardly causes any health problems in healthy individuals.
1.5.1.3.a Solid Organ Transplantation (SOT)
In solid-organ transplantation, HCMV activation in immunosuppressed recipients can lead to undesired outcomes such as viremia from HCMV and other opportunistic pathogens, damage to the graft leading to rejection, and mortality
- 45 - (Table 1.1). HCMV infection is a major risk factor for failure in graft adaptation in recipients resulting from direct and indirect clinical effects of HCMV activation leading to graft rejection. Additionally, preventive methods were found to reduce the danger of HCMV infection in SOT patients, however, late-onset HCMV viraemia can lead to graft rejection and mortality in liver transplantation (Eid and
Razonable 2010); (Helantera et al. 2006). The most significant risk group during transplantation is when transplanting an organ from an HCMV sero-positive donor to a recipient who is HCMV sero-negative (D+/R-) (Crough and Khanna
2009); (Limaye et al. 2002); (Rubin 2007). Generally, organ recipients acquire
HCMV infection as a primary type when the virus is transmitted through an infected organ from an HCMV sero-positive donor. This also occurs when the donor has active HCMV infection, or when latent HCMV reactivates in the recipient due to the immune-suppressed status (Eid and Razonable 2010).
Morbidity in SOT due to HCMV infection is high and several prevention methods have been introduced to reduce HCMV infection in transplanted organ recipients by using anti-viral prophylaxis and pre-emptive treatment (Eid and Razonable
2010); (Razonable and Paya 2003). In HCMV sero-negative recipients, nonexistence of previous immunity to HCMV assists the virus to replicate, when receiving an organ from HCMV sero-positive donor. This is also facilitated by the immune-suppressed status of the recipient (Eid and Razonable 2010). The high- risk group (D+/R-) continues to develop HCMV infection (38%), usually after completion of the anti-viral course, and this type of HCMV infection is called late-onset (Eid and Razonable 2010); (Arthurs et al. 2008); (Arthurs et al. 2007);
(Kijpittayarit-Arthurs et al. 2007), and several studies state that this type of late- onset infection is the reason behind a high proportion of cases of graft rejection
- 46 - and mortality (Eid and Razonable 2010); (Limaye et al. 2006, Arthurs et al. 2008).
This is also supported by the absence of HCMV-specific T-cell-immunity to counteract virus replication (Eid and Razonable 2010); (Kumar et al. 2009). Also, deficient T-cell mediated immunity against HCMV is a consequence of introducing immune-suppressive medication following the transplantation process that leads to depletion of lymphocytes for a period of time which HCMV uses to begin its replication; examples of these medications are Muromonab-CD3 (OKT-
3) and Anti-thymocyte Globulin (Eid and Razonable 2010); (Razonable and Paya
2003). The use of this immune-suppressive medication is essential in order to prevent immediate acute graft rejection, however HCMV activation is a major drawback of these medications (Razonable et al. 2001). Moreover, CMV was found to play a critical part in promoting acute graft rejection, immune- modulation and up-regulation of alloantigen, enhancing the rejection process.
Moreover, acute rejection of the graft promotes CMV activation; this is due to administration of anti-rejection treatment therefore to prevent CMV from re- activating an anti-CMV prophylaxis should be administered as well (Eid and
Razonable 2010); (Peleg et al. 2007a).
SOT recipients who show CMV activation can suffer major symptomatic disease or can show asymptomatic features and these depend on the degree of immune- suppression of the recipient where strong immune-suppression results in less capability to inhibit CMV activation (Eid and Razonable 2010). There are direct and indirect clinical impacts of CMV activation in SOT (Table 1.1).
- 47 - Table 1.1 Direct and indirect clinical effects caused by HCMV activation in
SOT (Eid and Razonable 2010).
Type of morbidity caused due to HCMV infection
Direct effects Indirect effects CMV disorder Acute rejection Fever, myalgia, Chronic rejection arthralgia Bronchiolitis obliterans myelo-suppression Transplant vasculopathy Tissue-invasive CMV disease Tubulointerstitial fibrosis Gastrointestinal disease Immunosuppression leads to co- Hepatitis infection with opportunistic pathogens Pneumonitis Fungal infections CNS disease Bacterial infections Retinitis (nocardiosis, listeriosis) Nephritis Viral infections (HHV-6, Pancreatitis HHV-7, HCV) Carditis Viral-associated neoplasia (PTLD) New-onset diabetes mellitus Mortality
The direct effects of CMV syndrome are symptoms resembling virulent mononucleosis infections such as fatigue, body pains, fever, diarrhea and myelosuppression (Table 1.1). Some SOT recipients suffer from CMV invasive disease and that can occur in all types of organ transplantation, and is common in the gastrointestinal tract, CMV-induced ulceration leading to hemorrhage, diarrhea, and perforation (Eid and Razonable 2010, Eid et al. 2010). Indirect clinical impacts of CMV infection in SOT recipients are caused by several subsidiary effects due to the modulation of the immune system (Razonable and
- 48 - Paya 2003). Therefore, opportunistic pathogens infect SOT patients at the same time of CMV infection activation, an example of these pathogens are fungi (Husni et al. 1998), bacteria such as Nocardia spp (Peleg et al. 2007b) HHV-6, HHV-7, hepatitis C and EBV (Eid and Razonable 2010). SOT patients who develop CMV infection or fail to obtain successful CMV preventive treatment are far more likely to undergo chronic or acute graft rejection in lung transplantation and in heart transplant recipients (Eid and Razonable 2010); (Fearon et al. 2007); (Hussain et al. 2007); (Ozdemir et al. 2007); (Thomas et al. 2009).
1.5.2 Immune response to CMV infection
1.5.2.1 Innate Immunity
The innate immune system differs from the adaptive immune system in being non-specific, generic and also for being rapid and represents the first line of defence against foreign pathogens. Also, the innate immune system assists in priming the response of the adaptive immune system. Pattern recognition receptors (PRRs) are involved in the recognition of CMV, they can detect pathogen associated molecular patterns (PAMPs), which are specific small motifs conserved in microbes such as bacterial lipopolysaccharide (LPS) that is recognized by TLR4.
1.5.2.2 Natural Killer (NK) cells
During HCMV infection, NK cell function and number have been reported to increase (Coudert et al. 2010); (Hokland et al. 1988); (Venema et al. 1994) and
- 49 - patients who suffer NK cell deficiencies are at risk of recurrent HCMV and HSV-
1 infection (Coudert et al. 2010); (Orange and Ballas 2006). When HCMV infects cells, it down-regulates the expression of class I MHC (HLA) on the surface of these cells, leading to NK cells initiating the cytotoxic process (Iversen et al.
2005); (Lodoen and Lanier 2005). Nevertheless, immune evasion strategies of
CMV have advanced to overcome NK cells killing strategies by producing UL-40 glycoprotein that can induce the expression of HLA-E which interact with the cognate inhibitory receptor of the NK cell, CD94-NKG2A (Iversen et al. 2005);
(Tomasec et al. 2000); (Wang et al. 2002). Adoptive transfer of NK cells led to postulation of an innate protection role against MCMV and assistance in the clearance of experimental MCMV infection (Crough and Khanna 2009); (Polic et al. 1998); (Bukowski et al. 1985); (Bukowski et al. 1983), furthermore, some mice strains have shown resistance toward MCMV and they become infected with
MCMV when NK cells are depleted (Figure 1.7) (Crough and Khanna 2009);
(Scalzo et al. 1990); (Scalzo et al. 1992).
- 50 -
Figure 1.7 MCMV infections in three different mice strains. (A) no signal for activation of NK cells is available. (B) reduction in MHC-I due to CMV infection leading to activation of NK cells and killing of the cell. (C) even the infected cell is recognized however due to the strong inhibitory signal of Ly49 no killing by
NK cells is started. (D) in this mouse strain NK cells lack Ly49, even though
MHC-I is reduced however activating receptors cannot recognize the infected cell therefore NK cell is not activated and the cell survives (Jensen and Berthold
2007).
- 51 - 1.5.2.2.a Missing self-hypothesis (Reduction in MHC-I expression)
NK cell function is regulated by a group of activating and inhibitory receptors.
Moreover, MHC-I is the traditional ligand for inhibitory receptors of NK cells.
Additionally, the inhibitory receptors that prevent NK cells from confronting self- cells are KIRs, LIR, NKG2A/CD94 and in mice Ly49 (Coudert et al. 2010). If the inhibitory receptors of NK cells were not able to detect MHC-I, this would activate NK cells (Coudert et al. 2010). CMV has the ability to encode several genes that modulate the expression of MHC-I, these genes are US2, US3, US6 and US11 (Table 1.2) (Coudert et al. 2010); (Hengel et al. 1999). In mice, MCMV encodes three viral genes that affect the surface expression of H-2 or MHC-I and they are M152, M06 and M04 (Coudert et al. 2010)
1.5.2.2.b Self cells expression of self-induced Ligands
The NKG2D receptor is expressed by NK cells in both mice and humans and is encoded by the KLRK1 gene; the function of this receptor is to activate NK cells after it binds to a family of ligands called the stress induced proteins. Moreover, stress induced proteins are expressed on cells that are infected or under genomic stress, such as in cancer. The stress induced proteins are for example in mice Rae-
1, Mult-1 and H-60, and MICA, MICB and ULBP 1 to 6 in humans (Coudert et al.
2010). NKG2D in humans is different in structure than mice where it exists with one long NKG2D isoform that initiates an activating signal through DAP10, and in mice as two isoforms (Coudert et al. 2010). CMV has evolved in mice and humans in order to evade the binding of NKG2D+ NK cells to the expressed ligands on CMV infected cells. These evasions strategies act to prevent NKG2D
- 52 - binding to the ligands by encoding immune evasion genes in human and mice’ also, by obstructing the surface expression of NKG2D ligands through the encoding microRNAs (Coudert et al. 2010).
1.5.2.3 Adaptive Immune response
The immune system has two mechanisms to fight the infection and protect the body. The specific immune system or the adaptive immune system comprises the humoral response mediated by B-cells and the secretion of antibodies and cell- mediated immunity that involves activation of T-cells, macrophages and cytokine secretions. The most important feature of the adaptive immune response is the building of immunological memory from previous infection, which protects the body from future recurrent infection.
1.5.2.3.1 Humoral Response to CMV
CMV glycoproteins gB and gH are the major targets for neutralizing antibodies
(Crough and Khanna 2009); (Rasmussen et al. 1991); (Mathew et al. 2001, Britt et al. 1990). Immunization with immune-affinity-purified gB induced the production of immunoglobulin to neutralize CMV (Gonczol et al. 1990), also, recombinant gB induced the production of neutralizing immunoglobulin but this immune- memory did not last for a long period (Frey et al. 1999); (Pass et al. 1999).
Antibodies against HCMV in babies have been transferred from sero-positive mothers to infants and have proven to protect the infants from HCMV infection
(Crough and Khanna 2009, Yeager et al. 1981).
- 53 - 1.5.2.3.2 T-cell-mediated immune responses
CMV is an opportunistic pathogen causing infection in immunocompromised individuals and patients. The immune system reacts mostly to CMV infection through cell-mediated-immunity and the participating immune cells are CD8,
CD4 and γδ T cells specific to CMV (Crough and Khanna 2009).
1.5.2.3.2.a γδT-cells in response to CMV
γδ T-cell levels were noticed to be elevated in renal transplant recipients during effective HCMV infection, in which the levels of γδ T-cells reached about 40% in blood, and the cells further expanded in response to increase in the severity of the infection (Crough and Khanna 2009); (Dechanet et al. 1999a); (Lafarge et al.
2001). In mouse models, γδ T-cell depletion resulted in a rise in MCMV infection
(Crough and Khanna 2009); (Ninomiya et al. 2000). Also, MCMV infection led to increased γδ T-cell numbers in salivary glands (Crough and Khanna 2009);
(Cavanaugh et al. 2003).
1.5.2.3.2.b CD8 + T cells in response to CMV
In mouse models, lymphocyte depletion led to reactivation of viruses such as
MCMV; adoptive transfer of CD8+ T-cells provided protection against MCMV which assisted in clarifying the significant role of CD8+ T-cells in preventing the reactivation of pathogens such as MCMV (Crough and Khanna 2009); (Mutter et al. 1988); (Polic et al. 1998); (Reddehase et al. 1985). In humans, it was noted that
HCMV could be reactivated in individuals deficient in CD8+ CMV-specific T
- 54 - cells (Crough and Khanna 2009) and HCMV specific-CD8+ T-cells have been noted to provide protection against viraemia in several groups of transplant patients such as BMT, lung and SOT (Riddell et al. 1992b); (Li et al. 1994);
(Quinnan et al. 1982). It is estimated that healthy CMV+ individuals can have about 10% HCMV-specific-CD8+ T-cells, and this percentage increase to 40% in the elderly in a process called memory inflation (Crough and Khanna 2009);
(Crough et al. 2007); (Gillespie et al. 2000); (Khan et al. 2004); (Sylwester et al.
2005). Additionally, CD8+ T-cells that are specific for the HCMV IE-1 protein were detected in lung and heart transplant recipients and found to provide a protective role against HCMV reactivation (Bunde et al. 2005). Furthermore, obtaining both HCMV-specific CD8+ and CD4+ T-cell immunity assisted in preserving successful lung allograft function and stability, in contrast to those who developed HCMV infection due to the failure of developing competent CD8+ T- cell immunity against HCMV (Shlobin et al. 2006). Primary HCMV infection in utero induced foetal CD8+ T-cells to expand in response to CMV infection
(Marchant et al. 2003). CD8+ T-cells specific for HCMV have been shown to have significant function in recovery from HCMV reactivation in bone marrow transplant patients (BMT) (Quinnan et al. 1982); (Radha et al. 2005, Reusser et al.
1991); (Li et al. 1994). Furthermore, HCMV specific CD8+ T-cells restored cellular specific immunity against HCMV when obtained from donors with
HCMV-specific-CD8+ T-cells and infused into allogeneic BMT recipients
(Riddell et al. 1992b); (Walter et al. 1995). In renal transplant patients, CD8+ T- cells specific to CMV provided protection against viral reactivation (Radha et al.
2005, Reusser et al. 1999, Sester et al. 2005). Distinct epitopes are targeted by
HCMV-specific-CD8+ T-cells that include immune modulators encoded by the
- 55 - virus such as pp28, pp150, and several structural and replication antigens such as
IE, early, early-late and late virus antigens and functions-associated-proteins such as capsids and immune evasion proteins (Elkington et al. 2003); (Manley et al.
2004); (Sylwester et al. 2005).
Figure 1.8 CD4 + and CD8+ T cell responses to HCMV. A) the pie graphs represent the response of CD4+ and CD8+ T cells to HCMV proteins encoded through the virus replication process (IE, early, early-late, and late; left) and HCMV proteins correlated with diverse functions (capsid, matrix/tegument, glycoprotein,
DNA/regulatory, and immune evasion; right). B) Schematic representation of the magnitude of CD4+ and CD8+ T-cell responses against immune-dominant HCMV- encoded ORFs such as UL123 (IE-1), UL122 (IE-2), and UL83 (pp65) (Crough and
Khanna 2009).
- 56 - Studies of CD8+ T cells during HCMV acute and chronic infection have revealed during acute infection CD8+ T cells are CD45RA-, CD45RO+, CD27+, CD28+/- and CCR7-. Moreover, during chronic infection two types of HCMV-CD8+ T cells co-exist, effector memory and terminally differentiated cells; the first are
CD45RA-, CD45RO+, CD27-, CD28-, CCR7- and the terminally differentiated cells are CD45RA+, CD45RO-, CD27-, CD28-, CCR7- (Appay et al. 2002);
(Gamadia et al. 2003). Additionally, a study indicated that the expansions of
HCMV-specific CD8+ T cells are oligo-clonal with some clones displaying the
CD28-, CD57+, and CCR7- phenotype of differentiated effector memory cells
(Khan et al. 2002). The clonal expansion of HCMV specific CD8+ T cells may lead to domination of these HCMV specific cells and may hinder the immune response toward other pathogens; this is supported by the findings connecting lower success of influenza vaccination with HCMV sero-positivity, and also
HCMV being a factor that assists the advance of AIDS (Sabin et al. 2000,
Webster et al. 1989).
1.5.2.3.2.c CD4+ T cells in response to CMV
T helper lymphocytes have significant and central roles in directing the adaptive immune system. They produce a variety of cytokines and chemokines with several immune functions, play an important role in B-cell antibody class switching, inducing phagocyte functions, assisting the growth of cytotoxic T-cells and attracting neutrophils, basophils and eosinophils to the infection site. The median frequency of HCMV-specific CD4+ T-cells in the blood is about 9% and in some individuals this rises to 40% (Sylwester et al. 2005); (Sester et al. 2002).
- 57 - In murine studies, repeated infection with MCMV resulted in mice undergoing
CD4+ T-cell depletion (Polic et al. 1998), also, the role of CD4+ T-cells to regulate MCMV infection was noted after depletion of CD8+ T-cells (Jonjic et al.
1994). The role of HCMV-specific- CD4+ T-cells was revealed when clinical complication of HCMV infection appeared in lung transplant recipients following acute deficiency of HCMV-specific-CD4+ T-cells, and this was also noted in renal transplant recipients (Sester et al. 2002); (Sester et al. 2005). During the latent status of the virus HCMV-specific-CD4+ T-cells assist indirectly to induce
B-cell production of antibodies and expansion of cytotoxic CD8+ T-cells (Walter et al. 1995); (Davignon et al. 1996). Also, the recovery of CD4+ T-helper cells is essential for the reconstitution of cytotoxic CD8+ T-cells (Reusser et al. 1991).
The importance of effector memory CD4+ T-cells to control HCMV was further studied in renal transplant recipients. A longitudinal study showed that the
HCMV-specific- CD4+ T-cell response was hindered in symptomatic compared to asymptomatic recipients following HCMV infection, suggesting a significant role of these cells in regulating recovery from HCMV infection (Gamadia et al.
2003). In BMT patients, helper T-cells were shown to help in protection from
HCMV infection as long they were present in detectable quantities (Hebart et al.
2002); (Krause et al. 1997); (Li et al. 1994). Another study suggested that CD8+
T-cell activation required the assistance of CD4+ helper T-cells; in this study predominantly HCMV-specific-CD4+ T-cell lines were infused into the seven patients and five showed progression in clearing HCMV infection, moreover, the infusion of HCMV-specific-CD4+ T-cells lines was followed by expansion of
CD8+ T-cells (Einsele et al. 2002). Furthermore, the study of HCMV-specific-
CD4+ T-cell specificity showed the capability of gB and gH and several other
- 58 - antigens to be recognised as frequent targets of HCMV (Figure 1.8) (Sylwester et al. 2005). Likewise, antigen presenting cells (APC) that present endogenous gB to
CD4+ T-cells resulted in production of granzyme-B (Hegde et al. 2005); an immune-dominant peptide epitope of gB-specific CD4+T-cells is
DYSNTHSTRYV restricted to HLA DRB*0701 (Elkington et al. 2004).
1.5.3 CMV immune-evasion
The Herpesvirus family has developed several strategies to evade both the innate and adaptive immune systems during the life cycle of the virus in the body. CMV specific evasion strategies assist in maintaining the existence of the virus inside the body between active and dormant states. These techniques of evasion can cause minor effects in healthy seropositive individuals when the virus reactivates, however, in immune-compromised or immune-suppressed individuals the virus can cause major health impacts and can lead to mortality. CMV has acquired genes homologous to host genes that occupy about 20% of the total CMV genome and can encode components of the host immune response such as MHC-I, cytokines and chemokines, extracellular and intracellular receptors (Loenen et al.
2001). MHC-II that is generally expressed by APCs is also affected by the CMV gene US2 via several methods; US2 can degrade DR-α and DM-α and also MHC-
II induced expression by IFN-γ is interfered with by the expression of the protein during immediate early and early CMV infection (Loenen et al. 2001); (Crough and Khanna 2009).
Non-classical HLA-E binds to CD94/NKG2A and suppresses the activation of
NK cells. CMV, however, reduces the expression of classical MHC-I on the surface therefore to evade CTL lysis, while the CMV gene UL-40 induces the
- 59 - expression of HLA-E on the cells infected with the virus and protects them from
NK cell attack (Loenen et al. 2001); (Tomasec et al. 2000).
1.5.3.1 CMV genes homologous to host genes
CMV has evolved to acquire genes that are homologous to host genes with potential roles in the immune response to the virus. MHC-I of the host has a homologous gene in CMV named UL18 that binds to leucocytes immunoglobulin like receptor 1 (LIR-1) and can inhibit several immune cells that express the ligands CD94/NKG2 such as NK cells and T-cells and including B-cells, DC and
γδ T-cells. UL33, UL78M, US28 and US27 are homologues to G protein coupled chemotactic cytokine receptor or chemokine receptor 1 (CCR1) that has an essential role in directing and migrating cells to the site of the infection and inflammation (Loenen et al. 2001); (Lalani et al. 2000); (Beisser et al. 1999).
UL111A is homologous to IL-10 and the potential functions of this gene are to inhibit cytokines IFN-γ and IL-12 and immune cells such as DC (Loenen et al.
2001). The viral gene UL144 encodes a tumor necrosis factor receptor super gene family (TNFR) homologue that stimulates several subsets of lymphocytes
(Loenen et al. 2001). Mutually UL146 and UL147 are homologous to IL-8 and have a role in lymphocyte trafficking to the site of inflammation and infection.
Research correspondingly is expecting to discover genes that induce FcR on cells invaded by CMV and can protect the cells from antibody mediated immune response that leads to elimination of cells hosting this virus (Loenen et al. 2001);
(Farrell et al. 1999).
- 60 - 1.5.3.2 CMV evading detection by Cytotoxic T cells
Peptides from intracellular antigen are degraded by proteasomes into short peptides 8 to 12 amino acids in length which are presented on MHC-I and can be recognized by CD8+ cytotoxic T cells. In order for HCMV to evade CTL it has evolved to acquire several different gene products: gpUS2, gpUS3, gpUS6, gpUS10 and gpUS11, furthermore, MCMV encodes three genes that interfere with MHC-I intracellular trafficking and their mechanisms of action are quite different from HCMV US genes; they are: m04, m06 and m152 (Loenen et al.
2001); (Crough and Khanna 2009). These gene products perform a variety of function and are expressed at several time points to interfere with MHC-I production and expression. US2 is an early gene that binds MHC-I in the ER and rapidly reduces the life-span of MHC-I from about six hours to less than two minutes (Loenen et al. 2001); (Tortorella et al. 1998). Next is US3, an immediate early gene that holds MHC-I inside the ER by binding to heavy chain/β2m complexes, though it does not block peptide binding to MHC-I (Loenen et al.
2001); (Jones et al. 1996). US6 appears later beyond the expression of US2 and
US3 and remains expressed by the virus through its life span, this gene affect the loading of the peptides on MHC-I inside the ER (Jun et al. 2000). Another early gene is US11 that functions to retain MHC-I in the cytoplasm of the cell (Wiertz et al. 1996).
1.6 Kidney Transplant Rejection
Renal transplantation is the favoured method of treatment for patients at end-stage renal failure or patients with established renal failure. This treatment provides
- 61 - many advantages that overcome the long repeated haemodialysis, which is high in cost and in general can transmit infection to the patients under repeated haemodialysis. Moreover, patients under repeated courses of haemodialysis are in danger of pain and mortality. The survival of the kidney is dependent on not being rejected by the immune system so to avoid short-term kidney rejection requires the use of immunosuppressive therapy (Issa et al. 2010). Rejection of a transplanted organ by the immune system is a general reason leading to graft failure. Even though the organ donor is selected after satisfying several criteria to match the recipient, rejection may occur and can lead to several adverse complications in both the graft and the body of the recipient.
The treatment to tackle allograft rejection was directed toward inhibiting T cell immune responses, due to the belief that T cell-mediated responses lead to inflammation and that have been alleged for more than half a century to be the main reason behind allograft rejection. These treatments managed to prolong graft survival but were not able to stop acute and chronic graft rejection which through immunohistochemical visualisation techniques provided evidence of cell- antibody- and complement-mediated damage leading to graft rejection (Gonzalez and Torres-Lopez 2012). The binding of receptors to their corresponding ligands such as LFA-1 (ICAM-1), CD40-CD40L and CD28-B7 are important to trigger the T-cell dependent immune response. Additionally, long term-survival of the graft can be achieved by blockade of these interactions, promoting graft tolerance
(Seino et al. 2001).
Donor MHC can be targeted by recipient T cells through two mechanisms of recognition, the first is through direct allorecognition in which T cells directly recognise donor MHC and target it. The other process is called indirect
- 62 - allorecognition in which recipient APC take up donor alloantigen and present it to
T cells, initiating an immune response against the graft (Gonzalez and Torres-
Lopez 2012). Also, alloantibodies targeting non-classical MHC-I alloantigens
MICA and MICB have been recognised as one cause of acute renal allograft rejection (Sumitran-Holgersson et al. 2002); (Zou et al. 2006, Mizutani et al.
2005). MHC-II DR and DQ are only expressed on APC and B cells and can stimulate the development of alloantibodies and T cells activation; MHC-II is also found on micro-vascular endothelial cells (Erlich et al. 2001).
The rejection mechanism of kidney transplants results from interactions between innate and adaptive immune systems. Additionally, the outcome of this interaction is hyperacute, chronic or acute graft rejection by the immune system (Issa et al.
2010).
1.6.1 T cells and B cells interface and generation of alloantibodies
The assistance of neighbouring T cells is essential for B cells in order to generate high affinity antibody. As APC present antigen to T cells, a general change in T cell shape arises and is followed by up-regulation of chemokine receptor CXCR5
(von Andrian and Mackay 2000, Ono et al. 2003). T cells then respond to chemokine CXC13 and migrate toward B cells located in primary follicles (Ono et al. 2003). This is followed by B cells becoming activated and acquiring CCR7 that interacts with both CCL19 and CCL2 chemokines and then B cells express antigenic peptide on MHC-II (Zinkernagel and Doherty 1997), that, specifically, in a transplantation situation is derived from donor MHC. B and T cells, after that, extend their contact surface area to allow thorough communications between their
- 63 - receptors and other ligands (Huppa and Davis 2003). B cell presentation of peptide to T cells through the MHC-II groove leads to generation of efficient antibodies against this peptide. The bridging between B and T cells is further improved due to other links to allow the production of first signal in which CD4+
T-cells bind to MHC-II molecules at the non-polymorphic region. This is followed by high affinity binding between LFA-1 (CD11a-CD18) of T cells and its ligands or intercellular adhesion molecule (ICAM-CD54) on B cells which strengthen the bridge between T and B cells (Joosten et al. 2002). After that, the antigen bridge relocates centrally to trigger the signal between T and B cells
(Scholl and Geha 1994).
1.6.2 T-cell role in graft rejection and graft tolerance
T-cell immune surveillance recognises the graft either directly or through allorecognition of donor peptides, followed by T-cell activation. Furthermore, effector responses triggered by T cells direct the immune system toward this graft leading to damage to or rejection of this graft (Issa et al. 2010). Allogeneic transplantation of organs is always under the direct danger of acute and chronic rejection that is directed by CD4+ and CD8+ T-cells (Issa et al. 2010). T-cells allorecognition of the mismatched MHC or alloantigen of the graft follows by their activation in the presence of the costimulation signal required by T-cells
(Issa et al. 2010). Activation of T-cells leads to their expansion to generate allo- reactive T-cells against the graft (Issa et al. 2010).
- 64 - 1.6.2.1 The allorecognition process
Mismatch in MHC between the graft and the recipient is the first trigger to initiate the immune response. Generally, allorecognition is defined as the process of detecting non-self-antigen although this antigen is from the same species as the host or graft recipient (Issa et al. 2010). In addition to polymorphic MHC, allorecognition can occur with minor histocompatibility antigens (miH). The latter are defined as peptides from allelically polymorphic proteins other than the donor’s MHC antigens, and further expressed by MHC of the host or recipients to
T-cells of the host (Issa et al. 2010, Game and Lechler 2002). Multiple miH mismatches between the graft and the recipient can evoke the host immune response against the graft leading to damage and rejection, which can speed up the rejection process if MHC is mismatched as well (Issa et al. 2010). Moreover, miH can evoke a slow rejection process even when MHC of the graft is compatible to the recipients (Dragun and Hegner 2009). Minor-histocompatibility antigens can be presented on both MHC-I and –II and have been encoded in both human (such as HA-2 antigen derived from myosin class I and restricted to HLA-A2) and mice
(about 40 miHs identified) on sex chromosomes, DNA of mitochondria and autosomes (Game and Lechler 2002). Instead of MHC-I presenting self antigen to
T cells to inhibit their cytotoxic activity, in case of transplantation the antigen from the graft is presented to T cells by self-MHC-I and this method is defined as indirect allorecognition. Moreover, APC from the donor migrating from the graft can present the donor self antigen to TCR of T-cells of the recipient which triggers the activity of T-cells and this way is defined as direct allorecognition.
The third pathway is defined as semi-direct allorecognition pathway where APC
- 65 - of the recipient present the donor antigen to TCR of T-cells of the recipient (Issa et al. 2010).
1.6.2.1.a. Direct allorecognition pathway
This allorecognition pathway was believed to be the single method of allorecognition of graft antigen, which led to immune responses toward the graft and maybe followed by rejection (Figure 1.9:A) (Afzali et al. 2007). The specific nature of the ligand that stimulates alloreactive T-cells is unclear however two theories have been suggested regarding the method of allorecognition of alloantigen on MHC-I molecule. The first hypothesis called high determinant density, which implies the capability of alloreactive T-cells to recognise the clear polymorphism of any allogeneic MHC and the peptide that is expressed in the groove of this allogeneic MHC is less priority (Afzali et al. 2007, Game and
Lechler 2002)
Therefore, any non-self MHC will be addressed by TCRs of alloreactive T-cells as ligands, this however results in high number of allogeneic non-self MHC as ligands capable of triggering the immune response against the graft. In addition, even TCRs of lower affinity will respond against non-self-polymorphic MHC
(Game and Lechler 2002, Afzali et al. 2007). What supports this hypothesis is the incidence of alloreactivity even in the absence of the peptides, moreover, there are several studies that show blocking (Schneck et al. 1989) or site mutation
(Villadangos et al. 1994) of TCR and MHC contacting regions would inhibit the generation of an alloresponse (Game and Lechler 2002, Afzali et al. 2007). The other hypothesis is the multiple binary complex model that proposes that T-cell
- 66 - alloreactivity is due to the non-self peptide expressed in the groove of MHC-I; this hypothesis claims that even a single non-self peptide is capable of inducing many T-cells against this peptide; this hypothesis was explained by the occurrence of alloreactive CD8+ T cells toward self-antigen expressed in the groove of a non- self MHC-I molecule (Whitelegg et al. 2005); Game and Lechler 2002, Afzali et al. 2007). Both hypotheses do not comply with the concept of thymic positive selection, but the resemblance of several alleles of MHC at the TCR contact region permit the probability of alloresponse in the context of positive selection
(Game and Lechler 2002).
Several attempts have been made to avoid direct allorecognition and expand the survival period of the graft, for example by purifying the graft from leucocytes via depleting the donor of leucocytes. In addition, rat kidney transplantation studies have managed to reach the point of thorough tolerance by the recipient immune system toward the kidneys; this was achieved after transplanting the kidneys into an intermediate recipient for the goal of depleting the kidneys of any donor derived leukocytes, however, the other way around by introducing DC from the donor into the recipient circulation resulted in acute rejection (Game and Lechler
2002).
1.6.2.1.b Indirect allorecognition pathway
This pathway is defined as expression of donor peptide by self-MHC on APC, which is generally different from the direct pathway in the process of antigen processing. Recipient APC obtain the alloantigen from the graft and treat it as a foreign antigen and express this peptide on MHC-II to CD4+ T-cells (Afzali et al.
- 67 - 2007), which CD4+ T-cells are essential for B-cell function to generate antibodies, or CTL (Figure 10:B). Furthermore, this pathway is believed to be responsible for the long-term rejection of the graft as the direct pathway depends on the availability of donor APC that disappear with time (Afzali et al. 2007).
This pathway is further supported by two approaches in which immunizing animals with allogeneic peptides resulted in strong rejection. Moreover, prolonged survival of the graft was detected upon intra-thymic injection of its peptide (Afzali et al. 2007).
1.6.2.1.c Semi-direct allorecognition pathway
This pathway of allorecognition implies that many surface expressed molecules even MHC can be moved and transferred between cells in order to stimulate the immune system of the recipient cells (Herrera et al. 2004). The methods involved molecular transfers through cell-to-cell contact (Game et al. 2005), and exosomes or small vesicles transfer (Morelli et al. 2004). This allorecognition pathway is directed by recipient APC receiving donor MHC-peptide complex via the process of MHC-transfer. Moreover, this APC after receiving MHC-peptide complexes are able to stimulate CD8+ T cells (the direct allorecognition pathway) and also process the peptide and express it to CD4+ T cells (indirect allorecognition pathway). Therefore, this method combines both allorecognition pathways and recruits both CD4+ and CD8+ T cells (Figure 10:C) (Afzali et al. 2007).
- 68 - 1.6.3 NK cell allorecognition
NK cell mediated allorecognition in solid organ transplantation is poorly identified. KIRs recognise self-MHC-I to stop the activation and killing of target cells by NK cells. Therefore, mismatched allografts can lead to graft targeting by
NK cells due to lack of self MHC-I on the graft (Afzali et al. 2007). Studies demonstrated evidence of NK cell infiltration as part of the innate immune response into the graft prior to T-cell infiltration; NK cells can secrete IFN-γ to induce both MHC-I and II expression (Pratschke et al. 2009). During development, NK cells undergo ‘licensing’ whereby those cells with inhibitory receptors for self-MHC class I alleles are allowed to mature. Hence, NK cells will recognise and kill target cells lacking self-MHC (Jonsson and Yokoyama 2009).
Alloreactivity can be triggered due to mismatches between the donor MHC and
NK cell inhibitory receptors against self-MHC expressed on recipient cells (Afzali et al. 2007). Moreover, depletion of NK cells has not been shown to prevent the occurrence of rejection, which is different from T-cell depletion that assisted in preventing rejection (van der Touw and Bromberg 2010). Moreover, following
NK cells after transplantation showed that they become activated due to the stress ligands of the allograft, therefore, activated NK cells produce IFN-γ and other proinflammatory cytokines that activate T-cells (Obara et al. 2005, Maier et al.
2001).
- 69 -
Figure 1.9 Three allorecognition pathways. A) the direct pathway where donor
APC express donor-self antigen on donor self-MHC and prime both CD4+ and
CD8+ T-cells. CD4+ T-cells assist CD8+ T-cells to inflict effector cytotoxic functions against the graft. B) Indirect pathways where recipient APC obtain donor antigens through necrotic and apoptotic cell material which are then processed and presented to CD4+ T cells. C) semi-direct pathway that includes molecule exchange between cells, (a) APC from both donor and recipient transfer membrane components such as allo-MHC-peptide complex. (b) Donor APC release endosomes containing MHC that fuse with membrane of the recipient
APC (c). Then donor APC stimulate the immune system through both direct and indirect allorecognition pathways (Afzali et al. 2007).
1.6.4 Development of the Alloresponse
The stress caused in the region of graft due to injuries caused by surgical procedures leads to local release of several cytokines and chemokines, moreover,
- 70 - these release of these immune mediators promotes the attraction of immune effector cells to this region and promotes rejection of the graft (Issa et al. 2010).
1.6.4.1 The alloresponse of CD4+ T-cells
It was thought that Th2 cells only mediate tolerance and Th1 cells promote graft rejection, however, recent studies suggested that both types of Th cells promote rejection (Issa et al. 2010). Th1 promote rejection through the release of IFN-γ and IL-2 that activates CD8+ T-cells, B-cells release IgG2 that activates complement and CD8+ T-cells also promote macrophage dependent delayed-type hypersensitivity. What is more, CD8+ T cells also may express Fas-L that promotes CD8+ T-cell cytotoxicity (Issa et al. 2010). Increases in Th2 cytokine gene expression were noticed in murine cardiac allograft rejection (Martinez et al.
1993a). Th2 cytokines IL-10, IL-4, IL-5, IL-13 and IL-9 promote rejection through activation of B-cells and eosinophils (Issa et al. 2010, Martinez et al.
1993b). Another study has shown Th2 cytokines within the graft are responsible for graft infiltration by eosinophils (Martinez et al. 1993b).
1.6.4.2 The alloresponse of CD8+ T-cells
CD8+ T-cells are believed to be the main participant leading to acute rejection, however, even after depletion of CD8+ T-cells acute rejection still occurs (Issa et al. 2010) although CD8+ T-cells are one of the main causes of apoptosis and graft rejection. In kidney graft rejection, CD4+ and CD8+ T-cells infiltrate the regions surrounding the tubules and express mediators promoting apoptosis: FasL,
- 71 - granzymes and perforin (Cornell et al. 2008, Robertson et al. 1996). Moreover, mRNA expression of granzyme B, FasL and perforin were elevated in the regions of tissue rejection (Issa et al. 2010). CD4+ T cells may mediate rejection without the participation of CD8+ T cells by a mechanism similar to delayed hypersensitivity by attracting macrophages into the graft (Cornell et al. 2008).
1.6.5 CD56+ T-cell response in allografts
The role of CD56+ T-cells in transplantation has not been established thoroughly.
Few studies have investigated their activities and whether these cells are promoting rejection or inducing tolerance of the kidneys or other grafts. In animal models, after liver transplantation donor NK and CD56+T-cells were detected in the blood of the recipients for two weeks after liver transplantation; however, these were not detected in renal transplantation (Moroso et al. 2010). In another study performed on mice, iNKT cells have been found to induce tolerance of the immune system towards cardiac allografts. In this study, iNKT cell deficient mice were not capable of developing long-term tolerance and acceptance to the cardiac graft, however, adoptive transfer of iNKT cells to the deficient mice assisted in development of long-term tolerance and acceptance of the cardiac graft.
Moreover, the blockade of LFA-1-ICAM-1 or CD40-CD40L interactions induced long-term survival of cardiac grafts in the presence of iNKT cells (Seino et al.
2001). In bone marrow transplantation in rodent models, CIK cells exerted graft- versus-host-tumour (GVT) effects (Edinger et al. 2003), which is due to their potent production of IFN-γ that exerts protective functions against GVHD (Baker et al. 2001).
- 72 - 1.6.6 The role of CMV in allograft rejection
In renal transplant patients CMV infection and disease can occur acutely or from reactivation of latent CMV. Acute primary infection can be triggered in the first months due to suppression of the immune system, however, it can be delayed until after the administration of prophylactic treatment (Weikert and Blumberg 2008).
In renal transplant recipients CMV infection can be asymptomatic, identified by
CMV replication or CMV disease may ultimately lead to organ rejection (Weikert and Blumberg 2008).
The activation of CMV triggers the host immune response, CMV releases immediate early antigens after it penetrates the cells; these genes are involved in the regulation of DNA replication. Following this by 6 to 24 hours, the virus releases late antigens to direct the production of nucleo-capsid proteins and the up-regulation of IL-2 (Brennan 2001, Weikert and Blumberg 2008).
Several studies on the association between CMV and kidney rejection, identified
‘CMV syndrome’ by the presence of CMV antigen in the blood and fever, muscle pain, leucopoenia, hepatitis, gastroenteritis, pneumonitis and thrombocytopenia
(Hartmann et al. 2006, Weikert and Blumberg 2008). A prospective study in patients who did not receive prophylactic treatments showed about two thirds of the patients developed CMV infection during the first 100 days, among the different groups of donor and recipient status. Moreover, D+R- patients had three times higher chance of developing CMV disease than other groups D-R+ and
D+R+ (Hartmann et al. 2006).
- 73 - 1.7. Aims and objectives
CMV infection has minor effects in healthy subjects; however, the activation of this virus in immunocompromised subjects can lead to serious clinical outcomes.
The reactivation of this virus in organ transplant recipients can lead to viraemia and graft rejection. CD56+ T-cells have many roles in different types of diseases, however the response of these cells to CMV has not been established. The main aim of the study is to examine the relationship between CD56+ T-cells and CMV, and in kidney transplant patients to assess whether this relationship affects CMV disease:
To compare the levels of CD56+ T-cells between CMV positive and CMV
negative healthy individuals.
To compare the levels of CD56+ T-cells in kidney transplant patients
according to CMV status of donor and recipient.
To investigate the phenotype of CD56+ T-cells in both healthy subjects
and kidney transplant patients according to CMV status.
To study the functional capabilities of CD56+ T-cells when stimulated by
CMV lysate. For this purpose several tests will be applied.
To investigate changes in gene expression in CD56+ T-cell when
stimulated by CMV lysate.
- 74 -
Chapter 2
Materials and Methods
- 75 - 2. Materials and Methods
2.1 CMV IgG ELISA
CMV IgG ELISA kits were purchased from Genway Biotech (Lot # 40-521-
475373 San Diego, Ca). This kit was used for qualitative detection of IgG antibodies against CMV in healthy donors participating in this study. The kit is suitable for detection of CMV IgG antibodies in human serum and plasma.
2.1.1 CMV IgG ELISA assay Principle
Detection occurs after the binding of CMV IgG antibodies to CMV antigen that is pre-coated in the microtiter strip wells. The washing of the wells is set to remove any background and unbound sample substances, then anti-human IgG conjugate that is labelled with horseradish peroxidase (HRP) binds to the CMV IgG antibodies and CMV antigen complex on the well. This complex is visualised via the addition of Tetramethylbenzidine substrate (TMB) to generate a blue colour in the wells. Sulphuric acid is further added to stop the reaction to generate a yellow endpoint colour (Figure 2.1), and the micro-well plate is analysed by Thermo
Labsystems Opsys MR ELISA at an absorbance of 450 nm as recommended by the company’s protocol.
2.1.2 Reagents and materials supplied and preparation
CMV coated wells, IgG sample diluents, stop solution (Sulphuric acid), washing solution, CMV anti-IgG conjugate (Peroxidase labelled antibody to human IgG),
- 76 - TMB substrate solution (Tetramethylbenidine), CMV IgG positive control, CMV
IgG cut-off control, CMV IgG negative control. All reagents are ready to use except the washing solution 20 x, this reagents was diluted 1:20 by adding sterile distilled water. Serum or plasma was prepared by diluting 5 µl serum or plasma with 500µl IgG sample diluent (1:100) and the mixture was suspended thoroughly by vortex.
2.1.3 ELISA assay procedure and measurement
ELISA was performed manually and five different controls were allocated and added in duplicate manner except the substrate blank; one well for substrate blank, two wells for the negative control, four wells for the cut-off control and two wells for the positive control. Donor serum or plasma was aliquoted into the microtiter plate in duplicate using sterilized pipette tips. At the beginning one well was left for substrate blank, then 100 µl controls were added and 100 µl diluted samples were added to their allocated wells, and the microtiter plate was covered with foil and incubated for an hour at 37 ᴼC. After that, the microtiter plate was aspirated from the contents and washed manually three times with 300 µl washing solution followed by tapping the microtiter plate on tissue paper to remove any remaining fluids that can result in poor precision and fluctuating absorbance value. The next step was to add 100 µl CMV anti-IgG conjugate to all wells except the substrate blank wells and the microtiter plate was incubated for 30 minutes at room temperature in the dark. Again the microtiter wells were washed manually with 300 µl washing solution three times and contents were aspirated and tapped on tissue paper. Then, 100 µl TMB substrate solution was added to the
- 77 - wells and the microtiter plate was incubated for 15 minutes in the dark. This followed by adding 100 µl stop solution into all wells and at this step all wells that previously emitted a blue colour well turned into a yellow colour. The measurement of the plate was performed at 450 nm and the substrate blank well was adjusted to zero (Figure 2.1).
Figure 2.1 ELISA assay principle. IgG from seropositive donors binds to wells pre-coated with CMV antigen, peroxidase conjugated antibody binds to CMV antigen-antibody complex. Subsequently, addition of TMB visualizes the immune complex and the reaction is stopped by the addition of stop solution that generates a yellow colour.
2.2 CMV lysate preparation
2.2.1 Fibroblast Cell Culture
Fibroblasts (SL fibroblast) were provided from Dr. Neil Blake, CIMI, Institute of
Infection and Global Health. The cells had been stored frozen in liquid nitrogen and thus were left to thaw quickly. Dulbecco’s Modified Eagle’s Medium
(DMEM) supplemented with 10 % FBS, 2 mM L-glutamine, 50 IU/ml penicillin,
- 78 - streptomycin and Amphotericin B was added to the vial slowly to avoid causing hypotonic shock to the cells. Then the cells were washed twice with DMEM at
400g for 10 minutes to remove DMSO from the cells. The cells were then incubated in a T-75 flask with 20 ml DMEM in a humidified 5% CO2 atmosphere at 37oC. The cells were checked every day for growth until the growth was >90% confluent, subsequently the fibroblasts were passaged by removing from the flask by using Cell Dissociation Solution (Sigma Aldrich, lot number C5914-100ML).
The removal of cells was performed in a class II cabinet as follows: cell dissociation buffer was warmed in a water bath at 37ᴼC for 10 minutes. The media was removed from the flask and the fibroblasts were washed with sterile PBS to remove all traces of DMEM from the flask. After that, 5 ml of cell dissociation buffer was added to the flask and the flask was rocked to distribute buffer around the flask, then it was incubated for 10 minutes in a humidified 5% CO2 atmosphere at 37oC. The flask was tapped firmly with the palm of the hand to remove the adherent cells and DMEM was added to inhibit cell dissociation buffer and the cells washed twice at 400g for 10 minutes. The cells were then counted and cultured in further flasks to acquire enough proliferative cells for all subsequent experiments.
2.2.2 CMV lysate preparation protocol-
CMV lysate was prepared by infecting cultured fibroblasts with live CMV of
Towne and AD169 strains (obtained from Dr. Naeem Khan). Fibroblasts were cultured according to the steps described earlier (2.2.1). Adequate cell numbers to perform infection at a multiplicity of infection (Wolf et al.) of 10 were obtained
- 79 - when confluence reached 75 % which is equal to ~ 1.5 × 10 6 cells. Then the cells were infected with live CMV, therefore, each cell was infected with a mean of 10 viruses, that is equal to 1.5 × 107 viruses diluted in 5 ml DMEM. Before adding the virus to the flask the media was removed, and then 5 ml of DMEM containing the live virus was added to the cells and incubated for 90 minutes in a humidified
o 5% CO2 atmosphere at 37 C, agitating the flasks every 15 minutes. After that, the media was removed and 20 ml fresh DMEM media was added to the flasks. The supernatant was collected daily after the cells began to show cytopathic effect
(CPE) for three days until the cells detached totally from the flask when a sterile scraper was used to harvest any adherent fibroblasts. The supernatants were washed at 400g for 10 minutes and the pellet was collected. Moreover, the pellets from all flasks were processed by freezing rapidly for 10 min at -80ᴼC then warming rapidly to 57ᴼC for 10min several times to rupture and destroy all cells in the pellet, then the lysate were exposed to UV light to eradicate all live viruses in the pellet. The lysate was then collected in several eppendorfs and stored at -
80ᴼC. The efficiency of the lysate to stimulate lymphocytes was tested by intracellular cytokine staining of IFN-γ production; furthermore, the cells were stimulated with different CMV lysate concentraions (0.1 µl, 0.5 µl, 1 µl, 1.5 µl, 2
µl, 2.5 µl, 3 µl) and the optimal concentration to stimulate 1 X106 PBMCs/ml was
2 µl CMV lysate.
2.3 Peripheral blood
2.3.1 PBMC Donors
PBMCs were collected from two groups of donors. The first group were healthy donors who were CMV seropositive and sero-negative. The second group were
- 80 - post-transplant kidney recipients. PBMCs were collected into 20ml universal tubes containing 600 units of preservative-free heparin (Wockhardt, Wrexham).
The patient PBMCs were divided into three groups according to their CMV sero- status as follows: donor negative / recipient negative (D-R-), donor positive / recipient negative (D+R-) and the third group was donor either donor negative or positive / recipient positive (R+). Patient PBMCs were provided by Mr A
Hammad, Transplant Unit, Royal Liverpool University Hospital. Blood samples from both healthy subjects and transplant patients were taken following informed consent.
2.3.2 PBMC Isolation
Isolation of PBMC from peripheral blood was carried out in a Class II microbiological safety cabinet. Blood samples obtained from both healthy subjects and transplant patients were slowly layered onto the same volume of
Ficoll-PaqueTM (Sigma, UK) in universal tubes in a diluted quantity 1:1. This was followed by centrifugation at 400g (1800 rpm) for 25 min at 18oC with the brake off. After centrifugation, different layers were formed as shown in Figure 2.2.
Before harvesting PBMCs, the plasma of healthy donors were collected into autoclaved eppendorf tubes and stored at -80oC for anti-CMV IgG ELISA testing.
The buffy coat interface containing PBMCs was harvested into universal tubes and washed twice with autoclaved PBS (Sigma) by centrifugation at these settings: 10 minutes, 400g (1800 rpm) and brake on. Following the washing step, the cells were resuspended in culture medium, counted and prepared for further analysis.
- 81 - Figure 2.2 PBMC separations. This figure illustrates the layers of cells after separation by Ficoll-
Paque centrifugation. This figure is adapted from
(www.Nature.com).
2.3.3 Counting of PBMCs
A TC10 Bio-Rad automated cell counter was used for counting the number of cells. 10 µl of cell suspension was loaded into TC10 counting slides, the loading being performed at an angle of 45° gently to permit capillary loading of the slide by the cell suspension. The loading was performed with care to avoid damaging the slide optical surface and to avoid the formation of bubbles that would interfere with the reading. TC10TM Counting slides are not reusable and they are disposed of as a biohazard waste.
2.4 Flow cytometry (FACSCalibur™ BD Biosciences)
A FACSCalibur was used to analyze cell surface and intracellular staining of lymphocyte populations with fluorescently conjugated mAb. Initially, labelled mononuclear cell populations were run through the FACS and the lymphocytes gated using a combination of FSC (Forward scatter) and SSC (Side scatter)
(Figure 2.3A). To define the position of CD56+CD3+ cells, PBMCs were stained
- 82 - by the mAbs anti-CD3 and anti-CD56 (Figure 2.3B). The FACSalibur is equipped with a mirror to allow the analysis of both APC and PerCP-Cy5.5.
2.4.1 Surface staining for phenotypic analysis
Mononuclear cells were aliquoted into FACs tubes for staining with different fluorochrome-conjugated monoclonal antibodies (Umeshappa et al.). These mAb were optimized according to the mAb manufacturer’s recommendations. To identify CD56+ T-cells both anti-CD3 mAb and anti-CD56 mAb were used to stain the cells (Figure 2.3B). Furthermore, several different mAb were also included as a third mAb for the purpose of identifying the phenotypic characteristic of CD56+ T-cells in CMV seropositive and seronegative healthy and kidney post-transplant patients (Table 2.1). Densty blot were used to measure the level of expression of the third marker on CD3+CD56+ cells after overlaying with the suitable isotype control antibody to allow for possible non-specific binding.
About 2–5 x 105 mononuclear cell aliquots were placed in FACs tubes, after which the cells were stained with mAb and placed at 4oC in the dark for 20 minutes. Following that, each tube was washed twice with sterile PBS for 10 minutes at 400g (1800 rpm) at 4oC to remove unbound fluorochrome-conjugated mAb. The cells were then resuspended in 300 µl PBS to analyze them by
FACSCalibur. The results were then analyzed by using WinMDI 2.9 software
(http://facs.scripps.edu/software.html; accessed 25/02/2014).
- 83 - Table 2.1 List of antibodies against surface markers and their suitable fluorochrome that have been used to phenotype CD56+ T-cells in healthy subjects and patients.
Monoclonal antibody Fluorochrome Manufacturer Catalogue no.
Anti-CD3 PerCP-Cy5.5 BD Pharmingen 560835
Anti-CD56 FITC BD Pharmingen 562794
Anti-CD4 PE BD Pharmingen 555347
Anti-CD8 PE BD Pharmingen 555635
Anti-CD57 PE BD Pharmingen 560844
Anti-CD58 PE BD Pharmingen 555921
Anti-CD94 PE BD Pharmingen 555889
Anti-NKG2C PE R&D Systems FAB138P-100
Anti-CCR7/CD197 PE eBioscience 12-1979
Anti-CD45RA PE BD Biosciences 66038
Anti-CD45RO PE BD Biosciences 64968
Anti-CD161 PE BD Pharmingen 556081
Anti-CD16 PE BD Biosciences 76180
Anti-CD127 PE BD Pharmingen 70755
Anti-CD28 PE BD Biosciences 75801
Anti-CD25 PE BD Biosciences 555432
Anti-CD62L PE BD Biosciences 65617
Anti-γδ-TCR PE BD Biosciences 562511
Anti-CD69 PE BD Biosciences 54963
Anti-TCR Vα7.2 PE Biolegend 351704
Anti-CD158a/h PE Miltenyi Biotec 130-092-684
Anti-CD158e PE Miltenyi Biotec 130-092-473
Anti-CD158f PE Miltenyi Biotec 130-096-200
- 84 - Figure 2.3 Gating strategies. These figures illustrate the gating strategy followed in this study to identify CD56+ T-cells and their phenotypic characteristics. A.
Density blot illustrates the gating of lymphocytes. B. This is a two colour density blot that shows the percentage of four different types of cell in the lymphocyte region after they were stained with anti-CD56 and anti-CD3. The percentage of
NK cells (CD56+CD3-) is 9.3%, the percentage of CD56+-T-cells (CD56+CD3+) is 8.4% and the percentage of T-cells (CD56-CD3+) is 66.4%. C. After gating
CD56+CD3+ cells, the level of expression of a third marker that is in this case
CD45RO was measured via density blot after compared to a negative control illustrated in D.
2.4.2 Functional analysis of CD56+ T-cells
2.4.2.1 Intracellular staining of CD56+ T-cells
For the purpose of studying the capability of CD56+ T-cells response to CMV,
PBMCs were intracellularly stained for the production of IFN-γ, TNF-α, IL-2,
Perforin, Granzyme-A and Granzyme-B when induced by CMV lysate.
Mononuclear cells were isolated according to 2.3.2. The cells were then washed
- 85 - twice using culture media RPMI 1640 (Sigma) containing (10% heat inactivated foetal calf serum + 2mM glutamine + 5000 unit Penicillin and Streptomycin). The cells were cultured in 48 well plates, in which some of the wells were treated as negative controls with culture medium alone, some wells were stimulated using
1:10 (prepared in the laboratory) or 200ng/ml CMV extract (The Native Antigen
Company, Upper Heyford) and for the positive control Staphylococcal enterotoxin
B (SEB) (Sigma Aldrich) was used. The cells were incubated overnight in a
o humidified 5% CO2 atmosphere at 37 C. The following day, 1 µl/ml of Brefeldin-
A (eBioscience, lot # 00-4506) was added to all wells of the plate and it was
o further incubated in a humidified 5% CO2 atmosphere at 37 C for 4 hours. Next, the cells were washed twice with culture media (RPMI 1640) for 10 min, 400g
(1800 rpm) and brake on. Then, 5 x 105 cells were harvested into FACs tubes and surface stained with anti-CD3 and anti-CD56 mAb using the same steps as in
Section 2.4.1. After that, the cells were washed twice with sterile PBS and fixed using 100 µl IC Fixation buffer (eBioscience) for 20 minutes in the dark at room temperature. The cells were then permeabilized by washing with 2 ml of 1 X
Permeabilization buffer (eBioscience) at 400g (1800 rpm) for 5 minutes with the brake on. Moreover, the permeabilization step was repeated three times. Each tube was stained with one of the intracellular antibodies (Table 2.2), and the tubes were incubated for 30 minutes at room temperature in the dark. Subsequently, the tubes were washed by 1X permeabilization buffer and further resuspended in 300 μl
PBS. The staining was analyzed using the FACSCalibur and subsequently analyzed by WinMDI 2.9 software. The production of these cytokines and expression of cytotoxic mediators by CD56+ T-cells was measured by the same procedure explained in Figure 2.3.
- 86 - IC mAb Manufacturer fluorochrome Isotype Surface mAb
Anti-IFN-γ Mouse IgG1 k CD3-FITC, CD56- BD Pharmingen PerCP-Cy5.5 560704 550795 APC
Anti-TNF-α Mouse IgG1 k CD3-FITC, CD56- BD Pharmingen PerCP-Cy5.5 560679 550795 APC
Anti-IL-2 Mouse IgG1 k CD3-FITC, CD56- BD Pharmingen PerCP-Cy5.5 560708 550795 APC
Anti- Perforin Mouse IgG2b CD3-FITC, CD56- PE eBioscience 12-9994 12-4732 APC
Anti-Granzyme A IgG1, k CD3-FITC, CD56- BD Pharmingen PE 78238 400137 APC
Anti-Granzyme B IgG1, k CD3-PerCP-Cy5.5, Biolegend FITC 515403 400137 CD56-PE
Table 2.2 List of intracellular mAb that have been used in these experiments.
Different types of surface mAb were used with IC mAb.
2.4.2.2 Degranulation assay of CD56+ T-cells; detection by LAMP-1
(CD107a) expression
2.4.2.2.1 MHC negative targets
K562 is a human erythroleukaemic cell line that lacks the expression of MHC-I that make them a susceptible target for NK cell detection. K562 cells were maintained in culture media, RPMI 1640 media, supplemented with 10% FBS, 2 mM L-glutamine, 50 IU/ml penicillin and the cells were incubated in a humidified
ᴼ 5% CO2 atmosphere at 37 C.
- 87 - NK cell cytotoxicity is mediated by two pathways, cross-linking of NK cell surface ligands such as FAS-L and the target cell death receptors expressed on their surface such as FAS. Additionally, the second pathway is dependent on lack of inhibitory signals via KIRs and is mediated via the releasing of perforin and granzyme (L. Wang et al. 2012, Ozoren and El-Deiry 2003, Trapani 2012).
CD107a is a marker for NK cell and CD8+ T-cell degranulation as part of the perforin-dependent cytotoxic process therefore it was investigated whether
CD56+ T-cells express this marker after their induction by CMV lysate and K562, and if there were any significant difference between healthy CMV seropositive and seronegative subjects.
2.4.2.3.2 PBMC stimulation by K562 cells and CMV lysate
PBMCs were isolated according to section 2.3.2. Samples from healthy adults participated in this experiment and their CMV sero-status was detected by ELISA as identified in section 2.1.3. The stimulation of PBMCs by K562 and CMV lysate was carried out in 48 well culture plates, at a ratio of 1:2 (PBMCs: K562); the cells numbers were counted according to section 2.3.3. Unstimulated PBMCs were used as a negative control and the levels of CD107a detected in the negative control were subtracted from those measured after K562 and CMV lysate stimulation. PBMCs were stimulated by K562 and CMV lysate for four hours in a
o humidified 5% CO2 atmosphere at 37 C, then 1 µl/ml Brefeldin A was added to the culture and the plate was left in the incubator for about 4 hours. The cells then were washed twice with sterile PBS in FACs tubes at 400g (1800 rpm) for 10 minutes. Following that, the cells were pelleted and stained with anti-CD3 PerCP-
- 88 - Cy5.5, anti-CD56 FITC and Anti-CD107a PE (555801, Isotype Mouse IgG1, k
555749) (the three antibodies were purchased from BD Pharmingen) and left on ice for 20 minutes, then the cells were washed with PBS and immediately analysed by BD FACSCalibur. The analyses were performed by using WinMDI
2.9 as mentioned in Figure 2.3.
2.4.2.4 Proliferation assay of CD56+ T-cells
2.4.2.4.1 Carboxy-Fluorescein Succinimidyl Ester (CFSE) assay
CFSE dye is frequently used to determine the capability of lymphocytes to proliferate and generate daughter cells in response to specific antigen. Each new generation of cells retains half the amount of CFSE of the parent cells. CFSE is integrated into the cells in a colourless and non-fluorescent condition; emission of
CFSE is triggered when intracellular esterases cleave the acetate group. The parent cells preserve the dye, moreover, during mitosis and proliferation to daughter cells the dye is halved with each new generation. The list of reagents used in this test is identified in Table 2.3.
- 89 - Reagent Manufacture Conjugated fluorochrome
CFSE Molecular ProbesTM Invitrogen fluorescein
Anti-CD3 BD Pharmingen PerCP-Cy5.5
Anti-CD56 BD Pharmingen APC
Table 2.3 Reagents that were used in proliferation assays of CD56+ T-cells.
2.4.2.4.2 PBMC CFSE labelling
The isolation and counting of the cells were performed as identified in sections
2.3.2 and 2.3.3 respectively. The labelling process was performed in universal tubes with sterile PBS to remove the incidence of generating backgrounds. CFSE was purchased from Molecular ProbesTM Invitrogen. At a concentration of 1 x 106
PBMCs were resuspended in 10ml of pre-warmed sterile PBS mixed with 5μM
CFSE in dimethyl sulphoxide, which was further incubated for 8 minutes in a
o humidified 5% CO2 atmosphere at 37 C. Consequently, the reaction was stopped by the addition of 5 volumes of ice cold PBS and centrifugation at 400g (1800 rpm) for 10 minutes. The washing step was repeated twice to remove any unbound CFSE. The cells were resuspended in culture medium, counted and 1 x
106 cells/well placed into 48-well culture plates.
2.4.2.4.3 PBMC stimulation by CMV lysate and SEB
To detect the ability of CD56+ T-cells to proliferate in response to CMV antigens,
PBMCs labelled with CFSE were stimulated by 2 µl CMV lysate for 7 days.
- 90 - Moreover, 1 µl SEB per 1X106 cells/ml was used as a positive control and
PBMCs labelled with CFSE were cultured in medium alone as a negative control.
At day 7, PBMCs were harvested into FACs tubes, which were further centrifuged twice in sterile PBS at 400g (1800 rpm) for 10 minutes. PBMCs were stained for surface markers to identify CD56+ T-cells; CFSE is FITC conjugated therefore anti-CD3-PerCP-Cy5.5 and anti-CD56-APC were used for staining and cells were incubated for 20 minutes on ice in the dark. PBMCs were washed twice with sterile PBS to remove any unbound mAb and resuspended in 400 µl of PBS. They were then analysed in a FACSCalibur, usingWinMDI 2.9 software for analysis.
The gating strategy used in the analysis of proliferated cells is identified in Figure
2.4.
Figure 2.4 Proliferation gating strategies. This figure illustrates the gating strategy followed in this study to identify proliferated cells. A. This figure shows the method followed to gate lymphocytes. B. This figure identifies the method of gating both NK cells and CD56+ T-cells. C. This figure shows the method of analysing proliferated cells at day 7. Cells to the left of the threshold had reduced
CFSE staining and were regarded as having proliferated.
- 91 - 2.4.2.5 CD56+ T-cell CMV specificity detection
T-cells are the essential part of cell-mediated immune responses toward antigens.
CD8+ cytotoxic T-cells (CTL) recognize antigenic peptides bound to major histocompatibility complex class-I (MHC-I), which is expressed on all nucleated host cells and there are three HLA class-I loci, A, B and C; however, MHC-II is expressed on antigen presenting cells (APC). When host cells are under stress such as virus infection, the host cells start to degrade the foreign antigen to smaller peptide fragments and begin to present it on MHC-I to allow the recognition by CD8+ cytotoxic T cells of the host cell’s abnormal status. CTL initiate the release of cytotoxic mediators such as perforin and granzyme which induce the abnormal host cells to undergo apoptosis.
To detect and enumerate the specific CD8+ T-cells recognizing target antigens,
MHC-I Pentamers or tetramers can be used. These multivalent reagents, made up of several soluble class I MHC molecules bound to specific antigenic peptide, have sufficiently high avidity to bind to T cells with antigen receptors specific for the peptide. In this study CD56+T-cell specificity toward CMV peptides were investigated by using Pro5 MHC-I Pentamers (Proimmune, Oxford). The first step was to identify possible candidates who were CMV seropositive and of the appropriate HLA tissue type. In this study, three of the common European alleles were studied: HLA-A*02, HLA-B*07, and HLA-B*08. After that, the second step was to staining the cells by CMV Pro5 pentamers to detect their specificity toward
CMV peptides. In Table 2.4, the list of reagent s used in this test is illustrated.
- 92 - 2.4.2.5.1 HLA tissue type
PBMCs from healthy donors were isolated according to section 2.3.2, centrifuged twice in PBS and aliquoted into FACs tubes. Monoclonal antibodies against HLA-
A2, B7 and B8 (Table 2.5) were used to stain PBMCs for 20 minutes and left on ice in the dark. Both anti-HLA-A2 and anti-HLA-B7 are FITC conjugated therefore after the end of staining period the tubes were washed and left on ice.
However, anti-HLA-B8 is conjugated to biotin therefore the tube was washed twice and stained with Streptavidin conjugated to FITC and left on ice for a further 20 minutes, then the tube was washed. The tubes were analysed by
FACSCalibur, and gating strategy is explained in Figure 2.5.
- 93 -
Figure 2.5 Tissue typing gating strategy. These figures explain the gating strategy followed to identify the tissue type of CMV seropositive donors.
Lymphocytes were gated in the scatter density plot and the tissue type was identified by using histograms of FITC staining. Representative positive and negative donors are shown the histograms are organised from the top as HL-A2 positive, HLA-B7 positive, HLA-B8 positive and the last one as HLA-B8 negative.
- 94 - PEPTIDE EPITOPE Conjugated Reagents Manufacture SEQUENCE ORIGIN fluorochrome
Anti-HLA A2 AbD Serotec - - FITC
Anti-HLA B7 AbD Serotec - - FITC
Anti-HLA B8 AbCam - - conjugated to
biotin
Streptavidin AbD Serotec - - FITC
A*02:01 NLVPMVATV HCMV APC Proimmune pp65 495-
504
B*07:02 Proimmune TPRVTGGGA HCMV APC
M pp65 417-
Pro5 Pentamers 426
B*08:01 Proimmune QIKVRVDMV HCMV APC
IE1 88-96
Anti-CD8 clone Proimmune - - FITC
LT8
Anti-CD56 Coulter, - - PE
Luton
Table 2.4 Reagents used in pentamer labelling.
- 95 - 2.4.2.5.2 CD56+ T-cells Pro5 Pentamers staining
PBMCs from CMV seropositive healthy donors who were tissue type positive for
HLA-A2, HLA-B7 and HLA-B8 were isolated and washed according to section
2.3.2. PBMCs were washed twice with sterile PBS then 1 x 106 cells/ml were allocated into FACs tubes for staining. The cells were re-suspended in 50 μl sterile PBS before staining the cells using the appropriate pentamer. Pro5 pentamers were centrifuged at 14,000g at 4 oC for 10 minutes. The purpose for this high-speed spin is removing any protein aggregates that reside in the vials, which can result in background and non-specific binding. Moreover, it is essentially to avoid pipetting the pentamers from the bottom of the vial. About 10
µl were used from each pentamer vials to stain PBMCs for 10 minutes shielded from light and at room temperature. The tubes were washed twice in 2 ml sterile
PBS at 400g (1800 rpm) for 10 minutes, supernatants from all tubes were discarded, tubes were stained with anti-CD8-FITC and anti-CD56-PE and the tubes left on ice shielded from light. Following the manufacturer’s recommendation, anti-CD19-PerCP-Cy5.5 was included to reduce the background due to the nonspecific binding of the pentamers to B cells. The tubes were washed twice, re-suspended in 400 μl PBS and the analysis was performed using a
FACSCalibur and the data analysed with winMDI 2.9, gating for CD19- CD8bright cells.
2.4.2.6 Identifying CD56+ T-cell gene expression profile in response to CMV
This experiment was designed to identify the gene expression profile of CD56+ T- cells in response to CMV lysate. The first step was to select four CMV
- 96 - seropositive candidates, then obtain enough peripheral blood to obtain sufficient
CD56+ T-cells when sorting this type of cells from PBMCs. Next PBMCs were isolated from the blood samples according to section 2.3.2, after that, cells were counted according to section 2.3.3 to guarantee an adequate number of cells for the sorting step. The cells were divided into two groups, the first to be stimulated with CMV lysate and cultured overnight in a humidified 5% CO2 atmosphere at
37oC, and the second group of cells to be cultured without stimulation overnight
o in a humidified 5% CO2 atmosphere at 37 C. The next day the two groups of cells were washed twice with sterile PBS at 400g (1800 rpm) for 10 minutes. Next, the cells were stained with anti-CD3 PerCP-Cy5.5 and anti-CD56-PE for 30 minutes and room temperature in the dark; following that, the cells were washed with sterile PBS twice and aliquoted in RPMI 1640 culture media to prepare them for the cell sorting step.
2.4.2.6.1 CD56+ T-cell sorting
CD56+ T-cell sorting was performed by Dr. Stuart Marshall-Clarke and Mrs.
Natalie O’Hara, in the Technology Directorate FACS facility, Faculty of Health &
Life Sciences. A FACSAria IIIu was used for cells sorting. In brief, the cells were stained by anti-CD56-PE and anti-CD3+-PerCP-Cy5.5 and the cells were sorted according to the mAb conjugated fluorochrome. Three groups of cells were sorted into RPMI1640 culture media: CD56+CD3- cells, CD56+CD3+ cells and CD56-
CD3+ cells. After that, the cells were washed with sterile PBS at 400g (1800 rpm) for 10 minutes, and prepared for immediate RNA extraction.
- 97 - 2.4.2.6.2 RNA extraction protocol
After sorting of the cells, RNA extraction from sorted CD56+ T-cells was done instantly using an RNeasy mini kit (Qiagen). At the beginning, concentrated RPE buffer was diluted with 4 volumes of 96% ethanol to prepare the working solution. The samples were adjusted to a volume of 1ml by the addition of 100 μl
RNase-free water plus 600 μl of buffer RLT and the mixtures resuspended thoroughly. Next, 250 μl of 96% ethanol was added and each sample mixed thoroughly by pipetting. Then, 700 μl from each sample was transferred to an
RNeasy mini spin column placed into 2 ml collection tube and centrifuged for 15 seconds at 8000g and the flow throughdiscarded. RNeasy columns were placed again into 2 ml collection tubes followed by the addition of 500 μl RPE buffer into the RNeasy mini spin columns and centrifuged for 15 seconds at 8000g.
Again the flow through was discarded from all tubes and again 500 μl RPE buffer added into RNeasy mini spin columns, however this time the tubes were spun for
2 minutes at 8000g. Next, the flow-through was discarded and RNeasy mini spin columns were centrifuged at 8000g for 1 minute to remove any traces of the ethanol. Then, RNeasy mini-spin columns were placed into 1.5 ml eppendorf tubes and 30 μl RNase-free water was added directly and carefully onto the gel- membrane of the RNeasy mini-spin column without touching the gel-membrane and centrifuged at 8000g for 1 minute. The eluted RNA was measured by
NanoDrop and placed in a -80oC freezer.
- 98 - 2.4.2.6.3 DNase-treatment of RNA
This step was to remove any residual DNA remaining after the isolation of RNA.
This was done by a RNase-Free DNase set (Qiagen, lot 79254). At the start, to prepare DNase I the stock was dissolved in 550μl RNase-free water using an
RNase free needle, then the vials were inverted to mix the solution, which was stored at -20 oC. DNase-I incubation mix was prepared by diluting 10μl of DNase-
I stock solution in 70μl RDD buffer to constitute 80μl.
The RNA eluate volume was brought up to 100μl by Rnase-free water. Then, about 350μl RLT buffer was added and mixed thoroughly. Next, 250μl 96%-
100% ethanol was added to dilute the RNA and mixed by pipetting gently. The whole quantity (about 700μl) was transferred to RNeasy-mini spin columns that were placed in 2 ml collection tubes, and the mixture centrifuged at 8000g for 15 seconds. The flow-through was discarded and 350 μl RW-1 buffer was added and centrifuged at 8000g for 15 seconds. After this, the 80 μl incubation mix was added directly onto the membrane of the column and left at room temperature for
15 minutes. This was followed by the addition of 350μl RW-1 buffer onto the membrane which was centrifuged at 8000g for 15 seconds, discarding the flow- through. About 500μl of RPE diluted with ethanol was added to the column and centrifuged for 15 seconds at 8000g, discarding the flow-through. 50μl RPE buffer and centrifugation for 2 minutes at 8000g. Then, the RNeasy mini-spin columns were placed into 1.5 ml eppendorfs and 30 μl Rnase-free water added to elute the RNA, the tube was centrifuged for 1 minute at 8000g, then RNA was measured by NanoDrop and stored at -80 oC.
- 99 - 2.4.2.6.4 One-Colour Microarray-Based Gene Expression
Microarrays were used to provide a comprehensive study of the gene expression profile of CD56+ T-cells in response to CMV stimulation. Agilent Human v2, 8 x
60K arrays were purchased from Agilent Technologies. The experimental work with the arrays was done by Dr. Lucille Rainbow, Centre for Genomic Research,
IIB. In brief, 8 total RNA samples (free from genomic DNA) were cleaned with
Ampure RNA beads and quantified by NanoDrop. The quality of RNA was assessed using Agilent 2100 Bioanalyser RNA 6000 Pico chips. Samples were prepared for hybridisation onto Agilent Human genome v2 arrays. Approximately
50ng of total RNA was used as initial input, and the Agilent Low-Input
Amplification Kit was used for One-Colour target preparation for microarray expression analysis according to manufacturer’s instructions. 600ng of Cy3 labelled aRNA (amplified RNA) was fragmented, and loaded onto the array which was hybridised for 17 hours at 650C and 10rpm in an Agilent hybridisation oven.
Following hybridisation the arrays were washed according to the manufacturer’s instructions and scanned using the Agilent Scanner (Figure 2.6).
Downstream analysis of Agilent microarray data was processed by using Agilent technology software GeneSpring GX 9.0.
- 100 -
Figure 2.6 Microarrays workflow flow-charts. The workflow flow-charts is adapted from Agilent Technologies. The steps from cDNA synthesis to feature extraction were processed by Dr. Lucille Rainbow, Centre for Genomic Research,
IIB.
2.4.2.6.5 Data Analysis of microarrays using GeneSpring GX9.0
The company recommended using Agilent technology default software for microarrays results; therefore, GeneSpring GX9.0 was used for this purpose. The software was developed by the company technology department and has the array design information and biological information about different entities of their array types. Data from Agilent single colour Expression data test were saved in text format (.txt). Steps for analysing the data are described as follows:
The text files containing microarrays results are loaded into GeneSpring
GX9.0.
GeneSpring GX9.0 will automatically extract the gene expression data and
perform standard normalisation and transformations by default, and these
- 101 - are threshold all values to 5, median shift normalisation to the 75
percentile and apply a baseline transformation by using the median of all
samples.
Files were divided into two groups according to the conditions and they
were the stimulated group and the un-stimulated group.
The two conditions of groups were compared together by using a
Moderated T-test with cut-off of 0.05.
Multiple testing corrections were applied to remove any false-positive
significantly expressed genes. The software default method for multiple
testing corrections was applied and it is called Westfall and Young
Permutation with cut-off 0.05 and fold changes was set to equal or more
than 2 folds change. Next, the significantly expressed genes were exported
into an excel file.
The results were clustered by hierarchical clustering to build a
phylogenetic tree to cluster genes together that has similar properties. This
step was also performed by GeneSpring GX9.0.
2.4.2.6.6 Validation of Microarrays results
RT-PCR analysis of gene expression was used for the validation of microarray experiments. Three genes of interest were picked for RT-PCR analysis: ISG-15,
USP-18 and IFN-γ. Moreover, for positive control or housekeeping gene L-32 was used. Both steps 2.4.2.6.7 and 2.4.2.6.8 were done with the help of Dr. Brian
Flanagan in Department of Women’s and Children’s Health, ITM, University of
Liverpool at Alder Hey Children’s Hospital.
- 102 - 2.4.2.6.7 Synthesis of cDNA (Reverse Transcription)
RNA samples were placed on ice to avoid degradation.. High Capacity cDNA
Reverse Transcription kit (Invitrogen catalogue number 4368814, UK) was used to reverse transcribe cDNA. First, 2 µl RT buffer, 2 µl dNTP nucleotides and 2 µl random primers were added into RNase-free eppendorfs. Then, 5 µl RNA
(approximately 1µg) followed by 1 µl reverse transcriptase enzyme (stored at -
200C) were added. The mixture was mixed gently by pipetting and incubated for 1 hour at 37 0C in an AccuBlock™ Digital Dry Bath. After that, cDNAs were diluted with 180 µl sterile water and stored at -800C.
2.4.2.6.8 RT-PCR gene expression analysis
Quantitative PCR analysis of the three candidate genes: IFN-γ, ISG-15 and USP-
18 were carried out by using RT-PCR. TaqMan® probes for gene expression assays were obtained from Applied Biosystems, UK. These TaqMan probes and the master mix details are identified in Table 2.5. The reaction was processed in an Applied Biosystems 7300 Real PCR system, and the software was also 7300
Systems software. The principle behind TaqMan probes is explained as follows, an oligonucleotide probe has a quencher on the 3′ end that prevent the fluorescent dye attached to the 5′ end from emitting, which is also called reporter dye.
However, when the target sequence is bound by the probe this will lead to cleavage of the reporter dye on the 5′ end by 5′ nuclease activity of the DNA polymerase. With the expansion of more complementary sequences there is cleavage of more reporter dye molecules from the TaqMan probes leading to the generation of more fluorescence. An increase in fluorescence is proportional to copy number of the target nucleic acid. Samples contain high levels of cDNA are
- 103 - detected before those containing smaller amounts. This results in a lower Ct, the cycle number at which fluorescence signal exceeds an arbitrary threshold. Hence, the Ct value, or cycle number is inversely proportional to the initial amount of target DNA. TaqMan probes are specific to their target sequence which makes them a better choice than conventional SYBR green methods.
Reagent Lot number UniGene ID
L32 TaqMan Probe 1093359 Hs.00388301_M1MRP
IFN-γ TaqMan Probe 666419 Hs.0989290_g1
ISG-15 TaqMan Probe 4331182 Hs00192713_m1
USP-18 TaqMan Probe 4331182 Hs00276441_m1
TaqMan Gene-Expression 1304177 - Master Mix
Table 2.5 Reagents used in RT-PCR analysis of gene expression. All the reagents were from Applied Biosystems, L32 TaqMan probe, IFN-γ TaqMan probe and TaqMan Gene-Expression Master Mix were provided by Dr. Brian
Flanagan.
The reaction mix for each probe was prepared as follow; first, four sterile eppendorfs were allocated in which each eppendorf was used for a single Taq-
Man probe. The quantity of probes and master mix were calculated by this method
(12.5 µl of master mix X number of wells) and for the probes (1.25 µl TaqMan probe X number of wells). These quantities were aliquoted into each well and
11.25 µl of cDNA of each sample were aliquoted into each of the 96 wells.
- 104 - The thermal cycle conditions applied for the reaction was the same as 7300
Applied Biosystems software default settings and the cycle is illustrated in Table
2.6.
Stage Steps Time and Temperature
Activation AmpErase® UNG 2 minutes 50ᴼC
Initial prior PCR Hot Start DNA Polymerase 10 minutes 95ᴼC activation
Melt 15 seconds 95ᴼC
PCR cycles
Anneal/Extend 1 minute 60ᴼC
Table 2.6 RT-PCR thermal conditions. The thermal condition applied to the
TaqMan gene expression assay was the same as 7300 Applied Biosystems software default, and the number of the cycles was 40 cycles. Activation
AmpErase® UNG principle was to prevent re-amplification of PCR wastes from previous experiments.
2.4.2.6.9 Protein expression analysis
Two proteins were studied futher to validate microarray data, also, to compare between CMV healthy groups.
- 105 - 2.4.2.6.9. a CD38 surface staining of CD56+ T-cells
Changes in CD56+ T-cell surface expression of CD38 in response to CMV lysate were further analyzed. Mononuclear cells were isolated according to step 2.3.2.
The cells then were washed twice by using culture media RPMI 1640 (Sigma) containing (10% heat inactivated foetal calf serum + 2mM glutamine + 5000 unit
Penicillin and Streptomycin). The cells were cultured in 48 well plates, in which some of the wells were treated as negative controls and other wells were stimulated using of 1:10 or 200 ng/ml (The Native Antigen Company, Upper
Heyford) CMV lysate. The cells were incubated for three days in a humidified 5%
o CO2 atmosphere at 37 C. Each day, 1 µl/ml of cells Brefeldin-A (eBioscience) was added to some of the wells of the plate and it was further incubated in a
o humidified 5% CO2 atmosphere at 37 C for 4 hours. Next, the cells were washed twice with culture media (RPMI 1640) for 10 min, 400g (1800 rpm) and brake on.
Then, 5 x 105 cells were harvested into FACs tubes and surface stained by anti-
CD3, anti-CD56, anti-CD38 PE mAb (Biolegend, 356604) using the same steps as in section 2.4.1. The staining was analyzed using a FACSCalibur and subsequently analyzed by WinMDI 2.9 software. The expression of CD38 by
CD56+ T-cells was measured by the same procedure explained in Figure 2.3.
2.4.2.6.9. b ISG-15 intracellular staining of CD56+ T-cells
PBMCs were intracellularly stained for the production of ISG-15 when induced by CMV lysate. Mononuclear cells were isolated according to 2.3.2. The cells were then washed twice using culture media RPMI 1640 (Sigma) containing (10% heat inactivated foetal calf serum + 2mM glutamine + 5000 unit Penicillin and
- 106 - Streptomycin). The cells were cultured in 48 well plates, in which some of the wells were treated as negative controls with culture medium alone, some wells were stimulated using 1:10 (prepared in the laboratory) or 200ng/ml CMV extract
(The Native Antigen Company, Upper Heyford). The cells were incubated
o overnight in a humidified 5% CO2 atmosphere at 37 C. The following day, 1
µl/ml of Brefeldin-A (eBioscience, lot # 00-4506) was added to all wells of the
o plate and it was further incubated in a humidified 5% CO2 atmosphere at 37 C for
4 hours. Next, the cells were washed twice with culture media (RPMI 1640) for
10 min, 400g (1800 rpm) and brake on. Then, 5 x 105 cells were harvested into
FACs tubes and surface stained with anti-CD3 and anti-CD56 mAb using the same steps as in Section 2.4.1. After that, the cells were washed twice with sterile
PBS and fixed using 100 µl IC Fixation buffer (eBioscience) for 20 minutes in the dark at room temperature. The cells were then permeabilize by washing with 2 ml of 1 X Permeabilization buffer (eBioscience) at 400g (1800 rpm) for 5 minutes with the brake on. Moreover, the permeabilization step was repeated three times.
The cells were blocked by 10 µl goat serum (Sigma-Aldrich, G9023) fot 10 minutes and room temperature, this followed by washing at 400g (1800 rpm) for
10 minutes with the brake on. The cells were labeled by 1 µl ISG-15 polyclonal antibody (Thermo Fisher, PA5-17461), and the tubes were incubated for 30 minutes at room temperature in the dark. Subsequently, the tubes were washed by
PBS at 400g (1800 rpm) for 10 minutes. The cells were labeled by 1 µl secondary antibody Pierce Goat anti-Rabbit IgG, H+L-FITC (Thermo-scientific, 31583), and the tubes were incubated for 20 minutes at room temperature in the dark.
Subsequently, the tubes were washed by PBS at 400g (1800 rpm) for 10 minutes.
- 107 - The staining was analyzed using the FACSCalibur and subsequently analyzed by
WinMDI 2.9 software. The production of these cytokines and expression of cytotoxic mediators by CD56+ T-cells was measured by the same procedure explained in Figure 2.3.
- 108 -
Chapter 3
Frequency and phenotype of
CD56+ T-cells in CMV seropositive and seronegative healthy subjects
- 109 - 3. Frequency and phenotype of CD56+ T-cells in CMV seropositive and seronegative healthy subjects
3.1 CMV sero-status by applying IgG ELISA assay
Sera from healthy subjects were analysed for the presence of IgG antibodies to
CMV. The results were calculated as follows: the mean of the cut off wells’ absorbance value was calculated according to this equation: [the sum of cut off values / 2]. The sample was considered clearly positive if the mean of the antibody titres in two wells was more than 10 % above the cut off mean absorbance value. The sample was considered clearly negative if the mean of the antibody titres in two wells was lower than 10 % below the cut off mean absorbance value (Table 3.1). When the mean of the antibody titrers was equal or less than 10 % above or below the cut off mean this was considered a grey zone and samples should be repeated and if they gave the same value again the result was considered negative, however, no sample has fallen between this grey zone values.
- 110 - 1 2 3 4 5 6 7 8
A 0.062 2.164 0.500 0.170 0.134 1.654 0.100 0.106
B 0.075 1.863 0.603 0.093 0.123 1.722 0.092 0.125
C 0.074 0.144 1.021 1.607 1.555 2.299 1.410 1.047
D 0.406 0.121 1.024 1.654 1.509 1.722 1.139 1.159
E 0.361 1.126 0.217 0.992 0.073 0.166 1.033 1.396
F 0.691 1.204 0.230 0.966 0.079 0.292 1.139 1.373
G 0.138 0.142 1.582 1.213 0.076 0.106 1.410 0.093
H 0.145 0.132 1.579 1.231 0.080 0.108 1.422 0.114
Table 3.1 IgG ELISA detection. This table illustrates the CMV sero-status of 29 healthy donors. Well A1 is the substrate blank. B1 and C1 are the negative control. D1 and E1 are the cut-off controls and their mean is 0.383. F1 is the positive control. Samples were allocated in duplicates, and any mean value more than 0.383+ 0.0383 (10%) was considered CMV seropositive. In addition, any sample less than 0.383- 0.0383 (10%) was considered CMV seronegative. No sample fell in the grey zone.
.
- 111 - 3.2 Frequency of CD56+ T-cells in CMV seropositive and seronegative healthy subjects
3.2.1 Introduction
CD56+ T-cells numbers have previously been studied in groups of patients with different clinical conditions. Generally, the numbers are reduced in some autoimmune diseases such as psoriasis and rheumatoid arthritis when the relative numbers of CD56+ T-cells were compared to healthy controls (Koreck et al. 2002,
Yanagihara et al. 1999). In addition, in patients infected with hepatitis E the numbers of CD56+ T-cells were also reduced (Srivastava et al. 2008). Moreover, chronic hepatitis C infected patients have reduced levels of CD56+ T-cells and they are also reported to be vulnerable to hepatic carcinoma (Kawarabayashi et al.
2000). Also, the numbers are reduced in smokers and patients with chronic obstructive pulmonary disease (Urbanowicz et al. 2009).
On other hand, CD56+ T-cell numbers have been reported to be expanded in infections conditions such as malaria, HIV and hepatitis B (Watanabe et al. 2003,
Paiardini et al. 2005, Barnaba et al. 1994). These cells were also expanded in lung cancer patients with both non-small-cell and small-cell lung cancer when the levels were compared to healthy blood donors (Al Omar et al. 2012).
The proportion of CD56+ T-cells was measured by collecting PBMCs from healthy donors, then double staining with anti-CD3 and anti-CD56 monoclonal antibodies. Relative number of CD3+CD56- T-cells and NK cells(CD56+CD3-) in CMV seropositive and CMV seronegative subjects were also measured and compared. During these analyses, the minimum number of accepted events using
- 112 - the FACS-Calibur was 20,000 events. 68 healthy donors were used in these analyses, of which 34 were CMV seropositive and 34 CMV seronegative.
The statistical analysis was performed by using Graph-Pad Prism and applying a
Paired T-test and the results are shown in a vertical scatter plot.
3.2.2 Aims
The aim of this section of the study was to compare the relative levels of CD56+
T-cells in CMV seropositive and seronegative healthy individuals. Moreover, the comparison was made by using two different methods, the first in relation to the total number of lymphocytes and the second to the total number of CD3+ T-cells only. Both age and gender effects were analysed and compared to test for any significant differences. Following that, relative levels of CD3+CD56- T-cells cells and CD3-CD56+ NK cells were compared between CMV seropositive and seronegative healthy individuals.
3.2.3 Results
3.2.3.1 CD56+ T-cells relative numbers in healthy CMV seropositive and seronegative subjects
Flow cytometric analysis of CD56+ T-cell population was applied to identify the relative numbers of these cells in healthy individuals. PBMCs were stained by anti-CD3 and anti-CD56 and expressed as a density plot. Numbers expressed as
- 113 - percentages of total lymphocytes and T-cells (Figure 3.1) were collected into
GraphPad prism; the two groups were compared by using a Paired T-test.
In Figure 3.2, the percentage of CD56+ T-cells was calculated from total lymphocytes and T-cells. The mean value of CD56+ T-cells in CMV seronegative subjects was 3.292%, and for CMV seropositive subjects the mean was 8.380%.
When these two groups were compared by applying a Paired t-test, there was a significant difference between them (P= <0.0001). This result indicates that
CD56+ T-cells were expanded to higher levels in healthy chronic CMV seropositive individuals compared to the level of CD56+ T-cells in CMV seronegative individuals.
- 114 -
Figure 3.1 CD56+ T-cells level in representative CMV + and CMV- subjects.
Density plots illustrate double staining of PBMCs by anti-CD3+ and anti-CD56+ with very clear-cut identification of these cells. Percentage of these cells was calculated from total lymphocytes and T-cells. The upper left density plot shows the percentage of CD56+ T-cells in a CMV seronegative healthy donor (1.8%).
The upper right density plot shows the percentage of CD56+ T-cells in a CMV seropositive healthy donor (8.5%). The lower left density plot shows the percentage of CD56+ T-cells from total T-cells in a different CMV seronegative healthy donor (1.6%). The lower right density plot shows the percentage of
CD56+ T-cells from total T-cells in a CMV seropositive healthy donor (21.4%).
CMV seronegative or seropositive plots were not obtained from the same donor. It is clear from the plots that all CD56+ T-cells are CD56+ dim and not bright.
- 115 - < 0.0001 < 0.0001 20 30 27 24 15 21 18 10 15 12 % T-cells 9 5 % Lymphocytes 6 3 0 0 CMV- CMV+ CMV- CMV+ CD3+CD56+ CD56+CD3+
Figure 3.2 Percentages of CD56+ T-cells in CMV+ and CMV- subjects. These vertical scatter plots compare the percentage of CD56+ T-cells obtained from total lymphocytes and CD3+ T-cells between CMV– and CMV+ subjects. The mean of
CD56+ T-cells in CMV seronegative subjects was 3.292± 0.3278, and in CMV seropositive subjects the mean was 8.380± 0.8006 (P <0.0001). Both scatter plots show clearly that CD56+ T-cells are expanded in CMV seropositive compared to
CMV seronegative subjects.
3.2.3.2 Relative numbers of CD56-CD3+ cells in healthy CMV seropositive and seronegative subjects
Relative numbers of total CD3+CD56- T-cells in CMV seropositive versus seronegative subjects were also evaluated and compared. CD3+ T-cells were compared by two different methods, the first method was to calculate CD3+ T- cells from the total lymphocytes and the second method was to calculate CD3+ T- cells from total cells expressing CD3 after excluding CD56+ T-cells (Figure 3.3).
- 116 - CD3+ CD3+ < 0.0001 100 0.0516 100 80 90 60
80
40 % CD3+CD56- % Lymphocytes % 20 70
0 60 CMV- CMV+ CMV- CMV+
Figure 3.3 Percentages of CD56- T-cells in CMV+ and CMV- subjects.
Frequency of CD56- T-cells was compared between CMV seropositive and seronegative healthy individuals. Both scatter plots showed more CD3+CD56-T- cells in CMV seronegative than seropositive subjects. In the left plot,
CD3+CD56- cells were evaluated from the total lymphocytes, the mean of
CD3+CD56- T-cells in CMV seronegative subjects was 59.98% ± 2.034, which was more than the mean of CD3+CD56- T-cells in CMV seropositive subjects
53.61% ± 1.475, although this was not quite statistically significant (P = 0.0516).
In the right scatter plot CD3+ CD56- cells were evaluated from the total CD3+ T- cells after subtracting CD56+ T-cells. The mean of CD3+CD56- T-cells in CMV seronegative subjects was 95.25% ± 0.4473, which was more than the mean of
CD3+CD56- T-cells in CMV seropositive subjects 83.84% ± 0.7806 which was significant (P<0.0001). The number of donors was 68 healthy individuals of whom 34 were CMV seronegative and 34 were CMV seropositive.
- 117 - It is clear from Figure 3.3 that a higher percentage of CD56- T-cells was present in CMV seronegative than CMV seropositive healthy individuals.
3.2.3.3 Relative numbers of NK cells (CD56+CD3-cells) in healthy CMV seropositive and seronegative subjects
Relative numbers of total CD56+CD3- NK cells in CMV seropositive versus seronegative subjects were also evaluated and compared. NK cells were found to be present in roughly equal proportions in CMV seronegative and CMV seropositive healthy individuals (Figure 3.4).
CD56+CD3- 0.8388 30
20
%
10
0 CMV- CMV+
Figure 3.4 Frequencies of CD56+CD3-NK cells in CMV+ and CMV- subjects.
This figure shows the frequency of CD56+CD3-NK cells in CMV seropositive versus seronegative individual donors. There was no significant difference between numbers of NK cells present in CMV- healthy individuals 15.14± 0.9879 compared with CMV seropositive 14.59± 0.7497 (P = 0.7873). 68 healthy donors were used of which 34 were CMV seronegative and 34 CMV seropositive.
- 118 - 3.2.3.4 CD56+ T-cell levels compared in different age groups
Relative numbers of CD56+ T-cells were evaluated in three different age groups:
20-34, 35-49 and >50 years. Relative CD56+ T-cell numbers were compared between each of these groups, then all groups were analysed by using One-way analysis of variance (Figure 3.5). It is clear that CMV seropositive subjects in all age groups have significantly expanded numbers of CD56+ T-cells than CMV seronegative healthy individuals.
One-way analysis of variance < 0.0001 20 0.0005
0.0002 0.0013 15
% 10
5
0 - + - + - + 20-34 35-49 50 CD3+CD56+
Figure 3.5 CD56+ T-cells compared by age group. This scatter plot illustrates the percentage of CD56+ T-cells present in the peripheral blood in 68 healthy donors; 34 were CMV seronegative and 34 were seropositive. It is clearly noticeable that more CD56+ T-cells were present in the PBMCs of CMV seropositive donors than CMV seronegative.
- 119 - 3.2.3.5 CD56+ T-cells compared according to gender
CD56+ T-cell numbers were compared between males and females. Levels of
CD56+ T-cells were compared between CMV seropositive and seronegative male donors and this was also done for female donors (Figure 3.6). After that, the levels of the cells were compared between CMV seropositive males and females, and
CMV seronegative males and females to see if there are any significant differences between the two genders (Figure 3.7).
0.0001 < 0.0001 20
15
% 10
5
0 - + - + Male Female CD3+CD56+
Figure 3.6 CD56+ T-cell levels according to gender. This figure shows the relative difference of CD56+ T-cells level within the same gender groups. Both male and female CMV seropositive subjects had significantly more than seronegative donors (P = 0.0001 and < 0.0001 respectively).
- 120 - 0.8632 20
15 0.0675 % 10
5
0 Male Female Male Female CMV+ CMV- CD3+CD56+
Figure 3.7 Levels of CD56+ T-cells in males and females with the same CMV sero-status. This figure shows the comparison between the levels of CD56+ T- cells in males and females with the same CMV sero-status. No significant differences were detected between genders in either group.
3.3 Characterisation of the phenotype of CD56+ T-cells in CMV seropositive and seronegative healthy subjects
3.3.1 Introduction
T-cell-mediated immunity has diverse characteristics in both function and phenotype. Both function and phenotype are adapted according to many factors.
The previous section 3.2 has shown clearly that CD56+ T-cells were significantly
(P <0.0001) expanded in CMV seropositive healthy donors.
The potential characteristics of CD56+ T-cells were also investigated through the detection of markers on these cells in CMV seropositive and seronegative healthy donors, moreover, the levels of these markers between both sero-statuses were
- 121 - compared to identify any significant variation. The expansion of these cells in
CMV seropositive healthy donors would be expected to lead to characteristic phenotypic differences from CMV seronegative individuals. These variations may involve both surface markers as well as functional differences.
Several markers were investigated in both CMV seropositive and seronegative healthy subjects. Naive T-cells are well known to express five markers CD45RA,
CD62L, CCR7, CD27 and CD28 (Hamann et al. 1997). Memory T-cells have a phenotype distinct from that of naive T-cells, instead of the CD45RA memory T- cells express CD45RO, however, a subtype of memory T-cells expresses high
CD45RA with low CD45RO and these are called TEMRA (Libri et al. 2011). This type of memory cell is highly differentiated in the case of both CD4 and CD8 T- cell compartments and starts to re-express CD45RA and increase in numbers in older individuals (Di Mitri et al. 2011). Moreover, memory T-cells are heterogeneous for marker expression and can be subdivided into functionally and phenotypically distinct subsets; first naive cells differentiate into central memory then effector memory and finally terminally differentiated effector memory TEMRA cells (Table 3.2). TCM or central memory cells express high CD45RO, CD62L and high CCR7, respond to CCL21 (secondary lymphoid tissue chemokine) and migrate centrally to lymph nodes; they secrete IL-2 after TCR triggering but not
IFN-γ and IL-4, though, when they proliferate and become effector cells they start producing IFN-γ and IL-4 (Sallusto et al. 2004). Moreover, CD62L and CCR7 are two essential receptors involved in cell extravasation via high endothelial venules
(HEV) toward T-cell areas in secondary lymphoid organ (Sallusto et al. 2004).
Furthermore, TEM express CD45RO, though they are CCR7 negative and have chemokine receptors and adhesion molecule to assist TEM migration to inflamed
- 122 - tissues. Moreover, they have the potential to secrete IFN-γ, IL-4 and IL-5 while
CD8+ T cells have a large arsenal of perforin in response to antigenic stimulation
(Sallusto et al. 2004).
Many markers were evaluated according to their function and possible role regarding CMV infection. The background regarding their role and functions in the immune mechanism has been explained in Table 3.3.
TCM TEM TEMRA Naive T-cells
CD45RA low low High High
CD45RO High High low low
CD28/CD27/ High High low High
CCR-7/CD62L
IL-2 High low low High
IFN-γ Low High High low
IL-4 Low High - Low
TNF-α High High - low
Table 3.2 Phenotypic characteristics of memory T-cells (Sallusto et al. 2004,
Hamann et al. 1997). TCM = central memory; TEM = effector memory; TEMRA = effector memory cells expressing CD45RA.
Another type of memory T-cell was identified and named TRM (tissue-resident memory T-cells). This type of memory T-cell is believed to remain at the site of infection after clearance of the pathogen. Moreover, the phenotype of these cells was found to be CD62L negative, CCR7 negative, CD11a high, CD44 high, CD69 positive and CD103 positive (Mueller et al. 2013).
- 123 - Table 3.3 Markers applied to phenotypic analysis of CD56+ T-cells in CMV seropositive and seronegative donors.
Marker Immune related function during CMV infection
Interacts with MHC-II on APC via extracellular domain, therefore
amplifying TCR signal to activate T-cells. Frequency of CD4+ T- CD4 cells specific for HCMV was detected to be between 9-40%
(Sylwester et al. 2005, Sester et al. 2002).
A co-receptor to TCR that targets MHC-I that is expressed on all
nucleated cells. Foreign peptide expressed on MHC-I lead to T-cell
CD8 activation leading to apoptosis of the cell under stress. Repeated
CMV viremia in humans who are deficient in CD8+CMV specific
T-cells (Crough and Khanna 2009).
Expressed on activated, memory T-cells, also on several other
CD45RO related immune cells. Improve TCR and BCR (B-cell receptor)
signalling and activation.
Generally expressed on Naive T-cells. Also, it has been found to be CD45RA expressed on highly on TEMRA memory T-cells (Libri et al. 2011).
Also called Lymphocyte function-associated antigen-1 (LFA-1). It
CD11a is a membrane glycoprotein that is involved in cell recruitment to
site of infection (Hamann et al. 1997).
Also called Lymphocyte function-associated antigen-3 (LFA-3),it is
CD58 up regulated due to CMV infection and assists in NK cells lysis of
CMV infected cells (Fletcher et al. 1998).
NK cells expressing CD57+ were expanded during acute HCMV CD57 episodes (Lopez-Verges et al. 2011).
- 124 - A killer lectin-like receptor provides an alternative activation
pathway for NK and CD8+ T-cells (Guma et al. 2005).
CD94/NKG2C CD94/NKG2C+ NK cells were found to expand in response to
CMV infection (Ugolini et al. 2001, Andre et al. 2004, Monsivais-
Urenda et al. 2010, Foley et al. 2012).
Killer cell lectin-like receptor super family B. It has been identified
that CD161 expression is up-regulated non-significantly on NK, CD161 CD56+ T-cells and CD3+ T-cells in CMV seropositive healthy
donors (Monsivais-Urenda et al. 2010).
Cell adhesion molecule known also as L-selectin and involved in
homing function of cells and also called lymph-node homing CD62L receptor (Hamann et al. 1997). TEMRA CD45RO+CD62L- cells were
found to be protective against CMV reactivation (Luo et al. 2010).
Expressed on the surface of T-cells and is a receptor for CD80. It
provides a co-stimulatory signal required for T-cell activation. CD28 Lower CD28 expression was found to be associated with CMV
sero-positivity (Prelog et al. 2013, van Stijn et al. 2008).
Interleukin-7 receptor involved in development of lymphocyte
activation. Involved in early viral-specific CTL becoming memory CD127 cells and down regulated in chronic viral infection such as CMV
(van Stijn et al. 2008, Golden-Mason et al. 2006).
CC Chemokine receptor type 7. Immune-suppressed patients have
CMV specific CD8+CD45RO+CCR7-CD27- cells (Gamadia et al. CCR7 2001). CCR7 expression is down regulated in CMV infection (van
Stijn et al. 2008).
- 125 - CD158a/h
(KIR2DL1/DS1) KIR family expressed on NK cells and some other T-cells. They are
CD158e named depending on the number of extracellular Ig-like domains.
(KIR3DL1) Also, on their cytoplasmic domain Long (L) which are inhibitory
CD158f KIR or short (S) which are activating KIR.
(KIR2DL5)
γδ-T-cells were expanded in peripheral blood of renal transplant
patients after few months due to CMV viremia (Dechanet et al. γδ-TCR 1999b). Non-significant increase was also found in seropositive
healthy donors than sero-negative donors (Alejenef et al. 2014).
Characteristic type of α TCR found mainly on MAIT cells. Using
TCRVα7.2 coloured conjugated antibodies for TCRVα7.2 and CD161 the
MAIT cells can be identified (Gerart et al. 2013).
Activation marker also called α chain of IL-2 receptor generally
found on activated T-cells and B-cells. CMV leads to activation of CD25 short term CD25+ CD4+ and CD8+ T-cells that produce cytokines
and are capable of lysing pp65 of CMV (Samuel et al. 2014).
Activation marker of T-cells and NK cells. Up-regulation of CD69
CD69 on T-cells occurs in response to CMV reactivation (Petersen et al.
2011).
Fc receptors FcγRIIIa (CD16a) and FcγRIIIb (CD16b). CD16 is
CD16 specifically upregulated on HCMV-induced γδ-T-cells (Couzi et al.
2012).
- 126 - 3.3.2: Aims of this study
The aims of this section of the study were to analyse and compare the phenotype of CD56+ T-cells in CMV seropositive and seronegative healthy individuals.
Different memory, naive and activation markers described in Table 3.3 were investigated through staining CD56+ T-cells with a third monoclonal antibody.
Moreover, the levels of these markers on CD56+ T-cells were compared between these two healthy groups by applying a Paired t-test.
3.3.3 Results
Overall, not all markers were evaluated from the same individual therefore the number of donors for each marker is different.
3.3.3.1 Expression of CD4 and CD8 by CD56+ T-cells
PBMCs from healthy CMV seropositive and seronegative were labelled with three colour monoclonal antibodies in order to phenotype CD56+ T-cells. The cells were stained by anti-CD3 PerCP Cy5.5, anti-CD56 FITC and anti-CD4 PE or anti-CD8 PE. The percentage of CD56+ T-cells expressing CD4 or CD8 were calculated after applying only the CD56+CD3+ region of cells to density plots demonstrating CD4 or CD8 cells (Figure 3.8). A total of 32 healthy donors were tested, 16 CMV seropositive and 16 seronegative. CD56+ T-cells were identified to express CD4+, CD8+.
- 127 -
Figure 3.8 Expression of CD4 and CD8 by CD56+ T-cells. This figure shows representative plots of the percentage of gated CD3+CD56+ cells expressing CD4 and CD8.
CMV seronegative CMV seropositive
In this section, the question was asked whether the majority of CD56+ T-cells expressed CD4 or CD8 T-cell receptors among CMV+ or CMV- subjects.
The mean of the expression of these two populations were compared by using
Paired t-test which is shown in Figure 3.9.
The results show that a mean of 74.79% of CD56+ T-cells in CMV seronegative healthy donors were CD8- (mean of CD8+CD56+T-cells 25.21 ± 3.316), while
- 128 - 87.12% of CD56+ T cells in seronegative donors were CD4- (mean of
CD4+CD56+T-cells 12.83 ± 2.361). Although four-colour staining with anti-CD4,
-CD8, -CD3 and -CD56 was not carried out, if no cells are double positive for
CD4 and CD8, this indicates that about 81% of CD56+ T-cells in CMV seronegative subjects were CD4- and CD8-.
CD8 CD4 < 0.0001 0.0167 100
80
60 % 40
20
0 CMV- CMV+ CMV- CMV+
CD56+ T-cells
Figure 3.9 Expression of CD4 and CD8 by CD56+ T-cells in CMV+/-. The percentage of CD8 and CD4 expressed on CD56+ T-cells was compared between
CMV seronegative and seropositive subjects. Both CD4+ and CD8+ CD56+ T- cells have expanded significantly (P= 0.0167 and <0.0001 respectively) in CMV seropositive than seronegative healthy subjects.
When the percentage of CD8+ CD56+ T-cells from both CMV seronegative and seropositive subjects were compared with each other, the results show clearly a significant expansion (P<0.0001) of CD8+CD56+ T-cells in CMV seropositive
(72.28% ± 3.619) compared to seronegative subjects (25.21%± 3.316; Figure 3.9).
- 129 - CD4+CD56+ T-cells were also expanded significantly (P = 0.0167) in CMV seropositive (29.83% ± 2.361) compared to seronegative subjects (12.83% ±
2.361).
Seropositive < 0.0001 100 Seronegative 80 0.0024 60 % 40
20
0 CD8 CD4 CD8 CD4 CD56+ T-cells
Figure 3.10 Expression of CD4 vs CD8 by CD56+ T-cells. The figure show that a greater proportion of CD56+ T-cells expressed CD8 than CD4 in both seronegative and seropositive subjects (P = 0.0024 and <0.0001, respectively).
A greater proportion of CD56+ T-cells were also found to express the CD8 receptor than CD4. This was found in both CMV seronegative and seropositive healthy individuals. Generally, CD8+CD56+T-cells were significantly more abundant (P= 0.0024) than CD4+CD56+T-cells in CMV seronegative subjects.
These CD8+CD56+T-cells were expanded significantly (P <0.0001) in the seropositive group more than CD4+CD56+T-cells (Figure 3.10).
- 130 - 3.3.3.2 CD56+ T-cells; naive and memory markers
T-cell mediated immune responses toward pathogens result in the development of antigen-specific T lymphocytes to eliminate this pathogen. Antigenic specificity depends on T-cell recognition via TCR of the unique peptides expressed on the
MHC molecules presented by APC. T-cell mediated immunity includes three aspects: the primary response via naïve T-cells, effector mechanisms via activated
T-cells and memory response via memory cells, also, T cells induce other cells to participate in the immune response through their secretion of immune modulators.
CD56+ T-cells were stained for naive and memory markers in CMV seropositive and seronegative healthy donors. The markers targeted were CD45RO, CD45RA,
CD62L, CCR7, CD28 and CD127. Levels of expression of these markers were compared between CMV seropositive and seronegative subjects to identify any significant differences between these two groups. Density plots illustrating the levels of expression of these markers are shown in Figure 3.11.
- 131 - Figure 3.11 Memory and naive markers expressed by CD56+ T-cells.
Representative density plots illustrating levels of memory and naive markers expressed by CD56+ T-cells in CMV seronegative and seropositive healthy individuals. The levels of expression were identified after each plot was compared to a negative control.
CMV seronegative CMV seropositive
- 132 -
- 133 -
CD45RO CD45RA CD62L CD28 CD127 CCR7 0.0014 0.1544 < 0.0001 0.0059 < 0.0001 0.0101 100
80
60 % 40
20
0 - + - + - + - + - + - + CD56+ T-cells
Figure 3.12 Comparison of memory and naive markers expressed by CD56+
T-cells. Scatter plot illustrating the comparison between the percentage of each memory and naive marker between CMV seropositive (+) and seronegative (-) subjects. Numbers of donors were as follows: for CD45RO 14 seropositive and 14 seronegative, CD45RA 19 seropositive and 19 seronegative, CD62L 14 seropositive and 14 seronegative, CD28 14 seropositive and 12 seronegative,
CD127 14 seropositive and 14 seronegative, and CCR7 5 seropositive and 7 seronegative.
- 134 - In Figure 3.12 the phenotypic analysis of CD56+ T-cells clearly showed significantly greater expression of CD45RO in seropositive (43.85% ± 4.733) than seronegative subjects (22.95% ± 3.066). Moreover, CD45RA showed almost the same level of expression between seropositive (98.88% ± 0.1436) and seronegative subjects (96.34% ± 2.189) clearly a substantial number of CD56+ cells from seropositive donors expressed both CD45RA and 45RO. CD62L was significantly expressed by more CD56+ T cells in seronegative (62.72% ± 4.611) than seropositive subjects (30.84% ± 6.150). Similarly, CD28 was also expressed on a significantly higher percentage of CD56+ T cells in seronegative (52.17% ±
7.314) than seropositive subjects (22.10% ± 2.852). Likewise, CD127 was expressed by significantly more CD56+ T cells in seronegative (78.73% ± 4.191) than seropositive subjects (23.88 ± SE 3.858) as was CCR7 (64.21% ± 9.713 vs.
20.23% ± 5.413).
CD56+ T-cells from CMV seronegative subjects have been shown to express lower proportions of CD45RO+ cells but higher proportions of CD45RA+,
CD62L+, CD28+ and CD127+ cells than CMV seropositive subjects. This pattern of surface marker expression is similar to that of naive and central memory T- cells. However, in CMV seropositive healthy individuals the CD56+ T-cell phenotype of high CD45RO, re-expressed CD45RA, low CD62L, low CD28, low
CD127 and low CCR7 was similar to that of TEMRA memory T cells and different from TEM in their re-expression of CD45RA as TEM do not re-express CD45RA.
- 135 - 3.3.3.3 CD56+ T-cell expression of Lymphocyte function-associated antigens
CD56+ T-cells were phenotyped for their levels of expression of LFA-1 and LFA-
3 and CMV seropositive and seronegative healthy donors compared. CD11a or
LFA-1 is involved in T-cell migration to the site of infection, CD58 or LFA-3 is upregulated in response to CMV infection and assists in NK cell lysis of CMV infected cells (Fletcher et al. 1998). In Figure 3.13 the proportions of CD56+ T cells expressing CD11a and CD58 are illustrated.
CMV seronegative CMV seropositive
Figure 3.13 LFA-1& 3 expressions by CD56+ T-cells. Representative density plots illustrating proportions of CD56+ T cells expressing CD11a/LFA-1 and CD58/LFA-3 in
CMV seronegative and seropositive healthy individuals. The levels of expression were identified after each plot was compared to a negative control.
- 136 - CD11a/LFA-1 CD58/LFA-3 5000 0.0228 0.6805 0.0101
100 4000
80 3000
60 MFI % 2000 40 1000 20
0 0 - + - + - + CD56+ T-cells CD11a+ CD56+ T-cells
Figure 3.14 LFA-1& 3 expressions by CD56+ T-cells between CMV+/-. On the left levels of LFA-1 and -3 expressed by CD56+ T-cells were measured and compared between CMV seropositive and seronegative healthy individuals. Number of donors for
CD11a were 19 seropositive and 19 seronegative, for CD58 the donors were 5 seronegative and 7 seropositive. The right scatter plot represents MFI levels of CD11a, it was found clearly MFI of CD11a forCMV seropositive subjects were significantly greater (P= 0.0228) than for CMV seronegative subjects.
Both CMV seropositive (98.85% ± 0.4308) and seronegative subjects (99.56% ±
0.1519) expressed similar proportions of CD11a+ CD56+ T cells, moreover, when
MFI levels of CD11a were compared, it was found that MFI of CD11a on cells from CMV seropositive subjects were significantly higher (P= 0.0228) than on those from CMV seronegative subjects. However, CD58 was expressed on a significantly lower proportion of CD56+ T cells (P = 0.0101) in CMV
- 137 - seropositive (88.83% ± 5.049) than seronegative donors (40.70% ± 12.36; Figure
3.14).
3.3.3.4 CD56+ T-cell expression of CD57, CD161, CD94/NKG2C and CD16
CD56+ T-cells were phenotyped for their expression of CD57, CD161,
CD94/NKG2C and CD16 and levels of expression compared between CMV seropositive and seronegative healthy donors. In Figure 3.15 the proportions of cells expressing CD57, CD161, CD94/NKG2C and CD16 on CD56+ T-cells are illustrated.
Figure 3.15 CD57, CD161, CD94/NKG2C and CD16 expression by CD56+ T- cells. Density plots illustrating levels of CD57, CD161, CD94/NKG2C and CD16 expressed by CD56+ T-cells in representative CMV seronegative and seropositive healthy individuals
CMV seronegative CMV seropositive
- 138 -
- 139 -
CD57 CD161 CD94 NKG2C CD16 0.0177 0.0022 0.0101 0.0015 0.5067 100
80
60 % 40
20
0 - + - + - + - + - + CD56+ T-cells
Figure 3.16 CD57, CD161, CD94/NKG2C and CD16 expression by CD56+ T- cells in CMV+ and CMV- subjects. Proportions of CD56+ T cells expressing
CD57, CD161, CD94/NKG2C and CD16 were measured and compared between
CMV seropositive and seronegative healthy individuals. Number of donors for
CD57 were 5 seropositive and 7 seronegative, CD161 were 17 seropositive and 16 seronegative, CD94 were 5 seropositive and 7 seronegative, NKG2C were 7 seropositive and 18 seronegative, and CD16 were 15 seropositive and 12 seronegative.
- 140 - In figure 3.16 a significantly higher percentage of CD56+ T-cells expressed CD57 in seropositive (77.43% ± 9.186) than seronegative subjects (29.30% ± 10.42).
Additionally, seronegative subjects (21.86% ± 4.622) had a significantly (P =
0.0022) higher proportion of CD161+ cells than seropositive subjects (6.857% ±
2.451). Moreover, significantly more CD56+ T cells expressed CD94 in seropositive (62.84% ± 8.261) than seronegative subjects (31.44% ± 7.281), and
NKG2C was also present on a significantly higher proportion of CD56+ T cells in seropositive (21.9% ± 4.94) than seronegative subjects (4.253% ± 1.496; P =
0.0015). However, CD16 was not expressed on a significantly higher proportion of CD56+ T cells (P= 0.5067) in seronegative (21.76% ± 4.596) than seropositive subjects (21.27% ± 2.905; P = 0.5067).
3.3.3.5 CD56+ T-cells expression of KIRs CD158 a/h, CD158e and CD158f
CD56+ T-cells were phenotyped for their expression of killer-cell immunoglobulin-like receptors (KIRs) CD158a/h, CD158e and CD158f and CMV seropositive and seronegative healthy donors compared (Figure 3.17). In figure
3.18 the levels of CD158a/h, CD158e and CD158f expression on CD56+ T-cells are illustrated.
Overall, no significant differences were found when CD56+ T-cells were compared between both CMV groups. Large variations were found in KIRs expression in all donor groups.
- 141 - Figure 3.17 KIR expressions by CD56+ T-cells. Plots illustrating expression of
KIRs CD158a/h, CD158e, and CD158f by CD56+ T-cells in representative CMV seronegative and seropositive healthy individuals.
CMV seronegative CMV seropositive
- 142 - a/h e f 0.8763 0.5303 0.2020 80
60
% 40
20
0 - + - + - + CD56+ T-cells
Figure 3.18 KIR expressions by CD56+ T-cells in CMV+ and CMV- subjects.
Proportions of CD56+ T cells expressing CD158a/h, CD158e and CD158f compared between CMV seropositive and seronegative healthy individuals.
Number of donors for CD158a/h were 5 seropositive and 7 seronegative, CD158e were 5 seropositive and 7 seronegative, CD158f were 5 seropositive and 7 seronegative.
3.3.3.6 CD56+ T-cell expression of CD25 and CD69
CD56+ T-cells were phenotyped for their expression of CD25 and CD69 and compared between CMV seropositive and seronegative healthy donors. In Figure
3.19 the proportions of CD56+ T cells expressing CD25 and CD69 are illustrated.
Overall, no significantly differences were detected in CD56+ T-cell phenotype when CMV seronegative was compared to seropositive healthy individuals.
However, CD56+ T-cells were expressing more CD69 in seropositive (2.375% ±
- 143 - 0.5830) than seronegative (2.845% ± 0.5092). Also, CD25 were expressed slightly more by seronegative (1.995% ± 0.4212) than seropositive (1.548% ± 0.3290) healthy donors (P= 0.5037).
Figure 3.19 CD69 and CD25 expression by CMV+/-. Proportions of CD56+ T cells expressing CD69 and CD25 in representative CMV seronegative and seropositive healthy individuals.
CMV seronegative CMV seropositive
- 144 - 15 CD69 CD25 0.2602 0.5037
10
%
5
0 - + - + CD56+ T-cells
Figure 3.20 CD69 and CD25 expression by CD56+T-cells in CMV+ and
CMV- subjects. Proportions of CD56+ T cells expressing CD69 and CD25 were measured and compared between CMV seropositive and seronegative healthy individuals. Numbers of donors for CD69 were 19 seropositive and 19 seronegative and for CD25 were 18 seropositive and 16 seronegative. Both markers were not tested from the same individual.
3.3.3.7 CD56+ T-cell expression of TCRVα7.2 and γδ-TCR
CD56+ T-cells were phenotyped for their expression of two different types of
TCRs and compared between CMV seropositive and seronegative healthy donors.
In Figure 3.21 the proportions of CD56+ T cells expressing TCRVα7.2 and γδ-
TCR in representative donors are illustrated.
- 145 -
Figure 3.21 TCRVα7.2 and γδ-TCR expression by CD56+ T-cells. Plots illustrating levels of TCRVα7.2 and γδ-TCR expressed by CD56+ T-cells in representative CMV seronegative and seropositive healthy individuals.
CMV seronegative CMV seropositive
- 146 - TCR V7.2 TCR 80 0.6428 0.8868
60
% 40
20
0 - + - + CD56+ T-cells
Figure 3.22 TCRVα7.2 and γδ-TCR expression by CD56+ T-cells in CMV+ and CMV- subjects. The scatter plot shows the proportions of CD56+ T cells expressing γδ TCR and Vα-7.2 TCR in healthy CMV seronegative and seropositive subjects. Overall, no significant difference was found between the two CMV groups. The numbers of donors for each group were 19 seropositive and 20 seronegative for γδ-TCR and 9 seropositive and 12 seronegative for Vα7.2
TCR.
In figure 3.22 γδ TCR expression on CD56+ T-cells was compared between CMV seropositive (21.39% ± 4.052) and seronegative subjects (21.15% ± 2.927) no significant difference resulted.
V α 7.2 TCR expressions by CD56+ T cells also did not differ significant when between seropositive (16.07% ± 3.463) and seronegative subjects were compared
(15.68% ± 3.626).
- 147 - 3.3.3.8 CD56+ T-cell expression of exhaustion markers
CD56+ T-cells were phenotyped for their expression of PD-1, CTLA-4 and
KLRG1 and compared between CMV seropositive and seronegative healthy donors. Proportions of CD56+ T cells expressing PD-1, KLRG1 and CTLA-4 in representative donors are illustrated in Figure 3.23. Moreover, the levels of expression In CMV seropositive and seronegative donors are illustrated in density plots (Figure 3.24).
0.7512 0.0435 0.4290
100
80
60 % 40
20
0 - + - + - + PD-1 KLRG-1 CTLA-4 CD56+ T-cells
Figure 3.23 PD-1, CTLA-4 and KLRG-1 expression by CD56+ T-cells in
CMV+ and CMV- subjects. The scatter plot shows the proportions of CD56+ T cells expressing PD-1, CTLA-4 and KLRG-1 in healthy CMV seronegative and seropositive. Only KLRG-1 has shown significant difference between CMV seropositive and seronegative healthy donors.
- 148 - Figure 3.24 PD-1, CTLA-4 and KLRG-1 expression by CD56+ T-cells in
CMV+ and CMV- subjects. Plots illustrating levels of CD56+ T cells expressing
PD-1, KLRG1 and CTLA-4 in representative CMV seronegative and seropositive healthy individuals.
CMV seronegative CMV seropositive
- 149 - Overall, no significant difference was found between the two CMV groups in PD-
1 (seronegative 8.187% ±2.064, seropositive 13.2% SE± 6.056) and CTLA-4
(seronegative 74.68% ±4.949, seropositive 74.25% ±4.777). However, KLRG-1 expression was significantly more in CMV seropositive (12.23% ±3.994) than seronegative (10.83% ±2.424) (P= 0.0435). Number of donors for seronegative 7 and seropositive 11 (Figure 3.23).
3.3.4 Discussion
In this part of the study relative numbers of the distinct subset of CD3+ T-cells that express the NK receptor CD56 (NCAM) were evaluated. This type of lymphocyte is also named CD56+ T-cells, CIK, NKT and CD56+CD3+ cells.
This part of the study aimed to identify any difference in levels of this subset of lymphocytes between CMV seropositive and seronegative healthy individuals, also, to evaluate both NK cell and CD3+CD56- T-cell levels between the same healthy individuals. A secondary aim was to identify if the levels were significantly different between age groups and gender groups. Section 3.2.3.1 has clearly identified that CD56+ T-cells were significantly expanded in CMV seropositive healthy donors compared to seronegative donors. Additionally, the mean levels of CD56+ T-cells in CMV seropositive subjects (8.38%) were 2.5 folds higher than the mean of CD56+ T-cells in CMV seronegative subjects
(3.29%).
Section 3.2.3.2 tested for differences in the levels of CD3+ CD56- T-cells between CMV seropositive and seronegative healthy donors. When comparing the levels of CD3+ CD56- T-cells as a proportion of total lymphocytes it was shown
- 150 - that CMV seronegative subjects had relatively more CD3+CD56- T-cells than seropositive subjects although this was not statistically significant (P=0.0516).
However, when CD3+CD56- T-cells has been compared between these two healthy groups after excluding CD56+T-cells there was clearly a significant difference (P<0.0001).
Any differences in NK cell levels between CMV seropositive and seronegative subjects were also evaluated and compared in section 3.1.3.3. NK cells were identified as cells that only express CD56 but without CD3. Mean levels of NK cells were the almost the same between CMV seropositive and seronegative healthy donors (P=0.7873).
Data were subdivided into different age groups which were compared in section
3.1.3.4. It was clear that CMV seropositive subjects had more CD56+ T-cells than
CMV seronegative subjects in all age groups. What is more, those healthy donors who were 50 years and over and CMV seropositive had more expanded CD56+ T- cells than the rest of the seropositive age groups. All CMV seronegative groups expressed similar levels of CD56+ T-cells. Comparing the levels of CD56+ T cells between genders in section 3.1.2.5 showed no significant difference between males and females.
All the previous sections clearly identified that CMV seropositive subjects had accumulated more CD56+ T-cells in their peripheral blood than CMV seronegative healthy individuals, which implies a role of these cells in response to chronic CMV carriage.
For the phenotypic analysis of this part of the study CD56+ T-cells were investigated. This was done through three colour surface staining of CD56+ T-
- 151 - cells in two groups of healthy donors; CMV seropositive and seronegative. The surface receptors that were targeted were: CD4, CD8, CD45RA, CD45RO,
CD62L, CCR7, CD28, CD127, CD11a/LFA-1, CD58/LFA-3, CD57, CD161,
CD94/NKG2C, CD16, CD158a/h, CD158e, CD158f, CD25, CD69, TCRVα7.2 and γδ-TCR. Percentage expression was calculated by density plot after the levels were compared to negative controls. Any differences in the percentage expression between CMV seronegative and seropositive subjects were compared by applying a Paired t-test. CMV status was determined by using ELISA as mentioned earlier in section 3.1.
CD56+ T-cells in CMV seropositive subjects were found to comprise a higher proportion of CD8+ and CD4+ cells than in seronegative subjects. Although four color staining with anti-CD3, CD56, CD4 and CD8 was not conducted, in some
CMV+ subjects the total % of CD56+ T cells expressing CD4 or CD8 was >100% suggesting that some expressed both CD4 and CD8. The higher incidence of CD8 expression on CD56+ T-cells was also found in several studies that identified that
CD8+ T-cells provide protection against CMV infection in both human and mice
(Crough and Khanna 2009). Moreover, HCMV detected to be reactivated in individuals who were deficient in CMV-specific CD8+ T-cells (Crough and
Khanna 2009). The proportionate increase in CD4+ CD56+ T-cells was also found in other studies in which HCMV-specific CD4+ T-cells increased between
9% to 40% (Sylwester et al. 2005, Sester et al. 2002).
Overall, CD8 was expressed significantly more by CD56+ T-cells than CD4 in both seronegative and seropositive subjects. This was also explained by other studies who found that CD8+ T-cells can expand to 17 folds to give rise to CD56+
T-cells, however, CD4+ cells failed to do so and even the generated cells were
- 152 - CD56 negative (Franceschetti et al. 2009). Both CD4 and CD8 tend to be stable markers of post-thymic T cells so any increases in response to CMV infection are presumably a result of expansion of CD4+ or CD8+ CD56+ T cells.
On other hand, some studies stated that CD56 acquisition is correlated with increased cytotoxicity of these cells. CD56 was acquired by lymphocytes in an ex vivo environment that already expressed the phenotype CD4+CD8+, CD4-CD8- and CD4-CD8+ after the cells were cultured with IL-2 and anti-CD3 mAb (Lu and Negrin 1994). However, it is not clear whether CMV infection in vivo leads to expansion of T cells already expressing CD56 or induction of CD56 expression by previously CD56- cells.
The phenotype of CD56+ T-cells in CMV seronegative subjects was found to be similar to that of naïve T cells, as low numbers expressed CD45RO, while high numbers expressed CD45RA, CD62L, CD28 and CD127. They only differed from TCM in their low expression of CD45RO while TCM have adequate expression of CD45RO (Sallusto et al. 2004). Furthermore, in analysis of CMV seropositive subjects it was found that numbers of CD56+T-cells expressing
CD45RO was increased significantly, together with a reduction in numbers of cells expressing CD62L, CD28, CD127 and CCR7. This surface expression is similar to TEMRA and different from TEM in re-expression of CD45RA as TEM do not re-express CD45RA (Sallusto et al. 2004). CD45RA re-expression was explained in several studies as an outcome of persistent chronic viral infection
(Henson et al. 2012).
CD11a is a component of the integrin LFA-1 and levels of expression on CD56+
T-cells were the same between each CMV group. However, MFI levels were
- 153 - increased significantly in CMV seropositive compared to seronegative subjects.
CD11a is essential in homing of immune cells to an infection site and also in extravasation and activation (E. Jiang et al. 2014). CD58, the ligand for
CD2/LFA-2 and essential in cell adhesion, was significantly increased in CD56+
T-cells in CMV seropositive compared to seronegative healthy individuals. This was also found in other studies indicating up-regulation of CD58 in response to
CMV infection, also, a strong correlation was found between NK cell lysis of
CMV-infected cells and the surface expression of CD58 (Fletcher et al. 1998).
CD57, CD94 and NKG2C were expressed by significantly more CD56+ T-cells in
CMV seropositive than seronegative healthy individuals. This is also consistent with another investigation that showed expansion of a CD94/NKG2C and CD57 expressing NK cell population during CMV infection (Lopez-Verges et al. 2011).
The ligand for NKG2C is HLA-E, which is expressed by CMV-infected cells and bound by the CMV encoded protein UL40. Also, HLA-E has a high affinity for the inhibitory receptor NKG2A, however this study identified that high
CD57/NKG2C NK cells lacked expression of NKG2A (Lopez-Verges et al. 2011) and another study has shown that culturing PBMCs with CMV-infected cells resulted in selective expansion of NK cell expressing the activation receptor
CD94/NKG2C over cells expressing CD94/NKG2A (Guma et al. 2006).
The significantly reduced numbers of CD161+ CD56+ T-cells in CMV seropositive subjects argued against any virus-associated expansion of Th17 cells, iNKT cells or MAIT cells, all of which are CD161+ (Cosmi et al. 2008,
O'Reilly et al. 2011, Dusseaux et al. 2011). Also, in CMV seropositive children
CD161 was expanded on NK cells (Monsivais-Urenda et al. 2010), however
CD161 was absent from CMV-specific CTL (Northfield et al. 2008). CD16 is
- 154 - expressed in the form of the Fc receptor FcγRIIIa (CD16a) on lymphocytes while
FcγRIIIb (CD16b) is expressed on neutrophils. CD16 expressing cells were found to be expanded in HCMV-induced γδ-T-cells (Couzi et al. 2012) while in this study CD56+ T-cells from CMV seronegative subjects did not have significantly less CD16 than in seropositive subjects. There were no significant differences in expression of KIRs CD158a/h, CD158e and CD158f on CD56+ T-cells from
CMV seronegative and seropositive subjects.
CD69 is an activation marker of T-cells and NK cells. Up-regulation of CD69 on
T-cells occurs in response to CMV reactivation (Petersen et al. 2011). However, this study has shown that levels of CD69 expression were almost the same between both seropositive and seronegative subjects and no significant difference resulted. CD25 is an activation marker of activated T-cells and B-cells and CMV leads to activation of short term CD25+ CD4+ and CD8+ T-cells that produce cytokines capable of lysing CMV pp65 expressing cells (Samuel et al. 2014).
However, no significant difference in CD25 expression was found between CMV seropositive and seronegative groups. About 21% of CD56+ T-cells in PBMCs were γδ-TCR positive however when CMV seropositive and seronegative donors were compared there was no significant difference, this argues against a study that had observed an expansion in γδ-T-cells in PBMCs in renal allograft patients due to CMV infection (Dechanet et al. 1999b). However, it in a recent paper from this group, a selective expansion of Vδ1+ γδ T cells in CMV positive healthy donors with increasing age was reported (Alejenef et al, 2014). It is possible that overall levels of γδ T cells remain unchanged following CMV infection as the normally more abundant Vδ2+ cells may not be expanded. Numbers of cells expressing Vα-
- 155 - 7.2 TCR, a marker of iNKT cells, also showed no significant differences between seropositive and seronegative subjects.
Overall, Figure 3.25 illustrates significantly increased while Figure 3.26 illustrates significantly decreased surface markers between CD56+ T cells from CMV seropositive and seronegative groups.
Figure 3.25 Phenotypic differences between CD56+ T cells from CMV seropositive and CMV seronegative healthy subjects. Cell surface markers that were mostly increased in CMV seropositive subjects. Results are expressed as mean percentage of CD56+ T cells with p values above indicating levels of significance between CMV seronegative and CMV seropositive subjects.
CD8+ CD4+ CD45RO+ CD45RA+ CD11A+ CD58+ CD57+ CD94+ NKG2C+ < 0.0001 0.0167 0.0014 0.1544 0.6805 0.0101 0.0177 0.0177 0.0015 100 90 80 70 60 % 50 40 30 20 10 0 - + - + - + - + - + - + - + - + - + CD3+CD56+
- 156 - Figure 3.26 Phenotypic differences between CD56+ T cells from CMV seropositive and CMV seronegative healthy subjects. Cell surface markers that were mostly decreased in CMV seropositive subjects. Results are expressed as mean percentage of CD56+ T cells with p values above indicating levels of significance between CMV- and CMV+ subjects.
CD161+ CD62L+ CD28+ CD127+ CCR7+ 0.0022 0.0004 0.0059 < 0.0001 0.0101 100
80
60 % 40
20
0 - + - + - + - + - + CD3+CD56+
CD56+ T-cell phenotypic characteristics in CMV seropositive subjects were consistent with memory cells (TEMRA) and different from TEM in their re- expression of CD45RA. While, in CMV seronegative subjects the phenotypic characteristic of CD56+ T-cells were similar to naïve T cells and different from
TCM in having low expression of CD45RO.
Exhaustion markers were included to validate the responsive functions of CD56+
T-cells. In chronic viral status the cells that are virus-specific such as CD8+ T- cells lose their cytotoxicity and acquire a non-functional exhausted status, starting
- 157 - by losing IL-2 production then followed by TNF-α and IFN-γ (Wherry et al.
2004). PD-1 (Programmed cells death) and PD-1-Ligands play a role in immuno- regulatory functions for T-cells activation and tolerance (Jin et al. 2011). No differences were noted in CD56+ T-cells between seropositive and seronegative healthy donors. KLRG-1or killer lectin-like receptor G1 recognizes the classical cadherin family (Grundemann et al. 2010). CD56+ T-cells in seropositive subjects expressed significantly more KLRG-1 than seronegative groups. This was also reported as KLRG-1 expression was induced in virus specific CD8+ T-cells against repetitive activation of chronic CMV, HIV, EPV infection which influences the function of these cells (Thimme et al. 2005). Also, another study showed that T-cells that express KLRG-1 preserve their function, however they demonstrate deprived proliferation capability (Voehringer et al. 2002), increase
KLRG-1 up-regulation of virus-specific CD8+ T-cells is an evidence of T-cell differentiation (Bengsch et al. 2010). On the other hand, CTLA-4 levels were roughly the same between sero-groups and no significant variation detected.
Surprisingly, levels of KLRG-1 and CTLA-4 were elevated, however, PD-1 was not which contradicts another study which claimed that PD-1 and KLRG-1 but not
CTLA-4 were elevated in virus specific CD8+ T-cells (Bengsch et al. 2010).
- 158 -
Chapter 4
Functional responses in CD56+ T cells from CMV seropositive and
CMV seronegative healthy subjects
- 159 - 4. Functional responses in CD56+ T cells from CMV seropositive and CMV seronegative healthy subjects
4.1 Cytokine responses of CD56+ T-cells
4.1.1 Introduction
It has been found that relative numbers of CD56+ T-cells have been expanded in
CMV seropositive healthy subjects. The next step was involving the evaluation of the capability of CD56+ T-cells to recognise and respond to CMV lysate. The cells were stimulated and evaluated for the immune expression of IFN-γ, TNF-α,
IL-2, Perforin, Granzyme A and Granzyme B.
IFN-γ is a type II interferon and involved in immune responses to viral infection.
The interferon were subtyped according to their sequence homology and receptor specificity (Schroder et al. 2004). Both IFN-γ and TNF-α (tumour necrosis factor- alpha) have been found to be expressed by CD8+ and CD4+ T-cells when induced by viral lysate (Radha et al. 2005). IL-2 is naturally produced in response to infection, and it is essential in several biological functions involving growth, differentiation and proliferation of cells. Moreover, TCR stimulation leads to production of IL-2 (Stern and Smith 1986). Perforin and Granzyme are part of one of the pathways of apoptosis induced by lymphocytes in response to virally infected or transformed cells. Both these molecules work in cooperative method in order to induce target cell apoptosis (Trapani and Smyth 2002).
PBMCs from both CMV seropositive and seronegative subjects were incubated
o overnight in culture medium in a humidified 5% CO2 atmosphere at 37 C. CD56+
- 160 - T-cell response was assessed by their content of IFN-γ, TNF-α, IL-2, Perforin,
Granzyme-A and -B as described in section 2.4.2.1.
4.1.2 Aims
This section is part of the functional investigations of the capabilities of CD56+
T-cells when stimulated by CMV lysate. Levels of cytokine expression from
CD56+ T-cells will be compared between cells from CMV seropositive and seronegative subjects. Also, CD56+ T-cell cytokines expression will be compared to NK cells and CD56- T-cells.
4.1.3 Results
PBMCs were incubated for 24 hours in 48 well plates; each well contained 1 X
106 cells/ ml. The cells were stimulated by CMV lysate at 2 µl/ml and SEB
(Staphylococcal enterotoxin B) at 1µl/ml. Moreover, un-stimulated cells were used as background and an isotype control was also applied to remove non- specific binding of monoclonal antibodies. Cells were labelled with anti-CD3 and anti-CD56 to identify CD56+CD3- NK cells, CD56+ T-cells and CD56- T-cells, the cells were further stained intracellularly as explained in section 2.4.2.1 for
IFN-γ, TNF-α, IL-2, Perforin, Granzyme A and B. Both results from CMV seropositive and seronegative donors were compared by using a Paired T-test, p<0.05 being regarded as statistically significant.
- 161 - SEB was used as a preferred positive control for intracellular staining for cytokines in response to CMV lysate and examples of the results are illustrated in
Figure 4.1.
Figure 4.1 SEB as a positive control for CD56+T-cells. Examples of the cells’ response to overnight stimulation with SEB on IFN- γ expression.
4.1.3.1 IFN-γ expression following stimulation with CMV lysate
IFN-γ expression by NK cells, CD56+ T-cells and CD56- T-cells after cells were stimulated overnight with CMV lysate. The levels of expression are illustrated by density plot in Figure 4.2.
Generally, PBMCs from healthy CMV seropositive donors expressed more IFN-γ in response to CMV lysate when incubated overnight in a humidified 5% CO2 atmosphere at 37oC. Following that, levels of IFN-γ expression were compared between CMV seropositive and seronegative by using a Paired T-test. The results are illustrated in Figure 4.3. The three groups of cells expressed significantly
- 162 - more IFN-γ in seropositive donors than seronegative subjects. IFN-γ expressed by
NK cells was significantly more (P = 0.002) in CMV seropositive (5.937%
±1.049, N=18) than seronegative subjects (2.332% ±0.4045, N=23). Also, IFN-γ expressed by CD56+ T-cells was significantly more (P = 0.0042) in CMV seropositive (5.482% ±0.6093, N=18) than seronegative subjects (2.585%
±0.3664, N=23). Moreover, IFN-γ expressed by CD56- T-cells was significantly more (P = 0.0399) in CMV seropositive (3.843% ±0.7222, N=18) than seronegative subjects (1.976% ±0.2981, N=22).
- 163 - Seronegative Seropositive
Figure 4.2 Levels of IFN-γ expression. In the representative populations of NK cells, CD56+ T-cells and CD56- T-cells after overnight stimulation with CMV lysate..
- 164 - 0.0020 0.0042 0.0399 20
15
10
% IFN- 5
0 - + - + - + NK CD56+ T-cells CD56- T-cell
Figure 4.3 Levels of IFN-γ expression in CMV+ and CMV- subjects. Paired T- tests were used to compare levels of IFN-γ expression by NK cells, CD56+ T- cells and CD56- T-cells after overnight stimulation with CMV lysate. IFN-γ was expressed by significantly more NK cells (P = 0.002) in CMV seropositive than seronegative subjects. Also, IFN-γ was expressed by significantly more CD56+
T-cells (P = 0.0042) in CMV seropositive than seronegative subjects. Moreover,
IFN-γ was expressed by more CD56- T-cells (P = 0.0399) in CMV seropositive than seronegative subjects.
4.1.3.2 TNF-α expression following stimulation with CMV lysate
Levels of TNF-α expression by NK cells, CD56+ T-cells and CD56- T-cells were studied after cells were stimulated by CMV lysate. The levels of expression were illustrated by density plot in Figure 4.4. TNF-α expressed by NK cells was not significantly different in CMV seropositive (3.008% ±0.6572, N=23) than seronegative subjects (2.264% ±0.4835, N=12). TNF-α expressed by CD56+ T- cells was significantly more (P = 0.0088) in CMV seropositive (3.912% ±0.6010,
N=23) than seronegative subjects (1.936% ±0.2648, N=20). Moreover, TNF-α
- 165 - expressed by CD56- T-cells was not significantly different (P = 0.3810) in CMV seropositive (1.926% ±0.2671, N=26) than seronegative subjects (1.369%
±0.3660, N=22), these results are illustrated in Figure 4.5.
Seronegative Seropositive
Figure 4.4 Levels of TNF-α expression. In a representative NK cells, CD56+ T-cells and CD56- T-cells from representative subjects after overnight stimulation with CMV lysate..
- 166 - 0.2017 0.0088 0.3810 15
10
% TNF 5
0 - + - + - + NK CD56+ T-cells CD56- T-cells
Figure 4.5 A Levels of TNF-α expression in CMV+ and CMV- subjects. Paired T- test was used to compare levels of TNF-α expression by NK cells, CD56+ T-cells and
CD56- T-cells after overnight stimulation with CMV lysate. TNF-α expressed by NK cells was not significantly different in CMV seropositive than seronegative subjects.
TNF-α expressed by CD56+ T-cells was significantly more (P = 0.0088) in CMV seropositive than seronegative subjects. Moreover, TNF-α expressed by CD56- T- cells was not significantly different (P = 0.3810) in CMV seropositive than seronegative subjects.
4.1.3.3 IL-2 expression following stimulation with CMV lysate
IL-2 expression by NK cells, CD56+ T-cells and CD56- T-cells was also studied after cells were stimulated by CMV lysate. The levels of expression are illustrated by density plot in Figure 4.6. IL-2 expressed by NK cells was not significantly different (P = 0.4086) in CMV seropositive (1.303% ±0.2354, N=14) than seronegative subjects (1.143% ±0.2076, N=17). Neither was IL-2 expressed by
CD56+ T-cells significantly different (P = 0.1151) in CMV seropositive (2.170%
- 167 - ±0.3907, N=10) than seronegative subjects (1.528% ±0.4528, N=17). Moreover,
IL-2 expressed by CD56- T-cells was also not significantly different (P = 0.1272) in CMV seropositive (0.6831% ±0.1183 N=25) than seronegative subjects
(1.401% ±0.3729, N=11), as illustrated in Figure 4.7.
Seronegative Seropositive
Figure 4.6 Levels of IL-2 expression. In NK cells, CD56+ T-cells and CD56- T- cells from representative subjects after overnight stimulation with CMV lysate.
- 168 - 0.4086 0.1151 0.1272 8
6
4
% IL-2
2
0 - + - + - + NK CD56+ T-cells CD56- T-cells
Figure 4.7 Levels of IL-2 expression in CMV + and CMV- subjects. Paired T- test was used to compare levels of IL-2 expression by NK cells, CD56+ T-cells and CD56- T-cells after overnight stimulation with CMV lysate. IL-2 expressed by NK cells was not significantly different (P = 0.4086) in CMV seropositive than seronegative subjects. IL-2 expressed by CD56+ T-cells was not significantly different (P = 0.1151) in CMV seropositive than seronegative subjects. Moreover,
IL-2 expressed by CD56- T-cells was not significantly different (P = 0.1272) in
CMV seropositive than seronegative subjects.
- 169 - 4.1.3.4 Perforin content following stimulation with CMV lysate
Perforin expression by NK cells, CD56+ T-cells and CD56- T-cells was tested after cells were stimulated by CMV lysate. The levels of expression are illustrated by the density plots in Figure 4.8. Perforin expression by NK cells was not significantly different (P = 0.6524) in CMV seropositive (90.43% ±2.094, N=10) than seronegative subjects (90.06% ±4.062, N=7). Perforin expression by CD56+
T-cells was not significantly different (P = 0.2335) in CMV seropositive (71.49%
±5.503, N=10) than seronegative subjects (54.90% ±11.28, N=7). Moreover, perforin expression by CD56- T-cells was not significantly different (P = 0.6592) in CMV seropositive (43.44% ±9.476, N=10) than seronegative subjects (30.03%
±7.742, N=7), these results are illustrated in Figure 4.9.
- 170 - Figure 4.8 Levels of perforin. Within NK cells, CD56+ T-cells and CD56- T- cells from representative subjects after overnight stimulation with CMV lysate.
Seronegative Seropositive
- 171 - Figure 4.9 Levels of perforin in CMV+ and CMV- subjects. Paired T-test was used to compare perforin expression by NK cells, CD56+ T-cells and CD56- T- cells after overnight stimulation with CMV lysate. Perforin expression by NK cells was not significantly different (P = 0.6524) in CMV seropositive than seronegative subjects. Perforin expression by CD56+ T-cells was not significantly different (P = 0.2335) in CMV seropositive than seronegative subjects. Moreover, perforin expression by CD56- T-cells was not significantly different (P = 0.6592) in CMV seropositive than seronegative subjects.
0.6524 0.2335 0.6592 100
80
60 % 40
20
0 - + - + - + NK CD56+ T-cells CD56- T-cells
4.1.3.5 Granzyme A expression following stimulation with CMV lysate
Granzyme A expression by NK cells, CD56+ T-cells and CD56- T-cells was tested after cells were stimulated by CMV lysate. The levels of expression were illustrated by density plots in Figure 4.10.
- 172 - Granzyme A expression by NK cells was not significantly different (P = 0.5976) in CMV seropositive (47.43% ±10.61, N=12) than seronegative subjects (38.44%
±11.78, N=7). Granzyme A expression by CD56+ T-cells was not significantly different (P = 0.3053) in CMV seropositive (49.25% ±6.776, N=12) than seronegative subjects (25.26% ±9.815, N=7). Moreover, perforin expression by
CD56- T-cells was not significantly different (P = 0.1830) in CMV seropositive
(19.98% ±7.183, N=12) than seronegative subjects (6.343% ±3.253, N=7), these results are illustrated in Figure 4.11.
- 173 - Figure 4. Levels of Granzyme A expression. In a representative NK cells,
CD56+ T-cells and CD56- T-cells from representative subjects after overnight stimulation with CMV lysate
Seronegative Seropositive
- 174 - Figure 4.11 Levels of Granzyme A expression in CMV + and CMV- subjects.
Paired T-test was used to compare levels of Granzyme A expression by NK cells,
CD56+ T-cells and CD56- T-cells after overnight stimulation with CMV lysate.Granzyme A expressed by NK cells was not significantly different (P =
0.5976) in CMV seropositive than seronegative subjects. Granzyme A expression by CD56+ T-cells was not significantly different (P = 0.3053) in CMV seropositive than seronegative subjects. Moreover, perforin expression by CD56-
T-cells was not significantly different (P = 0.1830) in CMV seropositive than seronegative subjects.
0.5976 0.3053 0.1830 100
80
60 % 40
20
0 - + - + - + NK CD56+ T-cells CD56- T-cells
- 175 - 4.1.3.6 Granzyme B expression following stimulation with CMV lysate
Granzyme B expression by NK cells, CD56+ T-cells and CD56- T-cells was tested after cells were stimulated by CMV lysate. The levels of expression are illustrated by density plots in Figure 4.12.
Granzyme B expressed by NK cells was not significantly different (P = 0.1416) in
CMV seropositive (78.35% ±3.897, N=8) than seronegative subjects (68.33%
±4.356, N=7). Granzyme B expression by CD56+ T-cells was significantly higher
(P = 0.0015) in CMV seropositive (65.66% ±3.034, N=8) than seronegative subjects (24.11% ±7.468, N=7). Moreover, Granzyme B expression by CD56- T- cells was significantly higher (P = 0.0037) in CMV seropositive (26.53% ±2.038,
N=8) than seronegative subjects (15.09% ±1.422, N=7), these results are illustrated in Figure 4.13.
- 176 - Figure 4.12 Levels of Granzyme B expression. In a representative NK cells,
CD56+ T-cells and CD56- T-cells from representative subjects after overnight stimulation with CMV lysate
Seronegative Seropositive
- 177 -
Figure 4.13 Levels of Granzyme B expression in CMV + and CMV- subjects.
Paired T-test was used to compare levels of Granzyme B expression by NK cells,
CD56+ T-cells and CD56- T-cells after overnight stimulation with CMV lysate.
Granzyme B expression by NK cells was not significantly different (P = 0.1416) in CMV seropositive than seronegative subjects. Granzyme B expression by
CD56+ T-cells was significantly higher (P = 0.0015) in CMV seropositive than seronegative subjects. Moreover, perforin expression by CD56- T-cells was also significantly higher (P = 0.0037) in CMV seropositive than seronegative subjects.
0.1416 0.0015 0.0037 100
80
60 % 40
20
- + - + - + NK CD56+ T-cells CD56- T-cells
- 178 -
4.2 Degranulation assay of CD56+ T-cells; detection by LAMP-1 (CD107a) expression
CD107a is a marker for NK cell and CD8+ T-cell degranulation as part of the perforin-dependent cytotoxic process therefore it was investigated whether
CD56+ T-cells express this marker after their induction by CMV lysate and the standard NK cell target K562, and if there were any significant differences between healthy CMV seropositive and seronegative subjects.
4.2.1 Aim
To assess levels of CD107a surface expression in NK, CD56+ T-cells and CD56-
T-cells in response to exposure to K562 and CMV lysate in CMV seropositive and seronegative healthy subjects.
4.2.2 Cell surface exposure of CD107a in response to stimulation
PBMCs were stimulated with K562 or CMV lysate in culture medium in a
o humidified 5% CO2 atmosphere at 37 C, following that the cells were labelled with anti-CD3, anti-CD56 and anti-CD107a. The levels of CD107a expression on the cells after exposure to K562 are illustrated in figure 4.14 and after exposure to
CMV lysate in figure 4.16.
- 179 - In Figures 4.15 and 4.17 proportions of cells expressing CD107a were compared between CMV seropositive and seronegative healthy subjects after exposure to
CMV lysate and K562.
When PBMCs were stimulated by K562 cells, levels of CD107a expression were found to be significantly more in NK cells (P = 0.0061) and CD56+ T-cells (P =
0.0202) in seropositive (15.00% ±2.653, 12.26% ±1.501 respectively) than seronegative (8.006% ±1.567, 7.186% ±1.138 respectively) healthy subjects after stimulation with K562 cells. However, CD56- T-cells did not express high levels of CD107a and the comparison between both CMV groups was non-significant (P
= 0.7089) (seronegative 0.7722% ±0.2411, seropositive 0.76% ±0.1676). The number of participants was for NK cells 11 seronegative and 15 seropositive,
CD56+ T-cells 12 seronegative and 12 seropositive, and for CD56- T-cells 9 seronegative and 18 seropositive
- 180 - .
Seronegative Seropositive
Figure 4.14 Levels of CD107a expression after PBMCs pulsed with K562.
Levels of CD107a expression on NK cells, CD56+ T-cells and CD56- T-cells from healthy subjects after exposure to K562 cells.
- 181 - 0.0061 0.0202 50
40
30 0.5297 20
% of CD107a 10
0 - + - + - + CD3-CD56+ CD3+CD56+ CD3+CD56-
Figure 4.15 Levels of CD107a expression after PBMCs pulsed with K562 in
CMV + and CMV- subjects. CD107a levels of expression were found to be significantly more in NK (P = 0.0061) and CD56+ T-cells (P = 0.0202) in CMV seropositive than seronegative healthy subjects after stimulation with K562 cells.
However, CD56- T-cells did not express high levels of CD107a and the comparison between both CMV groups was non-significant (P = 0.7089).
When PBMCs were stimulated by CMV lysate CD107a positive cells were found to be significantly more in NK (P = 0.0236) and CD56+ T-cells (P = 0.0376) in seropositive (5.742% ±1.185, 4.757% ±0.6900 respectively) than seronegative
(1.963% ±0.4527, 2.353% ±0.3082 respectively) healthy subjects after stimulation with CMV lysate. However, very few CD56- T-cells express high levels of
CD107a and the comparison between both CMV groups was non-significant (P =
0.1392) (seronegative 0.5750% ±0.1493, seropositive 1.467% ±0.4072). Numbers of healthy participants was 6 CMV seropositive and 6 seronegative subjects.
- 182 - Seronegative Seropositive
Figure 4.16 Levels of CD107a expression after PBMCs stimulated with CMV.
Representative plots of proportions of cells expressing CD107a on NK cells,
CD56+ T-cells and CD56- T-cells from healthy subjects after exposure to CMV lysate.
- 183 - 15 0.0236 0.0376
10 0.1392
5
% of CD107a %
0 - + - + - + CD3-CD56+ CD3+CD56+ CD3+CD56-
Figure 4.17 Levels of CD107a expression after PBMCs stimulated with CMV in CMV + and CMV- subjects. Proportions of cells expressing CD107a were found to be significantly more in NK (P = 0.0236) and CD56+ T-cells (P =
0.0376) in seropositive than seronegative healthy subjects after stimulation with
CMV lysate. However, CD56- T-cells did not express high levels of CD107a and the comparison between both CMV groups was non-significant (P = 0.1392).
Numbers of healthy participants was 6 CMV seropositive and 6 seronegative subjects.
- 184 - 4.3 Proliferative responses of CD56+ T cells to CMV antigens
PBMCs from CMV seropositive healthy donors were labelled with CFSE according to section 2.4.2.4.2. CSFE labelled PBMCs were incubated for 7 days in 48 wells culture plate in culture medium in a humidified 5% CO2 atmosphere at
37oC. PBMCs were incubated at three different conditions; PBMCs un-stimulated,
PBMCs with CMV lysate and PBMCs with SEB as a positive control. Cells proliferation was calculated for three different groups, NK cells, CD56+ T-cells and CD56- T-cells.
4.3.1 Aims
To detect the proliferative response of CD56+ T-cells from CMV seropositive subjects in response to stimulation with CMV.
4.3.2 Proliferative responses of CD56+ T cells to CMV antigens
Percentages of proliferated CD56+ T-cells at day 7 of the incubation period in response to CMV was compared by using a Paired T-test compared to un- stimulated cells. Representative density plots illustrating the percentage of proliferated cells are presented in Figure 4.18. Moreover, the percentage proliferation was compared in Figure 4.19. The percentage of proliferated cells in un-stimulated and CMV stimulated wells from the 5 CMV seropositive donors was compared by Paired T-test. Differences in NK cell proliferation did not quite reach statistical significance (P = 0.059) (seronegative 4.804% ±1.614, seropositive16.76% ±3.460). CD56+ T-cells (P = 0.0173) (seronegative 4.860%
- 185 - ±2.319, seropositive 21.32% ±5.539) and CD56- T-cells (P = 0.0203)
(seronegative 6.7% ±2.239, seropositive 21% ±4.170) proliferated significantly more in response to CMV than un-stimulated cells.
Figure 4.18 Representative plots of CFSE labelled PBMCs. PBMCs were stained by anti-CD56 and anti-CD3 to identify CD56+ T-cells, NK cells and
CD56- T-cells. CD56+ T-cells proliferated by 1.9% in the un-stimulated well,
7.1% in response to CMV lysate and 70.3% in response to SEB. NK cells proliferated by 2.5% in the un-stimulated well, 12.5% in response to CMV lysate and 52.1% in response to SEB. Last CD56- T-cells (CD3+CD56-) proliferated by
2% in the un-stimulated well, 15% in response to CMV lysate and 70.7% in response to SEB. Numbers of healthy CMV+ participants was 5.
- 186 -
Figure 4.19 The percentage of proliferated cells. Levels compared between un-stimulated and CMV stimulated wells from the 5 CMV seropositive donors were compared by Paired T-test. CD56+ T-cells (P = 0.0173) and CD56- T-cells
(P = 0.0203) significantly proliferated in response to CMV compared to un- stimulated cells.
- 187 - 4.4 Antigen specificity of CD8+CD56+ T cells in CMV+ subjects
Healthy CMV seropositive subjects were tissue typed for HLA-A2, -B7 and B8 according to section 2.3.2.5.1. Following that, HLA Pro5 Pentamers comprising these three common HLA tissue types loaded with immuno-dominant 9mer or
10mer CMV peptides (illustrated in Table 2.5) were used.
4.4.1 Aims
To study CD56+ T-cell (CD8bright+ CD56+) CMV antigen specificity toward CMV compared to CD8+ CD56- T-cells.
4.4.2 CD56+ T-cells Pro5 pentamer staining
PBMCs from CMV+ volunteers were labelled by Pro5 pentamers according to section 2.3.2.5.2. Lymphocytes were gated for CD8bright+ cells and both CD56+ and CD56- populations analysed for HLA pentamer staining. Pentamer-specific
CD8+ cells were detected in CD56+ and CD56- cells. Examples of Pentamer- labelled CD8+brightCD56+ T-cells and CD8+CD56- from candidates positive for either HLA-A2, -B7 and –B8 are illustrated in Figure 4.20.
The percentage of pentamer labelled CD8+brightCD56+ T-cells and CD8+ CD56- cells were compared by Paired T-test and the results are illustrated in Figure 4.21.
- 188 -
Figure 4.20 Representative plots of Pro5 CMV pentamer-labelleing. Both
CD8bright+ CD56+ and CD8bright+ CD56- cells were targeted in subjects positive for one of three different class I HLA types A2, B7 and B8. Some CD56+ T-cells showed specificity toward CMV peptides loaded on the pentamers, which can be visualised by the percentage of CD8+ CD56+ cells.
- 189 -
Figure 4.21 Comparison of the percentage of pentamer labelled cells in
CD56- and CD56+ T cells. Pentamer staining was compared between CD8 bright+CD56- and CD8bright+CD56+ T-cells. The lines join results from the same subjects. HLA-A2 and B7 pentamers labelled similar percentages of cells between
CD8bright+CD56- and CD8+bright+CD56+ T-cells. However, HLA-B8 pentamer labelled more CD8bright+CD56+ T-cells than CD8 bright+CD56- cells. No significant differenece detected.
4.5 Discussion
In Chapter 3, expansion of CD56+ T-cells in CMV seropositive healthy donors was detected. Also, these cells were shown to express memory surface markers in
CMV seropositive subjects while in CMV seronegative donors were more likely to express naive surface markers. Overall, this study has provided evidence that
CD56+ T-cells have dual functions where they preserved TCR specificity against
CMV and expanded in response to CMV stimulation while also performing NK- like cytotoxicity against MHC negative K562 cells. Additionally, this ex-vivo study aimed to analyse CD56+ T-cells’ potential immunological functions in response to CMV in CMV seropositive and seronegative PBMCs and to compare the results between them. Several experiments studied their capability of
- 190 - expressing intracellular cytokines IFN-γ, TNF-α and IL-2 and the cytotoxic mediators Perforin, Granzyme A and B. CD107a surface expression was also studied following stimulation with CMV lysate and K562. Moreover, CFSE labelling was used to detect proliferation in response to CMV and the polyclonal stimulator SEB. To study these functional properties, PBMCs from both sero- groups were incubated with CMV lysate and SEB as a positive control overnight.
Isotype and un-stimulated PBMCs were applied to remove any formed background. Paired t-tests were used to compare the mean of both sero-groups and the results were displayed as scatter plots. Also, the cells’ specificity to CMV was investigated by labelling the cells with Pro5 CMV Pentamers. The remaining possibility of cells response can be due to other part of the host cells (fibroblast) where CMV lysate were extracted.
CMV is life-long and latent virus and even though it is pathologically not risky in healthy seropositive subjects, however, it has been reported to disrupt the immune system and results in high levels of cytokine expression (Dunn et al. 2002).
CD56+ T-cells responded to CMV lysate by producing significantly more IFN-γ,
TNF-α, Granzyme B in seropositive than seronegative healthy donors. However,
CD56+ T-cell production of IL-2 and expression of Perforin and Granzyme A, when compared between sero-groups, were not significantly different and these results are illustrated in Table 4.1. The phenotype of CD56+ T-cells in CMV seropositive healthy subjects revealed memory characteristics therefore the significantly elevated proportions of cells expressing IFN-γ, TNF-α and granzyme
B can be argued as evidence of an immunological memory response of CD56+ T- cells against CMV. Even though perforin and granzyme A have not shown any
- 191 - significant variation from CMV seronegative donors however the levels are less than what have been detected in seropositive donors.
NK CD56+ T-cells CD56- T-cells
- + - + - +
IFN-γ 2.332 5.937* 2.585 5.482* 1.976 3.483*
TNF-α 2.264 3.008 1.936 3.912* 1.369 1.926
IL-2 1.143 1.303 1.528 2.170 1.401 0.6831
Perforin 90.06 90.43 54.90 71.49 30.03 43.44
Granzyme A 38.44 47.43 25.26 49.25 6.343 19.98
Granzyme B 68.33 78.53 24.11 65.66* 15.09 26.53*
Table 4.1 Cytokine and cytotoxic mediator detection. This table illustrates the mean proportions of cytokine and cytotoxic mediator expressing NK, CD56+ T- cells and CD56- T-cells in response to overnight stimulation with CMV lysate. (
* ) = Significantly more than seronegative group. + = CMV seropositive and - = seronegative.
- 192 - Among the differences between healthy subjects of different CMV sero-status were that a higher proportion of CD56+ T-cells produced IFN-γ in CMV+ subjects, the same as was found for NK cells and CD56- T-cells. Moreover, for
TNF-α significant production by CD56+ T-cells contrasted with data for NK cells and CD56- T-cells which showed no significant differences between both CMV sero-statuses. In addition, granzyme-B was expressed by a significantly higher proportion of both CD56+ CD56- T-cells from CMV+ subjects while NK cells showed no significant differences between both CMV sero-statuses.
CD56+ T-cell production of significant levels of IFN-γ has also been reported in other studies. CD56+ T-cells were reported to produce Th1 cytokines when stimulated non-specifically using PMA (Doherty et al. 1999). This was also found in patients with hepatitis B-associated liver disease (Barnaba et al. 1994).
CD56+T-cells were reported to produce significantly more IFN-γ when stimulated with K562 in chronic HIV-infected individuals and long-term non-progressor HIV infected patients than healthy donors (Y. Jiang et al. 2014). Significant levels of
TNF-α were also reported in CD56+ T-cells isolated from the human liver when they were stimulated by PMA and ionomycin (Doherty et al. 1999).
TEMRA and TEM both pre-store the serine protease Granzyme B and the pore- forming perforin, however this normally does not occur in TCM and naive cells
(Sallusto et al. 2004). Granzyme B was present in significantly more CD56+ T- cells and CD56- T-cells from CMV+ subjects, while perforin was not expressed by significantly more cells in CMV seropositive than seronegative subjects. This finding contrasts with a study that mentioned reduced levels of granzyme B and perforin in PBMCs of smokers and chronic obstructive pulmonary disease
(COPD) patients, however this study also reported reduced numbers of CD56+ T-
- 193 - cells in those individuals (Urbanowicz et al. 2009). However, this group have also measured CD56+ T-cells in induced sputum of both smokers and COPD patients and reported a significant increase in numbers and cytotoxicity of CD56+ T-cells in COPD patients than both smokers and healthy non-smokers; also smokers have significant more CD56+ T-cells than healthy non-smokers. Moreover, levels of granzyme B and perforin were significantly more in COPD patients than smokers and healthy non-smokers, also smokers have significant more granzyme B and perforin than healthy non-smokers (Urbanowicz et al. 2010). These increases may result from the greater incidence of pulmonary infections in COPD patients and smokers. CD56+ expression on CD8+ cells has been correlated with CD8+ effector functions and the cells were found to be enriched with perforin and granzyme B (Pittet et al. 2000, Santin et al. 2001).
CD107a or LAMP-1 is a marker for NK cell and CD8+ T-cell degranulation as part of the perforin-dependent cytotoxic pathway; CD107a can work as an activation marker as upon stimulation CD107a is transported to the surface of the cells. Significantly greater proportions of CD56+ T-cells and NK cells were found to express CD107a upon stimulation with K562 and CMV lysate in CMV seropositive than seronegative healthy donors. Increased expression of CD107a was also reported in CD56+ T-cells when stimulated by K562 and
PMA/ionomycin in chronic HIV-infected individuals and HIV long-lerm non- progressors (Y. Jiang et al. 2014). Moreover, the same was reported when hepatic
CD56+ T-cells collected from liver graft perfusates were stimulated with K562 and the SNU398 human hepatocellular carcinoma cell line leading to significantly greater expression of CD107a than in healthy PBMCs (Kim et al. 2013). In this study CD56- T-cells failed to express significant levels of CD107a which was also
- 194 - reported in a study on CD3+CD8+ levels of CD107a in response to CMV in females with recurrent abortions (Tarokhian et al. 2014).
PBMCs were labelled by CFSE and incubated for 7 days with CMV antigens and
SEB as positive control to detect the proliferation of NK cells, CD56+ T-cells and
CD56- T-cells. Levels of proliferated NK cells, CD56+ T-cells and CD56- T-cells were significantly more than un-stimulated cells. TEMRA have a defect in their proliferative capacity, however, CD56+ T-cells proliferated in response to CMV and SEB. This contrasts with a study which showed that CD56+ T-cells generated in vitro have no proliferative capacity (Franceschetti et al. 2009). In the present study, CD56+ T-cells proliferated to a similar extent as CD56- T cells in response to CMV antigens so it is clear that they do not lose their capability to proliferate in response to viral antigens as what have been reported in a study illustrated that T-
EMRA proliferate to some levels (Geginat et al. 2003).
CD8bright+CD56+ T-cells’ antigenic specificity was studied towards three common immuno-dominant 9mer or 10mer CMV peptides in healthy subjects of the appropriate HLA types. This was to reveal if TCR expressed by CD8bright+CD56+
T-cells can mediate specific binding and recognition towards CMV common peptides, and to compare their immuno-specificity with that of CD8+CD56- T- cells. The labelling with HLA-A*02:01-NLV that represents HCMV pp65 495-
503 showed no significant difference between CD8bright+CD56+ T-cells and
CD56-CD8 bright+ T-cells, the specificity of CD8bright+CD56+ T-cells and CD56-
CD8bright+ T-cells were both ranging between 0.2-0.3% up to 7.1%. Moreover, staining with a pentamer for HLA-B*07:02-TPR which is HCMV pp65 417-426 showed no significant difference between CD8bright+CD56+ T-cells and CD56-
CD8bright+ T-cells, the specificity of CD8bright+CD56+ T-cells was ranging from
- 195 - 0.5% up to 1.9% and CD56-CD8bright+ T-cells was ranging from 0.5% up to 2.5%.
Staining with the pentamer HLA-B*08:01-QIK 88-96 which is HCMV IE1 showed no significant difference between CD8bright+CD56+ T-cells and CD56-
CD8bright+ T-cells, the specificity of CD8bright+CD56+ T-cells was ranging from
1.9% up to 15.1% and for CD56-CD8bright+ T-cells was from 1.5% up to 6.2%.
CD8bright+CD56+ T-cells showed higher specificity toward QIK-HLA-B*08:01 than CD56-CD8bright+ T-cells, and this was also higher than both pp65 peptides
NLV-HLA-A*02:01 and TPR-HLA-B*07:02. The expression of CD56 (NCAM) has been correlated with anti-CD3 induced cytotoxic functions of CD8+ T cells in healthy subject (Pittet et al. 2000), and in human papilloma virus specific CD8+
T-cells (Santin et al. 2001). Moreover, these cells have been found to be effectors in protection against HIV, hepatitis C virus and Japanese encephalitis virus
(Sooryanarain et al. 2012, Hou et al. 2012, Doskali et al. 2011) as well as the autoimmune inflammatory condition Behçet’s disease (Ahn et al. 2005). Also,
CD4+CD28-CD56+ cells were expanded in patients with chronic hepatitis B virus infection (Wang et al. 2009) and mast cell stimulation by reovirus was found to lead to recruitment of CD56+ T cells in vitro (McAlpine et al. 2012). Overall, the results of Chapter 4 support the concept that CD56+ T cells are an important component of the anti-CMV response in healthy subjects.
- 196 -
Chapter 5
Frequency and phenotype of
CD56+ T-cells in CMV seropositive and seronegative Renal Transplant
Patients
- 197 -
5. Frequency and phenotype of CD56+ T-cells in CMV seropositive and seronegative Renal Transplant Patients
5.1 Introduction
CMV replication leading to viraemia is one of the major clinical infections leading to rejection, morbidity and mortality in kidney graft recipients. To prevent these complications the recipients are given prophylactic therapy where they receive antiviral medications such as valganciclovir (De Keyzer et al. 2011).
All patients studied had received a renal transplant at the Royal Liverpool and
Broadgreen University Hospital Trust Transplant Unit. They were receiving maintenance immunosuppression comprising tacrolimus or, in the case of older patients, cyclosporine. All patients are monitored regularly for post-transplant
CMV infection using standard PCR techniques. Since 2008, all patients who were
CMV- are given a kidney from a CMV+ donor and also any patients given
Campath 1 induction were given prophylactic treatment with valganciclovir for 90 days post transplantation. Prior to 2008, pre-emptive antiviral therapy was given to patients in whom CMV became detectable by PCR.
The risk of CMV viremia in transplant patients is correlated with the CMV sero- status of the donor and recipient. Moreover, the high risk group are those with D
(donors) CMV positive and recipients (R) CMV negative. Recognition of alloantigen by T-cells either directly or through indirect allo-recognition of donor
- 198 - peptide leads to activation of T-cells toward the graft and the expansion of allo- specific T-cells toward the graft, increasing the risk of acute or chronic rejection
(Issa et al. 2010).
5.2 Aim
The aim of this section of the study was to compare the relative levels and phenotype of CD56+ T-cells in Kidney transplant patients (KTPs) and to compare the results to what has been found in healthy donors.
5.3 Results
5.3.1 Frequency of CD56+ T-cells in CMV seropositive and seronegative renal transplant patients
Flow cytometric analysis of CD56+ T-cell populations was applied to identify the relative numbers of these cells in kidney transplant patients (KTP). PBMCs were stained with anti-CD3 and anti-CD56 and expressed as a density plot in the same way as in Figure 3.1. Numbers were expressed as percentages of both total lymphocytes and T-cells and were analysed using GraphPad prism; the CMV+ groups were compared with the D-/R- group by using a paired T-test. In Figure
5.1, the percentage of CD56+ T-cells was calculated from total lymphocytes and
T-cells. When the mean value of CD56+ T-cells from total lymphocytes was calculated, the D+R- patient group (mean 5.42% ±1.01) had significantly
(P=0.0247) more than in the D-R- group (mean 1.9% ±0.35). Moreover, in the R+ group (mean 5.11% ±0.69) levels were significantly higher (P < 0.0001) than in the D-R- group.
- 199 - When the CD56+ T-cell percentage was calculated from total CD3+ T-cells, the mean value of CD56+ T-cells from total CD3+ T-cells for D+R- patients (mean
9.74% ±1.12) was significantly (P=0.0037) more than for D-R- patients (mean
4.91% ±0.76). Moreover, levels for the R+ group (mean 9.72% ±0.97) were significantly (P=0.0003) higher than for the D-R- group.
0.0247 0.0037 < 0.0001 0.0003 20 25
15 20
15 10
10 % T-cells 5 % Lymphocytes 5
0 0 D-R- D+R- R+ D-R- D+R- R+ CD56+ T-cells CD56+ T-cells
Figure 5.1 Scatter plots comparing the percentage of CD56+ T-cells between different KTP groups. These scatter plots compare the percentage of CD56+ T- cells obtained from total lymphocytes and T-cells between different KTP groups,
D-R-, D+R- and R+. Clearly, D+R- (n=26) and R+ (n=26) KTP have significantly more CD56+ T-cells than D-R- (n=20) when CD56+ T-cells where calculated from total lymphocytes (P=0.0247 and < 0.0001 respectively) and when calculated from total CD3+ cells (P=0.0037 and 0.0003 respectively).
5.3.2 CD56+ T-cell levels during CMV viraemia
- 200 - Levels of CD56+ T-cells were investigated and compared between KTPs with or without CMV viraemia, as detected by the Medical Microbiology Laboratory,
Royal Liverpool University Hospital. In Figure 5.2, CD56+ T-cells from KTPs who have had a previous episode of viremia were compared to the levels of the cells in KTPs who have never had viraemia. When all patients were analysed together, those who had one or more episodes of viraemia had significantly more
CD56+ T cells than those who did not. Figure 5.3 shows levels of CD56+ T-cells in viraemic D+R- KTPs compared to non-viraemic D+R- KTPs. In those eight patients sampled after the episode of viraemia (pre-viraemia) levels were increased compared to patients without viraemia and this was statistically significant (P = 0.024). In the six D+/R- patients who had a viraemic episode after sampling (post-viraemia) CD56+ T cell levels were not significantly different from those in the group without viraemia (Fig. 5.3); no significant differences were found between other KTP groups.
25 0.0030
20
15 % 10
5
0 no viraemia viraemia CD56+ T-cells
Figure 5.2 KTPs studied who had an episode of viraemia. All KTPs studied who had an episode of viraemia (5.2% ±0.99) had significantly more CD56+ T-
- 201 - cells than KTPs without viraemia (4.3% ± 0.58; P=0.003).
D+R- 0.0351 25 0.0240 0.1798 20
15 % 10
5
0 no viraemia viraemia Pre-viraemia Post-Viramia CD56+ T-cells
Figure 5.3 D+R- patients studied who had an episode of viraemia. D+R- KTPs who had one or more episodes of viraemia (8.13% ±1.63) and pre-viraemia
(9.92% ±2.1) had significantly more CD56+ T-cells than KTPs without viraemia
(3.22% ± 1.20; P=0.03).
5.3.3 Relative numbers of CD56-CD3+ cells in KTPs
Relative numbers of total CD56- T-cells in KTPs were also evaluated and compared. CD3+ T-cells were compared by two different methods, the first method was to calculate CD3+ T-cells from the total lymphocytes and the second method was to calculate CD3+ T-cells from total cells expressing CD3 (Figure
5.4). No clear differences were observed between any of the groups. Frequency of
CD56- T-cells was compared between KTP groups. When evaluated from total lymphocytes the mean of CD56- T-cells in D-R- (N=20) was 42.49± 5.4, which
- 202 - was less than the mean of CD56- T-cells in D+R- (N=26) of 46.54% ± 4.85, although this was not quite statistically significant (P = 0.276). Moreover, The R+ group (N=26) at 49.10% ± 3.05 was at a similar level as the D-R- group
(P=0.3825). When CD56- T-cells were evaluated from the total T-cells, the mean of CD56- T-cells in the D-R- group was 94.34% ±0.95 which was greater than the mean of CD56- T-cells in the D+R- (N=26) of 91.34% ± 1.079, although this was not statistically significant (P=0.52). Moreover, the R+ group (N=26) at 90%
±0.95 was at a similar level as the D-R- group (N=20; P=0.24).
0.2760 0.5206 0.2438 100 0.3825 100
80 95
60 90
40 85 % T-cells
% Lymphocytes % 20 80
0 75 D-R- D+R- R+ D-R- D+R- R+ CD56- T-cells CD56- T-cells
Figure 5.4 Scatter plots comparing the percentage of CD56- T-cells between different KTP groups. Frequency of CD56- T-cells was compared between KTP groups. In the left plot, CD56- T-cells were evaluated from the total lymphocytes, the mean of CD56- T-cells in D-R- was less than the mean of CD56- T-cells in
D+R- and this was not quite statistically significant (P = 0.276). Moreover, The mean of R+ group was at a similar level as the D-R- group (P=0.3825). In the right plot, CD56- T-cells were evaluated from the total T-cells, the mean of
CD56- T-cells in the D-R- group was greater than the mean of CD56- T-cells in
- 203 - the D+R-, although this was not statistically significant (P=0.52). Moreover, the mean of R+ group was less than the D-R- group (P=0.2438).
5.3.4 Relative numbers of CD56+CD3- NK cells in KTPs
Relative numbers of total CD56+CD3- NK cells in KTPs were also evaluated and compared. NK cells were found to be present in roughly equal proportions in all three groups (Figure 5.5) which did not differ significantly from CMV seronegative and CMV seropositive healthy individuals (Figure 3.4).
0.2267 0.5113 50
40
30
20
% Lymphocytes % 10
0 D-R- D+R- R+ NK cells
Figure 5.5 Frequency of CD56+CD3-NK cells in KTPs. This figure shows the frequency of CD56+CD3-NK cells in KTPs. There were no significant differences between numbers of NK cells in any of the three groups. P values between groups are shown.
5.4 Characterisation of the phenotype of CD56+ T-cells in KTPs
Chapter 3 of this thesis investigated CD56+ T-cells in healthy CMV seropositive and seronegative donors. Many CD56+ T-cells were shown to express a memory
- 204 - phenotype in CMV seropositive healthy donors and in seronegative healthy donors most cells expressed naïve cell markers. In this section, CD56+ T cells were phenotyped in KTP groups D-R-, D+R- and R+.
5.4.1 Aims
The aims of this section of the study were to analyse and compare the phenotype of CD56+ T-cells in KTPs according to CMV status and also to compare the results with the CD56+ T cell phenotype in healthy CMV+ and CMV- donors.
5.4.2 Results
5.4.2.1 Expression of CD4 and CD8 by CD56+ T-cells in KTPs
PBMCs from KTPs were stained with three colours in order to phenotype CD56+
T-cells for their expression of CD4+ and CD8+. In Figure 5.6 the percentage of
CD4+ and CD8+ cells within the CD56+ T-cell population in KTPs groups is shown. In the D-/R- group, most CD56+ T cells were CD4-8- whereas in both
D+/R- and R+ groups almost all were either CD8+ or CD4+. This is quite similar to the pattern seen in healthy CMV- and CMV+ subjects. To determine if there were any significant differences between the surface expression of CD8 and CD4 on CD56+ T-cells in the different KTP groups the proportions were compared by paired t-test.
The percentage of CD8+ and CD4+ CD56+ T-cells was compared between KTP groups. CD8+ CD56+ T-cells were significantly more in D+R- (59.25% ± 6.36)
- 205 - and R+ (60.77% ± 4.55) than D-R- (26.32% ± 1.89; P=0.0011 and 0.0002 respectively). CD4+ CD56+ T-cells were significantly more in D+R- (42.83% ±
3.77) and R+ (38.57% ± 4.72) than D-R- (24.36% ± 2.61; P=0.0008 and 0.0434 respectively). Moreover, CD8 was expressed on significantly more cells than CD4 on CD56+ T-cells in both D+R- (P=0.025) and R+ (P=0.0335) groups.
CD8+ CD4+ 0.0011 0.0008 100 0.0002 0.0434 0.0251 0.0335 80
60 % 40
20
0 D-R- D+R- R+ D-R- D+R- R+ CD56+ T-cells
Figure 5.6 The percentage of CD8+ and CD4+ CD56+ T-cells compared between KTP groups. CD8+ CD56+ T-cells were significantly more in D+R- and R+ than D-R- (P=0.0011 and 0.0002 respectively). CD4+ CD56+ T-cells were significantly more in D+R- and R+ than D-R- (P=0.0008 and 0.0434 respectively). Moreover, CD8 was expressed on significantly more cells than CD4 on CD56+ T-cells in both D+R- (P=0.025) and R+ (P=0.0335) groups.
5.4.2.2 CD56+ T-cells; naive and memory markers in KTPs
- 206 - CD56+ T-cells were stained for naive and memory markers in KTPs. The markers used were CD45RO, CD45RA, CD62L, CD28 and CD127. Levels of expression of these markers were compared between KTP groups to identify any significant differences between these groups and in Figure 5.7 the results are illustrated.
Proportions of CD4RO+ cells were significantly more in the D+R- group (44.07%
SE± 7.619, N=15) and R+ group (41.78% SE± 4.467, N=12) than the D-R- group
(18.60% SE± 2.146, N=14) (P 0.0117 and 0.0007 respectively). Mean proportions of CD45RA+ cells were significantly (P 0.0341) more in D-R- (88.07%
SE±1.562, N=10) than R+ patients (78.47% ±3.197, N=15), however, there was no significant difference between D-R- and D+R- groups (85.88% ±2.539).
Proportions of CD62L+ cells were significantly more in D-R- (72.36% ±3.592,
N=10) than in D+R- (55.69% ±2.544, N=15) and R+ patients (45.14% ±4.904,
N=14) (P 0.0079 and 0.0001 respectively). Proportions of CD28+ cells were significantly more in the D-R- group (58.72% ±7.835, N=10) than in D+R-
(22.65% ±5.682, N=13) and R+ patients (21.66% ±4.463, N=19) (P 0.0192 and
0.0001 respectively). Finally, proportions of CD127+ cells were significantly more in D-R- (64.6% ±4.173, N=10) than D+R- (31.99% ±5.113, N=17) and R+ patients (33.50% ±4.820, N=13) (P 0.0035 and 0.0005 respectively). These differences are similar to those found between CDM- and CMV+ healthy subjects.
- 207 - 0.0117 0.4234 0.0079 0.0192 0.0035 0.0007 0.0341 0.0051 0.0001 0.0005 100
80
60 % 40
20
0 D-R- D+R- R+ D-R- D+R- R+ D-R- D+R- R+ D-R- D+R- R+ D-R- D+R- R+ CD45RO+ CD45RA+ CD62L+ CD28+ CD127+ CD56+ T-cells
Figure 5.7 Memory and naive markers expressed by CD56+ T-cells in KTPs.
This dot plot illustrates differences in proportions of CD56+ T-cells expressing memory markers in different KTP groups. Bar indicate levels of significance between groups (t-test).
5.4.2.3 CD56+ T-cells; CD11a, NKG2C and CD161 in KTPs
Percentages of CD56+ T cells expressing CD11a, NKG2C and CD161 were evaluated by using three colour staining. The results are illustrated in Figure 5.8, showing percentage positive cells and mean fluorescence intensity of CD11a expression in each group. No significant difference was found in proportions of
CD11a+ cells or MFI in all KTP groups. Proportions of NKG2C+ cells were significantly more in D+R- (32.96% ±8.522, N=7) and R+ (10.28% ±1.732, N=8) than D-R- patients (3.070 ± 1.017, N=6). Proportions of CD161+ cells were significantly more in D-R- (29.24% ±5.230, N=11) than D+R- (10.96% ±2.608,
N=18) and R+ groups (9.553% ± 3.579, N=11).
- 208 - 0.4841 0.0178 0.0138 0.3978 0.0657 0.0124 0.0261 4000 0.2744 100
80 3000
60 2000
% MFI 40 1000 20
0 D-R- D+R- R+ D-R- D+R- R+ D-R- D+R- R+ 0 D-R- D+R- R+ CD11a+ NKG2C+ CD161+ CD11a+CD56+ T-cells CD56+ T-cells
Figure 5.8 CD11a, NKG2C and CD161 expression on CD56+ T-cells from
KTPs.This dot plot illustrates the expression of CD11a, NKG2C and CD161 on
CD56+ T-cells from KTPs. No significant difference was found in proportions of
CD11a+ cells (left) or MFI (right) in all KTP groups. Bars show levels of significance between groups (T-test).
5.4.2.4 CD56+ T-cells; CMV HLA Pentamer labelling
PBMCs from KTPs of appropriate HLA type were labelled with Pro5 pentamers according to section 2.3.2.5.2. Lymphocytes were gated for CD8bright+ cells and both CD56+ and CD56- populations analysed for HLA pentamer staining.
Pentamers-specific CD8+ cells were detected in both CD56+ and CD56- cell populations. Representative examples of pentamer-labelled CD8+brightCD56+ T- cells and CD8+CD56- from patients positive for either HLA-A2, -B7 or -B8 are illustrated in figure 5.9. A total of four KTPs who were of HLA-A2 tissue type
- 209 - were tested, three who were HLA-B7 but only one who was HLA-B8 was tested.
The percentage of pentamer labelled CD8+brightCD56+ T-cells and CD8+ CD56- cells were compared by paired T-test and the results are illustrated in figure 5.10.
However, sample sizes were too small for adequate statistical comparison.
Figure 5.9 Representative density plots of HLA-pentamer-labelleing. Both bright
CD8bright+ CD56+ and CD8bright+ CD56- cells from D+/R- renal transplant patients of three different HLA tissue types, HLA-A2, -B7 and -B8. Upper panels,
CD56+CD8+ cells, lower panels CD56-CD8+ cells.
- 210 -
Figure 5.10 Comparision of HLA-pentamer-labelled cells. The percentages of
HLA pentamer staining was compared between CD8bright+ CD56- and
CD8bright+CD56+ T-cells in KTPs. The lines join results from the same patients.
5.5 Discussion
CMV reactivation in immuno-suppressed KTPs can lead to serious medical consequences such as graft damage, dysfunction, rejection and death. The level of danger behind CMV reactivation is correlated with the CMV status type of donors and recipients, as the high-risk group is D+R- patients. The previous chapters have revealed significant expansions in numbers of CD56+ T-cells in healthy
CMV seropositive compared to sero-negative subjects, this expansion being associated with increased immuno-functional activities against CMV. This chapter aims to identify the characteristic phenotype of CD56+ T-cells in KTPs and study their antigenic specificity against three common HLA immuno- dominant 9mer or 10mer CMV peptides. The comparison between CD56+ T-cell numbers in healthy subjects and KTPs is illustrated in Figure 5.11.
Levels of CD56+ T-cells when calculated from total lymphocytes or T-cells showed D+R- patients had significantly more than in the D-R- group. Moreover, in the R+ group levels were significantly higher than in the D-R- group. CD56+
- 211 - T-cell mean levels were more in healthy seropositive subjects than seropositive
KTPs, also healthy seronegative subjects had significantly more CD56+ T-cells than the KTP D-R- group. This could be explained as KTPs are under immunosuppression leading to reduced numbers of immune-effector cells. As cells were quantified as ‘percent total lymphocytes or T cells’, without knowing absolute lymphocyte numbers it is not possible to assess whether total cell numbers were different in different patient and healthy groups. Regarding NK and
CD56- T-cells, no significant differences were found between KTP groups and healthy subjects, while CD56- T-cells when calculated as a percentage of total
CD3+ cells in healthy subjects showed significant variation.
- 212 - * * * 0.0018 20 0.0012
15 0.0043
% 10
5
0 CMV- CMV+ D-R- D+R- R+ Healthy KPTs CD56+ T-cells
Figure 5.11 Levels of CD56+ T-cells in healthy subjects and KTPs.This graph compares levels of CD56+ T-cells in healthy subjects and KTPs. The lowest mean level was found in D-R- KTPs. Proportions of CD56+ T-cells were significantly more in CMV seronegative healthy subjects than in D-R- KTPs (P=0.0043). Also, healthy CMV+ subjects had significantly higher proportions of CD56+ T-cells than D+R- and R+ KTP groups (P 0.0018 and 0.0012 respectively) * indicates significant P value as shown in Chapter 3 and 5 section 5.3.1.
CD56+ T-cell levels were significantly expanded more in KTPs with than without viraemia. This was noted in specifically in the D+R- group where the cells significantly expanded more in patients with viraemia and who had viraemia after sampling than without viraemia. However, differences in levels in patients who had viraemia before sampling was not quite statistically significant. CD8+ CD56+
T-cells had reduced expression of CD56 when HIV patients were treated with prolonged antiviral therapy, however, in patients who were not receiving this prolonged therapy, CD56 expression was restored along with high perforin
- 213 - content (Poonia and Pauza 2014). This finding together with what has been detected in this study indicates that CD56+ T-cells expand and are involved in antiviral activity to control viraemia.
Relative numbers of CD56+ T-cells in healthy subjects have illustrated higher proportions of CD8+ and CD4+ cells in seropositive than sero-negative subjects.
Generally, proportions of CD56+ T-cells expressing CD8 were significantly more than those expressing CD4. Similar results were also detected in KTPs where significantly more CD56+ T-cells expressed CD8 in D+R- and R+ than D-R- groups, also, CD4 was expressed by significantly more cells in D+R- and R+ groups than in the D-R- group. CMV-sepcific-CD8+ and CMV-specific-CD4+ T cells have been reported to have protective roles against CMV in solid organ transplantation; deficiency of these CMV-specific cells resulted in clinical complications in lung, renal and BMT transplantation (Riddell et al. 1992a, Sester et al. 2002, Sester et al. 2005).
The characteristic surface phenotype of CD56+ T cells in KTPs was similar to what has been found in healthy subjects; more CD56+ T-cells tend to express memory markers in CMV+ than D-R- patients. In the case of CD11a no significant differences in MFI were detected although the mean in D-R- patients was higher, but not significantly more than in D+R- and R+ KTPs which is in contrast to what has been detected in healthy groups. This may be a result of the continuing immunosuppression in KTPs which may prevent up-regulation of activation molecules in T cell populations.
It has been reported in renal transplant patients that CMV-specific-CD8+T-cells provide a protective role against CMV activation (Radha et al. 2005, Reusser et al.
- 214 - 1999). CD8bright+ CD56+ T-cells were evaluated for their TCR specificity toward
CMV lysate and compared to CD8bright+CD56- cells in KTPs. For HLA-B*08:01
QIK unfortunately only one KTP was tested. PBMCs from four KTPs were labelled with HLA-A*02:01-NLV that represents HCMV pp65495-504 and showed no significant differences between CD8bright+CD56+ T-cells and CD8bright+CD56-
T-cells, the percent of peptide-specific CD8+brightCD56- T-cells ranging from
0.2% up to 3.8%, and CD8+brightCD56+T-cells between 0.5% up to 1.7% which is higher, but not significantly, in CD8+brightCD56-T-cells. However, these levels are less than in healthy CMV+ subjects, although levels are highly variable. Three
KTPs were labelled with HLA-B*07:02-TPR which is HCMV pp65417-426 and showed no significant difference between CD8bright+CD56+ T-cells and
CD8bright+CD56- T-cells, moreover, CD8bright+CD56- T-cells range between 0.2% up to 6.5% while CD8bright+CD56+ T-cells 0.1% up to 3.9%, reflecting a higher, but not significantly higher, percentage in CD8+brightCD56-T-cells, however these tended to be higher for both cell types than in healthy CMV+ subjects. The only
HLA-B*08:01 KTP showed 3.1% CD8bright+CD56- T-cells and 1.57%
CD8bright+CD56+ T-cells, and it is reported that reactivation of CMV in solid organ transplant patients triggered the induction of CMV-immediate early gene transcription (Reinke et al. 1999). Moreover, this specificity for CMV-IE-QIK can enhance CMV reactivation as CD56+ T-cells produce TNF-α in response to CMV
(Figure 4.5) and TNF-α can play a role in reactivation of CMV. This occurs through TNF-α indirect stimulation of surface expression of MIEP (major IE enhancer/promoter) via the activation of NF-kB (De Keyzer et al. 2011).
Additionally, the role of MIEP activation of CMV is not well understood
(DeMeritt et al. 2004). However, others reported CD8+ T-cells specific for CMV-
- 215 - IE-1 peptides provide a protective role against reactivation in lung and heart transplantation (Bunde et al. 2005). These data identified that CD8bright+CD56+ T- cells possess specificity toward CMV in KTPs, the levels of specificity are variable in healthy subjects and KTPs.
This chapter identified expansion of CD56+ T-cells in CMV positive KTPs as well as healthy donors. Also, CD56+ T-cells were more expanded in D+/R- KTPs with viraemia than those without viraemia. CD56+ T-cells tended to express memory markers in CMV positive and naïve markers in CMV negative KTPs as was also shown in healthy donors. TCR specificity of CD56+ T-cells also showed binding of pentamers labelled with common CMV epitopes. CMV antigen- specific CD56+ T cells were not selectively enriched when compared to the majority CD56- T cell population but still made up a significant proportion of the antigen-specific CD8bright+ T cell population.
Chapters 3 and 4 showed increased numbers and function of CD56+ T cells in healthy CMV+ subjects. Additionally, out of 72 KTPs evaluated in this study 25 had post-transplant viraemia (35%), followed by significant expansion of CD56+
T-cells in their PBMCs. This also was significant when levels of CD56+ T-cells in the high risk group D+R- were tested. With many CD56+ T-cells in CMV positive
KTPs expressing memory markers and keeping their TCR specificity, this can indicate that CD56+ T-cells, particularly CD8+ cells, are involved in immune responses towards suppressing CMV reactivation in KTPs. Mononuclear cells, especially CD8+ and CD4+ T-cells, spread and surround the tubules in kidney transplants and these cells are loaded with perforin, granzyme A and B (Colvin
RB 2006, Cornell et al. 2008), also, they express Fas-L (Robertson et al. 1996,
Cornell et al. 2008). Moreover, effector cells during acute rejection produce high
- 216 - levels of IFN-γ and TNF-α (Hoffmann et al. 2005) similar to CD56+ T-cell cytokine responses to CMV antigens in healthy subjects as found in this study. In
D+/R- patients in whom donor and recipient are matched for class I HLA, there is the possibility that some of these cells represent CMV-specific cytotoxic T cells.
- 217 -
Chapter 6
Study to identify CD56+ T-cell gene expression profile in response to CMV
- 218 - 6. Study to identify CD56+ T-cell gene expression profile in response to CMV
6.1 Introduction
In Chapters 3-5, evidence was provided for increased numbers and levels of activation of CD56+ T cells in healthy CMV+ subjects. When stimulated with
CMV antigens, CD56+ T cells proliferated and showed increased levels of both proinflammatory cytokines and molecules involved in cytotoxic function. The aims of this chapter were to conduct a wider investigation into the response of purified CD56+ T cells from healthy CMV+ subjects to antigenic stimulation.
Microarrays were used to provide a comprehensive study of the gene expression profile of CD56+ T-cells in response to CMV stimulation. This is a hybridization method in which nucleic acids from stimulated and unstimulated cells are screened against a large set of oligonucleotide probes to identify changes in levels of gene expression. Moreover, SurePrint G3 Human Gene Expression
Microarrays provide a comprehensive coverage of genes and transcripts with the most up-to-date content. Moreover, SurePrint G3 Human Gene Expression 8x60K
MicroarrayKit has 96 x 10 ERCC control probes and 10 x 32 E1A spike-in control probes. Both these controls are incorbrated into microarrays to improve performance assessment for platforms of gene expression.
PBMCs from healthy CMV+ subjects were stimulated overnight by CMV lysate, cells were then labelled by conjugated monoclonal antibodies anti-CD3 and anti-
CD56 then washed and sorted to isolate CD56+ T-cells. Following this, RNA was extracted and analysed by microarrays for gene expression. The results for some genes were validated by qPCR and protein expression methods. This part of the study aimed to detect differences in up-regulated and down-regulated expressed
- 219 - genes in CD56+ T-cells incubated under two different conditions. It was decided that gene expression have to be more than two-fold changed; the fold change is calculated by dividing the two detected intensities to give a ratio that is log 2 transformed.
6.2 Aim
To identify changes in the pattern of genes expressed by enriched CD56+ T-cells when stimulated by CMV lysate using microarray analysis. The results will be validated first by qPCR and second by detection of protein expression.
6.3 Results
6.3.1 One-Colour Microarray-Based Gene Expression
Agilent Human v2 8x60K arrays were purchased from Agilent Technologies. The experimental work with the arrays was done by Dr. Lucille Rainbow, Centre for
Genomic Research, IIB. The analysis was done according to section 2.4.2.6.5 and the 8X slide after the visualization is presented in Fig. 6.1. Analysis was performed by using a moderated t-test and Westfall-Young multiple testing corrections with cut off of p<0.05 as explained in the manufacturer’s reference guide.
- 220 -
Figure 6.1 Microarrray chips. Image showing Agilent Human v2 8x 60K arrays chip after the hybridization of cy3 labelled RNA from purified CD56+ T cells to the probes of the slide. Samples from four healthy CMV+ subjects were tested following stimulation with CMV antigens (upper row) or without stimulation
(lower row). Colours were inverted inside the regions from black to white to clearly illustrate those regions.
Normalisation of baseline signal levels was carried out between samples. This transformation of the gene signals ensure those which have not significantly changed to be around the area of zero is presented in Figure 6.2. The normalization was performed to minimize differences from non-biological sources such as technical variations, different RNA concentrations, hybridization, differences between the 8 areas of the chip and labelling the samples with Cy3 colour.
- 221 -
Figure 6.2 Gene normalization. All gene intensities of the samples were rescaled to the same relative abundance level centering on zero. S represents the stimulated
CD56+ T-cells and U for un-stimulated CD56+ T-cells, Red for up-regulated while blue for down-regulated genes.
Overall, 131 genes passed the statistical tests; genes with values less than or equal to a 2 fold change (FC) were removed to emphasize the aim of only detecting highly significantly changed genes (Figure 6.3). Out of these 131 the controls and duplicates were excluded to identify a total of 106 genes whose levels were significantly changed. The results are shown in Table 6.1 to indicate the genes and the level of significant change in expression.
Several gene banks such as Gene Cards (www.genecards.org) were used to analyse these 106 genes to identify their functions, as shown in the appendix.
- 222 -
Figure 6.3 Significantly expressed genes using the filtration method. In the upper volcano plot 131 Entities (red colour) passed the P-value multiple testing correction cut-off of 0.05 and FC ˃ 2 when comparing S= stimulated cells vs. U= un-stimulated cells. The table illustrates a summary of the results of the comparison between S and U cells. Clearly, 9610 entities have passed FC ˃ 2 while only 131 entities have shown to have a P-value cut off < 0.05.
- 223 - Table 6.1 Detailed information about genes whose expression significantly changed. This table shows the details of genes whose levels of expression were significantly altered in CD56+ T cells stimulated with CMV antigens after analysing microarrays results using gene spring. Up-regulated in pink and down- regulated in light blue. Log FC = fold change, X = Unknown.
Gene Symbol Gene Name Probe Name p. value Log FC Ensemble ID
X Unknown A_24_P15502 2.24E-04 3.1195118 Control
X Unknown A_33_P3246995 0.001442 4.4135036 Control
APOL6 apolipoprotein L, 6 A_24_P941167 0.001453 2.2074013 ENST00000409652
BST2 bone marrow stromal cell A_23_P39465 0.001407 2.6559215 ENST00000533098 antigen 2 C1orf173 chromosome 1 open reading A_24_P522998 4.39E-04 5.088724 ENST00000433746 frame 173 CACNA1A calcium channel, voltage- A_33_P3275846 9.76E-04 3.5303764 ENST00000357018 dependent, P/Q type, alpha 1A subunit CD38 CD38 molecule A_23_P167328 2.10E-04 5.3847494 ENST00000540195
CIB2 calcium and integrin A_33_P3308914 5.46E-04 2.5931766 ENST00000539011 binding family member 2 CMPK2 cytidine monophosphate A_33_P3401826 3.58E-05 3.621748 ENST00000478738 (UMP-CMP) kinase 2, mitochondrial CNP 2',3'-cyclic nucleotide 3' A_23_P21838 0.001426 2.5997334 ENST00000393892 phosphodiesterase CPSF7 cleavage and A_33_P3309039 2.86E-04 3.7240167 ENST00000439958 polyadenylation specific factor 7, 59kDa CREM cAMP responsive element A_23_P201979 6.44E-04 3.1300871 ENST00000460270 modulator CYP2J2 cytochrome P450, family 2, A_23_P103486 5.78E-04 3.0702367 ENST00000469406 subfamily J, polypeptide 2 DDX58 DEAD (Asp-Glu-Ala-Asp) A_33_P3418170 3.34E-04 3.6301575 ENST00000379882 box polypeptide 58 DDX60 DEAD (Asp-Glu-Ala-Asp) A_23_P41470 2.82E-04 2.6578708 ENST00000393743 box polypeptide 60 DDX60L DEAD (Asp-Glu-Ala-Asp) A_23_P30069 7.00E-04 2.754306 ENST00000511577 box polypeptide 60-like DHX58 DEXH (Asp-Glu-X-His) A_23_P38346 3.01E-04 2.8720646 ENST00000423748 box polypeptide 58 ECE1 endothelin converting A_24_P396375 2.70E-04 3.0791154 ENST00000415912 enzyme 1 EIF2AK2 eukaryotic translation A_23_P142750 1.83E-04 3.1597533 ENST00000395127 initiation factor 2-alpha kinase 2 EPSTI1 epithelial stromal A_23_P105794 1.34E-04 2.8132715 ENST00000476830 interaction 1 (breast) ETV7 ets variant 7 A_23_P42353 6.35E-04 3.7648585 ENST00000339796
FRMD3 FERM domain containing 3 A_33_P3363425 9.14E-04 3.2037926 ENST00000304195
- 224 - FTSJD2 FtsJ methyltransferase A_23_P374149 0.001427 2.3990216 ENST00000491004 domain containing 2 GALM galactose mutarotase A_33_P3273436 8.58E-04 3.285547 ENST00000272252 (aldose 1-epimerase) GBP1P1 Guanylate Binding Protein 1, A_19_P00808588 9.75E-04 3.7079072 ENST00000513638 Interferon-Inducible Pseudogene 1 GBP3 guanylate binding protein 3 A_21_P0010536 0.00109 3.8230472 ENST00000461384
GBP4 guanylate binding protein 4 A_24_P45446 0.001079 2.820499 ENST00000471938
GCH1 GTP cyclohydrolase 1 A_33_P3311439 6.30E-04 3.102622 ENST00000395524
GCH1 GTP cyclohydrolase 1 A_24_P167642 2.98E-04 3.2402992 ENST00000254299
GMPR guanosine monophosphate A_24_P277657 4.36E-04 3.7887647 ENST00000540478 reductase HERC5 hect domain and RLD 5 A_23_P110196 5.90E-06 4.295132 ENST00000502913
HERC6 hect domain and RLD 6 A_23_P250358 6.59E-04 3.5208874 ENST00000511939
HES4 hairy and enhancer of split A_33_P3342628 2.23E-05 6.210292 ENST00000304952 4 (Drosophila) HRASLS2 HRAS-like suppressor 2 A_23_P105012 4.36E-04 2.7677798 ENST00000255695
IFI35 interferon-induced protein A_23_P152782 0.001172 2.7933698 ENST00000538473 35 IFI44 interferon-induced protein A_23_P23074 9.08E-04 3.1116233 ENST00000485662 44 IFI44L interferon-induced protein A_23_P45871 5.49E-05 3.8292685 ENST00000476521 44-like IFI6 interferon, alpha-inducible A_23_P201459 4.59E-05 4.0179195 ENST00000362020 protein 6 IFIH1 interferon induced with A_23_P68155 4.96E-05 3.7843661 ENST00000543192 helicase C domain 1 IFIT1 interferon-induced protein A_23_P52266 7.29E-05 4.3839293 ENST00000546318 with tetratricopeptide repeats 1 IFIT2 interferon-induced protein A_23_P24004 2.02E-05 4.486988 ENST00000371826 with tetratricopeptide repeats 2 IFIT2 interferon-induced protein A_24_P304071 2.14E-05 4.2640696 ENST00000371826 with tetratricopeptide repeats 2 IFIT3 interferon-induced protein A_33_P3283611 2.26E-05 4.9602613 ENST00000543062 with tetratricopeptide repeats 3 IFIT5 interferon-induced protein A_24_P30194 0.001185 2.4392035 ENST00000371795 with tetratricopeptide repeats 5 IFITM1 interferon induced A_23_P72737 0.001419 3.144174 ENST00000452428 transmembrane protein 1 (9-27) IFITM2 interferon induced A_24_P287043 0.001107 2.621654 ENST00000527146 transmembrane protein 2 (1-8D) IFITM4P interferon induced A_24_P16124 2.07E-04 3.1359797 ENSG00000130487 transmembrane protein 4 pseudogene IFNG interferon, gamma A_23_P151294 1.22E-07 6.0913982 ENST00000229135
IGFBP7 insulin-like growth factor A_23_P353035 9.03E-05 4.926652 ENST00000295666 binding protein 7 IL15RA interleukin 15 receptor, A_23_P138680 5.43E-04 5.288265 ENST00000379972 alpha
- 225 - IL2RA interleukin 2 receptor, alpha A_24_P230563 4.74E-04 2.9039214 ENST00000447847
IRG1 immunoresponsive 1 A_33_P3319925 3.60E-05 5.714017 ENST00000449753 homolog (mouse) ISG15 ISG15 ubiquitin-like A_23_P819 2.79E-06 5.015381 ENST00000379389 modifier ISG20 interferon stimulated A_23_P32404 6.85E-05 3.4611702 ENST00000546338 exonuclease gene 20kDa KLHDC7B kelch domain containing 7B A_24_P117410 0.001145 2.7272 ENST00000395676
KLHDC7B kelch domain containing 7B A_23_P6535 4.02E-04 2.5850372 ENST00000395676
LAG3 lymphocyte-activation gene A_23_P116942 7.74E-04 3.5425837 ENST00000203629 3 LAP3 leucine aminopeptidase 3 A_23_P18604 0.001421 3.3443398 ENST00000226299
LOC79015 uncharacterized LOC79015 A_23_P254181 7.04E-04 3.0117376 ENST00000415889
MOV10 Mov10, Moloney leukemia A_23_P161125 0.001265 2.8278618 ENST00000471160 virus 10, homolog (mouse) MT1B metallothionein 1B A_23_P37983 2.07E-04 3.828361 ENST00000334346
MT1L metallothionein 1L A_23_P427703 0.001424 2.4807048 ENSG00000260549 (gene/pseudogene) MX1 myxovirus (influenza virus) A_23_P17663 2.92E-04 3.1825523 ENST00000455164 resistance 1, interferon- inducible protein p78 (mouse) MX2 myxovirus (influenza virus) A_23_P6263 0.001207 2.9111667 ENST00000330714 resistance 2 (mouse) NEXN nexilin (F actin binding A_23_P200001 3.39E-04 3.1112146 ENST00000480732 protein) NT5C3 5'-nucleotidase, cytosolic III A_33_P3393836 3.74E-05 3.8463078 ENST00000409467
NT5C3 5'-nucleotidase, cytosolic III A_23_P59547 3.97E-05 3.4802923 ENST00000405342
OAS1 2'-5'-oligoadenylate A_23_P64828 7.94E-04 2.9598832 ENST00000377508 synthetase 1, 40/46kDa OAS2 2'-5'-oligoadenylate A_23_P204087 2.40E-04 2.9682827 ENST00000392583 synthetase 2, 69/71kDa OAS2 2'-5'-oligoadenylate A_33_P3225522 7.10E-04 3.9064493 ENST00000449768 synthetase 2, 69/71kDa OAS3 2'-5'-oligoadenylate A_33_P3402489 1.44E-04 3.648745 ENST00000228928 synthetase 3, 100kDa OAS3 2'-5'-oligoadenylate A_24_P335305 1.73E-05 4.2969465 ENST00000323881 synthetase 3, 100kDa OASL 2'-5'-oligoadenylate A_23_P139786 2.53E-05 3.466197 ENST00000257570 synthetase-like PARP10 poly (ADP-ribose) A_24_P340853 8.34E-04 2.2641857 ENST00000527262 polymerase family, member 10 PARP12 poly (ADP-ribose) A_23_P111804 0.001137 2.4642165 ENST00000488726 polymerase family, member 12 PARP9 poly (ADP-ribose) A_23_P69383 8.58E-04 2.6494117 ENST00000452457 polymerase family, member 9 PGAP1 post-GPI attachment to A_33_P3312735 4.22E-05 3.343388 ENST00000422382 proteins 1 PI4K2B phosphatidylinositol 4- A_23_P18598 4.27E-04 2.6377878 ENST00000537420 kinase type 2 beta PLSCR1 phospholipid scramblase 1 A_23_P69109 4.13E-04 3.2515664 ENST00000448205
PML promyelocytic leukemia A_23_P358944 0.001168 2.2871413 ENST00000435786
- 226 - PML promyelocytic leukemia A_23_P334664 9.43E-04 2.3369055 ENST00000395132
PML promyelocytic leukemia A_24_P207139 4.88E-04 3.865312 ENST00000268058
PNPT1 polyribonucleotide A_23_P154488 4.71E-05 3.2472057 ENST00000402246 nucleotidyltransferase 1 PRIC285 peroxisomal proliferator- A_33_P3400374 1.69E-04 2.9332774 ENST00000427522 activated receptor A interacting complex 285 RGS16 regulator of G-protein A_23_P217845 9.38E-04 2.9532652 ENST00000367558 signaling 16 RNF43 ring finger protein 43 A_33_P3213029 7.71E-04 3.0697901 ENST00000407977
RRAGC Ras-related GTP binding C A_23_P97623 0.001363 4.112468 ENST00000474456
RTP4 receptor (chemosensory) A_23_P166797 5.34E-05 3.0634089 ENST00000259030 transporter protein 4 SAMD9L sterile alpha motif domain A_33_P3264846 1.49E-04 3.316527 ENST00000394472 containing 9-like SLC27A2 solute carrier family 27 A_23_P140450 6.37E-04 2.9749262 ENST00000544960 (fatty acid transporter), member 2 SLC38A5 solute carrier family 38, A_23_P84929 3.03E-04 2.8301091 ENST00000462359 member 5 SOCS1 suppressor of cytokine A_23_P420196 3.66E-04 3.0243402 ENST00000332029 signaling 1 SP140 SP140 nuclear body protein A_33_P3268555 7.23E-04 2.5678482 ENST00000544128
STAG3 stromal antigen 3 A_33_P3317058 8.01E-04 2.9011881 ENST00000379577
TMEM140 transmembrane protein 140 A_24_P372134 0.001084 2.490767 ENST00000456488
TNFSF10 tumor necrosis factor A_23_P121253 4.23E-05 4.3497934 ENST00000241261 (ligand) superfamily, member 10 TNFSF10 tumor necrosis factor A_33_P3226810 8.06E-05 3.9121034 ENST00000241261 (ligand) superfamily, member 10 TRIM5 tripartite motif containing 5 A_23_P356526 5.88E-04 2.5912266 ENST00000465634
USP18 ubiquitin specific peptidase A_23_P132159 9.73E-07 5.2692184 ENST00000215794 18 USP41 ubiquitin specific peptidase A_32_P54553 4.42E-07 5.602893 ENST00000454608 41 XLOC_000755 Uncharacterised A_21_P0001111 2.48E-04 3.4425192 ENST00000440492
XLOC_001609 Uncharacterised A_21_P0001893 2.51E-04 3.4356153 ENST00000452037
XLOC_001966 Uncharacterised A_21_P0002006 0.001414 3.5555193 ENST00000414795
XLOC_l2_000 Uncharacterised A_21_P0010531 7.63E-04 2.5990458 ENST00000438195
233
ZC3HAV1 zinc finger CCCH-type, A_23_P20122 0.001103 2.571025 ENST00000471652 antiviral 1 ZNFX1 zinc finger, NFX1-type A_24_P23034 0.001044 2.2538695 ENST00000537431 containing 1 X Unknown A_33_P3303742 0.001369 -2.6417236 ENST00000439905
BAIAP2 BAI1-associated protein 2 A_23_P315836 8.57E-04 -2.668449 ENST00000435091
C15orf52 chromosome 15 open A_23_P163467 0.001095 -2.8513255 ENST00000397535 reading frame 52 EGR3 early growth response 3 A_23_P216225 0.001465 -2.7386675 ENST00000317216
- 227 - FLJ40288 uncharacterized FLJ40288 A_21_P0014767 0.001468 -2.2612228 ENSG00000183470
FRY furry homolog (Drosophila) A_33_P3223488 2.19E-04 -2.8213084 ENST00000436046
GAS7 growth arrest-specific 7 A_24_P82466 0.001099 -3.479138 ENST00000323816
HSD17B1 hydroxysteroid (17-beta) A_23_P15542 9.83E-04 -2.394454 ENST00000225929 dehydrogenase 1 LOC1005057 uncharacterized A_21_P0000759 5.88E-04 -3.36895 LOC100505746 46
LOC388242 coiled-coil domain A_32_P135336 2.26E-05 -3.503117 ENST00000549950 containing 101 pseudogene MUC5B mucin 5B, oligomeric A_24_P102650 0.00107 -2.7954655 ENST00000349637 mucus/gel-forming MYLK4 myosin light chain kinase A_33_P3252939 0.001104 -2.545482 ENST00000274643 family, member 4 NAV2 neuron navigator 2 A_23_P52727 0.001392 -2.4784532 ENST00000396085
NRARP NOTCH-regulated ankyrin A_33_P3337272 0.001073 -2.4398358 ENST00000356628 repeat protein PLIN5 perilipin 5 A_23_P39251 9.74E-04 -2.6129234 ENST00000381848
PTGFRN prostaglandin F2 receptor A_33_P3307197 8.48E-04 -2.9451718 ENST00000544471 negative regulator SIGLECP3 sialic acid binding Ig-like A_32_P128258 6.71E-04 -3.44869 ENSG00000171101 lectin, pseudogene 3 UBXN11 UBX domain protein 11 A_24_P239811 7.93E-04 -3.1068769 ENST00000486658
VCAN versican A_23_P144959 2.76E-04 -5.141041 ENST00000512590
XLOC_001338 Uncharacterised A_21_P0002164 0.001206 -3.0045128
XLOC_011215 Uncharacterised A_21_P0008616 0.001288 -2.5846038
XLOC_011218 Uncharacterised A_21_P0008619 0.001492 -2.333941
XLOC_012522 Uncharacterised A_21_P0009199 0.001353 -2.5692058 ENST00000514726
XLOC_l2_012 Uncharacterised A_21_P0012998 0.001168 -2.9539542 ENST00000505712
135
XLOC_l2_015 Uncharacterised A_21_P0013601 5.38E-04 -2.766016
033
- 228 - Figure 6.4
Hierarchical clustering of the genes. Significant changes in gene expression between activated and inactivated CD56+ T cells were illustrated by using heat maps. A
Hierarchical clustering approach joins genes by dendrograms that have similar gene expression patterns between them.
- 229 - A comparison between CD56+ T-cells under stimulated and un-stimulated conditions revealed 131 probes that were significantly changed, hower, out of these probes only 106 genes were significantly changed. Hierarchical clustering of these genes was performed (Figure 6.4).
Overall, genes are arranged according to FC levels in Table 6.2. Two genes have
FC levels more than 6 and they are IFN-γ and HES4, and only VCAN was down- regulated to less than -5 folds between CMV-stimulated and un-stimulated CD56+
T-cells.
- 230 - FC Gene Symbol
6 ≤ HES4, IFNG
5 to 5.99 IRG1,USP41,CD38, IL15RA, USP18, C1orf173, ISG15
4 to 4.99 IFIT3, IGFBP7, IFIT2, IFIT1, TNFSF10, OAS3, HERC5, IFIT2, RRAGC
IFI6
TNFSF10, OAS2, PML, NT5C3, IFI44L, MT1B, GBP3, GMPR, IFIH1, ETV7,
CPSF7, GBP1P1, OAS3, DDX58, CMPK2, XLOC_001966, LAG3
3 to 3.99 CACNA1A, HERC6, NT5C3, OASL, ISG20, XLOC_000755, XLOC_001609,
LAP3, PGAP1, SAMD9L, GALM, PLSCR1,PNPT1, GCH1 FRMD3, MX1,
EIF2AK2, IFITM1, IFITM4P, CREM, IFI44, NEXN, GCH1 ECE1, CYP2J2,
RNF43, RTP4, SOCS1, LOC79015
SLC27A2, OAS2, OAS1, RGS16, PRIC285, MX2, IL2RA, STAG3, DHX58
SLC38A5, MOV10, GBP4, EPSTI1, IFI35, HRASLS2, DDX60L, KLHDC7B,
2 to 2.99 DDX60, BST2, PARP9, PI4K2B, IFITM2, CNP, XLOC_l2_000233, CIB2,
TRIM5, KLHDC7B, ZC3HAV1, SP140, TMEM140, MT1L, PARP12, IFIT5,
FTSJD2, PARP10, ZNFX1،APOL6
FLJ40288, XLOC_011218, HSD17B1, NRARP, NAV2, MYLK4,
-2 to -2.99 XLOC_012522, XLOC_011215, PLIN5, BAIAP2, EGR3, XLOC_l2_015033,
MUC5B, FRY, C15orf52, PTGFRN, XLOC_l2_012135
-3 to -3.99 XLOC_001338, UBXN11, LOC100505746, SIGLECP3, GAS7, LOC388242
-5 ≥ VCAN
Table 6.2 Significantly up- or down-regulated genes expressed by CD56+ T- cells in response to CMV categorised according to fold change (FC).
- 231 - Up-regulated genes grouped according to biological processes expressed by
CD56+ T-cells in response to CMV lysate are illustrated in bar charts with the FC levels of the genes shown. In Figure 6.5 the genes that are normally regulated by
IFN are illustrated.
SOCS1 PML OAS3 OAS2 Response to OAS1 IFN-ϒ IFNG GCH1 OASL PLSCR1 Response to PNPT1 IFN-β OAS1 Response to IFIT3 IFN-α IFIT2
0 2 4 6 8 Folds Change
Figure 6.5 Genes normally regulated by IFNs and up-regulated in CD56+ T cells following CMV stimulation. The bar chart shows more genes were regulated by IFN-γ than both α and β.
Genes expressed by CD56+ T-cells were sorted according to the effect of CMV lysate; these are shown in Figure 6.6. The biological processes involved are negative regulation of viral processing and genome replication, detection of the virus and response to the virus.
- 232 - PLSCR1 Negative regulation of OAS3 viral process and OAS1 genome replication IFIT1
IFIH1 Detection of virus DDX58
TRIM5 PML PLSCR1 OAS3 OAS1 MX1 ISG20 ISG15 IFNG Response to virus IFIT2 IFIT1 IFIH1 IFI44L IFI44 HERC5 EIF2A… DDX60 DDX58
0 2 4 6 8
Figure 6.6 Genes upregulated according to function by CD56+ T-cells due to an anti-CMV response. Three functions were included response to virus, detection and virus and negative regulaytion of viral process and genome replication.
- 233 - Genes upregulated by CD56+ T-cells due to immune response and defence response are shown in Figure 6.7.
Immune response Defence response USP18 SP140 TNFSF10 PLSCR1 EIF2A… SOCS1 USP18 PML SOCS1 OASL PML OAS3 OASL OAS2 OAS3 OAS1 OAS2 MX1 OAS1 ISG20 MX1 ISG15 ISG20 IL2RA ISG15 IFNG IL2RA IFNG IFIT3 IFIT3 IFIT2 IFIT2 IFIT1 IFIT1 IFIH1 IFIH1 IFI6 IFI6 HERC5 HERC5 GCH1 GCH1 DHX58 DHX58 DDX58 DDX58
2 2.5 3 3.5 4 4.5 5 5.5 6 6.5 7 2 2.5 3 3.5 4 4.5 5 5.5 6 6.5 7
Figure 6.7 Genes upregulated according to immune and defence response by
CD56+ T-cells due to an anti-CMV response. These two charts show genes upregulated in CD56+ T-cells in response to CMV lysate; TNFSF10 is involved in immune response, while SP140, PLSCR1 and EIF2AK2 in defence response.
According to the Gene Ontology Normal Usage Tracking System (GONUTS; http://gowiki.tamu.edu/wiki/index.php/Main_Page) the defence response is
- 234 - reactions triggered due to the incidence of a foreign body or due to an injury, while immune response is any immune system process in response to internal or external pathogen. Therefore, defence response is a wider definition.
Genes whose expression in CD56+ T-cells changed due to stress are shown in
Figure 6.8. Stress is due to the CMV lysate and other cell signalling processes leading to disturbance of metabolic or signalling processes and variation in temperature and pH; controlling stress is essential for cell homeostasis (Kultz
2005).
- 235 - VCAN Response to stress USP18 SP140 SOCS1 PNPT1 PML PLSCR1 OASL OAS3 OAS2 OAS1 MX1 ISG20 ISG15 IL2RA IFNG IFIT3 IFIT2 IFIT1 IFIH1 IFI6 HERC5 GMPR GCH1 EIF2AK2 ECE1 DHX58 DDX58 CD38 -6 -4 -2 0 2 4 6 8 Folds Change
Figure 6.8 Genes upregulated according to stress by CD56+ T-cells due to an anti-CMV response. Changes in expression of stress regulated genes in CD56+
T-cells. VCAN was the only gene that was down-regulated due to stress inCD56+
T-cells.
- 236 - 6.3.2 Validation of microarrays results
Results of microarray gene expression were validated by two methods, RT-PCR and protein expression by using flow cytometry. Three transcripts were quantified by RT-PCR in Dr. BF Flanagan’s laboratory (Women’s and Children’s Health,
ITM, Alder Hey Hospital) and they were IFN-γ, ISG-15 and USP-18. Two were analysed by detecting their protein expression by FACs analysis and they are ISG-
15 and CD38.
6.3.3 RT-PCR gene expression analysis
The experiment was described in section 2.3.2.6.8. The detected expression of these genes will be calculated and compared to microarrays. Quantitative RT-PCR is an accurate method to utilise and quantify the expression levels of genes by using Taq-man.
The protein ISGylation and de-ISGylation cycle involves ISG-15, USP18, E1 activating enzyme, E2 conjugating enzyme and E3 ligase (TRIM5, HERC5 and
HERC4). ISGylation is briefly explained as the covalent attachment of ISG-15 to proteins which is induced by IFN, while removal of ISG-15 is called de-
ISGylation via USP-18 (Schneider et al. 2014); this is similar to the ubiquitination pathway (Figure 6.9).
- 237 -
Figure 6.9 Ubiquitination cycle. Labelling proteins with ubiquitin or ubiquitin-like
modifier render them to be targeted for degradation within the 26s proteasome. The reaction is catalysed by E1, E2 and E3 (adapted from www.biochemie.charite.de)
(Takeuchi and Yokosawa 2008).
As monoclonal antibodies are available for IFN-γ, ISG-15 and USP-18, these were picked for quantification by RT-PCR and flowcytometry to validate microarrays results. In figure 6.10an illustration of two samples amplification blot is attached.
- 238 -
Figure 6.10 Amplification plot showing curves by RT-PCR. Samples were run in duplicates; the amplification blot shows curves of same products amplified at the same time. Delta (Δ) Rn used to show the magnitude of the signal produced when samples amplified. The green threshold line identifies the Ct value during the PCR exponential phase.
The results of RT-PCR of IFN-γ gene expression analysis are illustrated in Table
6.3, and for ISG-15 gene expression analysis are illustrated in Table 6.4, and for
USP-18 gene expression analysis are illustrated in Table 6.5.
- 239 - L32 - L32 Av IFN-γ Av Ct 2^Ct IFN- γ
Sample
S-1 26.3208 26.3934 26.3571 25.6838 25.8338 25.7588 0.5983
US- 7.532 185. 25.8969 25.4654 25.6812 32.7122 32.5192 32.6157 -6.9345 1
S-2 27.6131 27.2341 27.4236 26.6391 28.1414 27.3903 0.03335
US- 4.310 19.8 30.8165 30.6321 30.7243 34.925 35.0782 35.0016 -4.2773 2
S-3 29.3997 29.2063 29.303 27.9552 28.2704 28.1128 1.1902
US- 6.087 68.0 36.7006 36.8294 36.765 41.5109 41.8144 41.6627 -4.8976 3
S-4 33.9122 33.8688 33.8905 30.7599 30.9006 30.8303 3.06025
US- 7.177 144. 36.786 38.1902 37.4881 42.7882 40.423 41.6056 -4.1175 4
Table 6.3 RT-PCR results for IFN-γ. Av= average, L32 housekeeping gene, Ct= threshold cycle and the difference between stimulated and unstimulated was calculated by this equation (Ct s- Ct us), 2^Ct = 2 delta Ct to calculate relative gene expression and is expressed as fold change between non stimulated and stimulated cells.
- 240 - Table 6.4 RT-PCR results for ISG-15. Av= average, L32 housekeeping gene,
Ct= threshold cycle and the difference between stimulated and unstimulated was calculated by this equation (Ct s- Ct us), 2^Ct = 2 delta Ct to calculate relative gene expression. Expressed as fold change between non stimulated and stimulated cells. .
L32 - L32 Av ISG-15 Av Ct 2^Ct ISG-15
Sample
S-1 26.3208 26.3934 26.3571 21.0772 20.9384 21.0078 5.3493
US- 4.234 18.8
25.8969 25.4654 25.6812 24.4557 24.6773 24.5665 1.11465 1
S-2 27.6131 27.2341 27.4236 22.2968 21.5766 21.9367 5.4869
US- 5.931 61.3 30.8165 30.6321 30.7243 31.3468 31.0041 31.1755 -0.4512 2
S-3 29.3997 29.2063 29.303 26.5472 26.3905 26.4689 2.83415
US- 5.137 35.2
36.7006 36.8294 36.765 38.7396 39.3974 39.0685 -2.3035 3
S-4 33.9122 33.8688 33.8905 30.4878 30.3916 30.4397 3.4508
US- 6.118 69.4 36.786 38.1902 37.4881 39.6427 40.6693 40.156 -2.6679 4
- 241 - Table 6.5 RT-PCR results of USP-18. Av= average, L32 housekeeping gene,
Ct= threshold cycle and the difference between stimulated and unstimulated was calculated by this equation (Ct s- Ct us), 2^Ct = 2 delta Ct to calculate relative gene expression expressed as fold change between non stimulated and stimulated cells.
.
L32 - L32 Av USP-18 Av Ct 2^Ct USP-18
Sample
S-1 26.3208 26.3934 26.3571 24.8248 24.817 24.8209 1.5362 3.8107 14.04
US-1 25.8969 25.4654 25.6812 27.9286 27.9826 27.9556 -2.2745
S-2 27.6131 27.2341 27.4236 24.1246 24.1243 24.1245 3.29915 5.7899 55.33
US-2 30.8165 30.6321 30.7243 33.2039 33.2261 33.215 -2.4907
S-3 29.3997 29.2063 29.303 26.7736 26.9293 26.8515 2.45155 4.4915 22.49
US-3 36.7006 36.8294 36.765 39.1202 38.4896 38.8049 -2.0399
S-4 33.9122 33.8688 33.8905 32.5595 33.0314 32.7955 1.09505
US-4 4.5467 23.38 36.786 38.1902 37.4881 41.7594 40.1202 40.9398 -3.4517
- 242 -
The means of the gene expression values were compared to microarrays levels and illustrated in Figure 6.11.
Figure 6.11 Comparison between means of gene expression in RT-PCR and
Microarrays. Only USP-18 level of expression in microarrys were higher than
RT-PCT.
110 105 100 95 90 85
80
75 70 65 60 55 RT-PCR 50 45 40 Microarrays 35 30 Gene expression Gene 25 20 15 10 5 0 IFN-γ ISG-15 USP-18
6.3.4 ISG-15 and CD38 protein expression analysis by FACs
Monoclonal antibodies were only available for a few of the gene products found to be upregulated by microarray analysis. ISG-15 and CD38 protein expression were validated by flow cytometry. The samples from CMV negative and positive subjects were compared to analyse any possible significant differences.
- 243 - 6.3.4.1 ISG-15 protein expression by CD56+ T-cells
The method for detecting ISG-15 protein is described in section 2.3.2.6.9. A total of 10 CMV positive and negative subjects were analysed. ISG-15 expression levels were compared between three different cells groups: NK cells, CD56+ T- cells and CD56- T-cells after incubating PBMCs overnight with CMV lysate.
Results are illustrated in figure 6.12. The levels were calculated by density plot and presented in Figure 6.13.
0.0248 0.0152 0.0648 100
80
60 %
40
20
0 - + - + - + NK CD56+ T-cells CD56- T-cells
Figure 6.12 Levels of ISG-15 protein expression. ISG-15 in response to CMV extract in NK cells, CD56+ T-cells and CD56- T-cells were compared between
CMV positive and negative healthy donors.
- 244 -
Figure 6.13 Levels of ISG-15 protein expression by density blot. Density plots illustrating the levels of ISG-15 in different cell populations before and after overnight stimulation with CMV lysate. Results from a representative CMV+ subject are shown.
- 245 - Proportions of NK cells expressing ISG-15 following stimulation with CMV extract were significantly more in CMV positive (46.69% ± 9.179) than negative subjects (16.68% ± 3.371) (P=0.0248; Fig. 6.8), moreover proportions of CD56+
T-cells expressing ISG-15 were significantly more in CMV positive (47.550% ±
9.476) than negative subjects (14.64% ± 4.304) (P=0.0152). However, proportions of CD56- T-cells expressing ISG-15 were not significantly more in CMV positive
(36.67% ± 10.88) than negative subjects (11.74% ± 2.331) (P=0.0648).
6.3.4.2 CD38 protein expression by CD56+ T-cells
CD38 is an ectoenzyme and a marker of cell activation found on the surface of several immune cell types and functions in cell adhesion, signal transduction and calcium signalling. Ectoenzyme is situated on the outer surface of cells membrane and released upon stimulation. CD38 gene expression was found to increase significantly in CMV lysate stimulated CD56+ T-cells. Expression of CD38 protein was evaluated in NK cells, CD56+ T-cells and CD56- T-cells and compared between CMV sero-groups (figure 6.14).
- 246 -
Figure 6.14 CD38 expressions. CD38 expression by cells from CMV+ and
CMV- donors; no significant differences were found. The right scatter plot shows
NK cells to have the highest MFI for CD38 expression among the cell groups but no significant differences were seen between CMV groups.
6.4 Discussion
This part of the study aimed to identify changes in the gene expression of CD56+
T-cells in response to overnight CMV stimulation. The first step was to obtain purified CD56+ T cells which was performed using fluorescence activated cell sorting of double labelled cells. Although purities of sorted cell populations were
>90% but there was still some slight contamination, most likely with CD56- T cells. The possibility remains that these contaminating cells contributed in such a way that they significantly altered the pattern of changes in gene expression by
CD56+ T cells.
The gene expression experiments were done by using Agilent Human v2 8 x 60K arrays. The results were validated through RT-PCR and flow cytometric analysis
- 247 - of protein expression. Overall 106 genes were significantly changed following
CMV antigen stimulation of CD56+ T-cells. IFN-γ gene expression significantly increased and it is clear that most of the genes significantly changed were interferon stimulated genes; therefore, before proceeding to discuss their role it is essential to consider how the IFN antiviral response works. While the Type I interferons IFN-α and IFN-β are part of the innate immune response, IFN-γ is the sole representative Type II interferon (Schneider et al. 2014). In brief, the Type I
IFN mediated response is part of the innate immune response and participates in activation of the adaptive immune response. When a cell is invaded or detects the presence of a pathogen such as a virus, bacterium or parasite, there is consequent production of IFN that binds cell surface receptors and triggers a series of signal transducers also called Janus kinase signal transducer and activator of transcription pathway (JAK-STAT). This leads to transcriptional regulation of
IFN-regulated genes and preparation of the cells into an anti-pathogenic status
(Schneider et al. 2014), inducing the transcription of 500 to 1000 IFN-stimulated genes (ISGs) that encode proteins with several functions including resisting infection and shaping the immune response (Diamond and Farzan 2013).
Family groups of significantly changed ISGs are IFN-induced proteins with tetratricopeptide (IFIT) repeat domains which include IFIT1 (ISG-56), IFIT2
(ISG-54), IFIT3 (ISG-60) and IFIT5 (ISG-58) and also IFN-induced transmembrane proteins (IFITM) which include IFITM1, IFITM2 and IFITM4P
(Pseudogene) and Interferon induced with helicase C domain 1 (IFIH1). The latter exists in the cytoplasm and senses viral nucleic acid, induces the production of
IFN-α and β and is effective in triggering the innate immune response and NK cell activation (Cocude et al. 2003, Bamming and Horvath 2009) which in consistent
- 248 - with this study where it was found to increase in response to CMV lysate (Figures
6.6-6.8). This study showed IFIT1 and 2 to be upregulated in response to CMV which participate in negative regulation of viral processes and genome replication
(Figure 6.6). IFIT 1, 2 and 3 also participate in response to stress (Figure 6.8).
While IFIT5 is an essential enhancer for the innate immune system response against viral infection (Zhang et al. 2013), the IFIT family encodes proteins that are contained in the cytoplasm with no known enzymatic functions and are also named viral stress-inducible genes. Additionally, they contain tetratricopeptide repeats (TPR) that can participate in functions such as cell cycle progression, transcription, protein folding and transport (D'Andrea and Regan 2003, Diamond and Farzan 2013).
This family under normal condition are not expressed, however, during virus infection they are expressed rapidly (Sarkar and Sen 2004) and after ~2 hours of
IFN-α induction (Diamond and Farzan 2013) and were upregulated here (Figure
6.5). Their antiviral functions include suppression of translation initiation, binding of viral RNA and sequestering viral RNA and proteins in the cytoplasm; furthermore, studies suggest they can play roles in immune responses (Diamond and Farzan 2013). The IFITM family are similar to IFIT in lacking enzymatic functions, and they have effective antiviral functions against enveloped viruses.
IFITM proteins are found abundantly in late endosomes and lysosomes of the cell, and they function to block virus migration into the cells which make them effective against viruses which require entry into late endosomes and lysosome for their effective invasion, such as SARS-coronavirus and filovirus. However, they have no effect on HIV against which IFTIM2 is effective, this indicate selectivity for the virus they inhibit (Schneider et al. 2014). IFITM4P has been
- 249 - identified as a pseudo-gene (Diamond and Farzan 2013), which is known to be an outcome of accumulation of multiple mutations and their product is not required to perform biological functions.
2', 5'-oligoadenylate synthetase OAS genes 1, 2, 3 and L are induced by IFN-γ and are involved in generating immune and defence responses while OAS1 is induced by α as well (Figures 6.5 and 6.7). Also, OAS 1 and 3 are involved in negative regulation of viral processing and genome replication, and were found to be upregulated in response to CMV lysate (Figure 6.6). The OAS/RNase L system inhibits viral infection and is part of the innate immune pathway to degrade viral and cellular RNA (Silverman 2007). Mx1 is involved in response to viruses, immune and defence responses and response to stress (Figures 6.6, 6.7 and 6.8).
Myxovirus resistance (Mx) genes 1 and 2 are involved in inhibiting virus entry into cells, and are IFN-induced. Moreover, Mx1 acts at an early step of the virus cycle where it prevents viruses from reaching their cellular destination (Levy et al.
1986). Mx2 traps reverse-transcribed genomes from attaining a nuclear destination and is found to be effective against HIV and influenza-A (Silverman
2007). SOC1 is also induced by IFN-γ (Figure 6.5) and is involved in immune and defence responses and stress (Figures 6.7 and 6.8). It encodes SOCS protein that increases on early response to IFN induction and regulates IFN through JAK-
STAT (Schneider et al. 2014). LAG-3 is expressed by activated NK and T-cells and blocking LAG-3 and PD-1 at the same time induces T cell activation which reveals the inhibitory effects of these gene products on T-cells during chronic viral infection (Triebel et al. 1990, Blackburn et al. 2009).
Putative RNA helicases DEAD-box DDX58, DDX60, DDX60L and DEAH-box
DHX58 were up-regulated significantly in the response to CMV (Figure 6.6).
- 250 - DDX58 and DHX58 both involved in immune and defence responses and response to stress (Figures 6.7 and 6.8). DEAD-box and DEAH-box play roles in binding nucleic acids and unwinding RNA structure, also, in cell differentiation and are distinguished by their amino acids of the helicase 4-mer motif-II
(Abdelhaleem et al. 2003). PNPT1 and PLSCR1 are both induced by IFN-β and in response to stress (Figures 6.5 and 6.8), PLSCR1 is involved also in negative regulation of viral processing, genome replication and response (Figures 6.6 and
6.7).The IFN-induced Phospholipid scramblase 1 (PLSCR1) is involved in antiviral responses and cells with low levels of PLSCR1 are weakly responsive to antiviral-IFN responses (Dong et al. 2004). PNPT1 is an IFN-induced gene that encodes a protein involved in RNA processing and degradation (G. Wang et al.
2012). TNFSF10 encodes a protein of the TNF ligand family which induces apoptosis in transformed cells; this gene is involved in the immune response
(Figure 6.7).
VCAN was the only gene whose expression was found to be significantly decreased in response to CMV antigens and has been reported to be reduced due to stress (Figure 6.8). This gene encodes the extracellular matrix glycoprotein versican which is involved in cell adhesion, proliferation, migration and angiogenesis and plays a central role in tissue morphogenesis and maintenance.
Versican can bind L and P-selectin and CD44 (Wight 2002) and studies indicated
T-cells struggle to migrate in versican enriched media, while blocking of versican in this media induced cell migration (Evanko et al. 2012, Wight et al. 2014).
ISG-15, TRIM5 and HERC5 were all upregulated in response to CMV (Figure
6.6); these genes are involved in immune and defence responses and stress response (Figure 6.7 and 6.8).
- 251 - Figure 6.11 shows that expression levels of ISG-15, USP-18 and IFN-γ between
RT-PCR and microarrays correlated quite closely.I It is expected that RT-PCR would be more sensitive than microarrays and this was found for IFN-γ and ISG-
15, however, USP-18 microarrays expression levels was 10 folds higher than RT-
PCR. Moreover, genes involved in the innate immune response to viral infection
Type-I IFN responses; OAS, MX-1, MDA5 and TRIM5 were also increased in
CMV-specific-CD4+ T-cells (Hu, 2013).
In addition, ISG-15 and CD38 were further studied to validate microarrays and
RT-PCR results and to compare protein expression between CMV positive and negative healthy donors in response to CMV antigens. Figure 6.13 shows comparison between levels of ISG-15 after overnight stimulation. Clearly, CMV positive donors showed higher ISG-15 levels than CMV negative healthy donors and this was statistically significantly in NK cells and CD56+ T-cells although not quite for CD56- T cells.
However, CD38 was highly expressed among all cell groups with no significant differences between CMV- and CMV+ donors (figure 6.14). Mean fluorescence intensities (MFI) were increased in NK cells and CD56+ T-cells from CMV+ compared to CMV- donors, although this did not reach statistical significance.
CD38 functions as a signalling molecule and is expressed widely on several cell types; it is type II transmembrane glycoprotein and detected on activated T and B- cells. It has been argued in HIV patients that high levels of CD38 on CD8+ T- cells can be a negative prognostic marker (Liu et al. 1996). In EBV-induced infectious mononucleosis numbers of CD38 antigenic sites were higher than in
CMV-induced infectious mononucleosis, however, both were higher than in healthy control groups (Zidovec Lepej et al. 2003).
- 252 - To conclude this part of the study, CD56+ T-cell gene expression studies indicated high expression of genes involved in immune responses toward CMV.
Some genes were identified in the literature to be involved in responses against other viruses but not CMV such as DDX60, ISG15,ISG, MX1, MX2, OASL and
TRIM5 etc (Schoggins and Rice 2011). This is consistent with cellular studies in previous chapters indicating that CD56+ T-cells respond to CMV lysate and share some properties with NK cells and T-cells.
- 253 -
Chapter 7
General conclusion
- 254 - 7. General conclusions and future work
CMV is a common asymptomatic persistent infection and prevalence ranges between 30-90% in developed countries (Crough and Khanna 2009). HCMV infection is a potential threat leading to failure in graft adaptation in recipients sometimes leading to graft rejection. Moreover, CMV induces the immune system via triggering inflammation, immune cell infiltration, expression of adhesion molecules in the transplanted organ, the release of cytokines and chemokines, and growth factors connected to the process of acute rejection of the graft (Koskinen et al. 1999). Kidney rejection may be promoted via re-activation of CMV in the blood followed by fever, muscle pain, leucopenia, hepatitis, gastroenteritis, pneumonitis and thrombocytopenia (Hartmann et al. 2006, Weikert and Blumberg
2008).
CD56+ T-cells were first described by Lanier et al (Lanier et al. 1986) but shown by Schmidt-Wolf et al (1991) to be generated in vitro after the addition of anti-CD3, IFN-γ and IL-2 to PBMCs and incubation for 28 days. These cells were shown both to exert TCR specificity and to have NK cell function (Pievani et al.
2011). This study aimed to investigate any relationship between CD56+ T-cells and CMV both in healthy subjects and kidney transplant patients. CD56+ T-cells have the potential to promote cytopathic effects against transplanted kidneys upon
CMV activation.
CD56+ T-cell prevalence and function between CMV seropositive and seronegative healthy and KTPs were evaluated. Both healthy subjects and KTPs have significantly higher levels of CD56+ T-cells in seropositive than seronegative groups. In Figure 5.11 both healthy subjects and KTPs were
- 255 - compared to identify any significant differences between the groups studied.
Proportions of CD56+ T-cells were significantly higher in healthy CMV+ subjects than both CMV+ KTP groups, D+R- and R+. Additionally, healthy CMV- donors had significantly higher proportions of CD56+ T-cells than D-R- KTPs. This indicates that CD56+ T-cells expanded in response to the presence of CMV in
CMV + healthy and patient groups, and the significant lower levels in D+R- and
R+ KTPs than the healthy seropositive donors may be due to immune-suppression therapy consisting of Cyclosporine, Campath-1 or Tacrolimus; this was also found in CMV- healthy subjects and KTPs where the healthy subjects had significantly higher levels than D-R- patients. Still, with this immune-suppressive therapy the
CMV+ KTPs showed significantly higher levels of CD56+ T-cells than the D-R- patient group. If infection with viruses other than CMV induces expansion of
CD56+ T cells, in the case of D-/R- KTPs this may have been prevented by immunosuppressive drug treatment, giving rise to lower levels than in healthy
CMV- subjects. Other viruses such as HHV-6 and BK virus can also be re- activated in KTPs. Future work could examine CD56+ T cell levels and functional activity in KTPs in which donor and recipient were both negative for CMV but mismatched for these other viruses. This would show whether CMV is unique in inducing expansion of CD56+ T cells.
Both patients and healthy positive donors had increased CD4 and CD8 surface expression on CD56+ T-cells than in CMV- groups and CD8 was expressed on significantly more cells than CD4 in both seropositive healthy and KTP groups.
Furthermore, characterization of CD56+ T-cells in both CMV+ healthy subjects and KTPs revealed that these cells become memory cells with T-EMRA like phenotype and pre-store granzyme B as this study detected. The only difference
- 256 - detected was that in healthy donors MFI for CD11a was significantly higher in
CMV+ than CMV- subjects while in KTPs this was not statistically significant.
This also can be a result of administration of immune-suppressive therapy.
Overall, the results indicated that CD56+ T-cell prevalence and memory phenotype are almost the same between healthy subjects and KTPs depending on the sero-status of the individual.
The functional capacity of CD56+ T-cells was evaluated in Chapter 4 of this study. CD56+ T-cell populations have dual functions where some preserved TCR specificity against CMV and expanded in response to CMV stimulation while also performing NK-like cytotoxicity against MHC negative K562 cells. CD56+ T- cells from CMV+ donors reacted to CMV lysate by producing Th1 like cytokines
IFN-γ and TNF-α and Granzyme B in significantly greater proportions than in
CMV – donors (Table 4.1). Th1 cells promote an alloresponse to a kidney graft and graft rejection via the activation and recruitments of CD8+ T-cells, B-cells release IgG2a that activates complement and CD8+ T-cells also promote macrophage dependent delayed-type hypersensitivity. Additionally, CD8+ T cells also may express Fas-L that promotes CD8+ T-cell cytotoxicity (Issa et al. 2010).
Moreover, the CD107a degranulation assay showed that CD56+ T-cells from
CMV+ subjects express significantly more CD107a than CMV- donors after being pulsed by K562 and CMV lysate. KLRG-1 was expressed by significantly more
CD56+ T-cells in CMV+ than CMV- subjects, this marker was argued to increase on cells lacking proliferation capability (Voehringer et al. 2002).
However, this study showed that CD56+ T-cells obtained from CMV+ donors were able to proliferate in response to CMV stimulation which is consistent with another study (Geginat et al. 2003). TCR specificity was preserved by CD56+ T-
- 257 - cells obtained from both healthy subjects and KTPs, CD8bright+ CD56+ T-cells reacted specifically to pentamers loaded with common CMV peptides HLA-
A*02:01-NLV, HLA-B*07:02-TPR and HLA-B*08:01-QIK. CD8bright+CD56+ T- cell reactivity to these pentamers were compared to CD56-CD8 bright+ T-cells.
Furthermore, CD8bright+CD56+ T-cells from one HLA-B8+ healthy donor showed a higher proportion of HLA-B*08:01-QIK binding cells than CD8bright+CD56- T- cells. Comprehensive cellular work conducted in Chapters 3-5 and showed increased proportions and function of CD56+ T cells in CMV+ healthy subjects and KTPs, also CMV-specific cytotoxic T cells were found, which might indicate involvement toward the immune rejection of the kidney graft via CMV antigens on the graft especially in high risk groups D+R-. In future work, pentamer+ cells could be purified and tested for cytotoxic activity against both CMV-infected target cells and NK cell targets. This would indicate whether CD56+ T cells can retain both antigen-specific and non-MHC restricted cytotoxic function. This might be relevant to transplant rejection in D+/R- patients.
Gene expression profile of CD56+ T-cells purified from CMV+ PBMCs was evaluated as illustrated in Chapter 6. This study detected 106 genes that significantly changed following CMV antigen stimulation. The technique applied was Microarrays and in three cases was validated via RT-PCR and Flow- cytometry. CMV can trigger activation of signalling pathways that prime mRNA expression which in turn can be detected by Microarrays to provide quantitative study of the results of CMV lysate activation on CD56+ T-cells. The functions of these genes are involved in innate and adaptive immune responses. Interferon stimulated genes were significantly upregulated in CD56+ T-cells upon overnight stimulation with CMV lysate. Significantly expressed genes were sorted
- 258 - according to their biological process in Figures 6.5-6.8. Validation of microarrays has been processed by RT-PCR on IFN-γ, ISG-15 and USP-18.
However, a drawback of these studies is that there was some contamination with
CD56- T cells. This was not the case in flow cytometric studies as single cell expression could be investigated. CD38 and ISG-15 proteins expression and levels between CMV- and CMV+ healthy donors were studied via flow cytometry. Overall, ISG-15 was discovered in 1992 (Loeb and Haas 1992) and it is one of the early genes that response to viral stimulation and interferon induction, moreover, more than158 conjugation targets for ISG-15 have been detected; these targets are involved in several processes such as RNA splicing, stress response, transcription and translation (Zhao, 2005, Giannakopoulos, 2005).
ISG-15 was argued as essential in IFN-α/β induced antiviral response via maintaining antiviral protein stability, activity, localisation and association with other proteins during the antiviral response (Lu, 2006). ISG-15 knockout mice show increased susceptibility to viral infection (Lenschow, 2005, Lenschow,
2007), while CD56+ T-cells expressed more ISG-15 in CMV + than CMV – subjects, indicating potential immunological functions against CMV.In future work, CD56+ T cell clones could be derived and stimulated with CMV antigens to more definitively show their range of gene expression. Expression of a wider range of immune-related genes could also be studied at the protein level.
To conclude, the findings of this comprehensive study on CD56+ T-cells in relation to CMV infection increase the overall knowledge about these cells, and add significant information on their possible role in response to CMV. They suggest that CD56+ T-cells participate at some level in the immune response
- 259 - toward kidney grafts especially during CMV syndrome as there is the potential for increased levels of non-MHC restricted cytotoxic function.
Future work on these cells might involve study of their prevalence and functions against other pathogens in kidney or other organ transplant patients. Their involvement in graft rejection might also be investigated. This could most readily be done by obtaining pre-transplant blood samples and sequential samples post- transplantation and assessing CD56+ T cell numbers and activation status in relation to episodes of CMV viraemia or graft rejection. Moreover, following up to the functional studies of these genes there is a possibility of knocking out these genes in mice and looking at responses in an animal model or by knocking down genes in cultured cells to see if removing them changes the infectivity of CMV or changes the response of CD56+ T cells to CMV lysate.
- 260 -
Chapter 8
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- 293 - Appendix
Table of genes information collected from genes bank mainly www.genecards.org, otherwise the source is mentioned.
Gene Symbol Function
APOL6 Helps lipids cytoplasm
IFN-induced gene, Acts as a direct physical tether to block the release of BST2 enveloped viruses
C1orf173 Chromosome 1 Open Reading Frame 173
Voltage-gated calcium channels which mediate calcium influx in response to CACNA1A depolarisation.
Multifunctional ectoenzyme and functions in cell adhesion, signal CD38 transduction and calcium signalling.
CIB2 calcium homeostasis and participate in calcium regulation
CMPK2 participate in dUTP and dCTP synthesis in mitochondria
CNP RNA metabolism in the myelinating cell
CPSF7 Cleavage Factor
CREM cAMP-mediated signal transduction
CYP2J2 encodes a member of the cytochrome P450 superfamily of enzymes
Involved in viral double-stranded (ds) RNA recognition and the regulation of
DDX58 immune response, which encode innate immune receptor that detect and
sense viral nucleic acids
Positively regulates DDX58/RIG-1 and IFIH1/MDA5- Type 1 inducible DDX60 gene in response to viral infection.
DDX60L DEAD (Asp-Glu-Ala-Asp) Box Polypeptide 60-Like,
Encode DHX58 which regulate of DDX58/RIG-I and IFIH1/MDA5 mediated DHX58 antiviral signalling
- 294 - The protein encoded by this gene is involved in proteolytic processing of ECE1 endothelin precursors to biologically active peptides.
The protein encoded by this gene is an IFN-induced dsRNA-dependent
serine/threonine-protein kinase which plays a key role in the innate immune EIF2AK2 response to viral infection and is also involved in the regulation of signal
transduction, apoptosis, cell proliferation and differentiation
EPSTI1 Epithelial Stromal Interaction 1
ETV7 Transcriptional regulators
FRMD3 FERM Domain Containing 3, a tumour suppressor gene
FTSJD2 Interferon-Stimulated Gene
GALM Encodes an enzyme that catalyzes the epimerization of hexose sugars
GBP1P1 Interferon-Inducible Pseudogene
encoded protein interacts with a member of the germinal center kinase family GBP3 and exhibits antiviral activity against influenza virus
GBP4 Interferon-induced Gene and hydrolyze GTP to both GDP and GMP
GCH1 Encodes a member of the GTP cyclohydrolase family
Encodes an enzyme that catalyzes the irreversible and NADPH-dependent GMPR reductive deamination of GMP to IMP
Member of the HERC family of ubiquitin ligases and encodes a protein with
a HECT domain and five RCC1 repeats. Pro-inflammatory cytokines
HERC5 upregulate expression of this gene in endothelial cells. The protein localizes
to the cytoplasm and perinuclear region and functions as an interferon-
induced E3 protein ligase that mediates ISGylation of protein targets
Member of the HERC family of ubiquitin ligases, E3 ubiquitin-protein ligase
HERC6 which accepts ubiquitin from an E2 ubiquitin-conjugating enzyme in the form
of a thioester and then directly transfers the ubiquitin to targeted substrates
HES4 Encode a protein that function as transcriptional repressor
- 295 - Encode a protein that catalyzing the calcium-independent hydrolysis of acyl
HRASLS2 groups in various phosphatidylcholines (Upragarin et al.) and
phosphatidylethanolamine (PE)
IFI35, IFI44L, IFI44 Interferon-induced
IFI6 interferon Alpha-Inducible and encode a protein involved in apoptosis
interferon Beta-Inducible and part of innate immune response, cell growth IFIH1 and division
Also called ISG56, and it’s encoded protein involved in vrial replication IFIT1 inhibition
interferon Alpha-beta Inducible, also called ISG54, and it encodes protein IFIT2 involved in inhibition of viral mRNA
Interferon inducible also called ISG60 and encodes protein inhibits viral IFIT3 processes
Also called ISG58 and encodes protein involved in recognition of viral IFIT5 ssRNA
A protein encoding genes of interferon induced transmembrane protein 1 and IFITM1 prevents virus migration into the cells
A protein encoding genes of interferon induced transmembrane protein 2 and IFITM2 prevents virus migration into the cells
IFITM4P interferon induced transmembrane protein 4 pseudogene
IFNG Encodes IFN-γ protein
IGFBP7 Encodes insulin-like growth factor binding protein
IL15RA Encodes IL-15 cytokine receptor
IL2RA Encodes IL-2 cytokine receptor
Encodes and protein involved in negative regulation of TLR mediated IRG1 inflammatory immune response
ISG15 Encodes ubiquitin-like protein due to activation by IFN-α and β
- 296 - Encodes ubiquitin-like protein that involved in antiviral function of ISG20 interferon against RNA viruses
KLHDC7B Portion coding gene
LAG3 Lymphocytes activation gene -3
Leucine aminopeptidase and encodes protein involved in intracellular protein LAP3 processing
LOC79015 Uncharacterised
MOV10 Encodes protein included in RNA-mediated gene slicing
MT1B Metallothionein 1B
MT1L Metallothionein 1L
Myxovirus (Influenza Virus) Resistance 1, Interferon-Inducible Protein MX1 encodes protein involved in antiviral response
Myxovirus (Influenza Virus) Resistance 2, Interferon-Inducible Protein MX2 encodes protein involved in antiviral response
Encodes a filamentous actin-binding protein that may function in cell NEXN adhesion and migration
NT5C3 encodes a member of the 5'-nucleotidase family of enzymes
2'-5'-Oligoadenylate Synthetase 1 and it is interferon induced encodes OAS1 protein essential in anti-viral innate immune response
2'-5'-Oligoadenylate Synthetase 2 and it is interferon induced encodes OAS2 protein essential in anti-viral innate immune response
2'-5'-Oligoadenylate Synthetase 3 and it is interferon induced encodes OAS3 protein essential in anti-viral innate immune response
2'-5'-Oligoadenylate Synthetase-like and it is interferon induced encodes OASL protein essential in anti-viral adaptive immune response
Poly (ADP-Ribose) Polymerase Family, Member 10, and works as a PARP10 transcriptional factor
- 297 - Poly (ADP-Ribose) Polymerase Family, Member 12 and catalyses the post- PARP12 translational modification of proteins
Poly (ADP-Ribose) Polymerase Family, Member 9 and catalyses the post- PARP9 translational modification of proteins
PGAP1 Post-GPI Attachment To Proteins 1
PI4K2B Phosphatidylinositol 4-Kinase Type 2 Beta
PLSCR1 Enhance interferon antiviral response
Promyelocytic Leukemia, and function as transcription factor and tumour PML suppressor
Polyribonucleotide Nucleotidyltransferase 1, and involved in RNA PNPT1 processing and degradation
PRIC285 transcriptional coactivator (phosphosite.org)
RGS16 Regulator Of G-Protein Signalling 16, and inhibits signal transduction
Ring Finger Protein 43, E3 ubiquitin-protein ligase, is a HAP95 (AKAP8L; RNF43 MIM 609475) binding ubiquitin ligase that promotes cell growth
RRAGC Ras-Related GTP Binding C
RTP4 Receptor (Chemosensory) Transporter Protein, interferon responsive 4
SAMD9L Sterile Alpha Motif Domain Containing 9-Like
SLC27A2 Solute Carrier Family 27 (Fatty Acid Transporter), Member 2
SLC38A5 Solute Carrier Family 38, Member 5
SOCS1 Suppressor Of Cytokine Signaling 1
SP140 SP140 Nuclear Body Protein
STAG3 Stromal Antigen 3
TMEM140 Transmembrane Protein 140
Tumor Necrosis Factor (Ligand) Superfamily, Member 10, encodes cytokine TNFSF10 of TNF ligand family which induce apoptosis in stressed cells
TRIM5 Tripartite Motif Containing 5, function as a E3 ubiquitin-ligase and
- 298 - ubiqutinates itself to regulate its subcellular localization
Ubiquitin Specific Peptidase 18, belongs to the ubiquitin-specific proteases
USP18 (UBP) family of enzymes that cleave ubiquitin from ubiquitinated protein
substrates
Ubiquitin Specific Peptidase 41, encodes a protein that recognize and USP41 hydrolyze the peptide bond at the C-terminal Gly of ubiquitin
XLOC_000755 Alternative name lnc-IFI6-1, (LNCipedia.org) XLOC_001609 Alternative name lnc-mrps9-3, (LNCipedia.org)
XLOC_001966 Alternative name lnc-CMPK2-1, (LNCipedia.org)
Zinc Finger CCCH-Type, Antiviral 1, encodes a CCCH-type zinc finger ZC3HAV1 protein that is thought to prevent infection by retroviruses
ZNFX1 Inc Finger, NFX1-Type Containing 1
BAI1-Associated Protein 2, encodes a brain-specific angiogenesis inhibitor BAIAP2 protein
C15orf52 Chromosome 15 Open Reading Frame 52
Early Growth Response 3, transcription factor involved in muscle spindle EGR3 development and lymphocytes development
FLJ40288 Uncharacterized FLJ40288
FRY Furry Homolog (Drosophila)
Growth arrest-specific 7, encodes a protein play a role in promoting GAS7 maturation and morphological differentiation of cerebellar neurons
HSD17B1 Hydroxysteroid (17-Beta) Dehydrogenase 1
LOC100505746 Uncharacterised
LOC388242 Coiled-Coil Domain Containing 101 Pseudogene
Mucin 5B, encodes a member of the mucin family of proteins, which are MUC5B highly glycosylated macromolecular components of mucus secretions
MYLK4 Myosin Light Chain Kinase Family, Member 4
- 299 - Encodes a member of the neuron navigator gene family, which may play a NAV2 role in cellular growth and migration
NRARP NOTCH-Regulated Ankyrin Repeat Protein
Members of the perilipin family, such as PLIN5, coat intracellular lipid PLIN5 storage droplets and protect them from lipolytic degradation
PTGFRN Prostaglandin F2 Receptor Inhibitor,
SIGLECP3 Sialic acid binding Ig-like lectin 17, pseudogene
UBXN11 UBX Domain Protein 11
Versican, the protein encoded is involved in cell
VCAN adhesion, proliferation, proliferation, migration and angiogenesis and plays a
central role in tissue morphogenesis and maintenance
XLOC_001338 Inc RNA4563-3 (LNCipedia.org)
XLOC_011215 Inc-C15orf54-2 (LNCipedia.org)
XLOC_011218 Inc-EIF2AK4-1 (LNCipedia.org)
XLOC_012522 Inc-MBTD1-1 ,(LNCipedia.org)
XLOC_l2_012135 Uncharacterised , (LNCipedia.org)
XLOC_l2_015033 Uncharacterised, (LNCipedia.org)
- 300 - An example of percentage different cells expression before and after PBMCs stimulation overall no signifancnt difference was there:
NK cells CD56+ T-cells CD56-T-cells
Unst Stim Unst Stim Unst Stim
Seronegative 6 6.7 2.6 4.3 66 75
13.3 14 1.8 15 38.5 49.1
5.3 3.1 1.2 1.3 75 76.2
7.6 7.4 2.3 3.1 34 32
10.7 11 2.3 2.5 59 54
Seropositive 9.5 9.5 7.4 8 65 65.5
16.1 15.2 8.5 9.9 57.2 57.8
39.5 39 3.3 4 47 50.2
20 18 7.5 7.8 66.1 57.8
33 32 13.1 14.5 44.5 52.1
- 301 -