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Monitoring CD69 expression in circulating t-lymphocytes following renal transplantation Loo, Jimmy K.M. 1999

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MONITORING CD69 EXPRESSION IN CIRCULATING T-LYMPHOCYTES FOLLOWING RENAL TRANSPLANTATION by JIMMY K.M. LOO B.Sc, The University of British Columbia, 1995 A THESIS SUBMITTED IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF MASTER OF SCIENCE in THE FACULTY OF GRADUATE STUDIES (Department of Medicine) THE UNIVERSITY OF BRITISH COLUMBIA April 1999 © Jimmy K.M. Loo, 1999 In presenting this thesis in partial fulfilment of the requirements for an advanced degree at the University of British Columbia, I agree that the Library shall make it freely available for reference and study. I further agree that permission for extensive copying of this thesis for scholarly purposes may be granted by the head of my department or by his or her representatives. It is understood that copying or publication of this thesis for financial gain shall not be allowed without my written permission. Department of /rW£4f/]L ynejjiliAg" (f^U^cTy Z)f faebic,^) The University of British Columbia Vancouver, Canada Date DE-6 (2/88) ABSTRACT Acute graft rejection is the most frequent barrier to successful renal transplantation. The rejection process begins when transplant recipient T-lymphocytes become activated by graft-derived antigens, which causes these leukocytes to search for and eradicate the foreign organ. Studies were conducted to evaluate the surface expression of CD69, a T-cell activation marker, as an index of the immune status in renal transplant patients and as a potential diagnostic predictor of graft rejection. Peripheral blood lymphocytes were first stimulated in vitro with phytohaemagglutinin (PHA). The baseline and induced CD69 expression in princi-pal T-cell subsets was measured by flow cytometry and compared between normal healthy individuals, end-stage renal failure patients undergoing either haemodialysis (HD) or continuous ambulatory peritoneal dialysis (CAPD) therapy, and long-term stable kidney transplant patients (ST). New renal transplant recipients (NT) were monitored longitudinally during their first three post-transplant months. The temporal kinetics of CD69 expression and the effects of the immunosuppressive drug cyclosporin A (CsA) were also examined. Results indicated that circulating T-cells from healthy subjects expressed negligible levels of CD69. However, its expression was observed after 4 hours of PHA stimulation, with peak response occurring after 24 hours and declining to low levels rapidly after 72 hours. Based on this type of expression kinetics, CD69 was considered as an early marker of T-cell activation. CsA at concentrations that strongly suppressed T-cell blastogenesis only partially inhibited CD69 induction. Several significant variations in the CD69 expression of resting and 24-hr PHA-stimulated T-cells were observed between healthy controls and patients in HD, CAPD and ST groups. Most NT patients maintained a rejection-free post-transplant course, and their T-cells remained non-activated and CD69 l o w . During the first two post-transplant weeks, CD69 induction was reduced to half of the pre-transplant values possibly due to the intense immunomodulation required for rejection prophylaxis. In summary, CD69 emerged as a robust T-cell activation marker since potent immuno-suppression in vivo is required to curtail its induction effectively. Increased expression differentiated recently activated T-cells from resting ones. Hence, an increase in CD69-expressing T-cells in peripheral blood might represent an early indication of acute kidney allograft rejection. iii TABLE OF CONTENTS Abstract ii Table of Contents iv List of Figures vii List of Tables ix List of Abbreviations x List of CD Antigen Designations xiii Acknowledgements xiv Chapter One Introduction 1.1 End-stage Renal Disease (ESRD) 1 1.2 Dialysis as Renal Replacement Therapy 4 1.2.1 Haemodialysis (HD) 5 1.2.2 Continuous Ambulatory Peritoneal Dialysis (CAPD) 5 1.3 Kidney Transplantation 6 1.4 Acute Renal Allograft Rejection 7 1.4.1 Clinical Symptoms and Diagnosis 8 1.4.2 Immunological Monitoring of Peripheral Blood 8 1.5 Transplantation Immunopathology 9 1.5.1 T-lymphocyte Involvement 9 1.5.2 Allorecognition Mechanism 9 1.5.3 Membrane and Nuclear Signalling Pathways in Activated T-cells 12 1.5.4 Cell-mediated Allograft Destruction 16 1.6 Immunosuppressive and Anti-rejection Therapy 17 1.6.1 Cyclosporin A (CsA), Tacrolimus (FK506), and Sirolimus 17 1.6.2 Methylprednisolone (MP) and Prednisone (Pred) 18 1.6.3 Azathioprine (Aza) and Mycophenolate Mofetil (MMF) 19 1.6.4 Muromonab-CD3 (OKT3) and Antithymocyte Gamma-globulin (Atgam).. 19 1.7 Activation Inducer Molecule CD69 .20 1.7.1 Transcriptional and Translational Products of CD69 Gene 21 1.7.2 Signalling Components that Participate in CD69 induction 25 iv 1.7.3 Costimulatory Properties in T-cells 27 1.7.4 Detection in Immune-related Pathologies 27 1.8 Other Surface Markers of Activated T-lymphocytes 29 1.8.1 OX40(CD134) 29 1.8.2 Interleukin-2 Receptor a-chain (CD25) 30 1.9 Mitogenic Mechanisms of Carbohydrate-binding Lectins 30 . 1.9.1 Phytohaemagglutinin (PHA) 30 1.9.2 Concanavalin A (ConA) 31 1.10 Relevance 31 1.11 Objective 32 Chapter Two Materials and Methods 2.1 Study Subjects 34 2.2 Sample Collection 36 2.3 Reagents 38 2.4 Cell Culturing and Activation Experiments 38 2.4.1 Peripheral Blood Mononuclear Cell (PBMC) Isolation 38 2.4.2 P B M C Culturing Conditions 41 2.4.3 Mitogenic Stimulation 41 2.4.4 Co-cultivation with Immunosuppressive Drugs (ISDs) 41 2.4.5 Immunostaining 42 2.4.6 Whole Blood Culturing and Processing Technique 42 2.5 Fluorochrome-conjugated Monoclonal Antibodies (MAbs) 43 2.6 Flow Cytometric Analysis 45 2.7 Cell Proliferation Assay 46 2.8 Statistics 48 Chapter Three Results 3.1 Kinetics Studies 49 3.1.1 Expression Kinetics of Activation Markers in PHA-stimulated T-cells 49 CD69: Early Stage Molecule 49 OX40: Intermediate Stage Molecule 56 CD25: Late Stage Molecule 60 3.1.2 Effects of Mitogen Dose on Activation Antigen Expression 64 3.1.3 Comparison of PHA, ConA, and OKT3 in Inducing CD69 Expression 70 3.2 Effects of Immunosuppressive Drugs on CD69 Expression In Vitro 73 3.2.1 CsA 73 3.2.2 MP 78 3.2.3 Tacrolimus 78 3.3 Whole Blood Activation Marker Analysis 86 3.3.1 CD69 Assay of Circulating T-cells 89 Normal Controls 89 HD and CAPD Subjects 93 Stable Transplant Subjects 94 3.3.2 OX40 Profiles in Control, HD, CAPD, and ST Groups 95 3.3.3 Serial Immune Monitoring of New Allograft Recipients 99 CD69 Expression 102 OX40 Expression 112 PHA-induced Proliferation 118 Activation Marker Levels During Complications 120 Chapter Four Discussion 4.1 Healthy-state Circulating T-cells were CD69 l o w 122 4.2 Stimulation ofCD69 Expression by PHA 123 4.3 Kinetics of CD69 Expression During Polyclonal T-cell Activation 125 4.4 Different Lectins Induced Comparable Activation Phenotypes 127 4.5 Kinetics of OX40 and CD25 Surface Expression 128 4.6 Inhibition of Proliferation by ISDs Failed to Abrogate CD69 Expression 131 4.7 Whole Blood Technique of Measuring Activation Marker Expression 134 4.8 Immune Depression and Stimulation in Dialysis-treated ESRD Patients 137 4.9 Expression of Activation Molecules in Stable Transplant Recipients 140 4.10 Lymphocyte Activation Marker Expression in Early Post-transplant Period 142 Chapter Five Summary 146 Chapter Six References 149 vi LIST OF FIGURES Figure Page 1.1 Flow diagram depicting the immunological and physiological disturbances that occur in chronic kidney failure and replacement therapies 2 1.2 Schematic representation of the cellular and molecular events that precipitate T-cell-mediated renal allograft rejection . 10 1.3 Intracellular cascades linking membrane-proximal stimulatory signals to CD69 gene expression in activated T-cells 13 1.4 The molecular and genetic elements of the human CD69 glycoprotein 22 2.1 Culturing and phenotyping techniques for preparing whole blood lymphocytes and PBMCs for flow cytometric analysis 39 2.2 Double-gating strategy for analyzing activation marker expression 47 3.1 The kinetics of CD69 expression in T-cells 53 3.2 The kinetics of CD69 and OX40 expression in T-cells 58 3.3 The kinetics of CD69 and CD25 expression in T-cells 62 3.4 PHA dose-response curve based on CD69 expression in CD3+T-cells 66 3.5 PHA dose-response curve based on CD69 expression in CD4+T-cells 67 3.6 PHA dose-response curve based on CD69 expression in CD8+T-cells 68 3.7 PHA dose-response curve based on D N A synthesis in PBMCs 69 3.8 Comparing the kinetics of CD69 expression induced by PHA and ConA 71 3.9 Effects of varying CsA concentrations on CD69 kinetics in T-cells 75 3.10 Dose-dependent inhibition of D N A synthesis by CsA 77 3.11 Effects of varying MP concentrations on CD69 kinetics in T-cells 80 3.12 Dose-dependent inhibition of D N A synthesis by MP 82 3.13 Effects of varying tacrolimus concentrations on CD69 kinetics in T-cells 84 vii 3.14 Dose-dependent inhibition of D N A synthesis by tacrolimus 87 3.15 CD69 expression in circulating T-cells of healthy, dialysis and stable RTx subjects 91 3.16 OX40 expression in circulating T-cells of healthy, dialysis and stable RTx subjects 97 3.17 CD3 cell count in the peripheral circulation of new RTx patients 101 3.18 Longitudinal monitoring of CD69 expression in circulating T-cells of new RTx patients 108 3.19 Longitudinal monitoring of OX40 expression in circulating T-cells of new RTx patients 115 3.20 Longitudinal monitoring of D N A synthesis in PBMCs of new RTx patients 119 viii L I S T O F T A B L E S Table Page 2.1 ISD protocol in stable and new renal transplant recipients 35 2.2 Primary causes of ESRD among patient subjects, with their demographic characteristics compared to those of normal controls 37 2.3 Dual and triple combinations of monoclonal antibodies used in cyto-fluorometric phenotyping 44 3.1 CD69 expression kinetics in non-stimulated and PHA-activated T-cells 50 3.2 OX40 expression kinetics in PHA-activated T-cells 57 3.3 CD25 expression kinetics in PHA-activated T-cells 61 3.4 PHA dose-titration experiments based on 24-hr CD69 induction in T-cells 65 3.5 Influence of CsA on PHA-driven CD69 expression 74 3.6 Influence of MP on PHA-driven CD69 expression 79 3.7 Influence of tacrolimus on PHA-driven CD69 expression 83 3.8 CD3 cell counts and CD4/CD8 ratios in healthy, dialysis and stable transplant subjects 88 3.9 CD69 expression in healthy, dialysis and stable transplant subjects 90 3.10 OX40 expression in healthy, dialysis and stable transplant subjects 96 3.11 CD3 cell counts and CD4/CD8 ratios in new {de novo) RTx patients 100 3.12 CsA and prednisone dosage levels and plasma cyclosporine determinations in NT patients 103 3.13 Serum creatinine concentrations in the early post-transplant period 104 3.14 Longitudinal evaluation of CD69 expression in NT patients 105 3.15 Longitudinal evaluation of OX40 expression in NT patients 113 3.16 CD69 and OX40 expression during transplant complications 121 i x LIST OF A B B R E V I A T I O N S Actmark Activation marker ag antigen AICD Activation-induced cell death A I M Activation inducer molecule AP-1 Activation protein-1 APC Antigen presenting cell Asn Asparagine Atgam Anti-thymocyte Gamma-globulin A T L Adult T-cell leukaemia A U adenylate-uridylate Aza Azathioprine B-cell Bone marrow-derived lymphocyte BD Becton Dickinson b.i.d. two times a day BP Bandpass C a 2 + Divalent calcium cation [Ca2 +]i Intracellular concentration of free calcium ions CAPD Continuous ambulatory peritoneal dialysis CD Cluster of differentiation antigen C 0 2 Carbon dioxide ConA Concanavalin A cpm counts per minute C R A C Calcium release-activated channel CRD Carbohydrate recognition domain CRE Corticosteroid response element CsA Cyclosporin A CTL CD8+cytotoxic T-lymphocyte C-type Ca2+-dependent D A G Diacylglycerol DiOG 1,2-s72-dioctanoylglycerol D N A Deoxyribose nucleic acid dTTP deoxy-thymidine 5'triphosphate EA-1 Early activation protein -1 (CD69) ER Endoplasmic recticulum E R K Extracelluar signal-regulated kinase ESRD End-stage renal disease FCS Foetal calf serum Fig. Figure FITC Fluorescein isothiocyanate FK506 Tacrolimus FKBP FK506-binding protein FL Fluorescence FSC Forward scatter fyn X G a GTP-binding protein alpha subunit GFR Glomerular filtration rate GTP Guanosine 5'-triphosphate G V H D Graft-versus-host disease [3H]Thd Tritiated thymidine HD Haemodialysis HIV Human immunodeficiency virus H L A Human leukocyte antigen hrs hours HTLV-1 Human T-cell lymphotropic virus Type-1 ICAM-1 Intracellular adhesion molecule-1 IDDM Insulin-dependent diabetes mellitus IgA/G Immunoglobulin A/G-isotype I K B Inhibitory subunit of N F - K B IL-2(R) Interleukin-2 (receptor) IPs Inositol 1,4,5-triphosphate ISDs Immunosuppressive drugs IU international units INK c-jun N-terminal kinase kBq kilobecquerel kD kiloDalton lck P 56 l c k LFL Logarithmic fluorescence LiCi microcurie MAb Monoclonal antibody M A P Mitogen-activated protein M E K M A P or E R K kinase MFI Mean fluorescence intensity M H C Major histocompatibility complex min minute M L R Mixed lymphocyte reaction M M F Mycophenolate mofetil mOsm milliosmole MP Methylprednisolone mRNA Messenger RNA mSos Mammalian son of sevenless mTOR Mammalian target of rapamycin mW milliwatt N Normal healthy subject ND Not determined NF-AT Nuclear factor of activated T-cells N F - K B Nuclear factor of NF-icB/Rel family N K cell Natural killer cell NS Statistically non-significant NT New (de novo) renal transplant recipient o.d. everyday OH Hydroxy XI 0KT3 Murine anti-human-CD3s MAbs (Muronomab-CD3) PAF PBS + Azide + FCS PBMC Peripheral blood mononuclear cell PBS Phosphate-buffered saline PE R-phycoerythrin PerCP Peridinin chlorophyll protein PHA Phytohaemagglutinin Pi Inorganic phosphate PI3-K Phosphatidylinositol 3-kinase P I P 2 Phosphatidylinositol 4,5-bisphosphate PKC Protein kinase C PLCy Phospholipase C-gamma P M A Phorbol myristate acetate PMT Photomultiplier tube Pred Prednisone Pre-Tx Pre-transplant PTH Parathyroid hormone PTK Protein tyrosine kinase r correlation coefficient RA Rheumatoid arthritis raf-1 p74 r a f-' ras p21 r a s RBC red blood cell RNA Ribose nucleic acid rpm revolution per minute RPMI Roswell Park Memorial Institute RTx Renal transplantation SEM Standard error of the mean SH2 svc homologue domain 2 SI Stimulation index SLE Systemic lupus erythematosus src sacroma SSC Side scatter ST Stable, long-term renal transplant recipient STAT Signal transducer and activator of transcription T A T A D N A sequence of thymidine and adenine nucleotides T-cell Thymus-derived lymphocyte TCR T-cell antigen receptor Th CD4+helper T-lymphocyte TNFa Tumor necrosis factor-alpha V G H Vancouver General Hospital vs. versus v/v volume-to-volume wks weeks xii LIST OF C D A N T I G E N DESIGNATIONS CD2 Sheep red blood cell receptor CD3 Pan T-lymphocyte marker CD4 Helper T-cell co-receptor CD7 T-cell differentiation antigen CD8 Cytotoxic T-cell co-receptor CD14 Monocyte lipopolysaccharide receptor CD25 Interleukin-2 receptor a-chain CD28 T-cell costimulatory receptor for B7 ligands CD45 Leukocyte common antigen CD69 Activation inducer molecule CD134 OX40 Xlll 5 ACKNOWLEDGEMENTS I want to express my deepest thanks to my supervisor, Dr. Paul A . Keown, for giving me the opportunity to work in his laboratory. In addition to the time and effort he has donated to co-ordinate this project, his guidance and ongoing support have been vital in making this study possible. The suggestions and assistance offered by Drs. Jean LeRiche, David Landsberg and Bakul Dalai are very much appreciated. Equally important are the laboratory technicians who performed some of the clinical tests (serum creatinine level, TDx assay, osmolality check) in this study. I also appreciate the Clinical Research Ethics Board of the University of British Columbia for approving the investigation, and I own a huge debt to the physicians, nurses and phlebotomists working at the Vancouver General Hospital Renal Dialysis Unit, Peritoneal Dialysis Outpatient Clinic, Transplant Ward, and Solid Organ Transplant Clinic for their help in sample collection. Most importantly, I extend my gratitude to the study subjects for their donation of blood specimens and to the transplant donors for their invaluable "gifts of life," without which this study would have never been completed. Lastly, I am deeply indebted to my parents, family, friends, and colleagues for their support during the completion of this study. xiv Chapter One Introduction Chronic renal failure is characterized by the loss of excretory, endocrine, and homeostatic functions regularly carried out by the kidney (Figure 1.1) [1-4]. These physiologic dysfunctions require patients to undergo either dialysis treatment, which is inefficient and expensive [2,3,5-7], or renal transplantation (RTx), which is relatively cost-effective and often improves their quality of life [3,8-9]. The transplantation of functional kidneys from cadaveric or living-related donors now represents the treatment of choice for patients with end-stage renal disease (ESRD) [8,10-14]. In Canada, over 900 renal transplants are performed annually, with over 7,600 patients being maintained with functioning grafts [15]. Acute rejection remains as the main obstacle in RTx by causing irreversible graft destruction in the early post-transplant period [3,14,16-18]. A screening assay that assists in the early detection of impending acute rejection may enable prompt therapeutic intervention to prevent the onset of rejection and reduce the immunologic injury to the allograft [19-21]. 1.1 End-stage Renal Disease (ESRD) The pathologic hallmark of ESRD is the progressive destruction of kidney nephrons [2]. The gradual loss of renal function causes uraemia [1-4] and is manifested by a reduced glomerular filtration rate (GFR), an elevated serum creatinine level, the accumulation of metabolic toxins and medium-sized molecules [2,22-23], malnutrition [24-25], hyperparathyroidism [2-3], osteodystrophy caused by hypocalcaemia and the depressed synthesis of 1,25-dihydroxy-vitamin D 3 [3], and anaemia due to impaired erythropoietin synthesis [2-3] (Fig. 1.1). Without receiving renal replacement therapy such as dialysis or transplantation, ESRD patients will die quickly because of these disturbances. 1 Figure 1.1. F l o w d iagram depic t ing the i m m u n o l o g i c a l and phys io log ica l disturbances that occur in chronic k idney fa i lure and replacement therapies. B r o k e n arrows indicate the immune defects that are caused by phys io log ica l perturbat ions. Abbrev ia t ions : C a 2 + , ca lc ium cat ion; C A P D , cont inuous ambula tory per i toneal d ia lys is; C T L , C D 8 + cyto tox ic T -ce l l ; E S R D , end-stage renal disease; G F R , g lomeru lar f i l t ra t ion rate; H D , haemodia lys is ; I L - 2 ( R ) , in ter leuk in-2 (receptor) ; ISDs, immunosuppress ive drugs; O H , h y d r o x y ; P B M C , per ipheral b lood mononuc lear ce l l ; P T H , para thyro id hormone; R T x , renal t ransplantat ion; T C R , T-ce l l ant igen receptor; T h , C D 4 + h e l p e r T -ce l l ; T N F a , t u m o r necrosis factor-a lpha. 2 P R I M A R Y Progressive DISEASE renal dysfunction ESRD symptom complex: U R A E M I A Physiological ~1 •I 1 a-hydroxylase activity ^1,25-diOH vitamin D i renal reabsorption of and -l intestinal absorption of C a 2 + i •I plasma C a 2 + (hypocalcaemia) I t PTH production J Hyperpara-thyroidism Dialysis HD I 1 • i erythropoietin anemia 1 malnutrition L f - - - ' i glomerular f" filtration i t uraemic toxins e.g. guanidino compounds, middle-size molecules I • I GFR • T serum creatinine • t blood urea nitrogen Immunological t cell-mediated immunity depression 4- Th and CTL subsets u -V - i I P B M C proliferation ilL-2 synthesis I TCR/CD3 density activation t plasma TNFa T baseline IL-2R . T infection susceptibility t malignancy CAPD Complement Activation T circulating . CTL RTx I ISDs Peritonitis 3 The physiological changes associated with kidney failure also cause the immune system impairment that are frequently observed in ESRD. The immune dysfunction is manifested by a moderate lymphocytopenia, an increased susceptibility to infection, modulated immune effector functions, an increased incidence of neoplasia, and a poor vaccination response (Fig. 1.1) [22,24-25]. The deficiency is likely due to a state of defective cellular immunity in which the key host-defence cells, the thymus-derived (T-) lymphocytes, fail to produce adequate growth factors (e.g. interleukin-2 or IL-2) in response to antigenic challenge [23]. Anaemia, malnutrition and uraemic middle-sized molecules have been suggested to be causative factors of T-cell dysfunction [23-26], and some of them have been shown to inhibit the growth of in vzYro-stimulated lymphocytes [23]. Intriguingly, various signs of chronic activation such as elevated serum IL-2 levels are occasionally present. These may represent either the adaptive efforts of the immune system to compensate for its deficiency or chronic immune stimulation via contact with the dialysis membrane [27-29]. However, prolonged immune stimulation might lead to functional exhaustion and thus further depress immune competence [22,30-31]. 1.2 Dialysis as Renal Replacement Therapy The critical donor organ shortage makes transplanting every ESRD patient an unfeasible option [5]. Consequently, most patients rely on maintenance dialysis which primarily consists of two types: haemodialysis (HD) and continuous ambulatory peritoneal dialysis (CAPD). Both modalities act as "artificial kidneys" in the aim of removing metabolic and toxic wastes from the circulation. Selecting the dialytic technique suitable for a particular patient requires the consideration of the clinical benefits peculiar to each method and of 4 socio-economic issues [6,32-34]. Immunologic factors, however, are seldom considered since dialysis often fails to alleviate the ESRD-associated immune dysfunction [22,28]. 1.2.1 Haemodialysis (HD) Blood filtration in HD therapy involves the diffusion of low molecular weight substances across a synthetic membrane between the peripheral blood and a balanced salt solution [3,6,24]. A vascular access (e.g. an arteriovenous fistula) in the haemodialyzed patient allows the blood to enter into the capillaries of the dialyzer which are porous to urea, creatinine and electrolytes, and these substances are filtered into a surrounding bath of dialysate. Ultrafiltration regulates the body fluid content by removing excess water. A defective immune response can be demonstrated in many HD-treated patients, but whether this represents a treatment complication or the pathogenic effect of renal failure remains unclear. The various components of the dialyzer, such as the synthetic membrane, cause the immunologic stimulation of leukocytes, and complement activation can occur depending on the dialysis membrane composition [6,30-31]. The utilization of HD is also associated with a dampened activity of the autoimmune disorders that contribute to ESRD, an impaired blastogenic response in lymphocytes toward polyclonal stimuli, and a high incidence of infection [30,35]. The expression of certain surface adhesion and activation molecules in T-lymphocytes are upregulated, while others are decreased [30-31,35]. 1.2.2 Continuous Ambulatory Peritoneal Dialysis (CAPD) In CAPD, the semi-permeable peritoneum functions as the dialysis membrane. A concentration gradient is created between the circulation and peritoneal cavity following the 5 gravity instillation of dialysis solution into the abdomen through an implanted catheter [3,7]. Plasma urea and other waste products diffuse out from the peritoneal capillaries into the dialysate. The solution remains in the abdominal space for several hours before being drained, and continuous dialysis is achieved by immediate refilling with fresh dialysate. The most common complication associated with CAPD is bacterial peritonitis [3,7,29,36]. This occurs with a mean frequency of one episode per patient per year, and the residual scarring of the peritoneal membrane can preclude this method of dialysis [5]. The procedure itself facilitates the introduction of microbes into the peritoneal environment. The ability of abdominal leukocytes to co-ordinate host defence is impaired by the non-physiologic pH and high osmolality of the instilled dialysate and by the constant removal of immunoglobulins and immunocompetent cells [37-38]. In addition, lymphocytopenia, increased baseline intracellular calcium concentrations in peritoneal lymphocytes, and an abnormal upregula-tion of peripheral activated T-cells are also observed in these patients and might contribute to their immune deficiency [24,28-29,36]. 1.3 Kidney Transplantation The multiple advantages of RTx, including the restoration of kidney functions, the improvement of health and the elimination of dialysis requirement, make it the optimal treatment for renal failure. Prior to transplantation, donor-recipient genetic compatibility must be determined to minimize the possibility of graft rejection [39]. A high degree of match between their major histocompatibility complex (MHC) genes is desirable. The protein products of these genes, known as -the human leukocyte antigens (HLA), are 6 structurally similar when the donor and the recipients are compatible, thus reducing the potential of graft rejection [39-41]. Organ retrieval is performed by removing the left or right donor kidney (nephrectomy) along with the renal artery, vein, and ureter. After washing and perfusion with cold preservation fluid, the kidney is placed in the iliac fossa of the recipient. Subsequently, the donor renal vasculature is attached to the respective iliac blood vessels in the recipient while the ureter is anastomosed to the bladder. Renal allograft recipients require continuous immunosuppression to prevent the rejection of their transplants [42-43]. Current protocols rely on the use of cyclosporin A (CsA), predni-sone (Pred) and azathioprine (Aza) in a triple-drug regimen that blocks the immune response to the allograft while minimizing their individual toxicity [10-11,44-46]. Graft functions as indicated by serum creatinine levels are monitored to identify signs of transplant rejection. Although acute rejection represents the major cause of early graft loss, non-immunological complications can also precipitate graft dysfunction in the initial post-surgical period. For example, acute tubular necrosis is caused by ischaemia and reperfusion injury during the transplantation procedure [47]. Because of their immunocompromised state, transplant patients are also subject to infection, neoplasia and lymphoproliferative diseases [11,13]. 1.4 Acute Renal Allograft Rejection The immune system can differentiate between self-HLA antigens and non-self-HLA antigens. It can identify the allograft as non-self and mobilize its immunocompetent cells to 7 destroy it. Acute rejection is reported in almost half of the transplant patients, depending on the level of prophylactic immunosuppression and the degree of donor-recipient H L A matching [8,11,39,48-49]. It occurs predominantly within the first three post-transplant months and reduces one-year graft survival rate by 8 to 12% [10,16]. 1.4.1 Clinical Symptoms and Diagnosis The classical symptoms of acute kidney rejection are fever, oliguria, oedema, and graft tenderness [8]. An elevation of serum creatinine level often suggests rejection [17,50], and the histopathological examination of tissues obtained via core needle biopsy is the gold standard for diagnosis [12,21,51]. Unfortunately, biopsies are invasive, and often rejection can only be diagnosed with confidence when the inflammatory process has already caused extensive transplant injury [18,52-54]. 1.4.2 Immunological Monitoring of Peripheral Blood Strategies for monitoring the peripheral blood for signs of alloreactivity have been extensively studied, but their effectiveness in diagnosing and predicting transplant rejection remains controversial [19-21,51-52,55-63]. While some investigators suggested that the immunocompetent cells in the circulation are functionally and phenotypically distinct from those that mediate local inflammatory events such as transplant rejection [52,59], others have claimed that the peripheral cells are responsible for the allogeneic response [18,55-58, 60-62,64-66]. Naive cells may be alloactivated within the transplanted organ or in regional lymphoid tissues such as peripheral lymph nodes. They subsequently home to the sites of inflammation through the circulation and then undergo proliferation within the graft [67]. 8 Peripheral cellular immunity might also increase during immune processes such as infection [20,21]. In recent years, research has focused on examining the activation of recipient circulating lymphocytes, investigating the cause of this activation, and tailoring the treatment of rejection or non-rejection complications accordingly [18,20,21,52,55-58,60-61,64-66,68]. 1.5 Transplantation Immunopathology 1.5.1 T-lymphocyte Involvement Acute rejection is most commonly due to a cellular immune response directed against the transplant. The cardinal leukocytes that initiate and co-ordinate the anti-allograft process appear to be the T-lymphocytes [12,14,53,55,69-70]. These cells originate from the bone marrow and develop in the thymus to become mature T-cells that circulate in the peripheral blood. An estimated 2% of recipient circulating T-cells can be activated through allo-stimulation [71]. They identify foreign proteins (antigens) in the allograft or lymph nodes either directly or by interacting with professional antigen-presenting cells (APCs) such as dendritic cells and monocytes (Fig. 1.2). After recognizing non-self peptides, these T-cells become activated and may re-enter the circulation and home to the allograft, which is the source of the alloantigens [67]. Subsequently, they attach to the blood vessel endothelium and migrate into the transplant, where they co-ordinate various inflammatory reactions such as the secretion of pro-inflammatory cytokines. These cells also recruit other cytotoxic cells into the area which induce parenchymal damages (tubulitis) in the allograft. 1.5.2 Allorecognition Mechanism Both H L A class I and II molecules are presented on the surface of APCs. The clonotypic T-cell antigen receptor (TCR) on the surface of T-cells binds through its polymorphic regions 9 Figure 1.2. Schematic representat ion o f the cel lu lar and molecu lar events that precipi tate T-ce l l -media ted renal a l lograf t re ject ion. Shading = al logeneic. Abbrev ia t ions : ag, ant igen; A P C , ant igen present ing ce l l ; C T L , C D 8 + c y t o t o x i c T -ce l l ; I L - 2 , in te r leuk in -2 ; M H C , major h is tocompat ib i l i t y comp lex ; T C R , T -ce l l ant igen receptor; T h , C D 4 + h e l p e r T-ce l l . 10 Direct presentation Indirect presentation Migration of passenger leukocytes Shedding of alloantigens Donor M H C ' - T F T f Donor ag S^V T C R SelfMHC-Donor ag Allospecific T-cells (Th and CTL) I CD69 participation Activation phase t OX40 participation Tissue homing t Adherence to vascular endothelium I CD25 participation Allograft infiltration Arterial vascular endothelium t Proliferation phase t autocrine growth " IL-2 Pathological damages blood vessel wall destruction and haemorrhage IE tubulointerstitial and epithelial tissues t interstitial oedema, inflammation, and nephron tubular cell loss I Graft Rejection 11 to the amino acid residues of the H L A molecule and the bound peptide, thus conferring self-M H C restriction and antigen specificity [11]. The T-cell co-receptors, class II-binding CD4 and class I-binding CD8, bind to the same M H C protein as the TCR and differentiate T-cells into two functional subtypes known as the helper T-cells (Th) and the cytotoxic T-cells (CTL), respectively. Allorecognition can occur through the direct and indirect pathways (Fig. 1.2). According to the direct model [41,66], intragraft donor APCs present allo-HLA molecules with bound peptides on their surface directly to recipient T-cells. For indirect presentation, recipient APCs endocytose H L A antigens shed from the graft, process them into allopeptides and present them in the context of self-HLA molecules to alloresponsive T-cells. Current studies show that both pathways can elicit rejection [41,66]. 1.5.3 Membrane and Nuclear Signalling Pathways in Activated T-cells The functional TCR complex consists of the antigen receptor in association with a family of CD3 (y, 5, 8, C, and n) glycoproteins (Fig. 1.3). Upon antigen recognition, the CD3 molecules trigger intracellular signalling cascades by initiating the interaction of membrane-bound signal mediators and protein tyrosine kinases (PTK). Attached to the cytoplasmic tails of the T-cell co-receptors and CD3<^ chains are two principal src (sarcoma)-family PTK known as p56 l c k and p59 f y n [72]. The activation of these kinases requires the dephosphorylation of their negative regulatory phosphotyrosine residues by CD45 phosphatase [73], and the activated enzymes proceed to phosphorylate their cytosolic targets, including the tyrosine residues of C, chains. Situated in strategic sequence motifs, these 12 Figure 1.3. Intracellular cascades linking membrane-proximal stimulatory signals to CD69 gene expression in activated T-cells. Abbreviations: ag, antigen; AP-1, activation protein-1; APC, antigen presenting cell; Ca 2 + , calcium cation; [Ca 2 +] i 5 intracellular concentration of free calcium ions; C R A C , calcium release-activated channel; D A G , diacylglycerol; ER, endoplasmic recticulum; E R K , extracelluar signal-regulated kinase; fyn, p59f>'n; GTP, guanosine 5'-triphosphate; I K B , inhibitory subunit of NF-kB; IP 3, inositol 1,4,5-triphosphate; INK, c-jun N-terminal kinase; lck, p56 l c k; M E K , M A P or E R K kinase; M H C , major histocompatibility complex; mSos, mammalian son of sevenless; NF-AT, nuclear factor of activated T-cells; N F - K B , nuclear factor of NF-kB/Rel family; P,, inorganic phosphate; PI3-K, phosphatidylinositol 3-kinase; PIP 2 , phosphatidylinositol 4,5-bisphosphate; PKC, protein kinase C; PLCy, phospholipase C-gamma; raf-1, p74 r a f"'; ras, p21 r a s ; TCR, T-cell antigen receptor; TNFa, tumor necrosis factor-alpha. 13 A P C T-cell Nucleus c-jun gene expression • ^1 ( N F - A T ) t A P - i ; ( N F - K B ; activation C D 6 9 gene N F - A T A P - 1 N F - K B site site site • 14 phosphotyrosines act as the docking sites for SH2 (src homologue domain 2)-containing enzymes such as phospholipase C-gamma (PLCy) and allow them to be phosphorylated and activated by the src kinases [74]. The active PLCy hydrolyses a membrane lipid called phosphatidylinositol 4,5-bisphosphate ( P I P 2 ) into two second messengers known as the inositol 1,4,5-triphosphate ( I P 3 ) and diacylglycerol (DAG). IP3 is responsible for mobilizing calcium ions (Ca 2 +) from intracellular storage sites such as the endoplasmic recticulum (ER) into the cytosol [75]. The cytoplasmic free cations bind to a Ca2+-activated regulatory protein, calmodulin, which interacts with the serine/threonine phosphatase calcineurin [75-78]. The resultant Ca 2 + -calmodulin-calcineurin complex dephosphorylates a gene transcription regulator, the nuclear factor of activated T-cells (NF-AT), thereby activating it and permitting it to translocate into the nucleus. Nuclear NF-AT joins with the transcription activating protein-1 (AP-1), which consists of the proto-oncogenic proteins c-fos and c-jun. Docking of this complex onto the NF-AT binding sequence in the promoter of IL-2 gene activates its transcription [75-80]. The second messenger D A G and the cytosolic free calcium co-activate protein kinase C (PKC), a serine/threonine protein phosphatase [74]. This enzyme activates a signalling pathway involving the extracellular signal-regulated kinases (ERK 1/2) and their upstream mediator, the dual-specificity (serine/threonine- and tyrosine-phosphorylating) kinase called mitogen-activated protein (MAP)/ERK kinase (MEK). The transcription of c-fos gene is under the control of E R K [81-83]. An obligatory T-cell activation costimulatory signal is provided by the binding of CD28 in T-cells to a B7 molecule in accessory cells. This 15 interaction triggers the phosphatidylinositol 3-kinase (PI3-K), the PTK associated with CD28 [84], to stimulate the c-jun N-terminal kinase (INK) , which is responsible for enhancing the transcriptional activity of the c-jun gene [81-83]. The ERK-dependent signal transmission cascade is also mediated by a regulatory guanosine 5'-triphosphate (GTP)-binding protein known as p21 r a s (ras). It catalytic function becomes active in the GTP-bound state, which is controlled by the guanine nucleotide-exchanging protein mSos (mammalian son of sevenless). The mSos protein in turn relies on two SH2 adapter molecules known as She and Grb-2 to couple to the CD3 chains in order to become activated by CD3-associated PTKs. The consequence of p21 r a s activity is the activation of p74 ra f-1 (raf-1), a serine/threonine protein kinase upstream of the E R K enzymes [81,85-86]. 1.5.4 Cell-mediated Allograft Destruction After allorecognition, T-cells advance into their activation phase, progress through the cell cycle and transform into rapidly dividing lymphoblasts that are supported by the autocrine growth cytokine IL-2 [11,87-89]. During clonal expansion, the blast cells differentiate into effector T-cells that target the graft tissues [11,67,87-89]. The renal vasculature is the first site of their immune attack because of its accessibility to circulating effector cells, which mediate the direct cytolysis of vascular endothelial cells in arterioles and small arteries. The injury to the donor organ is also potentiated by lymphotoxins such as tumour necrosis factor-a (TNFa) that are synthesized and secreted by activated T-cells, macrophages and natural killer (NK) cells. These soluble and cellular mediators stimulate the endothelial cells to upregulate their expression of adhesion proteins such as the intracellular adhesion molecule-16 1 (ICAM-1) and M H C class-II proteins like HLA-DR [90-92], thereby facilitating antigen presentation and T-cell entry into the graft interstitum. The infiltrating mononuclear cells then invade the interstitial tissues, causing inflammation and oedema. As the renal tubular cells become injured, the nephrons lose their functional capacity [54,93-94]. Th and CTL cells displaying activation-associated markers such as the IL-2 receptor (IL-2R) are often found in high concentrations in the cellular infiltrates [53,65,91-92]. 1.6 Immunosuppressive and Anti-rejection Therapy Prophylactic immunosuppression normally involves the use of combination drug therapy [11,45]. Acute allograft rejection is normally treated first with intravenous methylprednisolone (MP); however, in steroid-resistant cases, muromonab-CD3 (OKT3) or antithymocyte (equine anti-human) gamma-globulin (Atgam) is used [11,44,95]. 1.6.1 Cyclosporin A (CsA), Tacrolimus (FK506), and Sirolimus CsA is a fungal cyclic undecapeptide capable of suppressing the activity of allostimulated T-cells in vitro and in vivo [96-98]. This agent binds to the cytosolic immunophilin cyclophilin [76,99]. The resultant complex then attaches to the phosphatase activity site of calcineurin 2"f* and impairs its ability to dephosphorylate and activate NF-AT even when Ca -calmodulin binding has occurred [77-78]. The resulting blockade in IL-2 synthesis prevents the autocrine growth of activated T-cells, thus accounting for the ability of this agent to suppress renal graft rejection and the proliferation of mitogen-stimulated T-cells [11,44]. Tacrolimus (FK506), derived from the fungus Streptomyces tsukabaensis, inhibits the proliferation of allostimulated T-cells through mechanisms similar to those of CsA 17 [11,44,100-101]. This compound binds to the immunophilin FK506-binding protein (FKBP)-12, and the drug-receptor complex attaches to the active site of calcineurin, hence preventing the activation and nuclear translocation of NF-AT. As a result, IL-2 gene expression and cell proliferation are inhibited [77-78,101-103]. Preliminary results from clinical trials demonstrated that this cyclic marcolide possesses equal or greater potency than CsA in preventing and reversing acute kidney rejection [95,104]. In contrast, sirolimus acts on the IL-2 receptor signalling pathways. This agent combines with an FKBP immunophilin and subsequently inhibits the protein kinase known as the "mammalian target of rapamycin" (mTOR), which participates in the activation of the protein translation regulator p70 S6 kinase and in the removal of cyclin inhibitors. The effect of this agent is manifested by the cell cycle arrest of IL-2-stimulated T-cells [105-107]. 1.6.2 Methylprednisolone (MP) and Prednisone (Pred) Corticosteroids, including MP and prednisone, mediate broad-spectrum anti-inflammatory effects that are capable of preventing and reversing rejection [102,108]. High-dose steroid administration induces T-lymphocytopenia, apoptosis, and the downregulation of IL-2 and TNFa synthesis in activated T-cells [44,64,102,109-110]. Corticosteroids and their cyto-solic glucocorticord receptors form drug-receptor complexes that bind to the corticosteroid response elements (CRE) within the enhancer regions of many cytokine genes, thus inhibiting their expression [111]. Additional effects such as interfering with the AP-1 and NF-AT binding that are necessary for the transcriptional initiation at the IL-2 gene promoter were also demonstrated in the work of Vacca et al. [112]. 18 1.6.3 Azathioprine (Aza) and Mycophenolate Mofetil (MMF) Aza interferes with the de novo nucleic acid synthesis required by activated T-cells to achieve proliferation [11,44,102]. This agent is an imidazole derivative of 6-mercaptopurine and is converted to its precursor in the liver and phosphorylated. The drug inhibits purine biosynthesis, deoxyribose nucleic acid (DNA) replication and ribose nucleic acid (RNA) production. Lymphocytes have limited ability in synthesizing nucleotides through the salvage pathways, which rely on recycled materials from nucleic acid breakdown. In contrast to Aza, M M F demonstrates lymphocyte specificity by specifically blocking the production of GTP, which is a type of nucleotide needed by activated T-cells to produce DNA, RNA, and glycosylated adhesion molecules [11,44,102]. 1.6.4 Muromonab-CD3 (OKT3) and Antithymocyte Gamma-globulin (Atgam) The murine OKT3 antibody specifically targets the human CD3 s-chain [11], resulting in the shedding of the CD3 molecules and the antigen receptors. T-cells thus become non-responsive to alloantigen stimulation [102]. Opsonization of OKT3-bound T-cells by the reticuloendothelial system causes lymphocytopenia, and modulated cells that reappear after OKT3 treatment remain transiently incapable of proliferation under in vitro mitogenic and allogeneic activation [113-116]. Atgam is a purified immunoglobulin G-isotype (IgG) fraction of the anti-lymphocyte serum isolated from horses immunized with human thymocytes. The administration of Atgam reduces the level of circulating T-cells via complement-dependent lysis and targeted destruction by marcophages, thus lessening the frequency and intensity of early graft 19 rejection [44,117-120]. The antibodies target a diverse array of epitopes, mask numerous T-cell proteins such as CD2, CD3, CD4, CD8, and IL-2R and block their signal-transduction capacities, hence suppressing cell activation [121-122]. 1.7 Activation Inducer Molecule CD69 Lymphocyte activation involves the participation of numerous signalling receptors that have minimal functional roles in resting cells [123]. The human CD69 molecule is an example of these signal-transducing membrane glycoproteins [124]. Corte et al. [125] in 1981 first isolated the anti-CD69 monoclonal antibody (MAb) from the antisera of mice immunized with human T-cells that were activated in a mixed lymphocyte reaction (MLR). This immunoglobulin reacts with the peripheral blood mononuclear cells (PBMC) of allogeneic bone marrow transplant patients with active graft-versus-host disease (GVHD) but not with those of healthy individuals. This target protein was later assigned the cluster of differentiation (CD) antigen number 69 (CD69) at the Fourth Leukocyte Typing Workshop in 1989 [126]. It is also designated as the activation inducer molecule (AIM), early activation antigen (EA-1), MLR-3, and Leu-23 [125,127-129]. The protein is transiently displayed in activated T-cells and appears to be the earliest surface marker expressed following their activation. Through binding to a not-yet-identified carbohydrate ligand, this receptor can trigger and potentiate various signalling cascades required for novel gene expression and activation-associated events [124]. Many haematopoietic cells express CD69 upon activation, including T-cells, bone marrow-derived (B-) cells, N K cells, eosinophils, and neutrophils [127,130-132]. Constitutive sur-20 face presentation during quiescence, however, is restricted to circulating monocytes, platelets, thymocytes, and epidermal Langerhans cells [133-136]. Similar to other cell-cycle-specific cytokine genes and proto-oncogenes, the CD69 gene remains dormant in resting T-cells and neither its transcripts nor protein products can be detected, indicating that quiescent lymphocytes do not require its participation. Stimulation of T-cells in vitro with antigens such as tetanus toxoid results in weak A I M expression (CD69 l o w), and profound in-duction (CD69 h ' s h) is achieved using mitogenic lectins such as phytohaemagglutinin (PHA) and concanavalin A (ConA), antibodies directed to CD2 or CD3, the P K C stimulator phor-bol myristate acetate (PMA), superantigen Staphylococcal enterotoxin B, and TNFa [114, 128,137-138]. Although the circulating lymphocytes of healthy persons rarely express CD69, it is detected in T-cells localized to tissues that are subject to constant antigen exposure, including endometrial mucosa, intestinal lamina propria, lymph nodes and tonsils [139-142]. 1.7.1 Transcriptional and Translational Products of CD69 gene Genomic library screening reveals that the human CD69 gene is non-polymorphic and composed of five exons, with the first and second exons encoding for the cytoplasmic and transmembrane portions of the core glycoprotein, respectively, and the last three exons for the extracellular structure, which contains the specialized carbohydrate recognition domains (CRD) for ligand binding (Fig. 1.4). This 15 kilobase gene is mapped to the band region 12.3-13.2 of the short arm (p) of chromosome 12 and postulated to be clustered with other lymphoid and N K cell-specific genes in a hypothesized " N K gene complex" [128-129,138,143-146]. Indeed, such a complex exists on mouse chromosome 6, and the CD69 21 Figure 1.4. The molecular and genetic elements of the human CD69 glycoprotein. C = carboxyl terminal, E = exon, N = amino terminal, P = phosphorylation, S-S = disulfide bond, black boxes = introns, gray boxes = transcription factor binding sites, black circles = serine/threonine residues, open circles = C a 2 + ions, and hexagons = carbohydrate moieties. Abbreviations: AP-1, activation protein-1; Asn, asparagine; Ca 2 + , calcium cation; Got, GTP-binding protein alpha subunit; IL-2(R), interleukin-2 (receptor); NF-AT, nuclear factor of activated T-cells; N F - K B , nuclear factor of NF-kB/Rel family; PLCy, phospholipase C-gamma; T A T A , D N A sequence of thymidine and adenine nucleotides. 22 "Asn-linked glycosylation CD69 ligand with carbo-hydrate moieties Ligand binding site Carbohydrate Recognition Domain Extracellular domain Extracellular Space Cytoplasmic membrane — T H — T A T A — P ~ i 1 N F - K B E l [ E2 [ E3 [ E4 [ E5 Human CD69 gene (Chromosome 12pl2.3-13.2) 23 genes are highly conserved between the two species [147]. Within the promoter region of the human gene are putative binding motifs of transcription factors (Fig. 1.4) [148]. CD69 expression in activated T-cells requires de novo biosynthesis, with its messenger RNA (mRNA) becoming detectable 30 to 60 minutes after stimulation through the TCR/CD3 complex [146-147]. These transcripts are rapidly degraded in 4 to 6 hours due to the adenylate-uridylate (AU)-rich sequences in their 3' untranslated regions that are specifically recognized by ribonucleases in cellular degradation pathways [149-150]. In contrast, the surface expression of CD69 is detected after 2 to 4 hours and persists for a longer period of time, reaching its peak level by 18 to 24 hours and declining afterwards [114,129,149-150]. The CD69 protein belongs to the lectin superfamily of Type II integral membrane receptors (amino termini intracellular, carboxyl termini extracellular, Fig. 1.4) [145-146,151]. This 60 kiloDalton (kD) surface molecule exists as a disulfide-linked hetero- or homodimeric transmembrane protein capable of binding sugar moieties in a calcium (Ca2+)-dependent manner (C-type). It consists of two subunhs with molecular weights of 28 and 33 kDa, and both are formed from 24 kDa core polypeptides [127,152-153]. Vance et al. [152] have suggested that the molecular weight heterogeneity is a consequence of differential glycosylation at asparagine (Asn) residues located within the Asn-glycan addition sites in the C-type CRD domains. Several serine residues in the intracellular domains are constitutively phosphorylated, an indication of their participation in cytosolic signalling [127,144]. 24 1.7.2 Signalling Components that Participate in CD69 induction Enzyme cascades that become functional during lymphocyte activation are crucial in linking membrane-proximal events, such as antigen recognition and ligand-receptor interaction, to nuclear changes, such as the induction of gene expression by activated transcriptional factors [74,81,154]. Many signalling components in these pathways have been demonstrated to be mediating the transcription of the CD69 gene. Among the various intracellular events in T-cell activation, a sustained elevation of intracellular free calcium concentration ([Ca ]j) is important in inducing CD69 expression (Fig. 1.3). The IP3-mediated release of C a 2 + from intracellular stores is short-lived and insufficient to stimulate the surface expression of this activation marker (Actmark) [129]. A prolonged increase in [Ca2+]j is achieved by influx through cytoplasmic membrane channels, such as the calcium release-activated channels (CRAC). Indeed, the opening of the C R A C 9-1-in the absence of other T-cell stimuli permits Ca entry and promotes CD69 expression, while both events are blocked by inhibiting these channels with lanthanum (La 3 +) or by removing extracellular C a 2 + using cation chelators [155-156]. A large body of evidence supports the hypothesis that the primary CD69 inductive event downstream of TCR/CD3 stimulation is PKC activation [128-129,155-157]. T-cells rapidly upregulate their CD69 expression when stimulated with PKC activators such as P M A and 1,2-sn-dioctanoylglycerol (DiOG), a synthetic D A G , while the induction is abrogated by staurosporine H-7, a potent PKC inhibitor [128,155,157]. Downstream of the PKC cascade is M E K , which appears to be the convergence point of the PKC-dependent pathway 25 responsible for CD69 upregulation and another pathway that relies on PTK but independent of PKC [158-160]. This alternative pathway involves tyrosine phosphorylation and is demonstrable through CD3 triggering. It appears to allow the activation of M E K and E R K to occur even when PKC function is blocked. Two independent groups have provided evidence that CD69 gene expression is indeed controlled by the signalling components under the PTK-dependent pathway, namely the oncoproteins p21 r a s and p74 raf"' [160-162]. They demonstrated that transfected leukaemic Jurkat T-cells expressing the constitutively active form of either p21 r a s or p74 raf"' are CD69 + without external stimulation, and those cells with the dominant negative form of either enzyme have impaired upregulation of this marker when triggered with anti-CD3 antibodies. In lymphocytes, p74 r a f _ 1 is found to activate M E K without relying on PKC [159]. The kinase activity and nuclear translocation of E R K enzymes extend the activation cascades into the nucleus, leading to the transcription of genes encoding for nuclear factors such as c-fos. Hetero- or homologous dimerization of c-fos and c-jun yields AP-1 [163], a transcriptional activator that has two putative binding sites within the CD69 gene promoter region. The ability of AP-1 to regulate CD69 expression is evidenced by the fact that antioxidants that bypass the antigen-receptor signalling pathways to activate AP-1 can also induce this marker [164]. The activity of this factor might also account for the ability of anti-CD28 MAbs to trigger CD69 expression (Fig. 1.3). NF-AT, the transcription partner of AP-1, appears to play a less significant role in controlling CD69 induction [165]. The N F - K B / R e l family of transcription factors are activated by inflammatory stimuli such as TNFa, which cause the dissociation of their inhibitory subunit I - K B and allow them to 26 translocate from the cytosol into the nucleus. Within the 5'promoter of CD69 gene are three N F - K B binding sequences, and TNFot-activated NF-KB/Rel proteins were found to interact with these motifs and participate in the transcriptional activation of CD69 expression [148]. 1.7.3 Costimulatory Properties in T-cells Most of the CD69 costimulatory functions have been elucidated by using anti-CD69 MAbs in conjunction with other T-cell polyclonal activators. Antibody-crosslinking of the CD69 receptors without other stimuli elevates [Ca2+]j via the transmembrane influx of extracellular calcium but fails to activate PKC or induce proliferation [166]. To generate competent agonistic signals, CD69 must act in synergy with other specific or non-specific stimulators. When PHA and PKC activators such as P M A are used with antibodies targeting this molecule, D N A synthesis, IL-2 production, and IL-2 receptor expression are augmented [167]. CD69 costimulation has also been shown to enhance NF-AT and AP-1 binding to IL-2 gene enhancer regions [168]. A study by Risso et al. [169] demonstrated that CD69 is physically associated with a Bordetella pertussis toxin-sensitive GTP-binding protein that might link the activation protein with PLCy. Its costimulatory action might also serve a pathogenic role in inflammation, since signalling through this protein augments TNFa synthesis, and this cytokine in turn positively regulates CD69 gene transcription [123]. 1.7.4 Detection in Immune-related Pathologies CD69 has been increasingly used to identify activated T-cells that are implicated in causing and aggravating both acute and chronic immune disorders. In inflammatory diseases such as rheumatoid arthritis (RA), most synovial T-cells have been shown to be CD69 + , indicating 27 that these lymphocytes are in a persistently activated state due to autoantigen challenge, pro-inflammatory mediators, or activated marcophages within the inflamed joint [170-174]. CD69+T-cells are also identified in the mononuclear cell infiltrates of halo naevi, ashy dermatitis, chronic active hepatitis, and bronchoalveolar lavage fluid after swine dust inhalation [175-178]. The tumour-infiltrating lymphocytes in human breast carcinoma and melanoma metastases exhibit CD69 expression, and this molecule is observed on circulating T-cells after cancer vaccine immunotherapy [179-181]. This early activation marker is also demonstrated in studies focusing on peripheral T-cells in human immunodeficiency virus (HlV)-seropositive patients and in those individuals with systemic lupus erythematosus (SLE) or G V H D [125,179-184]. The T-cells that are infiltrating acutely rejected heart and renal allografts in humans are identified immunohistochemically and cytofluorometrically to be CD69 + , suggesting the possible involvement of this marker in the pathogenesis of graft injury [53,65,185-187]. Alloreactive T-cells from the peripheral circulation of these transplant patients also up-regulate this marker in donor-specific allogeneic cultures, and similar phenotypic changes are demonstrated in the M L R of normal PBMC [66,188]. The utility of CD69 induction for detecting lymphocyte activation defects has been investigated in HIV infection. Circulating T-cells of HIV-positive individuals are defective in expressing the marker upon in vitro mitogen stimulation [115,189]. Similar defects were observed in human type-1 (insulin-dependent) diabetes mellitus (IDDM) and SLE [183,190]. 28 1.8 Other Surface Markers of Activated T-lymphocytes 1.8.1 OX40(CD134) Human OX40 plays an important role in regulating the adhesion of activated lymphocytes to vascular endothelium [191], an event which is critical for effector cells to initiate their extravasation into specific tissues [67,192]. Originally identified as a 50 kDa protein in rat CD4 +T-cell blasts, the human homologue belongs to the nerve growth factor receptor superfamily and was recently assigned as CD 134 [67,192-195]. This protein is only detected on activated T-cells and participates in T-cell costimulation as demonstrated by its ability to augment mitogen-induced proliferation [193,196]. Two research groups have identified its cognate ligand to be gp34, a surface glycoprotein expressed by vascular endothelial cells [196-197]. It is postulated that the binding of OX40 to gp34 facilitates the attachment of activated T-cells to the blood vessel wall and allows them to migrate into the interstitial space. Indeed, earlier reports showed that OX40-gp34 interaction mediates the adhesion of PHA-activated T-cells to the endothelial cells of human umbilical vein, aorta, and dermal microvasculature [191,198]. Phenotypic analysis of T-cells infected with the human T-cell lymphotropic virus Type-1 (HTLV-1) showed a strong surface expression of OX40 [191,198]. This virus causes adult T-cell leukaemia (ATL) in which leukaemic cells infiltrate the tissues in the intestinal tract, liver, lung, and spleen. The interaction between OX40 on these malignant cells and gp34 on endothelial cells might be pathogenic in ATL-mediated organ damage. OX40 expression is also detected in the synovial T-cells from R A patients and peripheral T-cells in allogeneic 29 G V H D [199]. This activation marker might also participate in human transplant rejection since rat anti-mouse OX40 MAbs prolong cardiac allograft survival in murine models [200]. 1.8.2 Interleukin-2 Receptor a-chain (CD25) The high-affinity heterotrimeric IL-2R is composed of a, p, and y subunits. The a-chain, designated as CD25, is co-expressed with the other two glycoprotein chains on the surface of activated T-cells and contributes to the stronger affinity of the entire receptor towards IL-2 [201]. Lymphocyte activation enhances the NF-KB-regulated expression of this growth factor receptor [201]. IL-2 signalling relies on the ability of the cytoplasmic domains of IL-2R(3- and y-chains to interact and activate various kinases including p56 l c k, p21 r a s and Janus kinase. The cascades initiated by these kinases promote the transcriptional activation of genes encoding for c-fos, c-jun, signal transducer and activator of transcription (STAT) and cyclins for cell cycle progression [88,89,201-204]. 1.9 Mitogenic Mechanism of Carbohydrate-binding Lectins 1.9.1 Phytohaemagglutinin (PHA) PHA is extracted from the red kidney bean Phaseolus vulgaris and is a member of the plant lectin family that exerts mitogenic activity in lymphocyte cultures [205]. The tetrameric protein is composed of two distinct subunits known as PHA-E for its erythrocyte-agglutinating ability and PHA-L for its lymphocyte-agglutinating and blastogenic capacity [206-207]. The lectin binds to the sugar moieties of cell surface proteins, thus causing the multivalent association and crosslinking of membrane glycoproteins that have signal-transmitting properties [208-209]. Lymphocyte proteins such as CD2, CD3, TCR, and 30 CD45 are implicated as cellular targets, thus suggesting that PHA stimulates T-cells by partly mimicking the antigen recognition process with the concomitant triggering of various activation signals [208-209]. Many studies have indicated that this mitogen induces activation events including the increase in [Ca ]j, c-jun expression, AP-1 binding, lymphokine gene expression, and cell cycle progression [80,210-212]. It requires accessory cells (e.g. APCs) to provide a stimulatory structural matrix and possibly cognate help in the form of surface proteins or cytokines to support its stimulation of T-cells [209,213]. The ability of PHA to induce CD69 expression in vitro is well documented [116,137,214-216]. 1.9.2 Concanavalin A (ConA) Similar mitogenic activity is exhibited by ConA, a plant lectin derived from the jackbean Canavalia ensiformis. This polyclonal activator also exists as a tetramer. It binds avidly with a-D-mannose, a type of monosaccharide that is attached to proteins during post-translational modification, and mediates cell activation by associating with and ligating these surface glycoproteins [206,208,217]. The action of ConA also relies on the intimate contact between T-cells and accessory cells, and this agent probably binds to B7-2 to deliver costimulatory signals through CD28 [218-219]. Putative T-cell receptors for this mitogen are CD7 and CD45, and its stimulatory effects include an increase of [Ca ]j, the activation of NF-AT, IL-2 synthesis, and cell blastogenesis [217-221]. 1.10 Relevance Because CD69 appears to be the earliest marker of lymphocyte activation, monitoring its expression might assist in the early diagnosis of the T-cell-mediated acute rejection of 31 kidney transplants. First, it would provide a simple and non-invasive means for immuno-logical monitoring. It might be used for the early detection of rejection, thereby permitting anti-rejection therapy to be initiated early and before the allograft sustains irreversible immunologic injury. Secondly, CD69 monitoring might be used to ascertain immune quiescence in stable patients so that the dose of immunosuppressants can be minimized. Thirdly, the assay might highlight periods of immunologic activation that require detailed investigation. Fourthly, it might also discriminate between immunological (e.g. rejection) and non-immunological (acute tubular necrosis and cyclosporine nephrotoxicity) causes of early graft dysfunction. Lastly, its utility might extend beyond renal transplantation and become a post-transplant monitoring tool in the field of solid organ transplantation. CD69 expression might also be useful in identifying increased or suboptimal T-cell activity in ESRD and RTx. This information might assist in manipulating the immune system in these patients to counteract abnormal activation or restore their immune competence. Understanding the expression kinetics of CD69 and other activation markers would provide significant insights into their functions in activated T-cells. Furthermore, their kinetics would shed light on their possible roles in the different stages of T-cell activation in immune diseases such as graft rejection (Fig. 1.2), and inhibiting their expression might become a therapeutic strategy. 1.11 Objective This study is based on five objectives. The first objective is to examine the optimal conditions for measuring CD69 expression on T-cell surface. The goal was to use the 32 techniques of triple-colour flow cytometry to analyze the kinetics of CD69 expression following in vitro mitogen stimulation. Cytofluorometric methods rely on fluorescent-tagged antigen-specific MAbs to identify cells bearing distinctive markers and examine the phenotypes of specific T-cell subsets. Polyclonal activators such as PHA, ConA, and OKT3 were used to induce CD69 expression in P B M C cultures. The second objective is to compare the expression kinetics of CD69 with those of OX40 and IL-2Ra. D N A synthesis acts as a positive indicator of cell proliferation and was measured using tritiated [3H]-thymidine ([ HJThd) incorporation assays. The third objective is to determine the influence of ISDs, including CsA, MP, and tacrolimus, on PHA-mediated CD69 induction in vitro to provide further insights into their inhibitory actions on the early phases of T-cell activation. The fourth objective is to use whole blood phenotyping to determine the baseline and PHA-induced CD69 and OX40 expression in the circulating T-cells of normal healthy individuals, chronic renal failure patients on HD or CAPD therapy, and long-term kidney transplant recipients. Whole blood cultures resemble closely to the in vivo micro-environments since they preserve, during culturing, the serum components, uraemic toxins, immuno-suppressants and other substances peculiar to the peripheral blood of the patients studied. The fifth objective is to monitor the CD69 expression of peripheral blood T-cells longitudinally following renal transplantation. The correlation between the occurrence of renal graft rejection and CD69 upregulation was evaluated. 33 Chapter Two Materials and Methods 2.1 Study Subjects A l l study participants provided written informed consent and the Clinical Research Ethics Board (Office of Research Services and Administration) at the University of British Colum-bia approved the project. Selection criteria were established for control and patient groups to exclude subjects with clinically diagnosed histories of immune deficiency, hyper-sensitivity disorders (e.g. atopy and allergy) and lymphoproliferative diseases and those who have received systemic anti-inflammatory, immunosuppressive (except for transplant patients) or immunostimulatory therapy in the preceding three months. Individuals who were less than 18 or more than 80 years of age were not included. Patients in the HD cohort (n = 10) underwent dialysis therapy 3 times a week (wk) and approximately 4 hours (hrs) per session at Vancouver General Hospital (VGH) Dialysis Unit. Treatment relied on single-use hollow-fibre dialyzers without specific membrane types. The other ten ESRD subjects depended on home CAPD and attended the Peritoneal Dialysis Outpatient Clinic. None of the dialysis subjects had kidney or other forms of transplantation previous to this study. The stable renal transplant (ST) patients (n = 10) participating in this study were on maintenance immunosuppression consisting of CsA, prednisone, Aza, and M M F in dual (n = 1) or triple (n = 9) combinations (Table 2.1). These subjects had been transplanted for more than one year (average 4.8 ± 1.9 yrs, ranging from 1 to 21 years) and were maintaining consistent graft functions (e.g. stable serum creatinine levels) without any signs of acute or 34 Table 2.1. ISD protocol in stable and new renal transplant recipients Patient cohorts Regimen ST (n= 10) NT (n= 10) CsA / Pred / Aza 8 2 CsA / Aza 1 -CsA / Pred / M M F 1 3 FK506 / Pred / Aza - 2 CsA / Pred / Sirolimus - 3 Dose range: CsA a 75 -125 mgb.i.d. 200- 1000 mgb.i.d. Preda 2.5 - 10 mgo.d. 10-90 mgo.d. Aza 100- 150 mgo.d. 100- 150 mgo.d. FK506 - 6-18 mg b.i.d. M M F 1 gb.i.d. 1 -2 gb.i.d. Sirolimus - 2 ml o.d. a) Table 3.12 summarizes the dosage levels of CsA and prednisone administered to NT patients during the monitoring period and also the whole blood trough concentrations of CsA in these subjects. Abbreviations: Aza, azathioprine; b.i.d., two times a day; CsA, cyclosporin A ; FK506, tacrolimus; ISD, immunosuppressive drug; M M F , mycophenolate mofetil; NT, new (de novo) renal transplant recipient; o.d., everyday; Pred, prednisone; ST, stable, long-term renal transplant recipient. 35 chronic rejection. Seven of these subjects received their transplants from cadaveric donors, and three from living relatives. Ten new renal allograft recipients (NT) were monitored longitudinally for three months after RTx. They were consecutively transplanted at V G H from June to September 1997 with either cadaveric (n = 8) or living (related: n = 1, parental: n = 1) donor kidneys and were immunosuppressed with triple therapy consisting of CsA, prednisone, Aza, FK506, M M F and sirolimus in various combinations (Table 2.1). Simultaneous pancreas-kidney trans-plantation was performed in two patients with IDDM to return them to normal glucose homeostasis and renal function. The primary causes of chronic renal failure in dialysis and transplant patients were listed in Table 2.2. Ten healthy volunteers who were matched for age and gender ratio with all other groups and who met the selection criteria were included as normal controls (Table 2.2). 2.2 Sample Collection Peripheral venous blood (-14 ml) was drawn via venipuncture into heparin-containing tubes (Becton Dickinson {BD} Vacutainer Systems, Franklin Lakes, NJ). Samples were collected between 8:00am to 11:00am to avoid the circadian variation of lymphocyte quantity and mitogenic response [222-224] and were processed within 24 hours of collection. Specimens were obtained predialysis for HD patients and during clinical visits for other patient subjects. NT subjects were monitored before their transplantation (pre-Tx) and on Days 2-4, 5-7, then every week in the first post-transplant month and every other week in the next two months during their follow-up transplant clinic visits until the end of the monitoring program. 36 Table 2.2. Primary causes of ESRD among patient subjects, with their demographic characteristics compared to those of normal controls Patient cohorts Demographics 'Normal HD CAPD ST NT Number of Subjects 10 10 10 10 10 Age (yrs)a'b 45 ± 4 58 ± 5 53 ± 4 44 ± 3 41 ± 3 Male:Female ratioc 5:5 7:3 5:5 6:4 6:4 Primary Aetiology of ESRD Glomerulonephritis (n = 6) 2 1 2 1 IDDM (n = 6) 1 - 1 4 Polycystic kidney (n = 5) 1 2 2 -Hypertension (n = 4) 2 2 - -Other diagnoses'1 (n = 8) 3 - 3 2 Unknown (n = 11) 1 5 2 3 a) each value is expressed as the mean ± SEM of each group b) no significant difference between normal and each patient group (unpaired Mest) c) no significant difference between study groups (Chi-square analysis) d) Other diagnoses include familial nephritis (n = 2), IgA (immunoglobulin A-isotype) nephropathy (n = 2), reflux nephropathy (n = 2), haemolytic uraemic syndrome (n = 1), and scleroderma (n = 1) Abbreviations: CAPD, continuous ambulatory peritoneal dialysis; ESRD, end-stage renal disease; HD, haemodialysis; IDDM, insulin-dependent diabetes mellitus; NT, new (de novo) renal transplant recipient; ST, stable, long-term renal transplant recipient; yrs, years. 37 2.3 Reagents Dulbecco's Ca2+-free, phosphate-buffered saline (PBS) powder from Gibco B R L (Grand Island, NY) was reconstituted with deionized water and sterilized by autoclaving. RPMI 1640 (Roswell Park Memorial Institute 1640), Dulbecco's 10X concentrated PBS solution, Trypan Blue dye, and foetal calf serum (FCS) were also obtained from Gibco. Medium additives including L-glutamine, penicillin, and streptomycin were from ICN (Aurora, Ohio), while human A B serum was from RJO Biochemicals (Kansas City, Missouri). Ficoll-Paque was purchased from Pharmacia (Uppsala, Sweden) and the fixative paraformaldehyde was from J .B .EM (Pointe Claire-Dorval, Quebec). Glacial acetic acid and ethanol were obtained from Canlab (Mississauga, Ontario) and Commercial Alcohol (Brampton, Ontario), respectively. [3H]Thd was purchased from New England Nuclear (Boston, M A ) and scintillation fluid was from Fisher Scientific (Fair Lawn, NJ ) . 2.4 Cell Culturing and Activation Experiments 2.4.1 Peripheral Blood Mononuclear Cell (PBMC) Isolation Peripheral blood mononuclear cells (PBMC) were isolated from heparinized blood by den-sity gradient centrifugation (Fig. 2.1) [225-226]. Blood was first diluted with an equal vol-ume of PBS, layered over 15 ml of Ficoll-Paque solution, and centrifuged at 1500 revolutions per minute (rpm) for 30 minutes. The mononucleocytes in the buffy coat interface were then extracted, washed twice with PBS and once in complete culture medium consisting of RPMI 1640 supplemented with, at their final concentrations, 4mM L-glutamine, 100 IU/ml penicillin, 100 pg/ml streptomycin, and 1% heat-inactivated human A B serum [214]. Differential cell counts were performed using Coulter Counter T660 38 Figure 2.1. Culturing and phenotyping techniques for preparing whole blood lymphocytes and PBMCs for flow cytometric analysis. Baseline levels were assessed without culturing. Centrifugation conditions are italicized. Abbreviations: hrs, hours; M A b , monoclonal antibody; min, minute; PAF, PBS + azide + foetal calf serum; P B M C , Peripheral blood mononuclear cell; PBS, phosphate-buffered saline; PHA, phytohaemagglutinin; rpm, revolution per minute; RPMI, Roswell Park Memorial Institute; v/v, volume-to-volume. 39 W H O L E B L O O D M E T H O D Heparinized whole blood (in autologous serum) + RPMI complete medium (blood:medium =10:1, v/v) I + 1.0 ug/ml PHA incubate at 37 °C for 24-72 hrs M A b labelling at 4 °C for 30 min + 2 ml erythrolytic solution for 20 seconds + 0.3 ml neutralizing solution 1500 rpm ^ 5 min wash with PAF solution i J O O r p w ^ 10 min 1 % paraformaldehyde j flow cytometric analysis P B M C M E T H O D Heparinized whole blood I + PBS (blood:PBS = 1:1, v/v) I Ficoll-Paque density isolation of P B M C 1500 rpm X 30 min 2 x PBS wash 1 x RPMI wash 1500 rpm ^ 10 min + 1.0 ug/ml PHA + 15% pooled serum I incubate at 37 °C for 4-120 hrs I 1 x RPMI wash 1500 rpm ^ 10 min M A b labelling at 4 °C for 30 min I wash with PAF solution 1500 rpm ^ 10 min 1 % paraformaldehyde i flow cytometric analysis 40 (Coulter, Hialeah, FL) and cell viability was assessed under a light microscope by means of Trypan Blue dye exclusion [227]. 2.4.2 P B M C Culturing Conditions Mononuclear cells were suspended in complete culture medium at 1.0 - 2.0 x 106 cells/ml and were placed in either 24-well flat-bottom plates or 14 ml polypropylene U-bottom tubes (both were BD Falcon products, Lincoln Park, NJ). Unstimulated or stimulated cells were cultured at 37°C in a humidified atmosphere of 95% air and 5% C 0 2 until being harvested. 2.4.3 Mitogenic Stimulation Lyophilized PHA mucoprotein (Gibco) derived from P. vulgaris was reconstituted with PBS, diluted with complete medium, and added to the P B M C cultures at a final concentration of 1.0 pg/ml. ConA powder (US Biochemicals, Cleveland, Ohio) was dissolved in medium and used at a final concentration of 0.2 ug/ml. OKT3 was immobilized on the surface of 24-well flat-bottom plates by incubating the antibody at 30 ng/ml in medium overnight at 4°C. The supernatant was subsequently removed and cells were added. At various times (after 0, 4, 12, 24, 48, 72, 96, or 120 hrs of incubation), cells were washed once with complete medium and were ready for antibody labelling. 2.4.4 Co-cultivation with Immunosuppressive Drugs (ISDs) The effects of immunosuppressants were studied by incubating them with PHA-stimulated PBMCs from the onset of culturing. CsA powder and MP solution were generous gifts from Dr. Nilufar Partovi (Pharmacy, VGH), and Dr. Paul Keown provided the FK506 crystalline powder. CsA and FK506 were each dissolved in a vehicle made up of 7 volumes of ethanol 41 to 3 volumes of deionized water. Further dilution with complete medium yielded stock solutions that were subsequently applied to cultures at final concentrations ranging from 0, 100, 200, 400 to 800 ng/ml for CsA [96] and from 0, 0.1, 2.5, 10.0 to 20.0 ng/ml for tacrolimus [228]. MP was mixed with complete medium and added to cultures at final doses of 0.0, 0.01, 0.1, 1.0, or 10 ug/ml [108]. The quantity of CsA in stock solutions and some transplant patient samples were determined using TDx fluorescence polarization immuno-assay (Abbott Laboratory, Abbott Park, IL.). Percent inhibition was calculated using the formula: %Inhibition = (untreated - treated) x 100% / untreated. 2.4.5 Immunostaining Medium-washed cells were incubated with specific and control antibodies for 30 minutes at 4°C in the dark. Following a single wash with cold PAF solution, which contained lx PBS, 0.1% sodium azide, and 1% heat-inactivated FCS, to remove unbound antibodies, the cells were fixed with 1% paraformaldehyde in PBS. 2.4.6 Whole Blood Culturing and Processing Technique Whole blood phenotyping was used to evaluate T-cell activation in study subjects (Fig. 2.1). Heparinized blood obtained from participants was mixed with complete medium in a 10:1 ratio under sterile conditions and cultured with 1.0 ug/ml PHA for 0, 24, 48, or 72 hrs at 37°C in 5% C O 2 humid air. After the appropriate incubation time, 100 ul aliquots were sub-jected to the "stain-lyse" procedure [229] in which cells were first stained with a pre-mixed panel of antibody combinations for 30 minutes at 4°C in the dark. Contaminating red blood cells (RBC) were removed by incubating these stained samples with 2.0 ml of acidic (pH 42 3.34 - 3.43), hypotonic (average 46 ± 1 mOsm) erythrolytic reagent for 20 seconds at room temperature. This lysing solution consisted of lx PBS diluted with distilled water (1:9) and 0.1% glacial acetic acid. Erythrocytes underwent hypotonic lysis, while lymphocytes were protected by their more robust cytoplasmic membrane. Immediately after erythrolysis, 0.3 ml of Dulbecco's 10X concentrated PBS solution (pH 6.79 - 6.80, hypertonic, average 2850 ± 10 mOsm) was added to restore the tonicity and pH of the cell suspensions. The cells were then pelleted, washed once with PAF and were fixed with 1.0% paraformaldehyde. 2.5 Fluorochrome-conjugated Monoclonal Antibodies (MAbs) Murine anti-human MAbs conjugated with either fluorescein isothiocyanate (FITC), R-phycoerythrin (PE) or peridinin chlorophyll protein (PerCP) were employed in dual or triple combinations (Table 2.3) for the direct detection of T-cell membrane proteins. The anti-bodies (colour, name of antibody, and antigen in parenthesis) were specific for CD3 (PerCP, Leu-4, pan T-cell marker), CD4 (FITC, Leu-3a, Th co-receptor), CD8 (FITC, Leu-2a, CTL co-receptor), CD69 (PE, Leu-23, activation inducer molecule), CD 134 (PE, anti-Hu-OX40, human OX40 molecule), and CD25 (PE or FITC, anti-IL2R, human IL-2R oc-chain). A l l MAbs were isotypically IgGi and were purchased from BD Immunocytometry Systems (San Jose, CA). Two tri-colour antibody cocktails produced by BD (Fastimmune System), the CD69PE+CD3PerCP+CD4FITC and CD69PE+CD3PerCP+CD8FITC combinations, were included for the differential analysis of CD69 expression in the helper and cytotoxic T-cell subsets, respectively. Background fluorescence due to non-specific binding and cell auto-fluorescence was determined using PE- or FITC-labelled murine isotype-matched (IgGi) control MAbs with irrelevant specificities. Monocyte contamination in the lymphocyte gates 43 Table 2.3. Dual and triple combinations of monoclonal antibodies used in cytofluorometric phenotyping M A b combination Target cell population IgGi-•PE + CD4-FITC + CD3-PerCP background fluorescence of helper T-cells IgGi -PE + CD8-FITC + CD3-PerCP background fluorescence of cytotoxic T-cells IgG 1-PE + CD3 -PerCP background fluorescence of T-cells CD69 -PE + CD4-FITC + CD3-PerCP CD69-expressing helper T-cells CD69 -PE + CD8-FITC + CD3-PerCP CD69-expressing cytotoxic T-cells OX40 -PE + CD4-FITC + CD3-PerCP OX40-expressing helper T-cells OX40 -PE + CD8-FITC + CD3-PerCP OX40-expressing cytotoxic T-cells CD25 -PE + CD4-FITC + CD3-PerCP IL-2Ra-expressing helper T-cells CD25 -PE + CD8-FITC + CD3-PerCP IL-2Ra-expressing cytotoxic T-cells CD25-PE + CD3-PerCP IL-2Ra-expressing T-cells CD14-PE + CD45-FITC + CD3-PerCP T-lymphocytes (CD14"CD45+) Abbreviations: FITC, fluorescein isothiocyanate; IgG, immunoglobulin G-isotype; IL-2Roc, interleukin-2 receptor a-chain; MAb, monoclonal antibody; PE, R-phycoerythrin; PerCP, peridinin chlorophyll protein. 44 during cytofluorometric analysis was assessed in a few samples using the CD14PE + CD45FITC panel [230]. The MAbs repertoire was titrated for optimal concentrations. 2.6 Flow Cytometric Analysis Stained PBMCs and whole-blood-lysed cells were analysed with an EPICS Profile I flow cytometer (Coulter) equipped with a 15 mW argon laser that generates a light beam at a wavelength of 488 nm for fluorochrome excitation. The light scattering characteristics of cells were registered linearly as forward scatter (FSC) for cell size and orthogonal (90°) side scatter (SSC) for granularity. After light absorption, each fluorescent tag generates a particular emission spectrum with maxima at 530 nm (green) for FITC, 575 nm (red) for PE, and 677 nm (purple) for PerCP. The signals from the excited FITC, PE, and PerCP were detected by fluorescence one (FL1) photomultiplier tube (PMT) through a 525/3Onm-bandpass (BP) filter, FL2 PMT through a 575/20nm-BP filter, and FL3 PMT through a 635/35nm-BP filter, respectively. Colour compensation adjustments were made to minimize the influence of overlapping spectra. The surface phenotype of T-cell subsets was analysed using the double-gating technique [214]. The first bitmap, the "lymphocyte gate," was set on the lymphocyte and lymphoblast populations based on their FSC vs. SSC properties [230], and the second bitmap, the "T-cell gate," was positioned on the CD3+T-cells according to their logarithmic FL3 (LFL3, CD3PerCP) vs. SSC profiles [189]. Cells that were simultaneously present in both bitmaps were analysed for their co-expression of activation markers and subset-differentiating co-receptors based on their logarithmic FL1 (LFL1, PE) and FL2 (LFL2, FITC) properties, 45 respectively, in a dual parameter cytogram. Quadrants were set to include < 1% of the background fluorescence based on cells incubated with control MAbs. A single parameter histogram was used to enumerate the CD3+T-cells that expressed the activation proteins. Figure 2.2 shows how the double-gating technique was used to determine surface marker expression. Results were reported as proportional expression, which was based on the percentages of cells expressing certain markers, and as mean fluorescence intensity (MFI), which was based on the fluorescence emitted by the cell-bound MAbs and estimates cell surface antigen density [231]. To calculate proportional expression, the percentage of cells specifically stained was recorded after subtracting the background signal from the percentage of positive cells. The logarithmic MFI was converted to linear fluorescent intensity using a 256-channel linear scale. The data of all five parameters (FSC, SSC, and LFL1 to LFL3) from 10,000 events were acquired for each sample and stored in listmode files. Further analysis was performed using the EPICS Elite Workstation software 4.01 (Coulter). Routine quality control procedures were performed before sample analysis to determine the optimal settings of the flow cytometer. First, the optical alignment of the argon laser was optimized using DNA-Check fluorospheres (Coulter), which are uniform-sized polystyrene beads tagged with a specific fluorescent dye. Then, the fluorescence intensity of each LFL parameter was standardized with Flow-Set fluorospheres (Coulter), which are also fluorescent beads with uniform size but were used to optimize the PMT voltages. 2.7 Cell Proliferation Assay The mitogen-driven D N A synthesis was quantified based on cellular [3H]Thd uptake [214]. Triplicate PBMC cultures in 96-well U-bottom microtitre plates (Nunc, Roskilde, Denmark) 46 A O T 1 1 1 1 1 1~~ 0 10 20 30 40 50 60 SSC B o -o • O " 0 10 20 30 40 50 60 SSC Figure 2.2. Double-gating strategy for analyzing activation marker expression. The cells examined in these cytograms were first activated with PHA for 24 hours in whole blood cultures and then stained with the antibody cocktail CD69+CD4+CD3 before undergoing erythrolysis and fixation. The two contour plots (A and B) in the upper panel illustrate how the lymphocyte and T-cell gates were positioned. Only cells that were simultaneously present in both bitmaps were analysed. Histograms (C and D) in the lower panel demonstrate how CD69 expression in the CD4 subset and CD3+T-cells was determined. Quadrants were set according to the background fluorescence of cells incubated with control antibodies. Abbreviations: FITC, fluorescein isothiocyanate; FSC, forward scatter; LFL, logarithmic fluorescence; PE, R-phycoerythrin; PerCP, peridinin chlorophyll protein; SSC, side scatter. 47 were unstimulated or activated for 72 hours at 37°C in 5% CO2 humid air. Afterwards, they were pulsed with 1 \xCi (37 kBq) of [ H]Thd for an additional 20 hours before being harvested onto glass fibre filters (Skatron, Sterling, V A ) with a cell harvester (Skatron). Radioactivity incorporation was measured in counts per minute (cpm) in scintillation fluid with a beta counter (LKB Wallac, Turku, Finland). The stimulation index (SI) was used to avoid interpersonal variability in mitogen response, and it represented the ratio of the mean cpm of the triplicate stimulated cultures to that of the triplicate unstimulated cultures. 2.8 Statistics A l l data were expressed as the mean ± standard error of the mean (SEM). The statistical significance of the differences between variable means was determined using unpaired or paired Student's Mest. The correlation between parameters was expressed as correlation coefficients (r) and was assessed with linear regression based on the method of least squares. Binomial distributions (e.g. male-to-female ratio) was evaluated with the Chi-square test. Differences of p < 0.05 were considered statistically significant. 48 Chapter Three Results 3.1 Kinetics Studies The mononuclear leukocyte fractions extracted from the peripheral blood of normal subjects contained 94.4 ± 1.2% viable cells as indicated by their exclusion of the Trypan blue dye. Flow cytometric analysis revealed that 89.5 ± 1.9% of the PBMCs located within the FSC vs. SSC bitmap were CD3-positive lymphocytes. The double-gating evaluation of several samples using the anti-CD 14/45 MAb pair showed that the CD3+lymphocyte populations consisted primarily of T-cells (%CD45+CD14" = 98.9 ± 0.6%) with minimal monocyte contamination (%CD45 +CD14 + = 0.08 ± 0.04%). The average CD4/CD8 ratio in normal PBMCs was 2.40 ± 0.49, with 60.3 ± 4.7% of CD3+T-cells identified as CD4-positive and 35.0 ± 3.1% as CD8-positive T-cells. The cell viability in PHA-stimulated cultures dropped non-significantly throughout the incubation period (24 hrs = 90.5 ± 1.7%, 72 hrs = 89.6 ± 1.9%, and 120 hrs = 78.6 ± 10.0%). The fraction of CD3+T-cells in the lymphocyte population remained constant after activation (24 hrs = 83.9 ± 2.4%, 72 hrs = 80.6 ± 6.0%, and 120 hrs = 84.6 ± 6.8%). The average stimulation index of these activated PBMCs after 92 hours of incubation was 352.1 ± 50.2. 3.1.1 Expression Kinetics of Activation Markers in PHA-stimulated T-cells CD69: Early Stage Molecule A negligible proportion of freshly isolated CD3+T-cells from healthy persons expressed CD69 (2.9 ± 1.1%; Table 3.1 and Fig. 3.1). This value was similar (p = statistically non-significant or NS) to the mean proportion of cells bound by isotype-matched control 49 Table 3.1. CD69 expression kinetics in non-stimulated and PHA-activated T-cellsa A) CD3+T-cells % positive cells MFI Time (hrs) Non-stimulated PHA Non-stimulated PHA 0 6.3 + 5.1 2.9+1.1 77 + 7 78 ± 2 4 7.3 + 2.6 25 .7±7.1 t 76 ± 7 83 ± 2 12 ND 42.0 ± 8.0t ND 103 ± 8 t 24 1.9 ± 1.1* 75.0±4.5 t 80 ± 6 * 128+ 6t 48 1.2 ±0 .5* 58.3 ±3.It 87 ± 3 * 109 + 7 72 2.2 ± 1.2* 43.2 ±7.U 86 ± 3 * 115 ± 9 96 1.1 ±0.7* 28.1 ±4.9t 86 ± 4 91 ±3A 120 1.2 ±0.6* 29.2 ±5.4t 82 ± 2 9 2 ± 4 § a) Cells were cultured in the absence (non-stimulated) or presence of 1 ug/ml PHA. Abbreviations: MFI, mean fluorescence intensity; ND, not determined; PHA, phytohaemagglutinin. * p < 0.025 vs. PHA after the same period of culture ** p < 0.05 vs. PHA after the same period of culture t p < 0.025 vs. its 0-hr level %p< 0.05 vs. its 24-hr level Ap < 0.025 vs. its 24-hr level §£> < 0.05 vs. its 72-hr level 50 Table 3.1. CD69 expression kinetics in non-stimulated and PHA-activated T-cells B) CD4 subset % positive cells MFI Time (hrs) Non-stimulated PHA Non-stimulated PHA 0 1.9 ± 1.5 1.0 ±0.5 97 ± 10 91 ± 3 4 2.3 ± 1.1* 36.0 ± 6.8t 87 ± 8 89 ± 4 12 N D 59.8±2.9t ND 108 ± 12 24 0.5 ±0.4* 84.9±2.6t 86 ± 8 * * 131 ± 7 t 48 0.9 ±0 .5* 54.7 ± 7.1$ 88 ± 10 92 ±7$ 72 0.8 ±0.4* 37.3 + 5.7$ 78 ± 3** 97 ±2$ 96 0.4 ±0.2* 29.2 + 1.7 86 ± 4 86 ± 5 120 1.4 ±0.8* 26.9±6.1§ 81 ± 3 88 ± 4 51 Table 3.1. CD69 expression kinetics in non-stimulated and PHA-activated T-cells C) CD8 subset . ^ = = = = = ^ = % positive cells MFI Time (hrs) Non-stimulated PHA Non-stimulated PHA 0 3.2 ± 1.9 1.5 ±0.4 88 ± 3 85 ± 3 4 4.5 + 1.9 12.9 ±5.8 86 ± 2 77 ± 2 12 N D 28.3 ±6.1 t N D 94 ± 7 24 1.9 ±0 .5* 63.1 ±6 .0 t 89 ± 8* 122 ± 6 t 48 1.3 ±0.7* 47 .0±2.8 t 82 ± 2 * 106 ± 6 72 3.0 ±2.2* 31.4 ±12.9t 85 ± 6 * 116 + 9 96 1.4 ±0 .5* 13.4 ±6.6 91 ± 7 9 0 ± 4 A 120 0.8 ±0.5* 7.8 ±0.5 89 ± 3 88 ± 2§ 52 Figure 3.1. The kinetics of CD69 expression in T-cells. Each value represents the mean ± SEM of 3 independent experiments with 3 healthy controls. A) CD3+T-cells. B) CD4 subset. C) CD8 subset. (*p < 0.025 vs. PHA after the same period of culture, tp < 0.025 vs. its 0-hr level, tp < 0.05 vs. its 24-hr level and **p < 0.05 vs. its 72-hr level) 53 o I H + Q U + ON >o Q V 0 s-B) CD4 subset 100.0 -r 90.0 + 12 24 36 48 60 72 84 Time (hrs) - • - PHA -o— Non-stimulated 96 108 120 C) CD8 subset 80.0 T 70.0 + 12 24 36 48 60 72 84 Time (hrs) - • - PHA - o - Non-stimulated 96 108 120 54 antibodies (0.2 ± 0.0%). The estimated CD69 epitope density, expressed in terms of the 256-channel-converted MFI values of CD69CD3 T-cells (78 ± 2), was also equivalent (p = NS) to that of the isotypic control (72 ± 3). The CD4 + and CD8+T-cells in general were C D 6 9 i o w ( C D 4 + : 1 < 0 ± 0.5% vs. 0.6 ± 0.6% with isotypic control, p = NS; CD8 + : 1.5 ± 0.4% vs. 0.5 ± 0.5% with isotypic control, p = NS) and weakly stained with the anti-CD69 M A b (MFI for CD4+:91 ± 3 vs. 75 ± 8 with isotypic control, p = NS; CD8 + : 85 ± 3 vs. 94 ± 22 with isotypic control, p = NS). The percentage values were normalized to the proportion of CD4 + or CD8 + subclass in the CD3 +T-cell population. CD69 expression was significantly upregulated (p < 0.05 vs. 0 hrs) in the CD3+T-cells and CD4 subset after 4 hrs of stimulation with 1.0 ug/ml PHA. The proportions of positive cells increased to 25.7 ± 7.1% and 36.0 ± 6.8%, respectively, although the changes in their MFI values (CD3 +: MFI = 83 ± 2; CD4 + : MFI = 89 ± 4) remained statistically non-significant. CD69 expression was also enhanced in CD8+T-cells (12.9 ± 5.8%, MFI = 77 ± 2) but did not reach statistical significance. With continuing incubation, more cells exhibited the CD69 + phenotype and the staining intensity increased, and the percentage and MFI values were all significantly increased after 12 hrs of activation. The proportions of cells expressing CD69 reached maximal levels after 24 hrs, when 75.0 ± 4.5% of CD3+T-cells (MFI = 128+6), 84.9 + 2.6% of CD4+T-cells (MFI = 131+7) and 63.1 + 6.0% of CD8+T-cells (MFI = 122+6) displayed this marker. These data were substantially higher than those of the resting and 4-hr-stimulated T-cells. After this point, the proportions of cells expressing CD69 declined rapidly and significantly 55 (p < 0.05 for 48 hrs vs. 24 hrs) with a concomitant reduction in CD69 surface density (p = NS). The reduction continued so that by 72 hrs the expression of CD69 on CD3 + (43.2 ± 7.1%, MFI =115 + 9), CD4 + (37.3 ± 5.7%, MFI = 97 ± 2) and CD8+T-cells (31.4 ± 12.9%, MFI = 116 + 9) was similar to those observed in the 4-hr and 12-hr cultures. By 120 hrs of activation, the cells appeared to maintain a low but constant level of CD69 expression, with 29.2 ± 5.4% of C D 3 + (MFI = 92 + 4), 26.9 ± 6.1% of CD4 + (MFI = 88 + 4) and 7.8 ± 0.5% of CD8+T-cells (MFI = 88 + 2) displaying this molecule on their surface. The phenotypic changes reported above were not observed in non-stimulated T-cells as indicated by their consistently low CD69 levels (% positive cells and staining) during the 5-day incubation period (Table 3.1 and Fig. 3.1). OX40: Intermediate Stage Molecule The PBMC approach was also used to analyse OX40 expression in normal subjects (Table 3.2 and Fig. 3.2). As observed with CD69 expression (Table 3.1), baseline OX40 levels (0 hrs) were very low, with 0.4 ± 0.2% of CD3 + , 0.0 ± 0.0% of CD4 + and 0.3 ± 0.2% of CD8+T-cells displaying this marker. OX40 expression remained undetectable in T-cells after 4 hrs of PHA activation (p = NS vs. 0 hrs) and was markedly lower (p < 0.05) than the CD69 response at this time. After 12 hrs of stimulation, however, the proportions of OX40+T-cells increased substantially to levels similar to those that were expressing CD69 and continued to rise after 24 hours thus reaching 46.3 ± 7.3% in C D 3 + and 66.1 ± 4.8% in CD4+T-cells. In contrast to the CD4 subclass, only a small portion of CD8+T-cells was stained positive for OX40 at this time (16.3 ± 7.0%; p <0.05 vs. 0 hrs), which was markedly 56 Table 3.2. OX40 expression kinetics in PHA-activated T-cells: C D 3 + CD4 + CD8 + Time (hrs) % MFI % MFI % MFI 0 0 .4±0.2 80 ± 4 0.0 ±0.0 80 ± 7 0.3 ±0.2 108 ± 21 4 o .o±o.ot 75 ± 5 0.1 ±0.1 t 91 ± 14 o .o±o.ot not-stainedb 12 28.0 ±10.7* 124 + 5* 44.6+13.4* 128 + 7* 8.7 ±2.5 106+10 24 46.3 ±7.3*t 131 ± 7 * 66.1 ±4.8*t 133 ± 6 * 16.3±7.0*t 131 ± 2 48 43.5 ±11.9 122 ± 7 67.4 ±10.9 128 ± 1 1 7.6 ± 5.2t 122 ± 6 72 38.8 ±14.6 105 ± 9 58.8 +7.0t 107+12 4 .2±0 .6 t 86 ± 3 96 34.6 ±7.6 104 ±4**t 50.1 ±4. It 107 ± 7 * * 2.9 ±5.1 90 ± 20 120 33.3 ±6.2 110±l**t 49.2 ±1.6f 113 ±l**t 8.3 ±5.2 121 ±0**t a) PBMC suspensions were stimulated with 1.0 pg/ml PHA. b) Cells were not stained due to the lack of anti-OX40 MAb binding Abbreviations: MFI, mean fluorescence intensity; PHA, phytohaemagglutinin. * p < 0.05 vs. its 0-hr level * * p< 0.05 vs. its 24-hr level t p < 0.05 vs. CD69 expression (Table 3.1) after the same period of culture %p< 0.025 vs. CD69 expression (Table 3.1) after the same period of culture 57 Figure 3.2. The kinetics of CD69 and OX40 expression in T-cells. Each value represents the mean ± S E M of 3 independent experiments with 3 healthy controls. A) CD3+T-cells. B) CD4 subset. C) CD8 subset. (*p < 0.05 vs. its 0-hr level and \p < 0.05 vs. OX40 after the same period of culture) 58 B) CD4 subset 100.0 T 12 24 36 48 60 72 Time (hrs) - • - C D 6 9 ^ - O X 4 0 84 96 108 120 C) CD8 subset 12 24 36 48 60 72 Time (hrs) - • - C D 6 9 - 6 - O X 4 0 84 96 108 120 59 lower than the proportion that was expressing CD69 (p < 0.05 in %, p = NS in MFI). The percentages of OX40 +CD8 +T-cells dropped further throughout the remainder of the incubation period and were significantly lower than those displaying CD69 at 48 and 72 hrs. The levels of these activation markers were comparable only after 96 hrs. Due to the lack of OX40 expression in CD8+T-cells, significantly fewer OX40-bearing CD3+T-cells were detected at 24 hrs compared to CD69 (p < 0.05), despite their equivalent surface densities. OX40 expression reached a plateau after 48 hrs in CD3 + and CD4+T-cells, at which point the proportions of cells expressing OX40 and CD69 were not markedly different. The OX40 + population reduced slightly at 72 hrs to 38.8 ± 14.6% for C D 3 + and 58.8 ± 7.0% for CD4 + T-cells, although this decline (p = NS between 24 and 72 hrs) was not as sharp as that for CD69. OX40 expression after 96 hrs was substantially greater than that of CD69, and 33.3 ± 6.2% of CD3 + and 49.2 ± 1.6% of CD4+T-cells were positively stained with the anti-OX40 MAbs after 120 hrs (p = NS vs. 24 hrs). CD25: Late Stage Molecule The PHA-induced IL-2Roc chain expression in normal controls was studied in CD3+T-cells over 5 days and in the CD4 + and CD8 + subpopulations at 0, 24, and 72 hrs (Table 3.3 and Fig. 3.3). CD69 expression was similar to previous results. CD25 was displayed by small fractions of circulating CD3 + (15.9 ± 1.1%, p < 0.001 vs. baseline CD69 level), CD4 + (15.7 ± 5.2%) and CD8+T-cells (0.7 ± 0.1%). Its upregulation lagged behind CD69 induction in CD3+T-cells after 4 hrs of PHA stimulation (5.8 ± 1.6% vs. 28.0 ± 8.6%, p < 0.05). More CD25+T-cells were detected by 24 hrs (CD3 +: 43.7 ± 7.3%, CD4 + : 73.1 ± 4.4% and CD8 + : 60 Table 3.3. CD25 expression kinetics in PHA-activated T-cellsa CD3 + CD4 CD8 Time (hrs) % MFI % MFI % MFI 0 15.9 ± 1.1* 83 ± 3 15.7 ±5.2 89 ± 2 0.7 ±0.1 96 ± 9 4 5.8 ±1.6**t 90 ± 5 N D N D N D N D 24 47.3±7.3***t 126 ± 5 73.1 ±4.4 t 138±12t 49.6 ± 5.4t 130±lOt 48 61.2 ±3.8 113 ± 6 N D ND ND ND 72 90.8±2.9***t 168±6*** 98.2±1.3***t 181±9***t 96.6±1.7***J 178±11***J 96 81.4+11.6*** 142+6*** ND ND N D ND 120 92.5±3.3*** 142+5*** ND ND ND ND a) PBMC suspensions were stimulated with 1.0 ug/ml PHA. Abbreviations: MFI, mean fluorescence intensity; ND, not determined; PHA, phyto-haemagglutinin. * p < 0.001 vs. CD69 baseline (0-hr) expression ** p < 0.05 vs. CD69 expression after the same period of culture ***£><0.025 vs. CD69 expression after the same period of culture \p< 0.05 vs. its 0-hr level tp< 0.05 vs. its 24-hr level 61 A) CD3+T-cells 100.0 x 0.0 H 1 1 1 1 1 1 1 1 1 1 0 12 24 36 48 60 72 84 96 108 120 Time (hrs) - • - C D 6 9 - A - C D 2 5 Figure 3.3. The kinetics of CD69 and CD25 expression in T-cells. Each value represents the mean ± S E M of 3 independent experiments with 3 healthy controls. A) CD3+T-cells. B) CD4 subset. C) CD8 subset. (*p < 0.001 vs. CD25 baseline (0-hr) level, **p < 0.05 vs. CD25 after the same period of culture, ***p < 0.025 vs. CD25 after the same period of culture, tp < 0.05 vs. its 0-hr level, and tp < 0.05 vs. its 24-hr level) 62 63 49.6 ± 5.4%) at which point the results in the CD4 and CD8 subsets were comparable to those of the CD69 expression. Although the CD69 expression and M A b staining intensity in CD3+T-cells declined sharply after 24 hrs, the CD25 levels of these cells continued to increase, with 90.8 ± 2.9% of CD3 + , 98.2 + 1.3% of CD4 + and 96.6 ± 1.7% of CD8+T-cells expressing the late marker by 72 hrs. These results were markedly higher (p < 0.025) than those of CD69 expression. By day 5 of culturing, almost all CD3+T-cells (92.5 ± 3.3%) maintained strong plateau levels of CD25 in contrast with their weak CD69 expression (p < 0.025) observed at this time. 3.1.2 Effects of Mitogen Dose on Activation Antigen Expression The effects of varying PHA concentrations on CD69 expression and D N A synthesis in healthy donor PBMCs were studied after 24 and 92 hrs of mitogen exposure, respectively (Table 3.4 and Fig. 3.4 to 3.7). The CD69 level induced by 0.2 ug/ml PHA was low (how-ever, p < 0.05 for both % and MFI when compared to non-stimulated cells), and it rose to a plateau at 1.0 ug/ml, at which concentration it was expressed in 60.4 ± 10.8% of CD3 , 81.0 + 6.2% of CD4 + and 52.0 ± 7.0% of CD8+T-cells. There was no significant increase in its expression at higher PHA concentrations (e.g. 3.0 ug/ml enhanced the proportions of CD69 + cells non-significantly to 69.2 ± 9.4% for CD3 + , 81.5 ± 6.6% for CD4 + and 61.2 ± 10.7% for CD8+T-cells). In fact, increasing the PHA concentration to 5.0 pg/ml led to a slight decrease in its expression. The background fluorescence also increased as higher mitogen doses were used. A PHA concentration of 1.0 ug/ml was therefore chosen for the activation assays. 64 Table 3.4. PHA dose-titration experiments based on 24-hr CD69 induction in T-cells1 PHA (p.g/ml) CD3 + CD4 + CD8 + % MFI % MFI % MFI 0 1.5 ±0.6 80 ± 3 0.4 ±0.2 80 ± 6 1.6 ±0.4 88 ± 7 0.2 27.7 ±5.2 94 ± 6 32.9 ±6.8 99 ± 7 16.9 ± 1.9 86 ± 8 1.0 60.4±10.8* 136 ±14* 81.0 ±6.2* 140 ±16* 52.0 ±7.0* 132 ±12* 3.0 69.2± 9.4* 130 ± 13 81.5 ±6.6* 131 ± 14 61.2±10.7* 129 ± 12 5.0 60.9 ±13.1 121 ± 9 72.4 ±11.8 124 ±11 45.5±12.6t 116 ± 12 a) PBMC suspensions were stimulated with PHA for 24 hours and then CD69 expression was measured. Abbreviations: MFI, mean fluorescence intensity; PHA, phytohaemagglutinin. * p < 0.025 vs. CD69 expression at 0.2 ug/ml tp < 0.05 vs. CD69 expression at 3.0 ug/ml 65 A) 100.0 -r 0.0 B) 100.0 - r 1.0 2.0 3.0 PHA concentration (ug/ml) 4.0 5.0 0.081x +25.69 r = +0.62 p =NS 100 200 300 Stimulation Index 400 500 600 Figure 3.4. PHA dose-response curve based on CD69 expression in CD3+T-cells. A) Percentage values. Each value represents the mean ± SEM of 3 independent experiments with 3 healthy controls. B) Correlation analysis of CD69 expression with P B M C proliferation between 0.2 ug/ml to 3.0 ng/ml of PHA. (*p < 0.025 vs. 0.2 ug/ml) 66 + Q a + ON Q U A) 100.0 -| 90.0 -80.0 -70.0 -60.0 -50.0 -40.0 - - - r 30.0 -20.0 -10.0 -0.0 -0.0 H 1 1 h-1.0 2.0 3.0 4.0 PHA concentration (ug/ml) 5.0 B) 100.0 x 0.11x + 28.63 r = +0.75 p < 0.025 100 200 300 400 Stimulation Index 500 600 Figure 3.5. PHA dose-response curve based on CD69 expression in CD4+T-cells. A) Percentage values. Each value represents the mean ± S E M of 3 independent experiments with 3 healthy controls. B) Correlation analysis of CD69 expression with P B M C proliferation between 0.2 pg/ml to 3.0 ug/ml of PHA. (*p < 0.025 vs. 0.2 pg/ml) 67 A) 80.0 T 0.0 B) 100.0 j 90.0 --80.0 --70.0 --H 60.0 --oo g 50.0 --S 40.0 --. Q ^ 30.0 --1.0 2.0 3.0 PHA concentration dug/ml) 4.0 5.0 0.086x+ 14.89 r = +0.66 p <0.05 100 200 300 400 Stimulation Index 500 600 Figure 3.6. PHA dose-response curve based on CD69 expression in CD8+T-cells. A) Percentage values. Each value represents the mean ± S E M of 3 independent experiments with 3 healthy controls. B) Correlation analysis of CD69 expression with P B M C proliferation between 0.2 ug/ml to 3.0 ug/ml of PHA. (*p < 0.025 vs. 0.2 ug/ml and tp < 0.05 vs. 3.0 ug/ml) 68 Figure 3.7. PHA dose-response curve based on D N A synthesis in PBMCs. Each value represents the ratio of the cpms between stimulated and non-stimulated cells and is expressed as the mean ± S E M of 3 independent experiments with 3 healthy controls. (*p < 0.05 vs. 0.2 ug/ml and tp < 0.025 vs. 1.0 ng/ml and 3.0 p-g/ml) 69 D N A synthesis increased with increasing PHA concentrations (SI of 1.0 ug/ml = 404.5 ± 44.5; 3.0 ug/ml = 476.9 + 42.9; 5.0 ug/ml = 669.3 ± 36.6, p < 0.025 vs. 1.0 and 3.0 ug/ml). CD69 expression and proliferation did not correlate significantly over the range of 0.2 to 3.0 Ug/ml PHA in CD3+T-cells (r = +0.62, p = NS), but they correlated positively in CD4 + (r = +0.75, p < 0.025) and CD8+T-cells (r = +0.66, p < 0.05). 3.1.3 Comparison of PHA, ConA, and OKT3 in Inducing CD69 Expression Cell activation with 0.2 ug/ml of ConA was compared to that with 1.0 ug/ml of PHA (Fig. 3.8). No significant difference was observed in the proportions of CD69 + cells after 4 and 24 hrs of incubation with PHA or ConA in the CD3 + (4 hrs: 25.7 ± 7.1% vs. 30.7 ± 4.3%, respectively; 24 hrs: 55.0 ± 7.4% vs. 50.9 ± 6.2%, respectively), CD4 + (4 hrs: 36.0 ± 6.8% vs. 35.3 ± 7.8%, respectively; 24 hrs: 73.3 + 3.3% vs. 57.4 ± 7.0%, respectively), or CD8 + T-cell populations (4 hrs: 12.9 ± 5.8% vs. 16.8 ± 8.1%, respectively; 24 hrs: 45.1 ± 8.9% vs. 38.3 ± 11.6%, respectively). The expression kinetics of CD69 was identical in both types of cultures, with a decline in its expression in CD3 + (43.7 ± 7.6% vs. 35.9 ± 8.6%), CD4 + (31.1 ± 4.8% vs. 27.2 ± 5.4%) and CD8+T-cells (14.2 ± 4.9 vs. 18.1 ± 8.1) at 72 hrs. D N A synthe-sis triggered by the two mitogens was comparable (SI: PHA = 363 ± 58, ConA = 321 + 46). A single experiment was performed to compare the effects of immobilized OKT3 and PHA. After 24 hrs, CD69 upregulation in OKT3-stimulated cultures was very low compared to that with PHA activation (CD3 +: 11.4% vs. 73.7%, CD4 + : 5.6% vs. 84.4% and CD8 + : 5.5% vs. 56.8%). The proportions of cells bearing CD69 increased only slightly after 72 hrs of culture (CD3 + : 11.4% vs. 32.1%, CD4 + : 9.1% vs. 35.5% and CD8 + : 18.0% vs. 14.8%). 70 A) CD3+T-cells 70.0 -r 60.0 + 0 12 24 36 48 60 72 Time (hrs) - • - P H A - a - C o n A Figure 3.8. Comparing the kinetics of CD69 expression induced by PHA and ConA. PHA was used at 1.0 ug/ml and ConA at 0.2 ug/ml. Each value represents the mean ± SEM of 3 independent experiments with 3 healthy controls. No significant difference between the two mitogens was observed. A) CD3+T-cells. B) CD4 subset. C) CD8 subset. 71 B) CD4 subset 90.0 -r 80.0 4-0 12 24 36 48 60 72 Time (hrs) - • - P H A - ^ - C o n A C) CD8 subset 60.0 -r 50.0 4-0 12 24 36 48 60 72 Time (hrs) - • - P H A — C o n A 72 D N A synthesis was also lower with OKT3 stimulation (SI: 11.0 vs. 157.8). 3.2 Effects of Immunosuppressive Drugs on CD69 Expression In Vitro 3.2.1 CsA The ethanohwater vehicle that was used to dissolve the CsA powder (Table 3.5 and Fig. 3.9) did not interfere with the 24-hr CD69 induction in CD3 + (70.8 ± 5.9%, MFI = 126 ± 3), CD4 + (82.9 ± 5.0%, MFI = 126 ± 3) and CD8+T-cells (56.0 + 9.6%, MFI = 117 ± 4). The addition of CsA in the range of 100 to 800 ng/ml did not significantly suppress CD69 expression. For instance, at 200 ng/ml, 55.4 ± 7.9% of CD3 + (MFI = 118 + 4), 79.8 ± 1.4% of CD4 + (MFI = 121 ± 5) and 52.4 ± 8.3% of CD8+T-cells (MFI = 102 ± 5) remained CD69 + (p = NS vs. untreated), and similar levels were observed at 800.0 ng/ml in these cells (56.2 ± 2.0% with MFI = 107 + 2, 73.5 ± 4.9% with MFI = 108 ± 3 and 46.9 ± 7.6% with MFI = 104 ± 4, respectively). The fluorescent intensities in CD8 + and CD3+T-cells appeared to be more sensitive to CsA inhibition (p < 0.05 for 800 ng/ml vs. untreated). Paradoxically, cyclosporine caused a dose-dependent increase in CD69 expression in 72-hr cultures. The expression in untreated controls was 42.6 ± 5.2% for CD3 + , 29.0 ± 4.8% for CD4 + and 36.0 ± 3.7% for CD8+T-cells. CsA at 200.0 ng/ml increased CD69 levels to 52.2 ± 4.1% for CD3 + , 51.7 ± 3.1% for CD4 + and 52.3 ± 12.8% for CD8+T-cells, and further to 60.4 ± 4.2% (p < 0.05 vs. controls), 66.2 ± 8.3% (p < 0.05 vs. controls) and 63.6 ± 10.7%, respectively, at 800.0 ng/ml. In contrast, CsA caused a clear and dose-dependent inhibition of D N A synthesis (Fig. 3.10). 73 Table 3.5. Influence of CsA on PHA-driven CD69 expression1 % C D 3 + %CD4 + %CD8 + CsA (ng/ml) 24 hrs 72 hrs 24 hrs 72 hrs 24 hrs 72 hrs 0 70.8 ±5.9 42 .6±5.2 t 82.9 ±5.0 29.0±4.8t 56.0 ±9.6 36.0 ±3.7 100.0 66.5 ±5.1 N D 85.3 ±3.8 N D 53.8 ±5.8 N D 200.0 55.4 ±7.9 52.2 ±4.1 79.8 ± 1.4 51.7+3.1t 52.4 ±8.3 52.3 ±12.8 400.0 59.4 ±5.1 57.5 ±4.7 81.0 ± 4.3 60.5 ±1.9* 45.3 ±4.3 56.5 ±6.0* 800.0 56.2 + 2.0 60.4 ±4.2* 73.5 ±4.9 66.2 ±8.3* 46.9 ± 7.6 63.6±10.7 MFI (CD69 +CD3 +) MFI (CD69 +CD4 +) MFI (CD69 +CD8 +) CsA (ng/ml) 24 hrs 72 hrs 24 hrs 72 hrs 24 hrs 72 hrs 0 126 ± 3 110 ± 2 t 126 ± 3 103 ±5§ 117 + 4 111 ± 10 100.0 126 ± 2 N D 125 ± 3 N D 114+1 ND 200.0 1 1 8 ± 4 t 109 ± 3 121 ± 5 102 ± 2§ 102 ±5J 110 ±3 400.0 107 ±2% 112 ± 2 119 ± 2 107 ± 1§ 102 ± 3J 108 ± 3 800.0 107 ±2% 115 + 5 108 ± 3 108 ± 4 * * 104 ±4* 115 + 7 a) Cells were stimulated with 1.0 ug/ml of PHA in the presence of CsA at the final concentrations indicated. The drug was dissolved in an ethanohwater (70:30) mixture, and untreated (control) cultures were incubated with the vehicle at a volume equals to those of the CsA-treated cultures. Abbreviations: CsA, cyclosporin A; MFI, mean fluorescence intensity; ND, not determined; PHA, phytohaemagglutinin. * p < 0.05 vs. untreated cultures at 72 hrs ** p < 0.025 vs. untreated cultures at 72 hrs t p < 0.05 vs. untreated cultures at 24 hrs \p< 0.025 vs. its 24-hr level §p < 0.05 vs. its 24-hr level 74 A) CD3+T-cells 80.0 T 0 10 20 30 40 50 60 70 80 Time (hrs) • 0 ng/ml - -a - 200 ng/ml --•©•-- 400 ng/ml - -A- - 800 ng/ml Figure 3.9. Effects of varying CsA concentrations on CD69 kinetics in T-cells. Each value represents the mean ± S E M of 4 independent experiments with 4 healthy controls. A) CD3+T-cells. B) CD4 subset. C) CD8 subset. (*p < 0.05 vs. untreated cultures at 72 hrs and tp < 0.025 vs. its 24-hr level) 75 B) CD4 subset 10 20 80.0 30 40 50 Time (hrs) — • — 0 ng/ml - -a - 200 ng/ml -••©•-- 400 ng/ml C) CD8 subset 60 70 80 - -A- - 800 ng/ml 70.0 4-60.0 O i H + 00 Q U + O N V O Q U 50.0 4-40.0 4-30.0 20.0 4-10.0 4-10 20 60 70 30 40 50 Time (hrs) 0 ng/ml - •• - 200 ng/ml --•©•-- 400 ng/ml - -A- - 800 ng/ml 80 76 90.0 j 80.0 --70.0 --60.0 --g 50.0 --H 40.0 --30.0 --20.0 --10.0 --0.0 +• 0-0 200.0 400.0 600.0 800.0 CsA concentration (ng/ml) Figure 3.10. Dose-dependent inhibition of D N A synthesis by CsA. Each value represents the mean ± S E M of 4 independent experiments with 4 healthy controls. The inhibition of [ H]Thd incorporation was assessed based on this formula: (cpm of untreated - cpm of CsA-treated) x 100/ cpm of untreated (* p < 0.05 vs. 100 ng/ml). 77 3.2.2 MP Control cells exhibited normal CD69 expression (CD3 +: 76.8 + 1.0%, MFI = 131+5; CD4 + : 86.7 ± 3.2%, MFI = 135 ± 6; CD8 + : 57.5 ± 6.1%, MFI = 114 ± 4; Table 3.6 and Fig. 3.11). MP did not influence the proportions of cells expressing CD69 but significantly reduced its surface density. At 1.0 ug/ml of MP, 72.7 ± 3.2% of CD3 + (MFI =112+1), 86.6 ± 1.1% of CD4 + (MFI =114 + 1) and 52.5 ± 3.7% of CD8+T-cells (MFI = 97 ± 3) displayed this marker (p < 0.05 for all MFI values). Higher doses of the drug (10.0 ug/ml) also failed to reduce the number of CD69 + cells significantly (CD3 +: 63.0 ± 1.4%, p < 0.05, CD4 + : 78.9 ± 1.3% and CD8 + : 50.4 ± 13.3%). The level of CD69 expression in control cells at 72 hrs declined to 29.4 ± 1.0% for CD3 + , 28.2 ± 6.6% for CD4 + and 21.0 ± 6.1% for CD8+T-cells. Under the influence of MP, the proportions of CD69-expressing CD4+T-cells were enhanced (1.0 ug/ml = 53.4 ± 5.5% and 10 ug/ml = 47.5 ± 6.4%, p < 0.05 vs. control cells), resembling the effects observed with CsA-treated cultures. The reductions between the 24 and 72-hr CD69 levels in MP-treated cells were, however, significant in both CD4 + and CD8 +T-cell subsets. Proliferation was strongly suppressed with increasing steroid doses (Fig. 3.12). 3.2.3 Tacrolimus The PBMCs in control cultures (Table 3.7 and Fig. 3.13) expressed normal levels of CD69 (CD3 +: 75.7 ± 2.6%; CD4 + : 84.6 ± 0.2%; CD8 + : 69.4 + 6.5%,) after 24 hrs of PHA stimulation. Higher concentrations of tacrolimus reduced the proportions of cells expressing this marker but without significantly affecting its surface density. At 10.0 ng/ml, CD69 78 Table 3.6. Influence of MP on PHA-driven CD69 expression1 % C D 3 + %CD4 + %CD8 + MP (Ug/ml) 24 hrs 72 hrs 24 hrs 72 hrs 24 hrs 72 hrs 0 76.8 ± 1.0 29.4 ±1.0t 86.7 ±3.2 28.2 ± 6.6t 57.5 ±6.1 21 .0±6.1 t 0.01 74.6 ± 1.0 ND 85.8 ±3.7 ND 57.0 + 6.5 N D 0.10 76.7 ±3.9 N D 86.0 ±2.5 ND 62.7 ±2.8 ND 1.0 72.7 ±3.2 40.6±6.5t 86.6+1.1 53.4±5.5*t 52.5 ±3.7 36.2±2.8*t 10.0 63.0±1.4** 35.4 ±2.4t 78.9 ±1.3 47.5±6.4*t 50.4±13.3 16.7±10.1t MFI (CD69 +CD3 +) MFI (CD69 +CD4 +) MFI (CD69 +CD8 +) MP (ng/ml) 24 hrs 72 hrs 24 hrs 72 hrs 24 hrs 72 hrs 0 131 ± 5 103 + 41 135 ± 6 102 ± 6 t 114 ± 4 101 ± 8 0.01 125 ± 6 ND 125 ± 7 N D 112 ± 4 N D 0.10 121 ± 3 N D 124 ± 2 N D 115 ± 8 ND 1.0 112 ± 1 * * 104 ± 3 114 ±1** 98 ± It 97 ± 3 10.0 107 ± 3 * * 101 ± 4 106 ±1** 9 4 ± 2 | 97 ± 6 * * 94 ± 3 a) Cells were stimulated with 1.0 [ig/m\ of PHA in the presence of MP at the final concentrations indicated. The drug was mixed with complete medium, and untreated (control) cultures were incubated with medium at a volume equals to those of MP-treated cultures. Abbreviations: MFI, mean fluorescence intensity; MP, methylprednisolone; ND, not determined; PHA, phytohaemagglutinin. * p < 0.05 vs. untreated cultures at 72 hrs **p < 0.05 vs. untreated cultures at 24 hrs t p < 0.05 vs. its 24-hr level 79 A) CD3+T-cells 90.0 T 80.0 + Time (hrs) — • — 0.0 mg/ml - -• - 1.0 mg/ml - - o- - -10.0 trig/ml Figure 3.11. Effects of varying MP concentrations on CD69 kinetics in T-cells. Each value represents the mean ± S E M of 3 independent experiments with 3 healthy controls. A) CD3 + T-cells. B) CD4 subset. C) CD8 subset. (*p < 0.05 vs. untreated cultures at 72 hrs, **p < 0.05 vs. untreated cultures at 24 hrs, and tp < 0.05 vs. its 24-hr level) 80 B) CD4 subset 100.0 T 0 10 20 30 40 50 60 70 80 Time (hrs) — • — 0.0 mg/ml - -a - 1.0 mg/ml - - o - • 10.0 mg/ml C) CD8 subset 70.0 x 60.0 + 0 10 20 30 40 50 60 70 80 Time (hrs) *—0.0 mg/ml - - a - 1.0 mg/ml --•©•--10.0 mg/ml 81 100.0 -r 90.0 4-80.0 70.0 60.0 4-£ 50.0 + 40.0 4-30.0 4-20.0 4-10.0 4-0.0 + 0.01 0.1 1 MP concentration (u,g/ml) 10 Figure 3.12. Dose-dependent inhibition of D N A synthesis by MP. Each value represents the mean ± S E M of 3 independent experiments with 3 healthy controls. The inhibition of [3H]Thd incorporation was assessed based on this formula: (cpm of untreated - cpm of MP-treated) x 100/ cpm of untreated (*p < 0.05 vs. 0.01 ng/ml). 82 Table 3.7. Influence of tacrolimus on PHA-driven CD69 expression3 % C D 3 + %CD4 + %CD8 + FK506 (ng/ml) 24 hrs 72 hrs 24 hrs 72 hrs 24 hrs 72 hrs 0 75.7 ±2.6 58.0 ±8.2 84.6 ± 0.2t 51.4+11.6 69.4 ±6.5 53.4 ±19.1 0.1 76.1 ±2.8 N D 85.9 + 2.1 N D 72.2 ± 5.6 N D 2.5 60.0 ±5.4* 49.0 ±11.2 69.5 ±9.2* 35.7±10.9 64.1 ±7.0 38.9 ±13.3 10.0 55.9 ±6.6* 55.6 ±5.7 67.8±4.8*t 39.4 ±10.9 50.3 ±9.7 54.6 ±11.4 20.0 44.5 ±2.8* 55.1 ±7.5 60.4±2.1* 34.7 ±14.1 31.2 ±4.9* 41.3 ±4.2 MFI (CD69 +CD3 +) MFI (CD69 +CD4 +) MFI (CD69 +CD8 +) FK506 (ng/ml) 24 hrs 72 hrs 24 hrs 72 hrs 24 hrs 72 hrs 0 124 ± 6 115 + 5 122 ± 6 t 101 ± 4 117 + 5 119 ± 7 0.1 127 ± 5 ND 122 ± 5 N D 130 ±11 ND 2.5 116 ± 4 108 ± 3 116 ± 7t 97 ± 2 124 ± 12 105 ± 6 10.0 109 ± 6 108 ± 4 111 ± 4 100 ± 3 109+11 102 ±5** 20.0 107 ± 7 106 ± 3 103 + 1 97 ± 7 97 + 4* 105 ± 6 a) Cells were stimulated with 1.0 ug/ml of PHA in the presence of tacrolimus at the final concentrations indicated. The drug was dissolved in an ethanohwater (70:30) mixture, and untreated (control) cultures were incubated with the vehicle at a volume equals to those of the tacrolimus-treated cultures. Abbreviations: FK506, tacrolimus; MFI, mean fluorescence intensity; ND, not determined; PHA, phytohaemagglutinin. * p < 0.05 vs. untreated cultures at 24 hrs **p < 0.05 vs. untreated cultures at 72 hrs \p< 0.05 vs. its 72-hr level 83 A) CD3+T-cells 100.0 j 90.0 --80.0 + Time (hrs) — • — 0.0 ng/ml - •• - 2.5 ng/ml • - • - -10 .0 ng/ml — -A- - 20.0 ng/ml Figure 3.13. Effects of varying tacrolimus concentrations on CD69 kinetics in T-cells. Each value represents the mean ± S E M of 3 independent experiments with 3 healthy controls. A) CD3+T-cells. B) CD4 subset. C) CD8 subset. (*p < 0.05 vs. untreated cultures at 24 hrs and tp < 0.05 vs. its 72-hr level) 84 B) CD4 subset 100.0 T 90.0 + Time (hrs) • 0.0 ng/ml - -o- 2.5 ng/ml --•©•--10.0 ng/ml - -A- - 20.0 ng/ml C) CD8 subset 90.0 T 80.0 4-0 10 20 30 40 50 60 70 80 Time (hrs) — • — 0.0 ng/ml - d - 2.5 ng/ml --•©•--10.0 ng/ml - -A- - 20.0 ng/ml 85 expression was reduced significantly to 55.9 ± 6.6% in C D 3 + (p < 0.05 vs. controls), 67.8 ± 4.8% in CD4 + (p < 0.05 vs. controls) and 50.3 ± 9.7% in CD8+T-cells. Marked inhibition of CD69 expression (p < 0.05 vs. controls) was also observed at 20.0 ng/ml in these cells (CD3 +: 44.5 ± 2.8%, CD4 + : 60.4 ± 2.1% and CD8 + : 31.2 ± 4.9%). In contrast to CsA and MP, tacrolimus did not increase but rather moderately reduced the CD69 expression in cells cultured for 72 hrs. Untreated cultures contained 58.0 ± 8.2% of CD3 + , 51.4 ± 11.6% of CD4 + and 53.4 ± 19.1% of CD8+T-cells that were CD69 + . At a concentration of 20.0 ng/ml, tacrolimus reduced CD69 expression (p < 0.05 vs. controls) to 55.1 ± 7.5% for CD3 + , 34.7 + 14.1% for CD4 + and 41.3 ± 4.2% for CD8+T-cells. The in-hibitory effect of tacrolimus at both 24 and 72 hrs appeared to be dependent on its con-centration, and the decline in MFI values were non-significant at almost all dosages tested. This drug also caused a marked and dose-dependent inhibition of D N A synthesis (Fig. 3.14). 3.3 Whole Blood Activation Marker Analysis The erythrolytic treatment of whole blood samples of healthy donors (N) significantly re-duced the number of RBC in these specimens (pre-lysis = 4.8 + 0.2 xl0 9 /ml vs. post-lysis = 0.5 ± 0.0 xl0 9 /ml , p < 0.001) with only a moderate reduction in the leukocyte count (pre-lysis = 6.9 ± 0.4 xl0 6 /ml vs. post-lysis = 4.2 ± 0.3 xl0 6 /ml , p < 0.025). The phenotypic examination of control subjects based on the whole blood method indicated that CD3 + T-cells constituted 66.0 ± 3.1% (Table 3.8) of their lymphocyte populations, and 99.8 ± 0.2% of the double-gated cells were T-cells (CD45+CD14"). The lymphocyte bitmap was well separated from debris and other contaminants such as residual erythrocytes, RBC ghost, and 86 80.0 T 70.0 4-60.0 4-50.0 4-c I 40.0 + 30.0 --20.0 -F 10.0 4 0.0 4 1 1 1 1 1 1 1 1 1 — 1 0.0 2.0 4.0 6.0 8.0 10.0 12.0 14.0 16.0 18.0 20.0 FK506 concentration (ng/ml) Figure 3.14. Dose-dependent inhibition of D N A synthesis by tacrolimus. Each value represents the mean + S E M of 3 independent experiments with 3 healthy controls. Inhibition of [3H]Thd incorporation was assessed based on this formula: (cpm of untreated - cpm of FK506-treated) x 100/ cpm of untreated (*p < 0.05 vs. 0.1 ng/ml). 87 Table 3.8. CD3 cell counts and CD4/CD8 ratios in healthy, dialysis and stable transplant subjects3 Patient cohorts Normal HD CAPD ST %CD3 66.0 ±3.1 67.4 ±5.6 64.4 ±3.5 86.1 ±3.8 t %CD4 48.2 ±5.7 45.6 ±5.9 59.7 ±4.3 47.8 + 3.7t %CD8 20.8 ±2.7 17.4 ±.2.4 11.5 ± 1.7* 23.3 ±2.2 Norm. %CD4 b 72.3 ±11.2 72.5 ±6.9 84.8 ±7.5 56.9 ± 7.1J Norm. %CD8 b 31.9 + 4.3 27.0 ±6.3 18.8 ±2 .3* 25.7 ±2.1 CD4/CD8 ratio 2.76 ±0.50 3.21 ±0.61 6.46 ± 1.21** 2.24 ±0.30 a) Each result is expressed as the mean ± SEM of all 10 participants of each group b) Each value is normalized to its CD3(%) population Abbreviations: CAPD, continuous ambulatory peritoneal dialysis; HD, haemodialysis; SEM, standard error of the mean; ST, stable, long-term renal transplant recipient. < 0.025 vs. controls, HD and CAPD t/?<0.05 vs. CAPD *p < 0.025 vs. controls and ST ** p < 0.05 vs. controls, HD and ST 88 platelets. The proportions of CD4 and CD8 subsets in the CD3 +T-cell populations were 72.3 ± 11.2% and 31.9 ± 4.3%, respectively, accounting for an average CD4/CD8 ratio of 2.76 ± 0.50 (Table 3.8). Table 2.1 and 2.2 summarize the clinical and demographic characteristics of the patient and normal subjects who enrolled in this study. 3.3.1 CD69 Assay of Circulating T-cells Normal Controls Peripheral blood contained low levels of CD69-expressing CD3+T-cells (1.4 ± 0.4%, MFI = 76 ± 3), CD4 + (0.3 ± 0.2%, MFI = 80 ± 5) and CD8 + (1.0 ± 0.2%, MFI = 86 ± 2) as shown in Table 3.9 and Fig. 3.15. These results were comparable to the data acquired by the PBMC technique (Table 3.1). Stimulation with PHA at 1.0 ug/ml induced a profound upregulation (p < 0.001 for % and MFI) of CD69 expression in 48.8 ± 2.8% of C D 3 + (MFI = 124 ± 2), 47.6 ± 3.3% of CD4 + (MFI = 129 ± 3) and 37.4 ± 3.8% of CD8+T-cells (MFI = 115 ± 2). The characteristic decrease in CD69 expression was also observed at 72 hrs, with 25.0 ± 2.1% of C D 3 + (MFI = 117 ± 2, p < 0.05 vs. 24 hrs for % and MFI), 24.1 ± 2.3% of CD4 + (MFI = 118 ± 2,p < 0.025 vs. 24 hrs for % and MFI) and 16.8 ± 1.8% of CD8+T-cells (MFI = 112 ± 3,p < 0.001 vs. 24 hrs for %) remaining CD69 positive. The mean percentage of CD69+T-cells in 24-hr-stimulated whole blood cultures was approximately half of that in the corresponding PBMC cultures (Table 3.1). Significant difference was only detected in the CD4 subset (p < 0.05). However, the kinetics profile of 89 Table 3.9. CD69 expression in healthy, dialysis and stable transplant subjects3 Patient cohorts Time (hrs)b Normal HD CAPD ST % CD69 + 0 1.4 ±0.4 2.8 ±0.5 3.0 ±0.4 3.1 ±0.5 CD3+T-cells 24 48.8 ±2.8* 50.2 ±5.6* 34.0 ±4.1*t 43.0 ±5.9* MFI 0 76 ± 3 77 ± 2 77 ± 2 8 1 ± 1 § 24 124 ± 2 * 119 + 5 117 + 3 116 + 4* % CD69 + 0 0.3 ±0.2 0.7 ±0.5 1.3 ±0.6 0.3 ±0.1 CD4+T-cells 24 47.6 ±3 .3* 47.3 ±7.0* 28.8±4.2*t 43.4 ±6.4* MFI 0 80 ± 5 85 ± 3 84 ± 3 83 ± 3 24 129 ± 3 * 120 ± 4 * 118 + 3*J 120 ± 4 * % CD69 + 0 1.0 ±0.2 1.3 ±0.2 2.9 ±0.7** 3.1±0.7*** CD8+T-cells 24 37.4 ±3.8* 39.9 ±6.2* 30.3 ±4.2* 31.8 ±5.6* MFI 0 86 ± 2 86 ± 3 85 ± 2 93 ± 2§ 24 115 + 2* 111 ± 4 * 110 ± 3* 107±3*A a) Each value is expressed as the mean + S E M of all 10 participants of each group. b) The time during which the whole blood sample was incubated with 1 ug/ml PHA at 37°C in 5%C0 2 humidified air. Abbreviations: CAPD, continuous ambulatory peritoneal dialysis; HD, haemodialysis; MFI, mean fluorescence intensity; PHA, phytohaemagglutinin; SEM, standard error of the mean; ST, stable, long-term renal transplant recipient. *p< 0.001 vs. its 0-hr level ** p < 0.05 vs. controls at 0 hrs *** p< 0.05 vs. controls and HD at 0 hrs §/? < 0.05 vs. C A P D at 0 hrs t p < 0.05 vs. controls and HD at 24 hrs tp< 0.025 vs. controls at 24 hrs Ap < 0.05 vs. controls at 24 hrs 90 C) CD3T-cells 70.0 -r N HD CAPD Subject groups a 0 hrs • 24 hrs ST Figure 3.15. CD69 expression in circulating T-cells of healthy, dialysis and stable RTx subjects. The data of resting and 24-hr-PHA-stimulated cells are expressed as the means ± SEM of all 10 subjects of each group. A) CD3+T-cells. B) CD4 subset. C) CD8 subset. (*p < 0.001 vs. its 0-hr level, **p < 0.05 vs. controls at 0 hrs, ***p < 0.05 vs. controls and HD at 0 hrs, and tp < 0.05 vs. controls and HD at 24 hrs) 91 60.0 B) CD4 subset N HD CAPD Subject groups B O hrs • 24 hrs ST C) CD8 subset 50.0 -r N HD CAPD Subject groups • 0 hrs • 24 hrs ST 92 CD69 expression was preserved in the whole blood cultures. HD and CAPD Subjects Pre-dialysis samples from HD patients were also subject to cytofluorometric analysis. A mean of 67.4 ± 5.6% of their lymphocytes was CD3+T-cells (Table 3.8), which was similar to the percentage observed in control subjects. No abnormal deviation in CD4/CD8 ratio (3.21 ± 0.61) was observed, with 72.5 ± 6.9% and 27.0 ± 6.3% of CD3+T-cells being CD4 + and CD8 + , respectively. The baseline CD69 level in HD subjects was comparable to that in healthy controls, with 2.8 ± 0.5% of CD3 + , 0.7 ± 0.5% of CD4 + and 1.3 ± 0.2% of CD8+T-cells positively stained for this marker (Table 3.9 and Fig. 3.15). Its upregulation in 24-hr-stimulated cultures appeared intact (CD3 +: 50.2 + 5.6%, CD4 + : 47.3 + 7.0% and CD8 + : 39.9 ± 6.2%) when compared to that of the controls (p = NS). The CD3 +T-cell count in CAPD patients (64.4 ± 3.5%) was similar to those of the control and HD groups (Table 3.8). However, there was a significant alteration of the CD4/CD8 ratio (6.46 ± 1.21, p O.05) that was apparently caused by a high CD4 count (84.8 + 7.5%) combined with a significantly low CD8 level (18.8 + 2.3%, p < 0.025 vs. controls). The peripheral T-cells in this cohort were equally quiescent as indicated by their CD69 i o w phenotype in C D 3 + (3.0 ± 0.4%) and CD4+T-cells (1.3 ± 0.6%; Table 3.9 and Fig. 3.15). In contrast, their CD8 subclass possessed a markedly higher fraction (2.9 ± 0.7%) of recently activated cells compared to controls (p < 0.05 for % but not MFI), but these cells displayed 93 normal upregulation of the early marker after 24 hrs of PHA stimulation (30.3 ± 4.2%, MFI = 110 + 3). The same activated whole blood cultures, however, contained considerably less CD69-expressing CD3 + (34.1 ± 4.1%; MFI = 117 ± 3) and CD4+T-cells (28.8 ± 4.2%, MFI = 118 + 3) relative to normal (p < 0.05 for % and p < 0.025 for MFI) and HD groups (p < 0.05 for % and MFI). The reduction was not caused by a delay in CD69 expression kinetics because a significantly smaller (p < 0.05 vs. 24 hrs) number of C D 3 + (18.1 ± 5.3%) and CD4+T-cells (15.1 + 5.2%) expressed this receptor at 72 hrs (CD8 +: 16.0 ± 3.8%). Stable Transplant Subjects The whole blood analysis of ST subjects (Table 3.8) revealed a significantly higher proportion of circulating CD3+T-cells (86.1 + 3.8%) compared to other groups (p < 0.025), while their CD4/CD8 ratio (2.24 ± 0.30, p < 0.05 vs. CAPD) and subset compositions (%CD4 = 56.9 ± 7.1%, p < 0.05 vs. CAPD; %CD8 = 25.7 ± 2.1%, p < 0.025 vs. CAPD) were equivalent to controls and haemodialyzed patients. Similar to other groups, ST patients possessed a small proportion of C D 3 + and CD4+T-cells (3.1 ± 0.5% and 0.3 ± 0.1%, respectively) that expressed CD69 before activation (Table 3.9 and Fig. 3.15). However, significantly higher numbers of CD69 +CD8 +T-cells (3.1 + 0.7%; MFI = 93 ± 2) were found in comparison to normal and HD subjects (p < 0.05), and their surface density was markedly higher than the CAPD group. Despite the high ISD concen-trations that were present in ST patient specimens (e.g. mean serum CsA level 2-3 hrs post-dosing = 968 ± 225 ng/ml, range = 564 - 1343 ng/ml, n = 3), early activation remained optimal as 43.0 + 5.9% of CD3 + , 43.4 ± 6.4% of CD4 + and 31.8 + 5.6% of CD8+T-cells in 94 whole blood cultures displayed CD69 upon 24 hrs of PHA stimulation. The PBMC isolation procedure removed nearly all the CsA in the sample supernatant (14+1 ng/ml, ranged from 12-17 ng/ml, n = 3), and the CD69 expression in the activated CD3+T-cells (54.9 ± 5.7%) in PBMC cultures was slightly higher (p = NS) than that in whole blood cultures. 3.3.2 OX40 Profiles in Control, HD, CAPD, and ST Groups Circulating T-cells were also assessed for their baseline and induced OX40 expression using the whole blood assays (Table 3.10 and Fig. 3.16). Similar to the observations made with PBMCs (Table 3.2), the CD4+T-cells of control subjects displayed low resting OX40 levels (0.2 ± 0.1%; MFI = 73 ± 4). Similar results were obtained in HD and ST patients without any significant intergroup differences. CAPD subjects revealed a different scenario in which the number of their OX40 +CD4 +T-cells (1.3 ± 0.4%, MFI = 78 ± 3) was abnormally higher than controls (p < 0.05). CD3 +T-cell analysis demonstrated the identical phenomenon (CAPD: 1.3 ± 0.4%, MFI = 77 ± 3; control: 0.4 ± 0.2%, MFI = 72 ± 3,p < 0.05 for %). PHA stimulation of healthy donor whole blood increased OX40 expression to 22.4 ± 2.0% for CD3 + (MFI =118 + 1) and 30.0 + 2.8% for CD4+T-cells (MFI = 124 ± 1), with comparable levels detected in both dialysis groups. Activated C D 3 + and CD4+T-cells from CAPD patients were capable of expressing competent levels of OX40 despite their impaired ability in upregulating CD69 expression. Compared to the control and the dialysis groups, ST subjects exhibited a mild reduction in both the proportion of stimulated CD4+T-cells expressing OX40 and staining intensity (22.3 ± 4.8%, MFI = 117 ±4, p < 0.05 vs. CAPD for MFI). OX40 induction in the CD3+T-cells of these transplant patients (16.0 ± 4.0%, MFI = 95 Table 3.10. OX40 expression in healthy, dialysis and stable transplant subjects3 Patient cohorts Time (hrs)b Normal HD CAPD ST % OX40 + 0 0.4 ± 0.2 0.9 ±0.4 1.3 ±0.4 t 0.4 ±0.2 CD3+T-cells 24 22.4 ±2.0* 24.7 ±7.6 27.0 ±3.2* 16.0±4.0*t MFI 0 72 ± 3 76 ± 2 77 ± 3 74 ± 1 24 118 ± 1* 114 + 7 124 ± 3 * 110±4*t %OX40 + 0 0.2 ±0.1 0.5 ±0.2 1.3 ±0.4 t 0.5 ±0.2 CD4+T-cells 24 30.0 ±2.8* 30.8 ±9.4 34.2 ±4.0* 22.3 ±4.8 MFI 0 73 ± 4 81 ± 5 78 ± 3 75 ± 2 24 124 ± 1 * 119 ± 9 129 ± 3 * 117 ± 4*t a) Each value is expressed as the mean ± SEM of 8 participants of each group. b) The time during which the whole blood sample was incubated with 1.0 ug/ml PHA at 37°C in 5%C0 2 humidified air. Abbreviations: CAPD, continuous ambulatory peritoneal dialysis; HD, haemodialysis; MFI, mean fluorescence intensity; PHA, phytohaemagglutinin; SEM, standard error of the mean; ST, stable, long-term renal transplant recipient. * p < 0.001 vs. its 0-hr level t p < 0.05 vs. controls at 0 hrs t p < 0.05 vs. CAPD at 24 hrs 96 A) CD3+T-cells 35.0 T 30.0 + HD CAPD ST Subject group HO hrs • 24 hrs Figure 3.16. OX40 expression in circulating T-cells of healthy, dialysis and stable RTx subjects. The data of resting and 24-hr PHA-stimulated cells are expressed as the means ± S E M of 8 subjects of each group. A) CD3+T-cells. B) CD4 subset. (*p < 0.001 vs. its 0-hr level, tp < 0.05 vs. controls at 0 hrs and tp < 0.05 vs. CAPD at 24 hrs) 97 B) CD4 subset 45.0 -r 40.0 + N HD CAPD Subject groups H 0 hrs • 24 hrs 98 110 ± 4) was similar to other subjects but lower than that in CAPD patients (27.0 ± 3.2%, MFI= 124+ 3,p< 0.05). The possibility of increasing OX40 expression in the CD8 + subset via whole blood stimulation was studied in healthy controls. These cells were OX40" prior to activation (0.0 + 0.0%, MFI = non-stained). The expression increased only slightly by 24 hours to 6.8 ± 1.9% (MFI = 101 +2), representing levels that were similar to those observed in PBMC cultures but remarkably lower (p < 0.001) than the CD69 induction in the same subset. 3.3.3 Serial Immune Monitoring of New Allograft Recipients Table 2.1 summarizes the ISD protocols of the new allograft patients in this study. The prophylactic administration of immunosuppressive antibodies was necessary for three patients in their early post-transplant weeks, with Atgam being used once in each of two patients and OKT3 in another subject for five consecutive days. One subject experienced an acute rejection episode (Day 30) characterized by an interstitial mononuclear cell infiltration and an absence of serum creatinine elevation. The episode was rapidly resolved with high-dose MP treatment for 3 days. No other acute rejection incidence was reported in the rest of group. Complications including oral Candida (Day 36), enterococcal infection (Day 63), and acute tubular necrosis (Day 2) were diagnosed in three patients and were all reversible. Prior to kidney transplantation, new transplant patients had a CD3 cell count (80.6 ± 2.2%, Table 3.11 and Fig. 3.17) that was significantly higher than those of the non-transplant groups but similar to that of the ST subjects. The proportion dropped substantially to 53.0 ± 9.6% (p < 0.05 vs. pre-Tx) within the immediate post-transplant days but returned gradually 99 Table 3.11. CD3 cell counts and CD4/CD8 ratios in new (de novo) RTx patients3 % Total lymphocytes % CD3+T-cells Post-RTx Norm. Norm. CD4/CD8 Course %CD3 %CD4 %CD8 %CD4 b %CD8 b ratio Pre-Tx 80.6 + 2.2t 50.5 ±4.0 26.1 ±3.7 t 62.9 ±7.1 34.0 ± 4.8{ 2.72±0.71t Days 2-4 53.0 ±9.6* 45.8 ±1.6 23.2 ±2.7 72.9 ±8.7 34.5 ±2.8 2.12 ±0.23 Days 5-7 61.9 ±9.7 50.1 ±3.6 21.4 ±3.8 68.5 ± 1.8 41.0 ±5.2 2.87 ±0.57 Week 2 63.5 ±8.0* 51.4 ± 5.5 17.3 ±3.2 69.6 ±13.1 24.8 ±5.1 3.90 ±1.04 Week 3 72.1 ± 2 . 1 * 45.8 ±4.3 14.2±2.4** 60.0 ±8.0 20.1 ± 3 . 1 * 4.24 ±1.79 Week 4 74.9 ±3.6 52.5 ±4.8 15.6 ±2.8* 68.7 ±8.2 21.5 ±4.0 3.44 ±0.88 Week 5 75.2+1.2* 56.0 ±3.3 21.6 ±3.7 75.2 ±6.8 28.4 ±6.1 3.55 ±0.96 Weeks 6-7 81.2 ± 3.7 51.3 ±2.9 14.3+2.2** 61.5 ±6.6 20.0 ±2 .5* 5.29 ±0.66 Weeks 8-9 78.2 ±4.1 51.1 ±5.4 15.6 ± 2.3* 61.6 ±10.4 20.8 ±3.0* 3.57 ±1.08 Weeks 10-12 81.8 ±3.4 46.7 ±7.0 19.2 ±3.2 50.2 ±10.4 26.5 ± 5.0 3.07 ±0.85 a) Each result is expressed as the mean ± SEM of all 10 subjects. b) Each value is normalized to its CD3(%) population Abbreviations: Pre-Tx, pre-transplant; RTx, renal transplantation; SEM, standard error of the mean. * p < 0.05 vs. pre-Tx level ** p < 0.025 vs. pre-Tx level \p < 0.025 vs. controls, HD and CAPD tp< 0.025 vs. CAPD 100 Figure 3.17. CD3 cell count in the peripheral circulation of new RTx patients. Each value represents the mean ± S E M of all ten subjects. The broken line indicates the mean pre-Tx value. (*p < 0.05 vs. pre-Tx level and tp < 0.025 vs. controls, HD, and CAPD) 101 to its pre-operative level within a few weeks (e.g. Weeks 6-7: 81.2 ± 3.7%, p = NS). This level was consistently maintained in the rest of the monitoring period. The average pre-transplant CD4/CD8 ratio (2.72 + 0.71) was similar to those of the controls and other subjects but lower than that of the CAPD patients. Although it fluctuated through-out the monitoring period (Table 3.11), no significant alteration was observed over time. The CD4 cell count remained constant after RTx when compared to its pre-operative level (62.9 ± 7.1%, p = NS vs. all cohorts). On the contrary, the CD8 cell count (pre-Tx: 34.0 + 4.8%, p < 0.025 higher than CAPD) was markedly depleted during Weeks 3, 6-7, and 8-9. Tables 3.12 and 3.13 summarize the serological and clinical parameters of NT patients relevant to this study. Serum creatinine was pathologically elevated before RTx (961 ±133 umol/1) and fell progressively within the initial post-transplant period to a mean of 322 ± 170 umol/1 by Week 5 and 192 ± 45 umol/1 by Weeks 10-12. The CsA dosages and whole blood levels were adjusted to conform to the established clinical ranges throughout the monitoring program, and the dose of prednisone administered was gradually reduced. These variables correlated weakly with CD69 and OX40 induction (Table 3.12 and 3.13). CD69 Expression Prior to RTx, 3.9 ± 1.2% of unstimulated CD3+T-cells expressed CD69 with an MFI of 82 ± 2 (Table 3.14 and Fig. 3.18). These cells maintained a similar degree of expression through-out the entire monitoring period, including Days 2-4 (3.1 ± 1.1%) and Weeks 10-12 (2.3 ± 0.5%). These results were all comparable to the data obtained in other groups. 102 Table 3.12. CsA and prednisone dosage levels and plasma cyclosporine determinations NT patients3 Dosage Levels Post-RTx Course Whole blood trough CsA (ng/ml) CsA (mgb.i.d.) Pred (mg o.d.) Pre-Tx N D 424±128 0 Days 2-4 445 ± 36 629 +121 63 ± 7 Days 5-7 393 ± 44 650±102 45 ± 13 Week 2 395 ±36 613 ± 52 26 ± 6 Week 3 471 ± 60 680 ± 90 20 ± 2 Week 4 368 ± 2 9 683 ±100 18 + 1 Week 5 391+ 64 730 ± 92 19+1 Weeks 6-7 379 ±35 555 ± 8 8 16 + 2 Weeks 8-9 347 ±35 490 ± 68 15 + 2 Weeks 10-12 320 ± 8 3 450±100 13 + 1 Correlation analysis r b %CD69 in CD4+T-cells +0.13 +0.03 +0.27 %CD69 in CD8+T-cells +0.12 +0.05 +0.30 %OX40 in CD4+T-cells +0.13 +0.07 +0.20 a) Each data is expressed as the mean ± SEM of 6 subjects. b) Correlation studies comparing each clinical variable to PHA-induced activation marker expression: p = NS for all analyses Abbreviations: b.i.d., two times a day; CsA, cyclosporin A; NT, new (de novo) renal transplant recipient; o.d., everyday; Pred, prednisone; Pre-Tx, pre-transplant; r, correlation coefficient; RTx, renal transplantation; SEM, standard error of the mean. 103 Table 3.13. Serum creatinine concentrations in the early post-transplant perioda'b Post-RTx Course Serum creatinine level (umol/1) Post-RTx Course Serum creatinine level (umol/1) Pre-Tx 961±133 Week 4 234 ± 97 Days 2-4 379 ± 8 0 Week 5 322 ±170 Days 5-7 311 ±143 Weeks 6-7 2821136 Week 2 131 ± 8 0 Weeks 8-9 199 ± 5 9 Week 3 312±172 Weeks 10-12 192145 a) Data are expressed as the means 1 SEM of 7 subjects. b) Correlation analyses comparing plasma creatinine levels to PHA-induced activation marker expression: %CD69 in CD4+T-cells: r = +0.12,p = NS %CD69 in CD8+T-cells: r = +0.1 \,p = NS %OX40 in CD4+T-cells: r = +0.25,p = NS Abbreviations: Pre-Tx, pre-transplant; RTx, renal transplantation; SEM, standard error of the mean. 104 Table 3.14. Longitudinal evaluation of CD69 expression in NT patients1 A) CD69 +CD3 +T-cells % MFI Post-RTx Course Baseline PHA Baseline PHA Pre-Tx 3.9+1.2 34.2±5.5*t 82 ± 2 110±3*t Days 2-4 3.1 ± 1.1 29.9 ±5.7 83 ± 3 96±4**§ Days 5-7 2.4 ± 0.7 22.1 ±7.1 79 ± 2 96 ± 5**§ Week 2 3.1+0.5 16.0±6.1*§ 85 ± 1 96±4**§ Week 3 1.9 + 0.4 29.8 ±6.4 84 ± 3 102 ± 5 Week 4 1.9 + 0.3 26.0 ±4.0 85 ± 3 105 ± 2 Week 5 1.7 ±0.4 28.4 ±6.1 84 ± 4 107 ± 4 Weeks 6-7 2.4 ±0.5 32.7 ±3.8 88 ± 3 106 ± 3 Weeks 8-9 2.0 ±0.6 42.8 ±7.1 88 ± 3 111 ± 2 Weeks 10-12 2.3 ±0.5 35.9 +3.1*tt 88 ± 4 109±4*t{ a) Baseline values were determined at 0 hrs; activated values were determined after 24 hrs of incubation with 1.0 ug/ml PHA at 37°C in 5%C0 2 humidified air. Each result is expressed as the mean ± S E M of all 10 subjects. Abbreviations: MFI, mean fluorescence intensity; NT, new (de novo) renal transplant recipient; PHA, phytohaemagglutinin; Pre-Tx, pre-transplant; RTx, renal trans-plantation; SEM, standard error of the mean. *p < 0.001 vs. its baseline level ** p < 0.05 vs. its baseline level *** p < 0.025 vs. its baseline level tp < 0.05 vs. controls at 24 hrs Ap < 0.025 vs. controls at 24 hrs tp < 0.05 vs. Week 2 at 24 hrs V p < 0.05 vs. controls and HD at 0 hrs §/? < 0.05 vs. pre-Tx, controls, HD, CAPD, and ST at 24 hrs 105 Table 3.14. Longitudinal evaluation of CD69 expression in NT patients B) CD69 +CD4 +T-cells % MFI Post-RTx Course Baseline PHA Baseline PHA Pre-Tx 0.6 + 0.3 32.6±5.6*t 84 ± 3 113 ±3*A Days 2-4 0.2 ± 0.3 25.8 ±5.5 84 ± 3 98 ± 4***§ Days 5-7 0.6 ± 0.5 18.6 ±6.5 76 ± 4 96 ± 6***§ Week 2 1.7 ±0.7 12.2±5.8**§ 77 ± 3 93 ±6***§ Week 3 0.7 ±0.4 28.2 ±6.5 79 ± 3 104 ± 6 Week 4 0.5 ±0.3 24.6 ±4.3 80 ± 2 107 ± 3 Week 5 0.1 ±0.1 24.6 ±5.8 79 ± 2 109 ± 4 Weeks 6-7 0.4 ±0.1 29.6 ±3.8 82 ± 4 107 ± 4 Weeks 8-9 0.3 ±0.1 38.3 ±6.9 83 ± 3 114 + 2 Weeks 10-12 0.5 ±0.2 32.2 ± 3.6*ft 81+3 112±4*A{ 106 Table 3.14. Longitudinal evaluation of CD69 expression in NT patients C) CD69 +CD8 +T-cells % M F I Post-RTx Course Baseline PHA Baseline PHA Pre-Tx 5.1 ± 1.8V 27.9 + 4.6* 8912 10213*A Days 2-4 4.812.1 26.615.1 8813 9 4 1 2 § Days 5-7 3.010.8 17.817.0 8213 8 9 1 4 § Week 2 4.1 10.8V 13.6 ± 5.2§ 8912 8 9 1 3 § Week 3 4.1 1 1.7 23.915.3 9013 9415 Week 4 4.611.5 21.514.5 9214 9712 Week 5 3.71 1.2 25.515.5 8814 100 + 4 Weeks 6-7 6.212.3 26.5 13.1 91 1 2 9613 Weeks 8-9 5.1 12.1 38.5 16.3 8714 101 12$ Weeks 10-12 4.1 1 1.0V 27.814.0*$ 8914 9814*A 1 0 7 A) CD3+T-cells 50.0 T 45.0 + Pre-Tx 2-4 5-7 8-14 15-21 22-28 29-35 36-49 50-63 64-84 (wk) (1) (1) (2) (3) (4) (5) (6-7) (8-9) (10-12) Post-transplant course HO hrs • 24 hrs Figure 3.18. Longitudinal monitoring of CD69 expression in circulating T-cells of new RTx patients. The data of resting and 24-hr PHA-stimulated cells are expressed as the means ± SEM of all 10 subjects. The broken line indicates the mean pre-Tx value. A) CD3+T-cells. B) CD4 subset. C) CD8 subset. (*p < 0.001 vs. its 0-hr level, **p < 0.05 vs. its 0-hr level, Vp < .0.05 vs. controls and HD at 0 hrs, tp < 0.05 vs. controls at 24 hrs, %p< 0.05 vs. Wk 2 at 24 hrs and §p< 0.05 vs. pre-Tx, controls, HD, CAPD and ST at 24 hrs) 108 B) CD4 subset Pre-Tx 2-4 (wk) (1) 5-7 8-14 15-21 22-28 29-35 (1) (2) (3) (4) (5) Post-transplant course II0 hrs • 24 hrs 36-49 50-63 64-84 (6-7) (8-9) (10-12) C) CD8 subset 50.0 -r Pre-Tx 2-4 5-7 8-14 15-21 22-28 29-35 (wk) (1) (1) (2) (3) (4) (5) Post-transplant course HO hrs • 24 hrs 36-49 50-63 64-84 (6-7) (8-9) (10-12) 109 The activated whole blood cultures from NT patients prior to RTx revealed a significant CD69 upregulation in CD3+T-cells (34.2 ± 5.5%, MFI = 110 ± 3, p < 0.001 vs. non-stimulated cells); however, the level was lower than that in the control group (p < 0.05). The ability of these cells to upregulate their CD69 expression declined progressively after RTx, as the CD69 induction was markedly reduced on Days 2-4 (29.9 ± 5.7%) and in Week 2 (16.0 ± 6.1%, MFI = 96 ± 4, p < 0.05 vs. pre-Tx % and MFI). Within Week 2, the proportion of T-cells that became CD69 + after stimulation was less than those of other subject groups. T-cells recovered their ability to express CD69 after Week 3 (29.8 + 6.4%; MFI = 102 ± 5). The mean percentage and MFI values were now close to the pre-transplant levels but remained lower than those of the control subjects. Similar levels were maintained for the rest of the monitoring period (e.g. Weeks 10-12: 35.9 ± 3.1%; MFI = 109 + 4). Analysis by linear regression revealed that CD69 expression before and after stimulation were not significantly correlated (%: r = +0.49; MFI: r = +0.34), but the activated MFI values were associated with the CD3 cell count (MFI: r = +0.92,/? < 0.025; %: r = +0.59, p = NS). The implantation of the renal allograft did not activate circulating CD4 +T-cells, as indicated by their lack of significant CD69 upregulation after transplantation (Table 3.14 and Fig. 3.18). No significant difference in the proportion or fluorescent staining of CD69 + CD4 + T-cells was observed between the pre-transplant (0.6 + 0.3%, MFI = 84 ± 3, p - NS vs. all other cohorts) and post-transplant periods from Days 2-4 (0.2 ± 0.3%, MFI = 84 + 3) to Weeks 10-12 (0.5 + 0.2%, MFI = 81 ± 3). 110 Stimulated lymphocytes revealed a significantly weaker CD69 induction in CD4 T-cells prior to RTx (32.6 ± 5.6%, MFI = 113 ± 3,p < 0.05 for % andp < 0.025 for MFI) compared to healthy controls. After engraftment, the proportion and fluorescent intensity of CD69 +T-cells began to decline progressively to 25.8 ± 5.5% on Days 2-4 (MFI = 98 ± 4,p < 0.025 vs. pre-Tx MFI) and to 12.2 ± 5.8% during 2nd week (MFI = 93 ± 6,p < 0.05 vs. pre-Tx % and MFI). The activation response during this time was markedly lower than those of other subject groups. The proportions of CD69+T-cells in stimulated cultures remained significantly higher than those of the non-stimulated cells. The CD69 upregulation in CD4+T-cells returned towards the pre-transplant level during Week 3 (28.2 ± 6.5%; MFI = 104 + 6) and remained stable until Weeks 10-12 (32.2 ± 3.6%; MFI = 112 ± 4). This level showed no signs of impairment compared to Week 2 (p < NS); however, similar to the pre-transplant state, it was markedly lower (p < 0.05) than the controls. Further evaluation showed that the percentage and MFI values of CD69 +CD4 +T-cells in resting and activated cultures were not correlated (%: r = +0.088; MFI: r = +0.59; both p = NS). However, both parameters in activated cells exhibited a positive correlation with CD3 levels (%: r = +0.64, p < 0.05; MFI: r = +0.86,p < 0.025). Peripheral CD8+T-cells expressed low baseline CD69 levels before and after RTx (Table 3.14 and Fig. 3.18). Significantly more pre-transplant CD8+T-cells expressed this marker (5.1 ± 1.8%, p < 0.05; MFI = 89 ± 2) compared to controls and HD patients. A similar pattern emerged during the immediate post-transplant months (e.g. Days 2-4: 4.8 ± 2.1%; Weeks 10-12: 4.1 ±1.0%), during which the percentage values were consistently higher than those in normal and haemodialyzed groups (p < 0.05). I l l The pre-transplant CD8+T-cells displayed normal upregulation of CD69 on activation (27.9 ± 4.6%, MFI = 102 ± 3, p = NS vs. all other groups for % and MFI). The PHA-induced expression started to decline on Days 5-7 (17.8 ±7 .0%; MFI = 89 ± 4, p < 0.05 vs. pre-Tx MFI) and was significantly impaired in the 2nd week (13.6 ± 5.2%, MFI = 89 ± 3, p < 0.05 vs. pre-Tx % and MFI). These levels were substantially lower than those observed in other groups. Normal levels of CD69 expression returned during Week 3 (23.9 ± 5.3%; MFI = 94 ± 5) and approached the pre-surgical level during Weeks 10-12 (27.8 ± 4.0%, MFI = 98 ± 4, p = NS vs. all other groups for % and MFI). The changes in CD69 expression between resting and activated CD8+T-cells did not correlate closely (%: r = +0.49; MFI: r = +0.34; bothp = NS). The receptor surface density, but not the proportion of CD69 + cells (r = +0.49, p - NS), was positively associated with the CD3 count (r = +0.72, p < 0.025). The baseline CD69 expression in isolated PBMCs of NT patients was also measured on several occasions (n = 5) during the post-transplant period. CD69 was expressed in 2.4 ± 0.3 in CD3 + , 0.8 ± 0.8% in CD4 + and 1.4 ± 0.8% in CD8+T-cells. These results were comparable to those of whole blood samples which were run in parallel (CD3 +: 2.4 ± 0.4, CD4 + : 1.1 ± 0.9 and CD8 + : 4.9 ± 1.4%), thus indicating that the low CD69 levels detected with the whole blood method were not due to the destruction of activated cells by the stain-lyse procedure. OX40 Expression OX40 expression was observed in 0.9 ± 0.4% (MFI = 80 ± 3) of the pre-transplant non-stimulated CD3+T-cells, and no significant increase was observed during the initial three months of RTx (Table 3.15 and Fig. 3.19). Following PHA activation, OX40 expression in 112 Table 3.15. Longitudinal evaluation of OX40 expression in NT patients3 A) OX40 +CD3 +T-cells % MFI Post-RTx Course Baseline PHA Baseline PHA Pre-Tx 0 .9±0.4 15.9 + 3.4*$ 8013 115 13**A Days 2-4 0.810.2 6.5 12.3*§ 7713 93 1 6***A Days 5-7 0.1 10.1 9.012.8 7212 105 15 Week 2 0.210.1 8.816.7 7616 103 111 Week 3 0.710.3 16.614.6 8012 10816 Week 4 0.5 1 0.2 8.210.3*§ 81 13 11018 Week 5 0.810.5 5.410.3*§ 7 6 1 2 10717 Weeks 6-7 0.910.3 10.011.8 7912 111 13 Weeks 8-9 0.910.4 13.3 13.0 7912 106 1 3**A Weeks 10-12 0.610.3 10.3 I4.0*t 81 13 10413**A a) Baseline values were determined at 0 hrs; activated values were determined after 24 hrs of incubation with 1.0 ug/ml PHA at 37°C in 5%C0 2 humidified air. Each result is expressed as the mean 1 SEM of 7 subjects. Abbreviations: MFI, mean fluorescence intensity; NT, new (de novo) renal transplant recipient; PHA, phytohaemagglutinin; Pre-Tx, pre-transplant; RTx, renal trans-plantation; SEM, standard error of the mean. *p < 0.025 vs. its baseline level ** p < 0.001 vs. its baseline level *** p < 0.05 vs. its baseline level \p< 0.05 vs. controls and CAPD at 24 hrs $ p < 0.05 vs. CAPD at 24 hrs §p < 0.05 vs. pre-Tx, controls, and CAPD at 24 hrs Ap < 0.05 vs. pre-Tx, controls, CAPD, and ST at 24 hrs 113 Table 3.15. Longitudinal evaluation of OX40 expression in NT patients B) OX40 +CD4 +T-cells % M F I Post-RTx Course Baseline PHA Baseline PHA Pre-Tx 1.5 ±0.7 23.2 ±4.4* 80 ± 3 120 ±4** Days 2-4 1.7 ± 1.0 10.4±3.2*§ 86 ± 4 96 ± 7§ Days 5-7 0.4 ± 0.2 13.8 ±4.3 75 ± 2 108 ± 5 Week 2 0.8 ±0.3 11.2 ± 6.4 74 ± 6 104 ± 12 Week 3 2.0 ± 1.2 24 2 + 4.6 83 ± 3 115 ± 7 Week 4 1.7 ± 1.0 15.3 ±2.0*t 79 ± 2 115 + 6 Week 5 0.1 ±0.1 7.9 ± 1.0*§ 75 ± 3 109 ± 6 Weeks 6-7 2.3 ± 1.1 15.9±2.7*t 78 ± 2 114 + 3*** Weeks 8-9 1.8 + 1.1 19.0 ±4.6 78 ± 2 112 + 3*** Weeks 10-12 2.3 ± 1.4 15.8±2.6*t 80 ± 2 109 ±4*** 114 A) CD3+T-cells 25.0 T 20.0 + 13 15.0 + o H + m Q O + o X o 10.0 + Pre-Tx 2-4 5-7 8-14 15-21 22-28 29-35 (wk) (1) (1) (2) (3) (4) (5) Post-transplant course • 0 hrs • 24 hrs 36-49 50-63 64-84 (6-7) (8-9) (10-12) Figure 3.19. Longitudinal monitoring of OX40 expression in circulating T-cells of new RTx patients. The data of resting and 24-hr PHA-stimulated cells are expressed as the means ± S E M of 7 subjects. The broken line indicates the mean pre-Tx value. A) CD3+T-cells. B ) CD4 subset. (*p < 0.025 vs. it 0-hr level, \p < 0.05 vs. controls and CAPD at 24 hrs, %p < 0.05 vs. CAPD at 24 hrs and §p < 0.05 vs. pre-Tx, controls, and CAPD at 24 hrs) 115 B) CD4 subset 35.0 T 30.0 + 25.0 4-Post-transplant course • 0 hrs • 24 hrs 116 pre-transplant cells (15.9 ± 3.4%; MFI = 115 ± 3) was normal but significantly (p < 0.05) less than that of C A P D group. Marked inhibition of OX40 expression (p < 0.05 vs. pre-Tx % and MFI) occurred on Days 2-4 (6.5 ± 2.3%; MFI = 93 ± 6) and during Weeks 4 and 5, when these results were significantly lower than those of normal and CAPD groups. The significant impairment of upregulation continued throughout the monitoring period despite the fact that the stimulated CD69 levels were normal (Table 3.14A). At the end of the program, OX40 induction (10.3 ± 4.0% and MFI = 104 ± 3) remained significantly lower (p < 0.05) than those in controls and CAPD groups. No correlation was observed between 0-hr and 24-hr levels (%: r = +0.29; MFI: r = +0.37) and between CD69 and OX40 expression in stimulated CD3+T-cells (%: r = +0.43; MFI: r = +0.62), but a significant and positive association was observed between the OX40 MFI of activated CD3+T-cells and CD3 cell count (MFI: r = +0.78,p < 0.025; %: r = +0.43, p = NS). The baseline and stimulated OX40 levels in the CD4 subset were closely parallel to those in the CD3+T-cells. A negligible proportion of circulating CD4+T-cells expressed OX40 before RTx (1.5 ± 0.7%, MFI = 80 ± 3). This level was similar to those of controls and other groups, and the post-transplant baseline expression closely resembled this level throughout the monitoring period (Table 3.15 and Fig. 3.19). Whole blood stimulation increased OX40 expression in CD4+T-cells to levels (23.2 ± 4.4%, MFI = 120 + 4) that were similar to those of controls and other patients. The number and surface density of activated cells were markedly modulated (p < 0.05 vs. pre-Tx % and MFI) after RTx on Days 2-4 (10.4 ± 3.2%, MFI = 96 ± 7) and in Week 5 (7.9 + 1.0%, MFI = 109 117 + 6). The reduction was significant when these levels were compared to those of controls and CAPD patients. Between Days 5-7 and Week 12, OX40 expression (% and MFI) was consistently but non-significantly lower than the activation responses before transplantation and in other subjects (except Weeks 3 and 5). Between Weeks 3 and 12, the OX40 upregulation was impaired when CD69 induction appeared normal (Table 3.14B). The quiescent and activated OX40 levels were not significantly associated (%: r = +0.48; MFI: r = +0.57). The MFI after induction and CD3 cell count were positively correlated (MFI: r = +0.81, p < 0.025; %: r = +0.48, p = NS), and a similar relationship existed between CD69 and OX40 in activated CD4+T-cells (MFI: r = +0.64, p < 0.025; %: r = +0.56, p = NS). PHA-induced Proliferation Cell proliferation was also evaluated in long-term PHA-activated cultures of new allograft recipients (Fig. 3.20). The pre-operative SI (436.8 ± 80.6) was not significantly different from that of healthy controls (352.1 ± 50.2). D N A synthesis was substantially reduced after RTx, as indicated by the lower SI values (p < 0.05 vs. pre-Tx) observed on Days 5-7 (116.5 ± 61.9) with a nadir on Days 8-14 (38.9 ± 16.7). It gradually increased towards the pre-transplant level within the following post-transplant month, reaching 540.4 ± 78.2 during Weeks 8-9 and 191.2 ± 81.8 during Weeks 10-12 (p = NS). Activation marker expression and proliferation appeared to undergo similar patterns of change throughout the monitoring period. Positive correlations (p < 0.025 for % and MFI) were observed between SI values and CD69 induction in C D 3 + (%: r = +0.76; MFI: r = +0.75), CD4 + (%: r = +0.73; MFI: r = +0.75) and CD8+T-cells (%: r = +0.82; MFI: r = 118 800 700 + 600 + Post-transplant course Figure 3.20. Longitudinal monitoring of D N A synthesis in PBMCs of new RTx patients. Stimulation indices are expressed as the means ± S E M of 6 subjects. The broken line represents the mean pre-Tx value. N D = not determined. (*p < 0.05 vs. pre-Tx level) 119 +0.81). In contrast, D N A synthesis did not correlate (p = NS) with OX40 induction in CD3+T-cells (%: r = +0.26; MFI: r = +0.33) and in CD4+T-cells (%: r = +0.23; MFI: r = +0.35). Proliferation and CD3 cell count were weakly associated (r = +0.52, p = NS). Activation Marker Levels During Complications The baseline and PHA-induced expression of CD69 and OX40 in peripheral blood samples obtained before and after immunological and non-immunological complications were summarized in Table 3.16. No abnormal upregulation was observed in the circulating T-cells of patients who had oral C a n d i d a , enterococcal infection and acute tubular necrosis when compared to the mean values of the entire NT population. The CD69 expression in stimulated T-cells was slightly depressed in several occasions. The peripheral T-cells of the patient who underwent a reversible acute rejection episode did not demonstrate any significant changes in their CD69 and OX40 expression. 120 Table 3.16. CD69 and OX40 expression during transplant complications CD69 OX40 Day of Complications Analysis %CD3 %CD4 %CD8 %CD3 %CD4 Acute Day 23 1.7 0.0 1.1 0.3 0.6 allograft (17.7) (15.7) (6.6) (8.5) (12.1) rejection (Day 30) Day 33 2.9 0.0 3.4 0.0 0.0 (16.1) (13.6) (9.9) (5.9) (9.6) Acute Pre-Tx 4.0 0.8 5.8 2.3 4.6 tubular (33.1) (26.9) (31.8) (20.1) (25.5) necrosis (Day 2) Day 6 2.1 0.2 5.7 0.3 1.0 (21.8) (16.1) (15.1) (14.2) (20.8) Enterococcal Day 48 0.8 0.0 1.0 2.0 4.4 infection (44.6) (40.5) (35.9) (14.9) (22.8) (Day 63) Day 63 0.7 0.0 1.1 2.6 1.8 (42.0) (38.3) (35.0) (14.5) (14.5) Oral Day 29 3.5 0.4 8.7 0.2 0.1 C a n d i d a (13.2) (9.9) (18.4) (4.8) (6.0) (Day 36) Day 43 6.5 0.8 10.2 0.2 0.6 (17.2) (12.7) (23.1) (5.6) (7.5) Mean valuesb Pre-Tx 3.9 ± 1.2 0.6 ±0.3 5.1 ± 1.8 0 .9±0.4 1.5 ±0.7 (all NT (34.2 ±5.5) (32.6 ± 5.6) (27.9 ±4.6) (15.9 ±3.4) (23.2 ±4.4) subjects) Post-RTx 2.3 ±0.5 0.5 ±0.2 4.1 ± 1.0 0.6 ±0.3 2.3 ± 1.4 (35.9 ±3.1) (32.2 ±3.6) (27.8 ±4.0) (10.3 ±4.0) (15.8 ±2.6) a) 4 NT patients experienced 4 separate episodes of complications during the monitoring period. The days of analysis represented the time when these patients were evaluated. Baseline values were evaluated at 0 hrs, and the values in parenthesis are the activation marker levels that were evaluated after 24 hrs of PHA induction. b) the mean values of the NT group before transplantation and Weeks 10-12 post-transplant (Table 3.14 and 3.15) Abbreviations: NT, new (de novo) renal transplant recipient; PHA, phytohaemagglutinin; Pre-Tx, pre-transplant; RTx, renal transplantation. 121 Chapter Four Discussion As lymphocytes undergo cellular activation, they express surface proteins that are involved in signal transduction, adhesion, growth factor binding, or other effector functions inherent to cycle-committed cells. The presence of these molecules can assist in identifying activated cells that might result from, or participate in, the pathogenesis of immune diseases. Conversely, their absence during stimulation might signify immune deficiency. This study focused on three activation markers, namely a membrane protein expressed in the early stage of T-cell activation (CD69), an intermediate-stage molecule (OX40) and a late-stage molecule (CD25), and on using these markers to study circulating T-cells in renal failure and kidney transplantation. These proteins have been shown to be upregulated in peripheral T-cells in diseases such as G V H D and cardiac allograft rejection [66,125,199,232], but they have not been studied together in renal transplantation. Monitoring de novo transplant recipients by assessing the peripheral whole blood CD69 and OX40 levels in their specific T-cell subsets appeared to be the first of its kind in the field of transplant research. Before proceeding to examine the behaviour of T-cells of dialysis and transplant patients, the kinetics profiles of the activation markers in the T-cells of normal subjects were analysed to delineate their functional significance and expression requirements during T-cell activation. 4.1 Healthy-state Circulating T-cells were CD69 l o w Consistent with earlier reports [116,129-130,137,183], the flow cytometric analysis of peripheral blood T-cells isolated from healthy individuals revealed low levels of CD69 expression (< 3.2% in the CD4 + or CD8 +T-cell subset in normal subjects), irrespective of the preparatory technique (whole blood or PBMC) that was used. These control subjects 122 reported normal renal function and immune status, and none have undergone any immunostimulatory (e.g. vaccination) or suppressive therapy recently. Although non-specific staining could not be ruled out, the small proportion of weakly stained CD69+T-cells observed might represent antigen-primed cells that re-entered the circulation from lymph nodes, tonsils, gastrointestinal mucosa and other areas where high proportions of T-cells display this marker [140-142]. The different steps in mononuclear cell isolation, such as saline dilution and gradient centrifugation, did not upregulate CD69 expression. A similar conclusion can be drawn with the whole blood phenotyping technique since the osmotic shock had no effect on its baseline levels. PBMCs were devoid of erythrocytes, platelets, and polymorphonuclear leukocytes according to previous studies [226,233] and as confirmed by light microscopy. Pooled human serum, complete medium, and culturing conditions did not induce CD69 expression or proliferation in mitogen-free cultures; the slightly higher levels observed at 4 hrs in non-stimulated cells might be due to a transient stimulation by serum growth factors. In addition, the double-gating technique was capable of focusing on T-cells (98.9 ± 0.6% purity) without including monocytes (0.1 ± 0.0%), a cell type that also expresses CD69 [133]. 4.2 Stimulation of CD69 Expression by PHA The tetrameric PHA crosslinks signal-transmitting surface proteins such as TCR, CD3, CD2, and CD45, thus activating T-cells and upregulating their CD69 expression. These PHA tar-gets are responsible for propagating intracellular transduction signals following antigen recognition and the provision of APC help [73-74,234]. Activation begins with an increase 123 of [Ca 2 +], which occurs within minutes of stimulation [211]. It is mediated by intracellular mobilization through IP 3 and sustained by extracellular influx (e.g. via CRAC) [210]. Then, the Ca2+-dependent pathways (e.g. the P K C / M E K / E R K cascade) become functional [157, 235]. The immediate rise of [Ca2+]j and the activation of PKC pathway trigger the early expression of CD69, which was already significantly increased by 4 hrs of culturing [129]. The signals generated through the antibody-crosslinking of CD3 and CD2 are synergistic in promoting transcription factor activation [234]. Similarly, through crosslinking with CD3, CD2, and other surface proteins, PHA enhances the expression and D N A binding activities of AP-1, NF-AT, and N F - K B [80,210], and these nuclear mediators have been proven in several studies to be the crucial regulators of CD69 gene transcription [148,164]. The activation and nuclear translocation of NF-AT allow it to bind to its specific sequence motif in the CD69 gene promoter region [164]. However, NF-AT appeared to play a less significant role than AP-1 and N F - K B in inducing CD69 expression as demonstrated by the lack of CD69 blockade in stimulated cultures by CsA in this study and in others [236]. PHA also activates p21 r a s, an oncoprotein capable of removing the inhibitory control on N F - K B [84] and inducing the expression of c-fos and c-jun through its downstream kinases p74raf"' and E R K (Fig. 1.4 and [159]). Indeed, the p21 r a s and p74 r a f _ 1 are critical kinases involved in CD69 expression, and the enhancement of c-jun expression and AP-1 binding inherent to PHA stimulation [80] is also responsible for the induction of the early marker. Corresponding to these findings is the presence of N F - K B and AP-1 binding sites in the promoter region of the CD69 gene [148,164]. 124 4.3 Kinetics of CD69 Expression During Polyclonal T-cell Activation In agreement with other investigations [80,114,137-138], the CD69 receptor was signifi-cantly upregulated after 4 hrs of PHA activation. Some studies suggested that its expression is maximal at 18 to 48 hrs [114,129], with most indicating 24 hrs [137,225,237]. This study confirmed that, based on the incubation times investigated, the highest number of CD69-expressing CD4 + and CD8+T-cells along with the strongest antibody staining occurred after 24 hrs of PHA activation. This time-course might reflect the contribution of CD69 in activated T-cells for enhancing IL-2 gene expression and overcoming phase barriers during cell cycle progression (e.g. Gi/S checkpoint) to achieve D N A synthesis [166-168]. The decline in CD69 expression after this 24-hour peak was also demonstrated in other kinetics studies [114,225,237]. In this investigation, the signals in both T-cell subsets fell to low levels by 96 to 120 hrs. The downregulation half-life was approximately 48 hrs, which was slightly longer than the reported value of 24 hrs [129]. The discrepancy might originate from the deliberate removal of stimuli from cultures by other research groups since the PBMCs in this study were incubated in the continuous presence of PHA. This expression pattern suggested that CD69 might not be involved in the later phases of T-cell development, when adequate IL-2 synthesis is achieved by other activation elements. In addition, the growth of lymphoblasts and their progeny is sustained by IL-2 signalling through its high-, affinity receptor, of which the a-chain (CD25) was strongly detected in almost all T-cells (> 92%) after 120 hrs, and by signalling proteins like OX40, whose expression was maintained in CD4 +T-cells. Phenotypic analysis was not performed beyond this time since the results would likely be complicated by cell death. A study by Iannone et al. [173] has shown that 125 the CD69 levels in PHA-stimulated cells remain low after 2 to 3 weeks of culturing. The reduction in CD69 expression is controlled by mRNA instability [149] and possibly also by proteolysis and protein recycling. Cell death was probably not responsible for the sharp decline since only a minor decrease in viable cells occurred in stimulated cultures. PHA causes activation-induced cell death (AICD) according to some studies, but this phenomenon is restricted to thymocytes, T-cell clones, and long-term (> 10 days) cultures [238-240]. The loss of surface targets of PHA also did not appear to be the causative factor of the reduction. First, the level of CD3+T-cells was consistent throughout culturing as assessed by flow cytometry. Secondly, although this study did not examine CD45 and CD2 expression during stimulation, other groups have indicated that activated lymphocytes did not exhibit modulated levels of these proteins [220,241]. Ijichi et al. [213] observed that PHA-mediated P B M C proliferation started to decline after 4 days of culturing and postulated that this was caused by the diminishing proportion of accessory cells. These cells are essential in provid-ing a surface matrix for PHA to stimulate T-cells. Because the decline in CD69 signal occurred as early as 48 hrs, accessory cell depletion was unlikely the cause of this reduction. A bi-directional dose relationship was observed between PHA and CD69 induction. From 0.2 to 3.0 p.g/ml, the extent by which CD4 + and CD8+T-cells upregulated CD69 after 24 hrs of stimulation correlated well with the D N A synthetic ability of PBMCs after 92 hrs of stimulation. Understandably, increasing mitogen concentrations might allow greater binding and crosslinking of surface proteins, thus increasing the cytosolic and nuclear activities that mediate CD69 expression. As suggested by others [114,137-138,236], CD69 induction 126 might provide a rapid, non-radioactive, subset-specific alternative to [ H]Thd uptake for assessing lymphocyte response in cultures. A PHA concentration of 5.0 ug/ml reduced the peak CD69 response, however. This effect was not a result of AICD because cell viability showed minimal reduction at 24 hrs (1.0 ug/ml = 93.1%, 3.0 ng/ml = 82.9%, and 5.0 ug/ml = 83.0%). Cell proliferation was significantly increased, suggesting the existence of inhibitory pathways that suppressed the expression of CD69 while leaving D N A synthesis intact. The current understanding of nucleotide metabolism maintains that lymphocytes rely on de novo enzymatic pathways that utilize uracil to produce deoxy-thymidine 5'-triphosphate (dTTP), a D N A nucleotide. The salvage pathway plays a less significant role and it depends on available thymidine such as [3H]Thd to generate dTTP [242]. The synthetic rate ofde novo pathways might be saturated at 3.0 (J.g/ml, and the additional SI increase at 5.0 ug/ml might be attributed to the salvage pathway utilizing exogenous thymidine to make new DNA. 4.4 Different Lectins Induced Comparable Activation Phenotypes ConA, another mitogenic stimulus, was used to determine whether the kinetics of CD69 expression differ according to the phytolectin employed. Several investigations have already shown that CD69 can be induced using this mitogen [137-138,237,242], and in this study, similar patterns of expression were observed between PHA and ConA. ConA was equally capable in upregulating CD69 levels by 4 hrs of incubation, and both lectins produced its peak expression in CD4 + and CD8 +T-cell subsets after 24 hrs of culturing with the subsequent characteristic decline. The levels of CD69 expression and cellular proliferation 127 induced by ConA, however, were lower than those induced by PHA. Similar results have been reported elsewhere [137-138,237,242]. While the respective doses used might be responsible for the difference, the alternative explanations are that ConA might have a less stimulating effect on T-cells and it might signal through pathways different from those of PHA. Although Con A is capable of binding more surface glycoproteins than PHA, its interactions with CD3 and CD45 do not appear to contribute to cell activation [211,220]. Although lectins such as PHA and ConA interact with various membrane proteins involved in the immunological functions of T-cells (e.g. TCR, CD3, CD45), their major drawback is the lack of target protein specificity. To examine the effect of a more specific stimulus, immobilized OKT3 at 30 ng/ml was employed as a trigger. This mitogen was less effective than either PHA or ConA for inducing T-cell activation, as indicated by the low CD69 expression at 24 and 72 hrs and the minimal P B M C proliferation. PHA was therefore selected as the stimulus of choice, since it has the advantages of being easy to handle, inexpensive and reliable in generating reproducible results. 4.5 Kinetics of OX40 and CD25 Surface Expression In this study, CD69, OX40, and CD25 were respectively labelled as early, intermediate, and late activation markers according to their expression kinetics in PHA-stimulated T-cells. Examining the expression of these molecules might increase the understanding of their functional contributions during T-cell development from quiescence to proliferation and identify the various stages of immune disease progression by quantifying the proportions of cells expressing them [115,137,189,237]. 128 As confirmed in other reports [191,196], the peripheral T-cells of healthy individuals were OX40". The lack of expression might serve to avoid non-specific costimulation or T-cell attachment to the vascular endothelium (via gp34), which they can otherwise gain entry into the interstitial tissues to mediate inflammatory reactions. Contrary to the rapid appearance of CD69, OX40 did not appear on the cell surface until 12 hrs after stimulation. The mechanisms accounting for this lag between the appearance of the two markers in activated T-cells are unclear since the signalling components controlling OX40 gene transcription have not been elucidated. This observation, however, supported the hypothesis that cellular activation precedes lymphocyte-endothelial interaction [192]. OX40 was therefore considered to be involved in the intermediate events during T-cell activation. OX40 expression in CD4+T-cells reached a maximum level between 24 and 48 hrs, when the proportions of CD69 + and OX40 + cells and the surface densities of these molecules were comparable. The proportion of OX40 +CD4 +T-cells subsequently declined, but approxi-mately half of them remained positive for this marker after Day 5, in contrast to the rapid disappearance of CD69. A similar kinetics pattern was also demonstrated in PHA-activated PBMC by Imura et al. [191] using a different anti-OX40 MAb. These findings indicated that while CD69 was removed from the cell surface after the initial activation phases, the adhesion and costimulatory properties of OX40 remained essential in the later stages. CD69 appeared to have a more important role in T-cell activation than OX40 based on two principal observations. First, the peak number of OX40 +CD4 +T-cells (67.4 ± 10.9%) never reached the level attained by that of CD69 +CD4 +T-cells (84.9 ± 2.6%) throughout the 5-day 129 activation period. This observation suggested that certain factions of activated, CD69-expressing CD4+T-cells did not express OX40. These cells might undergo activation and use adhesion molecules other than OX40 to attach to the vascular endothelium. Secondly, as indicated in this study and also in other reports [191,193,198], OX40 expression was restricted to a very small proportion of PHA-stimulated CD8+T-cells while CD69 was upregulated in most of these cells. OX40 has also been found to be expressed in CD4 + T-blastocytes and CD4 +CD8" adult T-leukaemic cells [191,198]. Altogether these findings lead to the conclusion that OX40 is probably not involved in the costimulation and endothelial binding of CD8 +T-cells. The small increase in OX40 expression in this subset at 24 hrs might simply represent an effect of polyclonal activation. This subset-restricted expression also accounted for the lower OX40 level in the CD3 +T-cell population as a whole. CD25 expression was found in less than 16% of resting CD3+T-cells, in agreement with other studies [137,141,225]. This result contrasted with the negligible baseline levels that were observed with the other two activation markers studied. It is normally considered that the restricted expression of the high-affinity IL-2R in naive cells serves to limit the antigen non-specific proliferation of peripheral lymphocytes in response to paracrine IL-2 secretion. While basal stimulation via this receptor may be necessary to maintain the viability of this naive population, it is possible that the CD25 + cells detected might represent the memory cells from previous immune stimulation. Basal CD25 expression was lower in peripheral CD8+T-cells than in CD4+T-cells, and whether this reflected a more stringent growth control mechanism on cytotoxic T-cells or differential memory cell survival remains to be determined. 130 The CD25 expression was significantly reduced during the early phase (at 4 hrs) of mitogen activation and then increased from 24 hrs. It is tempting to speculate that the reduction might temporarily arrest the later elements in cell activation and provide a brief window within which anergy can be established. The de novo expression of CD25 is mediated by the PHA-induced activation of N F - K B [210]. While CD69 disappeared rapidly from cell membrane after 3 to 5 days, more than 90% of CD4 + and CD8+T-cells remained positive for the late marker. This is consistent with the entry of cells into their proliferation phase during which their growth and replication are sustained by IL-2 signalling. The proliferation signals generated by the binding of IL-2 to its high-affinity receptor enhance PTK activity and promote transcription factor (e.g. STAT) and cyclin expression [88-89]. The progressive switch from early to late activation phenotypes and the maintenance of CD25 expression for over a week of continuous culturing have been demonstrated in previous reports [141,173]. 4.6 Inhibition of Proliferation by ISDs Failed to Abrogate CD69 Expression Since the discovery of cyclosporine, much research emphasis [78] has been placed on its ability to interact with cyclophilin, suppress NF-AT activation, inhibit IL-2 production and prevent allograft rejection. In this study, its influence on mitogen-induced CD69 upregulation in normal T-cells was examined by using it at concentrations equivalent to the plasma trough levels (~ 175 to 500 ng/ml) observed in RTx subjects in this study and patients studied elsewhere [243]. As shown in Table 3.5, CsA at these concentrations reduced only slightly the proportion of T-cells which were CD69 + . This drug was only marginally effective at the highest dose tested (800 ng/ml caused a 14% and 17% reduction in the proportion of CD69 + CD4 + and CD8+T-cells, respectively), although the surface 131 density appeared to be more sensitive to inhibition. Previous reports have shown that the 50% inhibitory dose (ID50) of CsA on CD69 expression is approximately 20,000 ng/ml (25x more than the maximal amount used in this study) while lower concentrations are incapable of interfering with its surface upregulation [243]. The lack of inhibition of CD69 expression in this study was not a result of reduced pharmacological activity of the drug since D N A synthesis was inhibited in a normal dose-dependent manner. CsA has been shown to inhibit NF-AT activation while allowing the activities of other transcription factors such as AP-1 and N F - K B to remain intact [78]. These non-inhibited nuclear mediators might be sufficient to maintain optimal CD69 gene expression when T-cell growth was suppressed by the lack of activated NF-AT and IL-2 synthesis [148,164]. In addition, a study by Feske et al. [244] indicated that while NF-AT, but not N F - K B and AP-1, is defective in D N A binding in individuals with severe combined immunodeficiency, CD69 expression is normally induced. Tacrolimus operates at the same molecular locus as CsA to inhibit calcineurin-mediated dephosphorylation of NF-AT. However, it binds to a distinct cytosolic receptor (FKBP) and is able to modulate T-cell activation at concentrations lower than those of CsA [100-101,104]. The results reported here revealed that FK506 inhibited CD69 in a dose-dependent manner beginning at a concentration of 2.5 ng/ml, with an ID50 close to 20.0 ng/ml in both CD4 + and CD8 +T-cell subsets. Unlike CsA, FK506 might be sufficiently potent to suppress all NF-AT activity [77], thereby influencing CD69 expression. Alternatively, it might inter-fere with other transduction cascades and transcription factors (e.g. c-fos and N F - K B ) that 132 are critical in CD69 expression [245,246]. Whether the inhibition of CD69 contributes to the in vivo effects of tacrolimus in preventing transplant rejection remains to be elucidated. Long-term (72-hr) stimulated T-cells significantly increased their CD69 expression when cultured with CsA but not with FK506. The mechanism accounting for this upregulation is unclear. Because cyclosporine did not completely block T-cell proliferation (< 70% in-hibition at the highest dose tested), it is possible that a low level of NF-AT activation was achieved through continued PHA stimulation, thus leading to the progressive accumulation of active transcription factors within the nucleus and resulting in the prolongation of CD69 surface expression. In this situation, costimulation through this receptor might represent a possible mechanism for acute rejection to occur in transplant patients that are under cyclosporine immunosuppression [186]. MP suppressed D N A synthesis but failed to reduce the proportion of CD69+T-cells following activation. The mechanism responsible for this bi-directional response is unknown. Other investigators have revealed that this drug and other corticosteroids bind to their cytosolic receptors and inhibit T-cell proliferation by directly interacting with the CRE regions of growth-promoting genes and modulating their expression [111,247,248]. It is possible that the CD69 gene promoter lacks these sequence motifs, thus allowing this marker to be expressed when other T-cell activities (e.g. proliferation) were suppressed by MP. Glucocorticords also block proliferation by antagonizing the synergistic co-operation of NF-A T and AP-1 binding sites in the IL-2 enhancer region and by inducing the synthesis of I K B , the inhibitor of N F - K B [112,249]. AP-1 and N F - K B are crucial transcription activators of 133 the CD69 gene [197,201]. It is unclear how CD69 expression was achieved when the activity of these nuclear factors was blocked. Whether CD69 expression can be activated by transcription factors that are not inhibited by MP remains to be determined. This study also demonstrated that the epitope densities, but not the proportions of CD69-positive cells, were susceptible to MP suppression. It remains unclear whether the steroid-mediated suppression of protein synthesis [250] was responsible for this inhibition. In addition, the drug enhanced the levels of this marker in 72-hr-activated lymphocytes, a phenomenon that was also observed with cyclosporine-treated cells. The mechanism underlying this increase in expression remains to be elucidated. These data collectively implied that CsA and MP were individually unable to block CD69 induction effectively. It is unknown if these agents also fail to suppress the expression of CD69 induced by allospecific activation in vivo [44,89,105,111-112], thus permitting T-cells to mediate graft rejection. However, both CD69 induction and cellular proliferation might be inhibited when these drugs were used in combination. Indeed, this study showed that CD69 expression was significantly reduced in new allograft patients in their early post-transplant periods, when high-dose ISDs in triple-drug regimens were administered. 4.7 Whole Blood Technique for Measuring Activation Marker Expression Erythrolysis has replaced PBMC isolation as the standard preparatory technique for flow cytometric immunophenotyping in clinical and research investigations [229,251,252]. Whole blood analysis reduces specimen manipulation, processing time, and the quantity of blood needed for evaluation. The "stain-lyse" protocol was adopted in this study for assess-134 ing lymphocyte surface phenotypes. Resting or activated lymphocytes were first incubated with specific antibodies and then subject to erythrolysis using the lysing reagent described (46 ± 1 mOsm and pH 3.34-3.43 vs. physiological: 287 mOsm and pH 7.4). According to Ashmore et al. [229], this "stain-lyse" method is superior to both P B M C isolation and the alternative "lyse-stain" technique in the detection of activated T-cells. One draw-back of the whole blood method was the reduction in the number of leukocytes available for analysis, compared with the selective lymphocyte enrichment and milder conditions in P B M C isolation which provided more CD3 T-cells to study (89.5 ± 1.9% vs. whole blood: 66.0 ± 3.1%). The CD4/CD8 ratios and subset compositions were similar in both methods. Although the in vivo microenvironment is difficult to reproduce under laboratory conditions, the whole blood method offers a close approximation to the peripheral circulation by retaining most of the cellular and soluble components in blood during culturing. This is particularly important in examining the circulating lymphocytes of ESRD and RTx subjects since uraemic toxins, ISDs and other substances peculiar to these patients and that might influence T-cell activation remained in the cultures. For instance, the average CsA concentration in the whole blood samples of ST subjects was 968 ± 225 ng/ml (as determined by the TDx assay), and most of it was removed by the P B M C isolation procedure (only 14 ± 1 ng/ml remained). Many studies have also maintained that the whole blood technique is suitable for analysing the surface phenotype of activated T-cells [229,253-255]. Whole blood CD69 induction assays have been used by others to investigate the early activation response of T-cells in both health and disease [189,236,253-254]. The erythrolytic 135 technique, similar to the mononuclear cell isolation method, demonstrated that resting T-cells from healthy controls were CD69 l 0 W When peripheral blood was cultured with 1.0 ug/ml PHA for 24 hrs, CD69 expression was upregulated in both CD4 + and CD8 +T-cell subsets. However, the mean percentages of cells positive for this marker in the whole blood cultures were less than those observed in the PBMC cultures that were run in parallel (CD4 +: 44.4% less; CD8 : 41.2% less). A delay in kinetics was unlikely the cause since the 72-hr whole blood cultures revealed a significant decline after the peak response at 24 hrs. Insufficient mitogen concentrations might explain the discrepancy between the whole blood and P B M C cultures, since the percentage differences of CD69+cells between the two methods were similar in both T-cell subsets. The actual amount of PHA that interacted with T-cells in whole blood cultures might be lower than that in P B M C cultures. This is due to the absorption of PHA by non-T-cells such as erythrocytes (which bind to the PHA-E subunit [206,207]), polymorphonuclear leukocytes and platelets which are otherwise removed during mononuclear cell isolation. Therefore, increasing the PHA dose to above 1.0 ug/ml might enhance the peak CD69 expression with the whole blood method. The hypo-tonic shock might also destroy a fraction of activated cells due to their lowered membrane integrity. An alternative explanation to the lower CD69 expression with the whole blood method was that the plasma components and cells present in whole blood might control (e.g. through cytokines or surface proteins) the levels of activation marker expression in T-cells. The OX40 molecule was expressed at very low levels in resting cells according to the whole blood analysis. This observation reinforced the idea that circulating T cells in healthy 136 individuals are quiescent and do not require the participation of costimulatory molecules such as CD69 and OX40. OX40 was induced after 24 hrs of PHA stimulation, and the proportion of positive cells was lower than that observed in stimulated PBMCs; inadequate stimuli in the whole blood cultures might again account for the difference. In addition, the mean percentage of OX40 +CD4 +T-cells was consistently lower than that of CD69 + CD4 + T-cells, which confirmed the in vitro results and indicated that certain fractions of CD4+T-cells did not upregulate this protein during polyclonal activation. As in experiments with isolated PBMCs, stimulation of whole blood samples failed to increase OX40 expression in CD8+T-cells substantially, suggesting that the functional importance of this molecule in this subset was less than that in CD4+T-cells. 4.8 Immune Depression and Stimulation in Dialysis-treated ESRD Patients There is abundant clinical and biological evidence of defective cellular immunity in ESRD [22-24,27-31,35-36,256-258]. The pathological changes associated with renal failure contribute to some of the immune impairments observed (Fig. 1.1). Although maintenance dialysis corrects some of the physiological problems, it fails to restore immune competence. In some studies, dialysis-treated patients exhibited both signs of immune suppression and abnormal activation [22,27,31]. For example, the peripheral lymphocytes of dialyzed subjects have been shown to be defective in PHA-induced proliferation and IL-2 synthesis but express abnormally high basal CD25 levels [22]. Because studies examining early T-cell activation events in these patients are lacking, this investigation evaluated the CD69 and OX40 expression in their peripheral T-cells. 137 The specimens of haemodialyzed subjects were drawn before the commencement of dialysis to avoid complement activation [30]. No abnormalities were observed in their CD3 counts or subset ratios, which contrasted with other studies [28,30,259] that reported ^lymphocyto-penia in this type of patients with a progressive decline in both of their CD4 + and CD8 + T-cell levels. The HD subjects in this study had undergone therapy for an average of 4.8 years, and they were not intended to reflect the patient population with exceptionally long-term therapy who might be expected to show more profound immunodeficiency. Similar to other reports [28,260], patients in this study demonstrated normal CD4/CD8 ratios. The CD69 and OX40 expression in resting and PHA-activated T-cells of HD subjects were similar to those of healthy controls. These results contrasted with those of an earlier study [30] which demonstrated that the baseline profiles of several activation markers are abnormal in this type of patients. Thus, this investigation revealed that the early phenotypic changes during T-cell activation appeared normal in haemodialyzed patients. The immunologic analysis was performed before the onset of routine HD treatment, and the significance of any changes in CD69 and OX40 expression after each dialysis session remains to be determined. In agreement with previous reports [24,25], CAPD therapy did not alter the level of circulating CD3+T-cells. Subset analysis, however, revealed an abnormally high CD4/CD8 ratio that resulted from an expanded CD4 population (increased in percentage) and a significantly diminished CD8 subset (decreased in percentage). These observations were in direct contrast with those made by Palop et al. [24], who demonstrated that the proportion of 138 CD8+T-cells in CAPD subjects was markedly increased while their CD4 +T-cell levels remained stable. However, the results in this study agreed with those obtained by Lewis et al. [29], who suggested that the increase in CD4+T-cells might represent a host response to the bacterial pathogens that are constantly introduced into the peritoneal cavity. CD4+helper T-cells are responsible for co-ordinating the immune defence against bacterial infection. Therefore, the increase in the proportion of circulating CD4+T-cells in CAPD subjects might be due to the high incidence of bacterial peritonitis in this group of patients. The immune system in this type of patients is depressed in many aspects. Enhancing the activation state of peripheral T-cells might be a mechanism to alleviate this depression and to counteract the loss of lymphocytes residing in the peritoneal cavity due to dialysis. This study demonstrated that the resting CD8+T-cells of these patients expressed an elevated CD69 level. Previous investigations also showed that other markers of activated cells (e.g. CD25) are abnormally upregulated in dialysis patients [29,261,262]. These observations together suggested that CAPD patients exhibit a chronic low level of lymphocyte activation. CD69 expression in the CD8+T-cells of CAPD patients increased normally following activation, despite the elevated resting level. On the contrary, CD69 expression in the PHA-stimulated CD4+T-cells of the same patients failed to increase to the levels attained by those cells of the normal subjects. This impairment in expression might be caused by uraemia, insufficient dialysis or other factors yet to be determined [22,23,25,257]. The suboptimal ability of CD4+T-cells to express CD69 and other activation markers might be one of the factors that lead to the increased susceptibility of CAPD patients to bacterial peritonitis. 139 Baseline OX40 expression in the CD4+T-cells of CAPD subjects was significantly higher than that of normal controls. It is possible that this increase in expression was due to the constant exposure of lymphocytes to bacteria and other pathogens introduced through the dialysis process [1,8,60,67]. Collectively, these data suggested that the expression of some lymphocyte activation markers was deregulated in CAPD-treated ESRD patients. These dialysis subjects demonstrated the simultaneous signs of chronic stimulation (e.g. elevated resting levels of CD69 +CD8 +T-cells) and hyporesponsiveness (e.g. depressed CD69 expression in PHA-stimulated CD4+T-cells). The expression of other activation markers (e.g. CD25 and H L A -DR) was also shown to be abnormal in CAPD subjects in other studies. The physiological perturbations caused by renal failure and the dialysis treatment were often cited as the sources of these abnormalities [22,25,31]. A clearer understanding of how renal disease and dialysis interfere with T-cell activation marker expression might assist in modifying dialysis therapies in order to maintain optimal host immune defence. 4.9 Expression of Activation Molecules in Stable Transplant Recipients Kidney function returns towards normal in stable graft recipients, and the persisting alterations in cell-mediated immunity reflect predominantly the effects of the immuno-suppressants used to prevent rejection [60]. Patients in this study were receiving conventional treatment with CsA, prednisone and Aza. The analysis of circulating lymphocytes of ST subjects revealed normal CD4 and CD8 subset distributions but abnormally high CD3 +T-cell levels. Whether this was related to continuous 140 allogeneic stimulation is not known. The resting CD4+T-cells of stable transplant patients exhibited normal CD69 expression. The PHA-induced CD69 expression was optimal despite their systemic immunosuppression. This observation agreed with the results from the in vitro experiments which showed that CsA and MP failed to suppress CD69 expression effectively. In contrast to the CD4 subset, the expression of this early marker was elevated in resting CD8+T-cells. It remained unclear whether they represented allospecific T-cells that achieved partial activation. This finding provided the first evidence that circulating T-cells in stable renal allograft recipients exhibited a chronically low level of activation. Other studies have demonstrated that low-grade allogeneic response occurred persistently for several years after successful RTx [65,263]. In addition, graft-infiltrating CD8+T-cells have been shown to express CD69 strongly during acute rejection [185-186]. The possible contribution of these peripheral CD69 +CD8 +T-cells in latent acute or chronic rejection warranted further investigations. Despite high baseline levels, this subset upregulated CD69 expression normally. Resting T-cells from these patients were OX40", and the upregulation of this marker upon stimulation was slightly less than that in healthy controls. This observation suggested that intermediate activation events, such as OX40 expression, might be impaired due to the effects of ISDs. Early events, like CD69 induction, were unaffected. The inhibition of OX40 upregulation might also reduce the possibility of endothelial adhesion by activated T-cells, thus limiting the migration of these cells into the transplant interstitum during acute rejection [109-110,185-186]. 141 4.10 Lymphocyte Activation Marker Expression in Early Post-transplant Period Previous studies have examined several peripheral immune parameters that might assist in diagnosing rejection [18,19,21,51-52,55,57-58,60,63,68]. The analysis of CD69 expression was investigated as a rapid and simple method for detecting the early presence of activated T-cells in newly transplanted patients that might indicate incipient allograft rejection [66,264] or infection. Although approximately half of the subjects studied were expected to experience graft rejection during the first three months after transplantation [8,49], only one patient underwent a single mild rejection episode. This may reflect both the small cohort analyzed (n = 10) and the high effectiveness of modern immunosuppression. Most of these subjects also participated in clinical trials involving newly developed immunosuppressants (e.g. tacrolimus and MMP) that might be more potent in suppressing alloreactivity than the conventional agents (e.g. CsA and prednisone). The expression of CD69 and OX40 was normal both 7 days before and three days after the event in that single patient who experienced rejection, although their expression was not measured during rejection. More frequent monitoring of peripheral blood or the aspirates of graft-infiltrating cells might provide more accurate information on the relationship between activation marker expression and acute rejection [68]. The pre-transplant lymphocytes of NT subjects exhibited several unusual features. The high percentage of circulating CD3+T-cells before transplantation was the most evident, and the mechanism responsible remains unclear. The reduction in peripheral CD3+T-cells during the 142 initial 2 to 3 weeks after transplantation might be a result of surgical trauma [265] combined with the effects of ISDs. For example, high-dose prednisone, which was administered within the first post-transplant week (Table 3.12), has been shown to cause lymphocyte apoptosis and redistribution [109,110]. Other agents such as Aza suppress the bone marrow production of leukocytes, and the anti-rejection antibodies (OKT3 and Atgam), which were administered in several NT patients, also deplete circulating lymphocytes rapidly [44, 117,120]. The recovery observed later in the monitoring period was likely due to the tapering of ISDs doses and recuperation from the transplant operation. Lymphocyte subsets and their relative ratios before RTx were similar to those of normal controls. CD8 +T-cell levels were significantly reduced on several occasions during the follow-up period. These observations contrasted with the findings of another longitudinal study [266] which showed that allograft recipients in the initial post-transplant months exhibit gradually decreasing levels of CD4+T-cells but an expanding CD8 +T-cell subset. The mechanism responsible for the reduction in CD8+T-cells observed in this study is unknown. Lymphodepleting agents such as OKT3 and Atgam do not show subset preference in removing circulating T-cells [117], and high-dose steroid therapy induced more profound apoptosis in CD4+T-cells than in CD8+T-cells [109]. Flow cytometry has been used in other studies to examine the increase in activated cells expressing markers such as CD25 and H L A - D R during acute rejection [12,20,55,60]. Consistent with another report [65] that demonstrated negligible CD69 expression during transplant quiescence, the baseline CD69 levels in both circulating T-cell subsets of NT 143 subjects remained low throughout the period of observation. The lack of significant expression was consistent with the absence of rejection in most of these patients; whether changes in CD69 expression occur during acute rejection remains to be determined. During the first two post-transplant weeks, lymphocyte activation was impaired as indicated by the reduced proportions and surface densities of CD69 + CD4 + and CD69 +CD8 +T-cells. Immunosuppression probably account for these defects. ISDs were administered at high doses during the initial weeks of RTx, and several patients received antilymphocyte antibodies (Atgam or OKT3) that were shown to suppress T-cell activity [113,121-122]. The specimens of NT subjects were obtained 2 to 3 hrs after the administration of ISDs, at which time the drug concentrations in these blood samples are normally very high (e.g. samples from ST patients 2-3 hrs post-dosing had an average CsA level of 968 ± 225 ng/ml). High doses of CsA produced a partial inhibition of CD69 expression as shown in the in vitro part of this study and others [236]. The reduction in CD69 surface density might also be mediated by prednisone, since MFI values appeared sensitive to MP inhibition. The impairment of CD69 expression after stimulation might represent another anti-rejection mechanism of these ISDs, since T-cells infiltrating renal allografts during acute rejection were found to express this marker [53,185,187]. The analysis of OX40 expression in the CD4+T-cells of NT patients provided further insight into the immune depression in these subjects. Resting T-cells from their peripheral blood expressed normal OX40 levels. This finding agreed with other reports which showed that quiescent T-cells are OX40" and T-cells present in rejected allografts of animal models and 144 in human G V H D upregulate this marker [199-200]. OX40 induction was impaired during the initial post-transplant weeks and continued at levels lower than the pre-transplant value. Similar non-significant impairment was also observed in ST subjects, suggesting that the transcriptional control of OX40 gene was more sensitive to the ISDs than that of CD69 gene. The blockade of IL-2 gene transcription by immunosuppressive therapy and the surgical trauma might be factors accounting for the reduction in PHA-induced blastogenesis during the early post-transplant months. In addition, the decrease in activation marker expression was also probably responsible for the reduction in proliferation since D N A synthesis and CD69 upregulation were positively correlated in these patients. As the dosages of ISDs were reduced in these patients (Table 3.12), the CD69 induction and D N A synthesis returned to their preoperative levels. At the end of the monitoring program, the CD69 levels were stable and similar to those of ST patients, indicating that the steady-state levels of activation marker expression in transplant patients might already be achieved after the first few weeks of transplantation. 145 Chapter Five Summary This study documented the unique expression kinetics of three signal-transmitting receptors, namely CD69, OX40 and CD25, in T-cells during mitogen stimulation. Several abnormalities in the surface expression of these activation markers were observed in the peripheral T-cells of ESRD patients treated with dialysis or kidney transplantation. CD69 was expressed early during the process of lymphocyte activation. Its expression reached a maximum level at 24 hours and declined rapidly thereafter. This early appearance is consistent with the possibility that CD69 plays an integral costimulatory role in the initial stages of lymphocyte activation, and its expression is down-regulated at later stages when this costimulatory function is no longer needed. In addition, the expression of CD69 preceded that of OX40. Maximal OX40 expression occurred around 48 hrs, suggesting that the activation of CD4+T-cells must be established before OX40-mediated endothelial adhesion can occur. The sharp decline in CD69 expression after peak response was not observed in OX40 and CD25 expression kinetics, suggesting that their functions remain essential in the later stages. Indeed, the IL-2R co-chain assists in the autocrine growth of proliferating T-cells, and this might account for CD25 expression being maintained at a strong level in long-term activated T-cells. PHA and ConA were comparable in their ability to trigger CD69 expression, although their mechanisms of triggering T-cell activation might not be identical. Signalling pathways that control CD69 induction and D N A synthesis were dissimilar in two aspects. First, increasing PHA concentrations induced a bi-directional response, in which higher doses decreased the 146 CD69 induction but enhanced proliferation. Secondly, ISDs inhibited PHA-induced lymphocyte proliferation at low doses, while CD69 expression was only partially suppressed at even high concentrations of the same agents. CsA and FK506 differed in their potency in suppressing CD69 expression but were similar in inhibiting D N A synthesis. This suggested that tacrolimus inhibits a broader range of early activation events than does CsA, thus providing an additional mechanism for tacrolimus to prevent and reverse acute rejection. The combination of these drugs with other ISDs was capable of suppressing CD69 induction as demonstrated by the reduction observed in the new allograft recipients. Whole blood measurement of CD69 expression yielded results similar to those with the P B M C technique. Although the cells in stimulated whole blood did not express the activation markers at the same levels as those in stimulated PBMCs, the expression kinetics was equivalent. The whole blood method was simple and rapid, with the added advantage of preserving the cellular and plasma compositions peculiar to the peripheral blood of the patient subjects. The immunological variables were normal in haemodialyzed patients; however, an abnormally high CD4/CD8 ratio, an expanded CD4 subset, a lower CD8 count, an elevated baseline expression of CD69 in CD8 T-cells and OX40 in CD4T-cells, and an impaired CD69 induction in CD4+T-cells were evident in CAPD subjects. These mixed signs of immune activation and deficiency might represent the immunologic perturbations caused by uraemia and dialysis, as suggested by other studies. Stable transplant recipients were similar 147 to healthy individuals in all the variables except for their increased levels of circulating CD3+T-cells. This study provided the first line of evidence that profound in vivo immunosuppression within the first few post-transplant weeks impaired CD69 and OX40 induction in T-cells. Taken together, the results indicated that the resting, peripheral blood T-cells in RTx patients undergoing a quiescent early post-transplant course did not express elevated levels of CD69. The reduction in PHA-induced CD69 expression was likely due to either the inhibition of signalling pathways by clinical immunosuppression or surgical operation or both. 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