Novel Immunological Aspects of CMV-Related Diseases: Pathogenesis, Diagnosis and Therapy (Intervirology, 5-6)

Novel Immunological Aspects of CMV-Related Diseases Pathogenesis, Diagnosis, and Therapy Proceedings of the 2nd Symposiu...

Author: C. A. Bruggeman | Hans Wilhelm Doerr | A. Ramon | M. Scholz

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Novel Immunological Aspects of CMV-Related Diseases Pathogenesis, Diagnosis, and Therapy Proceedings of the 2nd Symposium on CMV-Related Immunopathology September 17–18, 1999, Maastricht, The Netherlands

Guest Editors

Cathrien A. Bruggeman, Maastricht Hans Wilhelm Doerr, Frankfurt/Main Albert Ramon, Maastricht Martin Scholz, Frankfurt/Main

46 figures, 5 in color, and 22 tables, 2000

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Acknowledgements

The organizers of the 2nd Symposium on CMV-Related Immunopathology want to thank the following sponsors. Without their financial contribution the organization of this symposium would not have been possible. Main Sponsors Abbott Diagnostics, The Netherlands Roche Diagnostics, Belgium Roche Pharma, Belgium Roche Diagnostics, Germany Roche Diagnostics, The Netherlands Co-Sponsors Bayer Diagnostics Pharma, The Netherlands Beckman Coulter, The Netherlands Dépex, The Netherlands Meridian/Gull Laboratories, The Netherlands University Hospital Maastricht (azM), The Netherlands

S. Karger Medical and Scientific Publishers Basel N Freiburg N Paris N London New York N New Delhi N Bangkok Singapore N Tokyo N Sydney

Drug Dosage The authors and the publisher have exerted every effort to ensure that drug selection and dosage set forth in this text are in accord with current recommendations and practice at the time of publication. However, in view of ongoing research, changes in government regulations, and the constant flow of information relating to drug therapy and drug reactions, the reader is urged to check the package insert for each drug for any change in indications and dosage and for added warnings and precautions. This is particularly important when the recommended agent is a new and/or infrequently employed drug.

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All rights reserved. No part of this publication may be translated into other languages, reproduced or utilized in any form or by any means, electronic or mechanical, including photocopying, recording, microcopying, or by any information storage and retrieval system, without permission in writing from the publisher or, in the case of photocopying, direct payment of a specified fee to the Copyright Clearance Center (see ‘General Information’). © Copyright 2000 by S. Karger AG, P.O. Box, CH–4009 Basel (Switzerland) Printed in Switzerland on acid-free paper by Reinhardt Druck, Basel ISBN 3–8055–7049–X

Contents Vol. 42, No. 5–6, 1999

274 Acknowledgements 277 Preface Bruggeman, C.A.; Ramon, A. (Maastricht); Scholz, M. (Frankfurt am Main)

Part I. Immunopathology CMV Infection in the Immunocompromised Host 279 Time-Related Effects of Cytomegalovirus Infection on the Development of

Chronic Renal Allograft Rejection in a Rat Model Lautenschlager, I.; Soots, A.; Krogerus, L.; Inkinen, K.; Kloover, J.; Loginov, R.; Holma, K.; Kauppinen, H.; Bruggeman, C.; Ahonen, J. (Helsinki/Maastricht) 285 Overcoming the Problem of Cytomegalovirus Infection after Organ

Transplantation: Calling for Heracles? van Son, W.J.; de Maar, E.F.; van der Bij, W.; van den Berg, A.P.; Verschuuren, E.A.M.; The, T.H. (Groningen) 291 Clinical Significance of Cytomegalovirus-Specific T Helper Responses and

Cytokine Production in Lung Transplant Recipients Zeevi, A.; Spichty, K.; Banas, R.; Cai, J.; Donnenberg, V.S.; Donnenberg, A.D.; Ahmed, M.; Dauber, J.; Iacono, A.; Keenan, R.; Griffith, B. (Pittsburgh, Pa.)

Immune Escape and Reactivation 301 Human Cytomegalovirus Escape from Immune Detection Michelson, S. (Paris) 308 Human Cytomegalovirus Reactivation in Bone-Marrow-Derived

Granulocyte/Monocyte Progenitor Cells and Mature Monocytes Prösch, S.; Döcke, W.-D.; Reinke, P.; Volk, H.-D.; Krüger, D.H. (Berlin) 314 Human Cytomegalovirus Latency and Reactivation – A Delicate Balance

between the Virus and Its Host’s Immune System Söderberg-Nauclér, C. (Stockholm); Nelson, J.A. (Portland,Oreg.) 322 Measurement of Anti-Human Cytomegalovirus T Cell Reactivity in

Transplant Recipients and Its Potential Clinical Use: A Mini-Review Kern, F.; Faulhaber, N.; Khatamzas, E.; Frömmel, C.; Ewert, R.; Prösch, S.; Volk, H.-D.; Reinke, P. (Berlin)

CMV-Induced Pathomechanisms 325 Viral Inhibition of Interferon Signal Transduction Cebulla, C.M.; Miller, D.M.; Sedmak, D.D. (Columbus, Ohio) 331 Murine Cytomegalovirus Homologues of Cellular Immunomodulatory Genes Davis-Poynter, N.J. (Newmarket); Degli-Esposti, M. (Nedlands); Farrell, H.E. (Newmarket) 342 Molecular Mimicry by Cytomegaloviruses. Function of Cytomegalovirus-

Encoded Homologues of G Protein-Coupled Receptors, MHC Class I Heavy Chains and Chemokines Vink, C.; Beisser, P.S.; Bruggeman, C.A. (Maastricht)

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350 Cytomegalovirus-Induced Transendothelial Cell Migration. A Closer Look at

Intercellular Communication Mechanisms Scholz, M.; Blaheta, R.A.; Vogel, J.-U.; Doerr, H.W.; Cinatl, J., Jr. (Frankfurt am Main) 357 Altered Expression of Extracellular Matrix in Human-Cytomegalovirus-

Infected Cells and a Human Artery Organ Culture Model to Study Its Biological Relevance Schaarschmidt, P.; Reinhardt, B.; Michel, D.; Vaida, B.; Mayr, K.; Lüske, A.; Baur, R.; Gschwend, J.; Kleinschmidt, K. (Ulm); Kountidis, M.; Wenderoth, U. (Heidenheim); Voisard, R.; Mertens, T. (Ulm) 365 Human Cytomegalovirus Infection of Immature Dendritic Cells and

Macrophages Jahn, G.; Stenglein, S.; Riegler, S.; Einsele, H.; Sinzger, C. (Tübingen)

Part II. Diagnostics and Antiviral Therapy Diagnostics 373 Diagnostic Value of Nucleic-Acid-Sequence-Based Amplification for the

Detection of Cytomegalovirus Infection in Renal and Liver Transplant Recipients Goossens, V.J.; Blok, M.J.; Christiaans, M.H.L.; van Hooff, J.P. (Maastricht); Sillekens, P. (Boxtel); Höckerstedt, K.; Lautenschlager, I. (Helsinki); Middeldorp, J.M. (Boxtel); Bruggeman, C.A. (Maastricht) 382 Towards Standardization of the Human Cytomegalovirus Antigenemia

Assay Verschuuren, E.A.M.; Harmsen, M.C.; Limburg, P.C.; van der Bij, W.; van den Berg, A.P.; Kas-Deelen, A.M.; Meedendorp, B.; van Son, W.J.; The, T.H.; The Biomed 2 Study Group (Groningen) 390 New Advances in the Diagnosis of Congenital Cytomegalovirus Infection Lazzarotto, T.; Varani, S.; Gabrielli, L.; Spezzacatena, P.; Landini, M.P. (Bologna) 398 Significance of Qualitative Polymerase Chain Reaction Combined with

Quantitation of Viral Load in the Diagnosis and Follow-Up of Cytomegalovirus Infection after Solid-Organ Transplantation Vanpoucke, H.; Van Vlem, B.; Vanholder, R.; Van Renterghem, L. (Gent) 405 Viral Dynamics during Active Cytomegalovirus Infection and Pathology Emery, V.C. (London)

Antivirals 412 Inhibition of Cytomegalovirus in vitro and in vivo by the Experimental

Immunosuppressive Agent Leflunomide Waldman, W.J.; Knight, D.A. (Columbus, Ohio); Blinder, L.; Shen, J.; Lurain, N.S. (Chicago, Ill.); Miller, D.M.; Sedmak, D.D. (Columbus, Ohio); Williams, J.W.; Chong, A.S.-F. (Chicago, Ill.) 419 Proinflammatory Potential of Cytomegalovirus Infection. Specific Inhibition

of Cytomegalovirus Immediate-Early Expression in Combination with Antioxidants as a Novel Treatment Strategy? Cinatl, J., Jr.; Vogel, J.-U.; Kotchetkov, R.; Scholz, M.; Doerr, H.W. (Frankfurt am Main)

425 426 427 429 after 430

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Author Index Vol. 42, No. 5–6, 1999 Subject Index Vol. 42, No. 5–6, 1999 Author Index Vol. 42, 1999 Subject Index Vol. 42, 1999 Contents Vol. 42, 1999

Contents

Intervirology 1999;42:277–278

Preface

Fig. 1. From the left to the right. First row: Dr. A. Zeevi, Dr. S. Michelson, Dr. I. Lautenschlager, Dr. N. DavisPoynter, Dr. M.-P. Landini. Second row: Dr. S. Prösch, Mrs. F. Claus-Hahn, Dr. L. van Renterghem, Dr. L. van Poucke, Dr. C. Bruggeman, Dr. A. Ramon, Dr. D. Sedmak, Dr. H. Thé. Third row: Mrs. G. Grauls, Dr. V. Emery, Dr. J. Middeldorp, Dr. J. Neyts, Dr. R. Blok, Dr. C. Söderberg, Dr. V. Goossens, Dr. M. Scholz, Dr. T. Mertens, Dr. F. Kern. Fourth row: Dr. G. Jan, Dr. C. Vink, Dr. J. Waldman and Dr. J.-U. Vogel.

Cytomegalovirus (CMV) is usually harmless and without clinical significance. More than 50% of the world’s population are carriers of CMV and therefore host the virus their whole lives, most of them not even knowing that they are infected. So, what keeps us busy and curious is the fascination exerted by CMV, a herpesvirus that is significantly implicated in morbidity and mortality in immunocompromised patients, such as transplanted or AIDS patients. Moreover, evidence is accumulating that

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CMV may also be pathogenic in nonimmunosuppressed persons (autoimmunity?). The management of CMV disease may be an enormous challenge for every clinician when antiviral agents prove to be inefficacious in controlling viremia. It has been suggested that CMV induces aberrant immune reactions that are necessary for its survival and contribute to pathogenesis. However, we still do not know how CMV triggers the immune system in an obviously ‘intelligent’

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way that enables its escape from the immune system and lifelong persistence in the host. Moreover, in order to disseminate, CMV seems to drive white blood cells to attach to infected host cells, which allows it to live and to ‘grab’ a comfortable carrier leukocyte within the blood stream. The living and transportation costs of CMV are paid for by the host through modifications of immune responses, which may be subclinical in immunocompetent hosts but highly critical in hosts with naturally or iatrogenically modified immunity. Of course, the concept of unlimited virus production leading to disease (progression) as a result of a decrease in ‘controlling’ immune cells induced by immunosuppression is generally accepted. But besides the viremia-associated disease, CMV probably also acts ‘incognito’ through the ‘back-door’. More specifically, it has been proposed by several researchers that CMV manipulates the immune system during latency or the immediate early phase of replication. It may be speculated that this (prolonged?) immediate early phase drives the host cell and the immune system to support the persistence and dissemination of the virus. As it leads to tolerance or to aberrant immune responses – both predominant factors in pathogenesis – this modified behavior may be of immunological relevance. This novel philosophy on the impact of CMV on the host, and the frequent failure of standard antiviral drugs to halt the disease, led us to look for novel therapeutic agents. These drugs should be able both to stop viral replication and inhibit virus-induced immunomodulation. Since CMV immediate early proteins have transactivating properties for viral and cellular genes, it may be important to develop drugs that target CMV immediate early expression. To obtain clinical success in the treatment of CMV disease, one cannot restrict oneself to inhibiting pathomechanisms in experimental models. It is essential to combine the knowledge and know-how of scientists working in the areas of diagnosis, antiviral therapy, clinical and basic research to solve CMV-related problems in close collaboration. Two years ago, a small group of researchers from different disciplines gathered for the first time in Frankfurt, Germany, to discuss the multifold aspects of CMV-related immunopathology. The contributions to that meeting were published in Monographs in Virology, a Karger book series. To ensure continuity not only in our scientific work, we are pleased to publish the proceedings of this year’s meeting in Intervirology, which is also published by Karger publishers, whose extended and valuable cooperation we would like to acknowledge.

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Furthermore, we would like to express our appreciation to all the scientists who came to Maastricht: first of all, to those who already participated in the Frankfurt meeting and who came in order to discuss the progress accomplished during the past two years, as well as to the ‘new’ participants. A lot of new data and interesting findings were presented in this meeting, giving us new insights in this complex virus. We learned that during stress situations the virus reactivates, but also some new aspects of the effect of CMV infection on interferon signal transduction. New detection methods for the virus diagnostic laboratories were presented for detection of active CMV infection in transplant recipients and for the detection of congenital infections. Using basic experimental models (in vivo and in vitro) new insights in the pathogenesis of CMV disease and interaction with the immune system were given. Especially, immune escape mechanism of this virus is an intriguing topic. Important findings in the field of antiviral therapy are under development giving promising results for the near future. Some of these new antivirals were presented during this meeting. All these contributions can be found in this issue of Intervirology. We thank all contributors for the interesting discussions we had, and are looking forward to the next meeting in 2001. The sponsors who made this meeting possible are listed on a separate page. We would also like to sincerely thank them and hope that the collaboration between biomedical science and industry will be successful in developing novel strategies in fighting against CMV disease. Cathrien A. Bruggeman, Maastricht Albert Ramon, Maastricht Martin Scholz, Frankfurt/Main

Preface

Part I. Immunopathology CMV Infection in the Immunocompromised Host


Time-Related Effects of Cytomegalovirus Infection on the Development of Chronic Renal Allograft Rejection in a Rat Model Irmeli Lautenschlager Anu Soots Leena Krogerus Kaija Inkinen Jeroen Kloover Raisa Loginov Kaisa Holma Harri Kauppinen Cathrien Bruggeman Juhani Ahonen Departments of Virology, Surgery and Pathology, University of Helsinki and Helsinki University Central Hospital, Helsinki, Finland, and Department of Medical Microbiology, University of Maastricht, The Netherlands

Key Words CMV W Renal transplantation W Chronic rejection

Abstract Cytomegalovirus (CMV) infection is a risk factor for chronic allograft rejection. The histological findings of chronic renal allograft rejection include inflammation, vascular intimal thickening, glomerulosclerosis, tubular atrophy and fibrosis. We have developed a rat model of renal transplantation in which transplants, after an early inflammatory episode, end up with chronic rejection within 60 days. During the early phase of the process in this model, CMV increased and prolonged the inflammatory response, the expression of adhesion molecules, intercellular adhesion molecule-1 and vascular cell adhesion molecule-1 and their ligands, lymphocyte function antigen-1 and very late antigen-4 in the graft. Simultaneously, the production of various growth factors, such as transforming growth factor beta, platelet-derived growth factor and connective tissue growth factor was upregulated, which induce smooth muscle cell proliferation in the vascular wall and collagen synthesis by fibroblasts. Chronic rejection developed within 20 days in

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CMV-infected grafts. In summary, CMV infection accelerated and enhanced the early immune response, the induction of growth factors and collagen synthesis, and the development of chronic rejection in renal allografts. Copyright © 2000 S. Karger AG, Basel

Introduction

Cytomegalovirus (CMV) infection is a major problem in renal transplantation [1]. In addition to clinical symptoms, CMV infection has been suggested to cause glomerulopathy [2] and to trigger acute rejection of kidney allografts in clinical transplantation [3–5]. CMV infection is also thought to be one of the risk factors for chronic rejection [6]. Although CMV has been shown to be associated with chronic rejection of heart [7–9], lung [10] and liver transplants [11–13], little is known about CMV and chronic rejection in clinical renal transplantation. The inflammatory response and T cell activation are characteristic of alloresponse and seem to be necessary for the development of chronic rejection [6, 14]. Vascular adhesion molecules are important in the early phase of the alloresponse, cell-to-cell interactions, T cell activation

Dr. I. Lautenschlager, MD, PhD Transplant Unit Research Laboratory, Fourth Department of Surgery Helsinki University Central Hospital, Kasarmikatu 11–13 FIN–00130 Helsinki (Finland) Tel. +358 9 47188484, Fax +358 9 47188348, E-Mail [email protected]

and extravasation of inflammatory cells into the organ [15–17]. Firm adhesion of leukocytes to the endothelial cells is established by the binding of intercellular adhesion molecule-1 (ICAM-1) and vascular cell adhesion molecule-1 (VCAM-1), both members of the Ig superfamily, to the integrin molecules expressed on leukocytes, such as lymphocyte function-associated antigen-1 (LFA-1) and very late antigen-4 (VLA-4) [18, 19]. The induction of adhesion molecules is mediated by cytokines, such as IFN-Á, TNF-· and IL-1 [19], which are produced during the inflammatory process of the alloresponse. Various cytokines produced by activated T cells and macrophages, such as IL-1, IL-2, IL-4, IL-6, TNF-· and INF-Á, may also contribute to the development of glomerulosclerosis as well as stimulating proliferation of smooth muscle cells in the vascular wall or generation of fibrosis [14]. Inflammatory factors, such as platelet-derived growth factor (PDGF) and transforming growth factor beta (TGF-ß) are important in smooth muscle cell proliferation, but they have also been reported to be highly fibrogenic [20]. In particular, generation of fibrosis is also mediated by fibroblast growth factors (FGF) [20] and connective tissue growth factor (CTGF), which has been found to be important in renal fibrosis [21]. Histological findings of chronic renal allograft rejection are well defined and are characterized by focal interstitial lymphocytic inflammation and fibrosis, glomerular mesangial matrix increase and sclerosis, vascular intimal proliferation and tubular atrophy [22, 23]. These variables can be summarized by the chronic allograft damage index (CADI), which can be used as a predictive parameter for chronic rejection [24, 25]. Interstitial fibrosis is the most prominent histological finding associated with endstage chronic rejection. We have recently developed an experimental model in which, after an early inflammatory episode, rat renal allografts develop chronic rejection under triple-drug immunosuppressive therapy within 60 days after transplantation [26]. We studied the early phase of the process in detail and recorded that the peak of inflammation occurs 5–10 days after transplantation. The inflammation is associated with lymphoid activation and induction of vascular adhesion molecules, such as ICAM-1 and VCAM-1. However, while the chronic changes increased, early expression of adhesion molecules vanished together with immune activation and was no longer detectable at the end stage of chronic rejection [27]. Generation of fibrosis, which results from the synthesis of various collagen proteins, was also studied in this model [28].

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In order to study the processes involved in chronic rejection complicated by CMV infection, we have also used the appropriate rat virus (RCMV) in our experimental kidney transplantation model [29]. This article is a short summary of preliminary findings of our studies on the time-related effects of CMV on the inflammatory response, induction of the expression of adhesion molecules and growth factors, histological changes and the generation of fibrosis in a rat model of renal transplantation during the development of chronic rejection.

Materials and Methods Transplantations Renal transplantations were performed in a rat strain combination of DA(RT1a)/BN (RT1n) as described previously [26]. The animals received triple-drug immunosuppression with methylprednisolone (MP), azathioprine (AZA) and cyclosporine (CyA) (MP 2 mg/kg, AZA 2 mg/kg, CyA 5 mg/kg s.c. daily). One group of animals was infected with RCMV Maastricht strain (see below) and the other group was left uninfected. To quantify the inflammation associated with alloresponse, the rat allografts were monitored by frequent, ultrasound-guided, fine-needle aspiration biopsy specimens (FNAB) [30]. The inflammation was quantified by the increment method, and expressed in corrected increment units (CIU) as described previously [31]. Histology The grafts were harvested at 3, 5, 7, 10, 20, 30, 40, 50 and 60 days after transplantation and histological preparations were performed. Each experimental group contained 4 animals at each time point. The Banff criteria [32], without grading, were used to assess graft histology. The numberical CADI was used to quantify the chronic alterations characteristic of chronic rejection, as described previously [24, 25]: focal interstitial lymphocytic inflammation, fibrosis, glomerular sclerosis, mesangial matrix increase, vascular intimal proliferation and tubular necrosis. RCMV Infection One group animals receiving triple treatment were infected with RCMV, Maastricht strain [33], by intraperitoneal inoculation of 105 plaque forming units (PFU) 1 day after renal transplantation. Syngenic controls, with triple-drug treatment and CMV infection were also performed. The characteristics of the rat virus and RCMV infection, as well as inoculation of the virus, have been described in detail previously [34]. RCMV infection was confirmed by viral culture from the FNAB specimens, 6–7 days after inoculation. The viral cultures were performed on rat embryonal fibroblasts; standard virus culture conditions were used and the virus was demonstrated in the fibroblast cultures after the detection of cytopathic effects by immunofluorescence and monoclonal antibodies against RCMV early and late antigens [35]. Direct detection of the RCMV infection was performed in parallel from frozen sections of the transplanted grafts by immunofluorescence staining and using the same monoclonal antibodies against RCMV antigens.

Lautenschlager et al.

Demonstration of Adhesion Molecules, Their Ligands and Lymphoid Activation Markers The expression of adhesion molecules and the number of inflammatory cells expressing their ligands and activation markers were immunohistochemically evaluated from frozen sections of the kidneys. An indirect immunoperoxidase technique and monoclonal antibodies against rat ICAM-1, VCAM-1, LFA-1, VLA-4, IL-2R and rat MHC class II were used. The expression of adhesion molecules in the various structures of the kidney was assessed and the intensity of expression was graded from 1 to 3. The numbers of infiltrating inflammatory cells positive for the ligand molecules, IL-2-R and class II were counted per high-power visual field. Demonstration of Growth Factors The expression of TGF-ß and platelet-derived growth factor-AA (PDGF-AA) was demonstrated by immunoperoxidase staining and monoclonal antibodies from the frozen sections of the explanted renal allografts at various time points. CTGF expression was determined at the mRNA level by in situ hybridization using a digoxigenin-labeled RNA probe [21]. Determination of Total Collagen and Expression of mRNAs of Various Collagen Types Total collagen and DNA contents were measured from the tissue homogenate of the grafts removed at different time points after transplantation. The methods for total collagen, estimated from the amount of tissue hydroxyproline [36], and DNA [37] have been described previously. The generation of fibrosis was determined as total collagen content/DNA ratio. The total collagen/DNA ratio reflects the amount of extracellular matrix to the cell content of the allografts. For mRNA analysis, total RNA was extracted from the grafts [38]. For accurate quantification, collagen I and collagen III mRNAs slot blot hybridizations were employed. The amounts of pro·1 (I) and pro·1 (III) collagen mRNAs were estimated by densitometric scanning of the exposed films using a densitometer connected to a computer to quantify the bands [39].

Results

RCMV Infection of Renal Allografts The presence of RCMV in the grafts was demonstrated by viral culture and direct antigen detection from the kidney on day 7 after transplantation [29]. CMV-specific antigens were detected in occasional passenger leukocytes and in some tubular structures, but a strong positive staining was found in the endothelium of capillaries and arterioles. In addition, the vascular wall of some large arteries were positively stained for CMV antigens located in the smooth muscle cells of the media. The endothelial cells of the large arteries were negative for CMV. The glomerular structures did not show any positivity for CMV antigens. The viral infection subsided within a week, and after day 14 posttransplantation, the RCMV-specific antigens were no longer detectable in the grafts.

CMV Infection and Chronic Renal Allograft Rejection

Impact of RCMV on Intragraft Inflammation and Graft Histology In uninfected animals, a mild inflammatory episode (peak 3.3 B 1.4 CIU) with some lymphoid activation but no macrophages was seen in FNAB about 5 days after transplantation. All grafts survived and the inflammation subsided, but graft histology demonstrated characteristic changes of chronic rejection within 60 days after transplantation [28]. Animals with RCMV infection also showed lymphoid activation, but the inflammation was significantly stronger (peak 4.5 B 1.8 CIU, p ! 0.05) than in uninfected animals [29], including a remarkable macrophage response (peak 1.5 B 1.2 CIU vs. 0.0 B 0.0, p ! 0.05). The inflammation continued until chronic rejection was histologically diagnosed within 20 days after transplantation. In the early phase after transplantation, by days 5–7, RCMV-infected grafts showed signs of vascular damage and heavy interstitial inflammation which consisted predominantly of lymphocytes. In contrast, grafts from noninfected rats showed less intense inflammation at this time point and only mild vascular changes. Already at 20 days after transplantation, in grafts harvested from RCMV-infected animals, characteristic findings of chronic rejection were recorded with significantly higher CADI values than in the noninfected grafts (9.0 B 0.5 vs. 5.8 B 1.7, p ! 0.05) [Kloover et al., unpubl. results]. A remarkable interstitial fibrosis was characteristic for end-stage grafts of the RCMV-infected animals. Impact of RCMV on Adhesion Molecule Expression and T Cell Activation In the RCMV-infected group, expression of ICAM-1 and VCAM-1, seen in the endothelium of the larger vessels, increased to a maximum at 7 days and remained at this level during the follow-up until 30 days after transplantation. In the noninfected grafts, induction of the adhesion molecules was also seen shortly after transplantation, but the expression decreased during the development of chronic changes. In the capillary endothelium, prolonged VCAM-1 and ICAM-1 expression was also seen in CMV-infected grafts [Kloover et al., unpubl. results]. Inflammatory cells expressing the ligands LFA-1 and VLA-4 were abundantly present in noninfected animals at 3–5 days after transplantation and the number of these cells decreased steadily during the development of chronic histological changes. RCMV infection resulted in a significant increase in the number of LFA-1-positive


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and VLA-4-positive cells present in the allograft when compared to noninfected animals [Kloover et al., unpubl. results]. The number of cells positive for the lymphoid activation markers IL-2R and MHC class antigens in allografts harvested from RCMV-infected animals was significantly increased during the early phase of the process compared to the noninfected rats. The lymphoid activation markers reflected the increased T cell activation in the CMVinfected grafts [Kloover et al., unpubl. results]. Effect of RCMV on TGF-ß, PDGF-AA and CTGF in the Graft A significantly more intense expression of TGF-ß in the capillary and vascular endothelium of the graft was reached earlier (by days 3–5) in RCMV-infected rats compared to the noninfected group which reached a lower peak expression at day 20 after transplantation. Also a more intense expression of PDGF-AA in the capillaries was seen earlier (at day 7) in the RCMV-infected grafts, compared to the noninfected group, which reached a lower peak at day 14. The expression of PDGF-AA was low in the vascular endothelium, without differences between the groups [Holma et al., unpubl. results]. In situ hybridization demonstrated a strong CTGF mRNA expression in fibroblasts, located mainly between the cortex and medulla and in a few gromerular cells of the RCMV-infeted grafts, peaking at day 14 after transplantation. In noninfected grafts, a minor expression of CTGF mRNA was recorded in the fibroblasts and glomeruli. This expression was strongest at 40–50 days after transplantation [Inkinen et al., unpubl. results]. Effect of RCMV on the Development of Fibrosis The time-related increase of collagen concentration in the grafts correlated with the development of interstitial fibrosis demonstrated by graft histology both in uninfected and in CMV-infected renal allograft recipients. In the grafts of CMV-infected animals, fibrosis associated with chronic rejection developed earlier, and a significantly higher collagen/DNA ratio than in the uninfected animals (6.2 B 2.5 vs. 3.0 B 1.0, p ! 0.05) was already evident at 20 days after transplantation [40; Inkinen et al., unpubl. results]. Both types I and III procollagen genes were expressed at all analysed time points after transplantation in both groups of animals. In RCMV-infected grafts, expression of types I and III collagen mRNAs was more prominent up to 20 days after transplantation than in noninfected kidneys [40; Inkinen et al., unpubl. results]. This indi-

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cated a more intense collagen synthesis activation and generation of fibrosis in the RCMV-infected renal allografts than in the noninfected grafts.

Discussion

In this experimental model of renal transplantation which develops chronic rejection, we demonstrated that CMV infection caused increased inflammation and macrophage response, induction of adhesion molecules and T cell activation in the graft. The increased and prolonged expression of adhesion molecules was probably due to cytokines, IL-1, IFN-Á and TNF-·, produced during the increased inflammatory response [41]. On the other hand, CMV also induces the production of various cytokines. CMV has been found to upregulate IL-1ß gene expression, which may lead to production of IL-1 by mononuclear cells [42]. CMV may interfere with the alloresponse while the immediate early genes of CMV have been found to upregulate the IL-2 and IL-2R genes [43]. The central role of TNF-· has been demonstrated, as CMV induces the production of TNF-· in monocytes and macrophages [44] and also RCMV has been shown to induce TNF-· in vivo [45]. This may lead to other immunological events, such as induction of adhesion molecules and increase of the inflammatory response. However, the increased early inflammation in CMV-infected animals was related to accelerated and enhanced histological changes of chronic rejection, such as vasculopathy and generation of interstitial fibrosis of the kidney transplant. Proinflammatory cytokines and growth factors play an important role in smooth muscle cell proliferation and in the stimulation of collagen synthesis in the fibrotic process. Although a great number of studies have been published on the development on graft vasculopathy, mainly dealing with experimental models of aortic allografts as our previous studies [46], very little is known about the generation of fibrosis associated with the end stage of the process of chronic kidney allograft rejection. The early accumulation of collagen in CMV-infected grafts correlated with an increased number of macrophages. Macrophages are known to be able to release inflammatory growth factors, such as FGF, PDGF and TGF-ß, which are highly fibrogenic and stimulate collagen synthesis in fibroblasts [20, 47, 58]. CTGF is one of the growth factors that is found to be associated with renal fibrosis [21]. Cytokines associated with inflammation, such as IL-1 and TNF-·, play an important role, not only in the regulation of adhesion molecule expression, but also in the stimula-


tion of the synthesis and release of the growth factors PDGF and TGF-ß [49–51]. The central role of growth factors, especially TGF-ß, in the pathogenesis of CMV-related changes, can also be explained by the fact that CMV infection has been shown to induce the transcription and secretion of TGF-ß1 [52]. This may lead to further stimulation of CTGF gene expression, which is shown to be stimulated by TGF-ß [53]. CTGF is secreted by fibroblasts after activation with TGF-ß [53]. The increased fibrogenesis in the CMVinfected grafts can be mediated by TGF-ß-CTGF interactions. In conclusion, the time-related analysis of the effects CMV infection on the development of chronic rejection in a rat model of renal transplantation demonstrated that CMV is associated with increased early inflammatory response, T-cell activation and adhesion molecule expres-

sion in the graft. Simultaneously, the production of various growth factors, such as TGF-ß, PDGF-AA and CTGF is upregulated. This induces smooth muscle cell proliferation in the vascular wall and collagen synthesis by fibroblasts. CMV accelerates and enhances each of these steps in the development of chronic rejection.

Acknowledgments This work was supported by grants from the Sigrid Juselius Foundation, Helsinki University Central Hospital Research Funds, the Finnish Academy of Science and NWO, the Netherlands. The authors wish to thank Dr. Roy Lobb and Biogen Inc. for providing the anti-rat VCAM-1 antibody. The authors also thank Saara Merasto and Tarja Markov for technical assistance, Stephen Venn for correcting the English text and Kari Savelius for animal care.

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Overcoming the Problem of Cytomegalovirus Infection after Organ Transplantation: Calling for Heracles? Willem J. van Son Eltjo F. de Maar Wim van der Bij Arie P. van den Berg Erik A.M. Verschuuren T. Hauw The Departmentof Internal Medicine, University Hospital Groningen, The Netherlands

Key Words Cytomegalovirus infection W Organ transplantation W Pathophysiology W Symptomatology

Abstract Although diagnosis of CMV infections and treatment of CMV disease with effetive antiviral drugs have become much easier, the persistent problem of CMV infection after solid-organ transplantation still requires solid knowledge of the pathophysiology of its clinical manifestations in order to minimize the impact of CMV infections in the future. The complex symptomatology of CMV infection after solid-organ transplantation is reviewed as well as some of the new theories attempting to explain the myriad of symptoms seen after transplantation. Copyright © 2000 S. Karger AG, Basel

Introduction

Although a range of reliable and accurate diagnostic tests for CMV as well as effective antivirals are available, cytomegalovirus (CMV) infections remain a substantial clinical problem in modern organ transplantation even now at the close of this century. Compared to 10–15 years ago, a substantial part of the clinical problems caused by

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© 2000 S. Karger AG, Basel 0300–5526/99/0426–0285$17.50/0



CMV have been solved: for instance, CMV-related deaths are now relatively rare in solid-organ transplantation, but new problems have emerged. Aside from the warning signs of resistance of CMV to antivirals due to increasing (prophylactic?) use of those drugs in modern intensive immunosuppressive protocols, various complications have appeared in the aftermath of transplantation, some of which point to CMV as the culprit. For instance, CMV may be involved in the process of chronic transplant dysfunction (‘chronic rejection’), as reviewed by Lautenschlager [1], while others [2, 3] suspect a role for CMV in case of accelerated coronary atherosclerosis found in patients with CMV infection after heart transplantation. The necessity to solve old problems while at the same time new nuisances emerge reminds us somewhat of the Greek legend of the monster Hydra. According to this legend, Hydra was a monster with nine heads (the number varies), the center one being immortal. The monster’s haunt was the marshes of Lerna near Argos. The destruction of Hydra was one of the twelve Labors of Heracles, which he accomplished with the assistance of his nephew Iolaus. The problem with this monster was that if one of its heads was cut off, two grew in its place. Therefore, Heracles decided to burn out the roots with firebrands and at last severed the immortal head from the body. So, to overcome new problems attributed to CMV after solving old ones, we probably need a solution such as that

Willem J. van Son, MD PhD Department of Internal Medicine University Hospital AZG Hanzeplein 1 NL–9700 RB Groningen (The Netherlands)

used by Heracles in the legend. However, as we do not appear to be on the verge of evolving a method of killing the virus in its latent state (‘the immortal head’) without causing lethal damage to its host, CMV infections will remain a problem in clinical solid-organ transplantation.

Risk Factors for CMV Disease

As far as the symptomatology of CMV infection is concerned, it is known that a substantial part of patients with an active CMV infection are asymptomatic. Risk factors for CMV disease are well recognized and include the following: transplanting a CMV-seronegative patient with a CMV-seropositive donor organ (leading to a primary infection) and the recipient’s net state of immunosuppression, determined by the characteristics of the immunosuppressive protocol (type, dose, duration, timing) and various host factors (comorbidity, age, uremia, neutropenia, infections with other immunomodulating viruses) [4]. Bruning et al. [5] suggested an additional important role of allostimulation by the graft in the process of reactivation of CMV in a rat model. As far as the type of immunosuppression is concerned, especially antilymphocyte antibodies and monoclonal antibody preparations, such as OKT3, are well known to cause a high incidence of CMV disease when used for either induction or antirejection therapy. While the immunosuppressive potency of this class of drugs is usually suggested as the explanation for this high incidence of infection, other mechanisms might be involved. Recently, a novel mechanism of reactivation has been proposed: the proinflammatory cytokine TNF-· has been suspected to play a role in the reactivation of CMV by stimulation of the CMV major immediate early enhancer/promoter [6– 8]. Since the use of OKT3 is well known for its abundant cytokine release (i.e. TNF-·) this might explain the high incidence of active CMV infection during OKT3 treatment. Finally, viral load might be an important factor. Several authors stress the importance of a high viral load in relation to the risk of clinically relevant CMV disease [4, 9–13].

Symptomatology

Although in the era of rapid diagnostic tests for CMV and effective antivirals one might expect that the effect of CMV would fade away, CMV infections still have a substantial impact on graft and patient survival in solid-

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organ transplantation [4, 9, 14–22]. Of all patients who develop clinical manifestations of CMV infection, more than 90% do so within 1–6 months after transplantation, and 60% of the febrile episodes during this period are due to CMV infections [4, 9]. Because of the more potent immunosuppression used nowadays – with induction schemes including monoclonal and polyclonal anti-T-cell antibodies – timetable of infectious complications after transplantation tends to ‘shift to the left’ [4], resulting in earlier appearance of clinically significant CMV infections. When patients are symptomatic, symptoms may vary greatly. Most of the patients have a so-called ‘selflimiting syndrome’, consisting of fever (often spiking), arthralgia, leukopenia and/or thrombocytopenia, and abnormal liver enzymes [4, 9]. With tapering of the immunosuppressive therapy, the great majority of patients recover completely from the syndrome. Aside from this self-limiting syndrome, CMV may cause a myriad of symptoms in the grafted patient. Gastrointestinal symptoms during CMV infections are numerous. They include gastrointestinal ulcers that may bleed or perforate (i.e. gastric and colonic ulcers) [4, 9, 23–28]. CMV-associated vasculitis might be the common pathogenetic mechanism of this manifestation of the CMV syndrome in the immunocompromised patient [29]. Other gastrointestinal symptoms comprise pancreatitis [4, 9], granulomatous hepatitis [30] and pneumatosis intestinalis [4, 9]. The latter condition is of particular interest because the cysts may perforate, causing a sterile pneumoperitoneum. As a consequence, free air may be present under the diaphragm on chest X-ray; the awareness of the association of this condition with an active CMV infection may avoid an unnecessary surgical intervention [31, 32]. Gastrointestinal symptoms may be present without other major symptoms of infection, and CMV may be present in the gastroduodenal tract without symptoms. Franzin et al. [33] found evidence of CMV inclusion bodies in biopsies collected from the gastroduodenal mucosa of patients with a renal allograft in 9 out of 20 cases. The presence of these CMV inclusions was unrelated to viremia-induced or gastrointestinal symptoms at the time of endoscopy [33]. Several other symptoms have been attributed to CMV: lymphadenopathy, hepatosplenomegaly, pericarditis, myocarditis, encephalitis, retinitis, and skin ulcerations associated with vasculitis [4, 9, 34, 35]. CMV-induced Guillain-Barré syndrome is a wellknown phenomenon [36] and is seen relatively frequently in AIDS patients, but is a rarely encountered sequel of an active CMV infection in transplanted patients. In AIDS patients, direct infection of the nerves has been suggested

van Son/de Maar/van der Bij/van den Berg/ Verschuuren/The

to play a role in the pathophysiology of the syndrome, as well as autoimmune phenomena elicited by the virus [37, 38]. The reasons why the incidence of this condition differs so greatly between the AIDS- and the transplanted population are unknown, but one may speculate about the significance of some overt differences that exist between the two patient groups. For instance, although AIDS patients are also immunosuppressed, the nature of this suppression differs considerably, which might reflect differences in sequelae leading to a possible autoimmune process elicited by the virus. Another item that might be important in order to explain the dissimilarity is the duration of viremia. In contrast to AIDS patients, in whom prolonged CMV viremia is the rule, duration of viremia is usually short in transplanted patients, being mostly confined to the period of maximum immunosuppression, i.e. shortly after initiation of antirejection therapy. One might speculate about the importance of prolonged viremia as a prerequisite for the development of autoimmune phenomena. If the latter is true, this might be an important clue in the pathophysiology of a patient characterized by an unusually prolonged period of CMV viremia, very recently described by de Maar et al. [38a]. This patient presented with a chronic inflammatory demyelinating polyneuropathy after renal transplantation during recurrent CMV viremia. It is important, however, to stress that great caution should be exercised in designating CMV as the culprit in case of a given symptom: a single positive laboratory test may not be enough to consider the symptoms present as induced by CMV. Other possible causes must be excluded and laboratory signs for CMV infection have to be judged in concert with other signs of CMV infection. CMV pneumonia after solid organ transplantation is the manifestation that distinguishes serious illness from more benign disease, and this condition is associated with high mortality, especially when assisted ventilation is required [4, 9, 39]. However, although still feared in bone marrow transplantation, with the emergence of rapid diagnostic tests and the availability of effective prophylactic schemes as well as effective antivirals to treat CMV infection in an early stage, the incidence of CMV pneumonia has dramatically dropped. Renal involvement during CMV infection remains controversial as far as pathogenesis is concerned. It has been suggested that CMV could trigger the immune mechanism of acute rejection either via i.e. increased MHC class II expression in the graft or via mimicry since CMV was shown to have sequence homology and immunologic cross-reactivity with the HLA-DR ß-chain [40, 41]. More

recently, Reinke et al. [42] described a series of patients with asymptomatic CMV infections which they linked with late acute rejection. Alternatively, in 1981, Richardson et al. [43] described a distinctive pattern of glomerular injury in renal allografts that they associated with CMV viremia without relation with allograft rejection. The pathological features consisted of diffuse endothelial hypertrophy and necrosis, accompanied by an accumulation of fine fibrillar webs of periodic-acid-Schiff (PAS)positive material and mononuclear cells that resulted in obliteration of the glomerular capillaries. Fibrin, IgM as well as C3 were fond by immunofluorescence. No viral particles were detectable by electron microscopy or immunofluorescence using monoclonal antibodies directed to CMV early and late antigen [43]. The existence of this condition has been disputed by Herrera et al. [44] and Boyce et al. [45], who consider it to be a form of (vascular-type or ‘transplant glomerulopathy’) rejection. Aside from the directly attributable syndromes there are important indirect effects of CMV in the compromised host. CMV has important immunomodulating effects [4, 9] that contribute significantly to the ‘net immunosuppressive state’ of a given patient, making him or her prone to superinfections with i.e. Aspergillus species and Pneumocystis carinii [4, 9, 46]. In a recent multicenter study by the Boston Center for Liver Transplantation, a multivariate analysis showed that patients with CMV disease had significantly more invasive fungal disease and bacteremia [47].

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Pathophysiology of CMV Infection

How CMV causes symptomatology and organ dysfunction in the recipient is still rather enigmatic and the subject of much debate and research. The question is whether symptomatology is caused by the cytopathic effect of the virus itself or is brought about by the (innate as well as specific) immune response elicited by the virus or virusinfected cells (i.e. endothelial cells). On the one hand, a clear relation exists between the viral load and the likelihood and severity of CMV disease [4, 9–12], but on the other hand it has been known for years that notably the number of antigen-positive leukocytes (pp65 antigenemia), which has been shown to correlate well with viral load [48], is not always a reflection of the virtual severity of the clinical situation. The question remains unanswered whether this discrepancy is caused by a high viral load with a less virulent virus or a more mitigated immune

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response to the same virus. For instance, there exists a lot of data indicating that virus-induced pneumonitis is not only the result of uncontrolled virus replication in the immunocompromised host, who is unable to control viral replication, but is rather the result of a T-cell-mediated immune response induced by viral antigens [49]. So, the protean manifestations of CMV seen after solid-organ transplantation suggest that unraveling of the pathophysiology will most likely reveal a very complex and multifaceted mechanism. Is CMV-induced endothelial cell damage (‘vasculitis’?) – with or without secondary immune response of the infected endothelial cells – the key factor? The key role of the endothelial cell in the pathophysiology is in agreement with the work of Persoons et al. [50] using a rat model. They found that in this infection model multiple-organ involvement was associated with disseminated vascular pathology. CMV-infected endothelial cells might secondarily induce adhesion of leukocytes to the damaged endothelium [51], trigger the coagulation system [52] and induce cytokines such as IL-6 [53], which might contribute to the pathophysiology of CMV-induced multiorgan disease. CMV also infects endothelial cells of transplanted patients; these cells may even detach from the vessel wall and subsequently be released in the circulation [54]. In the mononuclear cell fraction of peripheral blood of a patient with a CMV infection after heart transplantation, Grefte et al. [54] unexpectedly found distinctly large cells (35–45 Ìm diameter), reminiscent of the classical ‘cytomegalic inclusion body cells’. In a subsequent study, Grefte et al. [55] showed that those cells could be demonstrated in a substantial part of patients with an active CMV infection after solid-organ transplantation, especially in those with a high viral load in their blood. Immunostaining revealed those cells to be endothelial cells positive for HCMV antigens of all three stages of the viral replication cycle (with the typical nuclear and cytoplasmic localization of the distinct antigens) [54], while transmission electron microscopic studies showed that viral capsids, viral particles and dense bodies were present in the nucleus and cytoplasm, respectively, indicating that those cells represent a site of virus production [55]. Whether those cytomegalic endothelial cells play a role in the dissemination of the virus and/or organ dysfunction is far from settled, but it surely fits in with the data obtained from animal models [50–53] that suggest a key role for the CMV-infected endothelial cells in the pathophysiology of CMV infection. CMV infection may thus be looked on as a systemic disease with a multifaceted symptomatology. In this respect, it is of interest that subclinical

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organ involvement can be demonstrated in the majority of patients after organ transplantation even with an asymptomatic CMV infection [56]. A majority of patients with an active CMV infection after renal transplantation show subtle pulmonary dysfunction (decreased diffusion capacity for carbon monoxide) without clinical pulmonary symptoms [56]. This may point to subclinical pneumonitis due to either a direct cytopathic effect of CMV on pulmonary tissue, or to cellular immune response directed to e.g. infected pulmonary endothelial cells, with or without serum complement activation by the classical or alternate activation pathway [56, 57]. Another explanation might be that large cytomegalic endothelial cells 35– 45 Ìm in diameter [54] plug into the lung capillaries, which have a diameter of only 5 Ìm. This was subsequently studied by De Maar et al. [58] using a method by which both components of diffusion of CO could be studied separately; the membrane factor Dm (i.e. affected by edema of the alveolar membrane) and the capillary (‘blood’) factor Vcap, which is most likely to be affected by plugging of those large cells into the capillary bed. We concluded from this study that, since both Dm and Vcap decreased during active CMV infection, the decreased diffusion capacity for CO was not caused by plugging of the cytomegalic endothelial cells alone. Other factors might be involved: i.e., since they contain active replicating virus [54, 55], the cytomegalic endothelial cells may lead to spreading of the virus into the capillary bed, which in turn together with complement activation and/or locally induced cytokine production [51, 53, 59], might cause edema of the alveolar membrane and hence to the decreased Dm found in these patients [58]. Recently, we have also shown subclinical involvement of the gastrointestinal tract [60]. Increased intestinal permeability indicating intestinal epithelial damage has been found in a substantial number of patients with an active CMV infection after renal transplantation [60]. Although this also points towards a more systemic nature of CMV infections, the exact reasons for those findings still have to be elucidated.

Summary and Conclusions

Even in modern transplantation, with the availability of rapid diagnostic tests and effective antiviral drugs, CMV infections remain the single most important infectious complication after solid-organ transplantation. In the last decade, much has been learned about its pathophysiology, although it requires much more thought to


comprehend the exact mechanism of the protean manifestations of CMV infection after organ transplantation. Since we continue to meddle in the inevitable marriage between CMV and host immunity, introducing new, nonselective immunosuppressive drugs, we probably need a ‘Heracles-like’ solution for solving the CMV problem in

modern transplantation. Since it is very unlikely that antiCMV drugs will become available in the near future to influence the latent state of this virus by nontoxic means, CMV will remain a serious problem, even at the dawn of the new millennium.

References 1 Lautenschlager I: Role of cytomegalovirus infection in the processes of organ allograft rejection; in Scholz M, Rabenau HF, Doerr HW, Cinatl J (eds): CMV-Related Immunopathology. Monogr Virol. Basel, Karger 1998, vol 21, pp 142–157. 2 Grattan MT, Moreno Cabral CE, Starnes VA, Oyer PE, Stinson EB, Shumway NE: Cytomegalovirus infection is associated with cardiac allograft rejection and atherosclerosis. JAMA 1989;261:3561–3566. 3 Koskinen PK, Kallio EA, Tikkanen JM, Shivola RK, Hayry PJ, Lemstrom KB: Cytomegalovirus infection and cardiac allograft vasculopathy (review article). Transplant Infect Dis 1999;1:115–126. 4 Rubin RH: Infection in the organ transplant recipient; in Rubin RH, Young LS (eds): Clinical approach to infection in the compromised host, ed 3. New York, Plenum Press, 1994, pp 629–705. 5 Bruning JH, Bruggeman CA, Van Boven CP, Van Breda-Vriesman PJ: Reactivation of latent rat cytomegalovirus by combination of immunosuppression and administration of allogeneic immunocompetent cells. Transplantation 1989;47:917–918. 6 Stein J, Volk HD, Liebenthal C, Kruger DH, Prosch S: Tumor necrosis factor alpha stimulates the activity of the human cytomegalovirus major immediate early enhancer/promoter in immature monocytic cells. J Gen Virol 1993; 74:2333–2338. 7 Docke WD, Prosch S, Fietze E, Kimel V, Zuckermann H, Klug C, Syrbe U, Kruger DH, Von Baehr R, Volk HD: Cytomegalovirus reactivation and tumor necrosis factors. Lancet 1994; 343:268–269. 8 Prosch S, Volk HD, Reinke P, Pioch K, Docke WD, Kruger DH: Human cytomegalovirus infection in transplant recipients: Role of TNF-· for reactivation and replication of human cytomegalovirus; in Scholz M, Rabenau HF, Doerr HW, Cinatl J (eds): CMV-Related Immunopathology. Monogr Virol. Basel, Karger, 1998, vol 21, pp 29–41. 9 Van Son WJ, The TH: Cytomegalovirus infection after organ transplantation: An update with special emphasis on renal transplantation. Transplant Int 1989;2:147–164.

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10 Van den Berg AP, van der Bij W, Van Son WJ, Anema J, Van der Giessen M, Schirm J, Tegzess AM, The TH: Cytomegalovirus antigenemia as a useful marker of symptomatic cytomegalovirus infection after renal transplantation: A report of 130 consecutive patients. Transplantation 1989;48:991–995. 11 Cope AV, Sweny P, Sabin C, Rees L, Griffiths PD, Emery VC: Quantity of cytomegalovirus viruria is a major risk factor for cytomegalovirus disease after renal transplantation. J Med Virol 1997;52:200–205. 12 Cope AV, Sabin C, Burroughs A, Rolles K, Griffiths PD, Emery VC: Interrelationships among quantity of human cytomegalovirus (HCMV) DNA in blood, donor-recipient serostatus, and administration of methylprednisolone as risk factors for HCMV disease following liver transplantation. J Infect Dis 1997;176: 1484–1490. 13 Hassan-Walker AF, Kidd IM, Sabin C, Sweny P, Griffiths PD, Emery VC: Quantity of human cytomegalovirus (CMV) DNAemia as a risk factor in renal allograft recipients: Relationship with donor/recipient CMV serostatus, receipt of augmented methylprednisolone and antithymocyte globulin. J Med Virol 1999;58:182– 187. 14 Fryd DS, Peterson PK, Ferguson RM, Simmons RL, Balfour HH, Najarian JS: Cytomegalovirus as a risk factor in renal transplantation. Transplantation 1980;30:436–439. 15 Rubin RH, Tolkoff-Rubin NE, Oliver D, Rota TR, Hamilton J, Betts RF, Paas RF, Hillis W, Szmuness W, Farell ML, Hirsch MS: Multicenter seroepidemiologic study of the impact of cytomegalovirus infection on renal transplantation. Transplantation 1985;40:243–249. 16 Bia MJ, Andiman W, Gaudio K, Kliger A, Siegel N, Smith D, Flye W: Effect of treatment with cyclosporine versus azathioprine on incidence and severity of cytomegalovirus infection posttransplantation. Transplantation 1985;40:610–614. 17 Lewis RM, Johnson PC, Golden D, van Buren CT, Kerman RH, Kahan BD: The adverse impact of cytomegalovirus infection on clinical outcome in cyclosporin-prednisolone treated renal allograft recipients. Transplantation 1988;45:353–359.

18 Schnitzler MA, Woodward RS, Brennan DC, Spitznagel EL, Dunagan WC, Baily TC: Impact of cytomegalovirus serology on graft survival in living related kidney transplantation: Implications for donor selection. Surgery 1997;121: 563–568. 19 George MJ, Snydman DR, Werner BG, Griffith J, Falagas ME, Dougherty NN, Rubin RH: The independent role of cytomegalovirus as a risk factor for invasive fungal disease in orthotopic liver transplant recipients. Am J Med 1997;103:106–113. 20 Schnitzler MA, Woodward RS, Brennan DC, Phelan DL, Spitznagel EL, Boxerman SB, Dunagan WC, Baily TC: Cytomegalovirus and HLA-A, B, and Dr locus interactions: Impact on renal transplant survival. Am J Kidney Dis 1997;30:766–771. 21 Bock GH, Sullivan EK, Miller D, Gimon D, Alexander S, Ellis E, Elshihabi I: Cytomegalovirus infection following renal transplantation – effects on antiviral prophylaxis: A report of the North American Pediatric Renal Transplant Cooperative Study. Pediatr Nephrol 1997;11:665–671. 22 Rosen HR, Corless CL, Rabkin J, Chou S: Association of cytomegalovirus genotype with graft rejection after liver transplantation. Transplantation 1998;66:1627–1631. 23 Ayulo M, Aisner SC, Margolis K, Moravec C: Cytomegalovirus-associated gastritis in a compromised host. JAMA 1980;243:1364. 24 Foucar E, Mukai KM, Foucar K, Sutherland DER, van Buren CT: Colon ulceration in lethal cytomegalovirus infection. Am J Clin Pathol 1981;76:788–801. 25 Cohen EB, Komorowski RA, Kaufman HM, Adams M: Unexpectedly high incidence of cytomegalovirus infection in apparent peptid ulcers in renal transplant patients. Surgery 1985; 97:606–612. 26 Shrestha BM, Parton D, Gray A, Shepard D, Griffith D, Westmoreland D, Griffin P, Lord R, Salaman JR, Moore RH: Cytomegalovirus involving gastrointestinal tract in renal transplant recipients. Clin Transplant 1996;10:170– 175. 27 Toogood GJ, Gillespie PH, Gujral S, Warren BF, Roake JA, Gray DW, Morris PJ: Cytomegalovirus infection and colonic perforation in renal transplant patients. Transplant Int 1996; 9:248–251.


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28 Halme L, Hockerstedt K, Salmela K, Lautenschlager I: CMV infection detected in the upper gastrointestinal tract after liver transplantation. Transplant Int 1998;11(suppl 1): S242–244. 29 Golden MP, Hammer SM, Wanke CA, Albrecht MA: Cytomegalovirus vasculitis. Case report and review of the literature. Medicine (Baltimore) 1994;73:246–255. 30 Clarke J, Craig RM, Saffro R, Murphy P, Yokoo H: Cytomegalovirus granulomatous hepatitis. Am J Med 1979;66:264–269. 31 van Son WJ, van der Jagt EJ, van der Woude FJ, Slooff MJH, Meijer S, The TH, Tegzess AM, van der Slikke LB, Donker AJM: Pneumatosis intestinalis in patients after cadaveric kidney transplantation: Possible relationship with an active CMV infection. Transplantation 1984;38:506–510. 32 Mannes GP, De Boer WJ, van der Jagt EJ, Meinesz AF, Meuzelaar JJ, van der Bij W: Pneumatosis intestinalis and active cytomegaloviral infection after lung transplantation. Groningen Lung Transplant Group. Chest 1994;105:929– 930. 33 Franzin G, Muolo A, Griminelli T: Cytomegalovirus inclusion bodies in the gastroduodenal mucosa of patients after renal transplantation. Gut 1981;22:698–701. 34 Dorfman LJ: Cytomegalovirus encephalitis in adults. Neurology 1973;23:136–144. 35 Minars N, Silverman JF, Escobar MR, Martinez AJ: Fatal cytomegalic inclusion disease. Arch Dermatol 1997;113:1569–1571. 36 Hart IK, Kennedy PGE: Guillain-Barré syndrome associated with cytomegalovirus infection. Q J Med (New Ser 67) 1988;253:425– 430. 37 Yu RK, Ariga T, Kohriyama T, Kusunoki S, Maeda Y, Miyatani N: Autoimmune mechanism in peripheral neuropathies. Ann Neurol 1990;27(suppl):S30–305. 38 Wucherpfennig KW, Strominger JL: Molecular mimicry in T cell-mediated autoimmunity: Viral peptides activate human T cell clones specific for myelin basic protein. Cell 1995;80: 695–705. 38a de Maar EF, Kas-Deelen DM, de Jager AE, The H, Tegzess AM, van Son WJ: Inflammatory demyelinating polyneuropathy in a kidney transplant patient with cytomegalovirus infection. Nephrol Dial Transplant 1999;14:2228– 2230. 39 Peterson PK, Balfour HH, Marker SC, Fryd DS, Howard RJ, Simmons RL: Cytomegalovirus-disease in renal allograft recipients: A prospective study of the clinical features, risk factors and impact on renal transplantation. Medicine 1980;59:283–300.

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40 Von Willebrand E, Petterson E, Ahonen J, Hayry P: CMV infection, class II antigen expression, and human kidney allograft rejection. Transplantation 1986;42:364–367. 41 Funjinami RS,Nelson JA, Walker L, Oldstone MBA: Sequence homology and immunologic cross-reactivity of human cytomegalovirus with HLA-DR ß chain: A means for graft rejection and immunosuppression. J Virol 1988;62: 100–105. 42 Reinke P, Fietze E, Ode-Hakim S, Prosch S, Lippert J, Ewert R, Volk HD: Late acute renal allograft rejection and symptomless cytomegalovirus infection. Lancet 1994;344:1737– 1738. 43 Richardson WP, Colvin RB, Cheeseman SH, Tolkoff-Rubin NE, Herrin JT, Cosimi AB, Collins AB, Hirsch MS, Mcluskey RT, Russel PS, Rubin RH: Glomerulopathy associated with cytomegalovirus viremia in renal allografts. N Engl J Med 1981;305:57–63. 44 Herrera GA, Alexander RW, Cooley CF, Luke RG, Kelly DR, Curtis JJ, Gockerman JP: Cytomegalovirus glomerulopathy: A controversial lesion. Kidney Int 1986;29:725–733. 45 Boyce NW, Hayes K, Ge D, Holdsworth SR, Thomson NM, Scott D, Atkins RC: Cytomegalovirus infection complicating renal transplantation and its relationship to acute transplant glomerulopathy. Transplantation 1988;45: 706–709. 46 Husni RN, Gordon SM, Longworth DL, Arroliga A, Stillwell PC, Avery RK, Maurer JR, Metha A, Kirby T: Cytomegalovirus infection is a risk factor for invasive aspergillosis in lung transplant recipients. Clin Infect Dis 1998;26: 753–755. 47 George MJ, Snydman DR, Werner BG, Griffith J, Falagas ME, Dougherty NN, Rubin RH: The independent role of cytomegalovirus as a risk factor for invasive fungal disease in orthotopic liver transplant recipients. Boston Center for Liver Transplantation CMVIG-Study Group. Cytogam, MedImmune, Inc Gaithersburg, Maryland. Am J Med 1997;103:106– 113. 48 van der Bij W, Torensma R, van Son WJ, Anema J, Schirm J, Tegzess AM, The TH: Rapid immunodiagnosis of active cytomegalovirus infection by monoclonal antibody staining of blood leukocytes. J Med Virol 1988;25:179– 188. 49 Grundy JE, Shanley JD, Griffiths PD: Is cytomegalovirus interstitial pneumonitis in transplant recipients an immunopathological condition? Lancet 1987;ii:996–999.


50 Persoons MCJ, Stals FS, van Dam-Mieras MC, Bruggeman CA: Multiple organ involvement during experimental cytomegalovirus infection is associated with disseminated vascular pathology. J Pathol 1998;184:103–109. 51 Span AHM, van Boven CPA, Bruggeman CA: The effect of cytomegalovirus infection on the adherence of polymorphonuclear leukocytes to endothelial cells. Eur J Clin Invest 1989;19: 542–548. 52 van Dam-Mieras MCE, Muller AD, van Hinsberg VWM, Mullers WJHA, Bomans PHH, Bruggeman CA: The procoagulant response of cytomegalovirus infected endothelial cells. Thromb Haemost 1992;68:364–370. 53 Almeida GD, Porada CD, St Jeor S, Ascensao JI: Human cytomegalovirus alters interleukin6 production by endothelial cells. Blood 1994; 83:370–376. 54 Grefte JMM, van der Giessen M, van Son WJ, The TH: Circulating human cytomegalovirus (HCMV)-infected endothelial cells in patients with an active HCMV infection. J Infect Dis 1993;167:270–277. 55 Grefte JMM, Blom N, van der Giessen M, van Son WJ, The TH: Ultrastructural analysis of circulating cytomegalic cells in patients with active cytomegalovirus infection: Evidence for virus production and endothelial origin. J Infect Dis 1993;168:1110–1118. 56 van Son WJ, Tegzess AM, The TH, Duipmans J, Slooff MJH, van der Mark TW, Peset R: Pulmonary dysfunction is common during a CMV infection after renal transplantation, even in asymptomatic patients: Possible relationship with complement activation. Am Rev Respir Dis 1987;136:580–585. 57 van Son WJ, van der Bij W, Tegzess AM, Anema J, van der Giessen M, van der Hem GK, Marrink JM, The TH: Complement activation during active cytomegalovirus infection after renal transplantation: Due to circulating immune complexes or alternate pathway activation? Clin Immunol Immunopathol 1989;50: 109–121. 58 de Maar EF, Kas-Deelen AM, The TH, Tegzess AM, Ploeg RJ, van Son WJ: Cytomegalovirus pneumonitis after kidney transplantation is not caused by plugging of cytomegalic endothelial cells alone. Transpl Int 1999;12:56–62. 59 Bergeron Y, Ouellet N, Deslauriers AM, Simard M, Olivier M, Bergeron MG: Cytokine kinetics and other host factors in response to pneumococcal pulmonary infection in mice. Infect Immun 1998;66:912–922. 60 de Maar EF, Kleibeuker JH, Boersma-van Ek W, The TH, van Son WJ: Increased intestinal permeability during CMV infections in renal transplant recipients. Transplant Int 1996;9: 576–580.




Clinical Significance of Cytomegalovirus-Specific T Helper Responses and Cytokine Production in Lung Transplant Recipients Adriana Zeevi Kathy Spichty Richard Banas Jane Cai Vera S. Donnenberg Albert D. Donnenberg Mamun Ahmed James Dauber Aldo Iacono Robert Keenan Barthley Griffith Divisions of Transplantation Pathology, Transplant Surgery, Medicine and Thomas E. Starzl Transplantation Institute, University of Pittsburgh Medical Center, Pittsburgh, Pa., USA

Key Words Cytomegalovirus W T helper responses W Cytokines W T cell activation W Allograft rejection

Abstract Cytomegalovirus (CMV) disease continues to be a major problem for lung transplant recipients. In CMV-seropositive individuals, we detected two types of CMV-specific responses: a self-restricted response stimulated by soluble CMV antigen (sCMV-Ag) and a non-self-restricted response induced by CMV-infected cells (cCMV-Ag). Lung transplant recipients who develop the CMV-specific self-restricted T helper response have a low risk of recurrent CMV disease. In contrast, during CMV disease, lung transplant recipients exhibit only the non-selfrestricted T helper responses. We characterized the T cell activation and the kinetics of cytokine production of sorted CD4+ and CD8+ T cells from PBLs of CMV seropositive donors. The two types of CMV antigens induced cytokine production in both T cell subsets. We also per-

Supported by NIH grant HL55986 and Pathology Educational Research Grant.

ABC Fax + 41 61 306 12 34 E-Mail [email protected] www.karger.com



formed competitive RT-PCR for Granzyme B (GB) in BAL cells of lung transplant recipients prior to, during and following CMV disease. CMV disease was associated with increase in GB gene expression when was accompanied by acute cellular rejection while it remained low in patients with CMV disease that did not have a complicated course. In summary, CMV-activated T cells within the allograft may produce inflammatory cytokines and effector molecules that may promote allograft rejection. Copyright © 2000 S. Karger AG, Basel

Introduction

Cytomegalovirus (CMV) disease continues to be a major problem for lung transplant recipients, who cannot generate an efficient immune response to control this viral infection. Immunocompromised individuals are particularly at risk from life-threatening CMV infection and disease [1, 2]. In lung transplant recipients, the loss of CMV-specific T helper (Th) memory response in pretransplantation seropositive individuals and the persistent lack of CMV-specific Th memory response following primary infection are associated with recurrent CMV disease [3]. The generation of Th response is dependent on

Adriana Zeevi, PhD Room W1551 Biomedical Science Tower University of Pittsburgh Medical Center Pittsburgh, PA 15261 (USA) Tel. +1 412 624 1073, Fax +1 412 624 6666, E-Mail [email protected]

the presentation of soluble CMV antigen (sCMV-Ag) by autologous or HLA class-II-matched antigen-presenting cells (APCs). Another type of CMV-dependent Th memory response was initially described by Waldman et al. [4]. CMV-infected endothelial cells that lack HLA class II expression could stimulate proliferation of CD3+ T cells isolated from CMV-seropositive individuals [4, 5]. CMV-infected lung epithelial A549 cells could also trigger the proliferation of CD3+ T cells in seropositive individuals in the absence of MHC-restricted APCs [6]. CD3+ T cells from bronchoalveolar lavages (BALs) and peripheral blood of lung transplant recipients exhibit CMV-specific Th memory responses when stimulated either with the sCMV-Ag presented by autologous APC (self-HLA-restricted response) or with the cellular CMVAg (cCMV-Ag), from CMV-infected A549 cells (non-self HLA-restricted response) [6]. The cCMV-Ag Th response was also observed in lung transplant recipients during CMV disease while they failed to respond to sCMV-Ag. These findings suggested to us that CMV-infected parenchymal cells within the lung may recruit and activate CD3+ T cells to release cytokines that can promote inflammation and allograft rejection. In this study, we further characterize the pool of CMVspecific memory Th cells and their potential role in promoting inflammation within the allograft. We investigated cell activation and cytokine production in CD4+ and CD8+ T cell subsets stimulated by these two types of CMV-Ags: sCMV-Ags and cCMV-Ags.

Materials and Methods Preparation of Cells Peripheral blood mononuclear cells from CMV-seropositive individuals were obtained by Ficoll-Hypaque gradient. CD3+ T cells (98% purity by flow cytometry) were obtained by T lympho-qwik (One Lambda, Inc.; Canoga Park, Calif., USA), a commercially available cocktail of monoclonal antibody (mAb) and complement. CD4+ and CD8+ T lymphocytes were sorted by flow cytometry (Becton Dickinson Flow Activated Cell Sorter). CD8+ bright cells were positively sorted, eliminating possible NK cell contamination, which are CD8 dim. CD8– cells with lymphocyte light scatter were considered as the CD4+ cell population. Source of CMV-Ag sCMV-Ag was prepared as previously described using a glycine buffer extraction method following infection of the human fibroblast cell line (MRC-5) with the AD169 strain of CMV [3]. Mock antigen was prepared from the same cells without infection. cCMV-Ag was prepared by infecting alveolar epithelial cell line A549 (human lung carcinoma from American Type Cell Collection) with CMV strain VHL/E [6].

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Four-Color Flow Cytometry Analysis of T Cell Activation following CMV Stimulation T cells from CMV-seropositive and seronegative individuals were stimulated with sCMV-Ag and cCMV-Ag for 2–5 days and were stained for expression of CD4+ and CD8+ T cell subset markers and T cell activation markers: CD25 bright (·-chain IL-2 receptor) and CD69 (early T cell activation antigen). The data were expressed as percent-activated T cells expressing CD25 or coexpressing CD25 and CD69 antigens. Data were acquired on a four-color Beckman-Coulter XL Cytometer and analyzed using WinList (Verity Inc, Topsham, Me., USA). T Cell Proliferation Assays Isolated CD3+ T cells and sorted CD4+ and CD8+ T cells at a concentration of 105 cells/well were cultured in RPMI-1640 plus 5% human serum with sCMV-Ag (1:200 dilution) and autologous irradiated (4,000 rad) peripheral blood mononuclear cells (5 ! 104 cells/ well). The same cell preparations were also cultured with irradiated (10,000 rad) CMV-infected epithelial cells (A549/CMV) or with uninfected cells (0.37 ! 104/well). All of the cultures were incubated for 6 days in 37 ° C with 5% CO2, pulsed with 1 ÌCi of 3H-thymidine/ well for 18 h, harvested and counted in a liquid scintillation counter. Results were expressed as counts per minute. Response to CMV stimulation was considered positive if the counts per minute were 3-fold over the background and greater than 1,000 [6]. Two-Color Flow Cytometry for Intracellular Cytokine Detection The procedure used in this study was adapted from a previously published method [7]. Briefly, unsorted CD3+ T cells and sorted CD4+ and CD8+ T cells were cultured for different periods with sCMV-Ags and cCMV-Ags. In the last 4 h of incubation, Brefeldin A (5 Ìg, Sigma) was added to the culture to prevent the cytokine release from the cells. For positive control of intracellular cytokine production, we stimulated the CD3+ T cells with PMA/Ca-ionophore for 4 h [8]. Cells collected after various types of stimulation were washed and stained for surface markers with fluorescein isothiocyanate (FITC)-labeled CD8 mAb (mouse anti-human IgG1Î, HIT8a). Then, the cells were fixed with 4% paraformaldehyde overnight, permeabilized with 1% saponin and stained for intracellular cytokines (all commercially available from Pharmingen). The anti-cytokine antibodies were labeled with phycoerythrin (PE) and consisted of IL-2 (rat anti-human IgG2a), IFN-Á (mouse anti-human IgG1Î, 4S.B3) and IL-4 (rat anti-human IgG1, MP4-25D2). Isotype control antibodies for intracellular cytokine staining consisted of FITC-conjugated mouse anti-human IgG1, PE-conjugated mouse anti-human IgG1 and PE-conjugated rat anti-human IgG2a, each used at comparable concentrations of the antibody of interest. The production of cytokines was determined by detection of PE staining. Quadrant statistics were based on the staining of the negative isotype controls. Results were expressed as percent activated CD4+ or CD8+ T cells. In addition, we calculated the total number of CD4+ and CD8+ T cells per million peripheral blood lymphocytes. Cytokine-producing cells were also subdivided on the basis of intensity of cytokine staining into ‘high’ and ‘low’ producers by dividing the positive quadrant into bright (high) and dim (low) areas. Quantitative RT-PCR (QRT-PCR) for Granzyme B mRNA Expression Quantitative gene analysis was performed by a competitive PCR method as described previously [9, 10]. RNA was extracted from

Zeevi et al.

Fig. 1. Expression of T cell activation marker CD25 on CD4+ and CD8+ T cells cultured for 5 days with sCMV-Ag

and autologous APCs. The frequency of CD4+CD25+ T cells was 6-fold higher than that of the CD8+CD25+ T cells.

BAL cells, measured and reversed-transcribed to cDNA. Patientderived cDNA (WT) was co-amplified with a known amount of a target gene competitor (CT), which is amplified with the same primers but is shorter. PCR products were separated by agarose gel electrophoresis, stained with ethidium bromide, and analyzed by densitometry. Standard curves were generated for granzyme B (GB) and the housekeeping gene GAPDH (GAP). Results were expressed as ratio of GB/GAP to control for the accuracy of RNA measurement and the efficacy of reverse transcription.

Cytomegalovirus-Specific T Helper Responses in Lung Transplant Recipients

Results

Expression of T Cell Activation Markers CD25 and CD69 Antigens on CMV-Stimulated T Cell Subsets A small proportion of CD4+ T cells (5–10%) stimulated with sCMV-Ag presented by autologous APCs expressed low levels of CD25 after 2 days in culture while after 5 days of sCMV-Ag stimulation, 69% of CD4+ T cells expressed CD25. In contrast, the expression of CD25


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Fig. 2. Expression of T cell activation marker CD25 on CD4+ and CD8+ T cells cultured for 5 days with cCMV-Ag

(CMV-infected A549 cells). The frequency of CD25+ T cells was low and similar in both T cell subsets.

on CD8+ T cells stimulated by sCMV-Ag remained low throughout the 5-day culture period (10% CD8+CD25+) (fig. 1). The majority (95%) of CD4+CD25+ T cells also expressed the early T cell activation marker CD69 by day 5 while only 60% of CD8+CD25+ T cells co-expressed the CD69 antigen (data not shown). The pattern of T cell activation following cCMV-Ag stimulation was different from the sCMV-Ag stimulation. The proportion of CD4+ and CD8+ T cells expressing CD25 following stimulation

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with cCMV-Ag was low (7 and 6%, respectively) (fig. 2). In addition, the coexpression of CD25 and CD69 T cell activation markers was mainly on the CD4+ T cell subset. 74% of CD4+ CD25+ T cells were CD69+ while only 16% of CD8+ CD25+ T cells coexpressed CD69 antigen (data not shown). T cells from a seronegative individual stimulated with either CMV-Ag did not express the T cell activation antigens CD25 and CD69.

Zeevi et al.

CD3 80,000

CD4

3H-Thymidine uptake (net cpm)

CD8 60,000

40,000

20,000

0 sCMV-Ag

sCMV-Ag

Fig. 3. Proliferatiaon of CD3+ T cells and of sorted CD4+ and CD8+ T cell subsets stimulated with sCMV-Ags and cCMV-Ags. The proliferative response was measured by 3H-thymidine uptake (y-axis). Significant proliferation was observed with sorted CD4+ T cells stimulated with sCMV-Ag + self-APC while the response of sorted CD8+ T cells was minimal. Both T cell subsets responded to cCMV-Ag stimulation.

Proliferative Responses of Sorted T Cell Subsets Stimulated with CMV-Ags We tested the proliferative responses of isolated CD3+ T cells and sorted CD8+ and CD4+ T cells stimulated with the two types of CMV-Ags: sCMV + autologous APC and CMV-infected A549 cells (cCMV-Ag). As shown in figure 3, sCMV-Ag stimulation resulted in significant proliferation of CD3+ and CD4+ T cells. The proliferative response of sorted CD8+ T cells stimulated by sCMV-Ag was minimal. These results indicate that the Th memory response triggered by sCMV-Ag resides in the CD4+ T cell subset. In contrast, stimulation with the cCMV-Ag induced a similar level of proliferation of CD4+ and CD8+ T cell subsets. Although both T cell subsets responded to CMV-infected (but not to uninfected) A549 epithelial cells, neither subset achieved the level of proliferation of CD4+ T cells stimulated with sCMV-Ag (fig. 3). Intracellular Cytokine Production in T Cell Subsets Stimulated with Two Types of CMV-Ag We compared the frequency of IFN-Á-producing CD4+ and CD8+ T cells following optimum T cell activation in vitro (4 h of stimulation with PMA/Ca-ionophore) vs. 5 day of coculture with sCMV-Ag or cCMV-Ag. As


shown in figure 4, most of CD4+ and CD8+ T cells (67 and 84%, respectively) were activated after in vitro stimulation with PMA/Ca-ionophore. The majority of CD8+ T cells were bright (65%) while only 27% of CD4+ T cells were bright (high producers). Following sCMV-Ag stimulation, the frequency of activated CD4+ T cells producing IFN-Á and the distribution of dim and bright cells was similar to the positive control (PMA/Ca-ionophore). In contrast, the percent CD8+ T cells producing IFN-Á after 5 days in culture with sCMV was 2-fold lower than in the positive control (46 vs. 84%, respectively) and the percent of bright CD8+ T cells was significantly diminished (28% vs. 65%, respectively) (fig. 4). The percent of T cell subsets stimulated with cCMV to produce IFN-Á was 3- to 2-fold lower than following PMA/Ca-ionophore and sCMV-Ag stimulation and most of the cells were dim (low producers) (fig. 4). The frequency of cytokine-producing T cells was assessed early following CMV activation (6– 48 h) and after 3–8 days in vitro stimulation. Early after CMV activation, we could not detect any intracellular cytokine production (data not shown). The peak response occurred between 5 and 7 days of coculture with CMVAgs. Both T cell subsets exhibited similar kinetics of cytokine production following CMV activation. However, the total number of sCMV-Ag specific CD4+ T cells positive


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750,000 CD4-IL2

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CD4-IFNg 450,000 CD4-IL4 300,000 CD8-IL2

150,000 CD8-IFNg

CD8-IL4

0 5

6 7 Days in culture with sCMV (ratio CD4/CD8 3:1)

8

Fig. 5. Kinetics of intracellular cytokine production in T cell subsets following sCMV-Ag

stimulation. Results were expressed as number of CD4+ or CD8+ cells per million peripheral blood lymphocytes (PBL) (y-axis). All three cytokines IL-2, IFN-Á and IL-4 were produced by both T cell subsets. The peak response was seen after 6–7 days in culture. The ratio of CD4/ CD8 cells in these cultures was 3:1 respectively and the total number of CD4+ T cells producing cytokines was also 4- to 5-fold higher than the CD8+ T cells.

for intracellular cytokine (IL-2, IL-4 and IFN-Á) production was 4- to 6-fold higher than the total number of cytokine-producing CD8+ T cells. Although the frequency of activated CD4+ and CD8+ T cells might be similar following sCMV-Ag stimulation (fig. 4), the total number of CD4+ T cells in these cultures was higher than the CD8+ T cells (ratio CD4/CD8 3:1) (fig. 5). To further characterize the contribution of T cell subsets to the cytokine pool, we determined the frequency of cytokine-positive CD4+ and CD8+ T cells sorted prior to stimulation with CMVAgs. Over 40% of the CD4+ T cell subset produced all three cytokines tested and there was no difference between the unsorted and sorted CD4+ T cells stimulated with sCMV-Ag (fig. 6). Similarly, the frequency of CD8+ T cells positive for the various cytokines in the unsorted and sorted populations did not change. There was a trend to a slight elevation in the IFN-Á-producing CD8+ sorted T cells (fig. 6). A similar pattern with unsorted and sorted

Fig. 4. Frequency of intracellular IFN-Á production in T cell subsets co-cultured with PMA/Ca ionophore for 4 h or with sCMV-Ag and cCMV-Ag for 5 days. Two-color flow-cytometric analysis was performed with FITC-labeled anti-CD8 mAb and PE-labeled anti-IFN-Á mAb. The percent of dim (low producers) and bright (high producers) was calculated for each condition. For each condition, an IgG control mAb was also used.


T cell subsets was observed following cCMV-Ag stimulation. Both T cell subsets were equally activated with the exception of sorted CD4+ IL-4 producing cells. Overall, the sCMV-Ag activation generated a 2-fold higher response in T cells than the cCMV-Ag stimulation (fig. 6). Expression of GB mRNA in BAL Cells during CMV Pneumonia and Correlation with Acute Lung Rejection We determined the mRNA levels for GB in BAL cells of lung transplant recipients before, during and after CMV pneumonia (CMV-P). We employed a quantitative RT-PCR method to determine the level of GB in BAL cells. The data were expressed as a ratio of GB to the housekeeping gene GAP. Two patterns were observed in lung transplant recipients and they are illustrated in the examples given in table 1. Patient A, who was CMV seronegative before transplantation and received a CMVpositive donation, was at high risk of developing CMV-P after transplantation. The patient had a low ratio of GB/ GAP (20 ! 10 –3) prior to diagnosis of CMV-P (day 119 after transplantation). During CMV-P, the GB/GAP ratio increased significantly (34-fold) and remained elevated as the patient experienced acute cellular rejection (ACR3) on day 161 after surgery. At this time, the histological diagnosis was mixed with evidence of CMV inclusions and cellular infiltrates, possibly associated with acute cel-


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IL-2

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CD8u+

CD8s+

Fig. 6. Intracellular cytokine production in T cell subsets stimulated with two types of CMV-Ags. CD3+ T cells were cocultured with CMV-Ags and then stained for CD8 and the specific cytokines (unsorted CD4u+ and CD8u+ T cells). Prior to stimulation, CD8+ and CD4+ T cells were sorted and then cultured for 5 days with the two types of CMV-Ags (sorted CD4s+ and CD8s+ T cells).

Reactive T cells (net) (%)

60

40

20 ND 0

CD4u+

lular rejection. In contrast, the second patient (B) had no cellular rejection before or immediately after the episode of CMV-P and his GB/GAP ratios remained low throughout the follow-up period (12 to 0 !10 –3). Since this patient was CMV seropositive prior to transplantation and received a CMV-negative donation, he might have experienced only a transient reactivation of his own CMV.

Discussion

In the current study we investigated the phenotype and functional characteristics of CMV-specific Th memory responses in seropositive controls and in lung transplant

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CD4s+ CD8u+ Unsorted (u) and sorted (s) T cell subsets

CD8s+

recipients. We have shown that in seropositive individuals, the pool of CMV-specific Th memory cells consists of two types of Th cells [3]. These are: the Th memory cells that are stimulated by sCMV-Ag and are restricted by selfAPC, and the Th memory cells that respond to cCMV-Ag in the absence of self-APC. The CMV-specific Th memory response initiated by nominal CMV-Ag (sCMV-Ag + self-APC) is primarily associated with CD4+ Th subset, as shown by the expression of T cell activation markers CD25 and CD69 (fig. 1). These findings are also supported by the proliferative responses of sorted T cell subsets. CD4+ T cells responded vigorously to sCMV-Ag while the sorted CD8+ T cells exhibited a minimal response (fig. 3). In contrast, the cCMV-Ag in the absence of self-APC stimulated both T cell subsets equally (fig. 2, 3). Although

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Table 1. Detection of GB mRNA in BAL

A Patient A: CMV–/donor CMV+

cells of lung transplant recipients before, during and after CMV-P

Postoperative day

Biopsy histology Immunosuppression

Lymphocytes in BAL, % GB/GAP ! 10 –3 a

119

135

161

neg. Neo ACsA imuran steroid 14 20

CMV-P/AR? Neo stop ACsA stop imuran steroid 18 680

AR3 Neo Sol ! 3 g steroid 28 150

B Patient B: CMV+/donor CMV– Postoperative day 57 Biopsy history Immunosuppression Lymphocytes in BAL, % GB/GAP ! 10 –3 a a

neg. tacrolimus steroid 2 12

80 CMV-P tacrolimus steroid 8 5

120 lymphocytic bronchitis tacrolimus steroid 1 0

The results are expressed as the ratio GB/GAP ! 10 –3. Neo = Neosporine; ACsA = aerosol cyclosporin; Sol = solumedrol.

the proliferative capacity of T cell subsets is different following CMV-Ag stimulation, both T cell subsets respond to CMV-Ags by producing various cytokines (fig. 4–6). These results support the contention that CMV infection within the allograft may promote the recruitment and activation of both T cell subsets. The release of proinflammatory cytokines from CMV-activated T cells (IL-2, IFN-Á) within the allograft may enhance graft antigenicity and promote allograft rejection. This enhanced alloreactive response in the presence of CMV infection is manifested by upregulation of cytotoxic granule gene expression GB (table 1). GB is expressed in activated cytotoxic T cells and is significantly associated with acute cellular rejection of kidney allografts [9, 11]. In lung recipients, we have also seen a strong correlation between the expression of GB mRNA greater than 40 (ratio of GB/GAP) and acute cellular rejection [unpubl. results]. In those patients in whom the process of rejection and CMV infection occurred concomitantly, the levels of GB increased significantly (table 1). Those patients also had more severe forms of allograft rejection. Conversely, lung recipients that had a suppressed immune system associated with disseminated CMV disease (retinitis, gastritis) did not ex-

press high levels of GB mRNA within the allograft. We have previously shown that the lack of CMV-specific Th memory response (sCMV-Ag + self-APC) was associated with a risk of recurrent CMV disease or disseminated CMV disease [3]. However, even in the absence of Th memory response to nominal CMV Ag, we could still demonstrate the presence of CMV-dependent Th response to CMV-Ag [6]. Potentially, infected allograft endothelial cells or epithelial cells that harbor low levels of CMV infection may recruit and activate host T cells. Waldman et al. [12] have shown that infected donor endothelial cells are resistant to cell-mediated cytotoxicity, but they promote the generation of allogeneic cytotoxic T cells. These allogeneic cytotoxic T cells can kill uninfected endothelial cells and induce allograft damage through cytokines and cytotoxic effector molecules (perforin, GB).



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References 1 Boland GJ, Hene RJ, Ververs C, de Haan MA, de Gast GC: Factors influencing the occurrence of active cytomegalovirus (CMV) infections after organ transplantation. Clin Exp Immunol 1993;94:306–312. 2 Meyers J, Flourny N, Thomas E: Risk factors for cytomegalovirus infection after human marrow transplantation. J Infect Dis 1986;153: 478–483. 3 Zeevi A, Morel P, Spichty K, Dauber J, Yousem S, Williams P, Grgurich W, Pham S, Iacono A, Keenan R, Duquesnoy R, Griffith B: Clinical significance of CMV-specific T helper responses in lung transplant recipients. Hum Immunol 1998;59:768–775. 4 Waldman WJ, Adams PW, Orosz CG, Sedmak DD: T lymphocyte activation by cytomegalovirus-infected, allogeneic cultured human endothelial cells. Transplantation 1992;54:887– 896.

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5 Sedmak DD, Roberts WH, Stephens RE, Buesching WJ, Morgan LA, Davis DH, Waldman WJ: Inability of cytomegalovirus infection of cultured endothelial cells to induce HLA class II antigen expression. Transplantation 1990;49:458–462. 6 Zeevi A, et al: Two types of cytomegalovirusspecific memory responses: Potential role of donor infected target cells in recruitment of activated T cells to the graft. Monogr Virol Basel Karger, 1997, vol 21, pp 120–133. 7 Anderson J, Anderson V: Characterization of cytokine production in infectious mononucleosis studies at a single-cell level in tonsil and peripheral blood. Clin Exp Immunol 1993;92: 7–13. 8 Zeevi A, et al: High frequency of intragraft IFN-g producing CD8+ T cells in predictive of lung rejection. J Heart Lung Transplant 1999; 18:82.


9 Pavlakis M, Strehlau J, Lipman M, Strom TB: Use of intragraft gene expression in the diagnosis of kidney allograft rejection. Transplant Proc 1996;28:2019–2021. 10 Pavlakis M, Strehlau J, Lipman M, Shapiro M, Maslinski W, Strom TB: Intragraft IL-15 transcripts are increased in human renal allograft rejection. Transplantation 1996;62:543–545. 11 Strehlau J, Pavlakis M, Lipman M, Shapiro M, Vasconcellos L, Harmon W, Strom TB: Quantitative detection of immune activation transcripts as a diagnostic tool in kidney transplantation. Proc Natl Acad Sci USA 1997;94:695– 700. 12 Waldman W, Knight D, Adams P: Cytolytic activity against allogeneic human endothelia: Resistance of CMV-infected cells and virally activated lysis of uninfected cells. Transplantation 1998;66:67–77.

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Part I. Immunopathology Immune Escape and Reactivation


Human Cytomegalovirus Escape from Immune Detection Susan Michelson Unité d’Immunologie Virale, Institut Pasteur, Paris, France

Key Words Cytomegalovirus W Immune escape W Chemokine sequestration W HLA

Abstract Human cytomegalovirus (CMV) has devised numerous means of escaping immune surveillance. The CMV genome encodes at least 4 genes involved in downregulating surface expression of HLA class I molecules. In addition, it sequesters CC chemokines, induces Fc receptors, interferes with induction of HLA class II antigens, and can inhibit natural killer cell activity. CMV can efficiently block the presentation of immediate early antigens, the first viral proteins to be produced. Together, these mechanisms probably contribute to the ability of CMV to persist in its host and may play a role in the immunopathology of CMV disease. Copyright © 2000 S. Karger AG, Basel

Many viruses have devised ways to avoid detection and elimination. This has allowed them to survive persistently or latently in their hosts over centuries. Herpesviruses are archetypes of such host-virus relationships. Over millions of years, herpesviruses have perfected their genome products to confront recognition and aggression by the immune system [1].

ABC

© 2000 S. Karger AG, Basel 0300–5526/99/0426–0301$17.50/0



The mechanisms of immune evasion discussed here will also address ways in which cytomegalovirus (CMV) can counter secondary effects of immune responses, such as induction of oxygen radicals and precipitation of apoptosis. Of course, the simplest way to avoid recognition is not to express any viral genes. Human CMV employs this technique. It establishes itself in latency in circulating monocytes [2], bone marrow progenitor populations [3–7] including circulating CD34+ [8] and CD14+ cells [9], and probably endothelial cells [10]. At these sites, it refuses to express itself as long as the host remains healthy or cells are stimulated by inflammatory cytokines [11]. Further evidence of this nonexpression is witnessed by the fact that a substantial number of seronegative, healthy individuals are nonetheless PCR positive [12]. ‘Latency transcripts’ have been described in different bone marrow progenitor cells of about 30–40% healthy individuals [4]. Detection of antibodies to proteins potentially encoded by ‘latency open reading frames’ has not been published. Thus, individuals who are CMV seronegative but PCR positive may actually be ‘seropositive’ for these and other proteins still to be studied. The significance and consequences of such ultra-low-level viral expression remain to be defined, both in the host and in the recipient of material from such carriers. Viruses may even take avantage of nonspecific responses to infection, like the presence of inflammatory

Susan Michelson Unité d’Immunologie Virale, Institut Pasteur 28, rue du Dr-Roux, F–75724 Paris Cedex 15 (France) Tel. +33 1 45 68 82 64, Fax +33 1 45 68 89 41 E-Mail [email protected]

cytokines, to maintain a low profile. Such would be the case when CMV infects a cell in which viral expression is aborted at the immediate early (IE) stage. We remind the reader that IE protein expression is not synonymous with full virus replication [13]. An example of abortive infection with IE expression is the inhibition of CMV replication by interferons (IFN), principally IFN-Á and IFN-ß [14–16]. Although full viral replication is blocked, expression of IE proteins occurs [14, 16]. This means that strategies of immune evasion must be called into play to avoid not only detection, but also destruction of infected cells. The inflammatory cytokines IFN-Á and IL-ß, when present together, activate the inducible form of nitrous oxide synthase in certain epithelial cells [16]. CMV can downregulate this enzyme activity, thereby protecting itself from destruction due to free-radical generation. CMV also appears to have found ways to avoid destruction from apoptosis, which results from induction of TNF during infection or from CMV-induced DNA damage [17, 18] and cell cycle arrest [19–22]. IE proteins appear to protect infected cells from destruction by apoptosis induced by TNF [23] or other pathways [24]. IE proteins also bind to and inactivate p53 in coronary smooth muscle cells [25]. In endothelial cells [26], CMV infection results in sequestration of p53 in the cytoplasm, thereby blocking apoptosis induced by serum starvation and mediated by p53 activation. There are some indications that CMV may manipulate p53-mediated apoptosis in vivo [27]. In various tissues of CMV-infected patients, no apoptosis was seen, and CMV cells positive by in situ hybridization or immunochemistry were seen to express p53. However, in CMV-bearing cells, there was simultaneous expression of MDM2 in the absence of p21/WAF1. This pattern of expression is compatible with inactivation of p53 [reviewed in ref. 28]. CMV has also developed at least four direct means to avoid detection of IE protein expression (discussed below). Some of these mechanisms also help CMV to evade immune detection as the viral cycle progresses. One of the first events is the nonpresentation of IE proteins at the cell surface. The phosphoprotein pp65 (UL83), which enters cells along with virions and dense bodies, is involved in the phosphorylation of IE1, thereby impeding its presentation at the cell surface [29]. We have studied the persistence of pp65 in fibroblasts in which de novo pp65 expression is blocked and found that protein persists in the nucleus for up to 16 h [unpubl. results]. Such persistence means that pp65 would be continually present from the time of viral entry until its de novo production, thereby assuring the nonpresentation of IE proteins.

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Since IFN can induce MHC class II presentation, CMV manages to block IFN induction of HLA II in CMV-infected cells ([30] and see Sedmak in this issue for details). A third means of avoiding detection during the IE period is an imposed block of surface expression of MHC class I by the product of US3. US3 is a short-lived, IE protein that is synthesized between 1 and 4 h after infection. It is an endoplasmic reticulum (ER) resident protein which retains stable, peptide-loaded MHC class I molecules in the ER compartment without increasing the degradation rate of class I [31]. Another CMV gene product which affects MHC class I presentation, US2, appears within 3 h after infection [32]. It causes rapid dislocation of newly formed heavy chains from the ER to the cytosol where they are degraded [32]. This involves a Sec61 translocation complex [33]. As viral genome expression increases, the virus calls into play additional genes to avoid detection. One of the main means to accomplish this is by rendering cells incapable of presenting antigens through continued downregulation of HLA class I and class II antigens [30]. Effectively, the CMV genome has two other genes whose products interfere with MHC class I surface expression, US6 and US11 (fig. 1). US6 [34] is another short-lived, early ER resident, transmembrane protein [35]. This protein coprecipitates as a TAP-ß2-microglobulin-class I complex. US6 associated with TAP prevents peptide translocation. US11, also an ER resident protein, causes rapid destruction of both free and ß2-microglobulin-associated class I molecules. The product of US11 dislocates class I molecules from the ER to the cytosol where they are attached by N-glycanase and proteasomes [36]. US2 and US11 gene products do not, however, affect expression of HLA-G and HLA-C on trophoblast cells infected with vaccinia viruses expressing these CMV genes [37]. US2 and US11 also fail to co-immunoprecipitate with the latter HLA molecules. There is further suggestion in the literature that US2 and US11 may differentially affect HLA class I breakdown depending on their allelic form [38] and/or the cell type [32]. While other CMV gene products may regulate surface expression of HLA-G and HLA-C, it would certainly be astute of CMV not to affect expression of these HLA classes since they are noted for their inhibitory effect on NK activity [39]. In addition, HLA-G appears to associate with the same peptides as does HLA class I, suggesting that the immune system might then become tolerant to such peptides. Perhaps CMV infection even stimulates HLA-G expression? However, since HLA-G expression is dependent on TAP func-

Michelson

Fig. 1. Immune escape mechanisms deployed by human CMV. Human CMV genes or indirect mechanisms, which interfere with immune detection of infected cells, are given in boxes surrounding the central cell. The numbers in parentheses indicate articles concerning each point.

tion [see references in Lanser, 39] and CMV interferes with TAP [34], HLA-G molecules may not escape from CMV-induced dysfunction of TAP. CMV downregulation of certain enzymatic activities might also affect antigen presentation. Phillips et al. [40] described downregulation of the cellular aminopeptidases CD10 and CD13 during CMV infection. These membrane enzymes are thought to play a role in trimming peptides during their insertion into the grooves of MHC class I and II molecules [41, 42]. Hence, reduction of these enzymatic activities could further affect surface expression and stability of MHC molecules. CMV codes for its own HLA-class-I like molecule [43], designated here CMV HLA (fig. 1). Only about 39% of the ß2-microglobulin contact points with the 2m ·-chain are conserved in CMV HLA, while those with the ß 2m contacts are conserved [44]. The CMV HLA can bind

peptides [45]. This gene is found expressed in peripheral blood monocytes in transplanted patients [46]. It can inhibit NK cell activity [47, 48] through rather selective binding [49] to an LIR-type (also called ILT) inhibitory receptor. Not all authors have found that CMV HLA downregulates NK lysis [50]. Refractoriness to NK lysis also apparently involves levels of expression of LFA-3 expression [51], CD94 [47], and LFA-1 [52] on CMVinfected cells. Clinical isolates differed in their ability to upregulate or downregulate LFA-3 expression, while all isolates studied downregulated HLA class I to the same extent [51]. Fletcher et al. [51] also demonstrated cell type variations and the probable implication of both IE/early and late gene products. The fourth approach CMV has adopted to escape immune surveillance is to impede infiltration and inflammation by interfering with CC chemokine secretion. Che-

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303

mokines are small (8–10 kD), secreted proteins which are responsible for attracting circulating leukocytes to sites of infection and injury [reviewed in ref. 53]. In addition, they are necessary for the full activation of lymphocyte effector mechanisms [54, 55]. Chemokines are divided into four subfamilies depending on the number and disposition Nterminal cysteine residues: CXC, CC, CX3C and C. Their receptors are correspondingly designated CXCR 1–5, CCR 1–9, CX3CR1 and CR1. With few exceptions, chemokines of a given subfamily bind only to receptors of the same subfamily. One exception is the Duffy antigen which binds CXC and CC chemokines; binding does not elicit an intracellular signal. Another notable exception is the receptor encoded by human herpesvirus-8 (KSHV-GPCR), which is constitutively active and binds both CXC and CC chemokines [56]. Lastly, the US28 encoded by CMV, which may also be constitutively active, binds CX3C as well as CC chemokines (see below). The CMV genome contains four genes (UL33, US27, US28 [57] and UL78 [58]) which show homology to chemokine receptors. US28 is transcribed just after IE genes and before pp65 [59] in vitro and is expressed in vivo [60]. UL33 is an early gene and US27 is expressed with late kinetics [61]. The ligand for receptors UL33 and UL78 are unknown. However, study of the murine [62] and rat [63] homologues of UL33 (M33 and R33, respectively) demonstrated that this protein plays a role in homing virus to salivary glands and establishment of viral latency at this site. Study of the rat homologue of UL78 shows that deletion of this protein significantly lowers viral replication in vitro, prolongs survival in vivo, and leads to the formation of syncytia [64]. In the context of the infected cell, US27 and US28 bind the CC chemokines, RANTES and MCP-1 with high affinity while US28 also binds MCP3 and MIP1· and ß [61]. In addition, US28 also binds the CX3C chemokine fractalkine, a membrane-bound chemokine with a cytoplasmic domain [65]. Finally, US28 is involved in cellto-cell fusion [66], which could presumably allow virus to pass intercellularly without being exposed to neutralizing antibodies. US27 and US28 sequester chemokines from the environment of infected cells [61, 67–69]. Although CMV infection of fibroblasts induces production of RANTES, this chemokine disappears from supernatants with progression of the viral cycle. Similary, MCP-1, which is constitutively produced by these cells, disappears from the medium. When cells are infected with virus deleted of US28 or US27 and US28, RANTES and MCP-1 are continually found in the supernatants.

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When RANTES is added exogenously to cells late after infection, it disappears from the supernatant. This observation, coupled with the intracellular accumulation of RANTES detected by immunofluorescence [67], suggested that RANTES may be sequestered within the cell as it is made and/or that it is being continually internalized from the extracellular medium. To investigate these possibilities, biotin-labeled RANTES was added extracellularly to fibroblasts infected for 72 h [61]. After stripping off external RANTES from the cell surface, cells were lysed and immunoblotted with peroxidase-labeled streptavidine to detect intracellular biotinylated RANTES. RANTES was internalized by cells infected with wildtype CMV, but not by cells infected with a US27/US28 deletion mutant or by uninfected fibroblasts. Surprisingly, infected cells expressing either US28 or US27 alone also internalized RANTES. These results demonstrate that US27 is also a functional receptor for RANTES and that both receptors are continually internalizing chemokine from the extracellular medium. The results also suggest that these receptors may be constitutively active. Their avidity is considerable since they can simultaneously sequester at least two chemokines, RANTES and MCP1, present at 1100 pM and 110 nM concentrations, respectively. Even when these chemokines are superinduced by TNF they disappear almost completely from supernatants of infected cells [61]. Results presented by Randolph-Habecker et al. [70] and our unpublished observations show that monocyte chemoattraction is substantially reduced to supernatants conditioned on CMV-infected fibroblasts. We have further observed that supernatants of cells infected with ¢US28 or ¢US27/US28 viruses retain chemotactic activity for monocytes unlike those from cells infected with wild-type CMV. These observations strongly suggest that chemokine sequestration by CMV-infected cells could indeed impede leukocyte infiltration. Several reports demonstrate that engagement of MHC class I and II by bacterial or viral antigen results in the production of chemokines (RANTES, MCP-1, IL-8, IP-10) [71–75]. This could be the consequence of prior induction of TNF and IL-1ß [76]. In any case, CC chemokines are necessary for activation of leukocyte effector mechanisms [54, 55, 77]. It therefore remains to be seen whether CMV-related sequestration of CC chemokines can also affect lymphocyte and monocyte effector mechanisms. CMV has also devised a simple strategy for combating and neutralizing cytotoxic CMV-specific antibodies through the induction of Fc receptors [78, 79]. Though originally described as requiring viral DNA synthesis

Michelson

[80], they were subsequently found to be expressed on human and mouse cells abortively infected with human CMV [81]. Infection leads to a 100-fold increase in Fc receptor expression on infected fibroblasts [82]. These receptors are also expressed on infected endothelial cells [83]. Fc receptors are found in the tegument of purified virus where proteins of 69 and 33 kD are involved in IgGFc binding [84]. The induced FcR are not recognized by antibodies directed against human FcÁRI, RI or RIII [83]. The CMV-induced receptors bind all 4 classes of IgG, but not IgA or IgM [85, 86]. The rank of binding affinities of these receptors for IgG subclasses is IgG16IgG41IgG21IgG3 [86]. The isotype with the highest affinity corresponds to that in which the majority of anti-CMV antibodies are found in human sera [87, 88]. More than 96% of CMV-reactive immunoglobulins are of the IgG1 isotype. Although CMV-reactive IgG3 constitute only about 3% of anti-CMV-reactive antibodies, this

isotype has 10-fold more neutralizing activity than IgG1 isotype antibodies [87]. Thus, CMV induction of Fc receptors may severely affect antibody-mediated immune responses. Human CMV, like all living organisms, strives to survive. The multiple mechanisms it uses to avoid being detected by the immune system has allowed it to survive in its host and evolve with it. Lytic CMV replication, except in the setting of AIDS, is probably not the main cause of disease. Anatomopathological studies using immunochemistry and in situ detection of viral DNA or RNA often reveal the presence of CMV at disease sites in the absence of late viral products. In murine CMV, disease may occur where little or no lytic replication occurs [89, 90]. One can only wonder what role the consequences of CMV immune evasion are playing in the development of disease.

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Human Cytomegalovirus Reactivation in Bone-Marrow-Derived Granulocyte/Monocyte Progenitor Cells and Mature Monocytes Susanna Prösch a Wolf-Dietrich Döcke b Petra Reinke c Hans-Dieter Volk b Detlev H. Krüger a Departments of a Virology, b Medical Immunology and c Nephrology/Internal Intensive Care Medicine, School of Medicine (Charité), Humboldt University, Berlin, Germany

Key Words Human cytomegalovirus W Virus reactivation W Monocyte/granulocyte progenitor cells W Systemic inflammation W TNF-· W Stress W cAMP

Abstract Monocyte/granulocyte progenitor cells of the bone marrow are a major site of human cytomegalovirus (HCMV) latency. The mechanisms of establishment and maintenance of HCMV latency are still unknown. Reactivation of the latent virus in bone-marrow-derived progenitor cells has been demonstrated in vitro and suggested to occur also in vivo. Clinical studies have shown that reactivation is a rather frequent event not only in immunosuppressed but also in nonimmunosuppressed patients and in healthy blood donors. At least three independent mechanisms of virus reactivation are discussed: systemic inflammation connected with strong tumor necrosis factor alpha release; application of cAMP-elevating drugs, and highly stressful events associated with increased plasma catecholamine levels. Copyright © 2000 S. Karger AG, Basel

ABC

© 2000 S. Karger AG, Basel 0300–5526/99/0426–0308$17.50/0



Latency of Human Cytomegalovirus

After primary infection, human cytomegalovirus (HCMV), like other herpesviruses, establishes life-long latency in the infected individual. Under particular circumstances, the virus can be reactivated, causing either clinically asymptomatic HCMV infection or HCMV disease depending on the constitution of the immune system. Based on seroconversion, about 50–60% of adults in Central Europe are latently infected with HCMV. In addition, screening of peripheral blood mononuclear cells (PBMC) from healthy blood donors for HCMV DNA by PCR has indicated that at least 5–20% of seronegative individuals also harbor latent virus [1–6]. Using monocyte-enriched PBMC, Larsson et al. [6] found that even up to 55% of seronegative blood donors harbor latent HCMV. Altogether, this suggests that at least 80% of adults may be latently infected with HCMV. Over the last few years it has been demonstrated that peripheral CD14+ blood monocytes, and in particular their progenitor cells, CD34+ granulocyte/monocyte progenitor cells in the bone marrow, are a major site of HCMV latency in vivo [1, 5, 7–11]. The frequency of latently infected cells in healthy blood donors is very low, ranging from 0.004 to 0.01% of mononuclear cells de-

PD Dr. Susanna Prösch Institut für Virologie, Universitätsklinikum Charité Campus Charité-Mitte, Humboldt-Universität Schumannstrasse 20/21, D–10098 Berlin (Germany) Tel. +49 30 2802 3117, Fax +49 30 2802 2180, E-Mail [email protected]

rived from granulocyte-colony-stimulating-factor-mobilized peripheral blood or from bone marrow cells with only 2–13 viral genome copies per latently infected cell [12]. During latency the viral genome persists as a circular plasmid [13]. While no lytic gene expression could be detected in latently infected monocytes or in progenitor cells from seropositive carriers which tested positive for viral DNA [9, 14], Kondo et al. [10, 15] and Kondo and Mocarski [16] have described novel latency-associated HCMV transcripts in progenitor cells latently infected in vitro as well as in progenitor cells from normal healthy blood donors. The latency-associated transcripts were found to be of sense and antisense orientation corresponding to the immediate early (IE) gene region and are suggested to encode three novel proteins. Antibodies against two of the three proposed proteins could be detected in the serum of healthy blood donors [10]. The role of these latency-associated transcripts and the predicted immunogenic proteins for maintenance of latency and reactivation is still unknown. Reactivation of latent HCMV from experimentally infected CD34+ granulocyte/monocyte progenitor cells as well as mature monocytes has been shown by different groups [5, 15, 17, 18]. Taylor-Wiedeman et al. [14] demonstrated partial HCMV gene expression (IE genes) in latently infected monocytes from healthy donors after differentiation in vitro. Similarly to HCMV, mouse CMV (MCMV) has also been found to establish latent infection in bone-marrowderived hematopoietic cells of virus-infected mice [19, 20], where the virus could be reactivated ex vivo [21]. There is no doubt that reactivation of latent HCMV is not a rare event in patients with severe immunosuppression, such as transplant recipients and HIV-infected individuals and that it leads to active HCMV infection and disease. However, temporary reactivation of HCMV can also be observed in people with diminished immune responsiveness, such as elderly people [our unpubl. results] and in nonimmunosuppressed patients with septic disease [4, 22, 23] or chronic liver disease [4], following trauma [24, 25], psoriasis [26] or myocardial infarction [our unpubl. results] or even in healthy individuals [6, 26, 27]. Screening PBMC samples from seropositive healthy blood donors over longer observation periods by a PCR technique not sensitive enough to detect latent virus demonstrated several short-term increases in HCMV DNA which may be the result of ‘oscillating’ low-level virus replication [6]. The more recent finding that healthy HCMV-seropositive blood donors have a high number of HCMV-specific effector T cells (0.1–3%) suggests a fre-

quent interaction between the virus and the immune system, supporting the notion that temporary reactivation of HCMV is a frequent event even in healthy probands [28].

Human CMV Reactivation in Monocyte/Granulocyte Progenitor Cells


HCMV Reactivation by Systemic Inflammation and Tumor Necrosis Factor Alpha

Clinical studies investigating the incidence of active HCMV infection and disease in immunosuppressed patients have indicated that reactivation of the virus is mainly dependent on the particular mode of immunosuppression. Transplant recipients treated with antilymphocyte antibodies such as OKT3 or ATG developed HCMV infection and disease more frequently in comparison with those treated with cyclosporin A, anti-CD4 monoclonal antibody or steroid bolus [29–36]. Systemic inflammation has been suggested to be the main reason for HCMV reactivation in many, but not all, patients. As shown in transplant recipients, plasma levels of inflammatory cytokines, especially tumor necrosis factor alpha (TNF-·), correlated with the subsequent appearance of active HCMV infection and development of HCMV disease [33, 37]. Very recent data from different laboratories working with human and murine models indicate that TNF-· plays a key role in virus reactivation in bone-marrowderived granulocyte/monocyte progenitor cells. Hahn et al. [18] were able to reactivate latent HCMV from experimentally infected human progenitor cells in the presence of TNF-·. Similarly, Henry [21] reported TNF-·-mediated ex vivo reactivation of MCMV from bone marrow myelomonocytes which were isolated from latently infected animals. MCMV reactivation has also been achieved in the lung of latently infected mice by TNF-· treatment in the absence of immunosuppression [38]; however, the cell type in which reactivation of the virus occurred unfortunately remains unclear. The hypothesis that TNF-·-dependent reactivation of latent HCMV occurs in bone-marrow-derived progenitor cells is strongly supported by our observation that HCMV antigenemia can be detected not earlier than 5–10 days after the occurrence of TNF-· peak levels in plasma, indicating that reactivation probably occurs in progenitor cells rather than in mature monocytes, since the latter have a half-life in the peripheral blood of only 1 day [4, 33]. Furthermore, temporary HCMV antigenemia can be observed in all seropositive and most seronegative patients with septic disease which is known to be associated with high TNF-· secretion, despite the absence of immunosuppression [4,

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22, 23]. In contrast to transplant recipients, HCMV antigenemia in septic patients is eliminated very efficiently after recovery from sepsis because the immune system is not severely or only temporarily suppressed. By in vitro transfection assays we have previously shown that TNF-· causes a strong stimulation of the major IE enhancer-promoter of HCMV, especially in the undifferentiated granulocyte-monocyte-progenitorlike cell line HL-60 [33, 39]. This major IE enhancer/promoter controls expression of the IE proteins IE1 and IE2 and is responsible for the initiation of HCMV replication as well as the transition from latency to productive infection [40–42]. Therefore, activity of the IE enhancer/promoter should be considered to be a critical event for reactivation of the virus. Stimulation of the IE enhancer/promoter by TNF-· was found to be mediated by its binding to the TNF-· receptor I (p55) and induction of a signal transduction cascade leading to activation of the eukaryotic transcription factor NF-ÎB. Activated NF-ÎB is translocated into the nucleus where it binds to the 18-bp repetitive sequence motifs in the IE enhancer and increases the transcriptional activity of the IE promoter [36, 43]. Reactivation of endogenous HCMV in CD14+ mature monocytes from peripheral blood of healthy seropositive blood donors was found to be dependent on the differentiation of promonocytic cells into permissive macrophages which could be induced by granulocyte-macrophage colony-stimulating factor (GM-CSF) and glycocortisone [14]. Treatment of the latently infected monocytes with GMCSF did induce IE gene transcription while early and late gene transcription could not be observed. In a different experimental context, Söderberg-Naucler et al. [44] were able to show that the formation of HCMV-permissive macrophages depends on the presence of TNF-· and interferon gamma (IFN-Á). A direct effect of TNF-· or IFN-Á on HCMV replication was not detected. From our in vitro experiments we know that the stimulatory effect of TNF-· on the IE enhancer-promoter is abrogated in differentiated monocytes [39]. Our preliminary results show that during the differentiation of monocytic cells a transcription factor is expressed which binds to the IE enhancer adjacent to an NF-ÎB binding site, which seems most important for TNF-·-mediated stimulation of the IE promoter. Binding of this factor interferes with the transcription-stimulating activity of NF-ÎB at the protein level [Prösch et al., in preparation]. This phenomenon may explain why TNF-· has no effect on HCMV reactivation in mature monocytes.

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HCMV Reactivation by Stress and cAMP-Elevating Drugs

As discussed above, reactivation of latent HCMV is not restricted to patients with systemic inflammation, suggesting additional mechanism(s) of HCMV reactivation independent of TNF-·-mediated stimulation. Toro and Ossa [27] observed that temporary reactivation of HCMV in healthy blood donors, as measured by a DNA PCR procedure not sensitive enough to detect latent virus, was associated with the occurrence of stressful events. To verify the relationship between HCMV reactivation and stress we initiated a clinical study to investigate the incidence of active HCMV infection in patients with acute myocardial infarction – a highly stressful situation. As hypothesized, in all HCMV-seropositive patients the virus was reactivated, as measured by HCMV antigenemia. Elevated TNF-· levels could not be detected in the plasma of these patients. However, HCMV antigenemia was preceded by a rise in plasma catecholamine levels. Moreover, the plasma level of catecholamines in individual patients correlated with the frequency of HCMVantigen-positive PBMCs [Prösch et al., in preparation]. This observation raised the question whether catecholamines directly induce reactivation of latent HCMV or stimulate expression or even replication of the virus which was reactivated by other as yet unknwon mediator(s). By in vitro transfection experiments it became obvious that catecholamines, similarly to TNF-·, stimulate the activity of the HCMV IE enhancer-promoter in monocytic cell lines. In contrast to TNF-·, this stimulatory effect is not restricted to undifferentiated HL-60 cells (which are a model for bone-marrow-derived granulocytemonocyte progenitor cells) but also occurs in THP-1 cells which resemble mature monocytes. This may correlate with the observation that in patients with myocardial infarction antigenemia is detectable earlier than in patients with systemic inflammation. The molecular mechanism by which catecholamines stimulate the IE enhancerpromoter of HCMV involves increased expression and/or activation of the transcription factor CREB-1/ATF-1 via the cAMP pathway. CREB-1/ATF-1 binds to 19-bp sequence motifs of the IE enhancer and increases the transcriptional activity of the IE promoter. Thus, the action of catecholamines on the HCMV IE enhancer-promoter follows a mechanism which is independent from that of TNF-· [Prösch et al., in preparation]. In vitro stimulation of the HCMV IE enhancer-promoter in monocytic cells via the cAMP pathway was also demonstrated for pentoxifylline (PTX), which promoted

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active HCMV infection and disease in vivo in transplant patients, despite its inhibitory activity on TNF-· secretion. Furthermore, PTX and TNF-· together cause a synergistic stimulation of the IE enhancer-promoter in monocytic cells as well as of virus replication in primary endothelial cells and human embryonal lung fibroblasts [36, 45]. Catecholamine-mediated (re)activation of latent HCMV may be the reason for active HCMV infections (as defined by HCMV antigenemia) not only in patients with acute myocardial infarction but also for the proposed temporary reactivation of HCMV in healthy virus carriers (see above). If so, stress-mediated HCMV reactivation follows a mechanism very similar to that known for ·-herpesviruses (HSV1 and 2, varicella-zoster virus) [for reviews, see ref. 46, 47]. Interestingly, intraoperative hypothermia during liver transplantation was found to increase the risk of HCMV infection [48], and as is known, hypothermia is an event which induces stress hormones.

HCMV replication s

Conclusion and Outlook

In summary, reactivation of latent HCMV from bonemarrow-derived progenitor cells as a main site of HCMV latency in humans seems to be a rather frequent event independent of immunosuppression itself. The constitution of the cellular immune system determines the outcome of reactivation. In the presence of a normal or mainly intact immune response, the active HCMV infection should be eliminated efficiently within 1–2 weeks, as observed in healthy blood donors, in patients with a septic disease or in patients after myocardial infarction [4, our unpubl. results]. In individuals with a diminished immune response, as evident in elderly people or transplant recipients under low immunosuppressive therapy, persistent antigenemia can be observed, indicating that the virus infection is only partially controlled. In patients with strong immunosuppression, the reactivated virus replicates in an uncontrolled manner, frequently leading to severe HCMV disease. One definitive mechanism of reactivation is based on the stimulatory effect of the proinflammatory cytokine TNF-· on IE gene expression of the virus in latently infected bone-marrow-derived progenitor cells. A second, independent mechanism may involve a direct stimulatory effect of stress-induced catecholamines on viral gene expression (summarized in fig. 1), a hypothesis which has to be proved now in an animal model. Increasing understanding of the molecular mechanisms involved in

Fig. 1. Potential mechanisms of HCMV reactivation in granulocytemonocyte progenitor cells and mature monocytes.



HCMV reactivation should lead to the development of strategies for therapeutic intervention, with the aim of preventing reactivation. This may be the only way to prevent HCMV-associated disease in immunosuppressed patients. Prophylactic or preemptive treatment of transplant recipients with the currently available antiviral drugs can decrease the incidence of severe HCMV diseases and delay the onset of disease, but has no influence on reactivation of the virus and its proinflammatory effects [49– 53]. An understanding of the molecular mechanisms of HCMV reactivation is an essential prerequisite for developing ways to prevent reactivation of latent HCMV.

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29 Singh N, Dummer JS, Kusne S, Breinig MK, Armstrong JA, Makowka L, Starzl TE, Ho M: Infections with cytomegalovirus and other herpesviruses in 121 liver transplant recipients: Transmission by donated organ and the effect of OKT3 antibodies. J Infect Dis 1988;158: 124–131. 30 Hooks MA, Perlino CA, Henderson JM, Millikan WJ Jr, Kutner MH: Prevalence of invasive cytomegalovirus disease with administration of muromanab CD-3 in patients undergoing orthotopic liver transplantation. Ann Pharmacother 1992;26:617–620. 31 Hibberd PL, Tolkoff-Rubin NE, Cosimi AB, Schooley RT, Isaacson D, Doran M, Delvecchio A, Delmonico FL, Auchincloss H Jr, Rubin RH: Symptomatic cytomegalovirus disease in the cytomegalovirus antibody seropositive renal transplant recipients treated with OKT3. Transplantation 1992;53:68–72. 32 Paya CV, Wiesner RH, Hermanns PE, LarsonKeller JJ, Ilstrup DM, Krom RA, Rettke S, Smith TF: Risk factors for cytomegalovirus and severe bacterial infections following liver transplantation: A prospective multivariate time-dependent analysis. J Hepatol 1993;18: 185–195. 33 Fietze E, Prösch S, Reinke P, Stein J, Döcke WD, Staffa G, Loening S, Devaux S, Emmrich F, von Baehr R, Krüger DH, Volk HD: Cytomegalovirus infection in transplant recipients: The role of tumor necrosis factor. Transplantation 1994;58:675–680. 34 Krogsgaard K, Boesgaard S, Aldershvile J, Arendrup H, Mortensen SA, Petterson G: Cytomegalovirus infection rate among heart transplant patients in relation to the potency of antithymocyte immunoglobulin induction therapy. Copenhagen Heart Transplant Group. Scand J Infect Dis 1994;26:239–247. 35 Portela D, Patel R, Larson-Keller JJ, Ilstrup DM, Wiesner RH, Steers JL, Krom RA, Paya CV: OKT3 treatment for allograft rejection is a risk factor for cytomegalovirus disease in liver transplantation. J Infect Dis 1995;171:1014– 1018. 36 Prösch S, Volk HD, Reinke P, Pioch K, Döcke WD, Krüger DH: Human cytomegalovirus infection in transplant recipients: Role of TNF· for reactivation and replication of human cytomegalovirus; in Scholz M, Rabenau HF, Doerr HW, Cinatl J Jr (eds): CMV-Related Immunopathology. Monogr Virol. Basel, Karger, 1998, vol 21, pp 29–42. 37 Humar A, St Loius P, Mazzulli T, McGeer A, Lipton J, Messner H, MacDonald KS: Elevated serum cytokines are associated with cytomegalovirus infection and disease in bone marrow transplant recipients. J Infect Dis 1999;179: 484–488. 38 Koffron A, Varghese T, Hummel M, Yan S, Kaufman D, Fryer J, Leventhal J, Stuart F, Abecassis M: Immunosuppression is not required for reactivation of latent murine cytomegalovirus. Transplant Proc 1999;31:1395– 1396.

Prösch/Döcke/Reinke/Volk/Krüger

39 Stein J, Volk HD, Liebenthal C, Krüger DH, Prösch S: Tumor necrosis factor alpha stimulates the activity of the human cytomegalovirus major immediate early enhancer/promoter in immature monocytic cells. J Gen Virol 1993; 74:2333–2338. 40 Iskenderian AC, Huang L, Reilly A, Stenberg RM, Anders DG: Four of eleven loci required for transition complementation of human cytomegalovirus DNA replication cooperate to activate expression of replication genes. J Virol 1996;70:383–392. 41 Mocarski ES, Kemble GW, Lyle JM, Greaves RF: A deletion mutant in the human cytomegalovirus gene encoding IE1(491aa) is replication defective due to a failure in autoregulation. Proc Natl Acad Sci USA 1996;93:11321– 11326. 42 Greaves RF, Mocarski ES: Defective growth correlates with reduced accumulation of a viral DNA replication protein after low-multiplicity infection by a human cytomegalovirus ie1 mutant. J Virol 1998;72:366–379. 43 Prösch S, Staak K, Stein J, Liebenthal C, Stamminger T, Volk HD, Krüger DH: Stimulation of the human cytomegalovirus IE enhancer/ promoter in HL-60 cells by TNF· is mediated via induction of NF-ÎB. Virology 1995;208: 197–206.


44 Söderberg-Naucler C, Fish KN, Nelson JA: Interferon-Á and tumor necrosis factor-· specifically induce formation of cytomegalovirus-permissive monocyte-derived macrophages that are refractory to the antiviral activity of these cytokines. J Clin Invest 1997;100:3154–3163. 45 Staak K, Prösch S, Stein J, Priemer C, Ewert R, Döcke WD, Krüger DH, Volk HD, Reinke P: Pentoxifylline promotes replication of cytomegalovirus in vivo and in vitro. Blood 1997; 89:3682–3690. 46 Steiner I: Human herpes viruses latent infection in the nervous system. Immunol Rev 1996;152:157–173. 47 Turner SL, Jenkins FJ: The role of herpes simplex virus in neuroscience. J Neurovirol 1997; 3:110–125. 48 Paterson DL, Steplefeldt WH, Wagener MM, Gayowski T, Marino IR, Singh N: Intraoperative hypothermia is an independent risk factor for early cytomegalovirus infection in liver transplant recipients. Transplantation 1999; 27:1151–1155. 49 Przepiorka D, Ippoliti C, Panina A, Goodrich J, Giralt S, van Besien K, Mehra R, Deisseroth AB, Andersson B, Luna M et al: Ganciclovir three times per week is not adequate to prevent cytomegalovirus reactivation after T-cell-depleted marrow transplantation. Bone Marrow Transplant 1994;13:461–464.

50 Winston DJ, Imagawa DK, Holt CD, Kaldas F, Shaked A, Busuttil RW: Long-term ganciclovir prophylaxis eliminates serious cytomegalovirus disease in liver transplant recipients receiving OKT3 therapy for rejection. Transplantation 1995;60:1357–1360. 51 Gomez E, de Ona M, Aguado S, Tejada F, Nunez M, Portal C, Diaz-Corte C, Sanchez E, Ortega F, Alvarez-Grande J: Cytomegalovirus preemptive therapy with ganciclovir in renal transplant patients treated with OKT3. Nephron 1996;74:367–372. 52 Canpolat C, Culbert S, Gardner M, Whimbey E, Tarrand J, Chan KW: Ganciclovir prophylaxis for cytomegalovirus infection in pediatric allogenic bone marrow transplant recipients. Bone Marow Transplant 1996;17:589–593. 53 Grundy JE: Current antiviral therapy fails to prevent the pro-inflammatory effects of cytomegalovirus infection, whilst rendering infected cells relatively resistant to immune attack; in Scholz M, Rabenau HF, Doerr HW, Cinatl J Jr (eds): CMV-Related Immunopathology. Monogr Virol. Basel, Karger, 1998, vol 21, pp 67–89.


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Human Cytomegalovirus Latency and Reactivation – A Delicate Balance between the Virus and Its Host’s Immune System Cecilia Söderberg-Nauclér a Jay A. Nelson b a Department b Department

of Biosciences at Novum Karolinska Institute, Novum Stockholm, Sweden, and of Molecular Microbiology and Immunology, Oregon Health Sciences University, Portland, Oreg., USA

Key Words Cytomegalovirus W Macrophages W Latency W Reactivation W Allogeneic stimulation

Abstract Human cytomegalovirus (HCMV) is a ubiquitous herpesvirus that still causes severe morbidity and mortality in immunocompromised individuals. During its evolution, the virus has developed sophisticated methods to evade immune recognition and to establish life-long persistence in its host. Today, we know that the virus establishes latency in myeloid lineage cells and that the virus is dependent on immune activation mechanisms to reactivate it from latency to produce a new viral progeny. During this process, a number of viral proteins are produced that interfere with different immune recognition pathways. The current knowledge of the delicate balance between the virus‘ continuous existence and its host’s immune system will be summarized in this chapter. Copyright © 2000 S. Karger AG, Basel

ABC

© 2000 S. Karger AG, Basel 0300–5526/99/0426–0314$17.50/0



Introduction

For many years, human cytomegalovirus (HCMV) was not considered to be a major human pathogen, since the virus only caused rare cases of HCMV inclusion disease in neonates. However, the importance and the interest of HCMV as a pathogen has increased over the past two decades, with the escalation in the number of patients undergoing immunosuppressive therapy following organ or bone marrow transplantation, as well as with the increasing amount of AIDS patients. Today, we know that approximately 70–100% of the population in the world are carriers of the virus. A primary HCMV infection is followed by a life-long persistence of the virus in a latent state, and reactivation may occur later in life. HCMV infection is usually subclinical in immunocompetent individuals, but the virus can cause fatal disease in immunocompromised patients. HCMV can infect virtually all organ tissues, but manifestations of organ involvement generally include symptoms from the liver, the lungs, the intestine and the CNS. In order to coexist with its host, HCMV has developed a number of mechanisms to avoid immune recognition and to utilize specific immune functions for reactivation and spread of virus. The current knowledge of these aspects will be reviewed in this chapter.

Cecilia Söderberg-Nauclér Department of Biosciences at Novum Karolinska Institute, Novum S–141 57 Stockholm (Sweden) Tel. +46 8 608 9146, Fax +46 8 704 8893

Clinical Manifestations of HCMV Infection

Depending on the country studied, between 60 and 100% of the population experience HCMV infection during their life-time [1, 2]. Generally, primary HCMV infection takes place during childhood, and the virus is transmitted through close personal contact. All bodily excretions are possible sources of HCMV transmission and most children become infected via breast milk [3] or via transmission from child to child in day-care centers and schools [2, 4]. HCMV can also be transferred by blood products, bone marrow grafts and solid-organ transplants, and several studies have identified leukocytes as the most important source of virus in blood products [5–7]. Although immunity against the virus does not prevent reinfections, maternal immunity and pretransplantation immunity against HCMV as well as the immunosuppressive regimen used have been shown to influence the incidence of HCMV disease [8]. While 50–90% of bone marrow and organ transplant patients experience HCMV infections in the posttransplantation period, the prevalence of HCMV approaches 100% in most HIV-infected groups of patients [1]. Furthermore, with an incidence of 0.2–2.2% per live birth, HCMV infection is the most common congenital virus infection [9–11]. Interestingly, HCMV causes different clinical symptoms in these different groups of patients [1, 12, 13]. While clinical abnormalities in HCMV-infected transplant recipients often include spiking fever, leukopenia, malaise, leukopenia, hepatitis, interstitial pneumonia and impaired graft function, HCMV retinitis and gastrointestinal disease are the most common disease manifestations among AIDS patients [as reviewed ref. 1]. Neonates who are born with congenital HCMV infections have a high prevalence of birth defects such as hearing loss and mental retardation [1, 14]. Furthermore, HCMV has been implicated in the development of atherosclerosis, chronic rejection in solid-organ recipients and bone marrow failure as well as chronic graft-versus-host disease in bone marrow transplant patients [15–19].

HCMV Infection in Epithelial Cells during Acute HCMV Infection

During the acute phase of HCMV disease, organ involvement may include symptoms from virtually all organ systems. Histological analyses of tissues obtained from patients with severe HCMV disease have demonstrated HCMV-infected cells in all of these tissues [1, 20].

Human Cytomegalovirus Latency and Reactivation

The most frequently infected cell types found during HCMV disease include endothelial cells (EC), epithelial cells, macrophages and fibroblasts. In addition, neuronal cells, smooth muscle cells and hepatocytes are occasionally virus positive in infected patients. The susceptibility of these cells to HCMV infection has also been confirmed in in vitro studies of the virus [1, 20]. During primary acute disease, HCMV predominantly infects epithelial cells, and infected cells have been found in a number of different organs including the lungs, liver, intestine, kidney, breast and salivary glands. Since epithelial cells constitute an interface between the host and its environment, these cells have been implicated as the initial site of infection from which the virus spreads to other tissues. The ability of the virus to infect epithelial cells facilitates the release of virus in bodily excretions especiall from epithelial cells in the bowel, bladder, breast and salivary gland, but the infection of epithelial cells may also facilitate viral transmission to underlying tissues. In support of the latter hypothesis, studies of tissue samples from organs obtained from patients with HCMV disease have demonstrated infected fibroblasts as the predominant cell type in tissues such as the lungs, the intestine and the placenta [1, 21–23]. HCMV infection of both epithelial cells and fibroblasts results in destruction of the infected cell, which may explain the development of ulcerations in tissues such as the intestine. Since epithelial cells and fibroblasts appear to be end-stage lytic target cells, this cell type has not been considered as a candidate for a reservoir of latent virus.

HCMV Infection in EC

Another cellular target for HCMV is the vascular EC. Early studies of tissues obtained from bone marrow transplant patients with acute HCMV disease demonstrated that endothelial cells commonly harbor HCMV without signs of cytopathology [20]. Further studies demonstrated that EC are frequently infected in tissues obtained from HCMV-infected patients [22, 24]. In particular, EC from the microvasculature in the brain, the lungs, the heart, the gastrointestinal tract and the placenta are often virus positive [22, 23]. HCMV has also been detected in the vessel walls of HCMV-seropositive individuals without active infection [25, 26]. However, the ability to establish HCMV infection in EC in vitro has been controversial. Early reports suggested that HCMV was unable to productively infect EC [25] while others have observed infection of a low percentage of EC in culture [26, 27]. The


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majority of these studies have utilized human umbilical vein EC (HUVEC) to examine replication of HCMV in EC. Although these cells are relatively easy to obtain, HUVEC are of fetal origin and may not represent HCMV infection in adult tissue. Since EC exhibit different physiological and biochemical differences which are dependent on the cellular origin (adult versus fetal), anatomical location, and vessel size (large vessel versus capillary) [28], EC in different organs may exhibit differences in the ability of HCMV to replicate in these cells. In support of this hypothesis, our group has demonstrated that HCMV infection of EC obtained from the microvasculature (MVEC) or the macrovasculature (aortic EC (AEC)) demonstrated strain variabilities as well as differences in the effects of the virus on the respective cells [29]. While HCMV infection in MVEC was lytic and resulted in destruction of infected cells within 1 week after infection, HCMV infection of AEC resulted in a persistent infection [29]. Virus infection in AEC did not result in cell lysis, and production of virus was demonstrated in these cells up to 40 days post infection. Thus, while HCMV-infected endothelial cells in large vessels such as the aorta may be persistently infected and release infectious virus, virus infection of the microvasculature may result in destruction of infected cells. These observations sugget that the endothelium on vessel walls may be persistent sites of virus and have generated an interest in the association of HCMV with the development of atherosclerosis [30]. This hypothesis has been further reinforced by animal models which have directly implicated CMV in the development of vascular sclerosis in transplanted organs (chronic rejection) [31, 32] and in restenosis in the arteries of patients following angioplasty [15]. In addition, these observations suggest that persistently infected EC release infectious virus into the blood and that these cells can transfer infection to circulating blood cells.

HCMV Infection in Macrophages

HCMV most likely spreads to different tissues either by trafficking of free virus particles in the blood, or by utilizing cells as a vehicle for spread of virus. Transmission of latent HCMV was early shown to occur through transfusion of blood products, bone marrow grafts and solid organs [1, 7, 33, 34], but the latent cellular reservoir has been difficult to identify. Studies of separated peripheral blood cell populations derived from patients with HCMV disease or asymptomatically infected individuals have identified monocytes as the predominant infected

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cell type [35]. However, HCMV infection of monocytes is infrequent and viral replication may be abortive or restricted to early events of gene expression [36–38]. In contrast, tissue samples from patients with HCMV disease demonstrate a high frequency of HCMV-infected macrophages which express late viral genes [39, 40]. In vitro studies of HCMV replication in primary monocyte/macrophage systems have shown that the ability of the virus to replicate in these cells is dependent on the state of cellular differentiation [36, 37]. Furthermore, HCMV also seem to have developed functions to remain persistent in macrophages, which include strategies to delay viral replication, avoid lysosomal degradation and avoid destruction of the infected cell. In essence, HCMV utilizes macrophages as a Trojan horse similar to lentiviruses [41, 42]. Since macrophage differentiation is extremely complex and diverse, it is important to dissect the components required for the formation of HCMV permissive macrophages to understand HCMV pathogenesis. For this purpose, a number of primary monocyte/macrophage systems have been established to study HCMV replication in vitro. Two of these systems are based on activation of T cells and production of cytokines for the differentiation of monocytes into macrophages, which are fully permissive for HCMV [36, 43]. These macrophages can be maintained for long periods of time without the addition of exogenous cytokines. Using concanavalin A stimulation of peripheral blood mononuclear cells to differentiate monocytes into macrophages, HCMV replication has been shown to be dependent on the presence of CD8+ T lymphocytes and the production of IFN-Á and TNF-· [43]. This finding was quite surprising since both TNF-· and IFN-Á are known to have antiviral effects. However, neither INF-Á nor TNF-· inhibited HCMV replication in infected macrophages [43]. The identification of IFN-Á and TNF-· as factors necessary for the production of HCMV permissive macrophages has important clinical implications, since an immune-mediated process which involves the activation of T cells and the production of these cytokines may be necessary for virus production in macrophages in vivo. In support of this hypothesis, elevated levels of both IFN-Á and TNF-· have been found in sera of patients with HCMV disease [44–48], and TNF-· stimulates the immediate early (IE) promoter in myeloid cells [49, 50]. In addition, activation of HCMV in vivo frequently occurs following allogeneic organ and bone marrow transplantation as well as during bacterial infections. Thus, T cell activation and subsequent production of cytokines may be necessary for virus production in HCMV-infected patients. Furthermore, allogeneic stimu-

Söderberg-Nauclér/Nelson

lation of peripheral blood mononuclear cells from HCMV-positive healthy donors results in reactivation of latent virus in macrophages in vitro [51]. These experiments conclusively identified myeloid lineage cells as a cellular reservoir for HCMV latency and described the first in vitro system to study different aspects of reactivation of latent virus from naturally infected cells. The recovery of HCMV from allogeneically stimulated macrophages suggests that virus production is intimately linked to activation of the immune system. Further studies confirmed this hypothesis since reactivation of latent HCMV was shown to be dependent on allogeneic activation of T cells and the production of IFN-Á [52]. Thus, immune activation of T cells and the production of IFN-Á resulting in macrophage differentiation is a key element in the reactivation of latent HCMV.

HCMV Establishes Latency in Myeloid Lineage Cells

Macrophages are a heterogeneous population of terminally differentiated myeloid lineage cells, whose precursors are monocytes, monoblasts and promonocytes. Since monocytes in the peripheral blood are cells with a short half-life, the ability to reactivate latent HCMV in macrophages implies that HCMV is maintained in a precursor population of the myeloid cell lineage. Myeloid cells would provide an ideal site of latency for a virus that is closely linked to the immune system for activation. HCMV has previously been reported to infect CD34+ pluripotent stem cells both in vitro [53–56] and in vivo [57, 58]. However, these findings are controversial and do not explain why virus is not consistently found in all mature peripheral cell lineages [35, 59]. One culture system which demonstrates HCMV infection in CD33+ myeloid progenitor cells in vitro suggests that myeloid cells may be infected early during hematopoiesis [60, 61]. In this model, in vitro infection of myeloid progenitor cells results in production of infectious HCMV after differentiation into CD14+ macrophages. Transcripts, which have been implicated for HCMV latency, were associated with the lack of infectious virus in myeloid progenitor cells. Although myeloid progenitor cells are an obvious site for HCMV latency, these cells have not been confirmed as a latent source of virus in vivo. A conclusive identification of an in vivo latent site requires reactivation of virus from cells obtained from healthy individuals.


The Importance of Immune Activation in the Reactivation of HCMV

In healthy HCMV carriers, the virus is believed to remain latent and the function of the immune system is critical in the control of virus latency. This hypothesis originates from the observation that immunosuppressed individuals often suffer from HCMV disease. In these patients, reactivation of virus rather than reinfection or primary infection is considered to be the major cause of disease. Thus, experimental evidence for immune activation of HCMV from the peripheral blood of healthy donors has important clinical implications. Transplant patients commonly reactivate HCMV betwen 1 and 4 months after transplantation [62]. HCMV infection has been associated with acute rejection in organ transplant patients and acute graft-versus-host disease in bone marrow transplant patients [63]. An allogeneic activation of cells in solid organs or bone marrow grafts would provide a microenvironment conducive to cellular differentiation and reactivation of HCMV, since immune activation of myeloid lineage cells by activated T cells facilitates reactivation of HCMV. Thus, the allogeneic activation of residual infected leukocytes in the solid organ or bone marrow recipient tissue may be the primary source of virus in transplant patients. Furthermore, since blood transfusions commonly are performed over histoincompatibility barriers, reactivation of HCMV through allogeneic stimulation of transferred blood cells would be expected if either the donor or the recipient were virus positive. Furthermore, when virus reactivates in transplant patients, the iatrogenic immunosupression would exacerbate the severity of the disease. Since the immunosuppressive state in the patient also impairs the ability of the host to respond against other pathogens, immunosuppressed individuals often suffer from infections by other pathogens. Although inefficient, the immune response against the microbe also results in immune activation of T cells, which may cause reactivation of latent virus in macrophages. This situation is exemplified by the increased incidence of HCMV disease following bacterial infections in different groups of patients [64] as well as in HIV-infected individuals who experience other opportunistic infections. The hypothesis of viral activation has also been implied for other viruses. For example, allogeneic stimulation of HIV-infected macrophages induces high levels of viral expression [65] and conjugates of dendritic cells and T cells have been suggested to be important sites for replication of HIV [66]. Thus, immune activation as a part of the host defense against a pathogen or non-self cells may result in reactivation of virus and HCMV disease.


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The Delicate Balance between HCMV and the Host’s Immune System

Since HCMV infection generally is silent in immunocompetent individuals, the balance between the virus and its host’s immune system is most likely accomplished. However, in immunocompromised patients, an imbalance between the virus and the immune system may lead to uncontrolled virus replication. T cells which are activated by recognition of foreign antigens in the context of MHC molecules or during inflammation, produce a number of different cytokines. Production of inflammatory cytokines in response to HCMV infection would activate effector cells that can eliminate virus-infected cells and exhibit antiviral effects. In support of this theory, HCMV specific CD4+ T cells have been shown to respond to HCMV antigens with the production of IL-2, TNF-· and IFN-Á [20]. In addition, HCMV infection induces the production of cytokines such as IL-6 and TGF-ß in fibroblasts or endothelial cells and TNF-·, IL-1 and IFN-· in infected monocytes [as reviewed in ref. 67]. In addition, IFN-Á and TNF-· are produced by lymphocytes which interact with HCMV-infected cells [68]. Both IFN-Á and TNF-· are thought to be antiviral cytokines since they interfere with viral replication. However, these cytokines have also been shown to be important for the development of HCMV-permissive macrophages [52], and IFN-Á was shown to be crucial for reactivation of HCMV in myeloid lineage cells [43]. Thus, while immune reactions contribute to the basic protection against HCMV, the virus has taken an advantage of activated immunological cells and the production of antiviral cytokines to obtain a state of macrophage differentiation which supports virus production and reactivation of latent virus. Previous animal studies have also demonstrated that these cytokines positively influence viral replication. For example, TNF· has been shown to promote CMV replication and pathogenicity in the rat model [69]. In contrast, IFN-Á has been shown to restore CMV antigen presentation, which implies a role for IFN-Á in the T-cell-mediated control of CMV infection [70–72]. Prophylactic IFN-Á treatment has been shown to reduce mortality in CMV-infected mice [73], and IFN-Á-depleted mice demonstrate increased murine CMV titers in multiple organs [74]. Furthermore, IFN-Á has been shown to be important for the clearance of murine CMV (MCMV) in infected animals [75, 76], and experiments performed in mice lacking the IFN-Á receptor demonstrate that IFN-Á reversibly inhibited reactivation of latent MCMV [77]. Thus, in vivo experiments suggest that the inflammatory cytokines are im-

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portant also in the basic protection against CMV. These observations imply that although CMV is dependent on immune activation to reactivate from latency, virus replication may be inhibited by the action of inflammatory cytokines. This hypothesis may explain why immunosuppressed individuals frequently suffer from severe HCMV infections. Another example of an immunological dysfunctin that indirectly may exacerbate HCMV disease in immunocompromised patients is bone marrow failure. This complication is common both among HCMV-infected bone marrow transplant recipients and AIDS patients, and HCMV has been implicated in this disease process by suppression of hematopoiesis, or possibly by direct viral effects on effector cells. HCMV-induced immunosuppression is exemplified by leukopenia, depression of lymphocyte responses to T cell mitogens and decreased NK as well as cytotoxic T lymphocyte activities [as reviewed in ref. 20]. T cells may be functionally impaired without being infected with HCMV, since HCMV infection has been shown to suppress various monocyte functions, including IL-1 production and T cell proliferative responses to mitogens such as concanavalin A or pokeweed mitogen as well as antigen presentation [20]. Another study suggests that HCMV induces TNF-· production, which mediates the release of arachidonic acid and the metabolite PGE2 in infected monocyte cultures. These compounds inhibit mitogenic T cell proliferative response in vitro and implicates these substances in HCMVinduced immunosuppression [20]. Thus, HCMV may have developed functions which inhibit T cell activation not only by interfering with peptide presentation in the context of MHC molecules (see below), but also by acting directly on the T cell level. Furthermore, although infection of monocytes and possibly T cells may contribute directly to immunological defects in these patients, HCMV infection could also, by infecting stromal cells in the bone marrow, indirectly affect normal myelopoiesis which may result in leukopenia. In support of this hypothesis, HCMV infection of stromal cells in vitro can reduce the capacity of these cells to support proliferation of primitive myeloid progenitor cells [16]. Thus, bone marrow failure may also be a result of HCMV infection in bone marrow stromal cells, which will fail to support normal hematopoiesis.


Chemokines

Together with cell adhesion molecules, chemokines direct target subsets of effector cells to specific tissue sites of infection and inflammation and are therefore believed to play important roles in wound healing and clearance of infectious pathogens. Chemokines are low-molecular cytokines, which are known to regulate hematopoiesis, induce extravasation of leucocytes and chemotaxis. There are two main subfamilies of chemokines; the ·-chemokines (CXC) and the ß-chemokines (CC), which are classified due to the position of the cysteins (C) in the Nterminus of the protein. Chemokines mediate their biological activity by binding to G-coupled receptors (GCR) which are numbered and designated according to their specificity; CXCR and CCR [78]. Since chemokines will recruit cells of the immune system to sites of inflammation, these molecules have also been implicated in mediating tissue injuries in both acute and chronic inflammatory processes, and are believed to play a pathogenetic role in HCMV disease by attracting T cells and monocytes to an inflammatory site. For example, HCMV-infected cells increase the production of chemokines such as RANTES and IL-8 [67]. RANTES expression has also been shown to be significantly increased in lung tissue from patients with HCMV pneumonia as compared to lungs affected by acute rejection [67]. In addition, macrophages obtained by bronchial alveolar lavage from patients with HCMV pneumonitis produced more RANTES than cells obtained from control individuals. Interestingly, HCMV encode for several putative proteins that may function as GCRs. In the HCMV genome, the open reading frames US27, US28, UL33 and UL78 have been identified by sequence homologies to known GCRs [67], but the functions of these receptors are unknown. Possibly, the virus utilizes these proteins to bind and neutralize chemokines which will inhibit recruitment of cells to tissue sites where HCMV-infected cells are located. US28 has been shown to bind a wide range of CC chemokines such as RANTES, MCP-1 and MIP-1·, but not CXC chemokines [67]. The GCR homologous do not appear to be essential for virus replication, since virus mutants which are lacking US27 and US28 from the HCMV genome do not appear to affect virus growth in vitro [67]. However, these receptors may have important functions in vivo. For example, studies of murine CMV have demonstrated that the UL33 homologue in mice may play an important role in targeting virus to specific tissues [67]. Furthermore, although the UL33 homologue in rats, R33, which is expressed at late times after infection does not exhibit an altered


growth in vitro, R33 knockout virus demonstrates lower mortality in infected animals [79]. In addition, similar to the murine UL33 homologue, the virus cannot replicate in the salivary glands of infected rats, which suggests an important function for this protein in the pathogenesis of CMV.

Concluding Remarks

HCMV establishes latency in myeloid lineage cells, and virus proteins have not been conclusively identified in latently infected cells. However, to secure its continuous existence, the virus must be reactivated and produce a new viral progeny and spread to other individuals. Therefore, HCMV is most likely reactivated from latency intermittently when myeloid lineage cells differentiate into macrophages and the virus then spreads to different tissues and to other hosts. During this process the virus becomes visible to the immune system which will attempt to eliminate it. When the virus becomes reactivated, the immune system is critical for the host to achieve control over the virus. Therefore, permanent existence of HCMV in an immunocompetent host has put an immense evolutionary pressure on the virus to develop strategies to avoid immune recognition. To evade recognition and killing, HCMV has developed unique strategies to block antigen presentation, to prevent eradication by different cells of the immune system, and to achieve life-long latency and persistence. Examples of these strategies include the inhibition of MHC class I expression by US2, US3, US6 and US11, the possible role of UL18 in the inhibition of NK cell killing of HCMV-infected cells as well as the inhibition of MHC class II expression by US2 and the observed IFN-Á irresponsiveness through the Jak/Stat pathway [as reviewed elsewhere in this issue, 80, 81]. Thus, a balance exists between HCMV and the immune system, otherwise either the virus or the host would be eliminated during the infection process. Since the virus persists in the host by a delicate balance with his/her immune system, a decreased immune function of the host often leads to an advantage for the virus, and uncontrolled virus replication often results in severe disease in immunocompromised patients. Increased knowledge of the basic nature of this virus will therefore provide us important information about basic virology and immunology, and will also help us develop new strategies for prophylaxis and treatment of HCMV infection.


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Measurement of Anti-Human Cytomegalovirus T Cell Reactivity in Transplant Recipients and Its Potential Clinical Use: A Mini-Review Florian Kern a Nicole Faulhaber a, b Elham Khatamzas a Claudia Frömmel a Ralf Ewert c Susanna Prösch d Hans-Dieter Volk a Petra Reinke b a Institut

für Medizinische Immunologie, Charité (Medizinische Fakultät der Humboldt-Universität zu Berlin), Klinik mit Schwerpunkt Nephrologie und Intensivmedizin, Charité, c Deutsches Herzzentrum Berlin (DHZB), and d Institut für Virologie, Charité, Berlin, Deutschland b Medizinische

Key Words Cytomegalovirus infection W Cytomegalovirus disease W Cellular immune defense W T cells W Intracellular cytokines W Immunosuppression

Abstract By allowing direct determination of the frequencies of antigen-specific memory T cells in peripheral blood, novel techniques based on flow cytometry provide new diagnostic opportunities in various clinical settings, including organ transplantation. While the importance of the T cell compartment for the anti-human cytomegalovirus (HCMV) immune response is undisputed, efficient monitoring of this response was previously impossible because the conventional methods for measuring CD4+ and CD8+ T cell responses are too time-consuming and cost-intensive. We analyzed how the rapid induction of anti-HCMV CD4+ and CD8+ memory T cells by HCMV viral lysate or HCMV-derived peptides, respectively, followed by a flow-cytometric detection step, may be used to monitor HCMV-specific CD4+ and CD8+ memory T cells in solid-organ recipients. We also discuss a number of preconditions for integrating such testing into the clinical routine. Copyright © 2000 S. Karger AG, Basel

ABC

© 2000 S. Karger AG, Basel 0300–5526/99/0426–0322$17.50/0



Measurement of the frequencies of antigen-specific memory T cells in peripheral blood by multiparameter flow cytometry has recently become an area of major interest in immunological research [1–4]. New methods allow the direct visualization of antigen-specific T cells in peripheral blood, their quantification (frequencies) and detailed phenotypic analysis. Such examinations may be useful in various clinical settings in which there is an increased risk of HCMV disease, such as, for example, solid-organ or bone marrow transplantation or HIV infection. Two major approaches for visualizing antigen-specific T cells exist to date. One of them is based on the rapid induction of cytokine synthesis in memory cells stimulated by complete proteins or protein lysates (CD4+ T cells) or small peptides (CD8+ T cells) and uses the detection of intracellularly retained cytokines (following treatment with the secretion inhibitor brefeldin A) as a readout parameter [1, 2]. The obvious advantage of this approach is that the T cells that are visualized are also functional, while a disadvantage is that T cells that are nonfunctional yet specific for the antigen used may be missed. The other, equally elegant technique is based on the direct staining of T cells which bind fluorescent labeled MHC class I/peptide complex tetramers [3, 4]. In order to be stained, the T cells must recognize the peptide included in the complex. This approach has the advantage

Florian Kern Institut für Medizinische Immunologie der Charité Campus Mitte, COZ D–10098 Berlin (Germany) Tel. +49 30 280 22858, Fax +49 30 280 25461, E-Mail [email protected]

of staining all T cells specific for one particular peptide; however, it does not examine T cell function. Because T cells are known to play an important role in the control of infection in human cytomegalovirus (HCMV) [5–8], the characterization and quantification (T cell function and frequencies of responding cells) of the HCMV-directed T cell response is a key issue in the search for efficient therapeutic and preventive strategies. While HCMV disease is a relatively rare complication after solid-organ transplantation, it is frequently following bone marrow transplantation [6, 9], and represents a major cause of morbidity and mortality in AIDS [10, 11]. However, there are no clinical signs to predict this dangerous complication. Recently, determinations of viral load have been used to predict the risk of developing HCMV disease [12]. The clinical usefulness of such determinations, however, has remained unclear. Because viral replication and, therefore, also viral load are likely to depend on the efficiency of cellular immune mechanisms [13], it may be useful to also analyze the HCMV-specific T cell response in these individuals. In fact, recent data indicate that there is an inverse correlation between the HCMVdirected cytotoxic T lymphocyte (CTL) response and HCMV antigenemia (immune cytology) [13]. These interesting results underline the potential benefit of testing the cellular immune defense; however, at the same time they call for a more rapid approach to this issue. The method for analyzing the CD8+ T cell compartment must be quicker than the CTL assay, so that clinical measures can be taken prior to the development of HCMV disease in patients where the T cell response is not sufficient. It is conceivable that by monitoring T cell reactivity towards HCMV in immunosuppressed individuals, clinically relevant lower limits of protective T cell reactivity, possibly related to viral load, may be defined. Values measured below such limits could then prompt antiviral therapy. The potential usefulness of such therapeutic indicators is documented by the widespread prophylactic use of antiviral agents following solid-organ or bone marrow transplantation. In establishing such a method for monitoring T cell reactivity several key issues will have to be addressed: Firstly, it has to be established whether the T cell responses measured following rapid cytokine induction assays represent a clinically relevant parameter. This will be the most critical point, since absence of a correlation with the clinical situation would make the assay useless. A particular problem in this regard is the fact that while the CD4+ T cell response can be measured by stimulation with complete viral lysate (which should contain all possi-

ble CD4+ T cell epitopes), peptides have to be used for stimulating CD8+ T cell responses (while the commonest length of CD8+ T cell epitopes is 9 amino acids, in our own experience peptides up to 15 amino acids in length can be used) [2]. Peptides are necessary for the external loading of MHC class I molecules on antigen-presenting cells (APCs). The processing of peptides following phagocytosis is not likely to result in class-I-associated presentation, and the transfection of APCs with complete or partial HCMV proteins would certainly be too time-consuming. Peptides used for stimulation, however, have to be selected from proteins against which CD8+ T cell responses are directed, and the choice of such proteins is extremely limited. Presently, the majority of known CD8+ T cell epitopes are found in the pp65 protein [14]. It is an additional disadvantage that responses to such select peptides may represent only a small proportion of the overall CD8+ T cell response to the respective or other HCMV proteins, because other potential epitopes are not accounted for. Judging the relevance of the CD8+ T cell response measured after stimulation with one particular peptide would therefore be very difficult. These limitations also apply to the tetramer technology. Secondly, and as a result of the aforesaid, it has to be determined whether both CD4+ and CD8+ T cell responses need to be taken into account. Current thinking favors a model where HCMV defense depends on cytotoxic CD8+ T cells which lyse HCMV-infected target cells. In analogy with the T cell response to hepatitis B virus [15], other CD8+ T-cell-dependent control mechanisms apart from cytotoxicity may exist in HCMV infection which have not yet been demonstrated. On the other hand, it has been shown that adoptive T cell transfer protocols are more effective in preventing HCMV disease in bone marrow recipients when CD4+ T cells are also transferred [16]. Thirdly, we need to know whether there is such a thing as a ‘protective’ level of T cell defense (CD4+ or CD8+) which can be defined in relative or absolute counts of responding T cells. For the monitoring of anti-HCMV T cell responses to be useful, minimum protective levels of T cell defense have to be established. Subpopulations of reactive T cells may also be considered in this regard, for example the CD57+/CD8+ T cell subset, which has been shown to contain the bulk of rapidly inducible HCMVspecific CD8+ T cells [17]. Also, the interindividual variations in such protective responses/frequencies must be analyzed, because if too much variation in protective levels exists, the assay may again be useless.

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Fourthly, the role of immunosuppression in such assays is important. A recent study using PMA/ionomycin stimulation of peripheral blood mononuclear cells (PBMC) followed by flow cytometric detection of T cell activation (intracellular cytokines), concluded that the effect of immunosuppression could be monitored by such testing [18]. In our own experience [unpubl. results], the effects of in vitro stimulation with PMA can be strongly reduced by preincubation of PBMC with cyclosporin A. It may therefore make a difference whether such tests are done in whole blood or in washed PBMC. As yet, no systematic study has addressed this question. Regardless of which of the two possibilities gives better results, it has to be established which of the two gives more relevant results. Using washed PBMC, we were able to observe that during the initial high-dose immunosuppressive in-

duction phase after kidney transplantation, the CD4+ T cell response (i.e. relative frequency of reactive HCMVspecific CD4+ T cells) declines. So, washing the cells did not wash away the effect of immunosuppression; nevertheless, it may have altered the response. Finally, the methods for measuring these responses must be standardized so that they can be compared between different laboratories. Important issues with regard to standardization are specimen handling, the stimulation protocol, use of suitable controls and instrument setup, to mention only a few. The European Work Group on Clinical Cell Analysis, a group of 16 core facilities and 20 associated laboratories (Chairman G. Schmitz, Regensburg) is presently conducting a multicenter study to address not only the issue of standardization, but also many other of the other above-mentioned issues.

References 1 Waldrop SL, Pitcher CJ, Peterson DM, Maino VC, Picker LJ: Determination of antigen-specific memory/effector CD4+ T cell frequencies by flow cytometry: Evidence for a novel, antigen-specific homeostatic mechanism in HIVassociated immunodeficiency. J Clin Invest 1997;99:1739–1750. 2 Kern F, Surel IP, Brock C, Freistedt B, Radtke H, Scheffold A, Blasczyk R, Reinke P, Schneider-Mergener J, Radbruch A, Walden P, Volk HD: T-cell epitope mapping by flow cytometry. Nat Med 1998;4:975–978. 3 Tan LC, Gudgeon N, Annels NE, Hansasuta P, O’Callaghan CA, Rowland-Jones S, McMichael AJ, Rickinson AB, Callan MF: A re-evaluation of the frequency of CD8+ T cells specific for EBV in healthy virus carriers. J Immunol 1999;162:1827–1835. 4 Altman JD, Moss PAH, Goulder PJR, Barouch DH, McHeyzer-Williams MG, Bell JI, McMichael AJ, Davis MM: Phenotypic analysis of antigen-specific T lymphocytes [published erratum appears in Science 1998;280:1821]. Science 1996;274:94–96. 5 Quinnan GV Jr, Burns WH, Kirmani N, Rook AH, Manischewitz J, Jackson L, Santos GW, Saral R: HLA-restricted cytotoxic T lymphocytes are an early immune response and important defense mechanism in cytomegalovirus infections. Rev Infect Dis 1984;6:156–163.

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6 Riddell SR, Walter BA, Gilbert MJ, Greenberg PD: Selective reconstitution of CD8+ cytotoxic T lymphocytes responses in immunodeficient bone marrow transplant recipients by the adoptive transfer of T cell clones. Bone Marrow Transplant 1994;14(suppl 4):S78–S84. 7 Walter EA, Greenberg PD, Gilbert MJ, Finch RJ, Watanabe KS, Thomas ED, Riddell SR: Reconstitution of cellular immunity against cytomegalovirus in recipients of allogeneic bone marrow by transfer of T-cell clones from the donor. N Engl J Med 1995;333:1038–1044. 8 Riddell SR, Watanabe KS, Goodrich JM, Li CR, Agha ME, Greenberg PD: Restoration of viral immunity in immunodeficient humans by the adoptive transfer of T cell clones. Science 1992;257:238–241. 9 Riddell SR, Greenberg PD: Cellular adoptive immunotherapy after bone marrow transplantation. Cancer Treat Res 1995;76:337–369. 10 Barry SM, Lipman MC, Johnson MA, Prentice HG: Respiratory infections in immunocompromised patients. Curr Opin Pulm Med 1999; 5:168–173. 11 Gatell JM, Marrades R, el-Ebiary M, Torres A: Severe pulmonary infections in AIDS patients. Semin Respir Infect 1996;11:119–128. 12 Boeckh M, Boivin G: Quantitation of cytomegalovirus: Methodologic aspects and clinical applications. Clin Microbiol Rev 1998;11:533– 554.


13 Reusser P, Cathomas G, Attenhofer R, Tamm M, Thiel G: Cytomegalovirus (CMV)-specific T cell immunity after renal transplantation mediates protection from CMV disease by limiting the systemic virus load. J Infect Dis 1999; 180:247–253. 14 Wills MR, Carmichael AJ, Mynard K, Jin X, Weekes MP, Plachter B, Sissons JG: The human cytotoxic T-lymphocyte (CTL) response to cytomegalovirus is dominated by structural protein pp65: Frequency, specificity, and T-cell receptor usage of pp65-specific CTL. J Virol 1996;70:7569–7579. 15 Guidotti LG, Rochford R, Chung J, Shapiro M, Purcell R, Chisari FV: Viral clearance without destruction of infected cells during acute HBV infection. Science 1999;284:825–829. 16 Riddell SR, Greenberg PD: Principles for adoptive T cell therapy of human viral diseases. Annu Rev Immunol 1995;13:545–586. 17 Kern F, Khatamzas E, Surel IP, Frömmel C, Reinke P, Waldrop S, Picker LJ, Volk HD: Distribution of human CMV-specific memory T cells among the CD8(pos.) subset defined by CD57, CD27, and CD45 isoforms. Eur J Immunol 1999;29:2908–2915. 18 van den Berg AP, Twilhaar WN, van Son WJ, van der Bij W, Klompmaker IJ, Slooff MJ, The TH, de Leij LH: Quantification of immunosuppression by flow cytometric measurement of intracellular cytokine synthesis. Transpl Int 1998;11(suppl 1):S318–S321.

Kern/Faulhaber/Khatamzas/Frömmel/ Ewert/Prösch/Volk/Reinke

Part I. Immunopathology CMV-Induced Pathomechanisms


Viral Inhibition of Interferon Signal Transduction Colleen M. Cebulla Daniel M. Miller Daniel D. Sedmak Department of Pathology, The Ohio State University College of Medicine, Columbus, Ohio, USA

Key Words Adenovirus W Cytomegalovirus W Ebola virus W Epstein-Barr virus W Hepatitis B virus W Human papilloma virus W Interferon signal transduction W Mumps W Varicella-zoster virus W JAK/STAT pathway

Abstract The type I and II interferons (IFNs) are potent stimulators of antigen processing and presentation and are essential in antiviral immunity. IFNs upregulate the transcription of major histocompatibility complex (MHC) class I and II molecules, associated antigen-processing proteins, and induce the production of direct antiviral effector molecules such as 2),5)-oligoadenylate synthetase, doublestranded-RNA-dependent protein kinase and Mx proteins. It is increasingly evident that viruses have evolved mechanisms to globally inhibit the actions of IFNs through disruption of their signal transduction pathways. Herein, we review the ability and novel mechanisms of several diverse viruses to inhibit IFN-induced JAK/STAT signal transduction. Copyright © 2000 S. Karger AG, Basel

Viral Inhibition of IFN Signal Transduction

Viruses have been in constant coevolution with the mammalian immune system, resulting in a diverse array of mechanisms by which they are capable of inhibiting the effects of innate and adaptive immunity on viral replica-




tion. In this review, we will focus on an emerging paradigm wherein the effectiveness of viruses, particularly human cytomegalovirus (HCMV), to escape direct and indirect antiviral effects is significantly enhanced by their ability to inhibit IFN-stimulated (JAK/STAT) signal transduction.

Antiviral Mechanisms of IFN

Both type I and II interferons (IFN ·/ß and Á, respectively) are major lines of defense against viral infections. IFNs, through activation of diverse antiviral pathways, block multiple steps of viral replication, including entry of the virus into the cell, transcription, translation, maturation, assembly and virion release [1, 2]. Three main proteins mediate these IFN-induced direct antiviral effects. (1) 2),5)-Oligoadenlyate synthetase (2),5)-OAS) interacts with double-stranded (ds) RNA to activate a ribonuclease (RNAse L) which degrades mRNA, preventing viral products from being synthesized [1, 2]. (2) dsRNA-dependent protein kinase (PKR) also inhibits translation of viral products by phosphorylating translation initiation factor eIF-2 [1, 2]. (3) Mx proteins disrupt influenza, vesicular stomatitis virus and herpes simplex virus (HSV) replication, working through an unknown mechanism [1, 2]. In addition to inhibiting viral replication, IFNs augment immune recognition and lysis of virally infected cells [2]. Virally infected cells are recognized by T cells via viral peptides presented in the context of major histocompatibility complex (MHC) class I and II molecules. Type I

Dr. Daniel D. Sedmak Department of Pathology 129 Hamilton Hall, 1645 Neil Ave Columbus, OH 43210 (USA) Tel. +1 614 292 4692, Fax +1 614 292 7072

IFNs upregulate MHC class I protein expression, and type II IFNs upregulate both MHC class I and II [3, 4]. Moreover, IFNs upregulate other antigen-processing machinery, such as subunits of the proteasome [5]. To underscore the importance of IFNs in controlling viral infections, in vivo studies, with a few exceptions, show that experimentally disrupting IFN action enhances viral disease. In studies with neutralizing antibodies to IFN-·/ß, mice infected with mouse hepatitis virus-3, influenza, Sindbis, encephalomyocarditis (EMC) virus, murine cytomegalovirus (MCMV), or HSV-1 were more susceptible to viral infection and pathology than nonIFN-·/ß-depleted controls [2]. In addition, symptomatic infections with Semliki Forest virus, EMC, HSV-1 and -2, Moloney sarcoma, Friend leukemia virus, or polyoma virus in naturally sensitive mice were exacerbated by IFN-·/ß depletion. Similar results were generated in experiments using IFN-·/ß receptor knockout mice [2]. Analyses of the role of IFN-Á in viral infection have produced analogous results, demonstrating that IFN-Á promotes recovery from and clearance of lymphocytic choriomeningitis and vaccinia viruses [2]. Several studies demonstrate the importance of IFNs in controlling human cytomegalovirus (HCMV) disease. Intramuscular IFN-· reduces the replication of MCMV in the spleen and liver of mice and IFN-· receptor knockout mice are 800-fold more susceptible to MCMV infection than their wild-type littermates [6, 7]. Numerous in vitro studies demonstrate that pretreating cells with IFN-· inhibits HCMV replication by decreasing transcription of the immediate-early (IE) HCMV gene products [8, 9]. The antiviral effects of IFN also extend to clinical therapy, as IFN-· treatment significantly reduces the incidence of serious HCMV infections in seropositive renal transplant recipients [10]. In addition to the type I IFNs, studies of IFN-Á in MCMV models demonstrate that IFN-Á is a critical cytokine in controlling acute and chronic CMV infection [7, 11]. In vivo, it has been documented that (1) IFN-Á accounts for the majority of natural killer cell-mediated antiviral effects during acute infection [12, 13]; (2) neutralization of IFN-Á prevents MCMV clearance from the salivary gland and prevents control of MCMV infection [14, 15], and (3) IFN-Á depletion increases MCMV titers in the liver and spleen [13]. In vitro it has been documented that (1) pretreatment of diverse cell types with IFN-Á inhibits HCMV replication [9, 16–19], and (2) IFN-Á pretreatment restores HCMV antigen processing and presentation to HCMV-specific CD8+ T lymphocyte clones [20, 21]. Recently, it has been reported that

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MCMV infection in IFN-Á receptor knockout mice leads to an uncontrolled persistent infection resulting in a severe vasculitis of large arterial vessels [7]. In contrast, there are very few studies demonstrating that IFN-Á positively influences CMV replication [22]. Furthermore, two animal models demonstrate that the IFN-stimulated JAK/STAT signal transduction pathway is critical for controlling CMV infections. STAT1 knockout mice and IFN-· receptor/IFN-Á receptor doubleknockout mice, which are deficient in IFN-stimulated signal transduction and biological responses, are exquisitely sensitive to viral infection [7, 23]. In these mice, acute MCMV infection proceeds unchecked and rapidly leads to death [7, 23].

IFN Signal Transduction

The diverse biological activities of the IFNs are mediated by a conserved signal transduction pathway [4]. IFN-· binds to its receptor (IFNAR1 and IFNAR2), which stimulates the activation of kinases JAK1 and Tyk2 (fig. 1) [24]. JAK1 and Tyk2 phosphorylate each other, the cytoplasmic tail of IFNAR1, STAT1 and STAT2 (signal transducers and activators of transcription). STAT1/STAT2 heterodimers unite with DNAbinding protein p48 to form the transcription factor complex ISGF3, which binds to the IFN-stimulated response element (ISRE) sites in many IFN-·-responsive promoters [4, 25–27]. Alternatively, phosphorylated STAT1 homodimers and STAT1/STAT2 heterodimers can move to the nucleus and bind to elements such as the inverted repeat (IR) element of the IFN regulatory factor-1 (IRF-1) gene to activate transcription in an ISGF3-independent manner (fig. 1) [28, 29]. In the IFN-Á signal transduction pathways, the receptor (IFN-ÁR1 and IFN-ÁR2) binds IFN-Á; associated JAK1 and JAK2 unite, triggering tyrosine phosphorylation (fig. 2) [4, 30]. Phosphorylated STAT1· forms a homodimer called IFN-Á-activating factor (GAF) that binds to the IFN-Á-activated sequence (GAS) elements in the promoters of IFN-Á-stimulated genes to activate transcription [31, 32].

Viral Disruption of IFN Signaling

Recent work demonstrates that viruses have evolved means of disrupting IFN-stimulated JAK/STAT signal transduction (table 1). Adenovirus E1A gene products

Cebulla/Miller/Sedmak

IFN-a/b

IFN-a/b

s

s

s

s s

Stat2 Stat1

s

s

s

s

IFNAR1

s

Stat1/Stat2 heterodimer

s

s

+

s

p48

s

s

IFNAR2

ISGF3

Stat1/Stat1 homodimer

s

s

ISRE

s

s

Fig. 1. Model of IFN-· signal transduction and gene expression. IFN-· binds to its receptor, which stimulates the activation of JAK1 and Tyk2. JAK1 and Tyk2 phosphorylate each other, the cytoplasmic tail of IFNAR1, and STAT1 and STAT2. STAT1/ STAT2 heterodimers unite with p48 thereby forming ISGF3, which binds to ISRE sites in many IFN-· responsive promoters (left side). Alternatively, phosphorylated STAT1 homodimers and STAT1/STAT2 heterodimers can move to the nucleus and bind to elements such as the IR element of the IRF-1 gene to activate transcription in an ISGF3independent manner (right side).

Tyk2

Jak1

IR

s

IFN-g

Jak2 s

s

s

s s s

s

IFN-gR1

s

s

Jak1

IFN-gR2

Stat1a GAF

Fig. 2. Model of IFN-Á signal transduction.

GAS element

s

s

Upper left shows signaling components. Upon binding, IFN-Á, IFN-ÁR1 and IFNÁR2 and associated JAKs unite (upper right) triggering tyrosine phosphorylation (black dots). Phosphorylated STAT1· forms a homodimer called GAF that binds to GAS elements in the promoters of IFN-Á stimulated genes to activate transcription.

inhibit IFN-· and IFN-Á signal transduction, thereby protecting E1A-expressing cells from IFN-stimulated antiviral and immunoregulatory responses, and E1A enhances replication by distinct viruses in IFN-treated adenovirusinfected cells [33–38]. Adenovirus E1A gene products decrease p48, and overexpression of p48 can restore IFN· signal transduction in E1A-transfected cells [35]. In addition, in some cell types (HeLa and 3Y1 rat fibroblasts

but not HT1080 cells), E1A also decreases STAT1 levels, further preventing IFN-· and IFN-Á-induced signaling [34, 35, 39]. However, the mechanisms which mediate these decreases are unknown. The ability of human papilloma virus (HPV) to inhibit IFN signal transduction has also been recently investigated since IFN-· treatments of HPV-infected patients have yielded only marginal clinical benefits [40]. The

Viral Inhibition of Interferon Signal Transduction


327

Table 1. Effects of viruses on IFN-induced

(JAK/STAT) signal transduction

Virus

IFN disrupted

Gene name/product Mechanism

Adenovirus

type I and II

E1A

decreases p48 decreases STAT1 (cell type dependent)

Ebola

type I and II

?

blocks formation of ISRE-binding signal transduction complexes; decreases GAS-binding complexes

EBV

type I

EBNA-2

blocks signaling downstream of ISGF3 activation and DNA binding; prevents antiproliferative but not antiviral effects of IFN-·

HBV

type I and II

terminal protein

disrupts ISGF3 formation

HCMV

type I and II

?

decreases JAK1 decreases p48

HPV

type I

E7

disrupts ISGF3 formation and p48 nuclear translocation

Mumps

type I

?

decreases STAT1·

HPV E7 oncoprotein was a candidate for disrupting IFN signaling since (1) HPV-transformed cells disrupted IFN· stimulated responses, (2) patients responding well to IFN-· treatment had lower levels of E7 oncoprotein, and (3) E7 shares sequence and functional homology with the IFN-·-signal-transduction-blocking adenovirus E1A protein [40]. Cells transfected with HPV E7 inhibited the induction of IFN-·-inducible genes, but not IFN-Á-inducible genes. Moreover, these cells lost ISGF3 formation and p48 nuclear translocation, without decreasing p48 levels. The E7 protein could potentially play a direct role in inactivating p48, since it binds p48 [40]. Mumps disrupts IFN-induced gene expression in infected cells through inhibition of the signaling pathway [41–44]. One recent study demonstrated that the lesion in the signaling pathway was an absence of STAT1·, mediated by a posttranscriptional mechanism [44]. This signaling defect prevented the antiviral effects of IFN against VSV superinfection [42]. Filoviruses, including the lethal Ebola virus, have proved resistant to IFN prophylaxis in infected monkeys [45, 46]. Recently, Ebola virus was shown to inhibit the induction of IFN-induced genes including MHC class I, IRF-1, and 2),5)-OAS. Gel shift experiments demonstrated that both IFN-· and IFN-Á signal transduction was blocked in Ebola-infected endothelial cells (EC); signaling complexes did not bind the ISRE, and only very low levels of GAS-binding complexes were formed [46]. The precise site of inhibition has not been determined,

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but likely candidates include JAK1, STAT1, or STAT2. This disruption of IFN signaling could contribute to the severe immunosuppression and high viral titers seen in Ebola infection. The hepatitis B virus (HBV) terminal protein was demonstrated to block the signaling response to IFN-· and IFN-Á by interfering with the formation of active ISGF3 complexes [47]. It is hypothesized that enough terminal protein may be generated in chronic HBV infections which could make IFN-· therapy less effective and antagonize infected hepatocyte antigen presentation to cytotoxic T lymphocytes. The Epstein-Barr virus (EBV) nuclear antigen 2 (EBNA-2) gene similarly prevents the activation of IFNstimulated genes by IFN-·; however, this inhibition appears to occur downstream of the activation of ISGF3 [48]. Moreover, unlike other viruses which interfere with IFN signaling, this inhibition of signaling prevents the antiproliferative effect of IFN-·, yet it does not prevent the antiviral actions of IFN [49]. HCMV employs many mechanisms enabling it to evade immunosurveillance and persist in the host [50]. Inhibiting IFN-Á and IFN-· signal transduction are some of those important strategies. HCMV is the first virus identified as inhibiting IFN-Á stimulated JAK/STAT signal transduction by targeting JAK1. This lesion results in blocked IFN-Á stimulated signal transduction and gene expression including the induction of MHC class II, class I, and associated antigen-processing genes in EC and


fibroblasts [51, 52]. HCMV IE and/or early (E) genes decrease JAK1 expression through a posttranslational mechanism involving the proteasome [52]. JAK1 is an essential component of type I IFN signaling, suggesting that HCMV could similarly inhibit IFN-· responsiveness by targeting JAK1. However, recent reports that a HCMV virion protein is capable of upregulating a subset of IFN-· responsive genes, independent of IFN treatment and signal transduction, cast doubt upon the ability of HCMV to block IFN-· responsiveness [53, 54]. Our experiments determined, however, that IFN-·induced ISGF3-dependent signaling was blocked, since IFN-· treatment was unable to upregulate MHC class I, 2),5)-OAS, or MxA RNA levels in HCMV-infected cells [55]. Further, ISGF3-independent IFN-·-induced signaling was inhibited, since IRF-1 mRNA induction was blocked in CMV-infected cells [55]. Gel shift experiments revealed that both ISGF3 and STAT1 homodimer formation were inhibited [55]. These findings could be explained by the discoveries that JAK1 as well as p48, an essential component of ISGF3, are both significantly decreased by HCMV [55]. Thus, CMV induces lesions in multiple levels of the IFN-· JAK/STAT signal transduction pathway, decreasing both JAK1 and p48. The ability for CMV to evade the type I and II IFN-stimulated antiviral and immunoregulatory responses in EC probably contributes to EC ‘reservoirs’ of infectious virus.

Another herpesvirus, varicella-zoster virus (VZV) has recently been shown to inhibit IFN-Á induction of MHC class II in fibroblasts, via IE or E gene expression [56]. It is hypothesized that like CMV, VZV may disrupt IFN-Á gene induction by inhibiting the JAK/STAT signal transduction pathway, yet this remains to be determined.

Conclusion

We propose that powerful IFN-mediated antiviral effects have led HCMV to evolve mechanisms of blocking IFN-signaling in infected cells – just as the intense selection pressure of cell-mediated antiviral immune responses has led HCMV to evolve multiple strategies to inhibibt MHC class I expression. Thus, further work directed toward the identification of viral proteins and mechanisms involved in disrupting IFNs may lead to therapies which will thwart these responses and ameliorate the complications associated with persistent infection.

Acknowledgment D.M.M. was a Howard Hughes Predoctoral Fellow. This work was supported by NIH Grant R01 AI38452–03.

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44 Yokosawa N, Kubota T, Fujii N: Poor induction of interferon-induced 2)5)-oligoadenylate synthetase (2-5 AS) in cells persistently infected with mumps virus is caused by decrease of STAT-1·. Arch Virol 1998;143:1985–1992. 45 Peters CJ, Sanchez A, Rollin PE, Ksiazek TG, Murphy FA: Filoviridae: Marburg and Ebola viruses; in Fields BN, Knipe DM, Howley PM (eds): Fields Virology. New York, LippincottRaven, 1996, vol 1, pp 1161–1176. 46 Harcourt BH, Sanchez A, Offermann MK: Ebola virus selectively inhibits responses to interferons, but not to interleukin-1ß, in endothelial cells. J Virol 1999;73:3491–3496. 47 Foster GR, Ackrill AM, Goldin RD, Kerr IM, Thomas HC, Stark GR: Expression of the terminal protein region of hepatitis B virus inhibits cellular responses to interferons · and Á and double-stranded RNA. Proc Natl Acad Sci USA 1991;88:2888–2892. 48 Kanda K, Decker T, Aman P, Wahlstrom M, von Gabain A, Kallin B: The EBNA2-related resistance towards alpha interferon in Burkitt’s lymphoma cells effects induction of IFN-induced genes but not the activation of transcription factor ISGF3. Mol Cell Biol 1992;12: 4930–4936. 49 Aman P, von Gabain A: An Epstein-Barr virus immortalization associated gene segment interferes specifically with the IFN-induced antiproliferative response in human B-lymphoid cell lines. EMBO J 1990;9:147–152. 50 Miller DM, Sedmak DD: Viral effects on antigen processing. Curr Opin Immunol 1999;11: 94–99. 51 Miller DM, Zhang Y, Rahill BM, Kazor K, Rofagha S, Eckel JJ, Sedmak DD: Human cytomegalovirus blocks interferon-gamma stimulated upregulation of MHC class I expression and the class I antigen processing machinery. Transplantation, in press. 52 Miller DM, Rahill BM, Boss JM, Lairmore MD, Durbin JE, Waldman WJ, Sedmak DD: Human cytomegalovirus inhibits major histocompatibility complex class II expression by disruption of the Jak/Stat pathway. J Exp Med 1998;187:675–683. 53 Zhu H, Cong JP, Shenk T: Use of differential display analysis to assess the effect of human cytomegalovirus infection on the accumulation of cellular RNAs: Induction of interferon-responsive RNAs. Proc Natl Acad Sci USA 1997; 94:13985–13990. 54 Navarro L, Mowen K, Rodems S, Weaver B, Reich N, Spector D, David M: Cytomegalovirus activates interferon immediate-early response gene expression and an interferon regulatory factor 3-containing interferon-stimulated response element-binding complex. Mol Cell Biol 1998;18:3796–3802. 55 Miller DM, Zhang Y, Rahill BM, Waldman WJ, Sedmak DD: Human Cytomegalovirus inhibits IFN-alpha-stimulated antiviral and immunoregulatory responses by blocking multiple levels of IFN-alpha signal transduction. J Immunol 1999;162:6107–6113. 56 Abendroth A, Arvin A: Varicella-zoster virus immune evasion. Immunol Rev 1999;168: 143–156.




Murine Cytomegalovirus Homologues of Cellular Immunomodulatory Genes Nicholas J. Davis-Poynter a Mariapia Degli-Esposti b Helen E. Farrell a a Virology

Section, Animal Health Trust, Newmarket, UK, and b Department of Microbiology, University of Western Australia, Nedlands, W. A., Australia

Key Words Herpesvirus W Cytomegalovirus W Immune evasion W G protein-coupled receptor W MHC class I W Chemokine

Abstract The study of ‘molecular mimicry’ or ‘genetic piracy’, with respect to the utilisation of cellular genes captured and modified during the course of virus evolution, has been an area of increasing research with the expansion in virus genome sequencing. Examples of cellular immunomodulatory genes which have been captured from hosts have been identified in a number of viruses. This review concentrates upon studies of murine cytomegalovirus (MCMV), investigating the functions of viral genes homologous to G protein-coupled receptors, MHC class I and chemokines. The study of recombinant MCMV engineered with specific disruptions of these genes has revealed their significance during virus replication and dissemination within the host. In the case of the latter two classes of genes, evidence suggests they interfere with cellular immune responses, although the detailed mechanisms underlying this interference have yet to be delineated. Copyright © 2000 S. Karger AG, Basel

ABC

© 2000 S. Karger AG, Basel 0300–5526/99/0426–0331$17.50/0



Introduction

Several decades of research into the mammalian immune system have revealed a diverse and complex array of interacting mechanisms for the recognition and eradication of viruses, bacteria and other parasites. These mechanisms can be broadly separated into ‘innate’ and ‘adaptive’ arms – the former responding to fairly non-specific signals indicative of infection/aberrant cell behaviour, the latter dependent upon highly specific antigenreceptor interactions. The hostile environment presented by these immune defences provides a strong impetus to the evolution of counteracting immune evasion mechanisms by parasite organisms. Characterisation of the evasion strategies employed by viruses has been the subject of several recent reviews [1–6]. For large DNA viruses, such as pox- and herpesviruses, examination of genomic sequences has revealed the presence of viral genes with striking homology to cellular genes with immunomodulatory functions. In this paper we discuss studies of such ‘hijacked’ genes encoded by murine cytomegalovirus (MCMV). MCMV is a member of the betaherpesvirus family, which also includes human, rat, guinea pig and porcine CMV and human herpesvirus type 6 and type 7 (HHV-6, -7). The CMVs conserve a number of characteristic biological features, which include a relatively long replicative

Nicholas J. Davis-Poynter Virology Section, Animal Health Trust, Lanwades Park Newmarket Suffolk CB8 7UU (UK) Tel. +44 1638 750 657, Fax +44 1638 750 794 E-Mail [email protected]

Table 1. Herpesvirus GCR homologues

CCR-like

CXCR-like UL33 family

UL78 family

UL33 M33 R33 U12 U12

UL78 M78 R78 U51 U51

Betaherpesviruses HCMV US28, US27 MCMV RCMV HHV-6 HHV-7 Gammaherpesviruses HVS HHV-8 MHV-68 EHV-2 E1 AHV-1 EBV

74 74 74 74

Other

E6 A5 BILF1

Underlining indicates where the gene products exhibit chemokine receptor-like functions. MHV-68 = Murine herpesvirus-68.

cycle, the ability to productively infect a variety of cell types (including monocytes and macrophages, which are important during virus dissemination and as sites of latency), the establishment of infection within salivary glands that serve as a major reservoir for virus shedding, and the ability to persist over the lifetime of an infected host [7, 8]. Infection by human CMV (HCMV) rarely causes disease in immunocompetent hosts, but if transmitted to a fetus in utero or following immunosuppression, it can result in potentially fatal disease ranging from multi-organ dysfunction to more localised disease such as pneumonia and retinitis. Experimentally, MCMV has provided a useful model for HCMV infection, as it displays many features of the human disease, including characteristic sequelae of immunosuppression, although unlike its human counterpart, MCMV is unable to cross the placenta [7, 9]. More recently, interest has grown in the use of MCMV as a natural infection system with which to analyse host-virus interactions. In this regard MCMV is particularly useful for studies of viral immune evasion/ subversion since the virus is well equipped with mechanisms finely tuned for interaction with the murine immune system, the viral genome can be manipulated to modify specific virus genes and studies of mouse immunity are more advanced than in any other species, enabling the use of sophisticated tools for the analysis of specific immune control pathways [10, 11].

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Determination of the complete sequence of MCMV has demonstrated its considerable colinearity and conservation with the HCMV genome, confirming the close relationship between these two viruses which had previously been inferred from their observed biological characteristics [12]. The arrangement of the two viral genomes is in accordance with the following scheme: the central region of each genome encodes the majority of the ‘housekeeping’ genes, which are conserved within the herpesvirus group as a whole; the majority of ‘CMV-specific’ genes likely to impart biologically conserved features flank the core region, and genes apparently unique to each virus are located towards the ends of each genome. The lack of primary sequence homology between MCMV and HCMV for genes located in the terminal regions indicates either that these genes were acquired independently by each virus or have diverged significantly from common ancestral genes as the two viruses have become adapted to parasitisation of murine and human hosts, respectively. Our group has concentrated upon analysis of the function of MCMV homologues of cellular immunoregulatory genes, specifically G protein-coupled receptors (GCRs), MHC class I molecules and chemokines.

Herpesvirus GCR Homologues

GCRs are a superfamily of multiply spanning, integral membrane proteins which, following binding to extracellular ligands, trigger intracellular second-messenger pathways via activation of G proteins associated with the intracellular domains of the receptor [13]. The majority of viral GCR homologues (vGCRs) identified to date have been found amongst the beta- and gammaherpesviruses, as indicated in table 1 [12, 14–22a]. Previous reviews and other papers in this issue describe various properties of vGCRs and these will not be covered in detail here [1, 6, 22b, c]. The majority of the herpesvirus vGCRs (CCRlike, CXCR-like and UL33 family) display sequence features characteristic of chemokine receptors and five of these have been shown to share functional characteristitics, namely US28 of HCMV, U12 of HHV-6, open reading frame (ORF)74 of herpesvirus saimiri (HVS) and HHV-8, and E1 of equine herpesvirus (EHV)-2 [19, 23– 26]. The remaining ORFs listed in table 1 show much lower homology to chemokine receptors. In particular, the Epstein-Barr virus (EBV) ORF BILF1, although lacking certain features indicative of a GCR, is conserved with EHV-2 E6 and alcelaphine herpesvirus (AHV)-1 A5, both of which have been postulated as vGCRs [1, 22a].

Davis-Poynter/Degli-Esposti/Farrell

Our studies have investigated the biological function of M33 and M78 of MCMV, vGCRs which are conserved for all betaherpesviruses characterised to date and are therefore likely to perform conserved functions, through the construction and characterisation of recombinant viruses with specific disruptions of these ORFs. During the characterisation of M33 we identified a splice near the 5) end of the transcript, which we found to be a feature conserved with HCMV UL33, and predicted similar splicing for HHV-6 and HHV-7 U12 [27]. The identification of splicing was significant since the resulting N termini of all four polypeptides conform more closely to chemokine receptors than the unspliced ORFs. Disruption of M33 had no effect upon MCMV replication in tissue culture, but resulted in dramatic attenuation in vivo, with an inability to recover infectious virus from salivary glands. Recently, we have similarly investigated the function of M78 and found that, while non-essential for replication in tissue culture, disruption of this ORF leads to significant attenuation in vivo [Fleming et al., unpubl. results]. Investigations of the functions of the related ORFs of rat CMV (RCMV) have been performed with similar results, as reviewed by Vink et al. [22c]. It appears, therefore, that genes in the UL33 and UL78 families may be important for the efficient replication of betaherpesviruses in vivo, although the mechanisms whereby they operate, and in particular the significance of chemokine binding/G protein interaction for their function, have yet to be determined.

CMV-Encoded MHC Class I Homologues

MHC class 1 molecules are pivotal in the control of cellular immune responses to intracellular parasites. Characterisation of CD8+ cytotoxic T cells (CTLs) has demonstrated that activation is tightly controlled by interactions between antigen-specific receptors on the T cell (TCRs) and MHC class I/peptide complexes displayed on the target cell surface. Following virus infection, the appearance of novel viral peptides in association with MHC class I triggers the activation of specific CTLs via their TCRs. These CTLs destroy infected cells and limit the further spread of infection via direct cytolytic mechanisms and the release of antiviral and pro-inflammatory cytokines. It has been noted that various viruses share the ability to interfere with the MHC class I/peptide display pathway. In the case of the CMVs, several distinct mechanisms have been delineated for both HCMV and MCMV, as discussed in recent reviews [5, 28]. Interference occurs

MCMV Piracy of Immunomodulatory Genes

at various steps in the pathway, including peptide generation (HCMV pp65), peptide translocation (HCMV gpUS6), class I maturation (HCMV gpUS2 and gpUS11), intracellular retention of class I (HCMV gpUS3; MCMV gpm152) and diversion of class I to lyosomes (MCMV gpm06). Interestingly, with the exception of pp65, all of the genes involved are members of glycoprotein gene families present at the left and right ends of the HCMV and MCMV genomes [12, 29]. The majority of these glycoprotein gene family members have yet to be ascribed functions, and it is likely that a variety of other mechanisms for interference with MHC class I, or other important immunomodulatory molecules, may be discovered through further characterisation of these genes. In view of the central role of MHC class I in controlling CTL responses, the identification of a virally encoded homologue in the genome of HCMV (UL18) triggered considerable interest, with theories concerning its function concentrating upon possible disruption of CTL recognition of infected cells [30]. It has been demonstrated that the UL18 gene product (gpUL18) shares the ability of cellular MHC class I to bind ß2-microglobulin (ß2m) and endogenous peptides, but is dissimilar in being highly glycosylated [31, 32]. It was initially proposed that gpUL18 may impair MHC class I expression by sequestration of ß2m, but studies of a gpUL18-null recombinant virus have demonstrated that class I downregulation occurs following infection in the absence of UL18 [33]. Subsequently, Fahnestock et al. [32] postulated that gpUL18 may interfere with natural killer (NK) cell recognition of HCMV-infected cells. These authors reasoned that HCMV-dependent downregulation of cell surface MHC class I may render infected cells susceptible to NK cell-mediated cytolysis, since NK cell inhibitory receptors engage cell surface MHC class I, and cells expressing low levels of MHC class I are generally good targets for NK cell attack [34–36]. Since gpUL18 was observed to bind both ß2m and peptide, they proposed that it may serve as an MHC class I ‘decoy’, with the ability to inhibit NK cell functions without activating CTLs. Subsequent studies of NK cell-gpUL18 interactions have yielded contrasting results, both supporting and conflicting with the above hypothesis. Reyburn et al. [37] reported that an MHC class I-deficient cell line, normally susceptible to human NK cell-mediated cytolysis, was protected following transfection with UL18. Furthermore, they demonstrated that in the presence of a blocking antibody directed against CD94 (an NK inhibitory receptor) the UL18-transfected cells were as sensitive as untransfected cells to NK-mediated cytolysis, suggesting


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that, in agreement with the decoy hypothesis, gpUL18 may engage CD94. CD94 possess a C-type lectin extracellular domain and forms a dimer with NKG2, the latter bearing intracellular immunoreceptor tyrosine-based inhibitory motif (ITIM) sequences which trigger cellular repression signals. The conclusions of this report have been questioned, however, on the basis that the UL18transfected cells were selected for cell surface ß2m expression (rather than for gpUL18 expression) and may therefore represent clones which have recovered cellular MHC class I expression. For example, the non-classical MHC class I protein, HLA-E, is known to require classical MHC class I signal sequence-derived peptides for stability and cell surface expression. Such a stabilising signal peptide could potentially have been provided by transfected gpUL18 (although the sequence of the signal peptide for gpUL18 does not conform to the consensus required for binding to HLA-E) [38]. Cosman et al. [39] employed expression cloning to identify potential receptors for gpUL18. In these studies, an interaction between gpUL18/Fc and CD94 could not be demonstrated. However, a novel receptor termed LIR-1 was identified. LIR-1 (also known as ILT-2) is an immunoglobulin superfamily gene member, with four extracellular Ig-like domains and four intracellular ITIM domains. Interestingly, variable subsets of human NK cells express LIR-1, whereas the molecule is expressed by the majority of monocytes, myeloid lineage dendritic cells and CD19+ B cells. The ability of LIR-1 to inhibit primary monocytes following activation via CD64 has been demonstrated, suggesting that gpUL18 may potentially serve as an MHC class I decoy to inhibit NK cells or indeed other cell types [40]. Leong et al. [41] raised objections to the role of gpUL18 as a decoy for NK cells, following studies investigating the susceptibility of gpUL18-transfected cells and fibroblasts infected with either gpUL18-null or gpUL18positive HCMV. In these studies, no protective effect of UL18 against NK cell-mediated cytolysis was observed. In contrast, it was found that transient, high-level expression of gpUL18 in a number of cell lines resulted in enhanced killing by NK cell clones compared with control transfectants. It was also observed that the relative susceptibility to NK cell-mediated lysis of wild-type and UL18-null HCMV-infected cells correlated with expression levels of the cellular adhesion molecule ICAM-1, suggesting that the sensitivity to NK cells is critically dependent on positive signals promoting NK cell recognition rather than negative signals such as those that could be provided by gpUL18. In these studies, however, it was not noted whether the NK cell clones and polyclonal popula-

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tions used as effectors were positive for expression of the gpUL18 ligand LIR-1. In an independent report, the susceptibility of human fibroblasts infected with several different HCMV isolates to NK cell-mediated lysis was found to vary significantly. Susceptibility correlated principally with different levels of expression of the cellular adhesion molecule LFA-3, suggesting that positive rather than negative signals were important in determining NK cell susceptibility, in agreement with Leong et al. [42]. The latter report also noted that NK cell susceptibility decreased at late times after infection, via a mechanism dependent upon the expression of virus late genes, although no candidate genes mediating this effect were reported. The identification of an MHC class I homologue encoded by MCMV suggested that class I homologues may be a distinguishing feature of cytomegaloviruses and has enabled the biological significance of these molecules to be investigated during infection of the natural host. Interestingly, while UL18 and m144 share a similar level of amino acid homology to cellular MHC class I, they display low homology to one another and are located at different positions in their respective genomes (left end for UL18, right end for m144). Furthermore, whereas UL18 has largely preserved the ·1, ·2, ·3 domain structure of cellular MHC class I, m144 carries a substantial deletion within the putative ·2 domain. Consistent with these sequence differences, gpm144 lacks the ability to bind endogenous peptides previously noted for gpUL18; however, both viral proteins conserve the ability to bind ß2m [32, 43]. These observations may be interpreted as evidence that UL18 and m144 have been acquired independently, following evolutionary segregation of the virus progenitors for HCMV and MCMV. However, the sequences of UL18 and m144 may have diverged from a common progenitor as a result of HCMV and MCMV adapting to their respective hosts and, since the genes appear to have diverged equally from cellular MHC class I, it is likely that they were captured from the host genome at approximately the same time. Moreover, there is evidence within the MCMV genome of a recombination event in an ancestral virus between the left and right ends of the genome which may explain the contrasting positions for UL18 and m144 [12]. Thus, in MCMV a single member of the right-hand glycoprotein gene family (m17) is located at the left end of the genome and m144 is positioned adjacent to the first gene of the right-hand gene family (m145). It has recently been reported that RCMV also encodes an MHC class I homologue which shares the relative genomic position and general features of m144


[Beisser et al., unpubl. results, 44]. Further sequencing studies of guinea pig and porcine CMV should indicate whether possession of an MHC class I homologue is a conserved feature of the CMV family and, if so, whether these ORFs are likely to have arisen independently or be derived from a common ancestral gene. In contrast to the conflicting results concerning the function of gpUL18, studies of the MCMV MHC class I homologue encoded by m144 have clearly demonstrated an important function of m144 in protecting against NK cell-mediated clearance. NK cells are known to be critical for the control of MCMV infection during the first few days following intraperitoneal inoculation, prior to the development of CTL-mediated clearance mechanisms [45–47]. An MCMV gpm144 null recombinant (K¢m144) was found to be highly attenuated for replication in the spleen, liver and lungs during the early acute phase of infection following intraperitoneal inoculation, but was able to replicate to wild-type levels at a later stage following dissemination to salivary glands [48]. Monoclonal antibody-mediated depletion experiments demonstrated that the observed attenuation of K¢m144 was due to more efficient clearance of the virus in an NK celldependent manner. Subsequent studies have indicated that both cytokine-mediated and direct cytolytic mechanisms are important for the increased clearance of K¢m144 compared with wild-type virus [44, Farrell et al., unpubl. results]. Additional studies have been directed at determining whether gpm144 acts directly to inhibit NK cell recognition of target cells or whether indirect effects upon NK cells may be involved. Kubota et al. [49] investigated whether gpm144 conferred protection on transfected cells in vitro against NK cell-mediated cytolysis. It was found that Raji cells (an EBV-transformed human B cell line) were afforded modest protection against antibody-dependent cell-mediated cytolysis by IL-2-activated murine NK cells. Independent studies using a murine TAP-2-deficient T cell lymphoma cell line, RMA-S, sensitive to NK cell recognition due to deficient MHC class I expression at the cell surface, have demonstrated the ability of gpm144 to protect against direct NK cell-mediated cytolysis [50]. In these studies, m144-transfected RMA-S cells were partially protected against lysis mediated by IL-2-stimulated, but not resting, NK cells. We have further demonstrated that gpm144 affords partial protection against the cytolytic activity of NK cells stimulated in vivo by MCMV infection [Degli-Esposti et al., unpubl. results]. The fact that pre-stimulated rather than resting NK cell activity was modulated by gpm144 suggests that the proportion of

cells expressing a putative receptor for gpm144 may increase following appropriate stimulation. Further studies comparing m144-transfected and untransfected cells in an in vivo tumour rejection mode have confirmed that gpm144 can protect against NK cell-mediated cytolytic clearance mechanisms [50]. Thus, m144-transfected RMA-S cells, injected intraperitoneally into C57B1/6 mice, were less susceptible to NK cell-mediated clearance than control lines. Additional observations made during these studies indicated that gpm144 may have effects in addition to directly inhibiting NK cell-mediated cytolysis. Thus, the number and relative cytolytic activity of NK cells recovered from the sites of tumour challenge were found to be reduced for m144 versus control transfected RMA-S cells. These results suggest that m144 may inhibit the production of inflammatory cytokines/chemokines responsible for the recruitment and activation of NK cells. Interestingly, it has been observed that a significantly enhanced inflammatory response follows infection with K¢m144 compared with wild-type MCMV, consistent with gpm144 having an anti-inflammatory effect. Further studies investigating this possibility are currently under way. Two potential mechanisms of action of viral MHC class I homologues (vMHCI) have been depicted schematically. These models, whereby a vMHCI may interfere with NK cell-mediated clearance of target cells, operate via distinct mechanisms but are not mutually exclusive. The first model (fig. 1a) represents the initial NK cell ‘decoy’ hypothesis, whereby cells susceptible to NK cell recognition due to decreased levels of cellular MHC class I (following MCMV infection or due to inherent mutation, e.g. TAP deficiency), are protected by virtue of vMHCI engagement of an inhibitory receptor expressed on the NK cell surface. In this model, the protection afforded by vMHCI is analogous to the protection afforded by cellular MHC class I via engagement of NK cell inhibitory receptors such as KIRs (primate), Ly49s (rodent) or CD94/ NKG2 (primate and rodent). The second model (fig. 1b) represents indirect inhibition of NK cells, whereby the activities of accessory cells required for efficient recruitment/activation of NK cells (e.g. via chemokine/cytokine release) are repressed by an interaction with vMHCI. It is known that cells of the monocyte lineage (in particular dendritic cells) are capable of stimulating NK cells, e.g. via the production of soluble mediators such as IL-12 or IFN-·/ß [51, 52]. Furthermore, it has recently been reported that dendritic cells are important for efficient NK cell function via an undefined mechanism requiring direct cell-to-cell contact and independent of IL-12 or IFN-



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their cellular distribution and potential to modulate NK cell function (either directly or indirectly) and by further characterisation of the effect of gpm144 upon immune responses generated in vivo.

Herpesvirus Chemokine Homologues

Fig. 1. Possible mechanisms for MHC class I inhibition of NK cells. a Direct inhibition. The virus-infected cell has downregulated cell

surface cellular MHC class I (M), but displays viral MHC class I (V). NK cells (NK) are directly inhibited via vMHC1 engagement of NK cell inhibitory receptors (IR). As a result, positive signals (e.g. mediated via engagement of NK activatory receptors, such as CD2 or NK1.1) are counteracted. b Indirect inhibition. vMHC1 displayed by the virus-infected cell engages inhibitory receptors present on accessory cells (AC). As a result, AC stimulation of NK cells (e.g. via cytokine release or direct contact) is blocked. Consequently, NK cell activation is inhibited.

·/ß [53]. The identification of LIR-1 as a potential inhibitory receptor expressed on monocytes/dendritic cells and able to engage both cellular class I and HCMV gpUL18, suggests that the potentiation of antiviral clearance mechanisms (e.g. NK cells) mediated by monocytes/dendritic cells may indeed be modulated via interactions with cellular or viral MHC class I. Current studies of m144 are directed at testing each of the above models, by attempting to identify receptors for gpm144 and determining

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Chemokines are typically small (7–15 kD), secreted polypeptides which are important modulators of inflammatory responses and lymphocyte trafficking [54, 55]. They are classified according to the arrangement of characteristic cysteine residues, with the majority identified to date being classified as either alpha (CXC) or beta (CC) chemokines, although more recently gamma (C) and delta (CX3C) chemokines have also been identified. Chemokines are active against a broad range of cell types, including eosinophils, neutrophils, monocytes, NK cells, T cells and B cells. The response of cells to chemokines is mediated by chemokine receptors (members of the GCR superfamily); thus the behavior of cells in the presence of specific chemokines is determined by the chemokinebinding characteristics of the chemokine receptors which they express. Typically, cells are triggered to cross vascular endothelia and migrate through tissues in response to a chemokine concentration gradient [56]. During virus infection, therefore, the concentration and profile of chemokines secreted at the site of infection are important in determining the numbers and characteristics of the cells recruited during the inflammatory response. The importance of chemokines during virus infection is evidenced by viral countermeasures identified in poxviruses and herpesviruses, which include chemokine receptor homologues (noted above), chemokine binding proteins [3, 6, 57, 58] and viral chemokine homologues. Amongst the herpesviruses, chemokine homologues have been identified within the genomes of Marek’s disease virus, HHV-8, HHV-6, HCMV and MCMV [15, 59–62]. Interestingly, laboratory passaged, avirulent strains of HCMV (e.g. AD169) possess sizeable deletions in their genomes when compared with low-passage clinical isolates (e.g. Toledo), resulting in the loss of a number of genes, including the chemokine homologues and indicating that although these genes are non-essential for virus replication in tissue culture they may be important for virus replication/pathogenesis in the host [61]. For the betaherpesviruses, the genes encoding chemokine homologues appear to have been acquired independently, since they occur at distinct positions (relative to conserved genes) in their respective genomes and display no primary


Fig. 2. Sequence comparisons between the MCMV chemokine

homologue (m131/129) and three cellular chemokines: human macrophage inhibitory protein beta (huMIP3ß), human MIP4 alpha (huMIP4·) (both CC chemokines) and rat lymphotactin (ratLTN, a C chemokine). For m131/129, the C-terminal 171 amino acids have been omitted; all other sequences are shown in their entirety. Conserved cysteins, characteristic of CC chemokines, are underlined.

Amino acid conservation between the aligned sequences is indicated by ‘*’ (identical amino acids) or ‘.’ (similar amino acids). Residues which are identical between m131/129 and one or more cellular chemokines are highlighted in black type. Sequence alignments were compiled using the program E CLUSTAL W and computing facilities provided by the Australian National Genome Information Service.

sequence homology to one another. Notably, whereas HCMVs have been found to encode three chemokine homologues [UL146 (Toledo), UL147 (Toledo, Towne), UL152 (Towne)], all of which are members of the CXC family, MCMV encodes a single, CC chemokine homologue. The MCMV chemokine homologue (also known as MCK-1) was initially identified by MacDonald et al. [62] as being encoded by the m131 ORF (also known as ORF HJ1) and was shown to be transcribed with the kinetics of a late gene. The predicted translation product of m131 is 81 amino acids, consistent with the size of cellular chemokines. Unexpectedly, however, it has subsequently been shown that m131 is expressed as a spliced transcript, resulting in fusion to the m129 ORF with the predicted addition of 199 amino acids to the C terminus [63, 64]. Moreover, antisera raised against gpm131/129 detect the expression of multiple polypeptide species of around 40 kD (in transfected or MCMV-infected cells) which, following deglycosylation, decrease in size to approximately 30 kD. This deglycosylated size is consistent with the predicted size (29.3 kD) of the m131/129 gene product (also known as MCK-2). An alignment between m131/129 and three cellular chemokines to which it is most similar (by FASTA analysis) is displayed in figure 2. To date, there is only one example of a cellular chemokine possessing a substantial additional sequence at the C terminus, namely fractalkine (a CX3C chemokine), which exhibits a mucinlike C-terminal stalk and transmembrane anchor to the

cell surface [65]. There is, however, no significant homology between m129 and the fractalkine C terminus, and m129 does not appear to encode a transmembrane region near the C terminus. Thus, the significance of m129 to the function of m131/129 is currently unknown. In order to investigate the function of m131/129 during infection we have characterised m131/129 null recombinants (¢m131/129) in comparison with wild-type virus [64]. Two days after infection by the intraperitoneal route, no differences were noted in the titres recovered from the spleen or liver between ¢m131/129 and wildtype virus. As the infection progressed, however, titres for the recombinant viruses were found to be significantly lower than those for wild-type virus in the spleen and liver, and at later time points in the salivary glands. Depletion studies were performed to determine whether the attenuation may be due to enhanced cell-mediated clearance of the ¢m131/129 viruses. These studies confirmed that ¢131/129 viruses are cleared more efficiently than wild-type from the spleen and liver through the action of NK cells and, to a lesser extent, T cells. In contrast, the depletion of NK cells or T cells did not have a significant effect upon virus titres recovered from the salivary glands. Unexpectedly, histological analysis of the foci of infection in the liver indicated a decreased rather than increased recruitment of inflammatory cells in the absence of m131/ 129, suggesting that m131/129 possesses pro-inflammatory activity.



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At least four distinct mechanisms can be proposed for the action of a viral chemokine (vCK) such as m131/129. Firstly, vCK may act as a chemokine antagonist (fig. 3a) by binding to chemokine receptors and thereby preventing interaction with cellular chemokines, without itself triggering a signal from the receptor. In this way vCK may exhibit anti-inflammatory activity, by delaying the onset and effectiveness of the cellular immune response. Chemokine receptor-inhibitory properties have previously been demonstrated for vCKs encoded by molluscum contagiosum (MC148) and HHV-8 (vMIP-II) [66, 67]. In contrast, a vCK may serve as a chemokine agonist by directly mimicking both the binding and receptor-activation properties of cellular chemokines (fig. 3b, c). Such activities may be advantageous for the virus by promoting the recruitment of susceptible target cells. This may be particularly important if myeloid or lymphoid cells able to cross vascular endothelia are required for virus dissemination (fig. 3b). Such a mechanism has recently been proposed for HCMV, following the demonstration that the vCK UL146 is chemotactic for neutrophils, which are permissive of virus replication [68]. Alternatively, a vCK may recruit cells which are ineffective at controlling virus infection or which actively repress anti-viral responses via the secretion of immune-suppressive cytokines and thereby promote immune evasion (fig. 3c). For example, HHV-8 vMIP-II has been found to trigger chemotaxis of

Fig. 3. Possible mechanisms for the action of vCK homologues. a vCK blockade of cellular chemokine receptors. Virus-infected cells

secrete vCK molecules (grey echelons) which bind to chemokine receptors on the surface of immune effector cells without inducing cellular activation. As a result, binding of chemokine receptors by cellular chemokines (black diamonds) is prevented. Consequently, chemotaxis of immune effector cells across the vascular endothelium to sites of virus infection is inhibited. b vCK recruitment of susceptible cells. Following engagement of vCK by receptors on the surface of susceptible cells, chemotaxis to the site of infection is promoted. Subsequently, susceptible cells become infected. c vCK recruitment of inappropriate cell types. Following engagement of vCK by receptors on the surface of immune effector cells, chemotaxis to the site of infection is promoted. The recruited cells subsequently inhibit antiviral clearance mechanisms, e.g. by the secretion of anti-inflammatory cytokines (–ve). d vCK modulation of infected cell behaviour. i vCK engagement of cellular (or viral) receptors triggers activatory signals leading to enhanced virus replication. ii vCK engagement of cellular (or viral) receptors triggers inhibitory signals leading to an inhibition of virus replication and a switch from the lytic cycle to latent infection.


eosinophils via interaction with CCR3, and HHV-8 vMIP-I has recently been reported to be chemotactic for Th2-type T cells via interaction with CCR8 [67, 69, 70]. A fourth possibility is that a vCK may modulate the behaviour of virus-infected cells via interaction with a cellular or viral chemokine receptor (fig. 3d), thereby promoting virus replication (fig. 3di) or (in the case of herpesviruses) triggering a switch between the lytic and latent phases of infection (fig. 3dii). In this regard it has recently been found that HHV-8 vMIP-II downregulates the activity of the viral chemokine receptor ORF 74 [71]. The available data for MCMV m131/129 suggest that the viral chemokine has immunosuppressive effects against NK and T cell-mediated clearance, in the absence of a general anti-inflammatory effect, and promotes virus dissemination to, or replication within, salivary glands (with the latter effect not apparently affected by NK or T cell depletion). Furthermore, it has been reported that m131 (in the absence of m129) is able to trigger a calcium flux in cultured monocytic cell lines and monocytes from MCMV-infected mice, consistent with it possessing proinflammatory activities [72]. Since monocytes are an important reservoir for MCMV infection and dissemination within the host, this may be linked to the observed replication defect of ¢m131/129 in salivary glands. All reported experimental observations may be explained by m131/129 acting as a chemokine agonist for cell types supporting replication-dissemination and also for cells which dampen anti-viral immune responses (fig. 3b, c). However, other activities, as outlined above, cannot be excluded. Indeed, it should be noted that HHV-8 vMIP-II exhibits both agonist and antagonist chemotactic properties against different cell types and is able to modulate intracellular activation via engagement of the viral chemokine receptor.

Conclusions

The diversity of immune evasion/subversion mechanisms exhibited by viruses attests to both the complexity of host immune responses and the adaptive pressures these impose upon viral pathogens. Large DNA viruses, such as poxviruses and herpesviruses, have been found to capture host immunomodulatory genes as a means of gaining a selective advantage. Studies of such genes, in addition to elucidating specific virus-host interactions and their impact upon pathogenesis, provide useful tools with which to dissect the molecular basis of cellular immune responses. Viruses that naturally infect mice,


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such as MCMV, are particularly useful due to the availability of reagents to characterise and modulate the immune responses of the host and thereby directly assess interactions between host immunity and viral immunomodulatory functions during infection.

Acknowledgments The authors’ research was supported by grants from the National Health and Medical Research Council (Australia) and the British Biotechnology and Biological Sciences Research Council (UK). N.D.-P. is supported by a Tetra Laval Fellowship.

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Molecular Mimicry by Cytomegaloviruses Function of Cytomegalovirus-Encoded Homologues of G Protein-Coupled Receptors, MHC Class I Heavy Chains and Chemokines

Cornelis Vink Patrick S. Beisser Cathrien A. Bruggeman Department of Medical Microbiology, Cardiovascular Research Institute, University of Maastricht, Maastricht, The Netherlands

Key Words Cytomegalovirus W G protein-coupled receptors W MHC class I W Chemokines W Pathogenesis W Recombinant virus W Animal models

Abstract Cytomegaloviruses (CMVs) are well known for their high prevalence rate within host populations as well as their ability to induce lifelong infections. To maintain a persistent and stable relationship with their host, CMVs have evolved various molecular mechanisms to both control host cell metabolism and evade immune surveillance. Among the viral gene products that are likely to be involved in these processes are homologues of cellular G protein-coupled receptors, MHC class I molecules and chemokines. The viral genes encoding these homologues have probably been pirated by the viruses during a long pathogen/host coevolution. In this report, we will discuss the possible functions of these homologues in the pathogenesis of CMV infections. Copyright © 2000 S. Karger AG, Basel

ABC

© 2000 S. Karger AG, Basel 0300–5526/99/0426–0342$17.50/0



Introduction

Cytomegaloviruses (CMVs) are species-specific ß-herpesviruses which cause acute, persisting and latent infections in both humans and animals. Although human CMV (HCMV) infections of immunocompetent individuals usually run an asymptomatic course, infections of immunocompromised individuals (e.g. AIDS patients, organ transplant recipients and neonates) can have lifethreatening consequences. A characteristic that is shared by all herpesviruses is the ability to induce a latent, lifelong infection. Obviously, such a lifelong interaction requires the virus to be highly adapted to its host and vice versa. Most importantly, the virus will have to employ strategies to remain hidden from the host’s immune system. In principle, this can be achieved by the virus going into a dormant state in which only a subset of viral genes is expressed and targets for host immune responses are not displayed. Recent studies have indicated that herpesviruses use a variety of strategies to interfere with the immune system of the host [1–4]. Most notably, several herpesvirus-encoded proteins have been identified that inhibit either synthesis, surface expression or peptide loading of cellular MHC class I molecules. In addition, various herpesvirus genes have been identified which encode homologues of immune effector

Dr. C. Vink Department of Medical Microbiology, University of Maastricht PO Box 5800 NL–6202 AZ Maastricht (The Netherlands) Tel. +31 43 387 6669, Fax +31 43 387 6643, E-Mail [email protected]

Table 1. CMV genes encoding homologues

of important immune effector molecules or regulatory proteins of the hosta

HCMV RCMV MCMV

GPCR genes

MHC class I genes

Chemokine genes

US27, US28, UL33, UL78 R33, R78 M33, M78

UL18 r144 m144

UL146, UL147, UL152 m131/129

a

The reported genes are from references [7] (HCMV GPCR genes); [39] (UL18); [35] (UL146, UL147, UL152); [10] (R33); [11] (R78), and [9] (M33, M78, m144, m131/129). The RCMV r144 sequence is from Beisser et al. [29a].

molecules or regulatory proteins of the host. In this report, we will focus on CMV genes which have the potential to code for homologues of G protein-coupled receptors (GPCRs), MHC class I heavy chains and chemokines. The possible functions of these genes and their products will be discussed.

CMV Homologues of GPCRs

Viral GPCRs GPCRs are members of a large and diverse family of receptors that function in signal transduction through cell membranes [5]. All GPCRs are composed of a central core domain consisting of seven transmembrane (TM) helices (TM-I to TM-VII) connected by three intracellular and three extracellular loops [6]. The majority of these receptors activate G proteins and are capable of transducing messages as different as photons, organic odorants, lipids, nucleotides, peptides and proteins. Thousands of GPCR variants are encoded by genes of eukaryotes as well as prokaryotes. In addition, some GPCRs are encoded by genes of poxviruses and herpesviruses. It is likely that these genes have been ‘hijacked’ by the viruses during the coevolution of pathogen and host.

to sequences of cellular chemokine-binding GPCRs (fig. 1) [4]. In addition, the US28 protein (pUS28) has been reported to be capable of binding CC chemokines (or ßchemokines), such as RANTES, MIP-1· and MCP-1, hence triggering the mobilization of intracellular Ca2+ [12–15]. It has been suggested that pUS28 is responsible for ß-chemokine sequestration in HCMV-infected cells [16]. Interestingly, pUS28 was recently shown to selectively recognize fractalkine (or neurotactin), which is a membrane-associated CX3C chemokine [17]. Since fractalkine is expressed on putative CMV target cells, such as endothelial cells, it was hypothesized by Kledal et al. [1] that the highly specialized pUS28-fractalkine interaction may be involved in the cell-to-cell transfer of HCMV.

HCMV US27 and US28 Genes Within the genomes of HCMV [7, 8], murine CMV (MCMV) [9] and rat CMV (RCMV) [10, 11], several genes have been identified that are capable of encoding homologues of host cellular GPCRs (table 1). HCMV carries four of these genes: US27, US28, UL33 and UL78 [7, 8]. Only two of these, UL33 and UL78, were found to have counterparts in RCMV (R33 and R78, respectively [10, 11]) as well as MCMV (M33 and M78, respectively [9]). Of the predicted amino acid sequences derived from the CMV GPCR-like genes, those encoded by HCMV US27 and US28 were found to have the greatest similarity

The UL33 Gene Family Due to the species specificity of HCMV, it is difficult to study the function of the HCMV US27 and US28 genes in vivo. Moreover, these genes do not have counterparts within the genomes of other (animal) cytomegaloviruses. In contrast, UL33- and UL78-like genes are conserved among all ß-herpesviruses (fig. 1), which illustrates their importance for the biological characteristics of these viruses. The functions of these genes can therefore also be studied in vivo, using animal models. Currently, the UL33 gene family consists of five members: HCMV UL33, MCMV M33, RCMV R33 and human herpesvirus 6 and 7 (HHV-6 and -7) U12 [18, 19]. The predicted amino acid sequences of the proteins encoded by the UL33like gene family have been found to comprise several features characteristic of chemokine receptors [10, 20]. In agreement with this, the HHV-6 U12-encoded protein (pU12) has been reported to be a functional receptor for ß-chemokines in vitro [21]. Whether the other members of the pUL33-like family are similarly capable of binding chemokines and/or other ligands has not yet been described. Nevertheless, the UL33, M33 and R33 genes have been found to be dispensable for the in vitro replica-

Molecular Mimicry by CMV


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Fig. 1. Relationship between host- and CMV-encoded GPCR-like amino acid sequences and MHC class I heavy chain-like sequences. a A phylogenetic tree based on a multiple alignment of GPCR-like amino acid sequences encoded by HCMV US27 (pUS27), US28 (pUS28), UL33 (pUL33) and UL78 (pUL78) [7, 8]; MCMV M33 (pM33) and M78 (pM78) [9]; RCMV R33 (pR33 [10]) and R78 (pR78 [11]); HHV-6 and HHV-7 U12 (pU12) and U51 (pU51) [18, 19]. In addition, the tree includes the amino acid sequences of human chemokine receptors CXCR4 [40] and CCR5 [41], and two nonchemokine receptors (other GPCRs): ß2-adrenergic receptor (beta 2-ad. rec.) [42] and rhodopsin [43]. CLUSTAL W pairwise alignment [44] was set to BLOSUM30 protein weight matrix, gap open penalty = 10, gap extension penalty = 0.1. Multiple alignment was set to BLOSUM series, gap open penalty = 10, gap extension penalty = 0.05, delay divergent sequences = 0.4. b A phylogenetic tree based on a CLUSTAL W multiple sequence alignment of MHC class I-like amino acid sequences encoded by HCMV UL18 (gpUL18) [39], MCMV m144 (gpm144) [9] and RCMV r144 (gpr144) [29a]. Also included are the amino acid sequences of three mammalian MHC class I proteins, rat RT1.AI [45], murine H2-Kd [46] and human HLA-A2 [47]. In the alignment, a PAM250 protein distance matrix was included. Pairwise alignment gap penalty = 3, multiple alignment gap penalty = 10, gap extension penalty = 10.

tion of HCMV [22], MCMV [20] and RCMV [10], respectively. In contrast, both M33 and R33 have been shown to be essential for virus replication in vivo. This was established by using recombinant viruses in which the M33 and R33 genes were deleted from the MCMV and RCMV genomes, respectively. Unlike wild-type (wt) RCMV, R33-deleted RCMV (RCMV¢R33) did not efficiently replicate in salivary gland epithelial cells of immunocompromised rats, which indicated that the recombinant virus is unable to either enter or replicate in salivary gland epithelial cells [10]. A similar observation was made in the murine model for M33-deleted MCMV [20]. Al-

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a

b

so, a significantly lower mortality was seen among RCMV¢R33-infected rats than among wt RCMV-infected rats [10]. Although these data underline the importance of the UL33-like genes in the pathogenesis of CMV infection, the exact function of these genes is still unknown. The UL78 Gene Family The UL78 gene family currently consists of five members: HCMV UL78 [7], RCMV R78 [11], MCMV M78 [9] and HHV-6 and -7 U51 [18, 19]. Similarly to the UL33-like genes, the positions of the UL78-like genes

Vink/Beisser/Bruggeman

a

b

c

Fig. 2. Deletion of the R78 gene from the RCMV genome results in a syncytium-inducing recombinant strain. The immunofluorescence micrographs (! 400) show uninfected rat embryo fibroblasts (a), and rat embryo fibroblasts infected with either wt RCMV (b) or recombinant strain RCMV¢R78c (c) [11]. The cells were stained with phalloidin-rhodamin (to detect F-actin fibers; red) as well as monoclonal antibody RCMV8 plus anti-mouse-fluorescein isothiocyanate (to detect RCMV early nuclear antigens; green).

within the ß-herpesvirus genomes are conserved. However, in contrast to the sequences of UL33-like genes, the sequences of members of the UL78 family are rather divergent (fig. 1). The predicted amino acid sequences derived from the UL78-like genes significantly resemble neither chemokine receptors nor any other of the thousands of GPCRs currently known. The assumption that UL78-like genes have the potential to encode GPCRs is based on three properties: (1) the presence of a 7-TM core domain within the predicted amino acid sequences derived from the UL78-like genes; (2) the presence of two conserved cysteine residues within these sequences, which may play a role in correct folding of the putative GPCRs, and (3) a stretch of amino acids within these sequences which bears similarity to a domain known to be required for G protein coupling [23]. Only a few studies have hitherto reported on the characterization of members of the UL78 gene family. In a study by Menotti et al. [24], the HHV-6 U51 gene product (pU51) was found to be expressed on the cell surface of infected cord blood mononuclear cells, in accordance with its putative function as a plasma membrane-associated protein. However, in cells transfected with a U51 expression construct, transport of pU51 to the cell surface was cell type depen-

dent: cell surface expression of pU51 was seen in activated T lymphocytes and T cell lines but not in other cell types [24]. The relevance of these observations is yet unknown. In our laboratory, we have recently started to study the RCMV member of the UL78 gene family, R78. We have generated two different recombinant virus strains: an R78 null mutant (RCMV¢R78a) and an RCMV mutant encoding an R78-derived GPCR from which the putative intracellular C terminus has been deleted (RCMV¢R78c). Interestingly, these recombinant viruses produced 10- to 100-fold lower virus titers than wt virus in vitro [11]. Also, unlike wt RCMV-infected fibroblasts, fibroblasts infected with the recombinants developed a syncytium-like appearance (fig. 2). The RCMV¢R78a and RCMV¢R78c strains can therefore be considered as the first syncytium mutant CMV strains reported [11]. We have hypothesized that R78 plays a role, either directly or indirectly, in the stabilization of cell-to-cell contacts [11]. In fibroblasts infected with R78deleted viruses, the contacts between neighboring cells may become unstable, leading to plasma membrane fusion. The RCMV R78 gene was also found to play an important part in the pathogenesis of virus infection in vivo: a lower mortality was observed among immuno-



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compromised rats infected with either RCMV¢R78a or RCMV¢R78c than among animals infected with wt virus [11]. Although we did not detect differences in replication between wt and recombinant viruses in vivo, it is possible that the attenuated phenotype that is seen after disruption of the R78 gene is correlated with the aforementioned efficiency of replication of the recombinant viruses in vitro, which was considerably lower than that of wt RCMV [11]. Taken together, our data indicate that RCMV R78, like R33, is important for the replication of RCMV in vivo. Future studies on the UL33- and UL78like genes will focus on detection of the proteins encoded by these genes, both in vitro and in vivo, and the elucidation of signalling pathways in which these viral GPCRs potentially function. Given the important in vivo functions of RCMV R33 and R78 and MCMV M33, it is likely that the corresponding genes of HCMV also serve a vital function in vivo. The gene products of UL33 as well as UL78 can therefore be considered as attractive targets for the development of novel anti-HCMV therapies.

CMV Homologues of MHC Class I Heavy Chains

Mammalian MHC class I proteins are members of the immunoglobulin superfamily. They are polymorphic, consisting of a membrane-bound, cell surface-expressed heavy chain, and an extracellular globulin-like light chain, ß2-microglobulin (ß2m). Two domains of the heavy chain form a groove in which small peptides (antigens) can be presented to cytotoxic T lymphocytes (CTLs) [25]. These peptides are generated by the degradation of intracellular proteins, which can be encoded either by the genome of the cell or by genes of intracellular parasites such as viruses. Upon the presentation of peptides by MHC class I molecules, CTLs can distinguish ‘foreign’ from ‘self’ peptides and thus discriminate between infected and uninfected cells. Subsequently, the infected cells are killed by the CTLs [25]. Many viruses, however, have developed ways to evade CTL-mediated killing. For instance, several herpesvirus proteins have been found to downregulate the antigen presentation of infected cells by interfering with either the synthesis or maturation of cellular MHC class I molecules [for reviews, see ref. 4, 26, 27]. As a consequence, these cells cannot be recognized by CTLs as being infected. Cells expressing class I molecules at a low level, however, are vulnerable to lysis mediated by natural killer (NK) cells. It has been hypothesized that this threat is overcome by both HCMV and MCMV through the expression of homologues of MHC class I heavy chains (en-

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coded by UL18 and m144, respectively), which might serve as decoys to protect infected cells from being killed by NK cells [28, 29]. Support for this hypothesis has come from a study by Farrell et al. [28] on an m144-deleted MCMV strain (¢m144). This recombinant strain was shown to be attenuated during the primary phase of virus infection in mice, whereas its virulence was comparable to that of wt MCMV in NK cell-depleted mice [28]. To study the function of the RCMV homologue of the m144 gene, r144, we generated an r144 knockout virus strain (RCMV¢r144). Like ¢m144 [28], RCMV¢r144 replicated with an efficiency similar to wt virus in various cell lines in vitro [29a]. However, in contrast to what was reported in the murine model [28], we did not detect differences between wt RCMV and RCMV¢r144 in their replication characteristics within infected rats. Firstly, the survival rate among groups of immunosuppressed rats infected with either RCMV¢r144 or wt RCMV was similar. Secondly, the dissemination of virus did not differ between RCMV¢r144- and wt RCMV-infected, immunosuppressed rats, both in the acute and latent phase of infection [29a]. We therefore concluded that the RCMV r144 gene did not play an important role in virus replication in our in vitro and in vivo experimental systems. It is possible that potential differences between RCMV¢r144 and wt RCMV in replication in vivo might be concealed due to the irradiation-induced immunosuppression which is applied in order to be able to study RCMV disease [10, 11]. If the r144 gene product (gpr144) was to serve as a decoy to evade immune surveillance, as was proposed for the MCMV m144-encoded protein (gpm144), an RCMV r144 knockout strain might be attenuated due to efficient, early clearance of this strain by rat NK cells. As a result of immunosuppression, however, the virulence of the r144 knockout strain may be preserved. In contrast to the RCMV/rat model, immunosuppression is not required in the MCMV/murine model in order to establish virusinduced disease. This difference could explain the apparent discrepancies between our findings with RCMV¢r144 and the results of Farrell et al. [28] with ¢m144. Nevertheless, after irradiation and RCMV infection of rats, NK cells are regenerated relatively efficiently and rapidly (between 10 and 15 days after infection) [30]. Consequently, if RCMV gpr144 functioned in the protection from NK cell cytotoxicity, the regenerated NK cell population should be able to eliminate RCMV¢r144 more efficiently than wt virus. However, we were not able to detect differences between wt and recombinant virus in replication in vivo, neither at day 4 nor at day 21 after infection [29a]. It is of course possible that the regenerated NK cell


population of immunosuppressed rats is still impaired in function due to the irradiation. A function similar to that proposed for gpm 144 [28] was attributed to the HCMV UL18 gene product (gpUL18) by Reyburn et al. [29]. They showed that a B lymphoblastoid cell line, 721.221, was protected from NK cytotoxicity after transfection with a vector expressing UL18. In contrast, Leong et al. [31] found that the expression of UL18 resulted in enhanced rather than reduced killing of target cells by NK cells. According to Leong et al. [31], the apparent discrepancies between their results and those of Reyburn et al. [29] could be due to the fact that in the latter study transfected cells were selected on the basis of surface expression of ß2m. This selection could have resulted in enrichment of a population of cells that express endogenous HLA-E, which was recently shown to protect 721.221 target cells from NK cell cytotoxicity [32]. Consequently, the protection of UL18-transfected 721.221 cells from lysis by NK cells could be result of the expression of HLA-E rather than UL18. It was previously suggested by Leong et al. [31] that viral MHC class I homologues may be more important in affecting the function of monocytes and dendritic cells than that of NK cells. This notion was supported by the finding that gpUL18 can interact with a membrane receptor, designated ILT2 [33] or LIR-1 [34], which is expressed predominantly on monocytes and B lymphocytes. Since ILT2/LIR-1 is expressed on only a minor subset of NK cells [34], the physiological relevance of the interaction of gpUL18 with this receptor on NK cells is unclear. Leong et al. [31] hypothesized that the binding of gpUL18 to ILT2/LIR-1 on monocytes or dendritic cells could suppress IL-12 production, which would limit the secretion of IFN-Á by NK cells, thereby altering the early immune response. Interestingly, such a mechanism could explain not only the severely restricted replication of ¢m144 in vivo [28], but also the similarity between RCMV and RCMV¢r144 in their replication characteristics within immunosuppressed rats. In analogy to gpUL18, gpr144 could interact with a receptor expressed predominantly on monocytes or macrophages, which could lead to an immune evasive effect. Interestingly, in support of a putative interaction between gpr144 and leukocytes other than NK cells, we found an increased influx of macrophages and CD8+ T lymphocytes in wt RCMV-infected tissue as compared to RCMV¢r144-infected tissue in a local (rat footpad) infection model [29a]. Whether the increased influx of these leukocyte subtypes is the result of either a direct or indirect interaction with gpr144 is currently under investigation. The identification of a putative re-

ceptor for either RCMV gpr144 or MCMV gpm144 on leukocytes might be crucial in the elucidation of the function of these proteins.



CMV Homologues of Chemokines

Chemokines belong to a family of chemoattractant cytokines that play a role in inflammatory responses to various pathogens. They promote the infiltration of leukocytes to the inflammatory site by both the modulation of leukocyte chemotaxis and upregulation of the expression of leukocyte adhesion molecules [35]. Chemokines bind to target cells via GPCRs and can be classified according to the position of conserved cysteine residues within their predicted amino acid sequences. Currently, four groups of chemokines can be distinguished: (1) the CXC (or ·-) chemokines, which have an intervening nonconserved amino acid residue (X) between the first and second conserved cysteine; (2) the CC (or ß-) chemokines, in which the first two cysteines are adjacent; (3) the C (or Á-) chemokines, which have only two of the four conserved cysteines, and (4) the CX3C (or ‰-) chemokines, which contain three irrelevant intervening amino acids between the first two conserved cysteines. To date, genes encoding homologues of chemokines have been identified in two CMV species only: HCMV (UL146, UL147 and UL152) [36] and MCMV (m131/129) [26, 37, 38]. Although we have recently determined the complete nucleotide sequence of the RCMV genome, we have as yet not been able to identify open reading frames (ORFs) that potentially encode chemokine-like polypeptides [unpubl. results]. The HCMV-encoded chemokine homologues show similarity to CXC chemokines, whereas the MCMV m131/129-encoded protein is more closely related to CC chemokines. Recent data indicate that the m131/129encoded chemokine homologue is considerably larger than the known cellular CC chemokines [26, 38]. This protein, also designated MCK-2 (for MCMV-encoded chemokine 2) [38], is produced from a transcript in which the m131 ORF is spliced at its 3) end to the downstream m129 ORF [26, 38]. To study the function of m131/129, a recombinant MCMV strain was generated in which the m131 ORF was disrupted [26]. This strain (¢m131) was reported to be cleared from the sites of in vivo infection more rapidly than wt virus. This effect was dependent on the function of both NK and CD4+/CD8+ T cells [26]. Also, wt MCMV was found to elicit a stronger inflammatory response than ¢m131 within spleen and liver tissue of infected mice [26]. This suggests that m131 may func-

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tion as a chemokine agonist by recruiting leukocytes to the sites of infection. The wt virus may profit from this if the cell types recruited are critical for virus replication and/or dissemination. The identification of a potential receptor for the m131/129-encoded polypeptide will be crucial in understanding the function of this protein in the pathogenesis of MCMV infection.

Acknowledgments The authors thank Suzanne Kaptein for her critical reading of the manuscript.

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32 Braud VM, Allan DS, O’Callaghan CA, Soderstrom K, D’Andrea A, Ogg GS, Lazetic S, Young NT, Bell JI, Phillips JH, Lanier LL, McMichael AJ: HLA-E binds to natural killer cell receptors CD94/NKG2A, B and C. Nature 1998;391:795–799. 33 Samaridis J, Colonna M: Cloning of novel immunoglobulin superfamily receptors expressed on human myeloid and lymphoid cells: Structural evidence for new stimulatory and inhibitory pathways. Eur J Immunol 1997;27:660– 665. 34 Cosman D, Fanger N, Borges L, Kubin M, Chin W, Peterson L, Hsu ML: A novel immunoglobulin superfamily receptor for cellular and viral MHC class I molecules. Immunity 1997;7:273–282. 35 Mantovani A: The chemokine system: Redundancy for robust outputs. Immunol Today 1999;20:254–257. 36 Cha TA, Tom E, Kemble GW, Duke GM, Mocarski ES, Spaete RR: Human cytomegalovirus clinical isolates carry at least 19 genes not found in laboratory strains. J Virol 1996;70: 78–83.


37 MacDonald MR, Li X-Y, Virgin HW 4th: Late expression of a ß-chemokine homolog by murine cytomegalovirus. J Virol 1997;71:1671– 1678. 38 MacDonald MR, Burney MW, Resnick SB, Virgin HW 4th: Spliced mRNA encoding the murine cytomegalovirus chemokine homolog predicts a ß-chemokine of novel structure. J Virol 1999;73:3682–3691. 39 Beck S, Barrell BG: Human cytomegalovirus encodes a glycoprotein homologous to MHC class-I antigens. Nature 1988;331:269–272. 40 Caruz A, Samsom M, Alonso JM, Alcami J, Baleux F, Virelizier JL, Parmentier M, Arenzana-Seisdedos F: Genomic organization and promoter characterization of human CXCR4 gene. FEBS Lett 1998;426:271–278. 41 Mummidi S, Ahuja SS, McDaniel BL, Ahuja SK: The human CC chemokine receptor 5 (CCR5) gene. Multiple transcripts with 5)-end heterogeneity, dual promoter usage, and evidence for polymorphisms within the regulatory regions and noncoding exons. J Biol Chem 1997;272:30662–30671. 42 Emorine LJ, Marullo S, Delavier-Klutchko C, Kaveri SV, Durieu-Trautmann O, Strosberg AD: Structure of the gene for human beta 2adrenergic receptor: Expression and promoter characterization. Proc Natl Acad Sci USA 1987;84:6995–6999.

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Cytomegalovirus-Induced Transendothelial Cell Migration A Closer Look at Intercellular Communication Mechanisms

M. Scholz R.A. Blaheta J.-U. Vogel H.W. Doerr J. Cinatl Jr. Institut für Medizinische Virologie, Johann-Wolfgang-Goethe-Universität, Frankfurt am Main, Deutschland

Key Words Cytomegalovirus W Transendothelial migration W Adhesion molecules W Cadherin-catenin complex W Metalloproteinases W VLA-5

Abstract A variety of cells such as leukocytes and tumor cells may adhere to endothelial cells and subsequently transmigrate into the solid tissue by involving specific intercellular molecular pathways. One important prerequisite for transendothelial migration is the loosening of endothelial cell-to-cell contact sites, which can be triggered by extravasating cells. Cytomegalovirus (CMV) has obviously evolved the ability not only to influence host cells floating in the blood stream to adhere to endothelial cells, but also to induce the formation of intercellular gaps within the endothelium, resulting in transendothelial migration. These features allow the virus to disseminate and evade the immune system. In coculture experiments with human endothelial monolayers and human CMV (HCMV)-infected neuroblastoma cells or leukocytes, changes in the integrity of the monolayer were observed and further analyzed on the molecular level. For example, HCMV may activate the integrin ß1·5 (VLA5) that triggers adhesion to endothelial cells with subse-

ABC

© 2000 S. Karger AG, Basel 0300–5526/99/0426–0350$17.50/0



quent focal disruption of endothelial cell-to-cell connections. It is hypothesized that a Ca2+-independent pathway following VLA-5 binding disconnects the cadherin-catenin-actin complex within the endothelial cells. The loss of cadherin function causes the loss of contact to the neighboring endothelial cells and thus could represent an important mechanism in HCMV-induced cellular transendothelial migration and disruption of the endothelial integrity. Copyright © 2000 S. Karger AG, Basel

Introduction

In spite of intensive research in cytomegalovirus (CMV) biology during the past decades, the exact localization of the virus is still a matter of debate. It has been reported that CMV-specific owl eye cells can be detected in all organs by histology [1]. Endothelial and epithelial cells especially seem to be infected. CMV probably travels through the body via the blood stream in order to settle down and persist latently in more comfortable sites. Evidence has been obtained that monocytes and granulocytes serve as carriers of the latent virus and thus help CMV to spread throughout the body via the blood stream [2]. The reason why CMV leaves the sites of latency in order to

Dr. Martin Scholz, Institut für Thorax und Kardiovaskuläre Chirurgie Klinikum der Johann-Wolfgang-Goethe-Universität Theodor-Stern-Kai 7, D–60590 Frankfurt am Main (Germany) Tel. +49 69 6301 7109, Fax +49 69 6301 7108 E-Mail [email protected]

Dissemination

Leukocyte manipulation

Fig. 1. Left: Leukocytes/tumor cells infected with CMV adhere to endothelial cells and may infect them. The infected leukocyte may also be triggered to transmigrate through the endothelial barrier. Thus the virus can infect other cells such as fibroblasts. Right: The infected endothelial cell may attract floating leukocytes, e.g. by secretion of chemokines. After cell-to-cell contact the virus can be transmitted to the leukocyte and dissemination may occur via the blood stream. N = CMV.

Adhesion Transmigration

infect other cells at different sites is not known. It is suspected that CMV-infected leukocytes exist in the blood stream not only during natural or iatrogenic immunosuppression but sometimes also in healthy persons without clinically overt symptoms. In addition, it is suggested that these CMV-infected cells are attracted to sites of inflammation (e.g. graft rejection) or that CMV induces leukocyte infiltration [3]. Both scenarios seem to be relevant for the in vivo situation. It can be speculated that CMV distinctly triggers communication between the host cell which floats in the blood stream and the endothelial or epithelial cell. However, an adequate in vitro model with infected leukocytes is not currently available. Experimentally, a neuroblastoma cell line (UKF-NB-4AD169) which is permanently and productively infected with CMV has been established [4]. Compared with the uninfected parental line (UKF-NB-4), the infected variant exhibits augmented metastasizing activity (following subcutaneous injection in BALB/c mice) and modified adhesion and transendothelial migration properties in vitro. Clinically, single case reports exist where CMV has been associated with neuroblastoma progression [5], and in one of our patients, histology revealed the presence of active virus within the tumor cells [unpubl. results]. The in vitro model with the permanently infected cell line UKF-NB-4AD169 may allow detailed studies on the way CMV manipulates cellular interactions during its voyage through the body. Moreover, knowledge about the mechanisms involved in CMV-induced transendothelial migration may be helpful in understanding not only CMV-associated leukocyte infiltration but also the recently discussed relationship between CMV and some tumors.

Several investigators have shown experimentally that CMV infection entails a variety of cellular modifications of the host cell. Moreover, as was recently published by Zhu et al. [6], more than 200 host cell genes are currently known to be modified by CMV. Some of these genes are involved in the transcription of immunorelevant molecules which are suspected to contribute to misleading immune reactions in the host. For example, infected fibroblasts or endothelial cells release enhanced or de novo-induced amounts of chemokines which in turn attract leukocytes towards the infected tissue [3]. The expression of cell membrane adhesion molecules is also influenced by CMV [7–9], which may thus trigger adhesion, rolling and transendothelial migration of the immune cell and also of tumor cells (as discussed further below). In this regard, CMV-induced expression of the adhesion molecule ICAM-1 has been the focus of a number of investigations [7, 10]. However, in coculture experiments it has been shown that CMV-induced ICAM-1 on infected endothelial cells is only partly involved in induced adhesion and migration of leukocytes since the maximum number of adherent cells was actually measured before enhanced membrane expression of ICAM-1 occurred [3]. The exact CMV-induced cellular interactions leading to adhesion and transmigration are not fully understood. In figure 1 the putative CMV-induced leukocyte-endothelial cell interactions are schematically summarized on the cellular level. It may be postulated that CMV manipulates the floating leukocyte to adhere to the endothelium

CMV-Induced Cell Migration


CMV Manipulates Intercellular Communication

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with a sufficiently high avidity to allow rolling and transmigration. The virus might find better living conditions in solid tissue and thus may infect cells such as endothelial cells or fibroblasts where it can persist for decades in a latent state. It has been reported that CMV reactivates when infected monocytes differentiate into macrophages, e.g. upon cytokine stimulation [11]. The exact signals between the endothelial cell and the CMV-carrying leukocyte that are important for this reactivation phenomenon are not known. Moreover, what are the mechanisms that allow the infected cell to transmigrate through the endothelial barrier? Is the transmigration a function of CMVinduced immune mechanisms that result in the loosening of the endothelial junction integrity? Waldman et al. [12] and Grundy et al. [2] reported that leukocytes may take up CMV from infected endothelial cells but also deliver virus to the endothelium. As shown in the right part of figure 1, contact between a leukocyte and a CMV-infected endothelial cell may result in the uptake of the virus by the leukocyte, a process which may be important for dissemination. Experimentally, it is quite difficult to study CMV-induced endothelial-leukocyte interactions due to the limited infectivity of leukocytes in vitro. In addition, the activity of the virus seems to be dependent on the differentiation state of the host cell and thus standardization of the experiments is difficult to achieve. In the next chapter, we describe recent and ongoing work with the long-term CMV-infected tumor cell line UKF-NB-4AD169, which elicits highly reproducible CMV-induced interactions with human endothelial cells.

Transendothelial Migration Model with Permanently Infected Tumor Cells

The neuroblastoma cell line UKF-NB-4 was obtained from a bone marrow metastasis of a patient with Evans stage IV [13]. From this cell line, the permanently and productively human CMV (HCMV)-infected variant UKF-NB-4AD169 was established [4]. The latter cell line exhibits enhanced expression of the tumor marker Nmyc, whereas expression of the physiological enzyme tyrosine hydroxylase is found to be reduced [4, 14–16]. Moreover, when inoculated subcutaneously into BALB/c mice, UKF-NB-4AD169 but not UKF-NB-4 induced neuroblastoma tumors with metastases [16]. Therefore, in vitro studies were conducted to investigate whether UKFNB-4AD169 evolved mechanisms that allow it to better escape the immune system of the host and to interact with

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the endothelium causing transendothelial migration with the subsequent formation of metastases. It should be mentioned that the infected variant exhibited reduced neural cell adhesion molecule (NCAM) expression, which is important for homophilic binding between neuroblastoma cells and for the activity of natural killer and lymphokine activated killer (LAK) cells that normally recognize and kill tumor cells. Indeed, in cytotoxicity assays with LAK cells, the UKF-NB-4AD169 line exhibited relative resistance compared with the uninfected parental cell line [unpubl. data]. In order to study the ability of UKF-NB-4 and UKFNB-4AD169 to adhere to and transmigrate through the endothelial barrier, tumor cells were cocultured with monolayers of endothelial cells [17]. It was found that the infected variant adhered to a higher degree than the uninfected variant. In addition, relative to the number of adherent cells the number of transmigrating cells was also higher with UKF-NB-4AD169. Interestingly, the coculture with UKF-NB-4AD169 but not with UKF-NB-4 resulted in focal disruption of endothelial monolayer integrity (fig. 2C, E, respectively). Video analyses showed that with UKF-NB-4AD169, disruption occurred only at the focal contact sites. It is suspected that this phenomenon may reflect an overshooting mechanism involved in the extravasation of tumor cells during the formation of metastases. CMV might induce these mechanisms in order to escape the immune system, to drive the host cell into solid tissues for latent persistence or to reach epithelial cells in exocrine organs, for example, from where the virus can easily be transmitted to other hosts. To investigate the underlying cellular mechanisms that lead to these CMV-induced modifications of the interactions with endothelial cells, the uninfected and infected cell lines were compared in relation to their phenotypic expression of adhesion molecules. In brief, there was no significant expression of ICAM-1, E-selectin or VCAM-1 in either cell line. As already mentioned, NCAM expression is reduced in the infected variant.

Fig. 2. Endothelial cells alone (A, B), and with cocultured UKF-NB4 (C, D) or UKF-NB-4AD169 (E, F). E Focal disruption of the endo-

thelial monolayer occurred after coculture with UKF-NH-4AD169. F When tumor cells were pretreated with an antibody against VLA-5 no focal disruption occurred. The integration of UKF-NB-4 into the endothelial monolayer (C) was also prevented by this antibody (D). B Antibody treatment alone had no influence on monolayer integrity.

Scholz/Blaheta/Vogel/Doerr/Cinatl



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Fig. 3. A Focal disruption occurs when di-

rect cell-to-cell contact between tumor cells and endothelial cells is provided. B Conditioned medium from endothelial cell/UKFNB-4AD169 coculture was added to otherwise untreated endothelial cells. C Coculture was performed with Boyden chambers where cell-to-cell contact was prevented by filter inserts. No focal disruption was detectable with conditioned medium or when cocultured in Boyden chambers.

In functional binding studies to various purified and immobilized molecules, UKF-NB-4 but not UKF-NB4AD169 bound to P-selectin. This finding correlated with the reduced expression of P-selectin glycoprotein ligand-1 on UKF-NB-4AD169. However, these findings did not explain the modified interactions with endothelial cells as described above. From the video analyses it was evident that the cell-to-cell contact between UKF-NB-4AD169 and endothelial cells is crucial for the development of focal gaps. To confirm this, conditioned medium from UKFNB-4AD169 was added to endothelial monolayers. In other experiments, cells were cocultured in Boyden chambers with endothelial cells in the lower compartment and tumor cells in the upper compartment (fig. 3). No changes in monolayer integrity were observed in these experiments. In contrast, when isolated and purified membrane fragments of UKF-NB-4AD169 were added gap formation occurred. However, this membrane-associated effect was

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reversible and could only be observed during the early phase of the experiments. From these results we suggest that CMV induces membrane-associated modifications which are responsible for gap formation in endothelial layers. A series of experiments was conducted to further define the intercellular pathways. Since it is known that the adherens junction between endothelial cells is dependent on extracellular Ca2+ (homophilic binding of VEcadherin), the role of Ca2+ was studied. For example, EGTA and equimolar amounts of Mg2+ were added to the culture medium in order to chelate Ca2+ without causing toxicity in the endothelial cells. However, gap formation within the endothelial layer after coculture with UKFNB-4AD169 appeared to be unmodified, thus ruling out a predominant role of extracellular disruption of the Ca2+dependent cadherin binding.


Leukocyte/tumor cell

EGFr?

s

Tyrosine phos.

s

s

s

c-erbB 2

s

Fig. 4. Adhesion of a CMV-infected leukocyte or tumor cell to the endothelium with proposed intracellular changes leading to the disruption of the cadherin-cadherin binding. The receptor that mediates the CMV-induced effects has not yet been defined. The EGF receptor (EGFr) is known to be involved in tyrosine phosphorylation and direct catenin association. In addition, the phosphorylated c-erbB-2 oncogene product may also associate with catenins and thus contribute to disruption of the complex. On the right it is indicated that the cytoskeleton is disrupted upon leukocyte contact.

VE-cadherin

Catenins

Another possible cause of disruption in the binding between endothelial cells may be the activity of proteases. Indeed, when endothelial cells and UKF-NB-4AD169 were cocultured in the presence of the proteinase inhibitor phenantroline, gap formation was partially prevented. Therefore, ELISAs were carried out to detect metalloproteinases in the culture medium of either tumor cell line. No differences in metalloproteinase levels or tissue inhibitors of metalloproteinases could be detected. Nevertheless, it may be possible that membrane type metalloproteinases could play a role in endothelial gap formation. Ongoing studies are dealing with this possibility. Because proteinases seem to play a minor role in gap formation it has been suggested that intracellular pathways in the endothelial cell (e.g. the protein kinase C-dependent pathway) might be of relevance. However, preliminary experiments with protein kinase C inhibitors failed to inhibit gap formation. Interestingly, when UKF-NB-4AD169 was pretreated with blocking antibodies against the ß1 chain and the heterodimer ß1·5 (VLA-5) of the integrin family, the UKF-NB-4AD169-induced effects were blocked (fig. 2F). When the uninfected UKF-NB-4 line was pretreated with a ß1 integrin-stimulating antibody, gap formation could be induced according to the infected cell line. It may be speculated that CMV triggers the functional activity of VLA-5 in the tumor cell to improve transendothelial migration. Blocking studies with a specific blocking peptide against fibronectin, which is the natural ligand of

VLA-5, failed to inhibit gap formation after coculture, suggesting that other yet undefined VLA-5-related intercellular communication mechanisms may exist.



Proposed Mechanisms

In the following we discuss a putative mechanism that might contribute to the phenomenon of focal disruption of the endothelial integrity by CMV-infected cells. This mechanism is the subject of ongoing investigations in our institute. Figure 4 schematically depicts the proposed intracellular pathways that might be involved. We hypothesize that intracellular mechanisms may disturb the cadherin-mediated adhesion of the endothelial cell to its neighboring endothelial cell. Since combined Ca2+ depletion and Mg2+ supplementation did not lead to gap formation, as described above, we assume that the function of the Ca2+ binding motifs of the extracellular domain of the cadherin 5 molecule are not essentially impaired, at least in our cell coculture model. The cytoplasmic domain of cadherin 5 was shown to be linked to molecules belonging to the group of catenins. The intact association of cadherins with catenins is a prerequisite for the stability of the cytoskeleton, since the cadherin-catenin complex in turn is linked to the actin filament [19]. It has been further shown that the stability of the cadherincatenin complex is dependent on the tyrosine phosphorylation status of ß-catenin and plakoglobin [20]. Further-

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more, phosphorylation of catenins seems to be initiated by the epithelial growth factor (EGF). In addition, an association between the EGF receptor and ß-catenin has been documented [21]. Moreover, the c-erbB-2 oncogene product has been found to be aberrantly phosphorylated and coimmunoprecipitated with the ß-catenin/plakoglobin-cadherin complex [22]. Interestingly, the c-erbB-2 gene product core region is highly homologous with the EGF receptor [23]. In our model, in conditioned medium of UKF-NB-4AD169 or in cocultures of UKF-NB-4AD169 cells with endothelial cells without cell-to-cell contact (Boyden chambers), focal disruption of endothelial integ-

rity did not occur, indicating that secretion of soluble factors such as EGF may not play a dominant role in our model. The challenge in our ongoing experiments is to confirm the hypothesis that CMV-induced VLA-5 activity may induce instability of the endothelial adherens junctions and to define the underlying intracellular pathways.

Acknowledgments This work was generously supported by the foundations Hilfe für krebskranke Kinder, Frankfurt, e.V., and Frankfurter Stiftung für krebskranke Kinder.

References 1 Bolck F, Machnik G: Leber und Gallenwege; in Doerr W, Seifert G, Uehlinger E (eds): Spezielle pathologische Anatomie. Berlin, Springer, 1978, vol 10. 2 Grundy JE, Lawson KM, MacCormac LP, Fletcher JM, Yong KL: Cytomegalovirus-infected endothelial cells recruit neutrophils by the secretion of C-X-C chemokines and transmit virus by direct neutrophil-endothelial cell contact and during neutrophil transendothelial migration. J Infect Dis 1998;177:1465–1467. 3 Scholz M, Vogel J-U, Blaheta R, Cinatl J Jr: Cytomegalovirus, oxidative stress and inflammation as interdependent pathomechanisms: Need for novel therapeutic strategies?; in Scholz M, Rabenau HF, Doerr HW, Cinatl J Jr (eds): CMV-Related Immunopathology. Monogr Virol. Basel, Karger 1998, vol 21, pp 90– 105. 4 Cinatl J Jr, Vogel J-U, Cinatl J, Weber B, Rabenau H, Novak M, Kornhuber B, Doerr HW: Long-term productive human cytomegalovirus infection of a human neuroblastoma cell line. Int J Cancer 1996;65:90–96. 5 Nigro G, Schiavetti A, Booth JC, Clerici A, Dominici C, Krysztofiak A, Castello M: Cytomegalovirus-associated stage 4S neuroblastoma relapsed stage 4. Med Pediatr Oncol 1995; 24:200–203. 6 Zhu H, Cong J-P, Mamtora G, Gingeras T, Shenk T: Cellular gene expression altered by human cytomegalovirus: Global monitoring with oligonucleotide arrays. Proc Natl Acad Sci USA 1998;95:14470–14475. 7 Scholz M, Hamann A, Blaheta RA, Auth MK, Encke A, Markus BH: Cytomegalovirus- and interferon-related effects on human endothelial cells. Cytomegalovirus infection reduces upregulation of HLA class II antigen expression after treatment with interferon-gamma. Hum Immunol 1992;35:230–239.

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8 Sedmak DD, Knight DA, Vook NA, Waldman WJ: Divergent patterns of ELAM-1, ICAM-1, and VCAM-1 expression on cytomegalovirusinfected endothelial cells. Transplantation 1994;58:1379–1385. 9 Craigen JL, Grundy JE: Cytomegalovirus induced up-regulation of LFA-3 (CD58) and ICAM-1 (CD54) is a direct viral effect that is not prevented by ganciclovir or foscarnet treatment. Transplantation 1996;62:1102–1108. 10 Burns L, Pooley JC, Walsh DJ, Vercelloti GM, Weber ML, Kovacs A: Intercellular adhesion molecule-1 expression in endothelial cells is activated by cytomegalovirus immediate early proteins. Transplantation 1999;67:137–144. 11 Soderberg-Naucler C, Fish KN, Nelson JA: Interferon-gamma and tumor necrosis factor-alpha specifically induce formation of cytomegalovirus-permissive monocyte-derived macrophages that are refractory to the antiviral activity of these cytokines. J Clin Invest 1997; 100:3154–3163. 12 Waldman WJ, Knight DA, Huang EH, Sedmak DD: Biodirectional transmission of infectious cytomegalovirus between monocytes and vascular endothelial cells: An in vitro model. J Infect Dis 1995;171:263–272. 12 Cinatl J Jr, Hernaı´z-Driever P, Cinatl J, Vogel J-U, Scholz M, Doerr HW, Jakobi G, Kornhuber B: Persistent human cytomegalovirus infection in a human neuroblastoma cell line enhances the malignant behaviour. Klin Pädiatr 1996;208:256. 14 Cinatl J Jr, Cinatl J, Vogel J-U, Rabenau H, Kornhuber B, Doerr HW: Modulatory effect of human cytomegalovirus infection on malignant properties of cancer cells. Intervirology 1996;39:259–269.


15 Cinatl J Jr, Cinatl J, Vogel J-U, Kotchetkov R, Hernaı´z-Driever P, Kabickova H, Kornhuber B, Doerr HW: Persistent human cytomegalovirus infection induces drug resistance and alteration of programmed cell death in human neuroblastoma cells. Cancer Res 1998;58:367– 372. 16 Cinatl J, Gussetis ES, Cinatl J Jr, Ebener U, Mainke M, Schwabe D, Doerr HW, Kornhuber B, Gerein V: Differentiation arrest in neuroblastoma cell culture. J Cancer Res Clin Oncol 1990;116(suppl):9. 17 Scholz M, Blaheta RA, Hundemer M, Doerr HW, Cinatl J: Die Cytomegalievirus-Infektion als möglicher Progressionsfaktor bei der Neuroblastomerkrankung. Klin Pädiatr 1999;211: 310–313. 18 Gianelli G, Falk-Marzillier J, Schiraldi O, Stetler-Stevenson WG, Quaranta V: Induction of cell migration by matrix metalloprotease-2 cleavage of laminin-5. Science 1997;277:225– 228. 19 Takeichi M: Cadherin cell adhesion receptor as a morphogenetic regulator. Science 1991;251: 1451–1455. 20 Matsuyoshi N, Hamaguchi M, Taniguchi S, Nagafuchi A, Tsukita S, Takeichi M: Cadherinmediated cell-cell adhesion is perturbed by vsrc tyrosine phosphorylation in metastatic fibroblasts. J Cell Biol 1992;118:703–714. 21 Hoschuetzky A, Aberle H, Kemler R: ß-Catenin mediates the interaction of the cadherincatenin complex with epidermal growth factor receptor. J Cell Biol 1994;127:1375–1380. 22 Kanai Y, Ochiai A, Shibata T, Oyama T, Ushijima S, Akimoto S, Hirohashi S: c-erbB-2 gene product directly associates with ß-catenin and plakoglobin. Biochem Biophys Res Commun 1995;208:1067–1072. 23 Ochiai A, Akimoto S, Kanai Y, Shibata T, Oyama T, Hirohashi S: c-erbB-2 gene product associates with catenins in human cancer cells. Biochem Biophys Res Commun 1994;205:73–78.




Altered Expression of Extracellular Matrix in Human-Cytomegalovirus-Infected Cells and a Human Artery Organ Culture Model to Study Its Biological Relevance Peter Schaarschmidt 1, a Barbara Reinhardt 1, a Detlef Michel a Bianca Vaida a Klaus Mayr a Anke Lüske a Regine Baur b Jürgen Gschwend c Klaus Kleinschmidt c Margaritis Kountidis d Ulrich Wenderoth d Rainer Voisard b Thomas Mertens a a Abteilung d Klinik

Virologie und b Abteilung Innere Medizin II-Kardiologie, c Abteilung Urologie, Universitätsklinikum, Ulm, für Urologie, Kreiskrankenhaus Heidenheim, Deutschland

Key Words HCMV W Extracellular matrix W Cytopathic effect W Human renal artery organ culture

tem for the investigation of molecular as well as functional consequences of HCMV infection in a more physiological microenvironment. Copyright © 2000 S. Karger AG, Basel

Abstract The influence of human cytomegalovirus (HCMV) on the transcription of 11 selected, representative extracellular matrix genes was investigated in cell culture. Northern blot hybridization indicated the downregulation of all mRNAs investigated. Based on our results and the known repression of other extracellular matrix transcripts and the ß-actin transcription during HCMV infection, we suggest that one molecular mechanism contributing to the cytopathic effect may be the transcriptional downregulation of genes encoding proteins involved in cell structure and intercellular connection. To further study the biological relevance of this and other pathogenetic mechanisms, we established a human renal artery organ culture system and characterized this new infection model for HCMV. Our model is a new suitable sys-

1

Both authors contributed equally to this work.

ABC

© 2000 S. Karger AG, Basel 0300–5526/99/0426–0357$17.50/0



Introduction

Besides well-established causative relationships between human cytomegalovirus (HCMV) infection and human pathology, this virus has recently been suspected of contributing to or even inducing diseases which had not been connected with viral infections before, e.g. atherosclerosis and chronic vascular rejection following solid organ transplantation [1–4]. The pathomechanisms of such manifestations cannot be explained exclusively by cell-destructive viral effects, but more likely by complex modulations of cellular gene expression. Increasing numbers of cell-encoded genes altered by HCMV infection are being identified. In this report we present data showing downregulation of selected representative extracellular matrix genes in HCMV-infected cells. The influence of HCMV on extracellular matrix might contribute to the development of the cytopathic effect (CPE) on a molecular basis and consequently might also lead to the detach-

Prof. Dr. Thomas Mertens Abteilung Virologie, Universitätsklinikum Ulm Albert-Einstein-Allee 11, D–89081 Ulm (Germany) Tel. +49 731 502 3341, Fax +49 731 502 3337 E-Mail [email protected]

Table 1. Influence of HCMV infection on the expression of cellular

genes Grouped cellular molecules or functions

Examples of regulated genes

Actin/tubulin/myosin Cell cycle Cell-surface and adhesion proteins

Actin-binding protein ↓ Gas-1 ↓ ICAM-1, LFA-3 ↑ CD10, CD13, MHC-1 ↓ TFPI-2 ↑ DAF ↑ IL-6, IL-8, RANTES ↑ ↑ ornithine decarboxylase, ↑ ·-type DNA polymerases ↑

Coagulation Complement Chemokines/receptors DNA metabolism enzymes Enzymes Extracellular matrix and cell adhesion GTP binding proteins Heat-shock and stress-inducible proteins Interferon-inducible genes Kinase/phosphate Laminins Ligands and receptors Prostaglandin synthesis Protein degradation Protooncogenes Splicing factors Transcription factors Translation factors

Fibronectin, integrins ↓ RanBP1 ↑ hsp 70 ↑ staf 50 ↑ RIP protein kinase ↑ ß2-laminin ↓ PDGF-receptor-· ↓ Cox2 ↑ E2-EPF ↑ BCL7B, p53 ↑ SFRS7 ↑ NF-ÎB, Sp1 Translation initiation factor ↑

ment of endothelial cells from the vessel wall. During active HCMV infection, circulating endothelial cells harboring large amounts of infectious virus have been found [5]. Since an animal model for HCMV infection does not exist and since studies employing cell monolayers do not sufficiently reflect the in vivo situation, there is a strong need to develop better model systems. In vitro studies in different cell cultures only allow the analysis of defined aspects in a very complex process of virus-host interactions. As a consequence, the relevance of HCMV infection for human pathology is difficult to study in vitro. Complex microenvironments, such as arterial vessels, which include cell-cell interactions cannot be examined adequately in cell culture systems. In addition, the results obtained by in vitro studies of cultured cells, e.g. human umbilical vein endothelial cells (HUVEC), should be interpreted with great care since there are significant differences between HUVEC and endothelial cells from vessels likely to be affected by atherosclerotic changes. In order to avoid these problems, a series of in vivo studies concerning the pathogenesis of graft vasculopathy using a rat model have been

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performed in recent years to evaluate potential influences of rat CMV on the development of atherosclerotic lesions [4]. The fact that HCMV is a very species-specific virus is a substantial drawback in all animal models, HCMV differs from animal CMVs in many biological properties [6], such as progression to clinical disease, distinct functions of socalled homologous genes [Wagner et al., unpubl. data] and, finally, in their susceptibility to antiviral drugs [7]. Moreover, differences in host cell reactions were observed. Whereas CMV infection in rat endothelial cells directly induced HLA class II expression on infected cells [8], no such changes could be observed following HCMV infection [9]. On the contrary, infected human endothelial cells were refractory to HLA class II expression following interferon-Á stimulation [10]. Moreover, the cytoarchitecture of the arterial wall differs significantly between humans and small mammals such as rats [11]. In order to avoid the problems mentioned above, we used a new human organ culture system [45]. This model should allow us to study functional virus-cell interactions, cell-cell interactions under the influence of HCMV infection relevant for atherosclerosis as well as virus spread.

Modulation of Gene Expression in HCMV-Infected Cells

Probably due to a very long coevolution HCMV interacts on multiple sites with its host cell during productive infection, and these interactions are both close and very complex. The virus has the potential to alter cellular gene expression through multiple pathways [12–23]. Increasing numbers of cell-encoded genes that are altered by HCMV infection are being identified (table 1) [24–40]. Zhu et al. [39, 40] demonstrated that differential-display RT-PCR and cDNA chip technology provide important new tools for the investigation of HCMV-induced modulation of host cell gene expression, but confirmation of these results by Northern blot and protein analysis is extremely important. The complex virus-host cell interaction may result in the modulation of different cellular functions. By influencing transcriptional control or cell signaling mechanisms, for instance, HCMV might have implications on cell differentiation and proliferation, and especially atherogenesis. By inducing the production of soluble cytokines or receptors, HCMV might even influence noninfected bystander cells. We are especially interested in elucidating the mechanisms of HCMV-induced CPE and pathogenetic processes in atherosclerosis.

Schaarschmidt et al.

Influences of Human Cytomegalovirus on Extracellular Matrix

Extracellular matrix connects different cells of the body. It compensates for the stress of movement, maintains shape, and can be considered as a composite of insoluble fibers, adhesion molecules and soluble polymers. The main fibers are collagen and elastin; laminin and fibronectin are important adhesion molecules and the soluble molecules include proteoglycans and glycoproteins [41] (fig. 1a). Apart from mechanical properties, extracellular matrices play key roles in signal transduction [42, 43]. The influences of HCMV on extracellular matrices are potentially connected with the development of CPE (fig. 1b). To extensively monitor the influences of HCMV on extracellular matrix gene expression, we performed systematic Northern blot analysis of 11 selected mRNAs encoding representative proteins of different extracellular matrix components [Schaarschmidt et al., unpubl. data] (fig. 1a). All investigated genes were transcriptionally downregulated in HCMV-infected fibroblasts (table 2). However, levels, time points and mechanisms of downregulation obviously differed, since some effects could even be observed using inactivated virus at very early time points.

Fig. 1. a Composition of extracellular matrix. b Effects of HCMV

infection on extracellular matrix.

Table 2. Downregulation of genes encoding

proteins of different components of extracellular matrix in HCMV-infected cells

Gene

GenBank Component accession numbers

Size of downregulated mRNAs, kb

·1-Collagen type 1 Elastin

M32798 A32707; X15603

fibers

4.7 2.2

Fibronectin (control) ß-Galactoside-binding lectin

P02751; X02761 X14829

adhesion molecules

7.7 0.5

Aggrecan Decorin

J05062; X80278 L01131

proteoglycans

Versican

P13611; X15998

7.1 1.9 1.6 8.2

Fibrillin Fibulin

S17064; X63556 P23142; P23143 P23144; X82494 P24821; X56160 P07996; X14787

glycoproteins

9.9 4.1 2.1 7.3 5.7

Tenascin Thrombospondin 1

Extracellular Matrix Altered by HCMV and an Artery Organ Culture Model


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Fig. 2. Example for the influence of HCMV on extracellular matrix genes. Downregulation of fibronectin transcription and translation and alteration in structure. The sizes of mRNA and protein are indicated. GAPDH transcription was used as control. m = Mock-infected; i = HCMV-infected (AD 169 strain).

Because of its known downregulation in HCMVinfected cells, fibronectin was used as a control. Suppression of the fibronectin mRNA and protein is shown in figure 2. Additionally, we were able to show changes in fibronectin structure in HCMV-infected fibroblasts by immunofluorescence (fig. 2).

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Taken together, the transcriptional downregulation of genes encoding proteins of different extracellular matrix components in cells infected with HCMV could be shown, documenting a substantial effort of the virus to reduce the extracellular matrix.


Fig. 3. Electron-microscopic examination of a renal artery segment 35 days after infection with the endotheliotropic HCMV isolate TB40E. N = Nucleus; Cy = cytoplasm; DM = plasma membrane; R = ribosomes; V = virus. a Uninfected cell. b Infected cell. c Corresponds to the box shown in b (higher magnification).

Since these investigations have been performed almost exclusively in cultured fibroblasts, it is important to further elucidate whether other cell types might be influenced in the same way. Concerning the pathogenesis of HCMV in vivo, fibroblasts are not as relevant as, e.g., endothelial cells, myocytes or other cell types. The virusinduced reduction of extracellular matrix might be of special importance in infected endothelial cells, potentially resulting in detachment from the vessels wall. Circulating endothelial cells allow productive infection and may serve as a vehicle for virus dissemination [44]. Besides being involved in virus spread, HCMV-infected endothelial cells might also be involved in atherogenesis by triggering vascular pathology.

A Human Renal Artery Organ Culture System for Studying the Pathogenetic Role of HCMV Infection

Segments of renal arteries were removed during routine nephrectomies. Maintained in culture, these vessel segments retain their proliferative capabilities for at least


28 days, as shown by BrdU incorporation. Neointimal thickening following mechanical injury could be demonstrated for 56 days, indicating the presence of a still viable system [45] (fig. 3). HCMV infection of renal artery segments was performed cell free with different clinical isolates and the laboratory strain AD 169. The characteristics of the progressing infection were evaluated by immunohistochemistry as well as by detection of infectious virus in the culture supernatant using a plaque assay. In this study, we demonstrated the feasibility of a human arterial vessel organ culture model for studying the influences of HCMV infection on the arterial vessel wall in an in-situ-like situation. Following infection with cell free virus, a long-lasting permissive HCMV infection could be induced in our renal artery model [Reinhardt et al., unpubl. data]. Productive infection in culture was observed after inoculation with clinical HCMV isolates as well as with the laboratory strain AD 169. HCMV early antigen could be detected in the intimal layer as early as 3 days following infection with the endotheliotropic strain TB40E (kindly provided by Dr. C. Sinzger, Tübingen). As the infection progressed, the virus penetrated into the vessel media. Infected foci could be observed within 2–5


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weeks after inoculation depending on the HCMV isolate applied. HCMV-antigen-positive cells in the lamina media were identified as myocytes by double staining. Additionally, HCMV-positive cells also appeared in adventitious tissue. The detection of increasing quantities of infectious virus in the culture supernatants proves the presence of a productive HCMV infection in this organ culture model. Peak levels of plaque-forming units (PFU) per milliliter supernatant were observed around day 30 after inoculation (fig. 4). After infection with different HCMV strains, distinct differences in the kinetics of infection and in cell tropism could be observed. However, no significant differences could be detected between renal artery segments obtained from different donors when they were infected with the same HCMV isolate (fig. 4). No expression of viral antigens was observed in mockinfected cultures, independent of the HCMV serostatus of the organ donor, indicating that the culture conditions used do not induce endogenous reactivation within the cultured artery segment.

Conclusion

We believe that applying techniques such as electron microscopy and in situ PCR methodology, this model will serve to elucidate various molecular and functional consequences of HCMV infection in a more physiological microenvironment (fig. 5). A number of urgent questions concerning vascular pathology possibly induced by HCMV could be addressed by using this model. Specifically, the biological importance of downregulation of extracellular matrix will be evaluated.

Acknowledgments Fig. 4. Detection of infectious virus in the culture supernatant. Renal

artery segments from different donors were infected in parallel with three different HCMV isolates. Results from segments obtained from the same renal artery donor are displayed by the same symbol. Infection was performed using 5 ! 105 PFU for AD 169 and 1 ! 105 PFU for TB40E and AN365. Virus yields are shown (PFU/ml).

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The authors thank I. Bennett and Th. Schmid for editorial assistance. The work was supported in part by grants from the Deutsche Forschungsgemeinschaft (SFB 451 A2), Bundesministerium für Bildung und Forschung (01 KS 9605/2 A8) and the European Community (PL 960471).


Fig. 5. Fictitious section through a renal artery segment. The organ

culture model should allow to study mechanisms and influences of HCMV infection relevant for atherogenesis in a more physiological microenvironment. Numbers indicate potential mechanisms to be elucidated: 1 = HCMV-induced detachment of endothelial cells from the vessel wall; 2 = adherence of latently infected monocytes to preexisting endothelial lesions: a potential mechanism of virus entry? 3 = HCMV-induced stimulation of smooth muscle cell (SMC) pro-

liferation and/or migration, e.g. by induction of platelet-derived growth factor ligand and/or platelet-derived growth factor receptor expression/activation; 4 = virus-induced production of various growth factors such as vascular endothelial growth factor; 5 = relevance of increased expression of adhesion molecules by HCMVinfected cells. EC = Endothelial cell; Mø = monocyte; MDM = monocyte-derived macrophages.

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Human Cytomegalovirus Infection of Immature Dendritic Cells and Macrophages Gerhard Jahn a Stephan Stenglein a Susanne Riegler a Hermann Einsele b Christian Sinzger a a Department

of Medical Virology, and b Department of Medicine, University of Tübingen, Tübingen, Germany

Key Words Macrophages W Immature dendritic cells W Human cytomegalovirus W Pathogenesis W Immunosuppression W Antigen-presenting cells

Abstract A central aspect of human cytomegalovirus (HCMV) pathogenesis is the interaction of the virus with different antigen-presenting cell (APC) types of the host. In principle, a number of various cell types have the potential of antigen presentation when MHC II expression is induced by appropriate stimuli. The most potent antigen presenters are monocytes/macrophages and dendritic cells (DCs), therefore called professional APCs. Interestingly, these cells seem to be targets of productive HCMV infection. The susceptibility of the monocyte/macrophage system has been analyzed intensively during the past decade. Investigation of the role of DCs during HCMV infection, however, has begun only recently. Copyright © 2000 S. Karger AG, Basel

ABC

© 2000 S. Karger AG, Basel 0300–5526/99/0426–0365$17.50/0



Introduction

Macrophages in organ tissue are terminally differentiated immune effector cells derived from monocyte precursors in the peripheral blood. While these precursor monocytes do not support the complete replication cycle of human cytomegalovirus (HCMV), the process of differentiation into macrophages during transmigration from the blood stream into tissues obviously induces a state of cellular permissiveness for productive HCMV infection in vitro [Fish et al., 1996; Söderberg-Nauclèr et al., 1998]. Thus, in vivo infection has previously been demonstrated in tissues [Sinzger et al., 1996]. The first publication about skin dendritic cells (DCs) appeared in 1868 and was entitled: ‘Über die Nerven der menschlichen Haut’ [Langerhans, 1868]. The interpretation of DCs as ‘skin nerve cells’ was followed by long-term speculation on their biological function. 106 years later, Steinman and Cohn discovered the lymphoid DCs in the mouse and pioneered the area of tissue-derived DCs as potent stimulators of immune responses [Steinman et al., 1974; Inaba et al., 1984]. Later on, a number of publications stated that DCs were also found in nonlymphoid tissues of rodents and humans [Hart and Fabre, 1981; Daar et al., 1983]. The DCs were also considered to play a major role in transplantation medicine [McKenzie et al.,

Prof. Dr. Gerhard Jahn Universitätsklinikum Tübingen, Medizinische Virologie Calwer Str. 7/6 D–72076 Tübingen (Germany) Tel. +49 7071 29 84921, Fax +49 7071 29 5790, E-Mail [email protected]

Fig. 1. Role and function of DCs.

Fig. 2. Some features of DCs. (+) Indicates low positivity, and + indicates high positivity. Modified from Banchereau and Steinman [1998].

1984]. Figure 1 illustrates some functional aspects of immature and mature DCs. For decades, a trial and error scenario dominated the characterization of markers for DCs, because of difficulties distinguishing DCs from monocytes/macrophages and problems in the purification steps of DCs. Figure 2 shows some markers of immature versus mature DCs. It should be mentioned that even experts in the field sometimes still disagree about the expression of specific markers on DCs (e.g. CD83, fig. 2). The heterogeneity of DCs – myeloic versus lymphoid, native versus immature, DC1 versus DC2, precursors of dendritic cell – does play a role in this aspect. It is now accepted that DCs are a leukocyte subpopulation of professional antigen-presenting cells (APCs) with an outstanding capacity for initiating primary and secondary T lymphocyte response [Steinman, 1991; Hart and Calder, 1994; Banchereau and Steinman, 1998; Hart, 1997; Reid, 1998]. The DCs that are present in the skin, mucosa, blood and tissue are the cell population most likely to have the initial contact with viruses during infection. For a number of viruses, such as HIV-1, HTLV-1, measles, influenza, it could be demonstrated that DCs are not only involved in triggering antiviral immune responses, but also in carriage or even propagation of viral infections [Schnorr et al., 1997; Bender et al., 1997, 1998; Bhardwaj et al., 1994; Patterson et al., 1998; Knight et al., 1993, 1997; Knight and Patterson, 1997; Klagge and Schneider-Schaulies,

1999]. In certain viral infections, the DCs therefore appear to have a dual, even paradoxical role – one as hosts (targets) for viruses and another one as stimulators and inducers of immunologic cellular and perhaps even humoral defense. The viral infection of DCs can lead to impairment of antigen presentation and consequently to immune dysfunction and progression of the viral infection. On the other hand, infected DCs can provide the viral agent with a safe haven as well as lead to immune activation. As a consequence, the infected DCs may therefore contribute to the pathogenesis of viral infections depending on the mode of interaction of virus with DCs in terms of hosting the pathogen in a silent way or in an acute replication phase. Investigations into HCMV cell tropism during acute infection in vivo have recently made remarkable findings: ubiquitously distributed infected cells such as epithelial cells, fibroblasts, endothelial cells, smooth muscle cells, hepatocytes, trophoblasts, macrophages and circulating cytomegalic endothelial cells in the blood as well as leukocytes carrying subviral material in the peripheral blood and in infected tissues. The findings of HCMV-infected cells in tissue with immediate-early through late viral antigen expression demonstrated by immunohistochemistry using double-staining techniques are in agreement with the data on HCMV infection in primary cell cultures [Plachter et al., 1996; Sinzger and Jahn, 1996; Sinzger et al., 1993, 1995, 1996, 1999; Halwachs-Baumann et al.,

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1998]. Relevant questions still open are the mechanisms of viral spread throughout the organism including the travel of HCMV from one compartment to another. In addition, the issue of impairment of the function of infected cells or organs by the virus is still open to discussion. Finally, the question of how HCMV handles or interacts with the host’s immune response leading to lifelong persistence is unanswered. To our knowledge, the first report dealing with DCs and HCMV was published in 1993 [Becker, 1993]. In this communication, the role of DCs in the pathogenesis of HCMV infection in immunocompetent and immunocompromised hosts was based on pure speculation. The next cascade of reports concerning HCMV infection and DCs came from different groups, particularly from the Northwest Pacific region [Söderberg-Nauclér et al., 1997, 1998]. These publications claimed that DCs were not permissive for HCMV infection in vitro, in clear contrast to the growth of HCMV in monocytes/macrophages. However, it was demonstrated by another group that HCMV remains latent in CD33+ cells as a precursor of DCs and myeloid cells [Hahn et al., 1998; Mendelson et al., 1996]. These authors presented evidence that CD33+ progenitors of DCs and monocytes/macrophages in healthy HCMV-seropositive individuals can function as important reservoirs of latent HCMV. Since during maturation immature DCs can migrate from various sites, such as the skin, liver, lung, mucosa to lymph nodes, it is important to screen for HCMV in the local lymphatic organs. Up to now, no investigations on the early phase of HCMV infection in vivo have been published and therefore no conclusive interpretation exists concerning the in situ situation in the lymph nodes [Younes et al., 1991].

free isolates. We used fibroblast-adapted strains and recent clinical isolates/endothelial cell propagated strains. Figure 3B shows a typical cytopathic effect of an HCMV endothelial-tropic strain in infected immature DCs 7 days after infection. These series clearly demonstrated that there is a significant difference between fibroblastadapted HCMV strains and endothelial-cell-propagated strains independent of the MOI [Riegler et al., 1999, 2000]. Fibroblast-adapted HCMV strains do not replicate in immature DCs whereas recent clinical isolates/endotheliotropic HCMV strains run through the complete replicative cycle. Our data clearly demonstrate that immature DCs are susceptible to HCMV infection in vitro, and the infected DCs show a cytopathic effect with enlarged nuclear inclusions typical of late-stage HCMV infection (fig. 3B) [Riegler et al., 1999, 2000]. In addition, it could be demonstrated by time kinetics of viral antigen expression using immediate-early, early and true-late specific monoclonal antibodies that the infected DCs are permissively infected by recent clinical HCMV/endotheliotropic isolates [Riegler et al., 1999]. The kinetics of viral antigen expression was slightly delayed in immature DCs in comparison to infected fibroblasts. Cell lysis of infected immature DCs appeared at a late stage in contrast to mock-infected DCs. In addition to these experiments, permissive HCMV infection could be demonstrated by single-step growth curves and infectious center assays [Riegler et al., 1999, 2000]. Taken together, our results clearly demonstrate that recent clinical isolates/ endothelial cell propagated HCMV isolates are efficient in infecting immature DCs in vitro, leading to the production of mature virions. Since there are no valid data on the HCMV infection of DCs in vivo, the consequences are highly speculative and possibilities are schematically illustrated in figure 4.

HCMV Infection of Immature DCs

Immature DCs from peripheral blood adherent mononuclear cells of healthy HCMV-seronegative individuals were generated as recently described [Brossart and Bevan, 1997; Brossart et al., 1997]. The adherent cells were cultured in RPMI-10% FCS, supplemented with 1,000 U/ml IL-4 and 100 ng/ml GM-CSF. The cytokines were repeatedly replenished on days 2, 4 and 6. The resulting homogeneous nonadherent cell population with typical DC morphology is shown in figure 3A. The DC cultures consisted of more than 90% pure populations displaying the phenotype of immature DCs (fig. 2). In experiments performed in our laboratory, these immature DCs were infected with different HCMV-cell-

Human Cytomegalovirus Infection of Immature Dendritic Cells and Macrophages

HCMV Infection of Macrophages

Whereas blood-derived monocytes cannot be productively infected by HCMV, these cells become permissive for the complete replication cycle after differentiation into macrophages [Taylor-Wiedeman et al., 1991; Ibanez et al.,1991; Lathey and Spector, 1991], and recent isolates seem to be more efficient than strains repeatedly passaged on fibroblasts [Minton et al., 1994; Sinzger et al., unpubl]. In monocyte-derived macrophages differentiated by overnight coculture with concanavalin-A-stimulated lymphocytes, we found cytopathic effects characteristic of


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Fig. 3. Cytopathic effect of HCMV-infected DCs. A Mock-infected DCs. B HCMV-infected DCs. Arrowheads indi-

cate late-stage infected cells with nuclear inclusions.

Fig. 4. Hypothetical role of HCMV-infected DCs. The speculative scheme illustrates possibilities and consequences of HCMV-infected DCs based on partial in vitro data.

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late-stage infection. In these cultures, numerous nuclear inclusions (denoted by arrowheads in fig. 5B) could be observed as a typical indicator of late-stage HCMV infection. The significance of these results for the in vivo situation was emphasized by our finding that tissue macrophages in various organs enable the expression of HCMV immediate-early, early and late proteins [Sinzger et al., 1996; Mazeron, et al., 1992]. Figure 6A shows the detection of HCMV early antigen p52 [Plachter et al., 1992] in infected lung tissue, whereas figure 6B shows the staining of HCMV late antigen pp150 [Jahn et al., 1987, 1990; Scholl et al., 1988; Plachter et al., 1992 a, b] in infected colon tissue.


Fig. 5. Cytopathic effect of HCMV in monocyte-derived macrophages. A Macrophages 6 days after mock infection. B Macrophages 6 days postinfection with HCMV strain TB40/E at an MOI of 5. Numerous nuclear inclusions are

visible.

Discussion

It is becoming clear that monocyte-derived macrophages and DCs have many roles in the immune system, particularly in viral infection. In addition to a role as mobile sentinels that bring antigens to T cells expressing costimulators for the induction of immunity, there is growing evidence that DCs do have a dual, even paradoxical role in some viral infections: one as hosts for the virus and another as inducers of immune stimulation or immunosuppression [Schnorr et al., 1997 a, b; Bender et al., 1997/1998; Bhardwaj et al., 1994; Patterson et al., 1998; Knight et al., 1993, 1997; Knight and Patterson, 1997;


Klagge and Schneider-Schaulies, 1999]. Previous work has demonstrated that macrophages sequester HCMV proteins in cytoplasmic vacuoles (which have not been unequivocally identified but are not lysosomes) [Fish et al., 1995]. Monocytes/macrophages may therefore help HCMV to avoid its destruction by the immune system and serve as a vehicle that carries HCMV into various organ tissues. Studies performed by Jay Nelson and his team have clearly demonstrated delayed kinetics of HCMV replication in this cell type [Fish et al., 1995]. For the first time, our group has obtained experimental data showing that HCMV can permissively infect immature DCs [Riegler et al., 1999, 2000]. We have first evi-


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Fig. 6. Immunohistochemical analysis of HCMV antigen expression in tissue macrophages. Detection of HCMV antigens by the immunoperoxidase technique yielded brown staining. Detection of macrophage marker CD68 by the immunoalkaline phosphatase technique yielded red staining. A Detection of HCMV early antigen p52. B Detection of HCMV late antigen pp150.

dence that HCMV infection of DCs in vitro rapidly leads to downregulation of class I molecules and to delayed downregulation of class II molecules. The markers for immature DCs do not change during the course of HCMV infection [Einsele, unpubl.]. Up to now, there are no data on HCMV infection of DCs in vivo, in contrast to the data on HCMV-infected macrophages [Sinzger et al., 1996] and the growing literature on HIV infection of DCs and macrophages [Grouard and Clark, 1997; Hladik et al., 1999]. Detection of HCMV-infected DCs in vivo will be technically demanding, since HCMV primary infection rarely leads to typical clinical symptoms, and since DCs, the smallest hematopoietic subpopulation, are very few. Our data on the in vitro infection of DCs suggest that they might play an important role in HCMV infection

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and in immune responses most likely leading to immune dysfunction (immunosuppression, impairment of DCs or of APCs; fig. 4). The lack of an appropriate model for HCMV latency/silent chronic infection has complicated the efforts to understand persistence versus reactivation of HCMV in healthy carriers. However, experiments on the permissiveness of specific types of blood cells including immature DCs for HCMV infection in vitro as well as the staining of viral antigens in peripheral blood cells that harbor viral DNA have directed our attention to the important role that cells of the myeloid lineage most likely play in the maintenance of HCMV in healthy carriers and in viral gene expression during the initial step of acute infection. Although this is only a hypothesis, it can be assumed that the DCs and/or macrophages play a domi-


nant role in these mechanisms of HCMV pathogenesis. Using mature DCs pulsed with viral peptide antigens, the authors of this paper have also recently shown generation of HCMV-specific T cells from HCMV seropositive and seronegative healthy donors for bone marrow transplantation [unpubl. results]. These peptide antigens were defined by epitope prediction and ELISPOT analysis of the frequency of HCMV peptide-reactive T cells in HCMVseropositive healthy donors. Thus, mature DCs can be used for the generation of HCMV-specific T cell lines and clones. Mature DCs might be ideal APCs for the genera-

tion of virus-specific immune effector cells to be used for adoptive T cell therapy in patients at high risk of developing HCMV disease, e.g. recipients of an allogeneic stem cell graft [Einsele et al., 1995, 1999, in press; Krause et al., 1997] or AIDS patients.

Acknowledgment This work was supported by the Deutsche Forschungsgemeinschaft (SFB 510, project B3). We thank H. G. Rammensee for stimulating discussion and constructive critique.

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Plachter B, Wieczorek L, Scholl B-C, Ziegelmaier R, Jahn G: Detection of cytomegalovirus antibodies by an ELISA using recombinant polypeptides of the large phosphorylated tegument protein pp150. J Clin Microbiol 1992b;30: 201–206 Reid CDL: The biology and clinical applications of dendritic cells. Transfus Med 1998;8:77–86. Riegler S, Sinzger C, Jahn G: Efficient productive infection of dendritic cells by endotheliotropic strains of the human cytomegalovirus (abstract 4.003). 24th Int Herpesvirus Workshop, Cambridge, Mass., 1999. Riegler S, Hebart H, Einsele H, Brossart P, Jahn G, Sinzger C: Monocyte-derived dendritic cells are permissive to the complete replicative cycle of human cytomegalovirus. J Gen Virol 2000; 81:393–394. Schnorr JJ, Seufert M, Schlender J, Borst J, Johnston ICD, ter Meulen V, Schneider-Schaulies S: Cell cycle arrest rather than apoptosis is associated with measles virus contact-mediated immunosuppression in vitro. J Gen Virol 1997a; 78:3217–3226. Schnorr JJ, Xantahkos S, Keikavoussi P, Kämpgen E, ter Meulen V, Schneider-Schaulies S: Induction of maturation of human blood dendritic cell precursors by measles virus is associated with immunosuppression. Proc Natl Acad Sci USA 1997b;94:5326–5331. Scholl B-C, v. Hintzenstern J, Borisch B, Traupe B, Bröker M, Jahn G: Prokaryotic expression of immunogenic polypeptides of the large phosphoprotein (pp150) of human cytomegalovirus. J Gen Virol 1988;69:1195–1204. Sinzger C, Bissinger AL, Viebahn R, Oettle H, Radke C, Schmidt CA, Jahn G: Hepatocytes are permissive for human cytomegalovirus infection in human liver cell culture and in vivo. J Infect Dis 1999;180:976–986.


Sinzger C, Grefte A, Plachter B, Gouw ASH, The TH, Jahn G: Fibroblasts, epithelial cells, endothelial cells and smooth muscle cells are major targets of human cytomegalovirus infection in lung and gastrointestinal tissues. J Gen Virol 1995;76:741–750. Sinzger C, Jahn G: Human cytomegalovirus cell tropism and pathogenesis. Intervirology 1996, 39:302–319. Sinzger C, Müntefering H, Löning T, Stöss H, Plachter B, Jahn G: Cell types infected in human cytomegalovirus placentitis identified by immunohistochemical double staining. Virchows Arch A Pathol Anat 1993;423:249–256. Sinzger C, Plachter B, Grefte A, The TH, Jahn G: Tissue macrophages are infected by human cytomegalovirus in vivo. J Infect Dis 1996;173: 240–245. Söderberg-Nauclér C, Fish KN, Nelson JA: Growth of human cytomegalovirus in primary macrophages. Methods 1998;16:126–138. Söderberg-Nauclér C, Fish KN, Nelson JA: Reactivation of latent human cytomegalovirus by allogeneic stimulation of blood cells from healthy donors. Cell 1997;91:119–126. Steinman RM: The dendritic cell system and its role in immunogenicity. Annu Rev Immunol 1991;9:271. Steinman RM, Lustig DS, Cohn ZA: Identification of a novel cell type in peripheral lymphoid organs of mice. III. Functional properties in vivo. J Exp Med 1974;139:1431. Taylor-Wiedeman J, Sissons JGP, Borysiewicz LK, Sinclair JH: Monocytes are a major site of persistence of human cytomegalovirus in peripheral blood mononuclear cells. J Gen Virol 1991;72:2059–2064. Younes M, Podesta A, Helie M, Buckley P: Infection of T but not B lymphocytes by cytomegalovirus in lymph node. Am J Surg Pathol 1991; 15:75–80.


Part II. Diagnostics and Antiviral Therapy Diagnostics


Diagnostic Value of Nucleic-Acid-SequenceBased Amplification for the Detection of Cytomegalovirus Infection in Renal and Liver Transplant Recipients Valère J. Goossens a Marinus J. Blok a Maarten H.L. Christiaans b Johannes P. van Hooff b Peter Sillekens e K. Höckerstedt d Irmeli Lautenschlager c Jaap M. Middeldorp e Cathrien A. Bruggeman a a Department

of Medical Microbiology and b Department of Internal Medicine, University Hospital Maastricht, Maastricht, The Netherlands; c Department of Virology and d Transplantation and Liver Surgery Unit, Department of Surgery, University Hospital Helsinki, Helsinki, Finland, and e Organon Teknika BV, Boxtel, The Netherlands

Key Words CMV W Screening W mRNA W Immediate early gene products W pp67 W PCR W Antigenemia

Abstract To evaluate the diagnostic value of nucleic-acid-sequence-based amplification (NASBA) for the detection of cytomegalovirus (CMV) infection in transplant recipients, we compared immediate early 1 (IE1) and late pp67 mRNA detection by NASBA with the antigenemia assay, PCR and viral culture in 72 renal transplant (RTx) recipients and with antigenemia and serology in 25 liver transplant (LTx) recipients. Antigenemia, viral culture and pp67 NASBA were almost equivalent for the detection of CMV in RTx recipients. In LTx recipients, antigenemia detected more positive samples and more positive recipients compared to pp67 NASBA. In RTx recipients, PCR detected more positive samples and positive recipients compared to pp67 NASBA, antigenemia and viral culture. Also the first day of detection was slightly earlier for PCR. However, IE1 NASBA was the most sensitive test The first two authors contributed equally to this work.

ABC

© 2000 S. Karger AG, Basel 0300–5526/99/0426–0373$17.50/0



and detected 96% of all positive samples and positive transplant recipients. In addition, IE1 NASBA preceded PCR and all other positive results. This makes IE1 NASBA a very attractive screening test for the early detection of CMV infection. Copyright © 2000 S. Karger AG, Basel

Introduction

Infection with human CMV is common and usually subclinical. However, serious CMV infection is possible in immunocompromised individuals, e.g. transplant recipients, HIV-infected patients and pregnant women [1]. After kidney, liver or other solid-organ transplantation, organ rejection occurs more frequently and a higher mortality rate is seen for CMV-positive recipients compared to CMV-negative recipients. The higher mortality rate as well as organ rejection can both be prevented by early antiviral treatment [2]. However, effective treatment requires an early diagnosis. In previous studies [3–5], we evaluated the diagnostic value of nucleic acid amplification tests for the early detection of CMV IE1 and late viral pp67 mRNA and viral DNA, using nucleic-acid-se-

Valère J. Goossens Department of Medical Microbiology, University Hospital Maastricht PO Box 5800 NL–6202 AZ Maastricht (The Netherlands) Tel. +31 43 38 76644, Fax +31 43 38 76643, E-Mail [email protected]

quence-based amplification (NASBA) and PCR, respectively. We compared these results with antigenemia, viral culture and/or serology. This comparison was done for either renal or liver transplant recipients. In this paper we report an overview of the results obtained for 97 transplant recipients of both groups and discuss the value of the methods used.

Patients and Methods Patients and Specimens Three groups of transplant recipients were studied. The first group consisted of 42 renal transplant recipients (RTx1). A total of 489 whole-blood samples were prospectively collected after kidney transplantations carried out at the Department of Internal Medicine, University Hospital Maastricht, between January 1995 and July 1996. The recipients could be grouped according to the CMV serostatus of the recipient (R) and donor (D) before transplantation. This resulted in the following distribution of recipients over the different serogroups: 13 D+/R+; 8 D+/R–; 13 D–/R+; 8 D–/R–. Heparinized whole-blood and serum specimens were collected weekly from inpatients and at every subsequent outpatient visit. Specimens were screened prospectively for the presence of CMV by viral culture and pp65 antigenemia. In addition, 1 ml of heparinized blood was added to 9 ml of NASBA lysis buffer and stored at –70° until analysis. In a second group of 30 renal transplant recipients (RTx2), a total of 291 EDTA blood specimens were prospectively collected after kidney transplantations carried out at the Department of Internal Medicine, University Hospital Maastricht, between July 1997 and November 1997. All these samples were collected twice weekly during the first 2 months after transplantation. The D/R serostatus distribution in RTx2 was 6 D+/R+; 9 D+/R–; 8 D–/R+; 7 D–/R–. The third group consisted of 25 liver transplant recipients (LTx). A total of 291 blood samples were prospectively collected after liver transplantations carried out at the Helsinki University Hospital between December 1996 and November 1997. The D/R serostatus distribution in LTx recipients was 12 D+/R+; 4 D+/R–; 6 D–/R+; 2 D–/R–; 1 D?/R+. pp65 Antigenemia Assay The pp65 antigen (ppUL83), a major early protein, was detected immunocytochemically in leukocytes isolated from whole-blood samples using antibodies directed against pp65. This assay was performed essentially as described by van der Bij et al. [6]. Briefly, leukocytes were isolated from whole blood and centrifuged onto a glass slide within 4 h after collection. The cells were subsequently fixed, and pp65 was demonstrated by a monoclonal antibody and immunoperoxidase staining. Finally, the slides were examined microscopically. The results were expressed as the number of positive cells per 50,000 leukocytes. Viral Culture For the detection of infectious CMV in blood, both conventional cell culture with detection of cytopathic effect (CPE) and shell vial culture with detection of early antigen fluorescent foci (DEAFF) were performed as described previously [7]. Briefly, leukocytes were purified from whole blood using dextran. Approximately 200,000 leuko-

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cytes were screened in duplicate for the presence of CMV by inoculation of human embryo fibroblast monolayers. Cell cultures were observed twice a week during a period of 6 weeks for the appearance of a typical CPE on replicating CMV. Also DEAFF was performed in duplicate after 2 days of culture, using the monoclonal antibody E13 (Biosoft, Paris, France) to the CMV immediate early (IE) antigen. For analysis, DEAFF and CPE results were combined to give one outcome for the viral culture. The viral culture assays were routinely performed for the RTx recipients but not for the LTx recipients. Serology Sera of RTx1 and LTx recipients were tested retrospectively to obtain more information about viral activity, in addition to antigenemia and other assays. For the qualitative detection of IgM antibodies against CMV in serum, the Im®X CMV assay (Abbott Laboratories, Abbott Park, Ill., USA) was used, while IgG was semiquantitatively measured by AXSym® (Abbott). Both tests are based on the microparticle enzyme immunoassay (MEIA) technology. NASBA for pp67 mRNA and IE1 mRNA Isolation, amplification and detection of pp67 mRNA, expressed from the UL65 open reading frame of human CMV, was done following the instructions of the manufacturer (NucliSens® CMV pp67 Organon Teknika, Boxtel, The Netherlands). The assay includes an internal system control (ISC) which serves as a positive control for isolation, amplification and detection; technical details are described elsewhere [3]. Isolation, amplification and detection of IE1 mRNA transcripts were done essentially as described for pp67 mRNA. Except that for the IE1 amplification two primers directed to exon 4 of the major IE gene were used. Also for IE1 NASBA, a specific ISC was developed [4]. Polymerase Chain Reaction Detection of CMV DNA was done with Viral Quant®, a quantitative PCR detection kit (BioSource, Etten-Leur, The Netherlands), following the instructions of the manufacturer. However, after adding 20,000 copies of the internal Calibration Standard (ICS) to each sample, nucleic acid isolation was performed essentially as described by Boom et al. [8]. The purified nucleic acids were resuspended in 200 Ìl of sterile water of which 5 Ìl was used for PCR. After an initial denaturation step at 95° for 2.5 min, the following temperature profile was used for 32 cycles: denaturing during 25 s at 94°, annealing during 25 s at 60° and extension during 1 min at 72°. Due to our special interest for detection of CMV as early as possible, we used this PCR and microplate detection system only qualitatively at the lowest recommended dilution of 1:20 for detection of the amplified DNA and a dilution of 1:80 for the specific detection of the ICS copies. Definition of CMV infection To allow a comparison of the results of the three groups of transplant recipients to be made, CMV infection was defined by at least one positive result for one or more of the following items: antigenemia, viral culture, IE1 NASBA, p67 NASBA. Mathematics Diagnostic values, including sensitivities, specificities, positive predictive values (PPV) and negative predictive values (NPV) of IE1 NASBA, pp67 NASBA, viral culture and PCR were calculated with antigenemia as reference test.

Goossens et al.

Table 1. RTx1, RTx2 and LTx recipients,

and samples positive for antigenemia, viral culture, pp67 NASBA, PCR and IE1 NASBA

Test

RTx1

RTx2

LTx

Total

Viral culture Positive samples Positive recipients, %

36 36 (15/42)

26 33 (101/30)

not tested

35 (25/72)

Antigenemia Positive samples Positive recipients, %

32 21 (9/42)

33 30 (9/30)

47 68 (17/25)

112 36 (35/97)

pp67 NASBA Positive samples Positive recipients, %

36 31 (13/42)

33 40 (12/30)

32 52 (13/25)

101 39 (38/97)

PCR Positive samples Positive recipients, %

not tested

49 43 (13/30)

not tested

43 (13/30)

IE1-NASBA Positive samples Positive recipients, %

164 62 (26/42)

66 67 (20/30)

132 88 (22/25)

362 70 (68/97)

Total Positive samples Positive recipients, %

167 64 (27/42)

74 73 (22/30)

136 88 (22/25)

377 73 (712/97)

Numbers of positive recipients are shown in parentheses. In 1 RTx recipient, at the moment of sampling of the first positive viral culture, IE1 mRNA was still undetectable. However, at the time the positive result of the conventional culture became available, IE1 NASBA was already positive in later samples. 2 Only 3 RTx recipients, 2 with only 1 sample positive for viral culture, and 1 recipient with only 1 sample positive for PCR, were IE1 NASBA negative. 1

Table 2. Maximal antigenemia levels (expressed as number of positive cells per 50,000 leukocytes) detected in 18 RTx and 17 LTx recipients with primary or secondary infection

D+R– recipients D+R+ recipients D–R+ recipients

RTx1

RTx2

LTx

67, 14, 8, 1, 0.33, 0.33 15, 3, 0.33 no positive results

259, 168, 78, 68, 10 58, 7, 4 1

100, 85, 50, 50 150, 100, 50, 40, 30, 30, 20, 20, 10 600, 150, 100, 8

Results

Antigenemia was positive in 21, 30 and 68% of 42 RTx1, 30 RTx2 and 25 LTx recipients, respectively (table 1). Antigenemia was also more frequently positive in primary (65%, 15/23 recipients) than in secondary infections (43%, 20/47 recipients). For RTx recipients, the maximal detected levels for antigenemia were higher in 11 primary infections than in 7 secondary infections. In 17 LTx recipients, the maximal detected levels for antigenemia were high in both primary and secondary CMV

NASBA for CMV Detection in Transplant Recipients

infections (table 2). In 18 RTx recipients, primary and secondary CMV infections were detected by antigenemia at a mean of 40 days (range 22–69) after transplantation (table 3). Viral culture was positive in 35% of RTx recipients (table 1) and was more frequently positive in primary (72%, 13/18 recipients) than in secondary infections (40%, 12/ 30 recipients). Compared to antigenemia, the sensitivity, specificity, PPV and NPV of viral culture were respectively 89, 86, 69 and 96% (table 4). In 16 RTx recipients positive for both viral culture and antigenemia, primary and


375

Table 3. The onset of CMV infection

(mean and range in parentheses), expressed as first day of positivity after transplantation, in RTx recipients as detected by viral culture (day of sampling), antigenemia, pp67 NASBA, PCR and IE1 NASBA

Test

RTx1 (n = 26)

RTx2 (n = 22)

Antigenemia+ (n = 18)1

Viral culture Antigenemia pp67 NASBA PCR IE1 NASBA

43 (19–69) 41 (28–69) 42 (21–75) not tested 33 (11–69)

35 (26–54) 39 (22–54) 43 (26–55) 35 (15–48) 30 (1–54)2

38 (23–69) 40 (22–69) 42 (26–75) 35 (22–48) 33 (15–69)

1

Selection of antigenemia-positive recipients in both RTx groups. IE1: 1 recipient was positive at day 1, another at day 4 and 18 recipients became positive between day 15 and 54 after transplantation.

2

Table 4. Diagnostic values of viral culture, pp67 NASBA, PCR and

IE1 NASBA compared to antigenemia in RTx1 (n = 42), RTx2 (n = 30) and LTx (n = 25) recipients Test and group Sensitivity, % Specificity, % PPV, %

NPV, %

Viral culture RTx1 RTx2 Mean

89 89 89

82 90 86

57 80 69

96 95 96

pp67 NASBA RTx1 RTx2 LTx Mean

100 89 71 87

88 81 88 86

69 67 92 76

100 94 58 84

PCR RTx2

100

81

69

100

IE1 NASBA RTx1 RTx2 LTx Mean

100 100 100 100

55 48 38 47

38 45 77 53

100 100 100 100

secondary CMV infections were detected by viral culture (day of sampling) at a mean of 38 days (range 23–69) after transplantation (table 3). In the RTx group, combination of conventional cell culture and shell vial culture resulted in culturing CMV in 62 samples (table 1). Of these 62 positive samples, 58 (94%) were detected by CPE and only 25 (40%) were detected by DEAFF. pp67 NASBA was positive in 31, 40 and 52% of RTx1, RTx2 and LTx recipients (table 1), respectively, and was more frequently positive in primary infections (70%, 16/ 23 recipients) than in secondary infections (47%, 22/47 recipients). Compared to antigenemia, the sensitivity,

376


specificity, PPV and NPV of pp67 NASBA were 87, 86, 76 and 84%, respectively (table 4). In 17 RTx recipients positive for both pp67 NASBA and antigenemia, the primary and secondary CMV infection was detected by pp67 NASBA at a mean of 42 days (range 26–75) after transplantation (table 3). For renal transplant recipients, the results obtained with the pp67 NASBA are similar to those from viral culture (day of sampling) and antigenemia. First, the number of samples and recipients that were positive with these 3 different tests were almost similar (table 1). In about 80% of these samples, a positive result for pp67 NASBA, antigenemia or viral culture was confirmed by at least one of the other two tests. Secondly, the distribution of the first day of positivity is similar for these 3 tests (table 3). However, a positive result of the viral culture will always take an additional delay of 2 days (for DAEFF) to 6 weeks (for CPE) due to the required incubation time. Of 112 antigenemia positive samples, 44 were negative for pp67 NASBA (table 5), predominantly in samples with low antigenemia levels. However, in 6 LTx recipients, all treated with ganciclovir, also some pp67 NASBA-negative samples with high antigenemia results (up to 100) were seen. PCR results were only available for the RTx2 group. Of the 30 RTx2 recipients, 13 (43%) were positive for PCR (table 1). Compared to antigenemia, the sensitivity, specificity, PPV and NPV of PCR were respectively 100, 81, 69 and 100% (table 4). In 9 RTx2 recipients positive for both PCR and antigenemia, primary and secondary CMV infections were detected by PCR at a mean of 35 days (range 22–48) after transplantation (table 3). The number of RTx2 recipients positive for PCR (13) was not significantly different compared to antigenemia (9), viral culture (10) and pp67 NASBA (12). However, in these recipients, PCR was positive in 49 samples compared to 33, 26 and 33 samples for antigenemia, viral culture and

Goossens et al.

Fig. 1. Typical examples of results obtained by the pp65 antigenemia assay, viral culture, serology, IE1 and pp67 NASBA in 3 RTx recipients (patients I, II and III) and 1 LTx recipient (patient IV). In RTx recipients, the indicated rejection therapy consisted of three doses of methylprednisolone. To indicate very low antigenemia levels, the antigenemia results are expressed as the number of pp65-positive cells per 150,000 leukocytes. Results from the ImX CMV assay were expressed as index values. Sera with IgM index values 60.500 were

considered positive. The levels of anti-CMV IgG were expressed as the number of antibody units per milliliter (AU/ml). Sera with IgG levels 615 AU/ml were considered positive. The viral culture results are presented by the day the blood sample was taken from the recipient (DEAFF and CPE results were combined to give a single outcome for viral culture). In patient III and IV, the black line indicates the period of antiviral therapy with ganciclovir.

pp67 NASBA, respectively (table 1). PCR detected even more positive samples (49) than the 44 positive samples obtained with viral culture, antigenemia and/or pp67 NASBA together. Fifteen RTx2 recipients were positive for viral culture, antigenemia and/or pp67 NASBA. Of these 15 recipients, 11 were positive for PCR including all (6) high-risk transplantations (D+R–) and all (9) antigenemia-positive recipients. IE1 NASBA was positive in 62, 67 and 88% of RTx1, RTx2 and LTx recipients, respectively (table 1), and was detected in almost all primary (96%, 22/23 recipients) and secondary (98%, 46/47 recipients) CMV infections. Compared to antigenemia, the sensitivity, specificity, PPV and NPV of IE1 NASBA were 100, 47, 53 and 100%, respectively (table 4). In 18 RTx recipients positive for both IE1 NASBA and antigenemia, CMV infection was detected by IE1 NASBA at a mean of 33 days (range 15– 69) after transplantation (table 3). In 94% (46/49) of pos-

itive RTx and in all 22 positive LTx recipients, IE1 NASBA was the first test to become positive. IE1 NASBA could detect CMV several days to several weeks before all other tests, and especially in individual RTx recipients, this diagnostic advantage was up to 28 days compared to viral culture (day of sampling), antigenemia and PCR and up to 35 days compared to pp67 NASBA. Of all samples positive for at least one test, 96% (362/377) were detected by IE1 NASBA (table 1). In the three groups of RTx1, RTx2 and LTx recipients, there were samples that were only positive for IE1 NASBA. In part, these positive IE1 NASBA results were confirmed later on by other tests, including pp67 NASBA, antigenemia, viral culture and PCR. In other recipients, positive IE1 NASBA results could not be confirmed by antigenemia, viral culture or pp67 NASBA, but were confirmed by a serological response with the appearance of anti-CMV IgM antibodies and/or a significant increase in anti-CMV



377

Table 5. Distribution of 134 samples positive for antigenemia (112) and/or pp67 NASBA (90)

Antigenemia negative pp67 NASBA-positive samples

RTx1

RTx2

LTx

Total

pp67+ pp67–

pp67+ pp67–

pp67+ pp67–

pp67+ pp67–

6

9

22

7

Antigenemia result between 0.1 and 3.0 pp67 NASBA-positive samples 11 pp67 NASBA-negative samples

14


0


0

Antigenemia result higher than 30.0 pp67 NASBA-positive samples pp67 NASBA-negative samples

0

Total

25

Discussion

Several methods are available for the early detection of CMV infection. The oldest one is the conventional cell culture with CPE detection. A disadvantage of this test is that the results are obtained only 1–6 weeks after starting the culture. Using shell vial culture with DEAFF, results can be obtained within 2 days, but the number of positive samples is low. This means that for the early detection of CMV in RTx recipients, detection by CPE takes too long for clinical use and detection by DEAFF is too insensitive. Especially for blood cultures, such a difference between DEAFF and CPE has already been described [1]. The sensitivity of shell vial culture can probably be improved by increasing the number of polymorphonuclear leukocytes inoculated [9]. It is also important to reduce the transport time of specimens prior to culture to avoid false-negative culture results [10].


0 3

8

7

19

3

8

13 6

17

4

0 33

20

15

1 8

14

19 3

2 3

1

IgG antibodies (see patient I in fig. 1). In patient II in figure 1, the first and the last positive IE1 NASBA results were not confirmed by other tests. Also after treatment with ganciclovir, for several weeks to months (see patients III and IV in fig. 1), samples were positive only for IE1 NASBA and negative for antigenemia and viral culture.

378

8

5 22

2 32

24

2 90

44

Due to its diagnostic accuracy, rapidity, quantitative nature and technical simplicity, the antigenemia assay is one of the cornerstone methods for diagnosis and management of active CMV infection in immunocompromised patients [11] and attempts are made at its standardization [12]. Besides this, the antigenemia assay was the only method used in both the RTx1 and RTx2 groups and in the LTx group. For both reasons, we used the antigenemia assay as the ‘golden standard’ to compare with viral culture, pp67 NASBA, PCR and IE1 NASBA. The described results of antigenemia-positive samples and recipients and a different distribution among RTx recipients with primary or secondary infection (table 5) are in agreement with others [13]. In our RTx and LTx recipients, a good correlation was also found between the number of pp65positive leukocytes and the clinical status of the recipient before and after treatment with ganciclovir (see for example patients III and IV in figure 1). Detection of CMV late viral pp67 mRNA using the NASBA methodology has been possible for several years already [14]. For RTx recipients, pp67 NASBA is at least equivalent to antigenemia and viral culture. However, in LTx recipients more than in RTx recipients, antigenemiapositive but pp67 NASBA-negative samples were seen (table 5). Two main situations are responsible for the presence of such pp67-NASBA-negative, antigenemia-positive samples. First, this can happen very early in the infec-

Goossens et al.

tion. However, this does not need to be a problem in clinical practice. This was demonstrated in 4 RTx recipients in our study with one sample being negative for pp67 NASBA, but in each recipient already the next sample, taken 3–4 days later, was positive for pp67 NASBA. Secondly, after antiviral therapy, there is a difference in the disappearance of the antigenemia and the pp67 NASBA signals. Upon initiation of antiviral therapy, there is a block in the viral replication cycle and in the production of pp67 mRNA, a true late (Á) marker. Due to the instability of mRNA, pp67 NASBA will soon become negative while the stabler pp65 protein is still present in polymorphonuclear leukocytes. In addition, the pp65 protein is transcribed from UL83, a gene with ß/Á characteristics [15], and cannot be considered a true late marker. Therefore, low levels of pp65 antigen cannot be used to monitor therapeutic efficacy. In general, for RTx recipients, pp67 NASBA is equivalent to antigenemia and viral culture for the early detection of CMV infection. In LTx recipients with primary CMV infection, all 4 recipients were positive for both antigenemia and pp67 NASBA. In LTx recipients with secondary CMV infection, pp67 NASBA was less frequently positive (47%, 9/19 recipients) compared to antigenemia (68%, 13/19 recipients). This difference between pp67 NASBA and antigenemia in RTx and LTx recipients is most probably due to more frequent and early use of antiviral products in LTx recipients. It also reflects the great difference in clinical status and immunosuppression between RTx and LTx recipients. Especially LTx recipients are heavily immunosuppressed and are much more easily treated for CMV, because they are more at risk of developing a life-threatening infection. In fact, antiviral therapy with ganciclovir was given to 72% (18/ 25) of the LTx recipients (including all 15 recipients with symptomatic CMV infection and 3 recipients with ganciclovir prophylaxis during antirejection treatment) compared to only 14% (6/42) of the RTx1 recipients. Within the RTx2 group, PCR detected all the (high) risk recipients. These findings are in agreement with other results obtained with the same quantitative CMV DNA detection method [16]. However, due to 27 samples and 9 recipients positive for other tests but negative for PCR, the analytical sensitivity of PCR was not as expected. This is probably also the reason why the manufacturer modified the method afterwards by lowering the number of added ICS copies and increased the number of cycles. With this modified method, samples containing at least 3 copies per amplification (600 copies/ml) were detectable [16]. However, in general, the diagnostic value of monitoring CMV DNA remains in dispute, considering the

life-long persistence of CMV in the circulatory compartment. IE1 mRNA NASBA is a very sensitive diagnostic method for the early detection of CMV activity after renal or liver transplantation. Of 22 LTx recipients and 45 RTx recipients positive for at least one test, only 3 were missed by IE1 NASBA. However, in these 3 RTx recipients, with only one sample positive for only one test, contamination of viral culture or PCR could not be excluded. In general, in RTx and LTx recipients, IE1 NASBA allows the quickest detection of an ongoing CMV infection, bringing an early alerting signal to the clinician. This permits an intensified diagnostic monitoring and possibly early antiviral treatment to be performed. Recently, with the quantification of IE1 and pp67 mRNA levels, differences between pp67 NASBA, antigenemia and IE1 NASBA became clearer [17]. In lung transplant recipients, IE1 NASBA became detectable days before pp67 NASBA and antigenemia. This was the result of a much higher level of IE1 mRNA. When antigenemia-positive cells or pp67 mRNA levels become detectable (x100 pp67 mRNA molecules per milliliter), IE1 mRNA levels already exceed 10,000 mRNA molecules per milliliter. In addition, relapsing infection and ineffective therapy were paralleled by persisting IE1 mRNA levels (15,000/ml), which disappeared only upon complete resolution of the infection. However, high IE1 mRNA levels were also observed in some patients whose CMV infection remained subclinical. The difference in the detectable level of 100 IE1 mRNA molecules per milliliter for NASBA [17] and in the detectable level of 600 copies of DNA molecules per milliliter for PCR [16] also contributes to higher numbers of samples and recipients being positive for IE1 NASBA compared to PCR. Although IE1 NASBA has advantages compared to other tests, the significance of a positive IE1 NASBA is not always clear. In all three transplant recipient groups, a positive signal with IE1 NASBA was not always an indication of an active or symptomatic infection with CMV (e.g. patient I in fig. 1). The question can be raised whether IE1 NASBA is perhaps too sensitive for practical use and detects too many asymptomatic or subclinical infections. Indeed, some IE1 NASBA results could not be confirmed by other tests. On the other hand, no IE1 mRNA is detectable in healthy virus carriers [Middeldorp et al., in preparation]. In addition, there is increasing evidence that the presence of IE and/or E gene products is a key factor in some aspects of the pathogenesis of CMV infection. CMV infections cause major infectious complications after organ transplantation, and a variety of clinical manifesta-



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tions, such as fever, leukopenia, thrombopenia, encephalitis, retinitis, pneumoniae, hepatitis and glomerulopathy, have been described [18]. In kidney allografts, vascular changes, glomerulosclerosis and tubular atrophy are associated with chronic rejection, but the most characteristic long-term finding is interstitial fibrosis [19]. Some of these complications and findings can not be explained by productive viral replication with synthesis of new viral DNA and late gene products as structural viral proteins and resulting in ‘virus plaques’ in tissues with consequent loss of tissue and tissue function. More important is the activity of the immediate early and/or early gene products inducing many proinflammatory activities along with cytokines, chemokines and upregulation of adhesion molecules [20]. Also immune escape phenomena are already present in the immediate early/early phase [21]. In addition, sequence homology and immunologic cross-reactivity of immediate early antigens and HLA-DR antigens have been demonstrated [22]. At this moment, the clinical significance of the detection of IE1 mRNA with the IE1 NASBA is not fully understood, especially in patients negative for antigenemia/viral culture with or without antiviral prophylaxis or treatment. However, in the near future with the increasing prophylactic and therapeutic use of DNA-polymerase-blocking antiviral products, IE1 and other immediate early/early gene products can ‘build up’ behind the block of viral DNA synthesis [20]. Thereby, this antiviral block could extend and accentuate rather than prevent inflammatory responses, possibly leading to delayed or more chronic complications in the posttransplantation period.

Conclusion

Viral culture, pp65 antigenemia and late pp67 mRNA detection by NASBA are almost equivalent in the detection of CMV in RTx recipients, Not only the first day of detection, but also the number of positive samples and positive recipients are similar for viral culture, pp65 antigenemia and pp67 NASBA. PCR detected CMV in more samples and in more recipients than did these three tests. Also the first day of detection was slightly earlier for PCR. However, PCR missed some recipients that were positive in the other tests. In RTx and LTx recipients, IE1 NASBA was the most sensitive test and detected 96% of the positive samples and positive recipients. In most recipients, IE1 NASBA precedes PCR and all other positive results. This makes IE1 NASBA a very attractive screening test for the detection of CMV infection and allows an intensified diagnostic investigation to be performed with possibly early antiviral treatment. With the high sensitivity and NPV of IE1 NASBA, a negative result rules out a CMV infection. However, the clinical significance of some positive IE1 NASBA results is not yet clear and needs further investigation. Therefore, at the moment, a positive IE1 NASBA should be used in combination with assays that are more specific for the detection of symptomatic CMV infection. As representative for immediate early gene and early gene products, IE1 NASBA may also provide more insight into the pathogenesis of CMV-related immunopathology.

References 1 Britt WJ, Alford CA: Cytomegalovirus; in Fields BN, Knipe DM, Howley PM (eds): Fields Virology. New York, Lippincott-Raven, 1996, pp 2493–2523. 2 Patel R, Snydman DR, Rubin RH, Ho M, Pescovitz M, Martin M, Paya CV: Cytomegalovirus prophylaxis in solid organ transplant recipients. Transplantation 1996;61:1279–1289. 3 Blok MJ, Goossens VJ, Vanherle SJV, Top B, Tacken N, Middeldorp JM, Christiaans MH, Van Hooff J, Bruggeman CA: Diagnostic value of monitoring human cytomegalovirus late pp67 mRNA expression in renal-allograft recipients by nucleic acid sequence-based amplification. J Clin Microbiol 1998;36:1341–1346.

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4 Blok MJ, Christiaans MHL, Goossens VJ, Van Hooff JP, Sillekens P, Middeldorp JM, Bruggeman CA: Early detection of human cytomegalovirus infection after kidney transplantation by nucleic acid sequence-based amplification. Transplantation 1999;67:1274–1277. 5 Goossens VJ, Blok MJ, Christiaans MHL, Sillekens P, Middeldorp JM, Bruggeman CA: Early detection of cytomegalovirus in renal transplant recipients: Comparison of PCR, NASBA, pp65-antigenemia and viral culture. Transplant Proc, in press. 6 Van der Bij W, Schirm J, Torensma R, Van Son WJ, Tegzess AM, The TH: Comparison between viremia and antigenemia for detection of cytomegalovirus in blood. J Clin Microbiol 1988;26:2531–2535.


7 Kraat YJ, Christiaans MHL, Nieman FHM, van den Berg-Loonen PM, van Hooff JP, Bruggeman CA: Risk factors for cytomegalovirus infection and disease in renal transplant recipients: HLA-DR7 and triple therapy. Transplant Int 1994;7:362–367. 8 Boom R, Sol CJA, Salimans MMM, Jansen CL, Wertheim-van Dillen PME, van der Noordaa J: Rapid and simple method for purification of nucleic acids. J Clin Microbiol 1990;28:495– 503. 9 Reina J, Saurina J, Fernandez-Baca V, Blanco I, Munar M: An increase in the number of polymorphonuclear leukocytes inoculated on shellvial culture increases the sensitivity of this assay in the detection of cytomegalovirus in the blood of immunocompromised patients. Diagn Microbiol Infect Dis 1998;31:425–428.

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10 Roberts TC, Buller RS, Gaudreault-Keener M, Sternhell KE, Garlock K, Singer GG, Brennan DC, Storch GA: Effects of storage temperature and time on qualitative and quantitative detection of cytomegalovirus in blood specimens by shell vial culture and PCR. J Clin Microbiol 1997;35:2224–2228. 11 The TH, Harmsen MC, Van der Bij W, Van den Berg AP, Van Son WJ: Relationship between monitoring the viral load in blood, human cytomegalovirus pathophysiology and management strategies of patients after transplantation. Monogr Virol. Basel. Karger, 1998, vol 21, pp 270–279. 12 The TH, Van den Berg AP, Harmsen MC, Van der Bij W, Van Son WJ: The cytomegalovirus antigenemia assay: A plea for standardization. Scand J Infect Dis 1995;suppl 99:25–29. 13 Sharma AK, Taylor JD, Tong W, Brown MW, Sells RA, Bakran A: Utility of the pp65 direct antigenemia test in the diagnosis of cytomegalovirus (CMV) in renal transplant recipients. Transplant Proc 1997;29:799.


14 Gerna G, Baldanti F, Middeldorp JM, Furione M, Zavattoni M, Lilleri D, Revello MG: Clinical significance of expression of human cytomegalovirus pp67 late transcript in heart, lung, and bone marrow transplant recipients as determined by nucleic acid sequence-based amplification. J Clin Microbiol 1999;37:902–911. 15 Grefte JM, Van der Gun BT, Schmolke S, Van der Giessen M, Van Son WJ, Plachter B, Jahn G, The TH: The lower matrix protein pp65 is the principal viral antigen present in peripheral blood leukocytes during an active cytomegalovirus infection. J Gen Virol 1992;73:2923– 2932. 16 Reagen KJ, Cabradilla C, Shuman B, Stollar N, Laudemann J, Bai X, Hosler G, Scheuermann RH: Analytical performance of a quantitative CMV DNA detection method. Monogr Virol. Basel, Karger, 1998, vol 21, pp 252–261. 17 Middeldorp J: Direct quantification of human cytomegalovirus (HCMV) immediate early and late mRNA levels in the blood of HCMVinfected individuals using competitive NASBA (abstract G0-21) 7th Int Cytomegalovirus Workshop, Brighton, April 1999. J Clin Virol 1999;12:103.

18 Rubin RH: Impact of cytomegalovirus infection on organ transplant recipients. Rev Infect Dis 1990;12(suppl 7):S754–S766. 19 Lautenschlager I: Role of cytomegalovirus infection in the process of organ allograft rejection. Monogr Virol. Basel, Karger, 1998, vol 21, pp 142–157. 20 Grundy JE: Current antiviral therapy fails to prevent the pro-inflammatory effects of cytomegalovirus infection, whilst rendering infected cells relatively resistant to immune attack. Monogr Virol. Basel, Karger, 1998, vol 21, pp 67–89. 21 Miller DM, Sedmak DD: Cytomegalovirus persistence: escape from cell-mediated immunosurveillance. Monogr Virol. Basel, Karger, 1998, vol 21, pp 1–11. 22 Fujinami RS, Nelson JA, Walker L, Oldstone MBA: Sequence homology and immunologic crossreactivity of human cytomegalovirus with HLA-DR chain: A means for graft rejection and immunosuppression. J Virol 1988;62:100– 105.


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Towards Standardization of the Human Cytomegalovirus Antigenemia Assay Erik A.M. Verschuuren Martin C. Harmsen Pieter C. Limburg Wim van der Bij Arie P. van den Berg Adriana M. Kas-Deelen Boelo Meedendorp Willem J. van Son T. Hauw The The Biomed 2 Study Group Department of Clinical Immunology, University Hospital Groningen, The Netherlands

Key Words Cytomegalovirus W Antigenemia W Standardization

Abstract The Human Cytomegalovirus antigenemia (HCMV-Agemia) test has been accepted worldwide as a clinical tool in the diagnosis and management of HCMV-associated syndromes in immunocompromised patients. The many modifications proposed since the first description by our laboratory make standardisation of the HCMV-Agemia assay necessary to enable multicenter clinical trials. We

For the Biomed 2 Study Group: Prof. Dr. M. Mach, Institut für Klinische und Molekulare Virologie, Universität Erlangen, Erlangen, Germany; Dr. C.A. Bruggeman, Dept. of Medical Microbiology, University of Limburg Maastricht, the Netherlands; Prof. Dr. H. Einsele, Med. Universitätsklinik II, Tübingen, Germany; Prof. Dr. G. Gerna, Servicio di Virologia, IRCCS Policlinico San Matteo, Pavia, Italy; Prof. Dr. J.E. Grundy, Dept. of Clinical Immunology Royal Free Hospital School of Medicine, London, United Kingdom, Dr. M.P. Landini, Policlinico S. Orsola, Instituto di Microbiologia, Bologna, Italy; Prof. Dr. Th. Mertens, Institut für Microbiologie, Abt. Virologie, Universität Ulm, Germany; Prof. Dr. G. Palu, Medical School, University of Padova, Italy, Prof. Dr. T.H. The, Dept. of Clinical Immunology, University Hospital Groningen, The Netherlands, and Dr. A. Volpi, Dept. of Public Health, School of Medicine, University of Rome ‘Tor Vergata’, Italy.

ABC

© 2000 S. Karger AG, Basel 0300–5526/99/0426–0382$17.50/0



report the initial work for standardization of the HCMVAgemia assay. A standard protocol is proposed, the optimal distribution conditions are investigated and the results of the shipment of positive and negative test slides as well as of two sets of coded internal standard slides are discussed. The main conclusions are that standard slides can be distributed at room temperature and that the results of participating laboratories with the coded internal standard slides were strikingly similar in spite of differences in HCMV-Agemia protocols used by participating laboratories. Copyright © 2000 S. Karger AG, Basel

Introduction

Since the Human Cytomegalovirus (HCMV) antigenemia assay (HCMV-Agemia assay) was first described by our laboratory more than 10 years ago [1], numerous studies have confirmed its clinical relevance for the diagnosis of active human HCMV infection. Its high diagnostic accuracy, rapidity and technical simplicity have made the HCMV-Agemia assay one of the cornerstone methods for the diagnosis and management of active HCMV infection in immune compromised patients [2–6]. However, a considerable number of modifications concerning every step in the protocol have been introduced

Erik A.M. Verschuuren Department of Internal Medicine University Hospital Groningen, PO Box 30.001 NL–9700 RB Groningen (The Netherlands) Tel. +31 50 3612945, Fax +31 50 3613151, E-Mail [email protected]

by different laboratories, e.g. the isolation method of polymorphonuclear cells (PMNs), the cell numbers used for preparing slides, the use of cytocentrifuged cell preparations versus methanol spreading, the use of different fixation methods, of different monoclonal antibodies, of chromogenic versus fluorescent detection of positive cells and the reading of the results by only counting the positive cells per slide versus the scoring of positive cells per 50,000 PMNs [7–12]. Introduction of these modifications was intended to improve the performance of the original assay, but this may lead to increased interlaboratory variation as well. These variations in methodology make comparison of published results difficult and form an obstacle to (future) multicenter trials of antigenemia-directed intervention studies in immune compromised patients with a high risk for fatal HCMV disease. Consequently, a concerted action was initiated to set up a standardization program for the HCMV-Agemia test in several collaborating laboratories and clinical centers in the European Community [13]. The primary goal was to come to similar test performances in participating laboratories. The first step was to make recommendations for a standard protocol and distribute it to the participating centers for modifying their current assay or adopting the standard protocol. Since participating laboratories were reluctant to abandon their local modifications, an alternative goal was set. This became the comparison of the HCMV-Agemia test performances in the participating laboratories on identical control slides. Only then could the decision be taken whether a standardized HCMVAgemia assay is justified and required for multicenter studies. The second step was the evaluation of the performance of the reference laboratory to see whether the reproducibility of the control slides was adequate. The third step was the investigation of optimal distribution conditions. Effects of temperature and fixation method were analyzed and subsequently positive/negative control slides were distributed to be tested in the participating laboratories. The last step discussed in this article is the evaluation of two sets of coded slides to evaluate their usefulness as an internal standard. One set of internal standard slides was made of pp65-positive baculovirus-transfected insect cells. A second set comprised pp65positive polymorphonuclear cells (PMNs) prepared by cocultivation of normal donor PMNs with productively infected endothelial cells [14, 15]. In the near future, the interlaboratory variation will be evaluated by distributing control slides made of patient material (external standard) with unknown numbers of positive cells.

Towards Standardization of the Human CMV Antigenemia Assay

Materials and Methods Isolation of Leukocytes Two milliliters of EDTA-treated blood was mixed with 1 ml PBS and 1 ml of a 5% dextran (MW 250,000) in 0.9% NaCl and allowed to settle at 1 g sedimentation force at 37° at a 60° angle for 10 min. The leukocyte supernatant was collected and centrifuged at 300 g for 2 min. The erythrocytes were lysed by resuspending the cell pellet in 2 ml cold erythrocyte lysing buffer (NH4Cl 155 mmol/l, KHCO3 10 mmol/l, Na2 W EDTA W 3H2O 0.1 mmol/l, pH 7.4) at 4° for 10 min. The cells were washed twice in PBS and then resuspended in PBS and counted. From this a cell suspension of 1.5 ! 106 cells/ml was made. Preparation of Cytospin/Slides Cytocentrifuge preparations were made using 100-Ìl cell suspensions (input 1.5 ! 105 cells) centrifuged at 550 rpm during 5 min (Cytospin 3; Shandon Southern products, Astmoor, United Kingdom). The slides were dried for 15–20 min with a cold blower, wrapped in aluminium foil and stored at –80° until use. Fixation and Staining Procedure Cells were fixed with formaldehyde and permeabilized with Nonidet P-40 solution as described before [11]. Slides were incubated in duplicate with 50 Ìl 1:5 diluted anti-HCMV-pp65 (C10/C11) [3] for 30 min in a 37° humid chamber, washed twice with PBS, and subsequently incubated with a 50-Ìl HRP conjugated goat Fab anti-mouse IgG (H+L) absorbed with human serum (Protos Immunoresearch, Burlingame, Calif., USA) per spot in a 37° humid chamber for 30 min. After two washings in PBS, the enzyme reaction was performed for 15 min with a 3-amino-9-ethylcarbazole (AEC) solution in 0.1 M acetate buffer (pH 4.9) (AEC 10 mg, Sigma Chemical Co., St. Louis, Mo., USA) dissolved in 4 ml N,N-dimethylformamide and then filled up to 80 ml with acetate buffer, filtered and supplemented with 75 Ìl H2O2 (30% v/v). The slides were then washed with acetate buffer for 10 min and counterstained with hematoxilin Mayer solution (see below for counting all cells), carefully rinsed with tap water and mounted in Kayser glycerin gelatin (Merck 9242). Quantification of Stained Slides All PMNs with a yellowish/brown nuclear staining were considered positive. Results were expressed as positive cells per 50,000 by counting all cells using a grid [1] or a semi-automated image analyser (Quantimet 500, Leica) [13]. Storage Test Fresh blood samples were taken from 9 solid-organ transplant patients positive in the diagnostic HCMV-Agemia assay, with scores between 10 and 250 per 50,000 PMNs. Six patients had HCMVAgemia levels between 10 and 50 per 50,000 PMNs, and 3 scored between 50 and 250 per 50,000 PMN. The fixed and unfixed slides of these patients were vacuum sealed and stored in duplicate at room temperature and at –80°, except for one slide of each patient which was stained on the day of sampling. Frozen slides were brought to room temperature with a cold blower before opening the vacuum seal to prevent the formation of condensation on the slides. Staining (and fixation if necessary) was done at days 1, 2, 7, 14 and 21 according to the standard protocol.


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Table 1. Variations in the HCMV-

antigenemia protocols in the 10 participating laboratories

Fixation method

MoAb

Staining

Quantification

PFA/NP-40 PFA/NP-40 PFA/NP-40 PFA/NP-40 PFA/NP-40 PFA/NP-40 PFA/NP-40 Acetone Acetone Methanol/Acetone

C10/C11 [3] BM2221 1C3, 2A6 and 4C1 [11] C10/C11 [3] CINApool2 C10/C11 [3] Clone 103 28/771 pp65-331 CINApool+4 Emmanuel pool4

PO PO IF IF IF IF IF PO PO IF

per 50,000 PMN per 50,000 PMN per 50,000 PMN per 50,000 PMN per spot per spot per spot per spot per spot per spot

Data are given as stated by participating laboratories. PFA/NP-40 = paraformaldehyde/ Nonidet P-40. References of monoclonal used are given when available. 1 Reference not available. 2 Argene Biosoft cod. 11-002. 3 Clone 10: developed by Dr. J. Booth. 4 Emmanuel pool: pool of monoclonal antibodies developed by Dr. Emmanuel.

Positive Negative Slides All standard slides were made on the day of blood drawing. Negative cytospots were made with PMN from a normal donor. HCMVpp65-positive insect cells (Baculovirus expressed) were made as described [13] and mixed with normal donor PMN at a ratio to obtain a score of 1 100. All slides were dried with a cold blower, wrapped in aluminum foil, vacuum-sealed and stored at –80° until distribution at room temperature (RT). Preparation of Internal Standard For the preparation of the reference standard containing pp65positive cells, two methods were used. Either HCMV-pp65-positive insect cells (Baculovirus expressed) [13] or pp65-positive cells made by coculture of donor PMNs with HCMV-infected endothelial cells [16] were used. To determine the percentage of positive cells in this cellular mixture, a cytospin slide was made and stained according to the standard protocol, and the pp65-positive cells were counted. Then the cellular mixture containing the pp65-positive cells was mixed at different ratios with normal donor PMN and cytospins were made (input 1.5 ! 105 cells). Ratios between pp65-positive cells and normal PMN were chosen to obtain slides with 0, 1–5, 6–10, 25–50, and 1 100 pp65–positive cells per 50,000 cells. Slides were dried with a blower, wrapped in aluminium foil vacuum-sealed and stored at –80° until use or distribution. A set of internal standard slides contained 5 coded slides with 2 cytospots each. Six cytospots of each preparation were stained at the reference laboratory to determine the actual number pp65-positive cells per 50,000 PMNs. Mailing by courier delivery to the participating laboratories took place at RT, advice was given to stain the slides upon arrival or to store them at –80° until use. Participating laboratories received two sets of coded slides A–E marked with the day of preparation without any further information.

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Results

Variations in the HCMV-Agemia Protocols Used by Participating Laboratories The fixation method of most (5/8) laboratories was formaldehyde/NP-40; 2 centers used acetone and 1 laboratory used methanol/acetone. The monoclonal antibody against the HCMV-pp65 antigen was a pool of C10/C11 (Biotest AG, Germany; IQ Products, the Netherlands) in 3 centers, 5 centers used an in-house monoclonal (or a pool of in-house monoclonals), 1 center used a mixture of an in-house monoclonal and Cinapool (Argene Biosoft) and 1 center a CINApool only (table 1). Storage Test Results of experiments on the influence of fixation and temperature on the quality of the slides evaluated at different time points after storage are shown in figure 1. No detectable loss of signal even after 3 weeks was noticed in any of the slides kept unfixed at –80° (fig. 1D). Remarkably a decrease of the scores was seen in a minority of the slides fixed with paraformaldehyde and subsequently stored at –80 ° C (fig. 1C). Similarly fixed slides lost their positivity when stored at room temperature (fig. 1A). When stored unfixed at room temperature (fig. 1B), no significant loss of signal was observed within the first 2 days of storage and the decline appeared less than with fixed slides stored at room temperature.

Verschuuren et al.

Fig. 1. Influence of fixation, storage temperature and time on slides of 9 CMV-Agemia-test-positive patients. All slides were wrapped in aluminum and vacuum-sealed. Each line represents the scores of one sample. A Fixed slides stored at RT. B Unfixed slides stored at RT. C Fixed slides stored at –80°. D Unfixed slides stored at –80°.

Distribution of Positive Negative Slides Based on the results described in the previous section, unfixed slides were aluminum wrapped, vacuum sealed and distributed at RT by courier delivery to ensure delivery within 2 days. All participating laboratories reported positive staining of baculovirus transfected HCMV-pp65 pos cells (data not shown). In general cell morphology was maintained well. No false-positive staining was reported by any laboratory. Internal Standard Slides In general, the participating laboratories were satisfied with the overall quality of the internal standard slides without problems with the staining procedure (fig. 2A, B).


Data are shown in table 2. Average scores of the slides by our laboratory showed that the internal standard preparations made with the baculovirus system were 0, 6, 8, 61 and 511. This was close to the objected number of pp65positive cells per 50,000 PMN (0, 1–5, 6–10, 25–50 and 1100). The scores of our laboratory with the internal standard slides made with pp65-positive PMN were all within objected ratios (average scores 0, 1, 9, 39 and 680). Seven out of nine participating laboratories returned their data of the coded internal standard slides. No false-positive results were reported. In general, all laboratories were able to detect the pp65-positive spiked cells on the cytospots. With the exception of center 2, all centers were able to detect an increasing number of pp65-positive cells with an


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Fig. 2. Photomicrographs of immunoperoxidase stained internal standard slides from the category of 1 100 pp65-positive cells mixed per 50,000 PMN. A Internal standard slide with pp65 baculovirus-infected insect cells. B Internal standard slide with pp65positive PMNs made by coculture of normal donor PMNs with HCMV-infected endothelial cells. Arrows indicate positively stained cells. Note the larger variation in both morphology and staining intensity in the slides spiked with pp65 baculovirus-infected insect cells (A) as compared to the slide spiked with pp65-positive PMNs.

increased number of spiked cells with both types of internal standard slides (table 2). There was a good correlation between the results of the different laboratories for each slide (fig. 3). The absolute scores of the slides with 0, 1–5, 6–10, 25– 50 and 1100 pp65-positive insect cells (baculovirus expressed) showed that, of the centers that quantified per 50,000 PMNs, center 3 (average scores, respectively: 0,

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13, 29, 123 and approximately 1450) scored higher than the reference center (center 1). The results of center 3 with the pp65-positive PMN made by coculture of donor PMN with HCMV-infected endothelial cells (average scores of 0, 2, 11, 79 and 785) were at the same level as the results of the reference center. The results of center 2 (average scores 0, 5, 0, 36 and 440 for the pp65-positive insect cells and 0, 0, 22, 4 and 172 for the pp65-positive PMNs) were

Verschuuren et al.

10,000

pp65 score

1,000

100

10

Fig. 3. Average results of all participating

laboratories with the distributed internal standard slides (table 2). Results are expressed as pp65-positive cells per 50,000 PMNs if available. For centers that quantified per cytospot, results per cytospot were used.

£1 Number of spiked cells per 50,000 PMN Type of spiked cell

1-5

6-10

25-50

>100

1-5

bac-pp65-infected insect cells

6-10

25-50

>100

in vitro pp65 positive PMN generated

Table 2. Results of the 8 responding laboratories of the coded unfixed internal standard slides distributed at room temperature to the 10 participating laboratories

Center

Number of HCMV-pp65-positive cells per 50,000 PMNs or per cytospot1 baculovirus-expressed pp65-positive insect cells

pp65-positive PMN made by coculture

0

1–5

6–10

0

1–5

6–10

25–50

1 (ref) 2 3

0 0/0 0/0

6B2 7/3 10/16

8B5 0/0 22/35

51B13 17/55 –/123

511B141 295/584 F1,500/F1,400

0 0/0 0/0

1B1 0/0 1/2

9B3 17/26 8/13

39B5 5/2 83/75

680B106 199/142 F670/900

11 41 51 61 71 81

0 0/0 0/0 0/0 0/0 0/0

8B3 13/8 0/7 19/15 15/13 5/2

10B5 13/11 17/15 35/27 23/24 22/12

66B13 114/102 44/19 107/127 124/120 56/37

376B74 890/677 646/420 ND/ND F1,200/F1,100 1,000/1,000

0 0/0 0/0 0/0 0/0 0/0

3B2 0/2 1/0 0/0 0/0 3/1

22B16 4/8 14/14 4/8 7/9 10/11

59B10 14/44 57/55 26/27 37/32 60/48

1,079B74 596/399 1,468/2,132 1200/1 200 F1,100/1,000 700/750

25–50

1 100

1 100

Staining was done according to the local protocol of the participating centre and scores of two cytospots (spot A/spot B) are shown. Results are expressed as pp65-positive cells per 50,000 PMNs or pp65-positive cells per cytospot. Two kinds of pp65-positive cells were used: baculovirus-expressed pp65-positive insect cells and in vitro generated pp65-positive PMNs. Results of the reference center are expressed as mean B standard deviation (n = 6) of both pp65-positive cells per 50,000 PMN and positive cells per cytospot. ND = not done. 1 Counts performed per cytospot.

neither in line with the expected results no with the results of the other participants for both the internal standard slides made with the pp65-positive insect cells and the pp65-positive PMNs. Of the centers that quantified per cytospot center 5 (average readings 0, 4, 16, 32 and 533 per cytospot) and center 8 (average readings 0, 3, 16, 47 and 1,000 per cytospot) were comparable with the reference center (average read-

ings 0, 8, 10, 66, 376 respectively per cytospot) and center 4 (average readings 0, 11, 12, 108, 784 per cytospot), center 6 (0, 17, 31, 117 and not done per cytospot) and center 7 (0, 14, 24, 122 and approximately 1,150 per cytospot) scored higher in the baculovirus-transfected pp65-positive cells than the reference center. With the pp65-positive PMNs, center 5 (readings of 0, 1, 14, 56 and 1,800 per cytospot) and center 8 (readings of



387

0, 2, 11, 54 and 725 per cytospot) had results comparable with those of the reference center (readings 0, 3, 22, 59 and 1,079 per cytospot), whereas center 4 (readings 0, 1, 6, 29 and 498 per cytospot), center 6 (readings of 0, 0, 6, 27 and 1200 percytospot) and center 7 (readings of 0, 0, 8, 35 and 725 per cytospot) had a lower result than the reference center. However, since these readings were not quantified per 50,000 PMNs, these results may reflect different cell numbers per spot. The ratios between the preparations with 6–10 and 25–50 pp65-positive cells were very similar (3–5) for the pp65-positive PMN in all laboratories that quantified per cytospot. The ratios were less similar (3–9) for the same centers when the baculovirus system was used. One center withdrew from the study, one center could not interpret the results due to high background staining.

Discussion

This study describes the preparation of pp65-positive standard slides and the evaluation of these slides by 8 different laboratories. Although differences in the absolute values were recognized, the results of the readings by the different laboratories were remarkably similar, despite distribution at RT and the differences in HCMV-Agemia protocols used by the participating laboratories. This was especially true for the in-vitro-generated pp65-positive PMNs made by coculture, for which their potential usefulness for standardization was recently described by Gerna et al. [16]. The main conclusion is that these standard slides are well suited for standardization purposes and can be distributed at RT, provided they are wrapped in aluminum, vacuum-sealed, and stored at –80° upon arrival until staining. This enables the different laboratories to evaluate and compare their own assay protocols including differences in fixation techniques. Since the participating laboratories were reluctant to abandon their own modifications of the protocol, it is of major importance to have an internal standard that performs well in all these different laboratories. Optimal storage conditions were evaluated to determine the distribution conditions. These proved to be storage of unfixed slides at –80°. Because of the logistic problems involved with distribution of material at –80° and as no apparent decrease in signal was seen with unfixed slides at RT, we concluded that, for standardization purposes, unfixed slides can be sent at RT. This was confirmed by the distribution of the positive negative slides and later by the results of the internal standard slides.

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Distribution and evaluation of these internal standard slides showed remarkable results. None of the responding laboratories had false-positive staining. Six out of 8 responding laboratories detected not only the positive cells but also detected the increasing number of spiked cells in the coded slides. This means that in spite of the differences in the protocols the results are remarkably similar. At the start of this concerted action only the baculovirus system was available to make pp65-positive cells. Therefore, this system was chosen to prepare the pp65positive cells for the positive negative slides of the internal standard. However, during the study it became possible to make pp65-positive PMN in vitro [14, 15]. Both methods were used to make internal standard slides. Besides morphological considerations, both types of internal standards have two important differences. The pp65 protein is actively produced in the pp65-baculovirus infected insect cells. During the course of infection, the polyhedrin promoter that drives the expression of pp65 is switched on, with maximal expression reached at late stages of baculovirus infection. As baculovirus infections generally are not synchronous, cells will harbor differing amounts of intracellular pp65. This was observed by the participants as differences in staining intensity of the baculovirus-spiked internal standards (fig. 2A). Thus the criterion for positivity appeared more difficult to set as compared to pp65-positive PMN and probably has led to more variation in the quantitation results (table 2). Furthermore, bac-pp65 is a recombinant protein which might influence the staining capacity of some of the anti-pp65 monoclonal antibodies. The use of in-vitro-generated pp65-positive PMN as internal standard may more closely resemble the patient situation. Nevertheless, these pp65 positive PMN are also a phenomenon generated in vitro in which the pp65 was acquired by co-cultivation of PMN with HCMV-infected endothelial cells. In contrast to infected cells, these PMN do not produce pp65 [17]. As it appears, PMNs generally seem to acquire similar amounts of pp65 upon cocultivation, which is reflected in a more homogeneous staining pattern of the pp65-PMN internal standard (fig. 2B). This allowed for a lower variation in the quantification of this type of internal standard, making it better suited than pp65 baculovirus-infected cells. When compared to the infection of insect cells with baculovirus, the production of pp65-positive PMNs generated in vitro is more laborious and the transfer protocol needs further optimization. Nevertheless, the use of pp65positive PMNs generated in vitro is to be the preferred method for production of internal slides.

Verschuuren et al.

Whether the sensitivity or the threshold of the laboratories differ cannot be decided. This will be investigated with the distribution of external (patient) standard slides in the near future. This will answer the question whether the assay is to be standardized for comparison of the results. In conclusion, this study describes the basic requirements towards the standardization of the HCMV-Agemia assay, and the main results are that unfixed standard slides can be distributed at RT provided they are wrapped in aluminum, vacuum-sealed, and stored at –80° upon arrival until staining. The pp65-positive PMNs generated in vitro made by coculture are better suited for standard-

ization purposes than the baculovirus-expressed pp65positive insect cells. Within individual laboratories, the relative differences between the standard samples were recognized correctly. However, in absolute terms, there were still differences between the participating centers. These differences can be diminished with standard slides as described here so an optimal concordance of test results between different laboratories can be obtained. Acknowledgments This study was supported by the European Community Grant No. ERB BMHMCT96-0471 (DG 12-SSMA).

References 1 van der Bij W, Torensma R, van Son WJ, Anema J, Schirm J, Tegzess AM, The TH: Rapid immunodiagnosis of active cytomegalovirus infection by monoclonal antibody staining of blood leucocytes. J Med Virol 1988;25:179– 188. 2 Boeckh M, Bowden RA, Goodrich JM, Pettinger M, Meyers JD: Cytomegalovirus antigen detection in peripheral blood leukocytes after allogeneic marrow transplantation. Blood 1992;80:1358–1364. 3 van den Berg AP, Klompmaker IJ, Haagsma EB, Scholten-Sampson A, Bijleveld CM, Schirm J, van der Giessen M, Slooff MJ, The TH: Antigenemia in the diagnosis and monitoring of active cytomegalovirus infection after liver transplantation. J Infect Dis 1991;164: 265–270. 4 The TH, van der Ploeg M, van den Berg AP, Vlieger AM, van der Giessen M, and van Son WJ: Direct detection of cytomegalovirus in peripheral blood leukocytes – A Review of the Antigenemia Assay and Polymerase Chain Reaction. Transplantation 1992;54:193–198. 5 Gerna G, Zipeto D, Parea M, Revello MG, Silini E, Percivalle E, Zavattoni M, Grossi P, Milanesi G: Monitoring of human cytomegalovirus infections and ganciclovir treatment in heart transplant recipients by determination of viremia, antigenemia, and DNAemia. J Infect Dis 1991;164:488–498.


6 The TH, van den Berg AP, van Son WJ, Klompmaker IJ, Harmsen MC, van der Giessen M, Sloff MJ: Monitoring for cytomegalovirus after organ transplantation: A clinical perspective. Transplant Proc 1993;25(suppl 4):5– 9. 7 Docke WD, Simon HU, Fietze E, Prosch S, Diener C, Reinke P, Stein H, Volk HD: Cytomegalovirus infection and common variable immunodeficiency (Letter): Lancet 1991;338: 1597. 8 Bein G, Bitsch A, Hoye J, Kirchner H, The detection of human cytomegalovirus immediate early antigen in peripheral blood leucocytes. J Immunol Methods 1991;137:175–180. 9 Revello MG, Percivalle E, Zavattoni M, Parea M, Grossi P, Gerna G: Detection of human cytomegalovirus immediate early antigen in leukocytes as a marker of viremia in immunocompromised patients. J Med Virol 1989;29: 88–93. 10 Jiwa NM, van de Rijke FM, Mulder A, van der Bij W, The TH, Rothbarth PH, Velzing J, van der Ploeg M, Raap AK: An improved immunocytochemical method for the detection of human cytomegalovirus antigens in peripheral blood leukocytes. Histochemistry 1989;91: 345–349. 11 Gerna G, Revello MG, Percivalle E, Morini F: Comparison of different immunostaining techniques and monoclonal antibodies to the lower matrix phosphoprotein (Pp65) for optimal quantitation of human cytomegalovirus antigenemia. J Clin Microbiol 1992;30:1232– 1237.

12 Wunderli W, Kagi MK, Gruter E, Auracher JD: Detection of cytomegalovirus in peripheral leukocytes by different methods. J Clin Microbiol 1989;27:1916–1917. 13 The TH, van den Berg AP, Harmsen MC, van der Bij W, van Son WJ: The cytomegalovirus antigenemia assay: A plea for standardization. Scand J Infect Dis Suppl 1995;99:25–29. 14 Grundy JE, Lawson KM, MacCormac LP, Fletcher JM, Yong KL: Cytomegalovirus-infected endothelial cells recruit neutrophils by the secretion of C-X-C chemokines and transmit virus by direct neutrophil-endothelial cell contact and during neutrophil transendothelial migration. J Infect Dis 1998;177:1465–1474. 15 Revello MG, Percivalle E, Arbustini E, Pardi R, Sozzani S, Gerna G: In vitro generation of human cytomegalovirus Pp65 antigenemia, viremia, and leukoDNAemia. J Clin Invest 1998; 101:2686–2692. 16 Gerna G, Percivalle E, Torsellini M, Revello MG: Standardization of the human cytomegalovirus antigenemia assay by means of in vitrogenerated Pp65-positive peripheral blood polymorphonuclear leukocytes. J Clin Microbiol 1998;36:3585–3589. 17 Grefte JM, van der Gun BT, Schmolke S, van der Giessen M, van Son WJ, Plachter B, Jahn G, The TH: Cytomegalovirus antigenemia assay: Identification of the viral antigen as the lower matrix protein Pp65 (letter). J Infect Dis 1992;166:683–684.


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New Advances in the Diagnosis of Congenital Cytomegalovirus Infection T. Lazzarotto S. Varani L. Gabrielli P. Spezzacatena M.P. Landini Department of Clinical and Experimental Medicine, Division of Microbiology and Virology, University of Bologna, Bologna, Italy

Key Words Cytomegalovirus W Diagnosis W Pregnancy W Congenital infection W PCR W Amniotic fluid W Antibody avidity W IgM

Abstract With the advances in anticytomegalovirus (anti-CMV) serology, the new recombinant IgM tests seem likely to become the screening tests for pregnant women whose prepregnancy serological status for CMV is unknown. When a woman is found to be IgM-positive, further diagnostic evaluation focused on determining whether this is due to a primary infection should be carried out. Maternal primary infections that were difficult to determine until a few years ago unless documented by seroconversion can now be readily diagnosed from the presence of low-avidity anti-CMV antibody which persists for approximately 20 weeks after primary infection. In primarily infected mothers prenatal diagnosis can be performed between 21 and 23 weeks of gestation, and the amniotic fluid (AF) represents the pathological material of choice to determine intrauterine virus transmission. In AF, the virus can be detected by culture and/or PCR. Both procedures differentiate uninfected from infected fetuses, but cannot predict fetal outcome. The determination of the viral load in AF carried out by quantitative PCR is more promising and could represent an important starting point for preemptive fetal therapy. Copyright © 2000 S. Karger AG, Basel

ABC

© 2000 S. Karger AG, Basel 0300–5526/99/0426–0390$17.50/0



Introduction

Cytomegalovirus (CMV) is the leading infectious cause of CNS malformations in children [1–4]. In developed countries, the virus is acquired congenitally in 0.3–2% of all live births [5–8]. Congenitally acquired CMV causes clinical damage in 90% of the newborns that are symptomatic at birth but also in approximately 10% of those who are asymptomatic at birth [9, 10]. Although mother-fetus transmission can occur at any time throughout gestation, infection during the first 16 weeks of pregnancy has been associated with a higher rate of damage [11]. Furthermore, although intrauterine transmission may be the result of either exogenous or endogenous maternal infection, a large study recently reported a 1.2 and 12.9% transmission in seropositive and seronegative women, respectively, indicating that preconceptional maternal immunity is protective against congenital CMV infection, decreasing the risk of infection by 90% [12]. Thus seropositive pregnant women (50–70% of the cases in industrialized countries) may encounter an endogenous viral reactivation or exogenous viral reinfection in the presence of specific immunity. In these subjects, the risk of transmission is approximately 1% and, even when transmission occurs, it is linked to a less virulent outcome than when the virus is transmitted in the absence of specific immunity [13]. Therefore, primary maternal infections are the infections to be diagnosed in pregnant women.

Maria Paola Landini, MD Department of Clinical and Experimental Medicine Division of Microbiology and Virology, Policlinico S. Orsola Via Massarenti n. 9, I–40138 Bologna (Italy) Tel. +39 051 346306, Fax +39 051 341632, E-Mail [email protected]

Serological Advances The Screening Test. A good screening test is characterized by its ability to identify all women undergoing an active CMV infection, its low cost, its completely automated nature and its worldwide distribution. The detection of CMV-specific IgM has traditionally been considered the most appropriate procedure for screening pregnant women. However, this procedure has always been hampered by the fact that the correlation of results obtained with different commercial kits is poor, and contradictory results are obtained if a serum sample is tested with two different kits [16, 17]. The recent introduction onto the market of new tests may lead to a rapid change in this situation. Detection of CMV-specific IgM is most commonly effected using preparations of the virus or viral lysate in an enzyme-linked immunosorbent assay (ELISA) [18– 24]. The key serological targets for detection of CMV-spe-

cific IgM include both structural (pUL32, pUL83, pUL80a) [25–29], and nonstructural (pUL57, pUL44) [24, 30–32] viral proteins. Variations in the relative amounts of these antigens produced during growth and purification of the virus can result in a different stoichiometric composition of the viral antigens used in the various IgM tests, leading to the discordant interassay results observed. Antigenic materials composed of peptides or purified recombinant proteins produced through molecular biology offer an attractive alternative for the detection of CMV-specific IgM [16, 18, 24, 25, 31–35]. We have shown that a cocktail of purified recombinant protein antigens containing portions of pUL32, pUL44, pUL83 and pUL80a is both necessary and sufficient to replace the virus in a microtiter ELISA for the detection of CMV-specific IgM [19, 36]. We have also developed and evaluated a fully automated recombinant antigen-based CMV IgM immunoassay which contains the same recombinant proteins necessary and sufficient in the microtiter ELISA [37]. The results obtained were in reasonably good agreement with the consensus interpretation of three commercially available CMV IgM ELISAs. Furthermore the new tests seem sensitive enough to detect all pregnant women undergoing an active infection, with an overall IgM positivity rate of 4.5% among both European and American pregnant women [37]. This percentage is consistent with the percentage of actively infected pregnant women as indicated by excretion of the virus [22]. A review of data from the literature indicates that 5–15% of randomly selected pregnant women from industrialized countries excrete infectious virus from one or more sites, the cervix being the most relevant [38–43]. In view of seroepidemiological data indicating that the seroprevalence in the same countries is approximately 60%, we can deduce that 2–6% of the virus excretion is due to primary infections, while 3–9% is due to viral reactivations. Furthermore, as IgM is produced in 100% of primary infections and in 70% of recurrences [44, 45], we should expect a prevalence of IgM-positive women of 4– 11%. Because of the sensitivity of the recombinant tests, their relatively low cost and their complete automation, they could represent the best screening method available today for pregnant women whose prepregnancy serological status for CMV is unknown. When anti-CMV IgM is detected in a pregnant woman, the diagnostic problem is not over. In fact, only 10% of IgM-positive women will have a congenitally infected fetus/newborn [46]. Therefore positivity for the IgM test should be considered solely as the starting point for

Diagnosis of Congenital CMV Infection


In general, over 90% of primary CMV infections in pregnant women, as in most otherwise healthy individuals, are asymptomatic and are likely to go undetected by the woman and her physician. Furthermore, the symptoms accompanying the rare symptomatic cases are fever, fatigue, headache, myalgia and sore throat, which are not specific for CMV. Therefore, laboratory techniques represent a decisive diagnostic approach.

Diagnosis of Primary Infection

The diagnosis of primary infection is relatively easy if a seroconversion is detected. However, in most cases pregnant women do not know their prepregnancy serological status for CMV. How can pregnant women undergoing an acute CMV infection be identified if this is asymptomatic? The only possibility is to screen all pregnant women. Many countries do not screen for CMV in pregnancy and accept that approximately 3 out of 1,000 newborns will suffer a congenital CMV infection [14, 15]. This is due mainly to the cost of screening, and because of evidence that the tests available for screening were unreliable. Furthermore, when CMV infections were detected in pregnant women, there was no way of identifying whether a fetus was at risk. If this was true until some years ago, during the last few years significant advances in both the serologic and virologic diagnoses of congenital CMV infections have been achieved which are changing the face of diagnosis of congenital infection. This review focuses on these advances.

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a comprehensive diagnostic evaluation to determine whether a primary infection was involved. Identification of Primary Infections. Several procedures have been described to differentiate primary from recurrent infections [27, 47–51]. Using a microneutralization assay, it was found that neutralizing antibodies first appear approximately 15 weeks after acute infection [52], making it possible to identify recent primary infection by the absence of neutralizing antibodies. This result is probably linked to the fact that antibodies to glycoproteins appear later than antibodies to other nonglycosylated viral proteins [53]. Another interesting and recent possibility is the determination of antibody avidity. The term IgG avidity is indicative of the low functional affinity of the IgG class antibody. During the first weeks following primary infection antibodies show a low avidity for the antigen. They progressively mature acquiring higher avidity. This characteristic is used at the diagnostic level to discriminate recent primary infections in several viral diseases [54–62]. Our own experience with anti-CMV-IgG-avidity testing (by ELISA) using a commercially available kit shows that a reliable distinction between primary and nonprimary CMV infection in pregnant women is possible. In fact more than 90% of primary infections and no recurrences revealed low-avidity IgG to CMV. In particular, low IgG avidity is a marker of primary infection for 18–20 weeks after the onset of symptoms in immunocompetent subjects [45]. Similar results were obtained by GrangeotKeros et al. [63]. IgM reactivity to different CMV proteins as detected by immunoblotting has also been shown to be useful in identifying pregnant women at risk of transmitting the virus to their fetuses [64]. In this respect, we determined the avidity index of anti-CMV IgG and the anti-CMV IgM profile in pregnant women at risk of transmitting CMV to their offspring [65]. The results obtained indicate that if the determination of anti-CMV antibody avidity is carried out before 18 weeks of gestation, it can identify all pregnant women who will give birth to an infected newborn. In contrast, IgM detected by blot showed a 69% sensitivity [65, 66]. Therefore it seems that the early determination of anti-CMV antibody avidity is a helpful tool in identifying a subgroup of women at risk of transmitting the infection. At this point, an interesting question arises regarding the possibility of using the avidity test as a screening test. We do not think that at present the avidity test could replace IgM detection as a screening test as its cost is approximately double that of the IgM test, it is not com-

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pletely automated and also because few companies produce it and distribution is not worldwide. Furthermore, even though the increase in CMV-specific IgM and IgG in response to CMV infection has been shown to occur simultaneously in most cases [67], in a recent study we detected CMV IgM prior to the detection of CMV IgG in 2 out of 39 women [37]. Virological Advances Although CMV excretion is a relatively common event during and after pregnancy, data from the literature indicate that the simple isolation of the virus during pregnancy, whether from the cervix or urine or both, is a poor indicator of the risk of intrauterine infection [38–43]. Perhaps because isolation of CMV or detection of CMV antigenemia from the blood of normal hosts is rarely successful since the viremia phase is much shorter than in immunocompromised subjects, the detection or quantitation of CMV in maternal blood as a prognostic marker of intrauterine transmission has received little attention until recently. An important result in this field was the finding that CMV could be detected in the blood of pregnant women only during ongoing or recent primary infections [68–70]. In particular, Balcarek et al. [68] reported a 10-fold higher risk of congenital CMV infection for the offspring of viremic mothers than those without evidence of viremia. In contrast, a more recent study showed that the maternal viral load in blood, determined by antigenemia, viremia and DNAemia, is low and does not correlate with the clinical course of infection, intrauterine transmission and the severity of outcome [70]. The same authors reported that the sensitivity of the antigenemia and viremia tests in the first month after the infection was low, detecting about 50 and 25% of positive patients, respectively, whereas after 2 months, only 25% of the patients remained antigenemia-positive and none were positive for viremia. Thus, antigenemia and viremia appear to be the only markers of acute primary CMV infection. LeukoDNAemia was detected in 100 and 90% of subjects examined, respectively 1 and 2 months after onset it remained positive for 6 months, but only about 50 and 25% of the patients were positive 3–6 months after infection. The persistence of CMV DNA in peripheral blood lymphocytes (PBL) for 3 months or more and the detection of viral DNA levels 650 GE/1 ! 105 PBL did not seem to be a factor associated with a higher risk for fetal infection. From these few studies, it is premature to draw any conclusion, but further investigations in this field are strongly recommended.

Lazzarotto/Varani/Gabrielli/Spezzacatena/ Landini

Prenatal Diagnosis

In cases where primary CMV infection or maternal infections with viremia are diagnosed during pregnancy, obvious concerns arise over the potential effects of intrauterine transmission to the developing fetus. Although the natural history of intrauterine CMV infection is not well understood, it is clear that some fetuses are irreversibly damaged by the virus before delivery. Those infants would not derive much benefit from postnatal therapy, but if infected fetuses could be detected before this irreversible stage has been reached, treatment in utero (when available) might have a significant effect on the course of the disease [71, 72]. In our opinion, prenatal diagnosis should be offered to pregnant women undergoing a primary infection or a high-viral-load infection during the first half of gestation. We also believe that if prenatal diagnosis is not offered or not available, the entire preliminary diagnostic phase (screening and avidity determination) is meaningless. Judging from our experience, approximately 70% of prenatal diagnoses would have a negative result, indicating the absence of fetal infection. This would prevent unnecessary termination of uninfected fetuses and let the woman continue her pregnancy with a higher level of confidence. Material to Be Used for Prenatal Diagnosis The first issue is the decision about what material is to be used for prenatal diagnosis. It is thought that CMV is transmitted when infected leukocytes cross the placental barrier to reach the fetal circulation via the umbilical cord vessels [73, 74]. The virus replicates in the fetal tissues and is then excreted into the amniotic fluid (AF) via fetal urine. Infected AF would be ingested by the fetus, and the virus could then replicate in the oropharynx and enter the fetal circulation to reach target organs. Therefore fetal blood and AF seem to be two logical choices of bodily fluid for the prenatal diagnosis of CMV transmission [46, 75–86]. We and others have compared AF and fetal blood and found similar results that strongly support the use of AF [46, 78, 81–86]. It is important to stress that the concomitant presence of viral DNA in maternal blood does not seem to represent a source of the viral DNA detected in AF through invasive procedures. This finding also implies that iatrogenic transmission of the infection seems unlikely, as described by Revello et al. [70].


When to Perform Amniocentesis A critical decision in prenatal diagnosis is when to perform amniocentesis. Two factors should be considered: firstly, some false-negative results have been reported before 20 weeks of gestation, as the fetus excretes CMV into the AF via urine and fetal diuresis only becomes established after 20–21 weeks of gestation [69]. Secondly, in most cases at least 6–9 weeks have to pass from the time of maternal infection before the virus can be detected in AF, and CMV transmission is correlated with severe fetal diseases mainly when it occurs during the first 12 weeks of gestation [82]. From the above considerations it seems that amniocentesis should be delayed as long as possible with respect to maternal infection, to reduce the risk of false-negative results due to delayed intrauterine transmission and, when indicated, a second procedure should be considered [86]. We routinely perform amniocentesis between 21 and 23 weeks of gestation [46]. We believe it is unlikely that infections occurring at the beginning of pregnancy could pass undetected because in analogy with what can be seen in congenitally infected newborns, fetal excretion could last for months. Test to Be Used on Amniotic Fluid We have recently reported the results of a large-scale study on prenatal diagnosis of congenital CMV infection carried out by both PCR and virus isolation from AF in pregnancies at risk of transmitting CMV [87]. Our results indicate that a positive PCR result in AF should be considered of very limited diagnostic value because its positive predictive value is approximately 50% [46, 87]. Low levels of viral DNA in AF in the absence of virus isolation and fetal infection is an intriguing and still unexplained finding. A positive virus isolation from AF is more indicative of congenital infection, but false negatives are not uncommon as intact viral particles are required to be infectious and to be detected in culture [46, 80, 82, 86, 87]. Intrauterine transmission does not necessarily mean that the fetus will suffer from the infection, and thus a further diagnostic problem arises as to how to identify fetuses at risk of developing severe CMV disease. The ultimate goal is the identification of an infected fetus at high risk of developing a severe CMV-related pathology, for two reasons: (1) to allow the mother to opt for pregnancy termination, and (2) with the advances in antiviral chemotherapy, the prenatal identification of a severely infected fetus could also allow preemptive fetal treatment [71, 72].


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Fig. 1. A management scheme for prenatal diagnosis of congenital CMV infection.

Until very recently, high risk fetuses were identified exclusively by fetal ultrasound scanning or other tests as part of a woman’s routine prenatal care. Findings in the fetus that should alert the clinician to the possibility of intrauterine CMV infection include the presence of oligohydramnios or polyhydramnios, nonimmune hydrops, fetal ascites, intrauterine growth retardation, microcephaly, cerebral ventriculomegaly or hydrocephalus, intracranial calcifications, pleural or pericardial effusion, hepatosplenomegaly, intrahepatic calcifications or pseudomeconium ileus. However, as other intrauterine infections (HSV, varicella-zoster virus, HIV, rubella, syphilis and

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toxoplasmosis) can cause similar clinical findings, a differential diagnosis should be made. It is obvious that in most cases clinical signs are identified too late to allow pregnancy termination. In those cases with documented viral transmission, the determination of the viral load in the AF by quantitative PCR seems to be able to identify fetuses at higher risk of developing a severe infection [87]. A management scheme for prenatal diagnosis of congenital CMV infection is proposed in figure 1.


Conclusions

The availability of more reliable IgM tests for screening pregnant women whose prepregnancy serological status for CMV is unknown, tests to determine the avidity index of anti-CMV IgG which can diagnose a primary CMV infection and PCR to detect the virus in AF have been very important steps torwards solving the diagnostic problem linked to congenital CMV infection. A few further steps, mainly in the prenatal determination of fetuses at risk of severe handicap, are foreseeable and should make it possible to resolve the diagnostic problem of congenital CMV infection.

Finally, it is extremely important that those who have the power to decide public health strategies should be acquainted with what has been discovered within the last 5 years and should act accordingly.

Acknowledgments The authors are grateful to the entire staff of the Laboratory of Molecular Virology for their continuous dedicated work and to Dr. B. Guerra (II Department of Obstetrics and Gynecology, University of Bologna) for the work with the patients. Personal investigations outlined in this review were partially supported by the Italian Ministry of Public Health (40 and 60%), the AIDS projects of the Ministry of Public Health, the Italian Ministry of Education and the University of Bologna.

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Significance of Qualitative Polymerase Chain Reaction Combined with Quantitation of Viral Load in the Diagnosis and Follow-Up of Cytomegalovirus Infection after Solid-Organ Transplantation H. Vanpoucke B. Van Vlem R. Vanholder L. Van Renterghem Laboratory Virology, University Hospital, Gent, Belgium

Key Words Qualitative PCR W Viral load W Cytomegalovirus W Transplantation

Abstract Quantitative PCR was evaluated in the monitoring of patients with ongoing posttransplantation cytomegalovirus (CMV) infection and antiviral therapy, compared to leukoDNAemia and serology. From January 1998 until May 1999, 61 patients were followed up weekly during 3 months after transplantation by a qualitative PCR. The quantitative PCR was performed on plasma samples from 21 selected patients, of whom 12 had a primary infection and 9 a reactivation or reinfection. Analysis of the viral load differences showed that the viral loads in patients with a primary infection were significantly higher than viral loads in patients with a reactivation (p ! 0.01). Based on the results of our study, we can state that qualitative PCR is a good marker for initiating preemptive therapy. In addition, viral quantitation is clinically useful for accurate diagnosis of established CMV disease, and monitoring of antiviral therapy. Copyright © 2000 S. Karger AG, Basel

ABC

© 2000 S. Karger AG, Basel 0300–5526/99/0426–0398$17.50/0



Introduction

Human cytomegalovirus (CMV) infection and disease are still a major problem in solid-organ transplant recipients. Because the intensity of immunosuppressive therapy influences the incidence and severity of CMV infection, infections usually occur during the first 4 months after transplantation [1]. Although antiviral prophylaxis led to a reduction in both the morbidity and mortality of CMV disease, the toxicity associated with the currently available antiviral agents (such as ganciclovir and foscarnet) remains a problem. This, together with overtreatment of many patients, the cost and the possibility of emergence of resistant CMV strains, implies that universal prophylaxis is not an ideal approach [2]. Therefore, efforts have been made to develop highly sensitive and quantitative detection methods to identify patients at risk for disease prior to its onset, thereby focusing antiviral treatment only to patients at risk for disease. Detection of early CMV infection relies on diagnostic methods, such as shell vial cultures, pp65 antigenemia and qualitative polymerase chain reaction (PCR) performed on peripheral blood samples [3]. The practical significance of culture-based assays has been limited by the relatively low sensitivity, the time-consuming nature of the methods, and the rapid loss of viability in stored spec-

Lieve Van Renterghem Laboratory of Bacteriology and Virology, Renal Division Department of Internal Medicine, Ghent University Hospital De Pintelaan 185, B–9000 Ghent (Belgium) Tel. +32 9 240 36 31, Fax +32 9 240 38 65, E-Mail [email protected]

imens. pp65 antigenemia is a sensitive method, but needs immediate and time-consuming processing and subjective slide interpretation [4]. Qualitative PCR on peripheral blood leukocytes is more sensitive and detects viral replication in the peripheral blood earlier than antigenemia, providing a longer period of time between detection of viral replication and onset of symptomatic infection and enabling earlier initiation of preemptive therapy [2, 5, 6]. High viral load is frequently correlated with clinical symptoms [7, 8]. A positive result for qualitative PCR on plasma is roughly correlated with higher viral load in the blood. However a quantitative assay is needed to determine the threshold values above which CMV-related clinical symptoms are likely to appear. Therefore, in a retrospective study, we have evaluated the possible clinical significance of qualitative PCR combined with the quantitation of the viral load in the diagnosis and follow-up of CMV infection after solid organ transplantation.

Materials and Methods Patients and Sample Collection From January 1998 until May 1999, 2 living-related and 59 cadaveric kidney transplantations were performed in our hospital. Thirteen of these transplantations were simultaneous kidney and pancreas transplantations and 1 was a kidney and heart transplantation. Twenty-one of them were receptor as well as donor CMV IgG negative and at low risk for a CMV infection. Fourteen were at high risk for a primary infection, being seronegative and obtaining a CMV-infected organ. Twenty-six were CMV seropositive before transplantation. In all these patients, EDTA blood was collected weekly during the first 3 months after transplantation. Immunosuppressive Therapy The immunosuppressive regimen of kidney and/or pancreas transplant patients consisted of triple therapy with induction. Induction was either with antithymocyte globulins or with IL-2 receptor antagonists (Basiliximab). Maintenance immunosuppressive therapy was started preoperatively: mycophenolate mofetil (MMF) and methylprednisolone and, only in combined kidney-pancreas transplantation, cyclosporine microemulsion. In the kidney-only transplant recipients, cyclosporine microemulsion was started at the time serum creatinine reached 50% of the initial value. One intravenous pulse of 160 mg methylprednisolone was given on day 0;, from day 1 onwards, only 32 mg was given, tapered towards 8 mg after 4 weeks. The maintenance dose of MMF was 3 g in kidney-pancreas transplants and 2 g in single-kidney transplants. Cyclosporine microemulsion was adjusted to trough levels, 250 ng/ml in kidney-pancreas and 150 ng/ml in kidney-only transplants. First rejections were diagnosed clinically and treated with pulse steroids (500 mg methylprednisolone on day 1, 2 and 4). Steroid-resistant rejections were biopsy confirmed and treated with anti-CD3 monoclonal antibodies (OKT3).

Diagnosis and Follow-Up of Cytomegalovirus Infection

Antiviral Therapy CMV prophylaxis in kidney-only transplant recipients depended on donor and recipient CMV serology and consisted of peroral acyclovir for 3 months, for D+/R+, D+/R– or D–/R+ patients. D–/R– patients received no prophylaxis. Pancreas and kidney transplant patients received 10 days of intravenous ganciclovir, regardless of donor or acceptor CMV serology. In case of PCR positivity for CMV, no change in prophylactic therapy was made nor was the patient treated preemptively. In case of a CMV-symptomatic infection, the patient was treated with intravenous ganciclovir, the first 12 days, continuing perorally in case of persistent positive PCR after intravenous therapy. Disease relapse or persistence was treated with an additional course of intravenous ganciclovir, and hyperimmune anti-CMV globulins were added. If this combination did not resolve all symptoms, ganciclovir was replaced by foscarnet. Definition CMV infection is defined as the detection of HCMV by qualitative PCR in at least two consecutive samples, and serological response (appearance of IgM and IgG for primary infection; at least duplication of IgG titer for reactivation and reinfection). Symptomatic infection is defined as otherwise unexplained pyrexia with or without leukopenia, thrombocytopenia and/or involvement of organs (hepatitis, pneumonia, encephalitis). Serology Patients were monitored weekly for active CMV infection by serology: IgM detection (Diasorin, Italy) and/or an increase in IgG (Eurogenetics, Belgium) titer at least twice between two samples in the same test run. Qualitative PCR Sample Preparation and DNA Extraction. The leukocyte fraction from 0.5 ml EDTA blood is isolated and DNA is extracted with the whole-blood DNA specimen preparation kit (Amplicor®) following the manufacturer’s instructions. DNA from 0.2 ml plasma is extracted using the QIAamp Blood Kit (Qiagen). Oligonucleotides. The primers used in the nested PCR assay are taken from the morphological transforming region II of Towne strain CMV [9]. The outer primer pair are 5)-GGT GAT GCT GTC GGT GAT GG-3), complementary to DNA strand nucleotides 290–309 and 5)-GCA TCC CCA ACA GCC TTT G-3), corresponding to DNA nucleotides 615–597. These primers amplify a 326-bp fragment of DNA. The inner pair of primers are 5)-CTA GGC GCT TCC GAG GAG GC-3), complementary to DNA strand nucleotides 342–361 and 5)-CAC GCG GAA AAG AAA GAC CGT-3), complementary to DNA nucleotides 525–505. These primers amplify a 184-bp fragment. PCR. The amplification reaction is done in a Perkon Cetus thermal cycler. Five microliters of target is amplified in a 25-Ìl reaction mixture containing final concentrations of 0.4 ÌM each primer (synthesized by Biosource, Europe), 0.2 mM each deoxynucleotide (Amersham Corp.), 10 mM Tris-HCl (pH 8.3), 50 mM KCl, 2.5 mM MgCl2, and 0.625 units of Taq polymerase (Perkin Elmer Cetus). The reaction solution is overlaid with 25 Ìl of light mineral oil to prevent evaporation. For the first set of primers, the amplification is initiated by preheating the reaction tubes for 10 min at 94° followed by 30 cycles of alternating denaturation for 15 s at 94°, primer annealing for 15 s at 57°, and primer extension for 15 s at 72°. Two microliters of the amplified material was used for the second nested DNA


399

amplification using 30 cycles, as described above, with the second primer set. As positive control, a mix of the DNA from 10 positive samples was made and diluted fourfould to the limit of detection. Aliquots of the diluted DNA are stored at –20°. Five dilutions around the detection limit are tested in each PCR batch to control internal consistency. Distilled water is used as negative control in the first and the last position to control for false-positive reactions due to contamination. Inhibition of the amplification is controlled by a PCR assay in the same conditions with a primer set that amplifies a fragment of the HLA DQ genome. Aliquots of the reaction mixture are analysed by electrophoresis in a 2% agarose gel containing 0.03 Ìg/ml ethidium bromide and visualized under UV light. Quantitative PCR In the second part of the study, a selection of 21 patients (11 kidney, 1 kidney-pancreas, 1 kidney-heart, 7 liver and 1 heart) with a proven CMV infection had their plasma retrospectively tested by a quantitative PCR, the Cobas Amplicor CMV Monitor Test. The quantitative PCR was performed on 210 plasma samples from peripheral blood, whose leukocytes were found positive with our inhouse qualitative PCR. This quantitative PCR detects CMV DNA in plasma and leukocytes and is based on four major processes: specimen preparation; PCR amplification of target DNA using CMV-specific complementary primers; hybridization of the amplified DNA to oligonucleotide probes specific for the target(s), and detection of the probe-bound amplified DNA by colorimetric determination. The quantitation of CMV viral DNA is performed using the CMV Quantitation Standard. The CMV Quantitation Standard is non-infectious plasmid DNA that contains the same primer-binding sites as the CMV DNA target and a unique probe-binding region that allows Quantitation Standard Amplicon to be distinguished from CMV amplicon. The Quantitation Standard is incorporated into each individual specimen at a known copy number and is carried through the specimen preparation, PCR amplification, hybridization and detec-

Table 1. Incidence of CMV infection and disease according to

donor-receptor CMV serology

CMV infection CMV disease

D+/R–

D+/R+

D–/R+

D–/R–

9/14 (64) 8/14 (57)

6/8 (75) 2/8 (25)

9/18 (50) 4/18 (22)

3/21 (14) 1/21 (4)

Figures in parentheses are percentages.

Table 2. Sensitivity, specificity, PPV and

NPV of qualtitative PCR to symptomatic infection

tion steps along with the CMV target and is amplified together with the CMV target. The Cobas Amplicor Analyzer calculates the CMV DNA levels in the test specimens by comparing the CMV signal to the Quantitation Standard signal for each specimen. In our study, the Cobas Amplicor CMV Monitor Test was performed on plasma according to the manufacturer’s instructions.

Results

Diagnosis of CMV Infection and CMV Disease by Qualitative PCR A total of 684 blood samples were analyzed by qualitative PCR. Thirty percent of leukocyte samples were positive, compared to only 20% of plasma samples. CMV infection developed in 27/61 patients (44%). Twelve patients (44%) were classified as asymptomatic and 15 (56%) as symptomatic or with disease.The incidence of CMV infection and disease with regard to CMV serology of the donor and recipient is summarized in table 1. The sensitivity, specificity and predictive values of detecting CMV DNA in leukocytes by qualitative PCR for the diagnosis of CMV symptomatic infection is shown in table 2. The median time for detection of CMV DNA in patients who developed CMV disease was 26 days posttransplantation (range: 7–171 days) and CMV DNA was detected after a median time of 22 days (range: 15–87 days) for the patients who did not develop CMV disease. The delay of detection could be overestimated because of the timing of the sampling (once a week). Table 3 summarizes the median time of first detection of CMV DNA and the predictive values according to the donor-receptor serostatus. Retrospective Testing of Plasma Samples from Solid-Organ Transplant Recipients Using Quantitative PCR To assess the reproducibility of the Cobas Amplicor CMV Monitor Test, 9 clinical samples were retested in duplicate in different runs. Reproducible results were obtained with samples having a coefficient of variation smaller than 20%.

Assays

Sensitivity

Specificity

PPV

NPV

CMV DNA leukocytes CMV DNA plasma

15/15 (100) 14/15 (93)

33/46 (72) 37/46 (80)

15/28 (54) 14/23 (61)

33/33 (100) 37/38 (97)


400


Vanpoucke/Van Vlem/Vanholder/ Van Renterghem

Table 3. Median time of first detection of

CMV DNA, PPV and NPV of qualitative PCR for the diagnosis of CMV disease

Donorreceptor

Patients with CMV disease

Sample

Median time of first detection of CMV DNA, days

PPV

NPV

D+/R–

8/14

leukocytes plasma

34 (range: 16–171)

8/9 (89) 7/8 (88)

5/5 (100) 5/6 (83)

D+/R+

2/8

leukocytes plasma

18 (range: 11–24)

2/6 (33) 2/5 (40)

2/2 (100) 3/3 (100)

D–/R+

4/18

leukocytes plasma

23 (range: 7–26)

4/10 (40) 4/7 (57)

8/8 (100) 11/11 (100)

D–/R–

1/21

leukocytes plasma

26

2/3 (33) 2/3 (33)

18/18 (100) 18/18 (100)


A summary of donor-receptor serostatus for HCMV, type of transplantation, peak viral load and symptomatology of the 21 selected patients is shown in table 4. Of these 21 patients, 12 had a primary infection and 9 patients, had a reactivation or reinfection. For the 12 patients with a primary infection, the peak viral load ranged from 3,040 to 111,000 copies of CMV DNA/ml plasma with a median viral load of 28,600 copies CMV DNA/ml. For the 9 patients with a reactivation or reinfection, the peak viral load ranged from !400 to 14,100 copies CMV DNA/ml plasma with a median viral load of 4,260 copies CMV DNA/ml. Analysis of the viral load differences between the two groups showed that the viral loads in patients with a primary infection were significantly higher than viral loads in patients with a reactivation or reinfection (p ! 0.01; Wilcoxon test). Development of a symptomatic infection or disease associated with HCMV in patients having a primary infection was 12/12 (100%) and only 5/9 (56%) patients with reactivation or reinfection developed a symptomatic infection. Representative Cases of CMV Disease after Transplantation Case 1. Case 1 is a patient who underwent cadaveric renal transplantation (D+/R–). (fig. 1) The qualitative CMV-PCR turned positive in leukocytes and plasma 5 weeks after transplantation; the infection was later proven by a seroconversion in IgG and IgM titer. A few days later, when the patient complained of high fever, ganciclovir therapy was initiated for 12 days. In the beginning of the 9th week after transplantation, he developed a pulmonary infection, probably a CMV pneumonia. Therefore ganciclovir therapy was resumed, but after 10 days of therapy


Table 4. Summary of 21 selected patients for quantitative PCR

Patients

Graft

Symptomatic infection

Peak viral load copies CMV DNA/ml

D+/R– patients 1 liver 2 kidney 3 kidney 4 liver 5 kidney 6 kidney 7 kidney 8 kidney 9 heart 10 kidney + pancreas 11 liver

yes yes yes yes yes yes yes yes yes yes yes

3,870 20,100 43,500 111,000 3,040 105,000 9,140 17,700 88,400 55,600 33,300

D+/R+ patients 1 kidney 2 liver 3 kidney 4 liver 5 liver 6 liver 7 kidney

yes no yes no yes no yes

9,750 ! 400 2,290 1,420 4,260 14,100 2,480

D–/R+ patients 1 kidney 2 kidney

yes no

7,660 6,650

D–/R– patients 1 kidney + heart

yes

24,200


401

Fig. 1. Clinical course of case 1.

the qualitative PCR still remained positive. Foscarnet was given because of suspected resistance of the virus to ganciclovir. Looking at the results of the quantitative PCR, done retrospectively, we can observe that during the first period of ganciclovir therapy, therapy possibly was stopped too early; at that moment, the viral load was still relatively high. The peak of the viral load can be correlated with symptoms of pulmonary infection. However, there was a rapid decline in the viral load during the second period of ganciclovir therapy; foscavir would not have been started, were it not for the remaining positive results of the qualitative PCR in leukocytes and plasma. Case 2. Case 2 is a female patient who also underwent a cadaveric renal transplantation (D+/R+) (fig. 2). The first week after transplantation, we found a positive qualitative PCR in the leukocytes in correlation with mild clinical symptoms. The infection was proven by an increase in IgG and IgM titers. Antiviral therapy with ganciclovir was started immediately and remission of the symptoms was achieved; also the qualitative PCR in the leukocytes became negative. At the 4th week after transplantation, CMV DNA was found again in the leukocytes and in the plasma. However, the patient was feeling well and therefore no therapy was started. Looking at the results of the

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quantitative PCR, done retrospectively, we can observe that the quantitative PCR was negative at the first week after transplantation and that there is a small peak of CMV DNA after stopping the therapy. Altogether, these results indicate that compared to the quantitative PCR, the qualitative PCR detects the infection earlier and remains longer positive after treatment. Case 3. Case 3 is a female patient who underwent a liver transplantation (D+/R+) (fig. 3). The qualitative CMV-PCR turned positive in leukocytes 4 weeks after transplantation and the infection was proven by an increase in IgG and IgM titers. The patient did not complain of any clinical symptoms and therefore no therapy was started. The quantitative PCR remained negative over the whole period. This also correlates with the patient’s asymptomatic clinical condition.

Discussion

Since the time when Van Der Bij et al. [10] described a method for early detection of CMV replication for the first time in 1988, which enabled a rapid diagnosis of active infection to be made before the appearance of






403

symptomatic CMV infection in transplanted patients, interest in such an approach led to further developments in other techniques. Based on the literature and our own experience in molecular biology, we selected and evaluated a qualitative PCR assay and introduced the most sensitive assay in our hands in the weekly follow-up of transplanted patients. Using serology as the ‘gold standard’ for diagnosis of CMV infection, qualitative PCR on leukocytes has proven to be a very sensitive test; it has a negative predictive value (NPV) of 100%: a negative result excludes viral replication or infection. However, it has a low positive predictive value (PPV) for the diagnosis of CMV disease, except in the D+/R– subgroup where a primary infection evolves more often to symptomatic disease (table 3). Due to the high sensitivity, a qualitative PCR test is not a useful marker for HCMV disease. Also for monitoring antiviral therapy, the assay is not helpful as CMV DNA can persist for a long time in the leukocytes and often also in the plasma of treated patients. A quantitative PCR could provide a more accurate indication of clearance under a certain threshold, excluding the risk of CMV disease. The approach of early detection of CMV activity by qualitative PCR would allow clinicians to target highrisk patients for preemptive therapy. By monitoring the infected patients by a quantitative PCR (viral load) test, a quantitative threshold can be established beyond which antiviral therapy has to be started or may be stopped, sparing the risk of the possible toxicity of the drug, such as neutropenia and its consequences. Therefore in the second part of the study, we evaluated the Cobas Amplicor CMV Monitor test retrospectively.

The aim was to determine whether the follow-up of the viral load is of additional clinical value in monitoring disease and antiviral therapy in patients in our transplantation units. The good reproducibility of the test with an interassay variability of less than 20% allows a reliable detection of fourfould differences in viral loads between two consecutive samples from the same patient. A significantly higher peak load is observed in the group at highest risk (D+/R–), all of which also turned out to become symptomatic infections. In our study population it is not possible to establish a correlation between viral load and severity of symptoms, since in most cases antiviral prophylaxis could have reduced the viral load. Nevertheless, as is demonstrated in the few representative cases, we can assume that quantitative PCR is a better marker of viral activity, allowing a distinction to be made between a high load with a risk for disease, and residual viral DNA that can be controlled by the immune reactivity of the patient. The results, expressed in copies of viral DNA, allow clinicians to feel more comfortable in making their decision, especially regarding cessation of therapy. In conclusion, we can state that a qualitative PCR is a good marker for the initiation of preemptive therapy. In addition, viral quantitation is clinically useful for accurate diagnosis of established CMV disease, and monitoring of antiviral therapy. Acknowledgments This study was supported by Roche Molecular Systems, Inc., who provided the equipment and reagents.

References 1 van Son WJ, The TH: Cytomegalovirus infection after organ transplantation: An update with special emphasis on renal transplantation. Transplant Int 1989;2:147–164. 2 Gane E, Saliba F, Valdecasas G JC, O’Grady J, Pescovitz M, Lyman S, Robinson C: Randomised trial of efficacy and safety of oral ganciclovir in the prevention of cytomegalovirus disease in liver-transplant recipients. Lancet 1997; 350:1729–1733. 3 Gerna G, Zipeto D, Parea M, et al: Monitoring of human cytomegalovirus infections and ganciclovir treatment in heart transplant recipients by determination of viremia, antigenemia and DNAemia. J Infect Dis 1991;164:488–498. 4 Boeckh M, Boivin G: Quantitation of cytomegalovirus: Methodologic aspects and clinical applications. Clin Microbiol Rev 1998;11:533– 554.

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5 Kidd IM, Fox JC, Pillay D, Charman H, Griffiths PH, Emery VC: Provision of prognostic information in immunocompromised patients by routine application of the polymerase chain reaction for cytomegalovirus. Transplantation 1993;56:867–871. 6 Barber L, Egan J, Lomax J, Yonan N, Deiraniya A, Turner A, Woodcock A, Fox A: Comparative study of three PCR assays with antigenaemia and serology for the diagnosis of HCMV infection in thoracic transplant recipients. J Med Virol 1996;49:137–144. 7 van den Berg AP, van der Bij W, van Don WJ, et al: Cytomegalovirus antigenemia as a useful marker of symptomatic cytomegalovirus infection after renal transplantation: A report of 130 consecutive patients. Transplantation 1989;48: 991–995.


8 Cope AV, Sweny P, Sabin C, Rees L, Griffiths PD and Emery VC: Quantity of cytomegalovirus viruria is a major risk factor for cytomegalovirus disease after renal transplantation. J Med Virol 1997;52:200–205. 9 Razzaque A, Jahan N, Mc Weeney D, Jariwalla RJ,Jones C, Brady J, Rosenthal LJ: Location of DNA sequence analysis of the transforming domain (mtr II) of human cytomegalovirus. Proc Natl Acad Sci USA 1988;85:5709–5713. 10 Van Der Bij W, Torensma R, van Son WJ, Anema J, Schirm J, Tegzess AM, The TH: Rapid immunodiagnosis of active cytomegalovirus infection by monoclonal antibody staining of blood leukocytes. J Med Virol 1988;25:179– 188.




Viral Dynamics during Active Cytomegalovirus Infection and Pathology Vincent C. Emery Department of Virology, Royal Free and University College Medical School, London, UK

Key Words PCR W Quantitation W Kinetics W Diagnosis

Abstract The central role that cytomegalovirus (CMV) load plays in its pathogenesis is being unravelled. In AIDS patients with active CMV replication, many months prior to the development of CMV disease, elevated CMV load in the blood and urine are significantly associated with an increased risk of disease progression. In addition, elevated load in blood is associated with an increased risk of death. Intervention with ganciclovir acts to rapidly inhibit CMV replication in vivo and has allowed estimates of the clearance/replication rate of CMV to be performed. These data indicate that CMV replicates dynamically in the human host with a doubling time of approximately 1 day. This knowledge has been used to determine the relative contribution of initial viral load and rate of change of viral load as predictors of CMV disease in organ transplant recipients. The data show that both these parameters have prognostic value in multivariate models and should allow the development of novel patient management strategies. Copyright © 2000 S. Karger AG, Basel




Introduction

My group amongst others has been instrumental in showing that cytomegalovirus (CMV) load in blood is a major determinant of CMV organ pathology and explains many previously identified risk factors [1–6]. Thus, in a series of prospective studies, we have identified peak CMV load during active infection as a major risk factor that correlates with the development of CMV disease [1– 4]. Furthermore, using multivariate statistical methods, we showed that the classical risk factors of donor/recipient serostatus were entirely explained by the viral load parameter [1–4]. Consequently, donor/recipient serostatus provides prognostic information because it identifies subgroups of patients destined to experience high CMV loads after transplantation. However, while this observation helped to explain the pathogenesis of CMV, it did not have immediate practical applications for individual patients because the peak viral load often coincided with the development of CMV disease. CMV has generally been described as a slowly replicating virus. Using a series of approaches detailed later, we have shown that CMV replicates dynamically in the human host with a doubling time of approximately 1 day [7]. These findings have implications for our understanding of CMV pathogenesis and for therapeutic intervention.

Vincent C. Emery Department of Virology, Royal Free and University College Medical School Royal Free Campus, Rowland Hill Street London NW3 2QG (UK) Tel. +44 171 830 2997, Fax +44 171 830 2854, E-Mail [email protected]

a

b

On the basis of the aforementioned findings, we have continued the analysis of data from prospective studies of transplant recipients, focusing on the early events surrounding the detection of viraemia to determine whether measurement of CMV viral load in the earliest available samples and calculation of the kinetics of viral replication can provide prognostic information [8].

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Association between Baseline CMV Load and CMV Disease in AIDS Patients

A virologic substudy of ACTG204 comparing valaciclovir and aciclovir for the control of CMV disease in patients with advanced HIV disease has allowed the influence of baseline CMV load on disease progression to be assessed [6]. Overall, 259 patients had baseline blood

Emery

c Fig. 1. Kaplan-Meier plot showing progression to disease (a, b) or death (c) stratified according to baseline viral load in blood (a, c) or urine (b).

loads, of whom 53 had detectable levels, and 269 had baseline urine loads, of whom 113 had detectable levels. The effects of baseline viral loads in blood or urine on progression to CMV disease are shown in figures 1a, b. In both analyses, increasing CMV loads in blood and urine were associated with an increasing risk of disease development (p = 0.009 and p = 0.02 for blood and urine, respectively). In blood (fig. 1a), disease progression was more rapid in patients with baseline HCMV loads greater than 10,000 genomes/ml with 20% of patients in this viral load strata progressing to disease by day 120 compared to 616 days for the same proportion of patients with undetectable viral loads at baseline. Similar Kaplan-Meier analysis revealed that elevated baseline HCMV loads in blood were also significantly associated with a shorter time to death (p = 0.0001, fig. 1c). The time to reach 40% mortality was 224 days in patients with greater than 10,000 genomes/ml, compared with 642 days for patients with undetectable viral loads at baseline. Interestingly, there was no relationship between baseline urine load and survival (p = 0.51). The effects of treatment allocation, baseline CD4 count and baseline viral loads in blood and urine were all significant in univariate proportion hazards analyses (table 1). In multivariate analyses, baseline viral loads in

Table 1. Analysis of baseline risk factors for progression to CMV

CMV Replication and Pathology


disease in patients enrolled in the virological substudy of ACTG 204 Parameter

Relative risk

95% CI

p value

Univariate analyses Baseline HCMV load in blood Baseline HCMV load in urine VACV CD4

1.47 1.49 0.58 0.89

1.11–1.96 1.16–1.90 0.34–0.99 0.80–0.99

0.007 0.002 0.05 0.03

Multivariate analyses Baseline HCMV load in blood VACV CD4

1.49 0.53 0.89

1.10–2.02 0.28–0.99 0.79–1.00

0.01 0.05 0.05

Baseline HCMV load in urine VACV CD4

1.44 0.55 0.91

1.12–1.85 0.30–0.99 0.81–1.01

0.005 0.05 0.12

Baseline HCMV load in blood Baseline HCMV load in urine VACV CD4

1.49 1.46 0.51 0.93

1.09–2.04 1.11–1.92 0.26–0.97 0.82–1.05

0.01 0.008 0.04 0.24

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7 CMV load (genomes/ml) 6

log10 CMV load

5 4 3 2 1 0

Fig. 2. Viral load-time plot of CMV load in

0

an AIDS retinitis patient following initiation of ganciclovir (GCV) therapy.

both blood and urine remained statistically significant after adjusting for treatment allocation, CD4 count and each other, with 1 log increases in baseline blood and urine loads being associated with 49 and 46% increases in the risk of disease progression, respectively. Treatment allocation remained significantly associated with disease progression after adjusting for baseline viral loads, although the effect of baseline CD4 count became non-significant.

Dynamics of CMV Replication in vivo

Using basic models for the viral dynamics of CMV [7] based upon those described for HIV and HCV [9–11], the decline of virus in the blood following ganciclovir therapy can be used to determine the clearance rate of CMVinfected cells. If the system is in equilibrium at the start of therapy with the rate of viral production equal to the rate of viral clearance, then after therapy with a drug that totally blocks viral production, the dynamics of the system follows the function y(0)e –at, where y(0) is the initial viral load and a the clearance rate constant. Plotting the change in CMV load with time followed by computation of the slope of decline after initiation of ganciclovir therapy allows the half-life of virus in the blood to be calculated according to the formula T½ = –ln2/slope. Analysis of the decline in CMV load following ganciclovir therapy of 5 AIDS retinitis patients with frequent

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5

10

15

20

25

Days of GCV

samples (median 5 samples over 21 days) allowed the half-life of decline to be calculated as 0.98 B 0.3 days in blood and 0.96 B 0.14 days in urine. A representative viral load decline pattern for one of these patients is shown in figure 2. The rapidity of this clearance indicates that CMV replication in the host is occurring rapidly and the virus exists in a dynamic equilibrium.

Application in Viral Load Kinetics to Identify Patients Developing CMV Disease after Transplantation

Since baseline CMV load in HIV-infected patients is important in the pathologic consequences of CMV infection (vide supra) and CMV replication in vivo occurs rapidly in the human host (approximate doubling time of 1 day), we were interested to ascertain whether these parameters would help explain why some individuals suffer CMV disease following transplantation. We have analysed 3,873 blood samples derived from prospective studies of 359 transplant recipients (renal, bone marrow and liver recipients) by PCR for CMV and identified 127 patients with DNAemia. CMV load in the samples was quantitated and the data used to calculate the rate of initial viral load increase by dividing the difference between the first quantifiable load and the preceding CMV-PCRnegative result (ascribed a value of 2.3 log10 genomes/ml blood) by the number of days between the two results.

Emery

Table 2. Logistic regression analysis to define the risk of CMV dis-

ease associated with a 0.25 log10 genomes/ml higher initial CMV load Patient group

Odds ratio 95% CI

p value

Liver transplant Renal transplant Bone marrow transplant

1.821 1.34 1.52

0.02 0.01 0.006

1.11–2.98 1.07–1.68 1.13–2.05

1

After adjustment for receipt of methylprednisolone, which remained significantly associated with disease in a multivariate model (odds ratio was 1.47 per each 1 g increase in methylprednisolone; 95% CI; 1.09–1.91; p = 0.01).

These data allowed comparison of the initial viral load with peak viral load attained, and the relative risk of the rate of increase in CMV load and its initial value as prognostic markers in univariate and multivariate statistical models. Fifty-five individuals (14 renal, 20 liver and 21 bone marrow transplant recipients) had distinct initial and peak viral loads (difference between initial and peak viral loads = 0.48 log10 genomes/ml; range 0.2–2.92 log10 genomes/ml). A strong relationship between initial and peak CMV loads (p = 0.0001) was present in these individuals, indicating that initial viral load levels were related to the ultimate peak viral load which we have previously shown to be associated with CMV disease [1–4]. This relationship remained highly significant (p = 0.0001) after adjusting for the number of days between the two measurements. The relationships between the development of CMV disease, initial CMV load, and other recognised risk factors for disease (including donor/recipient serostatus and receipt of augmented immunosuppression) were assessed using logistic regression analyses (table 2). For each patient group, initial viral load was the dominant risk factor for the development of disease. Among liver transplant recipients, the receipt of augmented methylprednisolone following liver transplantation was an additional risk factor for disease, both before and after adjusting for initial CMV load (adjusted odds ratio for methylprednisolone administration: 1.47; confidence interval 1.09–1.99; p = 0.01). Among renal transplant recipients, donor/recipient serostatus and receipt of antithymocyte globulin were of borderline significance (p = 0.04 and p = 0.05, respectively) in univariate analyses; however these became non-significant after adjustment for initial viral load. Similarly,

among bone marrow transplant recipients, the marginally significant relationship between donor/recipient serostatus and disease (p = 0.05) seen in a univariate model became non-significant after adjusting for initial CMV load. In the whole group of patients, the median time interval between the last CMV-PCR-negative result and initial CMV load was 7 days (range: 2–14 days) and was similar between the three patient groups (p = 0.14, Kruskal-Wallis). On average, viral loads increased at a rate of 0.24 log10 genomes/ml/day (range: 0.03–1.65 log10 genomes/ ml/day). The rate of increase was significantly correlated with the initial CMV load (Spearman’s r = 0.79, p = 0.0001) and the peak CMV load (Spearman’s r = 0.67, p = 0.0001). The rate of increase was also significantly higher in patients who developed CMV disease (0.33 [0.10–1.65] log10 genomes/ml/day) than in those who remained asymptomatic (0.19 [0.03–0.51] log10 genomes/ml/day, p = 0.0001, Wilcoxon, fig. 3).



Fig. 3. Rate of CMV load increase at the initial phase of active infec-

tion stratified according to whether the patients experienced CMV disease or remained asymptomatic.

409

Table 3. Univariate and multivariate logistic regression analyses

relating parameters of initial CMV load (per 0.25 log10 genomes/ml increase), peak CMV load (per 0.25 log10 genomes/ml increase) and rate of change in CMV load (per 0.1 log10 genomes/ml/day increase) to the development of CMV disease Parameter

Odds ratio 95% CI

p value

A. Univariate Initial CMV load Peak CMV load Rate of change of CMV load

1.39 1.55 2.04

1.22–1.58 1.34–1.81 1.44–2.88

0.0001 0.0001 0.0001

B. Multivariate model 1 Initial CMV load Rate of change in CMV load

1.28 1.52

1.06–1.53 1.06–2.17

0.01 0.02

C. Multivariate model 2 Peak CMV load Rate of change in CMV load

1.58 1.45

1.28–1.95 1.01–2.09

0.0001 0.04

D. Multivariate model 3 Initial CMV load Peak CMV load Rate of change in CMV load

0.77 1.92 1.65

0.56–1.05 1.37–2.68 1.07–2.52

0.1 0.0001 0.02

The results from univariate and multivariate logistic regression models incorporating the rate of increase in viral load, along with both the initial and peak CMV loads, are shown in table 3 for the whole group of patients. In the univariate analysis (table 2, panel A), each 0.1 log10 genomes/ml/day higher rate of CMV load increase was associated with an increased risk of CMV disease of 2.04 (p = 0.0001). The rate of viral load increase remained significantly associated with the development of disease after controlling in multivariate models (table 2, panels B, C) for either the initial or peak CMV loads, suggesting that knowledge of viral replication kinetics provides additional information about patient prognosis to that provided by either the initial or peak CMV load on its own. In models which inlcuded both the initial and peak CMV loads, the rate of viral load increase still remained significantly associated with the development of disease (p = 0.02), although the initial CMV load became non-significant (table 2; panel D).

associated with increased risks of CMV disease and death [6, 12]. Since CMV replication occurs over extensive time periods in AIDS patients prior to the development of disease [6, 12, 13], it appears that the viral load present many months prior to evidence of pathology encompasses much of the information dictating future events. This situation is reminiscent of the prognostic significance of HIV-1 plasma load following seroconversion [14]. In contrast to AIDS patients, CMV replication in transplant recipients usually occurs over relatively short time periods. Since peak viral loads are known to correlate with pathology and to be elevated in symptomatic individuals [1–4], the possibility that the replication dynamics of CMV in transplant recipients at early time points provides insight into future pathology was investigated. The conclusions of these studies show that both initial CMV load during the transition between PCR negativity to positivity and the rate of change in viral load are independent risk factors for disease after transplantation. These data will require refinement, especially with respect to the influence of the immune system, and basal and augmented immunosuppression on the rate of CMV load increase and through more frequent sampling in the very early phases of active infection. Nevertheless, such observations will facilitate the development of earlier pre-emptive therapeutic interventions and aid in the assessment of new anti-CMV agents in controlled clinical trials.

Acknowledgments I am grateful to my colleagues and collaborators who have been instrumental in providing the data summarised in this paper (Prof. P.D. Griffiths, Dr. E.F. Bowen, Dr. A.C. Hassan-Walker, Dr. A. Cope, Dr. M.A. Johnson, Dr. J.E. Feinberg, University of Cincinnati). Work in my laboratory is supported by the UK Medical Research Council, the Wellcome Trust, the National Institutes of Health, USA, the European Community (Biomed 2) and Glaxo-Wellcome.

Conclusions and Future Prospects

The central role of CMV load in the pathogenesis of CMV is now established. In AIDS patients, baseline elevated CMV load in whole blood and in plasma have been

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Emery

References 1 Cope AV, Sweny P, Sabin C, Rees L, Griffiths PD, Emery VC: Quantity of cytomegalovirus viruria is a major risk factor for cytomegalovirus disease after transplantation. J Med Virol 1997;52:200–205. 2 Cope AV, Sabin C, Burroughs A, Rolles K, Griffiths PD, Emery VC: Interrelationships among quantity of human cytomegalovirus (HCMV) DNA in blood, donor-recipient serostatus, and administration of methylprednisolone as riks factors for HCMV disease following liver transplantation. J Infect Dis 1997;176: 1484–1490. 3 Gor D, Sabin C, Prentice HG, et al: Longitudinal fluctuations between peak virus load, donor/recipient serostatus, acute GvHD and CMV disease. Bone Marrow Transplant 1998; 21:597–605. 4 Hassan-Walker AF, Kidd IM, Sabin C, Sweny P, Griffiths PD, Emery VC: Quantity of human cytomegalovirus (CMV) DNAemia as a risk factor for CMV disease in renal allograft recipients: Relationship with donor/recipient CMV serostatus, receipt of augmented methylprednisolone and anti-thymocyte globulin (ATG). J Med Virol 1999;58:182–187.


5 Bowen EF, Wilson P, Cope A, et al: Cytomegalovirus retinitis in AIDS patients: Influence of cytomegalovirus load on response to ganciclovir, time to recurrence and survival. AIDS 1996;10:1515–1520. 6 Emery VC, Sabin CA, Feinberg JE, Grywacz M, Knight S, Griffiths PD: Quantitative effects of valaciclovir on the replication of cytomegalovirus (CMV) in persons with advanced human immunodeficiency virus disease: Baseline CMV load dictates time to disease and survival. J Infect Dis 1999;180:695–701. 7 Emery VC, Cope AV, Bowen EF, Gor D, Griffiths PD: The dynamics of human cytomegalovirus replication in vivo. J Exp Med 1999;190: 177–182. 8 Emery VC, Sabin CA, Cope AV, Gor D, Hassan-Walker AF, Griffiths PD: Application of viral load kinetics to identify patients destined to develop cytomegalovirus disease following transplantation, submitted.

9 Ho DD, Neuman AU, Perelson AS, Chen W, Leonard JM, Markowitz M: Rapid turnover of plasma virions and CD4 lymphocytes in HIV-1 infection. Nature 1995;373:123–126. 10 Bonhoeffer S, May RM, Shaw GM, Nowak MA: Virus dynamics and drug therapy. Proc Natl Acad Sci USA 1997;94:6971–6976. 11 Neuman AU, Lam NP, Dahari H, Gretch DR, Wiley TE, Layden TJ, Perelson AS: Hepatitis C viral dynamics in vivo and the antiviral efficacy of interferon-alpha therapy. Science 1998; 282:103–107. 12 Spector SA, Wong R, Hsia K, Pilcher M, Stempien MJ: Plasma cytomegalovirus (CMV) DNA load predicts CMV disease and survival in AIDS patients. J Clin Invest 1998;101:497– 502. 13 Bowen EF, Sabin CA, Wilson P, et al: Cytomegalovirus (CMV) viraemia detected by polymerase chain reaction identifies a group of HIV-positive patients at high risk of CMV disease. AIDS 1997;11:889–893. 14 Mellors JW, Rinaldo CR, Gupta P, White RM, Todd JA, Kingsley LA: Prognosis in HIV-1 infection predicted by the quantity of virus in plasma. Science 1996;272:1167–1170.


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Part II. Diagnostics and Antiviral Therapy Antivirals


Inhibition of Cytomegalovirus in vitro and in vivo by the Experimental Immunosuppressive Agent Leflunomide W. James Waldman a Deborah A. Knight a Leonard Blinder b JiKun Shen b Nell S. Lurain c Daniel M. Miller a Daniel D. Sedmak a James W. Williams b Anita S.-F. Chong b, c a Department

of Pathology, The Ohio State University College of Medicine and Public Health, Columbus, Ohio, of General Surgery, Rush Presbyterian-St. Luke’s Medical Center, and c Department of Microbiology/Immunology, Rush University, Chicago, Ill., USA b Department

Key Words Cytomegalovirus W Leflunomide W Antiviral therapy W Endothelial cells W Fibroblasts

rently attenuate a major complication of immunosuppression, CMV disease, by a novel mechanism of antiviral activity. Copyright © 2000 S. Karger AG, Basel

Abstract Despite progress in antiviral chemotherapy, cytomegalovirus (CMV) remains a major cause of morbidity and mortality among pharmacologically immunosuppressed transplant recipients, frequently engaging the clinician in a struggle to balance graft preservation with control of CMV disease. Leflunomide, an inhibitor of protein kinase activity and pyrimidine synthesis, is an experimental immunosuppressive agent effective against acute and chronic rejection in animal models. Herein we summarize our recent studies demonstrating that leflunomide inhibits the production of multiple clinical CMV isolates (including multi-drug-resistant virus) in both human fibroblasts and endothelial cells. In contrast to all other anti-CMV drugs currently in use, leflunomide does not inhibit viral DNA synthesis, but rather appears to interfere with virion assembly. Finally, preliminary studies in a rat model suggest that this agent reduces viral load in vivo. These findings imply that leflunomide, an effective immunosuppressive agent, shows potential to concur-




Introduction

Despite widespread prevalence, cytomegalovirus (CMV) infection is rarely of significant consequence in the healthy individual with a competent immune system. However, the progression of the AIDS epidemic and experience with the rapidly growing population of transplant recipients emphasize that this ß-herpesvirus can be clinically problematic among immunosuppressed populations. HIV-induced erosion of cellular immunity promotes reactivation of endogenous latent CMV in AIDS patients and renders them highly susceptible to symptomatic primary infection. Pharmacologic immunosuppression required for preservation of allograft integrity similarly disables host defenses in transplant recipients. Thus, despite refinements in therapeutic intervention in these patients, CMV remains a source of a diverse constellation of serious, often life-threatening complications, including interstitial pneumonitis, diffuse gastrointestinal mucosal

W. James Waldman, PhD Department of Pathology, The Ohio State University College of Medicine and Public Health 166 Hamilton Hall, 1645 Neil Avenue Columbus, OH 43210-1218 (USA) Tel. +1 614 292 7772, Fax +1 614 292 7072, E-Mail [email protected]

cellular production of infectious virus. Thus we tested the hypothesis that leflunomide might exert inhibitory activity against CMV in human fibroblasts and endothelial cells, common targets of CMV infection in vivo [18, 19]. As summarized below, we have shown that this is indeed the case, and have further demonstrated the mechanism of viral inhibition by leflunomide to be unique among currently approved anti-CMV therapeutic agents. Finally, preliminary experiments in a rat model suggest that this agent is capable of reducing viral load in vivo.

ulceration, hepatitis, and retinitis, as well as destructive inflammatory lesions in a variety of other locations [1]. In addition, several studies have suggested a role for this virus as a contributing factor in allograft rejection [2, 3]. Currently, three compounds, ganciclovir (GVC), foscarnet (PFA) and cidofovir, are approved for clinical use in the control of CMV disease, none of which are free of toxic side effects. Drug-associated myelosuppression, metabolic toxicity, and nephrotoxicity limit the use of these agents in a significant number of patients [4–6]. Although specific mechanisms of action vary among these compounds, all three ultimately act by inhibition of viral DNA synthesis, either by impeding chain elongation following incorporation into nascent strands [4, 7], or by direct binding to viral DNA polymerase [6]. It is not surprising, therefore, that clinical strains of CMV have emerged which exhibit resistance to one or more of these drugs [8–10]. Clearly there remains a need for development of additional therapeutic approaches. Leflunomide [N-(4-trifluoromethylphenyl)-methylisoxazol-4-carboxamide; HWA 486] is an experimental immunosuppressive agent with demonstrated effectiveness in the prevention and reversal of acute allograft rejection in rats and dogs [11, 12], as well as in the reversal of chronic rejection when combined with cyclosporine [13]. It has also been shown to be effective against rat and murine graft-versus-host disease and against autoimmune disease in animal models [14]. Leflunomide has recently been approved for treatment of rheumatoid arthritis, and has been well tolerated in current phase I clinical trials in human transplant recipients [Dr. James W. Williams, Rush presbyterian St. Luke’s Medical Center, Chicago, Ill. pers. commun.]. This isoxazol derivative is metabolized to its active form, A77 1726 [N-(4-trifluoromethylphenyl)-2-cyano-3-hydroxycrotoamide], which is structurally unique among established immunosuppressive agents [14]. This metabolite exhibits two known mechanisms of action: inhibition of protein tyrosine kinase activity [15], and inhibition of dihydroorotate dehydrogenase, a key enzyme in the biosynthesis of pyrimidine nucleotide triphosphates (pyNTP) [16]. However, neither the complete spectrum of kinases targeted by leflunomide nor the specific contribution of each of these functions to its immunosuppressive activity in vivo has been fully resolved. Furthermore, additional mechanisms of activity may yet remain to be discovered. Based upon the fact that a number of CMV-encoded proteins are phosphoproteins, and that several CMV proteins themselves possess kinase activity [17], we speculated that protein kinase activity may be essential in intra-

Effect of Leflunomide upon the Production of Infectious CMV in vitro Our first evidene for an antiviral effect of leflunomide came from microscopic observation of human umbilical vein endothelial cell (HUVEC) monolayers inoculated at low titer with CMV VHL/E, a clinical isolate propagated in HUVEC to preserve its natural endothelial cytopathogenicity [20], and incubated in the presence or absence of A77 1726 (200 ÌM), the active metabolite of leflunomide [14]. This isolate induces obvious cytopathic change and, under normal circumstances, disseminates by direct cellto-cell transmission generating widespread cytopathology over a period of several days. In the presence of A77 1726, however, this restriction was dramatically restricted [21]. Similar patterns of dissemination and A77 1726-mediated restriction were evident in human fibroblast (HFF) monolayers inoculated with clinical isolate P8 (isolated from bronchial alveolar lavage obtained from a cardiac allograft recipient and propagated in HFF cells [10]). To quantitatively assess the impact of leflunomide upon the production of infectious virus in HUVEC and HFF, we performed standard plaque reduction and virus yield assays in the presence of A77 1726 over a range of concentrations equivalent to those which have been shown to attenuate immune activation by various stimuli in vitro [22], and equivalent to (or less than) well-tolerated serum levels measured in leflunomide-treated experimental animals [23] and human clinical trial subjects (270–450 ÌM) [Dr. James W. Williams, unpubl. data]. Data generated by these experiments demonstrated an A77 1726-mediated, dose-dependent reduction in infectious virus production in both cell types. The doseresponse curves of clinical isolate P8 in HFF and of clinical isolate VHL/E in HUVEC did not differ significantly, both indicating an IC50 of 40–60 ÌM [21]. Similar assays performed with CMV VHL/E and an additional clinical

Inhibition of CMV by Leflunomide


Methods and Results

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isolate, BUR/E [24], in the presence of cyclosporine A (CsA) or tacrolimus (FK 506), verified that neither of these commonly prescribed immunosuppressive agents possesses antiviral activity against CMV [21]. Effect of Exogenous Uridine upon the Antiviral Activity of Leflunomide Two mechanisms have been identified for the activity of A77 1726: inhibition of protein tyrosine kinase activity [15], and inhibition of dihydroorotate dehydrogenase [16], a key enzyme in the biosynthesis of pyNTP. To determine whether reduction of intracellular pyNTP pools was responsible for the antiviral effects of leflunomide, HUVEC were inoculated with CMV VHL/E, then incubated for 4 days in the presence or absence of 200 ÌM A77 1726, 200 ÌM exogenous uridine, or both. Uninfected HUVEC were included in each experiment as negative controls. Following harvest, cells were extracted by methods described by Khym [25] and intracellular NTP levels were analyzed by high-performance liquid chromatography. Small aliquots of cells from each group were reserved prior to extraction and assayed for virus yield by plaque assay. Data generated by these experiments demonstrated several trends, none of which reached statistical significance as determined by ANOVA. First, intracellular pyNTP levels were slightly reduced in untreated CMVinfected HUVEC as compared to uninfected control cells, but were not further reduced by A77 1726 treatment. Second, the addition of exogenous uridine increased pyNTP levels both in the presence and absence of A77 1726. Most importantly, however, the addition of exogenous uridine did not significantly reconstitute infectious virus production in A77 1726-treated, CMV-infected HUVEC [21]. Thus A77 1726-mediated inhibition of CMV activity appears to be independent of the inhibitory effects of this agent upon pyNTP synthesis. Effects of Leflunomide upon CMV Gene Transcription and Protein Expression We next sought to determine at which point in the viral replication cycle leflunomide exerts its inhibitory effects. Immediately upon viral entry of the host cell, CMV lower matrix protein pp65, a component of the viral tegument, translocates to the cell nucleus. Although the nuclear function of pp65 remains to be resolved, another tegument protein, pp71, which migrates in a similar manner, acts in concert with cellular proteins to promote rapid transcription of immediate early (IE) viral genes. IE gene products are primarily regulatory, activating early gene transcrip-

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tion, whose products are essential for viral DNA replication. Activation of late genes, which code primarily for structural protein components of the virion, occurs late in the replicative cycle and is dependent upon viral DNA replication [17]. To localize potential A77 1726-induced lesions in this temporal series of events, we first visualized a cross-section of CMV proteins in A77 1726-treated CMV-infected HUVEC by immunohistochemical staining with monoclonal antibody (mAb) specific for pp65, 72 kD IE1, or late structural glycoprotein B (gB). Staining patterns indicated that none of these viral proteins were prevented from being expressed in A77 1726-treated cells. Nuclear accumulation of pp65 was apparent in both untreated and A77 1726-treated monolayers within hours after inoculation, implying that leflunomide interferes neither with viral entry, nor with nuclear translocation of pp65. Nuclear IE1 expression also appeared to be unaffected by A77 1726. Finally, typical gB staining patterns were apparent at 72–96 h after inoculation in both treated and untreated cultures. In contrast, infected cells treated with PFA, an inhibitor of CMV DNA polymerase, were unable to express gB, while pp65 and IE1 expression in these cultures was unimpeded [21]. We next employed Northern blot analysis to determine whether A77 1726 quantitatively affected IE1 or gB transcription. Total cytoplasmic RNA was isolated from CMV-inoculated HUVEC incubated for 48 h in the absence or presence of 200 ÌM A77 1726, 1 mM PFA, or 1.2 mM ganciclovir (GCV). Each experiment also included RNA isolated from uninfected HUVEC (as negative control). Electrophoretically fractionated RNA was transferred to Nylon membranes and hybridized to [32P]labeled probes specific for CMV IE1, CMV gB, or cellular GAPDH (as a loading control), and bands were visualized by autoradiography. The results demonstrated that, in contrast to PFA and GCV which completely suppressed gB transcription, A77 1726 had no apparent effect upon the transcription of either IE1 or gB [21]. Effect of Leflunomide upon CMV DNA Synthesis Since expression of late structural proteins (such as gB) is dependent upon CMV DNA replication [17], our immunohistochemical and Northern blot analyses suggested that, in contrast to currently used anti-CMV therapeutics, leflunomide does not inhibit viral DNA synthesis. The potential novelty of such an alternative antiviral mechanism led us to employ three independent experimental approaches to test this hypothesis. First we measured the incorporation of [3H]thymidine in HUVEC or HFF

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which had been 100% infected with CMV VHL/E or P8, respectively, and incubated for 36 h in the presence or absence of various concentrations of A77 1726. While PFA inhibited radiolabel incorporation by CMV-infected cells in a dose-dependent manner, no inhibition was detected in the presence of A77 1726 [21]. We next generated dot blots of serially diluted DNA extracted from P8-infected HFF or VHL/E-infected HUVEC which had been incubated for 48 h after inoculation in the presence or absence of 100 ÌM (HFF) or 200 ÌM (HUVEC) A77 1726, or 1 mM PFA. Blots were hybridized to [32P]-labeled CMV-specific cDNA probe and visualized by autoradiography. These studies demonstrated that quantities of viral DNA synthesized in CMVinfected cells incubated in the presence of A77 1726 were approximately equivalent to those accumulated in untreated infected cells. In contrast, PFA-treated cells contained no detectable CMV DNA. The specificity of the probe was verified by the absence of hybridization to DNA extracted from uninfected cells [21]. Finally to directly test the effect of leflunomide upon viral DNA polymerase, crude protein extracts were prepared from CMV-infected (or uninfected) HFF or HUVEC, and viral DNA polymerase activity was measured in the presence of various concentrations of A77 1726 by biochemical assay of enzyme-catalyzed incorporation of [3H]TTP into nicked template DNA. While PFA reduced viral DNA polymerase activity in a concentration-dependent manner, A77 1726 showed no detectable inhibitory activity even at concentrations which dramatically reduced plaque formation. Experiments performed with extracts prepared from CMV VHL/E or BUR/E-infected HUVEC, or P8-infected fibroblasts generated essentially identical results. No enzyme activity was detected in extracts prepared from uninfected cells [21].

demonstrated typical herpesvirus capsids within the nuclei of A77 1726-treated cells, implying that neither nucleocapsid assembly nor viral DNA packaging is affected by this agent. However, we observed profound differences in the morphology of virions maturing in the cytoplasm. While tegument and external membrane were acquired normally in untreated cells, viral particles appeared not to mature beyond the 100 nm naked capsid stage in the presence of A77 1726 [21]. Activity of Leflunomide against Multi-Drug-Resistant CMV All currently approved anti-CMV chemotherapies focus upon inhibition of viral DNA synthesis, although specific mechanisms vary among different agents [4, 6, 7] (see Discussion). Thus it is not surprising that multi-resistant clinical strains have emerged [8–10]. Based upon our discovery of the apparently unique mechanism of viral inhibition by leflunomide, we tested the hypothesis that this agent might suppress activity of drug-resistant virus. CMV strain D16 was isolated from the same patient as strain P8. However, unlike P8, D16 exhibits resistance to GCV, PFA, and cidofovir [10]. Plaque reduction assays performed in HFF cultures revealed equivalent sensitivity of these two isolates to A77 1726 (IC50 F40–60 ÌM) [21]. Thus leflunomide-mediated inhibition of production of D16 is independent of resistance to current clinically applied chemotherapeutic agents.

Effect of Leflunomide upon Virion Morphology Collectively, results of our immunohistochemical staining, Northern blot analysis, viral DNA blots, and polymerase assays argue in favor of a leflunomide-associated antiviral mechanism which is unique with respect to other anti-herpesvirus compounds. Since three of the five major CMV tegument proteins are phosphoproteins [26], we hypothesized that leflunomide-mediated inhibition of protein phosphorylation might disable processes essential in the maturation and assembly of the complete viral particle. To address this issue, we employed transmission electron microscopy to directly examine virion morphology within A77 1726-treated or untreated CMV-infected cells at 4–7 days after inoculation. Electron micrographs

Antiviral Activity of Leflunomide in vivo To determine the effectiveness of leflunomide in the control of viral load in vivo, groups of immunodeficient nude rats were inoculated with rat CMV (RCMV Maastricht strain [27], 105 plaque-forming units/animal) and treated with either leflunomide (15 mg/kg/day for 14 days), GCV (10 mg/kg/day for 5 days), or drug-free vehicle. Experiments also included uninfected control animals. Following euthanization at 14 days after inoculation, viral infection was confirmed by histologic observation of typical cytomegalic ductal epithelium in salivary glands, and further verified by immunohistochemical detection of RCMV antigens in multiple tissues. Plaque assay of tissue homogenates prepared from salivary gland, spleen, and lung demonstrated 75–99% reduction in virus yield in organs harvested from leflunomide-treated animals, and 85–99% reduction in those harvested from ganciclovir-treated animals. No virus was ever recovered from uninfected control rats [28]. Thus in addition to its in vitro antiviral activity, leflunomide is capable of reducing viral load in vivo.



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Discussion

Despite substantial progress in antiviral chemotherapy and prophylaxis, CMV infection remains a significant complicating factor in the clinical course of up to 50% of organ transplant recipients [1]. Thus, fine tuning of pharmacologic immunosuppression to maintain a balance between graft preservation and control of CMV disease presents a persistent problem for the clinician. Results of the studies summarized herein suggest that leflunomide may help to alleviate this dilemma. Specifically, we have demonstrated that A77 1726, the active metabolite of leflunomide, inhibits infectious CMV production in a dose-dependent manner over a range of concentrations similar to those which inhibit in vitro T cell activation by mitogenic or allogeneic stimuli [22], and equivalent to (or less than) well-tolerated serum levels measured in leflunomide-treated experimental animals [23] and human clinical trial subjects [Dr. James W. Williams, unpubl. data]. Similar patterns of viral inhibition were observed for each of 4 individual clinical CMV isolates, and in both human fibroblasts and human endothelial cells, common targets of CMV infection in vivo [18, 19]. Furthermore no such inhbitory activity was observed for either CsA or FK 506 [21]. Which of the two known mechanisms of leflunomide, inhibition of pyrimidine synthesis or inhibition of protein tyrosine kinase activity, contributes most significantly to its immunosuppressive properties remains to be resolved. However, our experiments clearly show that the antiviral activity of this agent cannot be attributed to a reduction in intracellular pyNTP pools [21]. While previous studies have demonstrated profound pyNTP depletion in IL-2stimulated murine T cells [29] and murine leukemia cells [30] cultured in the presence of A77 1726, we observed little if any A77 1726-associated CTP or UTP reduction in CMV-infected endothelial cells. Although these outcomes might seem at first contradictory, it should be noted that in those previous studies the murine cells were vigorously proliferating prior to leflunomide treatment. In contrast, HUVEC assayed in our experiments were in a stationary, quiescent phase. Thus, with the exception of viral DNA replication, minimal intracellular NTP consumption would be expected. We note that, although addition of exogenous uridine increased intracellular pyNTP concentrations both in the presence and absence of A77 1726, it did not significantly reconstitute infectious virus production in A77 1726-treated, CMV-infected cells, nor did it correct A77 1726-associated defects in viral particle asssembly [21]. These data argue against

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the hypothesis that pyNTP depletion is responsible for the antiviral activity of leflunomide. The results of our experiments directed toward identification of specific viral processes targeted by leflunomide suggest mechanisms that are unique among anti-CMV therapeutic agents in use at present. GCV, currently the drug of choice for the treatment of CMV disease, is a guanosine analogue which is monophosphorylated in infected cells by the protein product of the CMV UL97 gene [31, 32]. Cellular kinases convert the monophosphate form to a triphosphate which is then incorporated by CMV DNA polymerase into the replicating viral DNA where its presence inhibits chain elongation [4]. PFA has been used increasingly as an alternative to GCV, in particular when GCV resistance is suspected [6]. This agent also inhibits CMV DNA replication but does so by binding directly to the viral DNA polymerase [6]. Cidofovir, the newest addition to the anti-CMV armamentarium, recently approved for use in the control of CMV retinitis [33], is a monophosphorylated cytosine analogue which, like GCV, is further phosphorylated by cellular kinases, ultimately inhibiting viral DNA synthesis following incorporation into nascent strands [7]. In contrast to these agents (as well as several others with similar activities against other herpesviruses), our studies indicate that leflunomide does not inhibit CMV DNA replication [21]. This was initially suggested by immunohistochemical staining and Northern blot analysis demonstrating that A77 1726 had no effect upon expression of CMV late structural protein gB or transcription of its mRNA, both of which are dependent upon viral DNA synthesis. Subsequent measurement of [3H]thymidine incorporation by infected cells, viral DNA dot blot analysis, and biochemical assay of viral DNA polymerase activity, provided proof that indeed this agent exerts no inhibitory effect upon the accumulation of viral DNA or upon the activity of the enzyme. Rather, the major A77 1726-mediated lesion in infectious virus production appears to be in the process of maturation and assembly of the complete virion, specifically (as demonstrated by transmission electron microscopy) failure to acquire tegument and external membrane in the cytoplasm of infected cells [21]. Although little is presently known of the structural organization of the CMV tegument, three of the five major tegument proteins are phosphoproteins (pp150, pp71, pp65) [26]. Thus we speculate that by inhibiting protein phosphorylation, leflunomide may interfere with tegument assembly. Experiments are currently in progress to test this hypothesis and to identify specific proteins whose phosphorylation is inhibited by A77 1726. Out-

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comes of these studies may implicate this compound as a useful reagent in further investigations of mechanisms of virion maturation. The emergence of drug-resistant variants of CMV in response to antiviral agents increasingly complicates treatment. Resistance to GCV is primarily due to mutation of the CMV UL97 gene which codes for the phosphotransferase responsible for the initial monophosphorylation of this compound [31, 32]. Several different mutations have been reported, all of which result in single amino acid substitutions or short deletions in the UL97 protein product [8]. In addition in the viral DNA polymerase gene (UL54) resulting in single amino acid substitutions have also been detected in GCV-resistant isolates [8]. Unlike GCV, PFA requires no anabolic processing to enable its binding to viral DNA polymerase [6]. Thus resistance to PFA appears to result exclusively from UL54 mutations [6, 8]. Similarly, the molecular basis for cidofovir resistance has thus far been traced exclusively to UL54 mutations [9]. While the mechanisms of action of these agents vary in detail, all three ultimately target viral DNA polymerization as a common endpoint. Moreover, it is not surprising that CMV variants which develop resistance to one of these drugs can often exhibit resistance to others [8–10]. It is also not surprising that leflunomide, as a consequence of its distinctly different mechanism of action, can exert inhibitory activity against drug-resistant CMV equal to that against drug-sensitive isolates. Leflunomide, recently approved for the treatment of rheumatoid arthritis, has been shown to be effective in the prevention and reversal of acute and chronic allograft rejection in several animal transplantation models [11– 13]. The reproducible success of those animal experiments and the documented low toxicity of this agent [34] have led to its current status as a candidate immunosuppressant in phase I clinical trials in human transplant recipients. Findings generated in the current investigation indicate that leflunomide also possesses an additional unexpected beneficial property that distinguishes it from other immunosuppressive drugs: antiviral activity against CMV. Furthermore this agent, likely by virtue of its unique mechanism of action, shows equivalent effectiveness against CMV variants which have developed resistance to approved antiviral drugs. Remaining to be further substantiated, however, is whether leflunomide can exert antiviral effects in vivo. Although additional studies are needed, our preliminary experiments in a rat model indicate it can [28]. Should subsequent large-scale animal studies support these preliminary results, this agent would show great potential in

the clinical setting. Although it seems overly optimistic to expect leflunomide to provide a single drug solution for both allograft rejection and CMV disease in the transplant recipient, an effective immunosuppressant that substantially reduces viral load would greatly simplify treatment protocols employing traditional antiviral agents. Specifically, additive or synergistic effects between leflunomide and GCV, PFA, or cidofovir could be expected to reduce the required dosage and/or duration of treatment, thereby reducing untoward side effects. In summary, by virtue of its low toxicity [34], its apparent bifunctionality as both immunosuppressant and antiviral agent, and its unique mechanism of action effective against even multi-drugresistant variants, leflunomide holds great promise in ultimately helping to alleviate the clinical struggle to balance graft-preserving immunosuppression with control of CMV disease.



Acknowledgments The authors thank Mr. Terry McBride for technical assistance in electron microscopy, Dr. Marshall Williams for providing polymerase assay reagents, Dr. Yingxue Zhang for technical support in the preparation of DNA cosmid probes, and Mr. Andy Guglielmi for technical assistance at the initiation of this project. This investigation was supported in part by National Institutes of Health/NHLBI grant HL56482 (W.J.W.), and project seed grants from the Department of Pathology, The Ohio State University College of Medicine and Public Health (W.J.W.) and Rush Presbyterian-St. Luke’s Medical Center (A.S-F.C., N.S.L.). Daniel M. Miller was a Howard Hughes Medical Institute Predoctoral Fellow.

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13 Xiao F, Chong A, Shen J, Yang J, Short J, Foster P, Sankary H, Jensik S, Mital D, McChesney L, Koukoulis G, Williams JW: Pharmacologically induced regression of chronic transplant rejection. Transplantation 1995;60: 1065–1072. 14 Bartlett RR, Dimitrijevic M, Mattar T, Zielinske T, Germann T, Rüde E, Thoenes GH, Küchle CCA, Schorlemmer H-U, Bremer E, Finnegan A, Schleyerbach R: Leflunomide (HWA 486), a novel immunomodulating compound for the treatment of autoimmune disorders and reactions leading to transplantation rejection. Agents Actions 1991;32:10–21. 15 Xu X, Williams J, Bremer E, Finnegan A, Chong A: Inhibition of protein tyrosine phosphorylation in T cells by novel immunosuppressive agent, leflunomide. J Biol Chem 1995; 270:12398–12403. 16 Williamson RA, Yea CM, Robson PA, Curnock AP, Gadher S, Hambleton AB, Woodward K, Bruneau J-M, Hambleton P, Moss D, Thomson TA, Spinella-Jaegle S, Morand P, Courtin O, Sautes C, Westwood R, Hecend T, Kuo EA, Ruuth E: Dihydroorotate dehydrogenase is a high affinity binding protein for A77 1726 and mediator of a range of biological effects of the immunomodulatory compound. J Biol Chem 1995;270:22467–22472. 17 Mocarski ES Jr: Cytomegalovirus biology and replication: in Roizman B, Whitley RJ, Lopez C (eds): The Human Herpesviruses. New York, Raven Press, 1993, pp 173–226. 18 Myerson D, Hackman RC, Nelson JA, Ward DC, McDougall JK: Widespread presence of histologically occult cytomegalovirus. Hum Pathol 1984;15:430–439. 19 Sinzger C, Müntefering H, Löning T, Stöss H, Plachter B, Jahn G: Cell types infected in human cytomegalovirus placentitis identified by immunohistochemical double staining. Virchows Archiv A Pathol Anat 1993;423:249– 256. 20 Waldman WJ, Roberts WH, Davis DH, Williams MV, Sedmak DD, Stephens RE: Preservation of natural endothelial cytopathogenicity of cytomegalovirus by propagation in endothelial cells. Arch Virol 1991;117:143–164. 21 Waldman WJ, Knight DA, Lurain NS, Miller DM, Sedmak DD, Williams JW, Chong AS-F: Novel mechanism of inhibition of CMV by the experimental immunosuppressive agent leflunomide. Transplantation 1999;68:814–825. 22 Chong AS-F, Finnegan A, Jiang XL, Gebel H, Sankary HN, Foster P, Williams JW: Leflunomide, a novel immunosuppressive agent. The mechanism of inhibition of T cell proliferation. Transplantation 1993;55:1361–1366.


23 Chong AS-F, Huang W, Liu W, Luo J, Shen J, Xu W, Ma L, Binder L, Xiao F, Xu X, Clardy C, Foster P, Williams JW: In vivo activity of leflunomide: Pharmacokinetic analyses and mechanism of immunosuppression. Transplantation 1999;68:100–109. 24 Waldman WJ, Knight DA, Huang EH, Sedmak DD: Bidirectional transmission of infectious cytomegalovirus between monocytes and vascular endothelial cells: An in vitro model. J Infect Dis 1995;171:263–272. 25 Khym JX: An analytical system for rapid separation of tissue nucleotides at low pressures on conventional anion exchanges. Clin Chem 1975;21:1245–1252. 26 Gibson W: Structure and assembly of the virion. Intervirology 1996;39:389–400. 27 Bruggeman CA, Debie WMH, Grauls G, Majoor G, Van Boven CPA: 1983. Infection of laboratory rats with a new cytomegalo-like virus. Arch Virol 1983;76:189–199. 28 Waldman WJ, Blinder L, Knight DA, Shen J, Lurain NS, Miller DM, Sedmak DD, Williams JW, Chong AS-F: Inhibition of CMV in vitro and in vivo by the experimental immunosuppressive agent leflunomide. J Clin Virol 1999; 12:174. 29 Elder RT, Xu X, Williams JW, Gong J, Finnegan A, Chong AS-F: The immunosuppressive drug metabolite of leflunomide, A77 1726, affects murine T cells through two biochemical mechanisms. J Immunol 1997;159:22–27. 30 Xu X, Williams JW, Gong H, Finnegan A, Chong AS-F: Two activities of the immunosuppressant, leflunomide: Inhibition of pyrimidine synthesis and protein tyrosine phosphorylation. Biochem Pharmacol 1996;52:527–534. 31 Littler E, Stuart AD, Chee MS: Human cytomegalovirus UL97 open reading frame encodes a protein that phosphorylates the antiviral nucleoside analogue ganciclovir. Nature 1992; 358:160–162. 32 Sullivan V, Talarico CL, Stanat SC, Davis M, Coen DM, Biron KK: A protein kinase homologue controls phosphorylation of ganciclovir in human cytomegalovirus-infected cells. Nature 1992;358:162–164. 33 Lalezari JP, Stagg RJ, Kuppermann BD, Holland GN, Kramer F, Ives DV, Youle M, Robinson MR, Drew WL, Jaffe HS: Intravenous cidofovir for peripheral cytomegalovirus retinitis in patients with AIDS. A randomized, controlled trial. Ann Intern Med 1997;126:257– 263. 34 Mladenovic V, Domljan Z, Rozman B, Jajic I, Mihajlovic D, Dordevic J, Popovic M, Dimitrijevic M, Zivdovic M, Campion G, Musikic P, Löw-Friedrich I, Oed C, Seifert H, Strand V: Safety and effectiveness of leflunomide in the treatment of patients with active rheumatoid arthritis. Arthritis Rheum 1995;38:1595– 1603.

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Part II. Diagnostics and Antiviral Therapy Antivirals


Proinflammatory Potential of Cytomegalovirus Infection Specific Inhibition of Cytomegalovirus Immediate-Early Expression in Combination with Antioxidants as a Novel Treatment Strategy?

J. Cinatl Jr. J.-U. Vogel R. Kotchetkov M. Scholz H.W. Doerr Institut für Medizinische Virologie, Johann-Wolfgang-Goethe-Universität, Frankfurt am Main, Deutschland

Key Words Human cytomegaloviruses W Chemoattractants W Antiviral drugs W Immunopathology W Antisense oligonucleotides

Abstract We observed the effects of antiviral therapy on CMV and/ or oxidative-stress-induced stimulation of proinflammatory molecules including interleukin-8 (IL-8), melanoma growth stimulatory activity-· (GRO-·) and intercellular adhesion molecule 1 (ICAM-1) using human foreskin fibroblasts. Ganciclovir, foscarnet or cidofovir completely suppressed virus replication, as demonstrated by CMV late (L) antigen production. These drugs did not influence CMV immediate-early (IE) antigen expression and had no effects on CMV-induced cellular changes in IL-8, GRO-· and ICAM-1 levels. Phosphorothioate oligonucleotide (ISIS 2922) suppressed both CMV IE and L antigen by 99%. ISIS 2922 completely suppressed CMVinduced upregulation of both chemokines and ICAM-1. Induction of oxidative stress by H2O2 upregulated IL-8 expression. Oxidative stress and CMV infection showed synergistic effects on IL-8 expression. ISIS 2922 only partially inhibited the upregulation of IL-8 in infected cells treated with H2O2, whereas cotreatment with ISIS 2922 and antioxidants inhibited the upregulation almost completely. The results showed that inhibition of CMV IE expression alone or in combination with antioxidants is promising for the treatment of CMV disease.

Introduction

The current anti-human cytomegalovirus (HCMV) treatment strategies are directed against virus DNA replication, but frequently fail to halt the disease [1]. This seems to be due to virus-induced ‘side effects’ that are not correlated to de novo production of virions and lysis of host cells. We and others suggested that the CMV immediate-early (IE) genes may play a predominant role in CMV-induced immunopathogenesis (inflammatory diseases). Moreover, redox mechanisms that are involved in inflammation may be important cofactors for activation of the virus, probably by stimulating CMV IE genes [2]. In order to develop novel treatment strategies which might supplement the therapeutic blocking of virus DNA replication, we promote the idea to target CMV IE in combination with antioxidants. In the following paper we focus on the role of (a) CMV IE in immunopathogenesis, (b) oxidative stress in CMV activation, and (c) ISIS 2922/n-acetylcysteine (NAC) in anti-CMV treatment, (d) ISIS 2922/ NAC in inhibiting CMV-induced immunomodulation.

HCMV (HCMV IE) and Immunopathogenesis

HCMV IE gene products have autoregulatory features and in addition are strong transactivators, known to stimulate the transcription of various viral and host cell genes [3]. The major immediate-early (IE1/IE2) promoter-enhancer is located upstream of the IE1/IE2 locus and con-

Copyright © 2000 S. Karger AG, Basel

ABC

© 2000 S. Karger AG, Basel 0300–5526/99/0426–0419$17.50/0



Dr. Jindrich Cinatl Jr. Institut für Medizinische Virologie, Zentrum der Hygiene Klinikum der Johann-Wolfgang-Goethe-Universität, Sandhofstrasse 2–4 D–60528 Frankfurt am Main (Germany) Tel. +49 69 6301 6409, Fax +49 69 6301 4302, E-Mail [email protected]

Table 1. Cytokines and growth factors stimulated by CMV infec-

tion Tumor necrosis factors (e.g. TNF-·) Interleukins (e.g. IL-1, IL-6) Interferons (e.g. IFN-·) CC chemokines (e.g. RANTES, MIP-1ß) CXC chemokines (e.g. IL-8, GRO-·) Transforming growth factors (e.g. TGFß) Angiogenic factors (e.g. bFGF) Hematopoietic growth factors (e.g. GM-CSF, G-CSF) IL = Interleukin; RANTES regulated on activation, normal T-cell expressed and secreted; MIP-1· = macrophage inflammatory protein; GRO-· = growth-related; bFGF = basic fibroblast growth factor; GM-CSF = granulocyte macrophage-colony-stimulating factor; G-CSF = granulocyte-colony-stimulating factor.

tains multifold binding sites for viral (e.g. IE2, pp71 tegument protein) and cellular transcription factors such as NF-ÎB, ATF/CREB, AP1, SP1, C/EBP, SREB, and NF1 [4–7]. These cellular transcription factors bind to repeated elements (16-, 18-, 19-, and 21-bp repeats). For example, in immunocompromised patients, such as transplant patients, aberrant secretion of the proinflammatory cytokine TNF-· may result in cytokine-mediated activation of NKÎB and subsequently to stimulation of the 18-bp repeat consensus region of the major immediate early promoterenhancer [8]. On the other hand, IE1 exerts an autostimulatory effect via activation of NF-ÎB which in turn stimulates the major immediate early promoter-enhancer and thus expression of IE1 [7, 9]. Particular IE2 gene products act as negative autoregulators by binding to the cis-repression signal (crs) sequence located immediately upstream of the IE1/IE2 transcription start site [10]. Therefore, IE2 gene products, especially IE2579aa and IE2338aa (late IE2 gene product), mediate the shut-off of IE1 and IE2 gene expression, probably at the transcriptional level by altering the RNA polymerase II [11]. However, IE2 proteins alone or in synergism with IE1 may be promiscous transactivators of both viral and cellular gene expression [12]. The transactivating properties of IE gene products may partly explain the significant pathogenicity of HCMV in immunocompromised patients such as transplant recipients and AIDS patients. These patients suffer from severe manifestations, such as graft rejection [13–15], pneumonia [16] or HCMV retinitis [17]. It is assumed that HCMV may alter the expression of genes coding for proteins with proinflammatory activity, probably without the requirement for virus replication [2, 18]. HCMV infec-

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tion has been associated with increased cytokine production, directly by the infected cells and/or indirectly by immune cells that interact with infected cells [19–21]. In table 1, a selection of HCMV-induced cytokines and growth factors that contribute to proinflammatory activity is shown. Because HCMV has been associated with inflammatory disorders, such as episodes of graft rejection after organ transplantation, it is assumed that HCMV triggers leukocytic infiltration of infected tissues. In vitro experiments performed in different laboratories have shown that HCMV induced the upregulation of several adhesion molecules such as intercellular adhesion molecule-1 (ICAM-1) or lymphocyte-function-associated molecule-3 (LFA-3) [18–22]. Moreover, HCMV infection was reported to be associated with increased functional binding of leukocytes to infected cells, which is a major step in leukocyte transmigration [23–26]. Recently, it has been shown that HCMV induces enhanced production of both CXC (i.e. IL-8 and GRO-·) and CC (i.e. RANTES and MIP-1·) chemokines in different cell types [2, 19, 27–29]. Importantly, secretion of CXC chemokines by HCMV-infected endothelial cells resulted in increased transendothelial neutrophil migration [29]. HCMV-induced chemoattraction of leukocytes in concert with enhanced expression of adhesion molecules may result in unspecific infiltration of the infected tissue and thus leads to HCMV-associated immunopathogenic effects. It has previously been shown that a variety of HCMVinduced alterations in cellular characteristics including expression of adhesion molecules and synthesis of cytokines or chemokines occur early after virus infection and may result from the transactivation activity of IE proteins [1, 2, 30]. The effects of CMV on the expression of the adhesion molecule ICAM-1 and the chemokines GRO-· and IL-8 are shown quantitatively in figure 1.

HCMV, Inflammation and Redox Mechanisms

Inflammatory mechanisms and oxidative stress both are discussed to be associated with CMV activation [2]. Interestingly, the proinflammatory cytokine TNF-· may induce oxidative stress and has been shown to directly induce CMV replication [8]. In HL-60 cells, it was shown that TNF-· stimulated the transfected CMV IE enhancerpromoter via induction of the transcription factor NF-ÎB [8], a pathway that is also involved in the activity of reactive oxygen intermediates. It has been reported that the

Cinatl/Vogel/Kotchetkov/Scholz/Doerr

3,000

2,500

Virus-free supernatant UV-inactivated virus Mock AD169

IL-8 (pg/ml)

2,000

1,500

1,000

500

a

0 500

GRO-a (pg/ml)

400

300

200

100

b

0

Fig. 1. Time course of IL-8 (a) and GRO-· secretion (b) as determined by means of ELISA, or ICAM-1 expression (c) by flow

cytometry 4, 24, and 72 h after CMV infection. Data are depicted as means B SD from one representative experiment performed in triplicates.

ICAM-1 (fluorescence intensity units)

500

400

300

200

100

0

c 4

24 Time after infection (h)

72

intracellular levels of reduced glutathion, the major radical scavenger, are inversely correlated with the permissiveness of CMV in different cell types [31]. In vitro, oxidative stress can be unspecifically induced by reactive oxygen species, such as H2O2. In contrast, glutathion lev-

els can be reduced specifically by treatment of the cells with buthioninsulfoximine, which inhibits Á-glutamylcysteine synthetase and leads to activation of CMV [32]. On the other hand, NAC, which increases the glutathion levels, was found to inhibit CMV infection in vitro [31].

Proinflammatory Potential of Cytomegalovirus Infection


421

s

s

Us

s

s

UL

MIE promotor

s

Exon No.

1

2

3

4

5

IE1 72 kD Alternatively spliced mRNA

IE1 55 kD

Fig. 2. Scheme of organization of the

IE2 86 kD

HCMV genome, its major IE (MIE) transcriptional unit and its exons. Shaded bars represent protein-coding regions. Alternatively spliced mRNAs encoding the major polypeptides and the complementary target sequence for ISIS 2922 are shown below.

IE2 RNA 5’-CGC AAG AAG AAG AGC AAA CGC-3’ ISIS 2922 5’-GCG TTC TTC TTC TCG TTT GCG-3’

Mock Mock + H2O2 AD169 AD169 + H2O2

70 60 5

IL-8 (ng/ml)

4

3

2

Fig. 3. Effects of the antiviral drug ISIS 2922 and/or antioxidant NAC on IL-8 production in CMV-infected fibroblasts after H2O2-induced oxidative stress as determined by means of ELISA. Data are depicted as means B SD from one representative experiment performed in triplicates.

1

0 Without treatment

ISIS 2922 (1 mM)

NAC (5 mM)

ISIS 2922 (1 mM) + NAC (5 mM)

Table 2. Effects of antiviral drugs on CMV-induced expression of IL-8, GRO-· and ICAM-1 Inflammatory Control molecule mock IL-8 GRO-· ICAM-1

GCV CMV

1.00B0.20 8.90B1.00 1.00B0.20 6.90B0.90 1.00B0.15 3.30B0.40

mock

PFA CMV

0.90B0.21 8.40B1.00 1.20B0.20 8.20B1.00 1.10B0.16 3.90B0.50

mock

HPMPC CMV

1.20B0.19 9.40B1.00 1.10B0.22 7.20B1.00 0.90B0.10 3.80B0.40

mock

ISIS 2922 CMV

1.10B0.22 9.10B1.00 0.90B0.19 6.80B0.80 0.80B0.10 3.40B0.40

mock

CMV

1.30B0.22 3.90B0.40 1.10B0.14 2.80B0.40 1.20B0.21 1.30B0.20

IL-8 and GRO-· production determined by ELISA; ICAM-1 expression measured by flow cytometry, 72 h after infection. Data relative to the mock-infected control are shown as means B SD from one representative experiment performed in triplicates.

422



Inflammation

s Antioxidants

IEA

EA

s

ROI

CMV disease

s

s

LA

s s

ISIS 2922 Combined with Antioxidants: A Novel Anti-CMV Treatment Strategy? The antiviral agents currently in clinical use, i.e. GCV, PFA and HPMPC, exert their actions by preventing viral DNA replication, and the subsequent synthesis of HCMV late proteins [34]. These drugs efficiently prevent virus replication, but have no influence on events (IE expression) that occur before viral DNA synthesis [35]. Thus, treatment with inhibitors of viral DNA synthesis may be insufficient to prevent these pathogenic features of HCMV infection. A phosphorothioate oligonucleotide, ISIS 2922 (Fomivirsen), has been developed for the treatment of CMV retinitis in AIDS patients [36, 37]. ISIS 2922 is currently

ISIS 2922

s

It has recently, been shown that chemokine secretion may be modified by CMV and oxidative stress [2, 27]. Interestingly, CMV-mediated chemokine secretion, as has also been demonstrated for ICAM-1 expression cannot be inhibited by the standard anti-CMV drugs ganciclovir (GCV), foscarnet (PFA), and cidofovir (HPMPC). On the other hand, ISIS 2922, an antisense oligonucleotide complementary to part of HCMV IE2 mRNA (fig. 2), almost completely inhibited CMV-induced ICAM-1 but only partly impaired CMV-induced IL-8 and GRO-· production (table 2) [33]. Moreover, CMV infection and concomitant oxidative stress exhibited synergy in terms of upregulation of IL-8 gene expression and secretion (fig. 3). The dramatic augmentation of IL-8 secretion elicited by the combination of CMV/oxidative stress, as documented in figure 3, might be more relevant in CMV disease than in CMV or oxidative stress alone. Experimentally, this synergistic effect was only partly reduced by NAC, but totally inhibited by combined treatment with ISIS 2922 and NAC. Therefore, it has been suggested that specific inhibitors of the IE gene products in combination with potent antioxidants may be an important additional option for the clinical treatment of CMV disease.

Induction of chemokines, cytokines, adhesion molecules

GCV HPMPC PFA

Fig. 4. Scheme showing interdependent mechanisms between in-

flammation and CMV infection. Effects of different antiviral targeting on suppression of virus-related immunopathology. ROI = Reactive oxygen species.

the only drug that targets the CMV IE mRNA by binding to complementary sense sequences on mRNA and thus blocks the production of CMV IE proteins (fig. 2). When administered intravitreally, ISIS 2922 significantly delayed disease progression [38,39]. However, the mode of antiviral action is not totally resolved yet. It is discussed that besides specific antisense inhibition (sequence dependent) nonantisense mechanisms (unspecific inhibition of virus adsorption) may also contribute to the antiviral effects of ISIS 2922 [40]. In conclusion, the strategy to inhibit IE expression of CMV alone or in combination with antioxidants seems to be promising for the treatment of CMV disease (fig. 4).

Acknowledgments This work was generously supported by the foundations: ‘Hilfe für krebskranke Kinder Frankfurt, e.V.’ and ‘Frankfurter Stiftung für krebskranke Kinder’. We are grateful to Sandra Dujardin, Karina Cordier and Gesa Meincke for technical assistance.

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Proinflammatory Potential of Cytomegalovirus Infection

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17 Holland GN, Pepose JS, Pettit DH: Acquired immune deficiency syndrome: Ocular manifestations. Ophthalmology 1983;90:859–873. 18 Craigen JL, Grundy JE: Cytomegalovirus induced up-regulation of LFA-3 (CD58) and ICAM-1 (CD54) is a direct viral effect that is not prevented by ganciclovir or foscarnet treatment. Transplantation 1996;62:1102–1108. 19 Grundy JE, Lawson KM, MacCormac LP, Fletcher JM, Yong KL: Cytomegalovirus-infected endothelial cells recruit neutrophils by the secretion of CXC chemokines and transmit virus by direct neutrophil-endothelial cell contact and during neutrophil transendothelial migration. J Infect Dis 1998;177:1465–1467. 20 Scholz M, Blaheta RA, Cinatl J, Encke A, Doerr HW, Markus BH: Lymphocytes induce enhanced expression of HLA class I antigens on cytomegalovirus-infected syngeneic human endothelial cells. Hum Immunol 1995;44:136– 144. 21 Scholz M, Hamann A, Blaheta RA, Auth MKH, Encke A, Markus BH: Cytomegalovirus- and interferon-related effects on human endothelial cells: Cytomegalovirus infection reduces upregulation of HLA class II antigen expression after treatment with interferon-Á. Human Immunol 1992;35:230–238. 22 Grundy JE, Downes KL: Up-regulation of LFA-3 and ICAM-1 on the surface of fibroblasts infected with cytomegalovirus. Immunology 1993;78:405–412. 23 Span AHM, Mullers W, Miltenburg AMM, Bruggeman CA: Cytomegalovirus-induced PMN adherence in relation to an ELAM-1 antigen present on infected endothelial cell monolayers. Immunology 1991;72:355–360. 24 Tuder RM, Weinberg A, Panatjotoboulos N, Kalil J: Cytomegalovirus enhances PBMC binding and HLA class I expression on cultured endothelial cells. Transplant Proc 1991; 23:91. 25 Grundy JE, Pahal GS, Akbar AN: Increased adherence of CD2 peripheral blood lymphocytes to cytomegalovirus-infected fibroblasts is blocked by anti-LFA-3 antibody. Immunology 1993;78:413–420. 26 Blaheta R, Hailer N, Harder S, Scholz M, Bereiter-Hahn J, Markus BH: Up- and downregulation of lymphocyte infiltration through allogeneic endothelial cell monolayers. Tex Med 1994;6:53–59. 27 Michelson S, Dal Monte P, Zipeto D, Bodaghi B, Laurent L, Oberlin E, Arenzana-Seisdesos F, Virelizier J-L, Landini MP: Modulation of RANTES production by human cytomegalovirus infection of fibroblasts. J Virol 1997;71: 6495–6500.


28 Murayama T, Ohara Y, Obuchi M, Khabar KSA, Higashi H, Mukaida N, Matsushima K: Human cytomegalovirus induces interleukin-8 production by a human monocytic cell line THP-1, through acting concurrently on AP-1 and NF-ÎB-binding sites of the interleukin-8 gene. J Virol 1997;71:5692–5695. 29 Craigen JL, Yong KL, Jordan JN, MacCormac LP, Westwick J, Akbar AN, Grundy JE: Human cytomegalovirus infection upregulates interleukin-8 gene expression and stimulates neutrophil transendothelial migration. Immunology 1997;92:138–145. 30 Burns L, Pooley JC, Walsh DJ, Vercelloti GM, Weber ML, Kovacs A: Intercellular adhesion molecule-1 expression in endothelial cells is activated by cytomegalovirus immediate early proteins. Transplantation 1999;67:137–144. 31 Vossen RC, Persoons MC, Slobbe-van Drunen ME, Bruggeman CA, van Dam-Mieras MC: Intracellular thiol redox status affects rat cytomegalovirus infection of vascular cells. Virus Res 1997;48:173–183. 32 Scholz M, Cinatl J, Gross V, Vogel J-U, Blaheta RA, Freisleben H-J, Markus BH: Impact of oxidative stress on human cytomegalovirus replication and on cytokine-mediated stimulation of endothelial cells. Transplantation 1996;61: 1763–1770. 33 Cinatl J Jr, Kotchetkov R, Weimer E, Blaheta RA, Scholz M, Vogel J-U, Gümbel HO, Doerr HW: The antisense oligonucleotide ISIS 2922 prevents cytomegalovirus-induced upregulation of IL-8 and ICAM-1 in cultured human fibroblasts. J Med Virol, in press. 34 Vogel J-U, Scholz M, Cinatl J Jr: Treatment of CMV diseases. Intervirology 1998;40:357– 367. 35 Cinatl J Jr, Kotchetkov R, Scholz M, Cinatl J, Vogel J-U, Herna´iz-Driever P, Doerr HW: Human cytomegalovirus infection decreases expression of thrombospondin-1 independent of the tumor suppressor protein p53. Am J Pathol 1999;155:285–292. 36 Anderson KP, Fox MC, Driver VB, Martin MJ, Azad RF: Inhibition of human cytomegalovirus immediate early gene expression by an antisense oligonucleotide complementary to immediate early RNA. Antimicrob Agents Chemother 1999;40:2004–2011. 37 Azad RF, Driver VB, Tanaka K, Crooke RM, Anderson KP: Antiviral activity of a phosphorothioate oligonucleotide complementary to RNA of the human cytomegalovirus major immediate-early region. Antimicrob Agents Chemother 1993;37:1945–1954. 38 Marwick C: First ‘antisense’ drug will treat CMV retinitis. JAMA 1998;280:871. 39 Perry CM, Balfour JAB: Fomivirsen. Drugs 1999:57:375–380. 40 De Clercq E: Fomivirsen: A viewpoint. Drugs 1999;57:381.


Author Index Vol. 42, No. 5–6, 1999

Ahmed, M. 291 Ahonen, J. 279 Banas, R. 291 Baur, R. 357 Beisser, P.S. 342 Berg, A.P. van den 285, 382 Bij, W. van der 285, 382 Biomed 2 Study Group 382 Blaheta, R.A. 350 Blinder, L. 412 Blok, M.J. 373 Bruggeman, C.A. 277, 279, 342, 373 Cai, J. 291 Cebulla, C.M. 325 Chong, A.S.-F. 412 Christiaans, M.H.L. 373 Cinatl, J., Jr. 350, 419 Dauber, J. 291 Davis-Poynter, N.J. 331 Degli-Esposti, M. 331 Döcke, W.-D. 308 Doerr, H.W. 350, 419 Donnenberg, A.D. 291 Donnenberg, V.S. 291 Einsele, H. 365 Emery, V.C. 405 Ewert, R. 322

Gabrielli, L. 390 Goossens, V.J. 373 Griffith, B. 291


Iacono, A. 291 Inkinen, K. 279 Jahn, G. 365 Kas-Deelen, A.M. 382 Kauppinen, H. 279 Keenan, R. 291 Kern, F. 322 Khatamzas, E. 322 Kloover, J. 279 Knight, D.A. 412 Kotchetkov, R. 419 Krogerus, L. 279 Krüger, D.H. 308

Prösch, S. 308, 322 Ramon, A. 277 Reinhardt, B. 357 Reinke, P. 308, 322 Riegler, S. 265 Schaarschmidt, P. 357 Scholz, M. 277, 350, 419 Sedmak, D.D. 325, 412 Shen, J. 412 Sillekens, P. 373 Sinzger, C. 365 Söderberg-Nauclér, C. 314 Son, W.J. van 285, 382 Soots, A. 279 Spezzacatena, P. 390 Spichty, K. 291 Stenglein, S. 365 The, T.H. 285, 382

Landini, M.P. 390 Lautenschlager, I. 279, 373 Lazzarotto, T. 390 Limburg, P.C. 382 Loginov, R. 279 Lurain, N.S. 412 Lüske, A. 357 Maar, E.F. de 285 Mayr, K. 357 Meedendorp, B. 382 Mertens, T. 357 Michel, D. 357 Michelson, S. 301 Middeldorp, J.M. 373 Miller, D.M. 325, 412

Farrell, H.E. 331 Faulhaber, N. 322 Frömmel, C. 322

ABC

Harmsen, M.C. 382 Höckerstedt, K. 373 Holma, K. 279 Hooff, J.P. van 373

Vaida, B. 357 Vanholder, R. 398 Vanpoucke, H. 398 Van Renterghem, L. 398 Van Vlem, B. 398 Varani, S. 390 Verschuuren, E.A.M. 285, 382 Vink, C. 342 Vogel, J.-U. 350, 419 Voisard, R. 357 Volk, H.-D. 308, 322 Waldman, W.J. 412 Williams, J.W. 412 Zeevi, A. 291

Nelson, J.A. 314



425

Subject Index Vol. 42, No. 5–6, 1999

Adenovirus 325 Adhesion molecules 350 Allogeneic stimulation 314 Allograft rejection 291 Amniotic fluid 390 Animal models 342 Antibody avidity 390 Antigenemia 373, 382 Antigen-presenting cells 365 Antisense oligonucleotides 419 Antiviral drugs 419 – therapy 412 Cadherin-catenin complex 350 cAMP 308 Cellular immune defence 322 Chemoattractants 419 Chemokine(s) 331, 342 – sequestration 301 Chronic rejection 279 Congenital infection 390 Cytokines 291 Cytomegalovirus 279, 285, 291, 301, 308, 314, 322, 325, 331, 342, 350, 357, 365, 373, 382, 390, 398, 412, 419 Cytopathic effect 357 Ebola virus 325 Endothelial cells 412 Epstein-Barr virus 325 Extracellular matrix 357

Immediate-early gene products 373 Immune escape 301 – evasion 331 Immunosuppression 322, 365 Interferon signal transduction 325 Intracellular cytokines 322 JAK/STAT pathway 325 Latency, human cytomegalovirus 314 Leflunomide 412 Macrophages 314, 365 Metalloproteinases 350 MHC class I 331, 342 Monocyte/granulocyte progenitor cells 308 mRNA 373 Mumps 325 Organ transplantation 285, 398 PCR 373, 390, 398, 405 pp67 373 Pregnancy 390 Reactivation, human cytomegalovirus 314 Recombinant virus 342 Renal transplantation 279 Stress 308 Symptomatology 285 Systemic inflammation 308

Fibroblasts 412 G protein coupled receptor(s) 331, 342 Hepatitis B virus 325 Herpesvirus 331 HLA 301 Human papillomavirus 325 – renal artery organ culture 357 IgM 390 Immature dendritic cells 365




T cell activation 291 – cells 322 – helper responses 291 Transendothelial migration 350 Tumor necrosis factor alpha 308 Varicella-zoster virus 325 Viral load 398 Virus reactivation 308 VLA-5 350

Author Index Vol. 42, 1999

Afione, S.A. 213 Aguilera, A. 37 Ahmed, M. 291 Ahonen, J. 279 Angeretti, A. 30 Aoki, H. 205 Arakawa, Y. 205 Astori, G. 221

Emery, V.C. 405 Encke, J. 117 Ewert, R. 322

Banas, R. 291 Barman, S. 238 Baur, R. 357 Beerheide, W. 228 Beisser, P.S. 342 Beltrame, A. 221 Berg, A.P. van den 285, 382 Bij, W. van der 285, 382 Biomed 2 Study Group 382 Blaheta, R.A. 350 Blinder, L. 412 Blok, M.J. 373 Bonaguro, R. 1 Bonino, F. 69 Borisova, G. 51 Boschetto, R. 1 Botta, G.A. 221 Breslin, J.J. 22 Bruggeman, C.A. 277, 279, 342, 373 Brunetto, M.R. 69 Buoro, S. 1 Caballero, E. 37 Cai, J. 291 Caudai, C. 1 Cavallo, R. 30 Cebulla, C.M. 325 Chong, A.S.-F. 412 Christiaans, M.H.L. 373 Cilla, G. 37 Cinatl, J., Jr. 350, 419 Cusan, M. 1 Dauber, J. 291 Davis-Poynter, N.J. 331 Degli-Esposti, M. 331 Döcke, W.-D. 308 Doerr, H.W. 350, 419 Donnenberg, A.D. 291 Donnenberg, V.S. 291 Edler, L. 228 Einsele, H. 365




Farrell, H.E. 331 Faulhaber, N. 322 Flotte, T.R. 213 Frazer, I.H. 43 Frömmel, C. 322 Fuente, J. de la 271 Fuller, F.J. 22 Gabrielli, L. 390 Gelderblom, H.R. 51 Goossens, V.J. 373 Gribaudo, G. 30 Griffith, B. 291 Grubhoffer, L. 9 Guggino, W.B. 213 Gutiérrez, M. 37 Guy, J.S. 22 Hansen, J. 17 Harmsen, M.C. 382 Hayashi, J. 205 Hertel, L. 30 Hilditch-Maguire, P.A. 43 Hino, O. 61, 205 HIV-2 Spanish Study Group 37 Höckerstedt, K. 373 Holguı´n, A. 37 Holma, K. 279 Hooff, J.P. van 373 Iacono, A. 291 Iino, S. 61, 166 Inkinen, K. 279 Irshad, M. 252 Ishii, K. 145 Jahn, G. 365 Jindra´k, L. 9 Jung, H.C. 263 Kairat, A. 228 Kas-Deelen, A.M. 382 Katz, E. 247 Kauppinen, H. 279 Keenan, R. 291 Kern, F. 322 Keywan, K. 247 Khatamzas, E. 322

Kim, C.Y. 263 Kim, J.S. 263 Kim, Y.T. 263 Kiyosawa, K. 185 Kloover, J. 279 Knight, D.A. 412 Knopf, C.W. 17 Koletzki, D. 51 Kopecky´, J. 9 Kotchetkov, R. 419 Kova´rˇ, V. 9 Koyama, T. 153 Krogerus, L. 279 Krüger, D.H. 51, 308 Kühn, F.J.P. 17 Lachmann, S. 51 Lambert, P.F. 43 Landini, M.P. 390 Landolfo, S. 30 Lautenschlager, I. 279, 373 Lazzarotto, T. 390 Lee, H.-S. 263 Lembo, D. 30 Lieppe, D.M. 43 Limburg, P.C. 382 Loginov, R. 279 Lurain, N.S. 412 Lüske, A. 357 Maar, E.F. de 285 Machuca, A. 37 Majumdar, S. 238 Matsuura, Y. 145 Mayr, K. 357 Meedendorp, B. 382 Meisel, H. 51 Mengoli, C. 1 Mertens, T. 357 Michel, D. 357 Michelson, S. 301 Middeldorp, J.M. 373 Miller, D.M. 325, 412 Mishiro, S. 153 Miyamura, T. 145 Mizokami, M. 159 Moritsugu, Y. 63 Moriya, T. 153 Murakami, S. 81 Muselmann, C. 51

427

Nassal, M. 100 Nelson, J.A. 314 Nishizawa, T. 196

Shen, J. 412 Sillekens, P. 373 Sinzger, C. 365 Smith, L.G. 22 Söderberg-Nauclér, C. 314 Son, W.J. van 285, 382 Song, I.S. 263 Soots, A. 279 Soriano, V. 37 Spezzacatena, P. 390 Spichty, K. 291 Stenglein, S. 365 Suzuki, R. 145 Suzuki, T. 145

Okamoto, H. 196 Orito, E. 159 Padula, M. 1 Palù, G. 1 Pellizzari, L. 1 Pipan, C. 221 Pizzighella, S. 1 Prösch, S. 308, 322 Pumpens, P. 51 Putlitz, J. zu 117

Tanaka, E. 185 Tanaka, J. 153 Tang, Z.-Y. 228 Taylor, J.M. 173 The, T.H. 285, 382 Totsuka, A. 63 Trepo, C. 125

Ramon, A. 277 Raphenon, G. 221 Reinhardt, B. 357 Reinke, P. 308, 322 Riegler, S. 265 Riera, L. 30 Rodriguez, U.A. 69 Schaarschmidt, P. 357 Scholz, M. 277, 350, 419 Schreiner, U. 17 Schröder, C.H. 228 Sedmak, D.D. 325, 412

428

Intervirology Vol. 42, 1999

Ukita, M. 196 Ulrich, R. 51

Vanpoucke, H. 398 Van Renterghem, L. 398 Van Vlem, B. 398 Varani, S. 390 Verschuuren, E.A.M. 285, 382 Vink, C. 342 Vogel, J.-U. 350, 419 Voisard, R. 357 Vokurkova´, D. 9 Volk, H.-D. 308, 322 Waldman, W.J. 412 Walsh, S. 213 Wands, J.R. 117 Wang, J. 213 West, D. 43 Williams, J.W. 412 Yarbough, P.O. 179 Yoon, J.-H. 263 Yoon, Y.B. 263 Yoshizawa, H. 153 Zeevi, A. 291 Zhou, G. 228 Zoulim, F. 125

Vaida, B. 357 Valensin, P.E. 1 Vanholder, R. 398

Author Index

Subject Index Vol. 42, 1999

·ACE6E7#19 43 Acute hepatitis 196 Acyclovir 247 – resistance 247 ADAR 173 Adeno-associated virus 213 Adenovirus 213, 325 Adhesion molecules 350 Affinity chromatography 17 Africa 221 Age-related markers 228 Allogeneic stimulation 314 Allograft rejection 291 ALT normalization 166 Amniotic fluid 390 Animal models 342 Antibody avidity 390 – column 17 Antigenemia 373, 382 Antigenicity 51 Antigen-presenting cells 365 Anti-idiotypic antibodies 9 Antisense oligonucleotides 419 Antiviral drugs 419 – therapy 125, 412 Blood donor 153 – transfusion 196 Cadherin-catenin complex 350 cAMP 308 Cellular immune defence 322 Chemoattractants 419 Chemokine(s) 331, 342 – sequestration 301 Chimaeric core particles 51 Choleraphage 238 Chronic hepatitis 196 – – B 125, 228 – transplant rejection 279 Circoviridae 196 Congenital infection 390 Coronavirus 22 Cytokines 291 Cytomegalovirus 30, 279, 285, 291, 301, 308, 314, 322, 325, 331, 342, 350, 357, 365, 373, 382, 390, 398, 412, 419 Cytopathic effect 357 Dihydrofolate reductase 30 Duck hepatitis B virus 100




E7 43 Ebola virus 325 Endothelial cells 412 Epitope mapping 17 Epstein-Barr virus 325 Extracellular matrix 357 Fibroblasts 412 Flaviviridae 185 Gene therapy 213 Genetic immunization 117 Genotypes 196 G protein coupled receptor(s) 331, 342 HBeAg minus HBV 69 HBx 228 Hepadnaviridae 125 Hepatitis 205, 252 –, anicteric 252 – A virus 63 – B virus 51, 69, 117, 125, 159, 228, 325 – – – encapsidation signal 100 – – – replication origin 100 – C virus 117, 145, 153, 159, 205 – – – carrier 153 – – – genotypes 1, 166 – – – RNA 1, 153, 263 – – – – level 166 – – – – negativity 166 – delta virus 173 –, epidemic 252 – E virus 179, 252 – G virus 185 – virus(es) 159, 196 Hepatocarcinogenesis 205 Hepatocellular carcinoma 205, 228 Herpes simplex 247 – – virus type 1 17 Herpesvirus 331 HIS tag 17 HIV-2 37 HLA 301 HPOL tag 17 HPV16 43 HSV-1 DNA polymerase 17 Human papillomavirus 325 – renal artery organ culture 357 IgM 390 Immature dendritic cells 365

Immediate-early gene products 373 Immune escape 301 – evasion 331 Immunosuppression 322, 365 Induced-fit activation 100 Infectious bronchitis virus 22 INNOLiPA 1 Interferon signal transduction 325 Intracellular cytokines 322 JAK/STAT pathway 325 Latency, human cytomegalovirus 314 Leflunomide 412 Liver damage pathogenesis 69 – disease 185 Macrophages 314, 365 Metalloproteinases 350 Methotrexate 30 MHC class I 331, 342 Molecular evolution 159 Monoclonal antibody 17 Monocyte/granulocyte progenitor cells 308 mRNA 373 Mumps 325 Mutants 69 Nucleoside analogues 125 – –, resistance 69, 125 Organ transplantation 285, 398 Papillomavirus 221 PCR 22, 30, 196, 373, 390, 398, 405 PCR-RFLP 221 Phage genome 238 Phage-specific proteins 238 Phylogenetic tree 159 Physical map 238 Positive, single-strand RNA virus 185 pp67 373 P protein, hepadnaviral, complexes 100 Precipitation reaction 263 Pregnancy 390 PreS1 epitope 51 Protein processing 63, 145 – tagging 17 Quiescent cells 30

429

Reactivation, human cytomegalovirus 314 Recombinant virus 342 Renal transplantation 279 Replication site 185 Replicative intermediates 238 Restriction fragment length polymorphism 1 Reverse transcription 100 Ribozyme 173 RNA editing 173 – maturation 228 – polymerase 81, 173 Self-cleavage 173 Sequence variation 221 Sequencing 221

430


Serology 179 Signal transduction 205 Stress 308 Symptomatology 285 Systemic inflammation 308 T cell activation 291 – cells 322 – helper responses 291 Thumb region 17 Tick-borne encephalitis virus receptor 9 Transendothelial migration 350 Transgenic mice 43 Truncated RNA 228 Tumor necrosis factor alpha 308 Turkey coronavirus 22

Vaccine escape mutants 69 Varicella-zoster virus 325 Viral heterogeneity 69 – load 398 – proteins 145 – resistance 125 Virology 179 Virus affinoblotting 9 – reactivation 308, 314 Virus-like particles 51 VLA-5 350 Western blot 17 X protein 81

Subject Index

Contents Vol. 42, 1999

81 Hepatitis B Virus X Protein: Structure, Function and Biology Murakami, S. (Kanazawa)

No. 1

100 Hepatitis B Virus Replication: Novel Roles for Virus-Host

Original Papers 1

Interactions

Typing of Hepatitis C Virus by a New Method Based on Restriction Fragment Length Polymorphism Buoro, S. (Padova); Pizzighella, S. (Verona); Boschetto, R.; Pellizzari, L.; Cusan, M.; Bonaguro, R.; Mengoli, C. (Padova); Caudai, C.; Padula, M.; Valensin, P.E. (Siena); Palù, G. (Padova)

9

A Putative Host Cell Receptor for Tick-Borne Encephalitis Virus Identified by Anti-Idiotypic Antibodies and Virus Affinoblotting Kopecký, J.; Grubhoffer, L.; KovárZ , V.; Jindrák, L. (CZ eské BudeZ jovice); Vokurková, D. (Hradec Králové)

17

22

30

37

A Novel Protein Tag from Herpes Simplex Virus Type 1 DNA Polymerase

125 New Antiviral Agents for the Therapy of Chronic Hepatitis B

Virus Infection Zoulim, F.; Trepo, C. (Lyon)

145 Processing and Functions of Hepatitis C Virus Proteins Suzuki, R.; Suzuki, T.; Ishii, K.; Matsuura, Y.; Miyamura, T. (Tokyo)

Schreiner, U. (Heidelberg); Hansen, J. (Wuppertal); Kühn, F.J.P.; Knopf, C.W. (Heidelberg)

153 Epidemiology of Hepatitis C Virus in Japan Moriya, T. (Hiroshima); Koyama, T. (Morioka); Tanaka, J. (Hiroshima); Mishiro, S. (Tokyo); Yoshizawa, H. (Hiroshima)

Sequence Analysis of the Matrix/Nucleocapsid Gene Region of Turkey Coronavirus

159 Molecular Evolution of Hepatitis Viruses Mizokami, M.; Orito, E. (Nagoya)

Breslin, J.J.; Smith, L.G.; Fuller, F.J.; Guy, J.S. (Raleigh, N.C.)

166 Problems in the Treatment of Hepatitis C with Interferon Iino, S. (Kawasaki)

Human Cytomegalovirus Stimulates Cellular Dihydrofolate Reductase Activity in Quiescent Cells Lembo, D.; Gribaudo, G.; Cavallo, R.; Riera, L.; Angeretti, A.; Hertel, L.; Landolfo, S. (Torino)

173 Hepatitis Delta Virus Taylor, J.M. (Philadelphia, Pa.)

Human Immunodeficiency Virus Type 2 Infection in Spain

179 Hepatitis E Virus. Advances in HEV Biology and HEV Vaccine

Machuca, A.; Soriano, V.; Gutiérrez, M.; Holguín, A. (Madrid); Aguilera, A. (Santiago); Caballero, E. (Barcelona); Cilla, G. (San Sebastián); and The HIV-2 Spanish Study Group

43

Nassal, M. (Freiburg)

117 DNA Vaccines Encke, J. (Boston, Mass./Heidelberg); zu Putlitz, J. (Boston, Mass./Freiburg); Wands, J.R. (Boston, Mass./Providence, R.I.)

T Cell-Mediated and Non-Specific Inflammatory Mechanisms Contribute to the Skin Pathology of HPV 16 E6E7 Transgenic Mice Hilditch-Maguire, P.A. (Woolloongabba/Charlestown, Mass.); Lieppe, D.M. (Madison, Wisc.); West, D. (Woolloongabba); Lambert, P.F. (Madison, Wisc.); Frazer, I.H. (Woolloongabba)

Approaches Yarbough, P.O. (Redwood City, Calif.)

185 GB Virus C/Hepatitis G Virus Kiyosawa, K.; Tanaka, E. (Matsumoto) 196 A Novel Unenveloped DNA Virus (TT Virus) Associated with

Acute and Chronic Non-A to G Hepatitis Okamoto, H.; Nishizawa, T.; Ukita, M. (Tochigi-Ken)

205 Hepatitis C Virus and Hepatocarcinogenesis Hayashi, J.; Aoki, H.; Arakawa, Y.; Hino, O. (Tokyo)

Short Communication 51

Characterization of Potential Insertion Sites in the Core Antigen of Hepatitis B Virus by the Use of a Short-Sized Model Epitope Lachmann, S.; Meisel, H.; Muselmann, C.; Koletzki, D.; Gelderblom, H.R. (Berlin); Borisova, G. (Riga); Krüger, D.H. (Berlin); Pumpens, P. (Riga); Ulrich, R. (Berlin)

211 Author Index 212 Subject Index

No. 4 Original Papers

No. 2–3

213 Delayed Expression of Adeno-Associated Virus Vector DNA Afione, S.A. (Baltimore, Md.); Wang, J. (Gainesville, Fla.); Walsh, S.; Guggino, W.B. (Baltimore, Md.); Flotte, T.R. (Gainesville, Fla.)

Viral Hepatitis Update

221 PCR-RFLP-Detected Human Papilloma Virus Infection in a

Guest Editors: Iino, S. (Kawasaki); Hino, O. (Tokyo)

Group of Senegalese Women Attending an STD Clinic and Identification of a New HPV-68 Subtype Astori, G.; Beltrame, A.; Pipan, C. (Udine); Raphenon, G. (Dakar); Botta, G.A. (Udine)

61 Editorial

Human Hepatitis Viruses – From Basic Research to Treatment Iino, S. (Kawasaki); Hino, O. (Tokyo)

63 Hepatitis A Virus Proteins Totsuka, A.; Moritsugu, Y. (Tokyo) 69 Hepatitis B Virus Mutants Brunetto, M.R.; Rodriguez, U.A.; Bonino, F. (Pisa)

228 Truncated Hepatitis B Virus RNA in Human Hepatocellular

Carcinoma: Its Representation in Patients with Advancing Age Kairat, A. (Heidelberg); Beerheide, W.; Zhou, G. (Heidelberg/Shanghai); Tang, Z.-Y. (Shanghai); Edler, L.; Schröder, C.H. (Heidelberg)

238 Intracellular Replication of Choleraphage w92 Barman, S.; Majumdar, S. (Calcutta) 247 Co-Infection of Acyclovir-Resistant and Acyclovir-Sensitive

Herpes simplex Type 2 Virus Strains in BS-C-1 Cells Keywan, K.; Katz, E. (Jerusalem)

ABC




252 Hepatitis E Virus: An Update on Its Molecular, Clinical and

Epidemiological Characteristics Irshad, M. (New Delhi)

263 A Precipitation Reaction Found in Patients with Hepatitis C

as a Marker for the Purification of Virus-Like Particles

CMV-Induced Pathomechanisms 325 Viral Inhibition of Interferon Signal Transduction Cebulla, C.M.; Miller, D.M.; Sedmak, D.D. (Columbus, Ohio) 331 Murine Cytomegalovirus Homologues of Cellular

Immunomodulatory Genes

Kim, C.Y.; Yoon, J.-H.; Kim, J.S.; Kim, Y.T.; Jung, H.C.; Lee, H.-S.; Yoon, Y.B.; Song, I.S. (Seoul)

Davis-Poynter, N.J. (Newmarket); Degli-Esposti, M. (Nedlands); Farrell, H.E. (Newmarket)

342 Molecular Mimicry by Cytomegaloviruses. Function of

Letter to the Editor

Cytomegalovirus-Encoded Homologues of G Protein-Coupled Receptors, MHC Class I Heavy Chains and Chemokines

271 A Unified Hypothesis for the Etiology of Epidemic Neuropathy de la Fuente, J. (Havana)

Vink, C.; Beisser, P.S.; Bruggeman, C.A. (Maastricht)

350 Cytomegalovirus-Induced Transendothelial Cell Migration.

A Closer Look at Intercellular Communication Mechanisms Scholz, M.; Blaheta, R.A.; Vogel, J.-U.; Doerr, H.W.; Cinatl, J., Jr. (Frankfurt am Main)

No. 5–6

357 Altered Expression of Extracellular Matrix in Human-

Cytomegalovirus-Infected Cells and a Human Artery Organ Culture Model to Study Its Biological Relevance

Novel Immunological Aspects of CMV-Related Diseases Pathogenesis, Diagnosis, and Therapy

Schaarschmidt, P.; Reinhardt, B.; Michel, D.; Vaida, B.; Mayr, K.; Lüske, A.; Baur, R.; Gschwend, J.; Kleinschmidt, K. (Ulm); Kountidis, M.; Wenderoth, U. (Heidenheim); Voisard, R.; Mertens, T. (Ulm)

Guest Editors: Bruggeman, C.A. (Maastricht); Doerr, H.W. (Frankfurt am Main); Ramon, A. (Maastricht); Scholz, M. (Frankfurt am Main)

365 Human Cytomegalovirus Infection of Immature Dendritic Cells

and Macrophages

276 Acknowledgements 277 Preface Bruggeman, C.A.; Ramon, A. (Maastricht); Scholz, M. (Frankfurt am Main)


Jahn, G.; Stenglein, S.; Riegler, S.; Einsele, H.; Sinzger, C. (Tübingen)

Part II. Diagnostics and Antiviral Therapy Diagnostics 373 Diagnostic Value of Nucleic-Acid-Sequence-Based

Amplification for the Detection of Cytomegalovirus Infection in Renal and Liver Transplant Recipients

279 Time-Related Effects of Cytomegalovirus Infection on the

Development of Chronic Renal Allograft Rejection in a Rat Model Lautenschlager, I.; Soots, A.; Krogerus, L.; Inkinen, K.; Kloover, J.; Loginov, R.; Holma, K.; Kauppinen, H.; Bruggeman, C.; Ahonen, J. (Helsinki/Maastricht)

Goossens, V.J.; Blok, M.J.; Christiaans, M.H.L.; van Hooff, J.P. (Maastricht); Sillekens, P. (Boxtel); Höckerstedt, K.; Lautenschlager, I. (Helsinki); Middeldorp, J.M. (Boxtel); Bruggeman, C.A. (Maastricht)

382 Towards Standardization of the Human Cytomegalovirus

Antigenemia Assay

285 Overcoming the Problem of Cytomegalovirus Infection after

Verschuuren, E.A.M.; Harmsen, M.C.; Limburg, P.C.; van der Bij, W.; van den Berg, A.P.; Kas-Deelen, A.M.; Meedendorp, B.; van Son, W.J.; The, T.H.; The Biomed 2 Study Group (Groningen)

Organ Transplantation: Calling for Heracles? van Son, W.J.; de Maar, E.F.; van der Bij, W.; van den Berg, A.P.; Verschuuren, E.A.M.; The, T.H. (Groningen)

390 New Advances in the Diagnosis of Congenital Cytomegalovirus

291 Clinical Significance of Cytomegalovirus-Specific T Helper

Infection

Responses and Cytokine Production in Lung Transplant Recipients Zeevi, A.; Spichty, K.; Banas, R.; Cai, J.; Donnenberg, V.S.; Donnenberg, A.D.; Ahmed, M.; Dauber, J.; Iacono, A.; Keenan, R.; Griffith, B. (Pittsburgh, Pa.)

Lazzarotto, T.; Varani, S.; Gabrielli, L.; Spezzacatena, P.; Landini, M.P. (Bologna)

398 Significance of Qualitative Polymerase Chain Reaction

Combined with Quantitation of Viral Load in the Diagnosis and Follow-Up of Cytomegalovirus Infection after Solid-Organ Transplantation

Immune Escape and Reactivation 301 Human Cytomegalovirus Escape from Immune Detection Michelson, S. (Paris)

Vanpoucke, H.; Van Vlem, B.; Vanholder, R.; Van Renterghem, L. (Gent)

405 Viral Dynamics during Active Cytomegalovirus Infection and

Pathology

308 Human Cytomegalovirus Reactivation in Bone-Marrow-Derived

Emery, V.C. (London)

Granulocyte/Monocyte Progenitor Cells and Mature Monocytes Prösch, S.; Döcke, W.-D.; Reinke, P.; Volk, H.-D.; Krüger, D.H. (Berlin)

Antivirals

314 Human Cytomegalovirus Latency and Reactivation –

A Delicate Balance between the Virus and Its Host’s Immune System

412 Inhibition of Cytomegalovirus in vitro and in vivo by the

Experimental Immunosuppressive Agent Leflunomide

Söderberg-Nauclér, C. (Stockholm); Nelson, J.A. (Portland, Oreg.)

Waldman, W.J.; Knight, D.A. (Columbus, Ohio); Blinder, L.; Shen, J.; Lurain, N.S. (Chicago, Ill.); Miller, D.M.; Sedmak, D.D. (Columbus, Ohio); Williams, J.W.; Chong, A.S.-F. (Chicago, Ill.)

322 Measurement of Anti-Human Cytomegalovirus T Cell

Reactivity in Transplant Recipients and Its Potential Clinical Use: A Mini-Review

419 Proinflammatory Potential of Cytomegalovirus Infection.

Specific Inhibition of Cytomegalovirus Immediate-Early Expression in Combination with Antioxidants as a Novel Treatment Strategy?

Kern, F.; Faulhaber, N.; Khatamzas, E.; Frömmel, C.; Ewert, R.; Prösch, S.; Volk, H.-D.; Reinke, P. (Berlin)

Cinatl, J., Jr.; Vogel, J.-U.; Kotchetkov, R.; Scholz, M.; Doerr, H.W. (Frankfurt am Main)

425 426 427 429

IV


Author Index Vol. 42, No. 5–6, 1999 Subject Index Vol. 42, No. 5–6, 1999 Author Index Vol. 42, 1999 Subject Index Vol. 42, 1999

Contents

Novel Immunological Aspects of CMV-Related Diseases: Pathogenesis, Diagnosis and Therapy (Intervirology, 5-6)

Mitochondrial medicine: diseases, diagnosis and therapy

The Diversity of Gastric Carcinoma: Pathogenesis, Diagnosis and Therapy

Immunological and molecular diagnosis of infectious disease

Autoimmune Diseases of the Skin: Pathogenesis, Diagnosis, Management, Third Edition

Autoimmune Diseases of the Skin Pathogenesis, Diagnosis, Management

Autoimmune Diseases of the Skin. Pathogenesis. Diagnosis, Management

Manual of Diagnosis and Therapy

alzheimers disease its diagnosis and pathogenesis

Molecular Diagnosis Of Genetic Diseases

Translational Vascular Medicine: Pathogenesis, Diagnosis, and Treatment

Malignant Mesothelioma: Pathogenesis, Diagnosis, and Translational Therapies

Translational Vascular Medicine: Pathogenesis, Diagnosis, and Treatment

The Diagnosis of Lymphoproliferative Diseases

Molecular Diagnosis of Infectious Diseases

Allergic diseases: diagnosis and treatment

Molecular Diagnosis of Genetic Diseases

Diagnosis Aging Skin Diseases

Pediatric Epilepsy-Diagnosis and Therapy

Aspects of the Novel

Epigenetic Aspects of Chronic Diseases

Aspects of the Novel

Epigenetic Aspects of Chronic Diseases

Signaling Pathways in Cancer Pathogenesis and Therapy

Aspects of the Novel

Caveolins in Cancer Pathogenesis, Prevention and Therapy

Handbook of Urology Diagnosis and Therapy

Neuroblastoma - Diagnosis, Therapy, and Prognosis

Neurochemical Aspects of Neurotraumatic and Neurodegenerative Diseases

Manual of Gastroenterology: Diagnosis and Therapy

Novel Immunological Aspects of CMV-Related Diseases: Pathogenesis, Diagnosis and Therapy (Intervirology, 5-6)

Mitochondrial medicine: diseases, diagnosis and therapy

The Diversity of Gastric Carcinoma: Pathogenesis, Diagnosis and Therapy

Immunological and molecular diagnosis of infectious disease

Autoimmune Diseases of the Skin: Pathogenesis, Diagnosis, Management, Third Edition

Autoimmune Diseases of the Skin Pathogenesis, Diagnosis, Management

Autoimmune Diseases of the Skin. Pathogenesis. Diagnosis, Management

Manual of Diagnosis and Therapy

alzheimers disease its diagnosis and pathogenesis

Molecular Diagnosis Of Genetic Diseases

Translational Vascular Medicine: Pathogenesis, Diagnosis, and Treatment

Malignant Mesothelioma: Pathogenesis, Diagnosis, and Translational Therapies

Translational Vascular Medicine: Pathogenesis, Diagnosis, and Treatment

The Diagnosis of Lymphoproliferative Diseases

Molecular Diagnosis of Infectious Diseases

Allergic diseases: diagnosis and treatment

Molecular Diagnosis of Genetic Diseases

Diagnosis Aging Skin Diseases

Pediatric Epilepsy-Diagnosis and Therapy

Aspects of the Novel

Epigenetic Aspects of Chronic Diseases

Aspects of the Novel

Epigenetic Aspects of Chronic Diseases

Signaling Pathways in Cancer Pathogenesis and Therapy

Aspects of the Novel

Caveolins in Cancer Pathogenesis, Prevention and Therapy

Handbook of Urology Diagnosis and Therapy

Neuroblastoma - Diagnosis, Therapy, and Prognosis

Neurochemical Aspects of Neurotraumatic and Neurodegenerative Diseases

Manual of Gastroenterology: Diagnosis and Therapy

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