University of Groningen Importance of molecular diagnostic of viral infections in renal transplant recipients Rurenga-Gard, Lilli

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Importance of molecular diagnostic of viral infections in renal transplant recipients

Rurenga-Gard, Lilli

DOI:

10.33612/diss.92267443

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Rurenga-Gard, L. (2019). Importance of molecular diagnostic of viral infections in renal transplant

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infections in renal transplant recipients

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L. Gard

ISBN: 978-94-028-1593-1

All rights reserved. No part of this thesis may be reproduced, stored or transmitted in any way or by any means without the prior permission of the author, or when applicable, of the publishers of the scientific papers.

Cover layout by Lilli Gard, inspired by Fritz Gard. Stockfoto ID: 511064444 Layout and design by Anna Bleeker, persoonlijkproefschrift.nl.

Printed by Ipskamp Printing, proefschriften.net

Financial support for the publication of this thesis was kindly provided by the University Medical Center Gronigen and Graduate School of Medical Sciences.

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Importance of molecular diagnostic

of viral infections in renal

transplant recipients

Proefschrift

ter verkrijging van de graad van doctor aan de Rijksuniversiteit Groningen

op gezag van de

rector magnificus prof. dr. C. Wijmenga en volgens besluit van het College voor Promoties.

De openbare verdediging zal plaatsvinden op woensdag 4 september 2019 om 14.30 uur

door

Lilli Gard

geboren op 29 november 1981 te Hamburg, Duitsland

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Prof. dr. W.J. van Son

Copromoteres

Dr. A. Riezebos-Brilman Dr. J.S. Sanders

Beoordelingscommissie

Prof dr A.C.M. Kroes, UMC Leiden Prof.dr. T van der Werf, UMCG Prof.dr. S. Bakker, UMCG

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Chapter 1 General Introduction 8

Chapter 2 A real time genotyping PCR assay for polyomavirus BK Journal virological methods 2015 Sep 1;221:51-6

28

Chapter 3 Longitudinal monitoring of BKPyV miRNA levels in renal transplant recipients with BKPyV related pathology reflects viral DNA levels and remains high in viremia patients after clearance of viral DNA. Manuscript in preparation

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Chapter 4 A delicate balance between rejection and BK polyomavirus associated nephropathy; a retrospective cohort study in renal transplant recipients.

PLoS One 2017 Jun 13;12(6):e0178801

60

Chapter 5 Incidence and outcome of BK polyomavirus infection in a multicenter randomized controlled trial with renal transplant patients receiving cyclosporine- mycophenolate sodium- or everolimus-based low-dose immunosuppressive therapy.

Transplant Infectious Disease 2017 Jun ;19(3)

84

Chapter 6 High Human Cytomegalovirus DNAemia Early Post-Transplantation Associates With Irreversible And Progressive Loss of Renal Function – a retrospective study Transplant

International 2017 Aug;30(8): 817-826

104

Chapter 7 Relevance of EBV load monitoring in renal transplant recipients; a retrospective cohort study

Manuscript submitted for publication

126

Chapter 8 Summary, discussion and future perspectives 144

Chapter 9 Nederlandse samenvatting 160

Chapter 10 Deutsche Zusammenfassung 170

Chapter 11 Dankwoord 180

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CHAPTER 1

General Introduction

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8 INTRODUCTION

Renal transplant recipients

For patients with end-stage renal disease, renal transplantation provides significant survival and quality-of-life benefits over dialysis. In the last decades graft survival in adult recipients has increased, due to improved surgical techniques, development of new and more potent immunosuppressive drugs and better post-transplantation care. However, worldwide waiting lists are growing partly because people are getting older 1_{. The use of expanded criteria donors}

(ECD) could contribute to reducing the waiting list. ECDs are donors over 60 years of age, having a history of hypertension, creatine level >1.5mg/dL or cerebrovascular cause of death. Although the risk of graft failure in recipients from ECD is >70% higher than after living or deceased donors, it still benefits over dialysis 2,3_{. Since the last century, the use of ECD has}

become common practice.

In the Netherlands, the number of patient on the waiting list, for kidney transplantation, increased with 9.0% in 2017 (n= 650) compared to 2016 (n= 595). Before 2017 a steady decrease was observed with 622 patients on the waiting list in 2014 and 544 patients in 2015 4_.

Immunosuppressiva

Over the last decades immunosuppressive drugs and management has improved, which has led to a transplant survival rate of 95% in the first year after transplantation. Especially, less acute rejection (<15%) is seen in the first year, while long-term survival rates and rejection episodes remained unaltered mainly due to the toxic side effects of the immunosuppressants and increased acceptance of more marginal donors and recipients 5,6_.

The regimens after kidney transplantation consist of a combination of different immunosuppressive drugs, as they have synergistic efficacy leading to lower dosages of individual drugs. Most transplantation centers use quadruple therapy, with an interleukin-2 receptor antibody for induction followed by immunosuppressive maintenance therapy (i.e. calcineurin inhibitor, mycophenolate and steroids). The function of induction therapy is to reduce the risk of rejection. Mostly basiliximab is used as interleukin-2 receptor antagonist, while T cell depleting agents as ATG, thymoglobulin or alemtuzumab are being used in high risk patients 6_{. Maintenance therapy can be categorized into calcineurin inhibitors (cyclosporine}

A, tacrolimus), costimulation blockers (betacepts), mammalian target of rapamycin inhibitors (sirolimus, everolimus), antiprofileratives (azathioprine, mycophenolic acid derivatives) and corticosteroids.

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To improve long-term outcome selection of appropriate immunosuppressive regimens is important and ideally should be patient tailored. Taking the patients pre-existing disease, risk of rejection, drug to drug interaction and interaction with other medication 6,7_{into consideration.}

Non-infectious complications after kidney transplantation

Short term complications of organ and patient survival after renal transplantation have decreased over the last decade, whereas long term (beyond 5 years) complications have remained unaltered 5,8_{. This progress is mostly the result of the introduction and use of new}

and more potent immunosuppressive drugs in the last two decades 9_{. The drawbacks, however,}

are the side effects, such as nephrotoxicity, development of diabetes, hypertension, increased susceptibility to tumor formation and infection 10,11_.

One of the major complications after renal transplantation is rejection. It can be classified in different stadia; hyper acute (occurring within minutes), acute (occurring within days to weeks), late acute (occurring after 3 months) or chronic (occurring months to years after transplantation)

5,12_{. Besides the different stadia, rejection can be divided into antibody-mediated or T-cell}

mediated rejection. The last one has become less frequent through the usage of effective T-cell directed induction/immunosuppression and is mostly reversible with limited impact on long-term outcome 9,13_{. Antibody-mediated rejection results more in graft loss, and no}

adequate immunosuppressive treatment is currently available. To reduce the immunological risk of humoral rejection 14_{, it is important to identify the HLA antibodies of the recipient and}

donor before and after transplantation 13_.

Whereas short term outcome has improved long-term outcome is still similar and an important long-term complication is chronic allograft dysfunction. Causes of allograft dysfunction could be immunological as highlighted above or non-immunological (chronic CNI toxicity, infection, chronic obstruction, recurrent or de novo glomerular diseases, recurrent or de novo diabetic nephropathy, hypertensive nephrosclerosis and renal artery stenosis). Chronic allograft dysfunction occurs slowly with progressive loss of renal function and is mainly diagnosed through slow rising serum creatinine levels, increasing proteinuria and worsening hypertension

15,16_.

Infectious complications

Adequate transplant function, hence prevention of organ rejection, depends on lifelong immunosuppression, making transplant recipients more prone for opportunistic infections, resulting in disease and mortality 17_.

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The highest risk of infections is during the first six months after transplantation, due to the peak of immunosuppression in this period 18_{. Viral and non-viral infections can occur}

simultaneously. Most common post-transplant opportunistic infections can be divided in viral (CMV, HSV, VZV, EBV and BKPyV), fungal (Candida species, Aspergillus species, Pneumocystis jiroveci, C. neoformans, and mucormycosis), bacterial (Nocardia species, L. pneumophila, L. monocytogenes, and mycobacteria) and parasitic (T. gondii and strongyloidiasis) 19_.

Many viral infections are the result of reactivation of latent viruses from either within the host or from the graft. Beside reactivations of latent viruses, also viral infections through community exposure may occur 19_.

Diagnosis is mainly through testing and imaging, as clinical signs and symptoms of these infections may overlap and are often atypical, subtle or absent 20_{. Pre-transplant, all potential}

donors and recipients are being screened, serologically, for most common viral pathogens . These results can subsequently be used to determine the appropriate prophylaxis and preventive strategies, i.e. update vaccination status and educate patient and family about preventing measures. Post-transplant screening by real-time polymerase chain reaction (real-time PCR) for viral pathogens is performed to detect and subsequently treat active infections.19_.

To obtain the best patient and graft outcome, it is challenging for the treating clinical physicians, to balance between over and under immunosuppression.

The most common viral infections in renal transplant recipients are with EBV, CMV and BKPyV. Therefore, the following paragraphs will focus on these viruses.

EPSTEIN-BARR VIRUS

Epstein-Barr virus (EBV) belongs to the gammaherpesviruses and is an ubiquitous DNA virus that infects up to 90% of the world’s population, mostly during childhood or early adolescence

21_{. Infection is often asymptomatic in immunocompetent individuals, but may result in a}

self-limiting, benign lymphoproliferative syndrome called infectious mononucleosis (Pfeiffer).

Following a primary infection, EBV establishes lifelong latency in memory B-cells. Asymptomatic reactivation in immunocompetent individuals is common 22_{. Cell-mediated immunity is}

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EBV infection

In solid organ transplant (SOT) patients, EBV-specific cell-mediated immunity is depressed by the use of immunosuppression, which subsequently may lead to the development of uncontrolled EBV-driven B-cell proliferation and malignant transformation. In addition, suppressive cytokines such as IL-10 have also been implicated as initiative factors supporting the development of this post-transplant lymphoproliferative disorder (PTLD) after SOT 24_{. It is a serious and often}

fatal complication, with mortality rates up to 70%. Of the PTLDs in immunosuppressed SOT recipients, 90% are EBV positive 25

PTLD may present itself with a diverse spectrum of clinical symptoms and signs. Symptoms may be similar to those seen with primary infection, whereas also other organs including the central nerves system, bone marrow and internal organs may be involved. PTLD may arise at any given time after transplantation. The highest incidence, however, is in the first year after SOT, i.e. early-onset PTLD, hence during the most immunosuppressed state, and is highly associated with a primary EBV infection. Late-onset PTLD is independent of the EBV sero-status (primary infection or reactivation) or immune suppression.

Overall incidence of PTLD is 1-2%. However, this incidence strongly depends upon the pre-transplantation EBV sero-status, the type of organ transplanted, and the immunosuppressive regimen. EBV naive recipients with a EBV positive donor have a 10 to 75-fold increased incidence of developing PTLD compared to EBV sero-positive recipients. This is due to the fact that they are unable to initiate an adequate EBV-specific CTL response 26_{. The incidence of PTLD varies}

between different SOT recipients and is relatively low in renal transplant recipients (RTR) (1-5%) compared to heart and lung (2-10%) and in intestinal and multivisceral transplant recipients (5-20%) 27_{. These percentages may be a reflection of the intensity of immunosuppressive regimens}

needed in these different groups and the quantity of lymphoid tissue present within the transplanted organs. As for the immunosuppressive regimen, the use of tacrolimus may give a 2 to 5 fold increase in the risk of developing PTLD 28-30_{. Use of OKT3 or ATG for prophylaxis against}

or treatment of acute rejection was associated with a 3 to 4 fold increase in the incidence of PTLD 29_.

Diagnosing EBV infection

There are several non-invasive diagnostic assays which can be used in diagnosing of EBV infection or PTLD development in renal transplant recipients. First, serological assays are most useful for determining the pre-transplantation sero-status. For this, enzyme-linked immunosorbent assays (ELISAs) or commercially available EBV enzyme immunoassays (EIAs) are available 31_{. Secondly, after transplantation the detection of EBV DNA viral load in whole}

blood, plasma or EBV genome in peripheral blood mononuclear cells by real-time PCR has been

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developed to aid in diagnosing an EBV infection and monitor the development of PTLD. The best assays use targets in a highly conserved ORF with no or few false-negatives 32_.

The implications of EBV viremia, however, after the first year of renal transplantation are not clear. Retrospective studies reporting on the prevalence, patterns and long term outcome of RTR demonstrate divergent associations of EBV viremia with graft loss, biopsy proven acute rejection (BPAR), renal function and other infections 33-35_{Recent studies focus on the progress}

on how EBV utilizes miRNA’s for immune invasion. These insights could help in the developing of using EBV miRNA’s as a new prognostic marker for risk stratification for PTLD 36,37_.

Besides EBV-diagnostics, 18F-FDG-PET scanning is now standard for PTLD staging by visualizing malignant lymphoma’s. 38_{The final diagnosis, however, is always based on histopathological}

examination. PTLD can be classified into four histological types ( early lesions, P-PTLD, monomorphic PTLD and classical Hodgkin lymphoma), whereby the first three are most common and develop the typical morphology 39_.

Treatment of EBV infection

Once a primary infection with EBV or reactivation is determined, several interventions could be considered. First-line treatment involves reduction of immunosuppressive therapy to restore the cell-mediated immune response, with the unfortunate consequence of increasing the risk of graft rejection. The immunosuppressive protocol can be switched to other immunosuppressive drugs, with potentially less lymphoma outgrowth. Several studies suggest that mTOR inhibitors-based immunosuppression, i.e. everolimus and sirolimus, displays potent inhibitory effect on PTLD-derived cells with maintenance of potent immunosuppressive properties 40-43_.

The treatment strategy which is standard of care in treating PTLD is the use of a chimeric human/mouse anti-CD20 monoclonal antibody, Rituximab. Since most of the PTLD is of B-cell lineage, the use of monoclonal antibodies specific for B-cells is a logical therapeutic approach. Rituximab binds to the CD20 antigen present on malignant B-cells, resulting in profound and long-lasting depletion of all B-cells (up to12 months) 44_{and it is also able to elicit strong}

anti-B-cell tumor responses. Mostly, it is given in adjunction to reduction of immunosuppression or chemotherapy, but also as first line treatment of PTLD. Results of treatment demonstrate a high efficacy 45_.

Although conclusive evidence of clinical utility of antiviral prophylaxis or therapy is lacking, anti-viral drugs are often considered, especially in recipients with a primary infection with EBV. Acyclovir and ganciclovir are capable of inhibiting lytic viral replication, suggesting that by diminishing the EBV viral load, the risk of PTLD may be reduced in EBV-naïve recipients 46,47_.

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Another way to restore the cell-mediated immune response is to administer either allogeneic or autologous cultured in vitro EBV-specific CTLs. Several preliminary clinical studies show that treatment with autologous EBV-specific CTLs may well be a very promising strategy, especially in EBV-seropositive recipients. These studies have demonstrated a considerable fall in EBV viral loads, a temporary increase in EBV-specific CTL precursors, and up to complete regression of established PTLD 48-50_{. More recently, Haque et al. demonstrated encouraging results in a phase}

II multicenter clinical trial using HLA-matched allogeneic CTLs for the treatment of PTLD. Upon treatment, a 52% response rate at six months was shown 51_.

EBV prevention strategies

A way to prevent EBV infection would be vaccination. Especially in EBV seronegative recipients, mostly children, exposure to EBV prior to transplantation is currently under investigation 52-54_.

An EBV vaccine is not yet available for general use and whether these vaccines are able to induce a sufficient immune-response in often already immunocompromised patients prior to transplantation, remains to be investigated 55,56_.

CYTOMEGALOVIRUS

Cytomegalovirus (CMV) is the fifth member of the human herpesvirus family and belongs to the betaherpesviruses. Seroprevalence of CMV in the adult population approaches 60% and increases with age 57,58_{. Most commonly, CMV is acquired during childhood to early adulthood.}

In immune-competent individuals, a primary infection usually manifests as a benign disease. After a primary infection, CMV establishes latency and is able to persist in numerous cellular sites, leukocytes, endothelial cells, renal epithelial cells and salivary glands 59_{. CMV infection is}

the most common opportunistic infection after renal transplantation. In absence of preventive measures, 40%-100% will develop CMV infection and 67% will develop CMV disease..

CMV infection

Lack of an effective CMV specific immunity predisposes RTR to develop CMV infection and CMV disease. Depending on the immunologic status of the transplant recipient, infection either occurs by reactivation of latent virus in response to immunosuppression or as primary infection due to donor-positive-recipient-negative mismatch 60_.

CMV infection comprises both direct and indirect effects. The direct effects, also known as CMV disease, are associated with high viral loads and can be further categorized as CMV syndrome or as tissue-invasive disease. CMV syndrome may manifest as mononucleosis-like syndrome with fever, malaise and myelosuppression. In about half of the cases, CMV invades an organ system and subsequently causes tissue-invasive disease. This may present as a gastrointestinal

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disease (colitis), pneumonitis, hepatitis, myocarditis, retinitis, with a predilection for the transplanted organ. As opposed to the direct effects, the indirect effects are a result of the immunomodulatory effects of CMV infection and independent of active CMV disease. These effects may present, in RTR, as graft rejection or dysfunction, opportunistic infections, co-infection with other viruses, development of cancer or PTLD and increased risk of mortality 61,62_.

Most frequently, CMV disease occurs within the first three months post-transplant 63_{. However,}

the current use of antiviral profylaxe against CMV in patients at risk, has resulted in a delayed onset of CMV disease. This so-called late-onset CMV infection occurs after the discontinuation of prophylaxis 61,64_{. It is most significant in seronegative recipient in combination with a}

seropositive donor; up to 30% develop late-onset CMV disease at a median of five months post-transplant 65,66_.

Diagnosing CMV infection

The diagnosis of CMV infection and disease has improved over the years, first by the introduction of the pp65 antigenemia assay and subsequently the quantitative real-time PCR assay. The antigenemia assay is a semiquantitative assay and can provide an estimate of the magnitude of viral load by counting the number of infected cells 67_{. However, the real-time}

PCR assays represent a major advance in the diagnosis of CMV, through the ability to quantify the amounts of CMV in a broad range of clinical samples which can be stored beforehand, the short turnaround time, and the reduced risk of contamination using a closed system. Results demonstrate that real-time PCR from whole blood is superior, for detection, monitoring and also for initiation of treatment, to either real-time PCR in plasma or peripheral blood leukocytes

66,68_.

CMV serology is useful to determine the CMV status during pre-transplantation screening of recipients and donors to determine the risk of either primary CMV infection or reactivation. Furthermore, seroconversion, in a formally seronegative RTR, can serve as a marker for immune protection against CMV, and may lower the risk of later CMV disease 69_{. Yet, due to}

the immunosuppressive medication, the role of serologic testing post-transplant is debated.

Another method is the detection of CMV specific cell-mediated immunity (CMI), as this has the ability to control viral replication. Detection of CMI can help to determine individuals at risk of an infection and to use as an indicator in therapeutic decisions. Several tests are available for measuring CMI, but only the Enzyme-linked Immunosorbent Spot (Elispot) and the Quantiferon Gamma Interferon-releasing assay are practical for diagnostic use. These tests are standardized, cost-effective and rapid, compared to the other tests, which are only used in specialized laboratories 70,71_.

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For diagnosing tissue-invasive disease histopathological examination of biopsy specimens can be used. By using a hematoxylin-eosin stain the presence of giant cells with typical intracellular viral inclusions, characteristic “owl’s-eye”inclusion bodies, may be demonstrated 72_{. This}

technique has a high specificity for disease, and hence a low sensitivity 73_.

Treatment of CMV infection

Current therapy for CMV disease consists of ganciclovir, foscarnet, cidofovir, valganciclovir, hyperimmune globulin, or combinations thereof. Foscarnet and cidofovir are being considered second-line therapies due to their associated toxicities. The first line of treatment for CMV disease remains ganciclovir intravenously, while in mildly asymptomatic cases of CMV disease and also in a prophylactic setting, valganciclovir has shown to be a effective alternative 74_.

Upon treatment, the viral load of CMV should decline. A “nonresponding” viral load may be a marker of drug-resistant CMV. To date, the incidence of ganciclovir-resistant CMV is low 0%-3% in recipients with CMV prophylaxis. In the population with a positive CMV donor and negative for CMV recipient the incidence is 5-12% 75-77_{. Resistant strains can be confirmed using genotypic}

and/or phenotypic methods. Most commonly, a mutation is found within the CMV UL97 (protein kinase) gene, making CMV resistant to ganciclovir, however, the virus remains susceptible to foscarnet and cidofovir. In the case of a mutation in UL54 (DNA polymerase) gene, CMV is resistant to all before mentioned drugs 78,79_.

Prevention of CMV

There are two main strategies to prevent CMV disease, universal prophylaxis and preemptive therapy. Universal prophylaxis is usually directed in patient where baseline risk of disease is high. The antiviral drugs are given from the time of transplant onwards, mostly for duration of three to six months.

The second strategy, preemptive therapy, is mainly reserved for patients at imminent risk for developing CMV disease. Peripheral blood samples are monitored on a weekly basis for the presence of CMV. Treatment is initiated after the first detectable CMV viral loads and before the onset of clinical symptoms.

Most transplant institutes give CMV prophylaxis after renal transplantation 80,81_{. Both strategies}

have proven to be effective in preventing CMV disease. However, there is increasing evidence that prophylactic treatment is beneficial and improves long-term graft and patient outcomes

82,83_{. In addition CMV prophylaxis with ganciclovir or valganciclovir suppress HHV6 replication}84_.

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Another issue that should be addressed is the optimal duration of prophylaxis. There is accumulating evidence that extension of the duration (200 days instead of 100 days) of prophylaxis may protect against this late onset CMV disease, by facilitating the development of CMV-specific immune responses 81,85,86_.

BK POLYOMAVIRUS

BK Polyomavirus (BKPyV) is a non-enveloped double-stranded DNA virus that belongs to the Polyomavirus family. It was discovered in 1971 87_{and has been associated primarily with}

nephropathy in RTR.

After a primary infection, which is usually asymptomatic or associated with mild upper respiratory tract symptoms, it remains latent in the epithelial cells of the reno urinary tract. The transmission route is still unknown, but likely involves the oral or respiratory route.

BKPyV is ubiquitous and widely spread in adult human population, whereby infection results in lifetime persistence of the virus, with seropositivity of up to 80% 88,89_{. It is divided into 4}

subtypes. Subtype I is the most prevalent throughout the world, while subtype IV can mostly be found in Asia and in Europe. Subtype II and III are more rare spread 90-92_.

BKPyV infection

Intermittent active replication and asymptomatic viral shedding occurs in up to 20% in immuncompetent individuals and up to 60% in immunocompromised. In immuncompromised individuals, reactivation of BKV can switch from subclinical replication to a lytic infection, which result in viruria in up to 50% and viremia in up to 20% 93_.

BKPyV reactivation is most common in RTR, where BKPyV infection could result in BKPyV nephropathy (BKPyVAN) 94_{. BKPyV viruria and viremia due to reactivation are found}

in, respectively, 80% and 40% of renal transplant patients, mainly in the first year after transplantation, and 1-10% of patients progress to BKPyVAN 95,96_{. Several risk factors are known}

for the developing of BKPyV replication and BKPyVAN, whereby immunosuppression plays an important role 97-99_.

Diagnosing BKPyV

Screening for BKPyV replication can be done by using the detection of decoy cells in urine or by screening urine and plasma viral load using real-time PCR. The last method is widely implemented in the laboratory as screening plasma for BKPyV replication is clinical relevant 100_.

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BKPyV serology is not performed routinely before transplantation, as about 90% of the human population is seropositive. Recent studies demonstrated that BKPyV IgG seroreactivity is stable over time and is associated with BKPyV replication after transplantation 101,102_{. This insight give}

new opportunities to predict patients at risk for developing BKPyVAN.

For the diagnosis of BKPyVAN, a biopsy is taken and by using SV40 staining the characteristic cytopathic changes in the tubular epithelial cells are visualized. This is necessary to differentiate between acute mediated rejection or polyomavirus infection.

A negative result cannot rule out BKPyVAN, therefore it is also necessary to screen the plasma and urine for BKPyV viral load. Urine viral loads of more than log 10 cp/ml and plasma loads of more than log 4 cp/ml are predictive values for developing BKPyVAN.

Recently, it has been demonstrated that BKPyV microRNA’s (miRNA) may play a role in induction of reactivation and the development of disease 103_{. These findings are interesting as they could}

help in identifying new antiviral strategies. Until now, a few clinical studies showed that BKPyV miRNA levels in renal transplant patients can be detected in plasma and urine and in recipients with BKPyVAN 104,105_{. However, other studies are necessary to find out the added value of}

measuring BKPyV mirRNA’s above BKPyV viral load.

Treatment and or prevention of BKPyV

In recipients with BKPyV viremia and risk of BKPyVAN , the first step in treatment is reduction, reduction of the immunosuppressive load. This could be done by reducing first the calcineurin inhibitors followed by reduction of the metabolite or reduce/discontinuous the anti-metabolite and thereafter the calcineurin inhibitors or reduce the anti-anti-metabolite as well as the calcineurin inhibitors 106_{. Although these strategies can lead to reduction of BKPyV plasma}

loads, the risk of transplant rejection increases and it is not effective in all patients 107,108_.

Antiviral agents, such as cidofovir, leflunomide, intravenous immunoglobulin and/or fluoroquinolones have been described.108-111_{. To date, however, the role of these specific antiviral}

therapy remains unclear. Adequately powered trials to define optimal treatment are therefore necessary.

Recent studies demonstrated screening pre-transplantation donors for BKPyV IgG seroreactivity and BKPyV replication could contribute to identify recipients at high risk. This could be used for the choice of the immunosuppressive regimen post-transplantation 101,112_{. Taking in}

consideration that an everolimus based regimen with reduced CNI exposure results in less CMV and BKPyV infection 113_.

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18 SCOPE OF THIS THESIS

The UMCG is a tertiary University Medical Centre which performs around 200 kidney transplantations per year. The implementation of new techniques in transplantations and the introduction of new immunosuppressive drugs, contribute to transplant kidneys not only under less optimal conditions, but also in older, more frail, recipients. This trend is, at one hand, good for the recipients. But at the other hand, infectious related complications increase, which makes screening pre- and post-transplantation more important.

The main focus of this thesis is to gain better insight in the clinical importance of regularly molecular monitoring of BKPyV, CMV and EBV DNA in adult RTR. Several research questions will be addressed in this thesis.

1. Development of additive diagnostic tools to gain more insight in BKPyV infection.

2. What is the optimal strategy and frequency for screening on BKPyV, CMV and EBV viral load after RTR?

3. What is the association between different immunosuppressive regimens and the detection of BKPyV, CMV and/or EBV?

The goal of this thesis is to improve patient management and transplantation outcome, which subsequently contributes to a better outcome of rental transplant recipients.

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19 REFERENCES

1. Hart A, Smith JM, Skeans MA, et al. Kidney. Am J Transplant. 2016;16 Suppl 2:11-46.

2. Ma MK, Lim WH, Craig JC, Russ GR, Chapman JR, Wong G. Mortality among younger and older recipients of kidney transplants from expanded criteria donors compared with standard criteria donors. Clin J Am Soc Nephrol. 2016;11(1):128-136.

3. Knoll GA. Kidney transplantation in the older adult. Am J Kidney Dis. 2013;61(5):790-797. 4. Nederlandse transplantatie stichting N. NTS jaarverslag 2017. . 2017.

5. Nankivell BJ, Alexander SI. Rejection of the kidney allograft. N Engl J Med. 2010;363(15):1451-1462. 6. Paliege A, Bamoulid J, Bachmann F, et al. Immunosuppression and its use in kidney transplantation.

Urologe A. 2015;54(10):1376-1384.

7. Lee RA, Gabardi S. Current trends in immunosuppressive therapies for renal transplant recipients. Am J Health Syst Pharm. 2012;69(22):1961-1975.

8. Moreso F, Hernandez D. Has the survival of the graft improved after renal transplantation in the era of modern immunosuppression? Nefrologia. 2013;33(1):14-26.

9. Bamoulid J, Staeck O, Halleck F, et al. The need for minimization strategies: Current problems of immunosuppression. Transpl Int. 2015;28(8):891-900.

10. Ekberg H, Bernasconi C, Noldeke J, et al. Cyclosporine, tacrolimus and sirolimus retain their distinct toxicity profiles despite low doses in the symphony study. Nephrol Dial Transplant. 2010;25(6):2004-2010.

11. Marcen R. Immunosuppressive drugs in kidney transplantation: Impact on patient survival, and incidence of cardiovascular disease, malignancy and infection. Drugs. 2009;69(16):2227-2243. 12. Sijpkens YW, Doxiadis II, Mallat MJ, et al. Early versus late acute rejection episodes in renal

transplantation. Transplantation. 2003;75(2):204-208.

13. Becker LE, Morath C, Suesal C. Immune mechanisms of acute and chronic rejection. Clin Biochem. 2016;49(4-5):320-323.

14. Loupy A, Lefaucheur C. Antibody-mediated rejection of solid-organ allografts. N Engl J Med. 2018;379(12):1150-1160.

15. Zhang R. Clinical management of kidney allograft dysfunction . 2014;4(2):7-14.

16. Goldberg RJ, Weng FL, Kandula P. Acute and chronic allograft dysfunction in kidney transplant recipients. Med Clin North Am. 2016;100(3):487-503.

17. Katabathina V, Menias CO, Pickhardt P, Lubner M, Prasad SR. Complications of immunosuppressive therapy in solid organ transplantation. Radiol Clin North Am. 2016;54(2):303-319.

18. Fischer SA. Infections complicating solid organ transplantation. Surg Clin North Am. 2006;86(5):1127-45, v-vi.

19. Cukuranovic J, Ugrenovic S, Jovanovic I, Visnjic M, Stefanovic V. Viral infection in renal transplant recipients. ScientificWorldJournal. 2012;2012:820621.

20. Pagalilauan GL, Limaye AP. Infections in transplant patients. Med Clin North Am. 2013;97(4):581-600, x.

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21. Cohen JI. Epstein-barr virus infection. N Engl J Med. 2000;343(7):481-492.

22. Thorley-Lawson DA, Gross A. Persistence of the epstein-barr virus and the origins of associated lymphomas. N Engl J Med. 2004;350(13):1328-1337.

23. Khanna R, Burrows SR. Role of cytotoxic T lymphocytes in epstein-barr virus-associated diseases. Annu Rev Microbiol. 2000;54:19-48.

24. Dierickx D, Habermann TM. Post-transplantation lymphoproliferative disorders in adults. N Engl J Med. 2018;378(6):549-562.

25. Paya CV, Fung JJ, Nalesnik MA, et al. Epstein-barr virus-induced posttransplant lymphoproliferative disorders. ASTS/ASTP EBV-PTLD task force and the mayo clinic organized international consensus development meeting. Transplantation. 1999;68(10):1517-1525.

26. Cockfield SM. Identifying the patient at risk for post-transplant lymphoproliferative disorder. Transpl Infect Dis. 2001;3(2):70-78.

27. Morscio J, Dierickx D, Tousseyn T. Molecular pathogenesis of B-cell posttransplant lymphoproliferative disorder: What do we know so far? Clin Dev Immunol. 2013;2013:150835.

28. Cao S, Cox KL, Berquist W, et al. Long-term outcomes in pediatric liver recipients: Comparison between cyclosporin A and tacrolimus. Pediatr Transplant. 1999;3(1):22-26.

29. Opelz G, Dohler B. Lymphomas after solid organ transplantation: A collaborative transplant study report. Am J Transplant. 2004;4(2):222-230.

30. Younes BS, McDiarmid SV, Martin MG, et al. The effect of immunosuppression on posttransplant lymphoproliferative disease in pediatric liver transplant patients. Transplantation. 2000;70(1):94-99. 31. Bruu AL, Hjetland R, Holter E, et al. Evaluation of 12 commercial tests for detection of epstein-barr

virus-specific and heterophile antibodies. Clin Diagn Lab Immunol. 2000;7(3):451-456.

32. Wadowsky RM, Laus S, Green M, Webber SA, Rowe D. Measurement of epstein-barr virus DNA loads in whole blood and plasma by TaqMan PCR and in peripheral blood lymphocytes by competitive PCR. J Clin Microbiol. 2003;41(11):5245-5249.

33. Bamoulid J, Courivaud C, Coaquette A, et al. Subclinical epstein-barr virus viremia among adult renal transplant recipients: Incidence and consequences. Am J Transplant. 2013;13(3):656-662.

34. Morton M, Coupes B, Roberts SA, et al. Epidemiology of posttransplantation lymphoproliferative disorder in adult renal transplant recipients. Transplantation. 2013;95(3):470-478.

35. Smith JM, Corey L, Bittner R, et al. Subclinical viremia increases risk for chronic allograft injury in pediatric renal transplantation. J Am Soc Nephrol. 2010;21(9):1579-1586.

36. Zuo L, Yue W, Du S, et al. An update: Epstein-barr virus and immune evasion via microRNA regulation. Virol Sin. 2017;32(3):175-187.

37. Gallo A, Vella S, Miele M, et al. Epstein barr viral microrna’s profiling of prognostic and diagnostic value in a PTLD patient cohort. . 2017(20th ESCV 2017):27.

38. Zijlstra JM, Lindauer-van der Werf G, Hoekstra OS, Hooft L, Riphagen II, Huijgens PC. 18F-fluoro-deoxyglucose positron emission tomography for post-treatment evaluation of malignant lymphoma: A systematic review. Haematologica. 2006;91(4):522-529.

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21

39. Mucha K, Foroncewicz B, Ziarkiewicz-Wroblewska B, Krawczyk M, Lerut J, Paczek L. Post-transplant lymphoproliferative disorder in view of the new WHO classification: A more rational approach to a protean disease? Nephrol Dial Transplant. 2010;25(7):2089-2098.

40. Jimenez-Rivera C, Avitzur Y, Fecteau AH, Jones N, Grant D, Ng VL. Sirolimus for pediatric liver transplant recipients with post-transplant lymphoproliferative disease and hepatoblastoma. Pediatr Transplant. 2004;8(3):243-248.

41. Majewski M, Korecka M, Kossev P, et al. The immunosuppressive macrolide RAD inhibits growth of human epstein-barr virus-transformed B lymphocytes in vitro and in vivo: A potential approach to prevention and treatment of posttransplant lymphoproliferative disorders. Proc Natl Acad Sci U S A. 2000;97(8):4285-4290.

42. Majewski M, Korecka M, Joergensen J, et al. Immunosuppressive TOR kinase inhibitor everolimus (RAD) suppresses growth of cells derived from posttransplant lymphoproliferative disorder at allograft-protecting doses. Transplantation. 2003;75(10):1710-1717.

43. Morales J, Fierro A, Benavente D, et al. Conversion from a calcineurin inhibitor-based immunosuppressive regimen to everolimus in renal transplant recipients: Effect on renal function and proteinuria. Transplant Proc. 2007;39(3):591-593.

44. Foran JM, Norton AJ, Micallef IN, et al. Loss of CD20 expression following treatment with rituximab (chimaeric monoclonal anti-CD20): A retrospective cohort analysis. Br J Haematol. 2001;114(4):881-883.

45. Zimmermann H, Trappe RU. EBV and posttransplantation lymphoproliferative disease: What to do? Hematology Am Soc Hematol Educ Program. 2013;2013:95-102.

46. Barkholt L, Linde A, Falk KI. OKT3 and ganciclovir treatments are possibly related to the presence of epstein-barr virus in serum after liver transplantation. Transpl Int. 2005;18(7):835-843.

47. Funch DP, Walker AM, Schneider G, Ziyadeh NJ, Pescovitz MD. Ganciclovir and acyclovir reduce the risk of post-transplant lymphoproliferative disorder in renal transplant recipients. Am J Transplant. 2005;5(12):2894-2900.

48. Savoldo B, Goss JA, Hammer MM, et al. Treatment of solid organ transplant recipients with autologous epstein barr virus-specific cytotoxic T lymphocytes (CTLs). Blood. 2006;108(9):2942-2949. 49. Comoli P, Labirio M, Basso S, et al. Infusion of autologous epstein-barr virus (EBV)-specific cytotoxic T

cells for prevention of EBV-related lymphoproliferative disorder in solid organ transplant recipients with evidence of active virus replication. Blood. 2002;99(7):2592-2598.

50. Burns DM, Crawford DH. Epstein-barr virus-specific cytotoxic T-lymphocytes for adoptive immunotherapy of post-transplant lymphoproliferative disease. Blood Rev. 2004;18(3):193-209. 51. Haque T, Wilkie GM, Jones MM, et al. Allogeneic cytotoxic T-cell therapy for EBV-positive

posttransplantation lymphoproliferative disease: Results of a phase 2 multicenter clinical trial. Blood. 2007;110(4):1123-1131.

52. Moutschen M, Leonard P, Sokal EM, et al. Phase I/II studies to evaluate safety and immunogenicity of a recombinant gp350 epstein-barr virus vaccine in healthy adults. Vaccine. 2007;25(24):4697-4705. 53. Elliott SL, Suhrbier A, Miles JJ, et al. Phase I trial of a CD8+ T-cell peptide epitope-based vaccine for

infectious mononucleosis. J Virol. 2008;82(3):1448-1457.

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22

54. Sokal EM, Hoppenbrouwers K, Vandermeulen C, et al. Recombinant gp350 vaccine for infectious mononucleosis: A phase 2, randomized, double-blind, placebo-controlled trial to evaluate the safety, immunogenicity, and efficacy of an epstein-barr virus vaccine in healthy young adults. J Infect Dis. 2007;196(12):1749-1753.

55. Cohen JI, Mocarski ES, Raab-Traub N, Corey L, Nabel GJ. The need and challenges for development of an epstein-barr virus vaccine. Vaccine. 2013;31 Suppl 2:B194-6.

56. Cohen JI. Epstein-barr virus vaccines. Clin Transl Immunology. 2015;4(1):e32.

57. Boeke CE, Pauly ME, Hatch-Stock H, Jackson JB. CMV antibody prevalence and seroincidence in plateletpheresis donors. J Clin Apher. 2008;23(2):63-65.

58. Munro SC, Hall B, Whybin LR, et al. Diagnosis of and screening for cytomegalovirus infection in pregnant women. J Clin Microbiol. 2005;43(9):4713-4718.

59. Griffiths P, Baraniak I, Reeves M. The pathogenesis of human cytomegalovirus. J Pathol. 2015;235(2):288-297.

60. De Keyzer K, Van Laecke S, Peeters P, Vanholder R. Human cytomegalovirus and kidney transplantation: A clinician’s update. Am J Kidney Dis. 2011;58(1):118-126.

61. Fishman JA. Infection in solid-organ transplant recipients. N Engl J Med. 2007;357(25):2601-2614. 62. Rubin RH. The pathogenesis and clinical management of cytomegalovirus infection in the organ

transplant recipient: The end of the ‘silo hypothesis’. Curr Opin Infect Dis. 2007;20(4):399-407. 63. Steininger C. Clinical relevance of cytomegalovirus infection in patients with disorders of the immune

system. Clin Microbiol Infect. 2007;13(10):953-963.

64. Husain S, Pietrangeli CE, Zeevi A. Delayed onset CMV disease in solid organ transplant recipients. Transpl Immunol. 2009;21(1):1-9.

65. Boeckh M, Riddell SR. Immunologic predictors of late cytomegalovirus disease after solid organ transplantation--an elusive goal? J Infect Dis. 2007;195(5):615-617.

66. Razonable RR, Humar A, AST Infectious Diseases Community of Practice. Cytomegalovirus in solid organ transplantation. Am J Transplant. 2013;13 Suppl 4:93-106.

67. Allice T, Enrietto M, Pittaluga F, et al. Quantitation of cytomegalovirus DNA by real-time polymerase chain reaction in peripheral blood specimens of patients with solid organ transplants: Comparison with end-point PCR and pp65 antigen test. J Med Virol. 2006;78(7):915-922.

68. Espy MJ, Uhl JR, Sloan LM, et al. Real-time PCR in clinical microbiology: Applications for routine laboratory testing. Clin Microbiol Rev. 2006;19(1):165-256.

69. Humar A, Mazzulli T, Moussa G, et al. Clinical utility of cytomegalovirus (CMV) serology testing in high-risk CMV D+/R- transplant recipients. Am J Transplant. 2005;5(5):1065-1070.

70. Kwon JS, Kim T, Kim SM, et al. Comparison of the commercial QuantiFERON-CMV and overlapping peptide-based ELISPOT assays for predicting CMV infection in kidney transplant recipients. Immune Netw. 2017;17(5):317-325.

71. Abate D, Saldan A, Mengoli C, et al. Comparison of cytomegalovirus (CMV) enzyme-linked immunosorbent spot and CMV quantiferon gamma interferon-releasing assays in assessing risk of CMV infection in kidney transplant recipients. J Clin Microbiol. 2013;51(8):2501-2507.

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23

72. Kasprzak A, Zabel M, Wysocki J, et al. Detection of DNA, mRNA and early antigen of the human cytomegalovirus using the immunomax technique in autopsy material of children with intrauterine infection. Virchows Arch. 2000;437(5):482-490.

73. Mattes FM, McLaughlin JE, Emery VC, Clark DA, Griffiths PD. Histopathological detection of owl’s eye inclusions is still specific for cytomegalovirus in the era of human herpesviruses 6 and 7. J Clin Pathol. 2000;53(8):612-614.

74. Razonable RR, Emery VC, 11th Annual Meeting of the IHMF (International Herpes Management Forum). Management of CMV infection and disease in transplant patients. 27-29 february 2004. Herpes. 2004;11(3):77-86.

75. Limaye AP, Corey L, Koelle DM, Davis CL, Boeckh M. Emergence of ganciclovir-resistant cytomegalovirus disease among recipients of solid-organ transplants. Lancet. 2000;356(9230):645-649.

76. Myhre HA, Haug Dorenberg D, Kristiansen KI, et al. Incidence and outcomes of ganciclovir-resistant cytomegalovirus infections in 1244 kidney transplant recipients. Transplantation. 2011;92(2):217-223. 77. Kotton CN, Kumar D, Caliendo AM, et al. Updated international consensus guidelines on the

management of cytomegalovirus in solid-organ transplantation. Transplantation. 2013;96(4):333-360. 78. Biron KK. Antiviral drugs for cytomegalovirus diseases. Antiviral Res. 2006;71(2-3):154-163. 79. Drew WL. Cytomegalovirus resistance testing: Pitfalls and problems for the clinician. Clin Infect Dis.

2010;50(5):733-736.

80. Singh N. Antiviral drugs for cytomegalovirus in transplant recipients: Advantages of preemptive therapy. Rev Med Virol. 2006;16(5):281-287.

81. Kotton CN, Kumar D, Caliendo AM, et al. The third international consensus guidelines on the management of cytomegalovirus in solid-organ transplantation. Transplantation. 2018;102(6):900-931.

82. Hodson EM, Jones CA, Webster AC, et al. Antiviral medications to prevent cytomegalovirus disease and early death in recipients of solid-organ transplants: A systematic review of randomised controlled trials. Lancet. 2005;365(9477):2105-2115.

83. Kalil AC, Levitsky J, Lyden E, Stoner J, Freifeld AG. Meta-analysis: The efficacy of strategies to prevent organ disease by cytomegalovirus in solid organ transplant recipients. Ann Intern Med. 2005;143(12):870-880.

84. Santos CA. Cytomegalovirus and other beta-herpesviruses. Semin Nephrol. 2016;36(5):351-361. 85. Doyle AM, Warburton KM, Goral S, Blumberg E, Grossman RA, Bloom RD. 24-week oral ganciclovir

prophylaxis in kidney recipients is associated with reduced symptomatic cytomegalovirus disease compared to a 12-week course. Transplantation. 2006;81(8):1106-1111.

86. Humar A, Limaye AP, Blumberg EA, et al. Extended valganciclovir prophylaxis in D+/R- kidney transplant recipients is associated with long-term reduction in cytomegalovirus disease: Two-year results of the IMPACT study. Transplantation. 2010;90(12):1427-1431.

87. Gardner SD, Field AM, Coleman DV, Hulme B. New human papovavirus (B.K.) isolated from urine after renal transplantation. Lancet. 1971;1(7712):1253-1257.

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88. Knowles WA. Discovery and epidemiology of the human polyomaviruses BK virus (BKV) and JC virus (JCV). Adv Exp Med Biol. 2006;577:19-45.

89. Stolt A, Sasnauskas K, Koskela P, Lehtinen M, Dillner J. Seroepidemiology of the human polyomaviruses. J Gen Virol. 2003;84(Pt 6):1499-1504.

90. Krumbholz A, Zell R, Egerer R, et al. Prevalence of BK virus subtype I in germany. J Med Virol. 2006;78(12):1588-1598.

91. Takasaka T, Goya N, Tokumoto T, et al. Subtypes of BK virus prevalent in japan and variation in their transcriptional control region. J Gen Virol. 2004;85(Pt 10):2821-2827.

92. Zhong S, Yogo Y, Ogawa Y, et al. Even distribution of BK polyomavirus subtypes and subgroups in the japanese archipelago. Arch Virol. 2007;152(9):1613-1621.

93. Hirsch HH, Vincenti F, Friman S, et al. Polyomavirus BK replication in de novo kidney transplant patients receiving tacrolimus or cyclosporine: A prospective, randomized, multicenter study. Am J Transplant. 2013;13(1):136-145.

94. Nickeleit V, Mihatsch MJ. Polyomavirus nephropathy in native kidneys and renal allografts: An update on an escalating threat. Transpl Int. 2006;19(12):960-973.

95. Egli A, Binggeli S, Bodaghi S, et al. Cytomegalovirus and polyomavirus BK posttransplant. Nephrol Dial Transplant. 2007;22 Suppl 8:viii72-viii82.

96. Hirsch HH, Steiger J. Polyomavirus BK. Lancet Infect Dis. 2003;3(10):611-623.

97. Hirsch HH, Knowles W, Dickenmann M, et al. Prospective study of polyomavirus type BK replication and nephropathy in renal-transplant recipients. N Engl J Med. 2002;347(7):488-496.

98. Dharnidharka VR, Cherikh WS, Abbott KC. An OPTN analysis of national registry data on treatment of BK virus allograft nephropathy in the united states. Transplantation. 2009;87(7):1019-1026. 99. Schold JD, Rehman S, Kayle LK, Magliocca J, Srinivas TR, Meier-Kriesche HU. Treatment for BK virus:

Incidence, risk factors and outcomes for kidney transplant recipients in the united states. Transpl Int. 2009;22(6):626-634.

100. Boothpur R, Brennan DC. Human polyoma viruses and disease with emphasis on clinical BK and JC. J Clin Virol. 2010;47(4):306-312.

101. Wunderink HF, van der Meijden E, van der Blij-de Brouwer CS, et al. Pretransplantation donor-recipient pair seroreactivity against BK polyomavirus predicts viremia and nephropathy after kidney transplantation. Am J Transplant. 2017;17(1):161-172.

102. Wunderink HF, van der Meijden E, van der Blij-de Brouwer CS, et al. Stability of BK polyomavirus IgG seroreactivity and its correlation with preceding viremia. J Clin Virol. 2017;90:46-51.

103. Martelli F, Giannecchini S. Polyomavirus microRNAs circulating in biological fluids during viral persistence. Rev Med Virol. 2017.

104. Li JY, McNicholas K, Yong TY, et al. BK virus encoded microRNAs are present in blood of renal transplant recipients with BK viral nephropathy. Am J Transplant. 2014;14(5):1183-1190.

105. Pietila T, Nummi M, Auvinen P, Mannonen L, Auvinen E. Expression of BKV and JCV encoded microRNA in human cerebrospinal fluid, plasma and urine. J Clin Virol. 2015;65:1-5.

106. Hirsch HH, Babel N, Comoli P, et al. European perspective on human polyomavirus infection, replication and disease in solid organ transplantation. Clin Microbiol Infect. 2014.

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107. Brennan DC, Aguado JM, Potena L, et al. Effect of maintenance immunosuppressive drugs on virus pathobiology: Evidence and potential mechanisms. Rev Med Virol. 2013;23(2):97-125.

108. Hirsch HH, Randhawa P, AST Infectious Diseases Community of Practice. BK polyomavirus in solid organ transplantation. Am J Transplant. 2013;13 Suppl 4:179-188.

109. Bernhoff E, Gutteberg TJ, Sandvik K, Hirsch HH, Rinaldo CH. Cidofovir inhibits polyomavirus BK replication in human renal tubular cells downstream of viral early gene expression. Am J Transplant. 2008;8(7):1413-1422.

110. Faguer S, Hirsch HH, Kamar N, et al. Leflunomide treatment for polyomavirus BK-associated nephropathy after kidney transplantation. Transpl Int. 2007;20(11):962-969.

111. Wu JK, Harris MT. Use of leflunomide in the treatment of polyomavirus BK-associated nephropathy. Ann Pharmacother. 2008;42(11):1679-1685.

112. Grellier J, Hirsch HH, Mengelle C, et al. Impact of donor BK polyomavirus replication on recipient infections in living donor transplantation. Transpl Infect Dis. 2018:e12917.

113. Pascual J, Berger SP, Witzke O, et al. Everolimus with reduced calcineurin inhibitor exposure in renal transplantation. J Am Soc Nephrol. 2018;29(7):1979-1991.

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CHAPTER 2

A Real Time genotyping PCR assay

for Polyomavirus BK

Lilli Gard,

a

_{Hubert G.M. Niesters}

a

_{, Annelies}

Riezebos-Brilman

a

a_{Department of Medical Microbiology, Division of Clinical}

Virology, The University of Groningen, University Medical Center Groningen, Hanzeplein 1, 9713 GZ Groningen, The Netherlands.

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28 ABSTRACT

Background

Polyomavirus BK (BKV) may cause nephropathy in renal transplant recipients and hemorrhagic cystitis in bone marrow recipients. We developed Real-Time PCRs (RT-PCR) to determine easily and rapidly the different BKV genotypes (BKGT) (I-IV).

Methods

On the VP1 gene a duplex of RT-PCRs was developed and validated to differentiate the four main BKGT. 212 BKV positive samples (21 plasma, 191 urine) were tested with these specific PCRs. Of these 212 samples, 55 PCR results were additionally confirmed by sequencing a VP1 gene fragment (nucleotide 1630-1956).

Results

For every genotype, a highly specific, precise and internally controlled assay was developed with a limit of detection of log 3 copies per ml. In 18 (8.5%) of these samples genotyping was not successful due to a low viral load. By sequence analysis, the genotype of 46 out of 55 and 2 out of 4 samples with double infection could be confirmed.

Conclusions

This study describes RT-PCRs for detection of the main BKGT. It proved to be rapid, cheap and sensitive compared to sequencing. Double infections can also be detected. This method will be of value to investigate the role of BKV infection in relation to the genotype.

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29 INTRODUCTION

In the early 1970s, Polyomavirus BK (BKV) was first isolated from the urine of a renal transplant patient by Gardner et al. 1_{It is a non-enveloped double stranded DNA virus belonging to the}

Polyomavirus family. BKV is ubiquitous and widely spread in the adult human population, with sero-prevalence rates up to 80%. Infection results in a lifetime persistence of the virus in which BKV establishes latency in the reno-urinary tract.1-3_{BKV can be classified into four}

genotypes (I-IV) using serological or genotyping methods.2,4_{Several studies investigated the}

geographical distribution of BKV. Genotype I is the most prevalent throughout the world, followed by genotype IV, which is mainly seen in Europe and Asia. Genotypes II and III occur rarely, while genotype III is mostly seen in Africa 5-7_.

Through the diversity of humans origin within the different continents, it is most likely that genotypes II, III and IV also occur in humans of different origin.So far, more than twelve other human polyomaviruses have been described. As far as known, only JC virus (JCV), Merkel cell virus and Trichodysplasia spinulosa virus can be associated with a human clinical disease. 8

Although the genome of BKV is closest related to JCV, both do cause different clinical features in immunocompromised patients. In bone marrow transplant recipients, BKV is associated with hemorrhagic cystitis whereas in renal transplantation patients it may cause nephropathy (BKVAN) with subsequent allograft loss. 9

Predictive markers for developing BKVAN are high titers in urine and plasma 10_{. At present}

reduction, changing or discontinuation of the immunosuppressive regimen is recommended, in patients with BKV viremia, 11,12_{through the lack of antiviral treatment for BKV diseases}13_.

Relating to the development of BKVAN, much is still unknown, like the association among the various BKV genotypes (BKGT) with their causative role in the development of clinical syndromes. Krautkramer et al. showed that high BKV viral load is independent of the mutations in BKV VP1 genes. 14_{On the other hand, Boldorino et al found a correlation between BKVAN}

and the frequency of single base-pair mutations in BKV DNA.15

The availability of a rapid, simple and cheap genotyping Real-Time PCRs (RT-PCRs) will be useful for studies to investigate the impact of the role of BKV infection in relation with BK genotype. Hence the aim of this study was to develop easy to use RT-PCRs.

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30 MATERIALS & METHODS

Clinical Specimens and plasmid

For this study, clinical specimens were tested for BKV DNA in the period January 2008 until September 2013 using an in-house developed quantitative BKV RT-PCR on the VP2 gene. From this period, 212 BKV positive (21 plasma and 191 urine) samples from 191 different patients (171 kidney Tx, 2 liver Tx, 16 hematology, 1 pancreas Tx and 1 other) were randomly selected with a DNA load ranging between log 2 copies per ml (cp/ml) and log 11cp/ml. Subsequently, 50 of these urine and plasma samples were randomly selected and sequenced for the determination of the BKGT.

For the validation of the assay urine and plasma from three patients with either genotype I, II or IV were used to prepare 10-fold dilution series. Since there was no clinical sample for genotype III a plasmid, using pDONR as vector, was constructed (Gene Art Life TechnologyRegensburg, Germany) using the VP1 gene sequence from Genbank accession number M23122.

BKV RT-PCR

Isolation of the BKV DNA was performed using the MP96 isolation station (Roche Diagnostics, Germany) according to the manufactures instructions. The DNA viral NA Blood SV 2.0 protocol was used with input of 200µl plasma or urine and elution in 100µl.

An internally controlled in-house developed quantitative BKV RT-PCR was used for the detection of BKV DNA. Currently, no international standard for BKV is available, therefore the validation and quantification of this BKV RT-PCR was done using the EZ Validation software (Life Technology, USA) using the AcroMetrix quality control materials. Furthermore, patient samples ranging from < log 2 to log 11 copies/ml were compared with the RealStar BKV PCR Kit (Altona Diagnostics, Germany) and the BKV Q-PCR Alert-kit ( ElitechGroup, France) to confirm that samples with a load of < log 2 with the in-house BKV RT-PCR are not false-negative (data not published). In short, our BKV RT-PCR is an internally controlled duplex based on the VP2 gene, with a detection limit of log 2 cp/ml. The reaction mixture was the same as described below for the specific BKGT RT-PCR. It consist of 300nM of each primer (Forward:5’- ATGGGTGCTGCTCTAGCACTTT-3’, Reverse 1:5’-GGATGCAATTTGAACTTCTATGGC-3’, Reverse 2: 5’-GGATGCAATTTGAACTTCTATAGCAG-3’) and 100nM of the probe (5’-FAM-CAGTGTATCTGAGGCTG-MGB-3’), which amplifies a 131bp of the VP2 gene from nucleotide position 624 to 755 using Genbank accession number NC_001538. As internal control, Phocine herpesvirus (PhHV) with minor modifications in the probe sequence (5’-Cy5-TTTTTATGTGTCCGCCACCATCTGGATC-BBQ-3’) was used as described by van Doornum et al. 16

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31

BKGT specific RT- PCR Primers and probe design

In total, 304 complete BKV genomic sequences that were available, up to February 2015, present in the Genbank database were aligned using Genedoc software to identify highly conserved regions in each BKGT. Primers and probes were designed on the VP1 gene, in a region with low variability between the subtypes of a genotype, but with high variability between the genotypes. The design was performed with unique sequences for the BKV subtypes I and IV and genotypes II and III using Primer Express v3.0 (Applied Biosystem, USA) and with consideration of the primer and probe design guidelines. Confirmation of the specificity as well as their homology was checked using BLAST software on the NCBI site. MGB (minor groove binding) probes were used for the discrimination between the 4 BKGT.

Validation

The four BKGT RT-PCRs are internally controlled and optimized for the primer/probe concentrations. PhHV, (see paragraph 2.3.1) , is used as an internal control. In all experiments. PCR reactions were performed in a 50µl reaction containing 20µl of isolated DNA, 2x Taqman Universal Mastermix (Life Technologies, USA), 5mg/ml Bovine serum albumin (Roche, Germany), 300nM of each PhHV primer, 100nM PhHV probe and primers and probes for the specific BKGT (Life Technologies, USA). Table 1 shows the concentration of the specific BKGT primers and probes.

The 4 internally controlled BKGT RT-duplex PCRs were performed on the ABI PRISM 7500 (Life Technologies, USA), with the following thermal conditions: 50°C for 2min, 95°C for 10min followed by 42 cycles of 95°C for 15sec, 60°C for 1 min.

Precision, Sensitivity and Specificity

The precision is validated by the intra/ inter assay variation, which was assessed by testing 10-fold serial dilutions in triplicate of each genotype.

Sensitivity was determined by using 10-fold serial dilutions of BKGT I to IV. All dilutions were tested in 10fold for the in-house BKV RT-PCR and the specific BKGT RT-PCRs.

For the specificity, 10-fold serial dilutions (ranging from log 2 to log 8 cp/ml) of each genotype were tested in every specific BKGT RT-PCR and the following viruses were tested with a dilution of log 4-5 cp/ml: B19, CMV, EBV, HHV6, HHV8, HSV 1, HSV 2, JCV and VZV. All viruses origins from patient samples Furthermore, the accuracy was determined by comparing the results of 55 positive BKV samples in the specific BKGT RT-PCRs with the sequence analysis of the 1630-1956 VP1 nucleotide fragment as described by Jin et al. 4

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32

Sequence analysis

In total, 55 samples of the 212 samples were also sequenced, in order to confirm the RT-PCR genotype resultsThese samples were randomly selected with a mix of urine and plasma samples and varying in load from log 2 cp/ml to log 11 cp/ml.

The PCR mixture consist of a total volume of 50µl, containing 5µl 10x PCR Buffer (Qiagen), 25mM MgCl₂(Qiagen), 50x dNTP plus (Roche, Germany) 500nM of primer 327-1 and 327-2 4_,

2.5 units Hotstar Taq Polymerase (Qiagen) and 10µl template DNA. The cycling conditions were as followed: 95°C for 15 min, followed by 50 cycles of 95°C for 30 sec, 55°C for 30 sec and 72°C for 1 min, with an final extension of 72°C for 10 min and cooling to 4°C. According to the manufacturer instructions the PCR products were sequenced using the the BigDye Terminator v3.1 Cycle Sequencing Kit (Life Technologies, USA) and the product was purified with the BigDye X Terminator Purification Kit (Life Technologies, USA). Sequence detection was performed on the ABI PRISM 3130 XL Genetic Analyzer (Life Technologies, USA). After alignment using the DNASTAR Lasergene 10 Core Suite software (DNASTAR, USA,) BLAST on the NCBI site was used to compare the BKV sequences.

RESULTS

Primer and probe design

Primers and probes were designed in the most conserved region for each genotype on the VP1 gene. This region was almost the same for genotypes II, III and IV, whereas for genotype I another region was used. Degenerated reverse primers were used for all assays and a degenerated probe for genotype III. Due to the melting temperature of the genotype I probe, it was not possible to use a degenerate probe, therefore the genotype I RT-PCR consist of 2 probes. Table 1 shows the sequences and locations of the primers and probes.