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A Prospective Longitudinal Study of Polyomavirus Shedding in Lung-Transplant Recipients

  1. Lora D. Thomas1,
  2. Regis A. Vilchez4,5,a,
  3. Zoe S. White5,
  4. Preeti Zanwar5,
  5. Aaron P. Milstone1,3,
  6. Janet S. Butel5 and
  7. Stephen Dummer1,2,3
  1. 1Department of Medicine, The Transplant Center, Vanderbilt University School of Medicine, Nashville, Tennessee
  2. 2Department of Surgery, The Transplant Center, Vanderbilt University School of Medicine, Nashville, Tennessee
  3. 3The Transplant Center, Vanderbilt University School of Medicine, Nashville, Tennessee
  4. 4Department of Medicine, Baylor College of Medicine, Houston, Texas
  5. 5Department of Molecular Virology and Microbiology, Baylor College of Medicine, Houston, Texas
  1. Reprints or correspondence: Dr. Stephen Dummer, 911 Oxford House, Vanderbilt University Medical Center, Nashville, TN 37232 (Stephen.Dummer{at}vanderbilt.edu).

  1. Presented in part: Infectious Diseases Society of America meeting, Boston, MA, 30 September–3 October 2004 (abstract 633).

  • ↵a Present affiliation: Department of Virology, Boehringer-Ingelheim Pharma-ceuticals, Inc., Ridgefield, Connecticut.

 
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Abstract

Background. Polyomavirus infection causes renal dysfunction after kidney transplantation, but it has not been thoroughly investigated in nonrenal solid-organ transplantation.

Methods. Fifty lung-transplant recipients provided prospective urine and blood samples over the course of 17 months. Samples were analyzed for BK virus (BKV), JC virus (JCV), and simian virus 40 (SV40) using conventional polymerase chain reaction (PCR), sequence analysis, and quantitative real-time PCR.

Results. Thirty-one (62%) of 50 patients had polyomavirus detected in at least 1 urine specimen, including 16 (32%) for BKV, 12 (24%) for JCV, and 6 (12%) for SV40. Mean BKV loads (5.0 log10 copies/mL) did not differ from those of JCV (5.7 log10 copies/mL; P=.38), but SV40 loads (2.5 log10 copies/mL) were lower than those of BKV (P=.006) and JCV (P=.002). Blood samples were negative. Infection with individual polyomaviruses or polyomavirus infection in aggregate was not associated with reduced creatinine clearance. Patients not shedding polyomavirus had better survival than patients shedding polyomavirus (P=.049).

Conclusions. Polyomaviruses BKV and JCV were commonly detected in urine from lung-transplant recipients. SV40 was found in 12% of patients but was shed at a lower frequency and with lower viral loads than the other viruses. Polyomavirus infection was not associated with renal dysfunction.

Organ-transplant recipients are at risk of polyomavirus infection and polyomavirus-induced renal dysfunction [1–6]. Nephropathy associated with polyomaviruses has been reported in 1%–10% of renal-transplant recipients, and it frequently leads to loss of the allograft [4]. BK virus (BKV) is the agent commonly implicated in renal transplantation, although renal dysfunction related to JC virus (JCV) or simian virus 40 (SV40) has also been described [7–9]. Polyomavirus nephropathy appears to be less common in recipients of other solid-organ transplants, although well-documented cases have been described [9–14].

SV40 was initially identified as a contaminant of vaccines used between 1955 and 1962, which is likely how it was introduced into human populations [15]. We recently described a lung-transplant recipient who developed end-stage renal failure related to SV40 infection [9]. This case and other reports suggested that SV40 circulates in the community and can cause infections in immunocompromised transplant recipients, but the prevalence and significance of infections in humans is unknown [16–20].

Only a few prospective studies have addressed the clinical significance of polyomavirus infection in non-renal solid-organ-transplant recipients [21–27]. Several studies failed to show an association between the detection of BKV or JCV and renal dysfunction [21–24], whereas one found an independent association between BKV reactivation and impaired renal function in a cohort of kidney, heart, and liver recipients [25]. Other studies that examined BKV in blood samples collected from kidney-, heart-, and liver-transplant recipients found that detection of polyomaviruses was more common in kidney recipients than in other transplant recipients [26, 27].

A recent editorial considered the need to determine whether BKV-related renal dysfunction is a problem in recipients of nonrenal solid-organ transplants [28]. The present article describes the results of the initial phase of a prospective longitudinal study of polyomavirus infection in 50 lung-transplant recipients over the course of 17 months of follow-up.

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Patients, Materials, and Methods

Study population. Adult lung- and heart-lung-transplant recipients receiving medical care at the Vanderbilt University Hospital were enrolled from April 2002 through October 2002 and were monitored until the end of August 2003. Patients were eligible if they had received a lung or a heart-lung transplant and were ambulatory. Patients were excluded if they had received a renal transplant or were <18 years old. Two patients undergoing dialysis were excluded because of the difficulty of getting urine specimens from them. All patients signed informed consent forms approved by the institutional review boards of each institution for the study.

Demographic data collected on patients included age, sex, underlying diagnosis, time since transplant, and cytomegalovirus (CMV) serological status. Donor information included age, sex, and CMV serological status. At each visit, the serum creatinine level and current immunosuppressive regimen were recorded. During the study period, we documented all episodes of acute rejection and the prevalence and time of onset of chronic graft dysfunction due to bronchiolitis obliterans (BO) or BO syndrome (BOS). Creatinine clearances were calculated with a standard Cockcroft-Gault formula using an ideal-bodyweight formula [29].

Sample collection, sample processing, and DNA extraction. Urine and blood samples were collected from patients at ∼3–4-month intervals when they came for routine clinic visits. Heparinized blood samples were placed upright for 2 h; plasma and ∼400 μL of cells at the interface were then pipetted off. This leukocyte-rich plasma was centrifuged at 600 g for 6 min, the plasma was removed, and the cell pellet of peripheral blood leukocytes (PBLs) was resuspended in 1 mL of PBS. Both cells and plasma were frozen at −70°C. Urine samples were obtained in sterile collection cups and spun at low speed (2000 g) for 10 min at room temperature to pellet cellular material, and the supernatants were separated from the pellets and stored at −70°C. The pellets were washed once with PBS, repelleted, and stored at −70°C. Frozen samples were shipped to Baylor College of Medicine (Houston, TX) for blinded laboratory analysis.

Sample processing was performed in a laminar flow hood in a biosafety level 3 facility free of polyomaviruses and plasmids in the Department of Molecular Virology and Microbiology at Baylor College of Medicine. These containment facilities ensured that human specimens were not at risk of being contaminated in the laboratory with viral plasmids during processing. As a further precaution, positive displacement pipetters and barrier-tip pipettes were used. PBLs and urine samples were processed using a standard proteinase K-phenol extraction protocol. PBLs and urine pellets were digested with proteinase K (Fisher) in a total volume of 1 mL at 55°C for 4 h. The digests were extracted with 500 μL of Tris-buffered phenol (pH 8), and the DNA was precipitated with sodium acetate/isopro-panol and washed once with 70% ethanol. The DNA pellets were then resuspended in Tris-EDTA buffer (pH 8.0) and stored at −20°C [30].

Conventional polymerase chain reaction (PCR), real-time quantitative (RQ)-PCR, and sequence analysis. The oligonucleotide primers used for conventional PCR and DNA se-quence analysis of polyomaviruses have been described elsewhere [9, 30, 31]. Each PCR contained 0.5–1 μg of PBL DNA (∼8×104–1.6×105 cell equivalents) or DNA extracted from a urine pellet (equivalent to 1.5–2.0 mL of urine) in a 100-μL reaction volume. PCR products were analyzed on a 2% ethidium-bromide-stained agarose gel. The limits of detection of the conventional PCR assays for polyomaviruses were ∼100–1000 genome copies/reaction [32].

Plasmids pBK-Dunlop, pJC-MAD-1, and pSVSph21-N served as positive control templates. Positive control plasmids were added to the control PCRs outside of the core facility after tubes that contained negative controls and test DNAs were closed. PBL DNA samples were tested for suitability for amplification using primers specific for the human β-hemoglobin gene (primers KM38/PC03) [33]. Only specimens from which cellular β-globin gene sequences were amplified were further examined. Urine specimens were not tested for cellular sequences, because those control assays did not predict whether urine samples contained viral sequences [34]. PCR amplifications were performed in a Perkin Elmer GeneAmp PCR System 2400 thermocycler or in an MJ Research PTC-200 Peltier Thermal Cycler for a total of 45 denaturation, annealing, and extension steps, using temperatures specific for each primer pair.

BKV, JCV, and SV40 loads were determined by RQ-PCR with fluorogenic probes for each virus using the ABI PRISM 7000 Sequence Detection System (Applied Biosystems). Specific primers and probes directed against the N termini of the large T-antigen (T-ag) genes of SV40, BKV, and JCV and reaction conditions have been described elsewhere [32]. RQ-PCRs were performed using the TaqMan Universal PCR Master Mix (Applied Biosystems). The limit of detection of each assay was 10 viral genome copies/reaction. Only samples that tested positive by conventional PCR were assayed for viral loads by RQ-PCR. Because of limited amounts of sample, quantitative assays were performed only for the specific polyomavirus(es) previously identified in a given specimen. DNA sequence analysis was performed on conventional PCR products as described elsewhere [35].

Definitions. Samples were first screened for polyomaviruses by conventional PCR using the universal primer set PyVfor/PyVrev [9, 36]. Specimens that yielded an amplicon were then tested with virus-specific primers for the regulatory region, T-ag, or VP1 sequences. Samples were considered to be positive for BKV on the basis of sequence analysis of regulatory region PCR products and for SV40 on the basis of sequence analysis of either regulatory region or T-ag products (in one case, a positive Southern-blot reaction was used in place of sequence confirmation). Samples were considered to be positive for JCV if they tested positive with 2 different JCV-specific primer sets (regulatory region and VP1). A patient was classified as positive for polyomavirus infection if at least 1 urine specimen tested positive.

Data analysis. Patients who tested positive for polyomavirus in urine were compared with those who tested negative for polyomavirus for the following variables: age, sex, time since transplant, diagnosis, type of immunosuppression (cyclosporine vs. tacrolimus and azathioprine vs. mycophenolate), death, the presence of acute rejection, CMV serological status (donor and recipient), and donor age and sex. If a patient received >1 immunosuppressive regimen during the follow-up period, the regimen was documented that the patient was receiving when he or she first started shedding any polyomavirus. For patients who did not shed any virus, the predominant regimen during the period of follow-up was used in data analysis. Because of small numbers of patients, we were able to perform dichotomous analyses only for treatment with cyclosporine versus tacrolimus and azathioprine versus mycophenolate. For each patient, we calculated a mean creatinine clearance using all creatinine clearances measured whenever a urine sample was collected. Each polyomavirus was analyzed independently. For example, patients with BKV infection were compared with those without BKV infection, even if they were infected with a different polyomavirus. Also, patients with any polyomavirus infection were compared with patients without polyomavirus infection.

The patients' mean creatinine clearances during the study period were correlated with their mean viral loads. If RQ-PCR was performed on >1 sample from a patient, the mean log10 value of the viral loads from all samples tested was used in the analysis. We also examined whether there was a relationship between urinary viral load and any of the following demo-graphic variables: age, sex, time since transplantation, and immunosuppressive regimen.

Proportions were analyzed using Fisher's exact test. The Mann-Whitney U test (for 2 groups) or the Kruskal-Wallis test (for >2 groups) was used to compare continuous variables between groups, including mean viral loads and mean creatinine clearances of individual patients. Survival curves (measured from the time of transplantation until death or the end of the follow-up period) were generated using the Kaplan-Meier method. Comparisons of survival curves were analyzed using log-rank statistics. Spearman rank correlation coefficients were used to compare patient's mean viral loads with continuous variables, including patient's creatinine clearances. Stata software (version 9.0; Statacorp) was used to conduct the statistical analyses. For all statistical analyses, P<.05 was considered to be significant.

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Results

Demographic characteristics of the patients are shown in table 1. The population included 28 men and 22 women; the mean age was 49.5±12.7 years. The most common underlying diagnoses were pulmonary fibrosis (28%), emphysema (26%), cystic fibrosis (16%), and pulmonary hypertension (12%). Patients were enrolled a mean of 3.7±2.9 years after transplantation (range, 25 days–11.5 years). A mean of 3.44±1.0 urine samples was collected per patient, over the course of an average follow-up time of 13.5±3.2 months. All but 24 urine samples (of 172 total) had an associated blood sample. Eighty-six percent of patients provided ≥3 samples during the follow-up period.

Figure 1
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Figure 1

Detection of polyomavirus DNA in urine samples from lung-transplant recipients. Examples are shown of polymerase chain reaction-amplified polyomavirus sequences visualized by agarose gel electrophoresis and staining with ethidium bromide. A, T-antigen products obtained using polyomavirus universal primers. B, BK virus (BKV)-specific regulatory region products. C, JC virus (JCV)-specific regulatory region products. D, Simian virus 40 (SV40)-specific regulatory region products. +, Positive control reactions with individual polyomavirus-specific plasmids; −, negative control reactions without added DNA template; M, molecular-weight markers.

Figure 2
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Figure 2

Quantitative viral load in urine, by virus. The mean viral load for each patient was used in generating the box plots. Patients who shed 2 viruses are included in both boxes. The nos. of patients in each box are as follows: BK virus (BKV; n=16), JC virus (JCV; n=12), and simian virus 40 (SV40; n=6) BKV vs. JCV, P=.38; BKV vs. SV40, P=.006; JCV vs. SV40, P=.002 (all Mann-Whitney U test). In each box and whisker plot, the lower and upper edges of the box indicate the 25th and 75th percentiles; the horizontal line within the box represents the median; the whiskers indicate either the minimum and maximum values or a distance of 1.5 times the interquartile range from the edge of the box (whichever distance is smaller).

Figure 3
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Figure 3

Effect of polyomavirus infections on survival of lung-transplant recipients. Solid line, Patients without polyomavirus infection (n=19); dashed line, patients with polyomavirus infection (n=31). Difference in survival, P=.049 (log-rank statistics).

Figure 4
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Figure 4

Calculated creatinine clearance by virus infection. The mean creatinine clearance for each patient is used in generating the box plots. Patients who shed 2 viruses are included in both boxes. The nos. of patients in each box are as follows: no virus (n=19), BK virus (BKV; n=16), JC virus (JCV; n=12), and simian virus 40 (SV40; n=6). Difference in creatinine clearances overall, P=.21 (Kruskal-Wallis test). BKV infection vs. no BKV infection, P=.52 (Mann-Whitney U test); JCV infection vs. no JCV infection, P=.14; SV40 infection vs. no SV40 infection, P=.20. In each box and whisker plot, the lower and upper edges of the box indicate the 25th and 75th percentiles; the horizontal line within the box represents the median; the whiskers indicate either the minimum and maximum values or a distance of 1.5 times the interquartile range from the edge of the box (whichever distance is smaller).

Table 1
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Table 1

Demographic characteristics of transplant recipients.

Dynamics of polyomavirus shedding. Extracts of urine pellets were tested for the presence of polyomaviruses BKV, JCV, and SV40. Examples of conventional PCRs are shown in figure 1. Overall, 64 (37%) of 172 urine specimens were positive for a polyomavirus, or an average of 2.0±1.1 positive samples for every patient who tested positive at least once. Patients with polyomavirus infection did not have more urine samples collected than those not infected with polyomavirus (3.5±1.0 for polyomavirus shedders vs. 3.3±1.1 for nonshedders; P=.53).

Thirty-one (62%) of 50 patients tested positive for polyomavirus in at least 1 urine specimen. This included 16 (32%) for BKV, 12 (24%) for JCV, and 6 (12%) for SV40. Three patients tested positive in urine for 2 viruses. These combinations included BKV and JCV (n=2) and BKV and SV40 (n=1). The frequency of shedding varied according to the virus detected. Seventy percent of the urine samples were positive for JCV in patients who had JCV detected at least once. The corresponding percentages for BKV and SV40 were 51% and 30%, respectively. The frequency of shedding was lower for SV40 than for BKV and JCV (P=.009). None of the 148 blood samples were positive for polyomavirus.

Quantitative analyses for viral load were done on 30 urine samples from 16 patients for BKV, 28 samples from 12 patients for JCV, and 7 samples from 6 patients for SV40. The viral loads of the individual viruses are demonstrated in a box plot (figure 2). Mean viral loads for BKV (5.0 log10 copies/mL) did not differ from those of JCV (5.7 log10 copies/mL; P=.38), but viral loads of SV40 (2.5 log10 copies/mL) were lower than those of BKV (P=.006) and JCV (P=.002).

Clinical correlations. Analysis of demographic variables (age, sex, time since transplantation, underlying diagnosis, donor age and sex, CMV serological status of donor or recipient, and diagnosis of acute rejection) did not show any significant association between patients who shed any polyomavirus in the urine and those who shed no polyomavirus. There was no significant association of these variables with detection of any of the 3 individual polyomaviruses in urine. Patients who shed 2 polyomaviruses in urine were significantly older than patients who did not shed 2 viruses (mean age, 62.3 vs. 48.7 years; P=.037). There was also no significant association between the immunosuppressive regimen and polyomavirus infection either for individual viruses or all viruses in aggregate.

We examined the year of birth of patients infected with SV40, to evaluate whether they would have been at risk of receiving vaccines contaminated with SV40. All 6 patients with SV40 infection were born before 1959 and, thus, could have received contaminated vaccines.

Six patients died during the follow-up period. Improved survival was seen in patients who did not shed any polyomavirus, compared with those who did shed polyomavirus (P=.049) (figure 3). There was no significant reduction in survival with relation to shedding of the individual viruses (data not shown). Causes of death included chronic graft dysfunction (n=3), lymphoma and respiratory syncytial virus infection (n=1), coronary artery disease (n=1), and cardiopulmonary arrest after back surgery (n=1). Other variables—such as age, sex, underlying disease, immunosuppression, and donor age and sex—were not associated with increased mortality. In a separate analysis, we examined the prevalence of chronic graft dysfunction due to BO or BOS. The prevalence of chronic graft dysfunction was 53% in patients who did not shed any virus and 63%, 42%, and 33% in patients who shed BKV, JCV, and SV40, respectively (P, not significant). The time to the development of chronic graft dysfunction did not differ among these 4 groups.

Patients' mean creatinine clearances, categorized by type of viral infection, are presented in figure 4. Creatinine clearances for the 4 groups were not significantly different overall (P=.21). Patients' creatinine clearances also did not differ when the polyomaviruses were analyzed individually.

There was no correlation between the viral load in the urine and the patient's mean creatinine clearance for any of the poly-omaviruses. There was no significant correlation between urinary viral load and the patient's age or the number of months after transplantation when the patient was enrolled. Urinary viral loads for any of the 3 viruses did not correlate with the patient's immunosuppressive regimen or sex.

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Discussion

The primary goals of the present study were to investigate the prevalence of polyomavirus (BKV, JCV, and SV40) infection and ascertain whether these viral infections are associated with renal dysfunction in lung-transplant recipients. The study is ongoing, and the present article analyzes data collected during the first 17 months in the first 50 patients enrolled in the study. We have shown that urinary shedding of polyomaviruses is common in lung recipients—62% of the patients demonstrated shedding at one time or another. BKV and JCV were detected more frequently and at higher viral loads than SV40. These observations are not likely to be due to laboratory contamination, given that negative controls analyzed in parallel in each assay were negative and none of the 148 patient blood samples were polyomavirus positive.

BKV is the major culprit of polyomavirus-associated nephropathy after kidney transplantation and is detected in up to one-half of those patients; however, only ∼5% of kidney recipients develop symptomatic illness, mostly in the form of tubulointerstitial nephritis [3, 37–42]. Studies have shown that biopsy-proven BKV nephritis led to allograft failure in 36% [1] and 46% [6] of affected patients. Augmented immunosuppression seems to play a role in polyomavirus reactivation: lowering of immunosuppression is associated with a decrease in BKV viral load and a reduction in allograft inflammation [1, 2, 4]. There is a concern that the use of tacrolimus and mycophenolate mofetil as immunosuppressive agents increases the risk of BKV reactivation after renal transplantation [1, 2, 43], but this may simply represent the widespread use of these medications. Our analysis did not find an association between immunosuppressant agents and polyomavirus shedding or between the presence of BKV and worse renal function in lung-transplant recipients.

JCV has been found to be a coinfecting agent in renal-transplant patients with BKV nephropathy [44]. The present results suggest that JCV is shed as commonly as BKV in transplant recipients. Whether JCV can cause nephropathy in kidney-transplant recipients without preexisting BKV-induced renal disease is unknown. JCV urinary shedding is also common among immunocompetent subjects, with JCV excretion being more frequent with advancing age [30, 45]. Our findings in lung-transplant recipients did not reveal any association between JCV infection and age. A similar lack of association between age and JCV shedding has been noted in patients infected with HIV-1 [34, 46].

Previous reports have indicated an association between SV40 infection and renal dysfunction among small numbers of transplant recipients and nontransplant patients [7, 9, 20]. SV40 sequences were detected and identified in the allografts of pediatric renal-transplant recipients [18] and in urine, blood, and kidney-biopsy samples from adult patients with focal segmental glomerulosclerosis [20], a known cause of end-stage renal disease. SV40 was also found in association with BKV in renal-transplant recipients with well-documented nephropathy, but its contribution to renal dysfunction was not characterized [7]. Moreover, a study in Italy reported the detection of SV40 DNA sequences in 2 of 7 children with hemorrhagic cystitis after bone-marrow transplantation [19]. In our study, patients with SV40 infection were all born before 1962, and they might have been exposed to vaccines contaminated by SV40. Other studies, however, have detected the presence of SV40 antibodies and DNA in individuals who were born after the discontinuation of SV40-contaminated vaccines [17–19]. Further studies in immunocompetent and immunosuppressed populations will be required to explore the dynamics and pathogenic mechanisms of SV40 infection.

Renal dysfunction is a common complication after nonrenal solid-organ transplantation [47–49]. Ojo et al. [47] documented that 7%–21% of nonrenal solid-organ-transplant recipients developed chronic renal failure within 5 years of transplantation, with the risk varying according to the organ transplanted. Twelve (24%) of the patients in our study had a creatinine clearance of <30 mL/min at some point during the follow-up period. There are several probable etiologies for reduced renal function after transplantation, including calcineurin inhibitor-related nephrotoxicity, diabetes mellitus, hypertension, and preexisting renal disease before transplantation [47]. An independent effect of polyomavirus infection on renal function in nonrenal solid-organ-transplant recipients has yet to be defined.

An unanticipated finding in our study was that patients who shed polyomavirus in the urine had worse survival than patients without polyomavirus shedding. This survival disadvantage was not explained by other demographic variables. The small number of deaths limited the extent of our analysis and also made it impossible to analyze individual causes of death. It is possible that polyomavirus reactivation is an indicator of increased underlying immunosuppression. In the future, it might be possible to correlate laboratory assays of immune function with polyomavirus infection. For now, the survival disadvantage associated with polyomavirus infection must be considered to be a preliminary finding requiring confirmation in larger studies.

Our study had certain limitations. Although our results show that polyomavirus excretion in the urine is quite common after lung transplantation, no renal biopsies were performed to investigate the presence of polyomavirus-induced nephritis. It is possible that sampling bias was introduced by collecting more specimens from patients who were more symptomatic or immunosuppressed. However, there was no difference in the mean number of specimens collected between subjects who shed polyomavirus in the urine and those who did not. The small number of subjects in our study prohibited multivariate analysis.

Molecular assays that are sensitive and specific are excellent tools for the analysis of polyomavirus infections in human populations. Serological assays are of minimal use for the diagnosis of active infections, because most illnesses related to these pathogens are thought to result from reactivation of latent viral infections, and cell culture for polyomavirus detection is cumbersome and inefficient because of slow viral growth and the requirement of specialized cell lines [37]. The present study illustrates the utility of molecular-based approaches to monitor reactivation of multiple polyomavirus types in clinical specimens in longitudinal studies.

In conclusion, these data show that shedding of all 3 polyomaviruses (BKV, JCV, and SV40) occurs among lung-transplant recipients; the frequency of shedding was similar for BKV and JCV, whereas shedding of SV40 was less common. Our ongoing longitudinal study will help determine the value of monitoring polyomavirus loads in recipients of nonrenal solid-organ transplants.

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Acknowledgments

We thank Stacy Kelley-Blackburn, Jean Barnes, and Haley Hoy, for their help with the study.

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Footnotes

  • Potential conflicts of interest: none reported.

  • Financial support. Leukemia and Lymphoma Society (grant 6147-03 to J.S.B. and R.A.V.); National Cancer Institute (grant CA104818 to J.S.B., R.A.V., and S.D.).

  • Received June 20, 2006.
  • Accepted September 20, 2006.
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  1. J Infect Dis. (2007) 195 (3): 442-449. doi: 10.1086/510625
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