Key PointsQuestion
What is the distribution and quantity of John Cunningham virus (JCV) in the tissues of the immunocompetent host and how does it correlate with JCV antibody levels?
Findings
In this study of multiple tissues obtained from 12 immunocompetent persons, JCV predominated in renal-reproductive tissues but was observed in other organs, including brain, in low copy numbers. Tissue distribution and viral copy numbers, particularly urine JCV copy numbers, appeared to correlate with JCV antibody levels.
Meaning
High JCV antibody levels may be indicative of more widespread JCV distribution and replication.
Importance
Although seroepidemiological studies indicate that greater than 50% of the population has been infected with John Cunningham virus (JCV), the sites of JCV persistence remain incompletely characterized.
Objective
To determine sites of JCV persistence in immunologically healthy individuals.
Design, Setting, and Participants
Tissue specimens from multiple sites including brain, renal, and nonrenal tissues were obtained at autopsy performed in the Department of Pathology at the University of Kentucky from 12 immunologically healthy patients between February 9, 2011, and November 27, 2012. Quantitative polymerase chain reaction was performed on the tissue specimens and urine. Serum JCV antibody status was determined by enzyme-linked immunosorbent assay.
Main Outcomes and Measures
The detection and quantification of JCV from the tissues by quantitative polymerase chain reaction illuminated sites of viral persistence. These results were correlated with JCV antibody levels.
Results
Autopsies were performed on 12 individuals, 10 men and 2 women, ranging in age from 25 to 75 years (mean, 55.3 years). Seven of 12 individuals were JCV antibody seropositive based on absorbance units. Serostatus was associated with amounts of JCV DNA in urine and its tissue distribution. John Cunningham virus DNA was found in 75% of genitourinary tissue samples from donors (18 of 24) with high JCV antibody levels, 13.3% of donors with low levels i(4 of 30), and 0% of seronegative persons (0 of 32). In nongenitourinary tissues, JCV DNA was detected in 45.1% of tissue samples of donors (32 of 71) with high JCV, 2.2% of donors with low JCV serostatus (2 of 93), and 0% of seronegative persons (0 of 43). Genitourinary tissues had higher copy numbers than other sites. John Cunningham virus DNA was detected in urine of seronegative individuals in a research-grade assay.
Conclusions and Relevance
Persistent (latent or actively replicating) JCV infection mostly predominates in genitourinary tissues but distributes in other tissues at low copy number. The distribution and copy numbers of the virus appear to correlate with urinary JCV shedding and serostatus.
Seroepidemiological data indicate that John Cunningham virus (JCV), the cause of progressive multifocal leukoencephalopathy (PML), is ubiquitous. Besides biological factors, differences in the frequency with which seropositivity has been demonstrated may in part be dependent on the assay used. By age 10 years, 40% to 60% of the population is JCV antibody positive.1-3 By adulthood, seroprevalence rates of 80% or higher have been reported.3,4 The mode of transmission of this virus is unknown. In the past 3 decades, the incidence of PML has risen sharply owing to the human immunodeficiency virus/AIDS pandemic, and since 2005, an increased incidence has been demonstrated with use of the immunomodulatory agent natalizumab, an α4β1 integrin inhibitor, and with other immunomodulatory regimens.
Endogenous reservoirs of JCV that contribute to PML are unknown. Polymerase chain reaction (PCR) or viral isolation techniques detect JCV in the urine of 10% to 70% of the population, and the frequency of urinary excretion may be higher in some populations such as pregnant women, older individuals, and organ transplant patients.5-14 At least 1 study found infection of the renal tubular epithelium.15 These observations suggest that the kidney is the chief site of JCV persistent infection. Whether JCV is persistent in the brain remains uncertain.10 However, the DNA sequence of the regulatory region from kidney or urine of individuals infected with JCV is markedly different from the sequence found in the brain of patients with PML16; JCV quasispecies with deletions can be demonstrated in the urine of some patients.17 Other sites of infection have been identified such as bone marrow,18 spleen, tonsils,19,20 and, rarely, lymph nodes. John Cunningham virus has also been detected in the lungs of immunosuppressed patients with and without PML.21
The frequency with which JCV is detected in tissues may be higher in the setting of immunosuppression, whether from infection or medication.21 In 1 study, 15 (31%) and 12 (24%) of 49 patients with multiple sclerosis treated with natalizumab had CD34+ lymphocytes and CD19+ cells, respectively, harboring the virus.22 In 2009, JCV was found in the cerebrospinal fluid of natalizumab-treated patients with multiple sclerosis without PML.23 Importantly, the same study found JCV DNA in CD34+ cells and monocytes at the same frequency in interferon and natalizumab-treated patients with multiple sclerosis, raising doubt that JCV in CD34+ equates to PML risk.
This study aimed to address the frequency, amount, and degree of expression with which persistent JCV infection is evident in a comprehensive set of body tissues from both JCV seronegative and seropositive persons.
This study was approved by the institutional review board of the University of Kentucky College of Medicine, Lexington. Tissue specimens were obtained at the time of autopsy performed in the Department of Pathology at the University of Kentucky on 12 nonimmunocompromised adults with written informed consent. Immunocompetence was defined by the absence of clinical conditions or laboratory findings consistent with an immunological disorder as determined by medical record review. Persons who were at known risk for PML, including those with congenital immunodeficiency syndromes such as AIDS, leukemia or lymphoma, sarcoidosis, and bone marrow or organ transplant, were excluded from analysis.
At autopsy, tissue specimens were obtained from brain (hemisphere cortex, white matter, cerebellar cortex, and white matter), mediastinal lymph node, lung, spleen, kidney, renal pelvis, urinary bladder, stomach, small and large intestine, liver, pancreas, testis or ovary, uterus or prostate, bone marrow, and skeletal muscle. To avoid cross-contamination of samples, separate scalpels were used for each tissue obtained, and the samples were otherwise kept in separate, sealed, sterile containers. One-third of excised tissue was fresh frozen, one-third was frozen in All Protect (Qiagen), and the remaining tissue was fixed and paraffin embedded. Whole blood was not assayed for viremia.
John Cunningham virus serology was performed at Biogen in Cambridge, Massachusetts, by capturing anti-JCV antibodies on MAD1 VLPs in a research-grade enzyme-linked immunosorbent assay similar to the method subsequently licensed to Focus Laboratories. That laboratory was blinded to the tissue sources. Whole blood was used to determine JCV antibody serostatus of all patients using a laboratory-based enzyme-linked immunosorbent assay. For purposes of this study, antibody absorbance was divided into 3 groups: absent, low absorbance level (<0.5), and high absorbance level (≥0.5) based on serum controls with known antibody serology.
Quantification of JCV DNA by Quantitative PCR
DNA was purified from 1.5-mL urine from all participants when available using the QIAamp Viral RNA Mini Kit (Qiagen) following concentration with Amicon ultracentrifugal filters (Millipore) and elution in 70 μL of water. DNA was purified from approximately 20 mg of (10-30) fresh frozen tissues, 200 μL of blood, and 100 μL of bone marrow from all seropositive patients and primarily from renal tissue of seronegative patients using the QIAamp DNA mini kit (Qiagen) and eluted in 200-μL Buffer AE (ie, 1-μL purified DNA from 0.1-mg tissue). Nonrenal tissues were examined in 2 seronegative patients for comparative purposes. Ten microliters of purified DNA were subjected to quantitative PCR (qPCR) using primers and probes specific for JCV detection, ie, the large tumor antigen as described by MacKenzie et al24 and reported as JCV DNA copies/μL purified DNA. The JCV probe was labeled at the 3′ end with the quencher fluorochrome, 6-carboxytetramethyl-rhodamine (PE Applied Biosystems). The 5′ end of the probe was labeled with the reporter fluorochrome, 6-carboxy-fluorescein. Real-time PCR was performed on an ABI Prism 7700 Sequence Detection system (PE Applied Biosystems). Cycling parameters were 50°C for 2 minutes, 95°C for 10 minutes, and 40 cycles of 95°C for 15 seconds and 60°C for 1 minute. Each PCR run contained negative controls, including reaction mixtures without DNA template and several specimens that were known to contain no JCV DNA. Positive controls consisted of a 10-fold dilution series (1 × 10° to 1 × 104 genome equivalents per reaction) of cloned JCV (MAD1 plasmid) sequences kindly provided by E. O. Major (National Institute of Neurological Diseases and Stroke). Each specimen was analyzed in duplicate. Results were scored as positive if either reaction yielded a threshold cycle value greater than the limit of detection for the standards. In the few cases in which amplification was detected in only 1 of the replicate reactions, the qPCR assay profiles were manually inspected to ensure true logarithmic amplification curves. Amplification curves were not obtained in hundreds of negative control assays (data not shown). Polymerase chain reaction product was reliably detected in the control standards containing 1 copy/µL of purified JCV DNA template, allowing for high-confidence quantification of samples at or greater than this concentration such that the limit of quantification was 10 copies/μL of JCV DNA.
To ensure quality in the PCR, 110 blinded DNA samples from the autopsy tissues were sent to the Major Laboratory at the National Institutes of Heatlh. This included 92 unique blinded samples (73 JCV DNA positive; 19 JCV DNA negative); 11 JCV DNA–positive duplicate samples; and 7 serial 10-fold diluted samples (from 1 urine JCV+ specimen with high copy number that was diluted 10 to 107 fold). Ninety-two tissue samples were also independently evaluated at the Biogen research laboratory and the Biogen development laboratory by qPCR. All samples presented as JCV DNA positive were positive in at least 2 laboratories. Mean values were derived collectively from means determined in each laboratory.
The Herculase II Fusion Enzyme system (600677; Agilent Technologies Inc) was used to amplify VP1 and the NCCR regions from DNA isolated for qPCR. The VP1 coding region was amplified with either primers 5′-GCAGCCAGCTATGGCTTTAC-3′ and 5′-GCTGCCATTCATGAGAGGAT-3′ or 5′-GTTGCCCAAAGGGAGGGAACCTATATTTC-3′ and 5′-AGCCTCTGGTGCAGACACACAGGAA-3′, and the NCCR was amplified with either 5′-CCTCCACGCCCTTACTACTTCTGAG-3′ and 5′-AGCTGGTGACAAGCCAAAACAGCTCT-3′ or 5′-GATTCCTCCCTATTCAGCACTTTG-3′ and 5′-TCCACTCCAGGTTTTACTAA-3′. The PCR products were cloned using the TOPO TA Cloning Kit (K4575J10; Life Technologies). Ligation products were transformed and plated, and each colony sequenced on an Applied Biosystems 3730XL using version 3.1 chemistry.
Quantification of JCV Large T-Antigen Expression by Quantitative Reverse Transcription PCR
Total RNA was purified from select replicate tissues that were stored frozen in AllProtect using the AllPrep DNA/RNA Mini Kit (Qiagen). Equal volumes of RNA samples were reverse transcribed and evaluated for spliced JCV large T-antigen message and human β-actin message.
JCV Antigen Detection by Immunohistochemistry
Tissues found to have JCV DNA by PCR were probed for the viral coat protein, VP1, and the T antigen after EDTA for formalin-fixed paraffin-embedded slides and acetone permeablization/fix of frozen sections. VP1 was determined by PAB597 staining at 1-ug/mL concentration; T antigen was stained using Santa Cruz’s V300 rabbit polyclonal sera at 1:50 for formalin-fixed paraffin-embedded sections and 1:100 for frozen sections.
Twelve autopsies were performed. Individuals were not immunosuppressed based on the medical records, although the status of 1 JCV seronegative individual who died of unknown cause could not be ascertained. Seven individuals were JCV seropositive based on controls with known JCV serology (3 high absorbance and 4 low absorbance) including 6 men and 1 woman with an age range of 25 to 67 years (mean, 47.9 years). Five individuals were JCV seronegative including 4 men and 1 woman ranging in age from 59 to 75 years (mean, 65.8 years). The underlying illnesses, causes of death, and JCV status are listed in Table 1.
John Cunningham virus DNA was detected in 8 of 12 individuals. The highest copy numbers of JC viral DNA were in urine, renal, and reproductive tissues; however, JCV DNA was detected in a wide variety of other tissues in low copy number (Table 2). Among the tissues in which JCV DNA was detected were brain, heart, lung, liver, intestine, spleen, lymph node, and thymus gland. John Cunningham virus DNA was not detected in the bone marrows of any of the 4 patients in whom it was studied.
Seven of 12 individuals were JCV seropositive. Serostatus was reflective of the amount of JCV DNA detected, with most JCV DNA–positive tissue samples being found in individuals with JCV antibody high absorbance (Figure 1 and Table 1). In fact, JCV DNA was detected in 53% of tissue samples (50 of 95) from JCV antibody high-seropositive persons, while JCV DNA was present in only 4.9% (6 of 123) and 0% (0 of 75) of tissue samples from JCV antibody low-seropositive and seronegative individuals, respectively. The presence of JCV DNA in urine also was associated with serostatus because quantifiable amounts of JCV were detected in urine from all 3 high–JCV serostatus individuals. In contrast, JCV DNA was present in amounts less than the level of quantification in the urine from 3 JCV-seronegative individuals.
Tissues that were positive by qPCR for JCV DNA were subjected to PCR amplification to examine the sequence of VP1 and the noncoding control region (95 samples from 4 individuals). Only tissues with high viral loads, including the left and right renal pelvis, urinary bladder, and urine from case 2 and urinary bladder and urine from case 6, were successfully amplified, cloned, and sequenced. Viruses from both cases were strain 1B with wild-type VP1. NCCR sequences were either archetype or archetype-like variants. Virus detected in the urine from case 2 exhibited a 14-base pair deletion that spanned the B/C regions, whereas the renal pelvis and urinary bladder did not. Virus from case 6 had an insertion in the F region of 5 bases that were a direct repeat of the immediately 5′ nucleotides (GCCAA) found in both urine and bladder.
To evaluate JCV gene expression, case 2 was evaluated. This case represented an example of large JCV distribution and high copy numbers in tissues (Figure 1). DNA and RNA were sequentially purified from 12 anatomic sites and evaluated for JCV large T-antigen message. John Cunningham virus DNA was not detected in left and right thalamus, left and right cerebellar cortex, or the left primary visual cortex. Low levels (less than the limit of quantification) of JCV DNA were detected in left and right frontal cortex, right primary visual cortex, peripheral nerve, thymus gland, and left kidney, while high levels (1 × 104 copies/μL) were detected in the left renal pelvis. Equal volumes RNA purified from these same tissues were used to generate cDNA. While greater than hundreds of thousands copies of human β-actin message were detected in the cDNA samples, JCV large T-antigen message was only detected in the samples derived from the left renal pelvis.
In an attempt to detect virus infection, immunohistochemistry was performed on either paraffin-embedded or frozen tissues for T-antigen and VP1-protein expression using antibodies raised to SV40 that recognize both JCV and BK virus proteins. The right and left frontal cortex, primary visual cortex, thalamus, mediastinal lymph node, thymus, peripheral nerve, skeletal muscle, prostate, testis, and right and left renal pelvis and urinary bladder from case 2; the kidney, renal pelvis, prostate, and skeletal muscle from case 3; the renal pelvis, bladder, and prostate from case 6; the spleen from case 7; and the renal pelvis and bladder from case 11 were tested. Only case 2 was positive. In this case, the renal pelvis was positive for T antigen (Figure 2) and the bladder was positive for T antigen and VP1, respectively. Control PML case tissue was routinely positive for both T antigen and VP1. Case 2 had the highest urine JCV DNA copy number, suggesting that immunohistochemistry can only detect replicating virus under conditions where large amounts of virus are involved.
This study systematically explored sites of JCV persistence by comprehensively sampling tissues obtained from autopsy in 12 individuals. Among the salient findings from the study is the confirmation that the kidney is the predominant site of viral persistence. Quantitative PCR revealed JCV concentrations in the kidney, renal pelvis, and bladder that, when detected, were 3 to 4 orders of magnitude higher than in other tissues. Viral copy numbers in the urine were often substantially higher than in the renal system tissues. As expected, viruses detected in urine were primarily archetype or archetype-like and did not exhibit the NCCR duplications associated with PML based on the sensitivity of the current PCR method. The current working hypothesis is that viral transmission, presumably with the archetype virus, typically occurs prior to age 20 years when seropositive rates for JCV begin to plateau.25 Because acute illness is not associated with JCV acquisition, the precise timing of the infection is difficult to determine and the mode of transmission remains unclear; however, urine-oral transmission is likely.26 Following infection, the virus develops a persistent infection of the renal system with the archetype virus, with subsequent mutation to the prototype form rendering it capable of causing PML.27
The presence of JCV in the brain in the absence of PML has been controversial. Some investigators28,29 have been unable to detect JCV in the brains of normal controls using PCR methods. Others have detected JCV DNA in 20% to 50% of normal brain tissue from immunocompetent individuals obtained from biopsy or at autopsy.28-33 Although viral DNA may be identified, in the absence of PML, JCV proteins have not been.34 As in some of the earlier studies using PCR to detect viral presence by qPCR,32,33 we detected JCV DNA in low copy numbers in 2 brain tissue samples (18.2%) harvested at autopsy in this nonimmunosuppressed cohort. Similarly, we were unable to convincingly confirm the presence of the virus in the brain using alternate techniques, such as large T-antigen RNA detection or by immunohistochemistry.
As previously noted,35 there was a positive association between JCV copy numbers in the urine and antibody levels with the virus dissemination to nonrenal tissues. Because the antibody assay used was a research-grade assay, it should not be equated with the currently available enzyme-linked immunosorbent assay through FOCUS Laboratories nor should the absorbance values be mistaken for antibody titers. Nonetheless, dividing the groups into seronegative and arbitrary low and highabsorbance values resulted in an intriguing correlation between antibody absorbance and the presence and quantity of virus detected. Virus was identified in 5 to 25 sampled tissues from the 3 individuals in the high JCV antibody absorbance group, with copy numbers ranging from just at the level of quantification to 9 780 102 copies/µL in comparison with only 0 to 5 tissues from the low JCV antibody absorbance, in which copy numbers did not exceed 129 copies/µL. The JCV was not detected in any tissue in the JCV seronegative group; however, as noted, 3 of 4 urine specimens from the latter group had detectable JCV in keeping with an earlier observation.36 Similarly, viral copy numbers were significantly higher in the high antibody group vs the low and negative groups. The average copy number in the high-antibody group was 3 000 000 copies/µL in the urine, and the average urinary tract tissue copy number was 19 500 copies/µL. The average copy numbers in the urinary tract tissue of the low-antibody group was 38 copies/µL, and virus was not detected in the urinary tract tissues of the seronegative group. This observation suggests that higher antibody levels may reflect more widely distributed virus and greater degrees of viral replication. Furthermore, it comports well with the observed association between JCV antibody titers (derived subsequent to this study) and risk of PML.37 The positive association between JCV copy numbers in the urine and the number of tissues infected and viral copy numbers suggests that quantitative assays of urine for JCV in the high-antibody patients may help in identifying those people who are persistently infected and at risk of PML.
This study has the strength of independent JCV DNA detection and quantification from multiple laboratories. However, it is not without its shortfalls. First, the number of persons in this retrospective study was not large. Second, not all 34 tissues were available from each person, and urine results were not always present for analysis. Third, while contamination is a potential issue, we minimized that risk with scrupulous methods including new scalpels for obtaining the tissue for each site.
We would anticipate that in the setting of immunosuppression, the frequency with which JCV could be detected would be significantly greater than what we observed in this study. The mechanism of immune modulation that not only increases the likelihood of JCV expression in peripheral blood6,38 but also predisposes to the development of PML are not fully understood.39,40 We found that persistent (latent or actively replicating) JCV infection mostly predominates in genitourinary tissues, but distributes in other tissues at low copy number. However, we found no evidence of replication outside of the kidney in these particular individuals. The distribution and copy numbers of the virus appear to correlate with urinary JCV shedding and serostatus.
Corresponding Author: Joseph R. Berger, MD, Department of Neurology, Perelman School of Medicine, University of Pennsylvania, 3400 Convention Ave, Perelman Center for Advanced Medicine, Room 765, Philadelphia, PA 19104 (joseph.berger@uphs.upenn.edu).
Accepted for Publication: November 17, 2016.
Published Online: February 27, 2017. doi:10.1001/jamaneurol.2016.5537
Author Contributions: Dr Berger had full access to all the data in the study and takes responsibility for the integrity of the data and the accuracy of the data analysis.
Concept and design: Berger, Miller, Danaher.
Acquisition, analysis, or interpretation of data: All authors.
Drafting of the manuscript: Berger, Miller, Danaher.
Critical revision of the manuscript for important intellectual content: All authors.
Statistical analysis: Doyle, Hack.
Obtained funding: Berger.
Administrative, technical, or material support: Berger, Doyle, Simon, Gorelik, Cahir-McFarland, Singhal, Hack, Owens, Neltner.
Supervision: Berger, Danaher, Simon, Gorelik, Cahir-McFarland.
No additional contributions: Miller, Norton.
Other–helped with the neuropathological issues: Nelson.
Conflict of Interest Disclosures: Dr Berger reports grants from Biogen during the conduct of the study; personal fees from Amgen, personal fees from Astra Zeneca, personal fees from Janssen, Millennium/Takeda, Novartis, Biogen, Roche, Genentech, Genzyme/Sanofi, Inhibikase, Forward Pharma, Johnson and Johnson, Pfizer, and Eisai outside the submitted work. Ms Doyle is an employee of Biogen and owns stock in Biogen. Dr Simon is an employee of Oncorus. Dr Simon has patents 15/063 777, 61/294 048, and 61/316 193 pending to Biogen Inc and was formerly employed by and received salary and stock compensation from Biogen Inc. Ms Norton is an employee of Biogen and owns stock in Biogen. Dr Gorelik is an employee of Fortress Biotech. Dr Gorelik reports personal fees from Biogen Idec during the conduct of the study and personal fees from Biogen Idec outside the submitted work; in addition, Dr Gorelik has patent WO2010090757 A1 pending for detection of John Cunningham virus. Dr Cahir-McFarland is an employee of Biogen and owns stock in Biogen. No other disclosures are reported.
Funding/Support: This study was supported by a grant to Dr Berger by Biogen.
Role of the Funder/Sponsor: The funding source had no role in the design and conduct of the study, the approval of the manuscript, or the decision to submit the manuscript. However, investigators from Biogen, the funding source, collaborated in the performance of the study, in particular, the performance of John Cunningham virus antibody levels on blood and enzyme-linked immunosorbent assay and isolated systolic hypertension studies on tissue specimens and provided assistance in the preparation and review of the manuscript.
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