Failure to detect human herpes simplex virus, cytomegalovirus, and Epstein–Barr virus viral genomes in giant cell arteritis biopsy specimens by real-time quantitative polymerase chain reaction

Cardiovascular Pathology 15 (2006) 280 – 286

Original Article

Failure to detect human herpes simplex virus, cytomegalovirus, and Epstein–Barr virus viral genomes in giant cell arteritis biopsy specimens by real-time quantitative polymerase chain reaction Milena Cankovic4, Richard J. Zarbo Department of Pathology, Henry Ford Hospital, Detroit, MI 48202, USA Received 24 March 2006; received in revised form 10 May 2006; accepted 31 May 2006

Abstract A study provided evidence of human herpes simplex virus (HSV) DNA in giant cell arteritis (GCA) biopsy specimens. This prompted us to study our own GCA biopsy specimens using real-time quantitative polymerase chain reaction for the detection of HSV1, cytomegalovirus, and Epstein–Barr virus DNAs. Our study failed to confirm an association between HSV1 and GCA, revealing no viral genome in 35 biopsy specimens of histologically positive temporal arteries. D 2006 Elsevier Inc. All rights reserved. Keywords: Cytomegalovirus; Epstein–Barr virus; Human herpes simplex virus

1. Introduction Human herpes simplex virus 1 (HSV) has been recently implicated in the etiology of giant cell arteritis (GCA) in a recent study [1], while others failed to find such an association [2]. GCA or temporal arteritis (TA) is a relatively common systemic vasculitis. It is a chronic vasculitis of large and medium-sized arteries, especially the extracranial branches of the carotid arteries [3]. Giant cell arteritis represents a highly complex immune response that is driven by interferon-gproducing T lymphocytes, macrophages in the vascular wall, and maladaptive arterial response to tissue injury [4,5]. Arterial lesions are composed of T lymphocytes and macrophages, which are often arranged in granulomas. The inflammatory response leads to excessive cytokine production, particularly of interleukin (IL)-1 and IL-6, which induces systemic inflammation. In addition, interferon-g, which is released by T cells, activates macrophages, leading to tissue Presented in part at the 95th Annual Meeting of the United States and Canadian Academy of Pathology, February 14, 2006. 4 Corresponding author. Department of Pathology, Henry Ford Hospital, 2799 W. Grand Blvd., Detroit, MI 48202, USA. Tel.: +1 313 916 9448; fax: +1 313 916 9113. E-mail address: [email protected] (M. Cankovic). 1054-8807/06/$ – see front matter D 2006 Elsevier Inc. All rights reserved. doi:10.1016/j.carpath.2006.05.007

injury. Vessel wall inflammation leads to luminal occlusions and tissue ischemia. Cranial arteries are particularly susceptible, but other vessels can be affected as well [3]. Temporal artery biopsy remains to be the diagnostic procedure of choice to detect arteritis in cranial vessels, and temporal arteries routinely undergo biopsies to establish a histologic diagnosis. The incidence of GCA is 15–25 per 100,000 in at-risk populations [6]. Susceptible persons are usually older than 50 years, and incidence rates increase with advancing age [7]. The cause of GCA is unknown. Some familial accumulation and the association with certain HLA-DR haplotypes seem to indicate a genetic predisposition. Variations in the incidence of GCA have been reported in different parts of the world, suggesting that environmental or genetic factors may be involved [8,9]. The disease appears to be most prevalent in Caucasians, especially those of Scandinavian origin [10]. As with many autoimmune diseases, infectious agents have been suspected as possible initiators of the inflammatory process in GCA [11]. Flu-like symptoms are often seen at the onset of the disease and are followed by an acute phase response, and molecular studies on GCA have suggested a possible link between infection and GCA. Except for the histopathology of the arterial wall [12,13], no diagnostic test

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that can be widely used for diagnosing GCA exists. In addition, a high percentage of patients present with nonspecific symptoms that do not conform to any specific set of criteria. Although some studies have presented evidence for a relationship between GCA and infection with several infectious agents such as Parvovirus B19, Chlamydia pneumoniae, and human herpes viruses [14–17], others have failed to detect the presence of these same infections by qualitative polymerase chain reaction (PCR)-based assays in GCA specimens [2,18]. Other studies have attempted to identify factors associated with GCA. In a retrospective study on 250 patients diagnosed with TA and followed for 27 years, Liozon et al. [19] found no association between TA and microbial infection. Narva´ez et al. [20] retrospectively studied a group of 143 patients [85 with polymyalgia rheumatica (PMR), 22 with TA, and 36 with both PMR and TA] to determine if environmental factors such as infectious agents and seasonal effects played a role in the etiology of these diseases. They were unable to confirm a seasonal pattern or an association between infection and the onset of either PMR or TA over a period of 13 years. Direct evidence for the detection of HSV in association with GCA was recently reported by Powers et al. [1], who were able to detect the presence of HSV DNA sequences by PCR in archival temporal artery biopsy specimens. This finding prompted us to confirm these intriguing results by applying two clinically validated PCR-based methods. We used realtime quantitative PCR (QPCR) to see whether HSV DNA could be detected in archival temporal artery biopsy specimens from patients with pathologically confirmed GCA or if cytomegalovirus (CMV) or Epstein–Barr virus (EBV), both of which have been implicated in the etiology of cardiovascular disease [21–23], might be involved. In addition, and to further increase the sensitivity of detection of HSV DNA, we used the minor groove binding (MGB) Eclipse HSV1/2 ASR (Nanogen, San Diego, CA) assay, which allowed us to detect the presence of HSV1 and/or HSV2 DNA in the temporal artery biopsy specimens. Finally, to ensure that our assays targeted portions of HSV genomes that remained preserved during tissue fixation, we used the PCR primers provided to us by Powers et al. [1] for qualitative HSV detection.

2. Methods 2.1. Case selection The study samples consisted of 35 cases of histologically confirmed GCA and 6 cases of temporal artery biopsy

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specimens with no inflammation. In addition, we selected 5 control cases as follows: giant cell myocarditis, n = 2; granulomatous inflammation of the skin, n = 2; and soft tissue inflammation, n = 1. When selecting controls, we tried to include cases with similar inflammatory reactions, with unrelated inflammation, and arterial biopsy specimens with no evidence of inflammation. The control tissues had been processed in the same manner as the arterial biopsy tissues under study were. All samples were formalin-fixed and paraffin-embedded biopsy specimens from the surgical pathology files of the Henry Ford Hospital collected from 1998 to 2005. The surgical pathology reports were reviewed to extract demographic parameters and pathologic characteristics of all samples (Table 1). For GCA cases, arterial biopsy specimens consisted of serially sectioned specimens. Hematoxylin– eosin-stained slides were reviewed, and histologic diagnoses were confirmed in all cases. 2.2. DNA isolation DNA was isolated using a QIAamp DNeasy Mini Kit (Qiagen, Valencia, CA). Briefly, five to six 10-Am sections were cut from paraffin-embedded biopsy material and placed into microcentrifuge tubes. Sections were dewaxed with xylene, treated with absolute ethanol, and dried as well as digested overnight at 568C with proteinase K (Qiagen). After column purification, DNA was eluted in 85 Al of an elution buffer (Qiagen). 2.3. Real-time QPCR To evaluate latent infections, we used QPCR assays and dual-labeled TaqMan probes for the detection of genomic sequences in HSV1, CMV, and EBV viral genomes. Specific TaqMan primers and fluorescent probes for the detection of HSV1 glycoprotein G were designed as originally described by Rycarz et al. [24] The forward HSV1 typing primer was 5V-TCC TG/CG TTC CTA/C ACG/T GCC TCC C-3V, and the reverse HSV1 typing primer was 5V-GCA GIC AC/TA CGT AAC GCA CGC T-3V. Primers and probes for CMV and EBV detection were designed as originally described by Kubar et al. [25]; the primer sequences were 5V-TGA GCC CGG CGG TGG T-3V as well as 5V-AGC TCA CCG ATC ACA GAC AC-3Vfor CMV and 5V-CCT GGT CAT CCT TTG CCA-3V as well as 5V-TGC TTC GTT ATA GCC GTA GT-3V for EBV.

Table 1 Patient demographics Diagnosis

Vessel involved

Cases (n)

Age (y; mean)

Female/male ratio

GCA Arterial biopsy (no inflammation) Giant cell myocarditis Skin (granulomatous inflammation) Soft tissue inflammation

Temporal artery Temporal artery Heart explant Skin biopsy specimen Soft tissue

35 6 2 2 1

81 70 36 11 and 65 71

24:11 5:1 2:0 1:1 0:1

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The TaqMan probes were labeled with the 6-carboxyfluorescein (FAM) reporter dye at the 5Vend and with the 6-carboxytetramethyl-rhodamine (TAMRA) quencher dye at the 3Vend. The TaqMan probes were as follows: 5V-FAMCGT CTG GAC CAA CCG CCA CAC AGG T-TAMRA-3V for HSV1; 5V-FAM-AGA GAA GCG CCA CAT ACA GCG C-TAMRA-3Vfor CMV; and 5V-FAM-CAG TAC GAG TGC CTG CGA CCA-TAMRA-3Vfor EBV. To test the general integrity of DNA extracted from our archival samples, we amplified the cyclophilin gene using previously published primer and probe sequences [26]. The HSV1, CMV, and EBV primers and probes were used as previously described [24,25]. Each 25-Al PCR mix contained 5 Al of extracted DNA, 800 nM of each primer, and 100 nM of each probe. All three PCRs were performed separately using Rotor Gene 3000 (Corbett Research, Sydney, Australia) under the following conditions: incubation for 2 min at 508C and 10 min at 958C, followed by 45 cycles of 15 s at 958C and 1 min at 608C. The fluorescent dye intensities were read automatically during PCR cycling. The threshold cycle number (C t value) represented the cycle number at which a positive amplification reaction was measured. Each specimen was run in duplicate; only those specimens for which the values in both replicates were above the cutoff were considered positive. 2.4. Nanogen MGB Eclipse protocol Minor groove binding Eclipse HSV1/2 ASR (Nanogen) was additionally used to test all study samples for the presence of HSV DNA. The MGB Eclipse detection reagents, which included HSV1/2-positive controls, were used according to the manufacturer’s protocol, and 10 Al of sample DNA was used for each PCR. The primers and the MGB probe in the kit have been designed to target genomic sequences of HSV1 and HSV2 glycoprotein D. Quantitative PCR was performed on Rotor Gene 3000. The reaction mixture was subjected to initial denaturation of 958C for 2 min followed by 45 cycles of 958C for 5 s, 568C for 20 s, and 768C for 20 s. Data were collected during the annealing step. Standard curves were generated, and C t values of patients’ samples were compared with those of positive controls.

precast 1.2% agarose gels (Cambrex Bio Science, Rockland, ME, USA). 2.6. Assay controls and contamination prevention During the initial stage of this study, we established the sensitivity parameters of the assay by isolating viral DNA from the HSV1 culture supernatant, kindly provided by Dr. E. Burd of the Microbiology Division of the Henry Ford Hospital. The sensitivity of the real-time TaqMan QPCR assay was determined using serial dilutions of this viral preparation. Copy numbers were assigned by comparing C t values of the laboratory-isolated HSV1 DNA with those of commercially purchased controls (Nanogen). Laboratoryprepared viral DNA was then diluted to obtain viral copy numbers that ranged from 1 million copies to 1 copy to establish the sensitivity of detection. The limit of detection achieved was fewer than 5 viral copies per reaction. Five concentrations of this viral DNA were selected to be used as standards for the generation of standard curves (Fig. 1). To test whether viral DNA remained preserved after tissue processing, we obtained formalin-fixed and paraffinembedded tissue sections that were already in use as positive controls for HSV, CMV, and EBV staining in the immunohistochemistry section of our pathology department. These samples had been fixed and processed in a manner similar to how the study samples were. Viral DNA from these positive controls was isolated in a manner identical to that observed for the study specimens. Copy numbers for each of these viral isolates were again assigned by comparing their C t values with those of commercially purchased controls (Nanogen). As expected, high viral copy numbers were detected in these controls. A positive control for the MGB Eclipse HSV1/2 ASR assay was included in the kit. For negative controls in all assays, we used a PCR mix with no DNA added. To prevent possible contamination of test samples, we observed standard precautions during all testing steps. Each tissue block was cut separately by a gloved histotechnologist, and equipment were disinfected between different blocks. The subsequent DNA isolation and PCR setups were performed in designated areas, and no more than six specimens were processed at one time. A negative control (no DNA tube) was included with every PCR run.

2.5. Human herpes simplex virus detection by qualitative PCR 3. Results Primers for HSV detection were kindly provided to us by Powers et al. [1]. These primers were designed to target the viral polymerase and were identical to the primers used in the study by Powers et al. [1]. Polymerase chain reactions were set up following the protocol of Powers et al. [1]. We used our low-level positive HSV1 control as the positive control in the PCR assay; a PCR mix with no DNA added was used as the negative control. All the study samples were tested in duplicate. The PCR products were detected using

Initially, we evaluated the ability of the TaqMan real-time QPCR to detect viral load over a wide range of viral copy numbers by testing HSV1 isolated from the culture supernatant. Precise separation on a logarithmic scale between high- and low-titer specimens (Fig. 1A) indicated that the precision of the technique and the instrumentation were acceptable and that linear quantitation was achieved (Fig. 1B). Clear discrimination between the results for the positive

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Fig. 1. (A) Amplification plots of serially diluted viral standards amplified by QPCR. Concentrations range from 1 million viral copies to 10 viral copies per reaction. The threshold cycle number (C t value) represents the cycle number at which a positive amplification reaction is measured. (B) Calibration curve for the detection of viral copy numbers. The log quantity of control samples is plotted versus cycle threshold (C t) to generate the standard curve from which unknown samples are quantified.

samples and those for the negative samples was seen with every run. We then demonstrated that viral DNA remained preserved in formalin-fixed and paraffin-embedded tissues as processed in our surgical pathology laboratory. Positive controls for HSV, CMV, and EBV staining were obtained from the immunohistochemistry section and were subjected to the same DNA isolation protocol as used for the study group. Viral copy numbers of 105 per 25-Al reaction for HSV, 105 per 25-Al reaction for CMV, and 103 per 25-Al reaction for EBV were recovered. Because of a high viral load in these

positive controls, the viral copy numbers detected in these fixed samples were higher than could be reasonably expected to be present in the study samples. However, because the sensitivity of detection of the QPCR assay was fewer than five viral copies per reaction, even very low viral copy numbers, as would be expected to be present in biopsy samples, would still be detected by our protocol. Once the assay parameters had been established, all the patient samples were tested in duplicate. Serially diluted positive controls were included in all runs, and standard

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Fig. 2. Amplification plots of representative patient samples amplified by QPCR and MGB Eclipse ASR. Flat lines indicate negative results. (A) HSV1 detection by QPCR; (B) CMV detection by QPCR; (C) EBV detection by QPCR; (D) HSV1 detection by MGB Eclipse HSV1/2 ASR and QPCR.

curves were generated from these. The C t values for patient’s samples were then compared with those from the standard curve. Furthermore, to ensure that DNA had been extracted sufficiently and that the quality as well as quantity of isolated DNA were adequate for PCR amplification, we amplified aliquots of isolated DNA for human cyclophilin gene fragments. All samples, including controls, gave positive signals, indicating that isolated DNA was of sufficient quantity and quality for PCR amplification. The following QPCR results were obtained for the study samples: 1. We detected no HSV1 viral DNA in any of the GCA samples tested. 2. CMV was detected in one of the GCA biopsy specimens.

3. The two skin specimens were both positive for the presence of EBV DNA, whereas all the other samples were negative. Fig. 2 shows representative patient results for the detection of HSV1 (A), CMV (B), and EBV (C). The summary of results for the entire study group is presented in Table 2. We additionally tested all the study samples in duplicate for the presence of HSV viral DNA using HSV1/2 ASR (Nanogen). A representative set of patient results is shown in Fig. 2D. Again, no evidence of HSV infection was found in any of the GCA or control samples tested (Table 2). To address the possibility that the viral glycoprotein G and glycoprotein D, which were targeted with our QPCR and MGB Eclipse primers, became destroyed during the fixation process and therefore lost as PCR targets, we used a

Table 2 Viral genome detection summary Diagnosis

HSV1 TaqMan (+/ )

HSV1/2 Eclipse MGB (+/ )

CMV (+/ )

EBV (+/ )

GCA Giant cell myocarditis Arterial biopsy (no inflammation) Skin (granulomatous inflammation) Soft tissue inflammation

0/35 0/2 0/6 0/2 0/1

0/35 0/2 0/6 0/2 0/1

1/35 0/2 0/6 0/2 0/1

0/35 0/2 0/6 2/2 0/1

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set of primers targeting HSV viral polymerase. These primers were kindly provided to us by Powers et al. [1] so that we could address the discrepancy issues in the two studies. None of the samples in our study showed reactivity when we used the viral polymerase primers. Our low positive HSV1 control, which was diluted to a level of 10 viral copies per reaction, showed a distinct positive band.

4. Discussion Vascular lesions of GCA are a consequence of inappropriate activation of the immune system, particularly T lymphocytes [5,27–29]. As with some other autoimmune diseases, infectious microorganisms have been suspected as possible instigators and the spectrum of suspects ranges from viral agents to bacterial organisms [30 – 32,14]. In the study by Powers et al. [1], HSV DNA was detected in 88% of histologically positive and 53% of histologically negative temporal artery biopsy specimens. These investigators studied formalin-fixed and paraffin-embedded tissue sections with PCR amplification followed by Southern blot detection of HSV DNA. Because viral DNA had been detected in such a large percentage of specimens, this study suggested that HSV either was a causative agent of GCA or played a major role in the pathogenesis of the disease. In this study, we used diagnostic methods of viral detection that are currently considered to be very sensitive and reliable; we were unable to reproduce the results observed by Powers et al. [1]. We found no evidence that the continuous presence of viral agents such as HSV1, CMV, and EBV plays any role in the pathogenesis or molecular diagnosis of GCA. We used real-time QPCR, a method of detection adopted by many clinical laboratories, and detected no evidence of viral infection in 35 cases of active GCA and 6 cases of clinically suspected GCA with no evidence of inflammation, with the exception of 1 GCA case, which was positive for CMV infection. Our results are in agreement with those of Helweg-Larsen et al. [2], who used frozen biopsy specimens as well as qualitative PCR-based testing and failed to detect evidence of human herpes virus DNA (HSV1, CMV, and EBV) in 30 temporal artery biopsy specimens from patients suspected of having GCA. One reason for the marked discrepancy that we report on might be explained by the fact that our method of detection differed from the one used by Powers et al. [1] They used PCR in combination with Southern blot analysis for the detection of HSV DNA, whereas we used real-time QPCR and MGB Eclipse real-time PCR. The PCR primers and probes in our study were designed to target genomic sequences for HSV glycoprotein G (QPCR) and glycoprotein D (MGB Eclipse) rather than the genomic sequence for viral polymerase that was used as the target in the study by Powers et al. [1]. In our study, the QPCR protocol had been validated for clinical use by two laboratories [24,33]. We additionally incorporated the MGB Eclipse protocol to

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confirm the results obtained by the QPCR protocol and to possibly increase the level of detection. Minor groove binding probes have been shown to have a high sensitivity of detection and are often recommended for use with difficult-to-amplify samples. In fact, MGB Eclipse HSV1/2 ASR reagents are available commercially and are recommended for clinical use for HSV detection. The results obtained with the MGB Eclipse protocol were 100% concordant with the results observed with the QPCR protocol in our study, providing a further reassurance of the validity of our results. To address the issue of different methodologies, Powers et al. [1] kindly offered to provide us with their primers so that the primer specificity can be addressed. We detected our lowlevel positive control (10 viral copies/reaction) as positive; however, we were unable to detect the evidence of HSV genomes in any of our study samples tested in duplicate. Although we did not proceed with the Southern blot detection step, we feel that the sensitivity of detection of qualitative PCR alone was sufficient to make us conclude that the negative results observed in our study were valid. We do not know whether the slight difference in methods used affected the outcome of the two studies or whether study samples differed in some way. The fact that we were repeatedly able to detect presence of HSV DNA in our controls by all three methods would argue that the sensitivity of detection in our study was sufficient to allow us to detect presence of HSV DNA even at low viral copy numbers. We did deviate from the manufacturer’s instruction in one aspect when using MGB Eclipse ASR. The results were obtained by analyzing the standard curves from real-time PCRs only, and no melting curve analysis was done because of technical limitations. Although we did observe a 100% concordance between our QPCR results and our MGB Eclipse results, we do not know whether the additional step of melting curve analysis would have allowed even greater sensitivity. One factor that might have affected the overall sensitivity of detection in our study was the fact that the amount of DNA recovered for 10 (24%) of 41 arterial biopsy specimens tested was low (b200 copies of genomic DNA per 25-Al reaction). This percentage is still much lower than the percentage of HSV-positive cases reported by Powers et al. [1] and would not explain our inability to detect HSV DNA in GCA biopsy specimens at the rate reported by these authors. In conclusion, using sensitive real-time QPCR assays for the detection of HSV1, CMV, and EBV DNAs that consistently achieved detection levels of fewer than five viral copies per 25-Al reaction, we cannot confirm the presence of HSV1 in biopsy specimens of histologically positive temporal arteries and that in biopsy specimens of temporal arteries with no evidence of inflammation. This study does not imply that microbial pathogens do not play a role in the pathogenesis of GCA. Although we failed to show the presence of HSV1, CMV, and EBV viral genomes in temporal artery biopsy specimens, only three viruses were included in this study. There is a possibility that persistence or

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recurrence of infection by other microbial pathogens, including Parvovirus B19 and Chlamydia pneumoniae, might be involved in GCA etiology.

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