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1 Department of Pathology and Animal Health, Faculty of Veterinary Medicine, Naples University Federico II, Naples, Italy
2 Laboratory of Virology, Regina Elena Cancer Institute, Rome, Italy
3 Department of Experimental Medicine and Clinics, Catanzaro University Magna Graecia, Catanzaro, Italy
Correspondence
Sante Roperto
sante.roperto{at}unina.it
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
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A supplementary figure showing sequence data is available with the online version of this paper.
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
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; Chambers et al., 2003; de Villiers et al., 2004). BPV-2 infection in the presence of environmental carcinogens, such as ptaquiloside (PT) of bracken fern (Pteridium aquilinum), has been associated with urinary bladder neoplastic lesions in adult cattle, in which chronic enzootic haematuria (CEH) is the most important clinical sign (Campo, 1997; Campo et al., 1992; Hopkins, 1986).
The effect of the route of viral infection, and the synergistic relationship between BPV-2 and immunosuppressive and oncogenic compounds present in the bracken fern in the malignant progression of bladder lesions (Campo, 1997; Campo et al., 1992; Jarrett et al., 1978; Reddy & Fialkow, 1983; Stocco dos Santos et al., 1998) are not well-known, thus deserving further investigations.
To date, the BPV-2 genome has been detected in lymphocytes during latent papillomavirus infection in cattle (Campo et al., 1994). In addition, the occurrence of horizontal transmission of BPV-2 has been reported in healthy cattle experimentally inoculated with peripheral blood from haematuric animals (Stocco dos Santos et al., 1998). More recently, BPV-2 DNA was detected in seven of 12 urinary bladders and 10 of 14 blood samples obtained from Brazilian cattle suffering from CEH. No histological diagnosis of any tumours was performed. In addition, BPV-2 was also detected in one urinary bladder and in one whole blood sample from asymptomatic cattle (Wosiacki et al., 2005).
In an attempt to gain insights into understanding the role and significance of BPV-2 presence in the blood stream, an analysis of blood samples was carried out on a large number of haematuric cattle grazing on bracken fern-infested lands, where urinary bladder tumours occur endemically (Borzacchiello et al., 2003).
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METHODS |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
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Samples of neoplastic and control bladder mucosa were split in two. One half was immediately fixed in 10 % neutral formalin and embedded in paraffin. Ticked sections (5 µm) were cut and stained with haematoxylin and eosin. Neoplastic lesions were classified according to the criteria reported in the recent World Health Organization (WHO) Blue Book on the pathology and genetics of tumours of the urinary system and male genital organs in humans (Lopez-Beltran et al., 2004; Sauter et al., 2004). Tumours of the urinary bladder in cattle share many morphological similarities to their human counterparts, and the WHO histological classification appears to encompass all microscopic patterns of the urinary bladder tumours observed in cattle (Roperto et al., 2007). The other half of the sample was immediately frozen in liquid nitrogen and stored in dry ice.
BPV-2 DNA detection and sequencing.
Unfractionated whole blood samples and frozen samples of bladder tumours were analysed by PCR as described previously (Borzacchiello et al., 2003). Briefly, venous blood was collected in heparinized vacutainers (BD Biosciences) and then stored at 4 °C; DNA was extracted with QIAamp DNA mini kit (Qiagen) according to the manufacturer's instructions. Frozen bladder samples were minced and incubated at 55 °C overnight with 60 µg proteinase K ml–1 in the appropriate buffer (Manos et al., 1989) and then at 95 °C for 10 min to inactivate the enzyme. Each sample (10 µl) was amplified in 50 µl reaction mixture containing 3 mM MgCl2, 1 U Platinum Taq (Invitrogen), 25 pmol each primer and 200 µM dNTPs. The reaction was carried out in an iCycler (Bio-Rad Laboratories) using forward (5'-TTGCTGCAATGCAACTGCTG-3') and reverse (5'-TCATAGGCACTGGCACGTT-3') primers that amplify a DNA fragment encompassing part of the E5 and L2 open reading frame (ORF) of BPV-1 (311 bp, from nt 3915 to 4226) and BPV-2 (306 bp, from nt 3919 to 4225) (Otten et al., 1993). PCR conditions were as follows: denaturation for 3 min at 95 °C, followed by 35 cycles of denaturation at 95 °C for 45 s, annealing at 50 °C for 45 s and extension at 72 °C for 1 min. The final PCR products were electrophoresed in 2 % agarose gel and visualized by ethidium bromide staining.
To confirm the PCR data, the amplified band was excised from the gel and purified through silicagel membranes by using the QIAquick PCR quantification kit, according to the manufacturer's instructions (Qiagen). Then, the amplified DNA was subjected to direct sequencing in an automated apparatus (Biogen).
Detection of circular BPV-2 DNA.
BPV-2-positive DNA samples from tissues and blood were analysed by rolling-circle amplification (RCA) according to the method developed by Rector et al. (2004). The multiple-primed RCA is a method that utilizes the 
29 DNA polymerase with random hexamer primers to amplify the complete circular genome of papillomaviruses without the need for prior knowledge of their DNA sequences. Briefly, multiple-primed RCA was performed with the TempliPhi 100 amplification kit (Amersham Biosciences) according to the manufacturer's instructions. Extracted DNA (2 µg) was transferred into a 0.5 ml tube with 5 µl TempliPhi sample buffer containing 450 µM extra dNTPs, and 0.2 µl TempliPhi enzyme mix containing the 29 DNA polymerase and exonuclease-protected random hexamers in 50 % glycerol. The samples were denatured at 95 °C for 3 min and then placed on ice. The reaction mixtures were incubated overnight (approx. 16 h) at 30 °C. Afterwards, the reaction mixtures were placed on ice, subsequently heated to 65 °C for 10 min to inactivate the 29 DNA polymerase, and stored at –20 °C until further analysis. The multiply primed RCA products (2 µl) were digested with 10 U EcoRI, a single cutter of the BPV-2 complete genome, and resolved in an ethidium bromide-stained agarose gel. As a negative control, water was amplified by multiple primed RCA and 2 µl was digested with EcoRI.
BPV-2 mRNA in blood samples.
Blood samples were collected from 15 animals with cancer and stored in PAXgene blood RNA tubes (Qiagen) containing a proprietary reagent that immediately stabilizes intracellular RNA. Purification of total RNA from bovine whole blood was performed with the PAXgene blood RNA kit (Qiagen) according to the manufacturer's instructions. RNA (300 ng) was reverse transcribed using the Superscript III First Strand kit (Invitrogen) in a final volume of 20 µl. To rule out the presence of contaminating DNA in RNA samples, all the assays were performed both with and without reverse transcriptase. The synthesized cDNA was analysed by PCR with specific primers for the E5 ORF (forward primer, 5'-CACTGCCATTTGTTTTTTTC-3'; reverse primer, 5'-GGAGCACTCAAAATGATCCC-3') using the PCR conditions described above, but using an annealing temperature of 48 °C. The final amplified products were electrophoresed in a 2 % agarose gel and visualized by ethidium bromide staining. The amplified band was excised from the gel and subjected to direct sequencing as above.
Protein isolation and E5 expression.
Fifteen bladder samples stored in dry ice were homogenized in lysis buffer containing 50 mM Tris/HCl pH 7.5, 150 mM NaCl, 1 % Triton X-100. Immediately prior to use, the following were added: 1 mM DTT, 2 mM PMSF, 1.7 mg Aprotinin ml–1, 25 mM NaF, 1 mM Na3VO4 (Sigma-Aldrich) by Ultra-turrax T8 Ika-Werke. The proteins were removed by centrifugation at 12 300 g for 30 min at 4 °C. The protein concentration was measured using the Bradford assay (Bio-Rad).
Blood samples (30 ml) were harvested from cattle suffering from chronic enzootic haematuria. An equal volume of 1x PBS was added to the blood and the resulting solution was transferred on a 2 : 1 Ficoll gradient (Biochrom AG). After centrifugation at 352 g for 20 min at room temperature, the recovered lymphocytes were homogenized with 50 µl lysis buffer (described above for bladder samples). The protein concentration was measured using the Bradford assay (Bio-Rad).
Proteins derived from tissues (1 mg) or from lymphocytes (500 µg) were immunoprecipitated by using 2 µg antibody anti-E5 (a kind gift from Dr M. S. Campo, University of Glasgow, Scotland) and 30 µl G-Sepharose (GE Healthcare). Immunoprecipitates were washed four times in complete lysis buffer (above), finally we added 4x LDS loading buffer (Invitrogen) and then heated at 70 °C for 10 min. Immunoprecipitates were separated on 4–12 % polyacrylamide gels and transferred to nitrocellulose filter membranes (Bio-Rad) for 16 h at 25 mA in 192 mM glycine/25 mM Tris-HCl (pH 7.5)/10 % methanol. Membranes were blocked for 1 h at room temperature in 5 % non-fat dried milk and incubated with primary antibody overnight at 4 °C. After three washes in Tris-buffered saline, membranes were incubated with rabbit anti-sheep IgG–horseradish peroxidase (HRP) (Santa Cruz) for 30 min at room temperature. Proteins were visualised by enhanced chemiluminescence (Amersham Biosciences).
Cytospin preparations and immunocytochemistry.
Blood from cows with haematuria was collected in heparin-coated tubes. Blood (100 µl) was lysed with 700 µl erythrocyte lysis buffer containing 0.8 % ammonium chloride and then leukocytes were washed twice by centrifugation at 250 g in 1 % PBS. A 100 µl aliquot of each sample was put into the appropriate well of a cytospin chamber (Thermo Scientific) and centrifuged at 164 g for 5 min at 4 °C. After fixing in methanol, slides were blocked for endogenous peroxidase activity in 0.3 % H2O2 in methanol for 20 min. Slides were then incubated overnight at room temperature in a humidified chamber with a 1 : 500 dilution of polyclonal sheep anti-E5 antibody (kindly provided by Dr M. S. Campo). The slides were washed three times with PBS, then incubated for 30 min with a 1 : 100 dilution of the appropriate biotinylated rabbit anti-sheep IgG (Santa Cruz). They were washed three times with PBS and then incubated with streptavidin-conjugated HRP (LSAB kit; DakoCytomation). Colour was developed by treatment with diaminobenzidine (DakoCytomation) for 5–20 min. Sections were counterstained with Mayer's haematoxylin.
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RESULTS |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
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BPV-2 DNA detection
PCR analysis, validated by direct sequencing of the amplified product, demonstrated the presence of true BPV-2 sequences; only the sequenced samples were defined as being positive. BPV-2 sequences were detected in 39 of the 50 tumour samples (78 % of examined cases) and in seven of 18 normal bladder control samples (39 % of the examined cases). In contrast, none of the analysed samples showed the presence of BPV-1 sequences. The difference in the presence of BPV-2 DNA between pathological and normal samples was highly significant (Fisher's exact test, P<0.004), suggesting a strong association of BPV infection with bladder tumours. Direct sequencing detected some false-positive samples arising from bovine genomic sequences (about 2 % of all analysed samples), as reported previously (Borzacchiello et al., 2003).
BPV-2 sequences were detected in 61 of 78 blood samples of haematuric animals (78.2 % of the examined samples) and in two of 14 healthy samples (
14.2 % of the examined samples) (Fig. 1a). The difference in the presence of BPV-2 DNA between pathological and normal blood samples was highly significant (Fisher's exact test, P<0.001).
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Sequence analysis
Sequence analyses of the positive bladder and blood samples from the same animal were done using CLUSTAL W (http://www.ebi.ac.uk/clustalw/) and BLAST [http://www.ncbi.nlm.nih.gov/BLAST (Altschul et al., 1997)] programs. This revealed the presence of DNA sequences matching the BPV-2 complete genome from the GenBank database, and in one case revealed the presence of two mutations in the non-coding region between the E5 and L2 genes (Supplementary Fig. S1, available in JGV Online). Alignment of this DNA sequence with those in the database revealed that the same mutations were present in the DNA sequence of a BPV-2 isolate from Hungarian equine sarcoids (GenBank accession number AF102551
[GenBank]
).
BPV-2 physical status and mRNA expression
Episomal (circular) forms of the BPV DNA were detected in 39 bladder tumour and blood samples collected from the same animals by using the multiple primed RCA (Fig. 2). Samples were digested with EcoRI, which recognizes a single cut site in the BPV-2 genome, and the detection of a single 8000 bp band indicated the presence of episomal forms. Despite the fact that this was a non-quantitative method, it is noteworthy that the band intensity of circular BPV DNA recovered from blood samples was always lower than that detected in tumour samples. Unfortunately, RNA from these samples was not available for the detection of viral transcripts, so we were unable to verify the expression of the viral genes. Nevertheless, the presence of viral transcripts for the E5 ORF was detected in two of three blood samples from other animals with BPV-2-positive bladder tumours (Fig. 1b
).
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) and in peripheral blood cells from animals with cancer (Fig. 3b). Analysis of cytospin preparations of peripheral blood leukocytes from infected animals demonstrated a strong cytoplasmic E5 immunoreactivity in numerous lymphocytes (Fig. 4). No E5 oncoprotein expression was detected in healthy animals.
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
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). These facts should be taken into account when evaluating a positive result from a PCR using consensus primers if this is the only diagnostic method used. In this analysis, only sequenced samples were considered; this revealed the presence of BPV-2 in whole blood samples. Moreover, RCA analysis suggested that the complete genome of BPV-2 was present in circular form in blood samples as well as in bladder tumours.
Our data on a large number of animals with histologically confirmed neoplastic disease not only clearly corroborate previous reports (Campo et al., 1994; Stocco dos Santos et al., 1998; Wosiacki et al., 2005) but also raise a number of questions about the source, role and significance of the presence of BPV DNA and viral oncoproteins in blood.
The simultaneous presence of BPV-2 DNA sequences in most of the blood and bladder tumour samples suggests that BPV-2 DNA could be derived from common sources, thus implying that the virus might spread via a haematogenous route. Recently, human papillomavirus (HPV) DNA was detected in peripheral blood mononuclear cells (PBMCs) obtained from paediatric patients and healthy blood donors. It has been speculated that PBMCs may serve as a source of HPV in the infection of epithelial cells (Bodaghi et al., 2005). If this is also the case in cattle, BPV-2 could reach the urothelial tissue not only from the paragenital area, as already suggested (Campo, 2002), but also via the blood stream, and could become involved in urinary bladder tumour development independently or jointly with several biological and/or chemical co-factors. PT, the major carcinogen of the bracken fern, is activated in alkaline environments such as the urine of herbivores (Borzacchiello et al., 2006; Prakash et al., 1996), it then appears to act synergistically with BPV-2, thus resulting ultimately in tumours of the urinary bladder in cattle. Recently, BPV DNA was simultaneously detected in different tissues of the same animals, thus suggesting a haematogenous virus spread (Freitas et al., 2007). In our study, the detection of circulating BPV DNA in only a limited number of healthy cattle that share grazing lands with animals with cancer suggests the plausibility of this hypothesis. We not only detected circulating BPV DNA sequences but also showed the presence of episomal DNA by RCA analysis. The presence of circular BPV DNA implies that at least E1 and E2 proteins are expressed, as they are absolutely required for the maintenance and replication of the episomal BPV DNA (Tonon et al., 2001). Moreover, mRNAs encoding the E5 gene were detected in a number of blood samples from animals suffering from BPV-positive bladder tumours. In addition to this, the E5 oncoprotein was detected in the blood of 13 of 15 cows suffering from urothelial tumours, in which E5 expression was documented in immunocytochemical analysis. Taken together, these data may indicate a biological activity of BPV-2 in the blood of these animals, reinforcing the hypothesis that the bloodstream may work as reservoir for BPV infection. In particular, it appears that lymphocytes are infected by BPV-2, as clearly demonstrated by the strong immunocytochemical positivity for the E5 protein.
An alternative explanation for circulating BPV DNA is that it might, at least in part, originate from primary bladder tumours. The presence of viral DNA in the peripheral blood fraction (PBF), plasma and serum is not an unusual event in tumour patients. Recently, HPV DNA was detected in serum and/or PBF of individuals with cervical, head/neck or schistosomiasis-associated bladder cancer (Yang et al., 2005) and women suffering from cervical carcinomas (Ho et al., 2005; Widschwendter et al., 2003). It has been postulated that HPV DNA is involved in carcinogenesis in breast tissue in some patients suffering from cervical cancer (Widschwendter et al., 2004). Very recently, sequences of the HPV types 16 and 18, known to be involved in cervical carcinogenesis, were detected in 86 and 48 % of breast cancer samples, respectively (de Villiers et al., 2005; Kan et al., 2005). It is noteworthy that HPV DNA has been detected both in lung and breast cancers of patients with a history of cervical cancer, thus suggesting that HPV DNA might be transported from the original site of infection to pulmonary and breast tissues by the bloodstream and subsequently be involved in secondary tumour development (Cheng et al., 2001; Widschwendter et al., 2004). However, it is well-known that urinary bladder cancers in cattle are characterized by a relatively low incidence of apparent metastases (about 8–10 %) (Pamukcu et al., 1976; Roperto et al., 2005). In addition, non-metastatic tumours in other organs of cattle with a history of urothelial cancer are known to occur very rarely. Therefore, it is reasonable to suggest that if the presence of BPV-2 E5 oncoprotein in PBMC is the result of a passive spread, the oncoprotein doesn't appear to play an important role in neoplastic events in distant organs. However, detection of E5 in blood may be utilized as an additional diagnostic and/or prognostic marker and as a target for introducing immunotherapeutic procedures to reduce deaths and economic losses due to BPV activity.
In conclusion, we provide evidence that BPV-2 may persist and be maintained in a replicative status in the bloodstream, in particular in the lymphocytes, and act as a reservoir of viral infection that in the presence of biological and/or chemical co-carcinogens can be involved in bladder tumour development.
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ACKNOWLEDGEMENTS |
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REFERENCES |
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Bodaghi, S., Wood, L. V., Robby, G., Ryder, C., Steiberg, S. M. & Zeng, Z. M. (2005). Could human papillomavirus be spread through blood? J Clin Microbiol 43, 5428–5434.
Borzacchiello, G., Iovane, G., Marcante, M. L., Poggiali, F., Roperto, F., Roperto, S. & Venuti, A. (2003). Presence of bovine papillomavirus type 2 DNA and expression of the viral oncoprotein E5 in naturally occurring urinary bladder tumours in cows. J Gen Virol 84, 2921–2926.
Borzacchiello, G., Russo, V., Gentile, F., Roperto, F., Venuti, A., Nitsch, L., Campo, M. S. & Roperto, S. (2006). Bovine papillomavirus E5 oncoprotein binds to the activated form of the platelet-derived growth factor β receptor in naturally occurring bovine urinary bladder tumours. Oncogene 25, 1251–1260.[CrossRef][Medline]
Brandt, S., Haralambus, R., Schoster, A., Kirnbauer, R. & Stanek, C. (2008). Peripheral blood mononuclear cells represent a reservoir of bovine papillomavirus DNA in sarcoid-affected equines. J Gen Virol 89, 1390–1395.
Campo, M. S. (1997). Papillomavirus and cancer. Vet J 154, 175–188.[CrossRef][Medline]
Campo, M. S. (2002). Animal models of papillomavirus pathogenesis. Virus Res 89, 249–261.[CrossRef][Medline]
Campo, M. S., Jarrett, W. F. H., Barron, R. J., O'Neil, B. W. & Smith, K. T. (1992). Association of bovine papillomavirus type 2 and bracken fern with bladder cancer in cattle. Cancer Res 52, 6898–6904.
Campo, M. S., Jarrett, W. F. H., O'Neil, B. W. & Barron, R. J. (1994). Latent papillomavirus infection in cattle. Res Vet Sci 56, 151–157.[Medline]
Chambers, G., Ellsmore, V. A., O'Brien, P. M., Reid, S. W., Love, S., Campo, M. S. & Nasir, L. (2003). Association of bovine papillomavirus with the equine sarcoid. J Gen Virol 84, 1055–1062.
Cheng, Y. W., Chiou, H. L., Sheu, G. T., Hsich, L. L., Chen, J. T., Chen, C. Y., Su, J. M. & Lee, H. (2001). The association of human papillomavirus 16/18 infection with lung cancer among nonsmoking Taiwanese women. Cancer Res 61, 2799–2803.
de Villiers, E. M., Fauquet, C., Broker, T. R., Bernard, H. U. & zur Hausen, H. (2004). Classification of papillomaviruses. Virology 324, 17–27.[CrossRef][Medline]
de Villiers, E. M., Sandstrom, R. E., zur Hausen, H. & Buck, C. E. (2005). Presence of papillomavirus sequences in condylomatous lesions of the mamillae and in invasive carcinoma of the breast. Breast Cancer Res 7, R1–R11.[CrossRef][Medline]
Fernandez-Contreras, M. E., Sarria, C., Nieto, S. & Lazo, P. A. (2000). Amplification of human genomic sequences by human papillomaviruses universal consensus primers. J Virol Methods 87, 171–175.[CrossRef][Medline]
Freitas, A. C., Silva, M. A. R., Carvalho, C. C. R., Birgel, E. H., dos Santos, J. F., Beçak, W. & Stocco dos Santos, R. C. (2007). Papillomavirus DNA detection in non epithelial tissues: a discussion about bovine papillomavirus. In Communicating Current Research and Educational Topics and Trends in Applied Microbiology, vol. 2, pp 697–703, Edited by A. Méndez-Vilas. Badajoz: Formatex.
Ho, C. M., Yang, S. S., Chien, T. Y., Huang, S. H., Jeng, C. J. & Chang, S. F. (2005). Detection and quantitation of human papillomavirus type 16, 18 and 52 DNA in the peripheral blood of cervical cancer patients. Gynecol Oncol 99, 615–621.[CrossRef][Medline]
Hopkins, N. C. G. (1986). Aetiology of enzootic haematuria. Vet Rec 118, 715–717.[Abstract]
Jarrett, W. F. H., McNeil, P. E., Grimshaw, W. T. R., Selman, I. E. & McIntyre, W. I. M. (1978). High incidence area of cattle cancer with a possible interaction between an environmental carcinogen and a papillomavirus. Nature 274, 215–217.[CrossRef][Medline]
Kan, C. Y., Iacopetta, B. J., Lawson, J. S. & Whitaker, N. J. (2005). Identification of human papillomavirus DNA gene sequences in human breast cancer. Br J Cancer 93, 946–948.[CrossRef][Medline]
Lopez-Beltran, A., Sauter, G., Gasser, T., Hartmann, A., Schmitz-Dräger, B. J., Helpap, B., Ayala, A. G., Tamboli, P., Knowles, M. A. & other authors (2004). Infiltrating urothelial carcinoma. In Pathology and Genetics of Tumours of the Urinary System and Male Genital Organs. Edited by J. N. Eble, G. Sauter, J. I. Epstein & I. A. Sesterhenn. Lyon: IARC Press.
Manos, M. M., Ting, J., Wright, D. K., Lewis, A. I., Broker, T. R. & Wolinsky, S. M. (1989). Use of polymerase chain reaction amplification for the detection of genital human papillomavirus. Cancer Cells 7, 209–214.[CrossRef]
Otten, N., von Tscharner, C., Lazary, S., Antczak, D. F. & Gerber, H. (1993). DNA of bovine papillomavirus type 1 and 2 in equine sarcoids: PCR detection and direct sequencing. Arch Virol 132, 121–131.[CrossRef][Medline]
Pamukcu, A. M., Price, J. M. & Bryan, G. T. (1976). Naturally occurring and bracken fern-induced bovine urinary bladder tumors. Clinical and morphological characteristics. Vet Pathol 13, 110–122.[Medline]
Prakash, A. S., Pereira, T. N., Smith, B. L., Shaw, G. & Seawright, A. A. (1996). Mechanism of bracken fern carcinogenesis: evidence for H-ras activation via initial adenine alkylation by ptaquiloside. Nat Toxins 4, 221–227.[Medline]
Rector, A., Tachezy, R. & Van Ranst, M. (2004). A sequence-independent strategy for detection and cloning of circular DNA virus genomes by using multiply primed rolling-circle amplification. J Virol 78, 4993–4998.
Reddy, A. L. & Fialkow, P. J. (1983). Papillomas induced by initiation promotion differ from those induced by carcinogen alone. Nature 304, 69–71.[CrossRef][Medline]
Roperto, S., Ambrosio, V., Borzacchiello, G., Galati, P., Paciello, O., Russo, V. & Roperto, F. (2005). Bovine papillomavirus type-2 (PBV-2) infection and expression of uroplakin IIIb, a novel urothelial cell marker, in urinary bladder tumors of cows. Vet Pathol 42, 812–818.[CrossRef][Medline]
Roperto, S., Borzacchiello, G., Cesellato, R., Galati, P., Russo, V., Sonnino, S. & Roperto, F. (2007). Sialic acid and GM3 ganglioside expression in papillomavirus-associated urinary bladder tumours of cattle with chronic enzootic haematuria. J Comp Pathol 137, 87–93.[CrossRef][Medline]
Sauter, G., Algaba, F., Amin, M. B., Busch, C., Cheville, J., Gasser, T., Grignon, D. J., Hofständer, F., Lopez-Beltran, A. & Epstein, J. I. (2004). Non-invasive urothelial tumours. In Pathology and Genetics of Tumours of the Urinary System and Male Genital Organs. Edited by J. N. Eble, G. Sauter, J. I. Epstein & I. A. Sesterhenn. Lyon: IARC Press.
Stocco dos Santos, R. C., Lindsey, C. J., Ferraz, O. P., Pinto, J. R., Mirandola, R. S., Benesi, F. J., Birgel, E. H., Bragança Pereira, C. A. & Beçak, W. (1998). Bovine Papillomavirus transmission and chromosomal aberrations: an experimental model. J Gen Virol 79, 2127–2135.[Abstract]
Tonon, S. A., Picconi, M. A., Bos, P. D., Zinovich, J. B., Galuppo, J., Alonio, L. V. & Teyssie, A. R. (2001). Physical status of E2 human papilloma virus 16 viral gene in cervical preneoplastic and neoplastic lesions. J Clin Virol 21, 129–134.[CrossRef][Medline]
Widschwendter, A., Blassnig, A., Wiedemair, A., Müller-Holzner, E., Müller, H. M. & Marth, C. (2003). Human papillomavirus DNA in sera of cervical cancer patients as tumor marker. Cancer Lett 202, 231–239.[CrossRef][Medline]
Widschwendter, A., Brunhuber, T., Wiedemair, A., Müller-Holzner, E. & Marth, C. (2004). Detection of human papillomavirus DNA in breast cancer of patients with cervical cancer history. J Clin Virol 31, 292–297.[CrossRef][Medline]
Wosiacki, S. R., Barreiro, M. A. B., Alfieri, A. F. & Alfieri, A. A. (2005). Semi-nested PCR for detection and typing of bovine papillomavirus type 2 in urinary bladder and whole blood from cattle with enzootic haematuria. J Virol Methods 126, 215–219.[CrossRef][Medline]
Yang, H., Yang, K., Khafagi, A., Tang, Y., Carey, T. E., Opipari, A. W., Lieberman, R., Oeth, P. A., Lancaster, W. & other authors (2005). Sensitive detection of human papillomavirus in cervical, head/neck, and schistosomiasis-associated bladder malignancies. Proc Natl Acad Sci U S A 102, 7683–7688.
Received 26 May 2008;
accepted 18 July 2008.
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