Replication and interaction of herpes simplex virus and human papillomavirus in differentiating host epithelial tissue

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Virology 315 (2003) 43–55

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Replication and interaction of herpes simplex virus and human papillomavirus in differentiating host epithelial tissue Craig Meyers,1,* Samita S. Andreansky,1,2 and Richard J. Courtney Department of Microbiology and Immunology, The Pennsylvania State University College of Medicine, Hershey, PA 17033, USA Received 22 January 2003; returned to author for revision 27 February 2003; accepted 3 June 2003

Abstract We have investigated the interactions and consequences of superinfecting and coreplication of human papillomavirus (HPV) and herpes simplex virus (HSV) in human epithelial organotypic (raft) culture tissues. In HPV-positive tissues, HSV infection and replication induced significant cytopathic effects (CPE), but the tissues were able to recover and maintain a certain degree of tissue integrity and architecture. HPV31b not only maintained the episomal state of its genomic DNA but also maintained its genomic copy number even during times of extensive HSV-induced CPE. E2 transcripts encoded by HPV31b were undetectable even though HPV31b replication was maintained in HSV- infected raft tissues. Expression of HPV31b oncogenes (E6 and E7) was also repressed but to a lesser degree than was E2 expression. The extent of CPE induced by HSV is dependent on the magnitude of HPV replication and gene expression at the time of HSV infection. During active HSV infection, HPV maintains its genomic copy number even though genes required for its replication were repressed. These studies provide new insight into the complex interaction between two common human sexually transmitted viruses in an in vitro system, modeling their natural host tissue in vivo. © 2003 Elsevier Inc. All rights reserved.

Introduction Worldwide, cervical cancer is the third most prevalent female cancer. Between 400,000 and 500,000 new cases are reported each year with nearly 200,000 deaths (Bai et al., 2000; Mohar and Frias-Mendivil, 2000). Human papillomaviruses (HPV) have been associated with over 90% of all cervical cancers examined and they are considered the most important risk factor (Broker and Botchan, 1986; Pfister, 1987b; zur Hausen and Schneider, 1987). Infection with HPV is necessary but not sufficient for the development of cervical cancer. It is hypothesized that HPV infection leads to the transformation and the subsequent expression of the malignant phenotype that influences the deregulation of the

* Corresponding author. Department of Microbiology and Immunology H107, The Pennsylvania State University College of Medicine, Hershey, PA 17033. Fax: ⫹1-717-531-4600. E-mail address: [email protected] (C. Meyers). 1 These authors contributed equally to this work. 2 Present address: Department of Immunology, St. Jude Children’s Research Hospital, Memphis, Tennessee 38105. 0042-6822/03/$ – see front matter © 2003 Elsevier Inc. All rights reserved. doi:10.1016/S0042-6822(03)00466-5

proliferating cell. This is a multistep process and the long period of latency from the initial time of infection to the development of disease suggests that other determinants may be involved as possible cofactors in the pathogenesis of HPV-related genital cancers. These include smoking (Brock et al., 1989), steroid hormones (Brinton, 1991; Brinton et al., 1990; Thomas, 1991a; WHO Collaborative Study, 1993), herpes simplex virus type 2 (HSV-2) (Jones, 1995; zur Hausen, 1982), human cytomegalovirus (Jones, 1995), human herpesvirus 6 (Chen et al., 1994; Del Vecchio et al., 1992; DiPaolo et al., 1993), human immunodeficiency virus (Vernon et al., 1992, 1993, 1994a, 1994b), and chlamydia (Anttila et al., 2001; Koskela et al., 2000; Smith et al., 2002; Tamim et al., 2002). Of particular interest is the association of HSV with HPV and cervical carcinogenesis. HSV-2 is transmitted sexually (similar to HPV) and can cause recurrent, painful ulcers (Sherman et al., 1991). Since HSV and HPV are transmitted sexually and infect the same cell type, both viruses have the potential to interact with each other, impacting neoplastic progression. Additionally, epidemiological studies on the association of cervical carcinoma with HPV and HSV have

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been reported (Di Luca et al., 1987, 1989; Eglin et al., 1981; Hildesheim et al., 1991; Macnab et al., 1986; Manservigi et al., 1986; McDougall et al., 1980, 1984; Olsen et al., 1998; Prakash et al., 1985; Rotola et al., 1986; Smith et al., 2002). Initially in the pathogenesis of HSV the virus infects and replicates at the mucosal or epidermal surfaces, and HSV is subsequently transported to the ganglia where it replicates and then becomes latent. The development of the organotypic epithelial culture system has provided a potentially powerful tool for the study of infectious agents that infect the epithelium. Previously we have demonstrated the facility of this system to study the replication and pathogenesis of HPV (Bromberg-White and Meyers, 2002; Mayer and Meyers, 1998; Meyers et al., 1992b; Ozbun and Meyers, 1996, 1997, 1998a, 1999b; Sen et al., 2002; Southern et al., 2001) and HSV (Visalli et al., 1997). In this present article we report the use of the organotypic epithelial raft culture system to investigate the interactions of two important human sexually transmitted viruses, HPV and HSV. The use of organotypic raft cultures is the only in vitro system that can provide a tissue culture system that will enable the complete replication cycle of both viruses.

Results Cytopathic effects and spread of HSV-1 (KOS) and HSV-2 (186) in organotypic HPV-positive epithelial culture tissues HPV-positive CIN-612 9E organotypic (raft) culture tissues were assayed for their ability to support HSV infection. CIN-612 9E raft tissues were allowed to grow and differentiate for 3, 6, or 9 days at the air–liquid interface and then were infected by application of 103 plaque-forming units (PFU) of either HSV-1 (KOS) or HSV-2 (186) to the top of the tissue. Raft tissues grown for 3, 6, and 9 days were used to study HSV infection of tissues which had stratified and differentiated to different extents with concomitant differences in the progress of the HPV life cycle. Mock-infected tissues were also included. Tissues infected with HSV following 3 days of growth and differentiation at the air–liquid interface were harvested as described under Materials and methods for histological examinations. Hematoxylin and eosin staining was done to examine the effect of HSV infection on the HPV-positive epithelial morphology (Fig. 1). Changes in the normal CIN612 9E morphology were clearly seen 24 h post-infection with both HSV-1 (KOS) and HSV-2 (186) (Figs. 1A, F, and K). Evidence of HSV-induced cytopathic effects (CPE) is visible 24 h post-HSV infection and increases until the tissue was completely decimated 5 days post-infection (Figs. 1E, J, and O), whereas mock-infected tissue displayed a morphology representative of CIN-612 9E raft tissues (Mayer et al., 2000; Mayer and Meyers, 1998; Meyers et al., 1992; Ozbun and Meyers, 1996, 1997). Tissues examined at

time periods greater than 5 days post-HSV infection (data not shown) remained devoid of tissue, suggesting that infection with HSV-1 or HSV-2 was capable of completely destroying the HPV-positive CIN-612 9E raft culture tissues. These results are similar to what we previously observed when we infected HPV-negative raft culture tissues with HSV-1 or HSV-2 (Visalli et al., 1997). HPV-negative raft culture tissues infected with HSV-1 or HSV-2 at any time point following the lifting of the raft culture to the air–liquid interface for growth and differentiation resulted in active HSV replication and total tissue decimation within 3 days of HSV infection of the tissues (Visalli et al., 1997). Thin sections of HPV-positive CIN-612 9E raft tissues infected with HSV were immunohistochemically analyzed to determine the location and spread of HSV-1 and HSV-2 within the epithelial architecture (Fig. 2). An HSV ICP4 polyclonal antibody was used to identify cells that were undergoing the early stages of HSV replication events (Figs. 2A–J). As early as 24 h post-HSV infection, some nuclei of the basal cell layer showed an ICP4-positive staining in both HSV-1 (KOS) and HSV-2 (186) infected tissues (Figs. 2A and F). Expression of ICP4 increased with time, whereas mock-infected tissue was negative for ICP4 expression at all time points (data not shown). Similar kinetics of ICP4 expression was previously obtained when we infected primary keratinocyte raft cultures HSV (Visalli et al., 1997). To identify cells within the epithelial cross sections that were undergoing late stages of the HSV life cycle, we used a monoclonal antibody to the HSV major capsid polypeptide, VP5 (Fig. 2K–T). Similar to the expression of ICP4 in the HSV-infected tissues, VP5 expression was observed as early as 24 h post-HSV infection (Figs. 2K and P). However, VP5 expression increased to a greater level with time than what was observed for ICP4, suggesting a steady increase in the synthesis of the major capsid polypeptide of HSV-1 and HSV-2. Again, mock-infected tissues were uniformly negative for VP5 expression at all time points (data not shown). From these initial studies we concluded that infecting HPV-positive raft tissues after 3 days of growth and differentiation at the air–liquid interface led to HSV replication and spread, CPE, and complete destruction of the tissue. Our next set of experiments was then designed to determine if the stage of the HPV life cycle had an effect on HSV infection. Based on our previous studies describing HPV gene expression (Mayer and Meyers, 1998; Ozbun and Meyers, 1996, 1997, 1998a, 1999a,b), HSV-1 (KOS) or HSV-2 (186) was added to the CIN-612 9E raft cultures at two particular stages of the HPV life cycle. Three days of growth and differentiation at the air–liquid interface represents a time when HPV gene expression is at a low point but just beginning to increase in preparation for viral replication. Six days of growth and differentiation at the air–liquid interface represents a time when HPV gene expression is at a midpoint in its linear logarithmic increase and is just at the boundary associated with maximum virion synthesis. Nine

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days of growth and differentiation at the air–liquid interface represents a time when HPV gene expression has reached its plateau and maximum virion synthesis is occurring. Having previously infected HPV-positive raft tissue with HSV after 3 days of growth, when HPV gene expression is low, we decided to test the effect of infecting with HSV after 6 and 9 days of growth when HPV gene expression is significantly higher. We therefore performed the same set of experiments but infecting CIN-612 9E raft tissue with HSV-1 (KOS) and HSV-2 (186) on the sixth day of growth and differentiation at the air–liquid interface. Raft cultures were harvested 1, 2, 3, 4, and 5 days post-HSV infection and thin sections were prepared and stained with hematoxylin and eosin to examine tissue morphology. Similar to CIN-612 9E (HPV31b) raft cultures grown for 3 days and then infected with HSV, HPV- positive cultures were grown for 6 days and infected with HSV and exhibited progressive tissue destruction over the first 5 days following HSV infection (Fig. 3). Mockinfected HPV-positive CIN-612 9E raft tissues grew and differentiated with similar kinetics and morphology as previously observed for CIN-612 9E raft tissues (Figs. 3A–F) (Mayer et al., 2000; Mayer and Meyers, 1998; Meyers et al., 1992b; Ozbun and Meyers, 1996, 1997). We then asked if the HSV-infected CIN-612 9E (HPV31b) tissues could continue to grow past day 5 post-HSV infection or would the tissue be eventually destroyed as seen previously in tissues that only grew for 3 days at the air–liquid interface before being infected with HSV (compare Figs. 1 and 3). In contrast to raft tissue infected on the third day of growth at the air–liquid interface, tissues infected with HSV following 6 days of growth at the air–liquid interface recovered from the destruction elicited by the infection and replication of HSV-1 (KOS) and HSV-2 (186) (Fig. 3). To determine the location and spread of HSV-1 and HSV-2 within the raft epithelial cells, we performed immunohistochemistry analysis to visualize the expression of the HSV ICP4 and VP5 (Fig. 4). The continued presence of VP5 strongly suggests that the virus was still present in the tissues (Figs. 5O and T). To further examine the long-term effects of the HSV infections on HPV-positive tissues, CIN-612 9E tissues infected with HSV after growth for 6 days at the air–liquid interface were harvested at 8, 10, 12, 14, and 16 days post-HSV infection, sectioned, and stained with hematoxylin and eosin. The tissue displayed CPE due to the HSV infection but was never completely destroyed (Fig. 5). Pictures of mock-infected tissues 12 and 16 days p.i. are included in Figs. 5F and L for comparison. The tissue appeared to go through cycles of high levels of CPE and then would recover before another round of high CPE. ICP4 and VP5 were expressed throughout the time period investigated, suggesting that HSV replication was continuous (Fig. 6). HPV-positive CIN-612 9E raft cultures were allowed to grow and differentiate for 9 days at the air–liquid interface before infection with HSV-1 (KOS) or HSV-2 (186) gave

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results similar to those seen in cultures grown for 6 days at the air–liquid interface and then were infected with HSV (data not shown). Additionally, infecting the HPV-positive tissues with only 102 PFU of HSV-1 (KOS) or HSV-2 (186) produced similar results as using 103. PFU of HSV for infection, except for slightly slower kinetics of CPE development and the expression of ICP4 and VP5 (data not shown). Replication kinetics of HSV-1 (KOS) and HSV-2 (186) in organotypic HPV-positive epithelial culture tissues To more directly demonstrate the ability of HPV-positive organotypic tissues to support HSV replication, HSV titrations were performed. HPV-positive CIN-612 9E raft culture tissues were allowed to grow and differentiate for 6 and 9 days at the air–liquid interface and then 102 or 103 PFU of either HSV-1 (KOS) or HSV-2 (186) was applied to the top of the epithelial tissue. Tissues were harvested at 24-h periods post-HSV infection as described under Materials and methods. Virus titrations were performed using Vero cells at each time point and the data were tabulated and graphed as total PFU per infected organotypic culture (Fig. 7). HSV infections exhibited typical eclipse and logarithmic viral infection phases followed by a steady decrease in the HSV titer. After longer incubation times relatively low titers of HSV-1 (KOS) and HSV-2 (186) were maintained with fluctuations of a log or two in the total amount of virus per raft tissues that correlated with the levels of the viral proteins, ICP4 and VP5, and with the state of the tissue CPE. Titers appeared higher immediately preceding time periods when the greatest CPE was observed. State of the HPV DNA in the HSV-infected tissue Previous studies have shown a strong association of HPV DNA integration into the host genome with cervical malignancy (Jeon et al., 1995; Jeon and Lambert, 1995). These studies strongly suggested that integration of HPV DNA into the host chromosomes can be an important milestone in the progression of cervical cancer. We have hypothesized that if HSV infection occurs during a specific stage in the HPV life cycle, then HSV has the potential to act as a cofactor and influence progression to cervical cancer possibly by inducing the integration of HPV genomic DNA into the host chromosomes. To test this hypothesis, we examined the status of HPV DNA at various times post-HSV infection. Total cellular and viral DNA was isolated and analyzed by Southern blot hybridization. Differences in HPV genomic copy number between mock-infected and HSVinfected HPV-positive raft tissues were then approximated by comparisons with copy number controls (Fig. 8). HPV-positive CIN-612 9E raft cultures were allowed to grow and differentiate for 6 or 9 days and then were infected with 103 or 102 PFU of HSV-1 (KOS) or HSV-2 (186). Raft grown tissues were allowed to grow for another 2, 4, 6, and

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Fig. 1. Hematoxylin and eosin staining of herpes simplex virus (HSV) infected HPV31b-positive CIN-612 9E organotypic raft culture tissues. CIN-612 9E raft culture tissues were allowed to grow and differentiate for 3 days at the air–liquid interface and then were mock-infected (A–E), infected with HSV-1 (KOS) (F–J), or infected HSV-2 (186) (K–O). To each tissue 10 PFU of virus were applied as described under Materials and methods. Infected tissues were incubated at 37°C harvested at 24-h time periods post-HSV infection, and fixed in 10% buffered formalin. Thin sections of tissue from each time point postinfection (pi) were stained with hematoxylin and eosin. (A, F, K) 1 day (1d) pi; (B, G, L) 2 days pi; (C, H, M) 3 days pi; (D, I, N) 4 days pi; and (E, J, O) 5 days pi. Fig. 2. Immunostaining of herpes simplex virus (HSV) infected HPV31b-positive CIN-612 9E organotypic raft culture tissues. CIN-612 9E raft culture tissues were allowed to grow and differentiate for 3 days at the air–liquid interface and then were infected with HSV-1 (KOS) (A–E and K–O) or infected HSV-2 (186) (F–J and P–T). To each tissue 103 PFU of virus were applied as described under Materials and methods. Infected tissues were incubated at 37°C harvested at 24 h time periods post-HSV infection, and fixed in 10% buffered formalin. Thin sections of tissue from each time point postinfection (pi) were immunostained with an HSV-1 ICP4 monoclonal antibody (A–J) or a monoclonal antibody to the HSV major capsid polypeptide, VP5 (K–T). (A, F, K, P) 1 day (1d) pi; (B, G, L, Q) 2 days pi; (C, H, M, R) 3 days pi; (D, I, N, S) 4 days pi; and (E, J, O, T) 5 days pi. Areas staining black represent positive staining for the HSV proteins ICP4 and VP5. Arrows point to representative areas of positive staining. Fig. 3. Hematoxylin and eosin staining of herpes simplex virus (HSV) infected HPV31b-positive CIN-612 9E organotypic raft culture tissues. CIN-612 9E raft culture tissues were allowed to grow and differentiate for 6 days at the air–liquid interface and then were mock-infected (A–F), infected with HSV-1 (KOS) (G–L), or infected HSV-2 (186) (M–R). To each tissue 103 PFU of virus were applied as described under Materials and methods. Infected tissues were incubated at 37°C, harvested at 24-h time periods post-HSV infection, and fixed in 10% buffered formalin. Thin sections of tissue from each time point postinfection (pi) were stained with hematoxylin and eosin. (A, G, M) 1 day (1d) pi; (B, H, N) 2 days pi; (C, I, O) 3 days pi; (D, J, P) 4 days pi; (E, K, Q) 5 days pi; and (F, L, R) 6 days pi.

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Fig. 4. Immunostaining of herpes simplex virus (HSV) infected HPV31b-positive CIN-612 9E organotypic raft culture tissues. CIN-612 9E raft culture tissues were allowed to grow and differentiate for 6 days at the air–liquid interface and then were infected with HSV-1 (KOS) (A–E and K–O) or infected HSV-2 (186) (F–J and P–T). To each tissue 103 PFU of virus was applied as described under Materials and methods. Infected tissues were incubated at 37°C harvested at 24-h time periods post-HSV infection, and fixed in 10% buffered formalin. Thin sections of tissue from each time point postinfection (pi) were immunostained with an HSV-1 ICP4 monoclonal antibody (A–J) or a monoclonal antibody to the HSV major capsid polypeptide, VP5 (K–T). (A, F, K, P) 1 day (1d) pi; (B, G, L, Q) 2 days pi; (C, H, M, R) 3 days pi; (D, I, N, S) 4 days pi; and (E, J, O, T) 5 days pi. Areas staining black represent positive staining for the HSV proteins ICP4 and VP5. Arrows point to representative areas of positive staining. Fig. 5. Hematoxylin and eosin staining of herpes simplex virus (HSV) infected HPV31b-positive CIN-612 9E organotypic raft culture tissues grown for extended time periods post-HSV infection. CIN-612 9E raft culture tissues were allowed to grow and differentiate for 6 days at the air–liquid interface and then were infected with HSV-1 (KOS) (A–E), infected with HSV-2 (186) (G–K), or mock- infected (F, L). To each tissue 103 PFU of virus was applied as described under Materials and methods. Infected tissues were incubated at 37°C harvested at different times post-HSV infection, and fixed in 10% buffered formalin. Thin sections of tissue from each time point postinfection (pi) were stained with hematoxylin and eosin. (A, G) 8 days (8d) pi; (B, H) 10 days pi; (C, F, I) 12 days pi; (D, J) 14 days pi; and (E, K, L) 16 days pi. Fig. 6. Immunostaining of herpes simplex virus (HSV) infected HPV31b-positive CIN-612 9E organotypic raft culture tissues grown for extended time periods post-HSV infection. CIN-612 9E raft culture tissues were allowed to grow and differentiate for 6 days at the air–liquid interface and then were infected with HSV-1 (KOS) (A–B) or infected HSV-2 (186) (C–D). To each tissue 103 PFU of virus was applied as described under Materials and methods. Infected tissues were incubated at 37°C harvested at different times post-HSV infection, and fixed in 10% buffered formalin. Thin sections of tissue from each time point postinfection (pi) were immunostained with an HSV-1 ICP4 monoclonal antibody (A–B) or a monoclonal antibody to the HSV major capsid polypeptide, VP5 (C–D). (A, C) 8 days (8d) pi; and (B, D) 12 days pi. Areas staining black represent positive staining for the HSV proteins ICP4 and VP5. Arrows point to representative areas of positive staining.

8 days post-HSV infection and then were harvested for Southern blot analysis. We observed no decrease in the levels of HPV DNA following HSV infection as compared

to mock-infected cultures (Fig. 8). In general there was an increase in HPV genomic copy number in HSV-infected cultures as compared to mock-infected cultures. HPV

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Fig. 7. HSV-infected HPV31b-positive CIN-612 9E raft tissue HSV growth curves. HPV31b-positive CIN-612 9E raft tissues were infected with 102 or 103 PFU of either HSV-1 (KOS) or HSV-2 (186) as described under Materials and methods. Raft tissues were allowed to grow and differentiate at the air–liquid interface for 6 or 9 days before HSV was applied. HSV-infected tissues were incubated at 37°C and harvested at various times post HSV infection, and HSV replication was titered on Vero cells as described under Materials and methods. HSV replication was graphed as total PFU per infected organotypic raft culture.

genomic copy numbers fluctuated at different time points post-HSV infection. The effect of the HSV infection was the same whether the raft tissues were allowed to grow for 6 or 9 days at the air–liquids interface before infecting with HSV. Additionally, in these tissues we were unable to detect the presence of integrated HPV DNA which would have appeared as a band just below the position of the loading wells on the Southern blot coincident with high molecular weight cellular DNA. This data demonstrate, at least at the time points analyzed, that infection with either HSV-1 (KOS) or HSV-2 (186) did not affect the episomal maintenance of HPV genomic DNA. Therefore, if HSV does act as a cofactor for cervical cancer progression, it functions separately from the induction of HPV genomic integration or requires a longer period of time to impart this effect, suggesting an indirect mechanism. The effect of HSV infection on HPV gene expression It has been suggested that down regulation of HPV E2 expression and/or derepression of the HPV oncogenes E6 and E7 expression are important steps in the progression of cervical cancer (Jeon et al., 1995; Jeon and Lambert, 1995; Lambert and Howley, 1988; Romanczuk and Howley, 1992; Schiller et al., 1989). A potential cofactor role for HSV could therefore be an effect on the transcription of HPV genes E2, E6, or E7. To test this, we compared the expression levels of HPV31b E2, E6, and E7 in mock-infected, HSV-1-infected, and HSV-2-infected tissues.

To measure HPV31b E2, E6, and E7 gene expression in mock- and HSV-infected CIN-612 9E tissues, we employed the RNase protection assay (RPA). To quantify changes in HPV31b E2, E6, and E7 expression following HSV infection, CIN-612 9E raft cultures were grown as described for 6 days at the air-liquid interface, and then were mock-, HSV-1-, or HSV-2-infected, and RNA was harvested 2, 4, or 6 days postinfection. To analyze E2 expression, RPA was performed using a 276 nt E2- specific riboprobe, which will result in a 169-nt protected fragment representing E2 transcripts. Two days post-HSV infection we observed a decrease in HPV31b E2 transcript expression particularly in tissues infected with HSV-2 as compared to mock-infected tissues (Fig. 9). This decrease in HPV31b E2 expression was increasingly more prominent at 4 and 6 days post-HSV infection, for both HSV-1 (KOS) and HSV-2 (186) as compared to the control mock-infected tissues. A decrease in HPV31b transcripts utilizing the E1 splice donor is clearly seen at 4 (Fig. 9, 6/10) and 6 days (Fig. 9, 6/12) post-HSV infection as compared to mock- infected samples (Fig. 9). The HPV31b E1 splice donor is predominately utilized to create the E1^E4 (Ozbun and Meyers, 1999a, 1998a) spliced in-frame gene and to a lesser extent the E1*I and E1*II spliced transcripts (Ozbun and Meyers, 1999a, 1998a). This data suggest that HSV infection induces an inhibition of HPV31b transcripts. A potential mechanism for this HSV infection associated effect could be attributed to the HSV virus host shutoff (vhs) gene, which inhibits host transcription and RNA processing by destabilizing host cell

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Fig. 8. Southern blot hybridization of HPV31b genomic DNA post-HSV infection in raft culture. HPV31b-positive CIN-612 9E cells were grown in raft culture and infected with HSV-1 (KOS), HSV-2 (186), or mock-infected after 6 (6/8 – 6/14) and 9 days (9/11–9/17) of growth and differentiation at the air–liquid interface as described under Materials and methods. Raft tissues were then harvested for DNA after 2 days (2d) (6/8 and 9/11) post-HSV infection (pi); 4 days pi (6/10 and 9/13); 6 days pi (6/12 and 9/15); and 8 days pi (6/14 and 9/17). Five micrograms of total cellular DNA was digested with BamHI (odd numbered lanes), which does not cut the HPV31b genome, and with HindIII (even numbered lanes), which cuts the HPV31b genome once linearizing it. Restriction enzyme digests were electrophoresed, Southern blotted, and probed with an HPV31- specific probe. The lowest band indicates form I (FI, supercoiled) HPV31b DNA (odd numbered lanes). The upper bands indicate form II (FII, relaxed circle) HPV31b DNA and dimeric HPV31b genomes (odd numbered lanes). The middle band indicates form III (FIII, linearized) HPV31b DNA (even numbered lanes). Experiments were performed at least three times.

mRNAs (Fenwick and Walker, 1978; Hill et al., 1985; Kwong and Frenkel, 1987; Reed and Frenkel, 1983; Schek and Bachenheimer, 1985). What is surprising is that studies have shown a requirement for the HPV E2 protein in the replication (Del Vecchio et al., 1992; Frattini and Laimins, 1994; Ustav and Stenlund, 1991) and episomal maintenance (Frattini and Laimins, 1994; Hubert et al., 1999; Ustav and Stenlund, 1991) of the HPV genome. Our data suggests that in the presence of a productive HSV infection, HPV E2 gene expression is not required for the replication and episomal maintenance of HPV DNA. To analyze HPV31b E6E7 oncogene expression during coinfection with HSV-1 (KOS) and HSV-2 (186), we used a 548-nt E6E7-specific riboprobe, which will result in a 365-nt protected fragment representing the expression of E6E7 transcripts. Four and six days (Fig. 10, 6/10 and 6/12) post-HSV infection, expression of E6E7 was undetectable in the mock-infected tissues but remained detectable in HSV-infected tissues (Fig. 10). This is in contrast with the rapid repression of the HPV31b E2 gene, suggesting differential specificity in the kinetics and magnitude of HSVinduced repression of HPV gene expression.

DISCUSSION Previously we reported the facility of the organotypic tissue culture for investigations concerning the initial infection and spread of HSV in its host tissue squamous epithelium (Visalli et al., 1997). Both HSV-1 and HSV-2 followed the standard infection kinetics in organotypic raft tissues derived from epidermal keratinocytes and ectocervical keratinocytes. We have now extended these studies to investigate the interaction and consequences of superinfecting HPV-positive raft culture tissue with HSV, mimicking a potentially important in vivo situation. HPVs have been associated with over 90% of all cervical cancers examined and are considered the most important risk factor (Broker and Botchan, 1986; Pfister, 1987a; zur Hausen and Schneider, 1987). Infection with HPV is necessary but not sufficient for the development of cervical cancer. It is hypothesized that HPV leads to the transformation and the subsequent expression of the malignant phenotype that influences the deregulation of the proliferating cell. This is a multistep process and the long period of latency from the initial time of infection to the development of malignancy

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Fig. 9. Ribonuclease protection analyses quantifying HPV31b E2 and E1 splice transcripts. HPV31b-positive CIN-612 9E raft cultures were allowed to grow and differentiate for 6 days at the air–liquid interface and then were mock-infected, or infected with 103 PFU of HSV-1 (KOS), or HSV-2 (186). Raft tissues were then harvested for RNA after 2 days (2d) (6/8); 4 days pi (6/10); and 6 days pi (6/12) post-HSV infection. Ten micrograms of DNase I treated total RNA or yeast RNA (Y) were analyzed. Location and size of RNA Century Markers (Ambion) are shown to the left in nucleotides (100 and 200). Experiments were performed at least three times.

for the development of cervical carcinoma compared to women infected with only one virus (Hildesheim et al., 1991). Early studies showed a prevalence of higher titers of HSV-2-specific antibody in women who developed cervical cancer (Rawls et al., 1976). To assess possible interactions between HSV-2 and high-risk HPV in the pathology of cervical carcinoma, cervical tumors were examined for the presence of both viral DNAs (Di Luca et al., 1987, 1989). It was reported that six of the eight cervical cancer biopsy specimens that contained HSV-2 sequences also contained HPV DNA. The influence on HPV gene expression by HSV has also been studied. Using heterologous systems, HSV-1 immediate-early transactivators such as ICP4 and ICP0 enhance transcription of the upstream regulatory regions of HPV16 (McCusker and Bacchetti, 1988). Furthermore, HSV-1 transactivators such as VP16 and ICP0 activated HPV18 expression in a variety of cells (Gius and Laimins, 1989). Infection with HSV-1 resulted in the amplification of bovine papillomavirus type 1 (BPV-1) sequences (Schmitt et al., 1989). Similar results were obtained by Hara et al. (1997) when they used HeLa cells and A431 cells where HPV DNA was transiently episomal following introduction by transfection. Infection of HeLa cells with HSV-1 strongly modified HPV gene expression by reducing the amounts of specific mRNAs (Karlen et al., 1993). This was attributed to

suggests that other determinants may be involved as possible cofactors in the pathogenesis of HPV-related genital cancers. These include smoking (Brock et al., 1989), steroid hormones (Brinton, 1991; Brinton et al., 1990; Thomas, 1991; WHO Collaborative Study, 1993), herpes simplex virus type 2 (HSV-2) (Jones, 1995; zur Hausen, 1982), human cytomegalovirus (Jones, 1995), human herpesvirus 6 (Chen et al., 1994; Del Vecchio et al., 1992; DiPaolo et al., 1993), human immunodeficiency virus (Vernon et al., 1992, 1993, 1994a, 1994b), and chlamydia (Anttila et al., 2001; Koskela et al., 2000; Smith et al., 2002; Tamim et al., 2002). Of particular interest with regard to our present study is the association of HSV-2 with HPV. HSV-2 infection is transmitted sexually (similar to HPV) and can cause recurrent, painful genital ulcers. A recent study of HSV-2 infections in the United States concluded that since late 1970s the prevalence of HSV-2 infections has increased 30 –32%, and HSV-2 is detectable in one of five persons 12 years of age or older nationwide (Brugha et al., 1997). Since HSV and HPV are transmitted sexually and infect the same cell type, both viruses have the potential to interact with each other and play a role in neoplastic progression. To date, there have been no studies in which the interaction of HPV and HSV has been studied in which both viruses could accomplish a complete replication cycle. Epidemiological studies on the association of cervical carcinoma with HPV and HSV-2 have been reported. Women infected with HSV-2 and HPV are at a greater risk

Fig. 10. Ribonuclease protection analyses quantifying HPV31b E6 and E7 transcripts. HPV31b-positive CIN-612 9E raft cultures were allowed to grow and differentiate for 6 days at the air–liquid interface and then mock-infected, or infected with 103 PFU of HSV-1 (KOS), or HSV-2 (186). Raft tissues were then harvested for RNA after 2 days (2d) (6/8); 4 days pi (6/10); and 6 days pi (6/12) post-HSV infection. CIN-612 9E monolayer RNA was also harvested for comparison. Ten micrograms of DNase I treated total RNA or yeast RNA (Y) were analyzed. Location and size of RNA Century Markers (Ambion) are shown to the left in nucleotides (200, 300, and 400). Experiments were performed at least three times.

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a virion component, notably the virion host shutoff protein, vhs. Our present studies were designed to define the interaction of these two sexually transmitted viruses (HPV and HSV) using epithelial organotypic raft culture model systems. Experiments were designed to determine the level of CPE induced by HSV infection, HSV’s ability to replicate and spread, and HSV’s effect on HPV replication at specific stages in the HPV life cycle. In vivo, HPV and HSV appear to infect the same genital tissues, suggesting the possibility that HSV acts as a cofactor in the multistep process of HPV-associated cervical cancer. Independent of the length of time the HPV-positive CIN-612 9E raft cultures were allowed to grow and differentiate at the air–liquid interface, HSV-1 (KOS) and HSV-2 (186) were able to infect the raft tissue, induce significant CPE, and replicate. Evidence for HSV replication was demonstrated by the expression of the immediate-early HSV protein ICP4 and the HSV late capsid protein VP5. Additionally, direct evidence was provided by titering HSV directly from the infected raft tissues. Whereas the magnitude and kinetics of HPV31b replication and gene expression did not significantly affect the ability of HSV to replicate, it did affect the ability of the tissue to recover from the HSV-induced CPE. HPV-positive raft tissues were infected with HSV-1 (KOS) or HSV-2 (186) after 3, 6, or 9 days of growth and differentiation at the air–liquid interface. HSV infection of HPV- positive raft tissue after 3 days at the air–liquid interface resulted in significant and complete CPE, decimating the tissue by 5 days post-HSV infection. HPV-positive raft tissues that were infected with HSV after 6 or 9 days at the air–liquid interface also resulted in significant CPE, but these tissues were able to recover and maintain a certain degree of tissue integrity and architecture. We examined the raft culture tissue for up to 16 days post-HSV infection and the tissue was still able to maintain a certain degree of integrity and architecture. The tissue went through a period of significant CPE, giving the appearance that it would succumb, but would always recover usually within another 24 h. Expression of HSV ICP4 and VP5 protein expression accompanied these cycles of extreme CPE and recovery. It is interesting that the ability of the tissue to “survive” HSV infection and CPE correlates with the increased magnitude of HPV31b replication and gene expression occurring at days 6 and 9 during growth and differentiation. It appears that HPV is able to elicit a survival mechanism maintaining vestiges of the host tissues architecture for its own replication. We directly examined the effects of HSV infection on the HPV31b life cycle by assaying the episomal maintenance and copy number of the HPV31b genome and the expression of the HPV31b E2 and E6E7 transcripts. Interestingly, HPV31b not only maintained the episomal state of it genomic DNA but also maintained its genomic copy number even during times of extensive HSV-induced CPE. HSV has been hypothesized as a potential cofactor for HPV- associated cervical carcinogenesis. One of the mile-

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stones associated with the development of the carcinogenic event is the integration of the HPV genome into the host chromosomes. We found no evidence for the induction of HPV genomic DNA to integrate into the host chromosomes. This does not rule out the possibility that HSV can still induce HPV DNA integration following greater time periods than what we have examined or that other host or external factors are also required. Additionally, it is interesting that HPV appears to maintain its ability to replicate normally. The replication of the related small DNA virus, SV40, following HSV superinfection adopts a large complex branched structure similar to the replication of the HSV DNA (Blumel et al., 2000). Studies have suggested that down regulation of HPV E1 and E2 expression and/or derepression of the HPV oncogenes E6 and E7 expression are important steps in the progression of cervical cancer (Jeon et al., 1995; Jeon and Lambert, 1995; Lambert and Howley, 1988; Romanczuk and Howley, 1992; Schiller et al., 1989). A potential cofactor role for HSV could therefore be an effect on the transcription of HPV genes E1, E2, E6, or E7. To test this we compared the expression levels of HPV31b E2, E6, and E7 in mock-infected, HSV-1-infected, and HSV-2-infected tissues. When the steady-state levels of the HPV31b E2 transcript were analyzed, we observed a complete loss of expression after 6 days post-HSV infection. The requirement for the HPV E2 gene product coupled with the E1 gene product has been shown for HPV DNA replication to occur (Del Vecchio et al., 1992; Ustav and Stenlund, 1991). This result was surprising in light of the previous result where HPV31b maintained the copy number of genomic DNA. Because the raft tissue is constantly replenishing itself, especially with HSV-induced CPE, HPV must continuously replicate its genome. This would suggest that HPV31b is able to replicate its DNA in HSV-infected raft tissues even when E2 gene expression is repressed. When we analyzed HPV31b oncogene expression, we observed that while expression was repressed, RNAs for E6E7 were still expressed even after 6 days post-HSV infection. Our studies have found no direct evidence for a role of HSV as a cofactor in HPV-associated cervical carcinogenesis. Although we should point out that these studies only examined the effect of HSV infection during one portion of the natural history of HPV infections, the replicative cycle. Our studies do however demonstrate an important dynamic relationship between two sexually transmitted viruses that commonly coinfect the human genital track. An increased understanding of the interaction between HPV and HSV provides an important addition to our knowledge of how sexually transmitted agents interact and provides a system whereby therapeutic interventions can be tested. Additionally, knowledge gained from studies to understand the interaction between HPV and HSV may help us focus on the portion of the natural history of HPV infections where HSV can act as a cofactor. In summary, to our knowledge we have developed the

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first system to study the interaction between HPV and HSV, two important human pathogens in a system where both viruses replicate. HSV-1 (KOS) and HSV-2 (186) are both able to replicate to high titers in HPV-positive host tissue. The extent of the CPE induced by HSV is dependent on the magnitude and kinetics of HPV replication and gene expression at the time of HSV infection. When HPV replication and gene expression are at high levels, then HSV does not destroy the tissue but the tissue instead goes through rounds of high CPE and recovery. During active HSV infection HPV is able to maintain its replication even when expression of HPV gene products required for its replication are repressed, suggesting that HSV may provide the necessary functions for HPV replication.

ing HSV to the top of the organotypic tissue four times at 15-min intervals for 1 h. Tissues were harvested at various times and frozen at ⫺80°C. Infected organotypic tissues were processed for virus titrations by adding 1 ml of medium per culture and sonicating the tissue for 30 s in a water-filled cup horn sonicator accessory to allow for the sterile disruption of the differentiated cell layers away from the collagen matrix. Tissue sonicates were frozen and thawed three times in a dry ice/ethanol bath for 15 min each. The 30-s sonication was repeated and the fractured organotypic tissues were spun at low speed to pellet the remaining collagen/cell debris. Aliquots of each organotypic tissue time point were titered on Vero cells and graphed as total PFU per infected organotypic culture.

Materials and methods

Immunohistochemistry

Cell and organotypic cultures

Stratified and differentiated organotypic culture tissues derived from HPV31b-positive CIN-612 9E cells were infected with either HSV-1 (KOS) or HSV-2 (186). Tissues were infected with 102 or 103 PFU of HSV as described above. Infected organotypic tissues were harvested at various times, fixed in 10% buffered formalin, and paraffinembedded. Sections of tissue from various time points were stained with hematoxylin and eosin or immunostained with an HSV-1 ICP4 polyclonal antibody (Yao and Courtney, 1989) or a monoclonal antibody to the HSV major capsid polypeptide, VP5 (Advanced Biotechnologies, Columbia, MD) using the VECTASTAIN Elite ABC Kit (Vector Laboratories, Burlingame, CA) according to the manufacturer’s instructions.

The HPV31b-positive cell line, CIN-612, was established from a cervical intraepithelial neoplasia (CIN) type I biopsy (Bedell et al., 1991). The CIN-612 9E clonal derivative maintains approximately 50 episomal copies per cell of the HPV31b genome (Bedell et al., 1991) and is capable of reproducing the complete HPV life cycle in culture (Meyers et al., 1992a). CIN-612 9E cells were maintained in monolayer culture with E medium containing 5% fetal bovine serum (FBS) in the presence of mitomycin C treated J2 3T3 feeder cells (McCance et al., 1988; Meyers, 1996). Organotypic (raft) CIN-612 9E tissue cultures were grown as previously described (Meyers, 1996; Meyers et al., 1992a; Meyers and Laimins, 1994). Briefly, CIN-612 9E cells were seeded onto collagen matrices containing J2 3T3 fibroblasts. After the CIN-612 9E cells grew to confluence, collagen matrices were lifted onto stainless steel grids and the cells were fed by diffusion from under the matrix. Once lifted to the air–liquid interface, organotypic cultures were fed with E medium plus 5% FBS. The CIN-612 9E cells were allowed to stratify and differentiate at the air–liquid interface. Infection and titering HSV infections and titering were done as previously described (Visalli et al., 1997). Briefly, differentiating organotypic culture tissues were infected with HSV-1 (KOS strain) or HSV-2 (186 strain) at different time points after lifting the organotypic cultures to the air–liquid interface. Medium was removed from each plate and approximately 102 or 103 PFU in a volume of 0.5 ml was applied and reapplied to the top of the organotypic tissues four times at 15-min intervals for 1 h. Infected tissues were allowed to continue to grow for varying lengths of time using E medium plus 5% FBS as before and incubated at 37°C. Mockinfection was done by applying and reapplying media lack-

Nucleic acid extraction HPV31b viral replication in organotypic raft cultures was measured by Southern blot analysis. Total cellular DNA in raft tissue was extracted as previously described (Ozbun and Meyers, 1998a). Total RNA was extracted from raft tissues by using TRIzol reagent according to manufacturer’s directions (Gibco-BRL, Bethesda, MD). To remove copurifying viral and cellular DNA, the RNA samples were treated with DNase I (Ozbun and Meyers, 1996, 1997). Southern blotting Five micrograms of total cellular DNA was digested with XbaI, which linearizes the HPV31b genome at nucleotide 4998, and separated by 0.8% agarose electrophoresis followed by transferring to GeneScreen Plus membranes (New England Nuclear Research Products, Boston, MA), as previously reported (Ozbun and Meyers, 1996, 1997). A 32Plabeled total HPV31 genomic DNA probe was generated by random primer extension and used to probe the membranes as described (Ozbun and Meyers, 1996, 1997).

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RNase protection assays (RPAs) Antisense RNA probes were synthesized as previously described (Ozbun and Meyers, 1996, 1997, 1998a, 1998b, 1999b). The E7,E1*I,E2 probe was derived from HPV31b cDNA clone pCR31b-E7 and is specific to 169 nt of E7,E1*I,E2 (HPV31 nt 747– 877^2646 –2683) and 131 nt of E7, E1 (HPV31 nt 747– 877). Probes were purified and hybridizations were performed as previously reported (Ozbun and Meyers, 1996, 1997). An antisense E6E7 probe was synthesized using the pAccI construct (a kind gift from Laimonis Laimins, Northwestern University College of Medicine). The 548-bp AccI fragment of HPV31b, corresponding to nt 200 –747, was amplified by PCR and subcloned into pBluescript (Hummel et al., 1992). The antisense RNA probe was created by AflIII digestion and by using the T3 promoter. This RNA probe yielded a 365-nt probe predicted to protect 325 nt of the HPV31b E6E7 mRNA. Samples were analyzed by electrophoresis through 5% polyacrylamide–7 M urea gels followed by autoradiography. RNA Century standards were prepared as per manufacturer’s recommendations (Ambion Inc., Austin, TX). Acknowledgments This work was supported by National Cancer Institute Grants RO1 CA79006 (C.M.) and RO1 CA42460 (R.J.C) and by the American Cancer Society Grant PF-98-16201-MB (S.S.A.). Funding was also received from the PA Department of Health, Health Research Formula Funding Program as a part of the PA Tobacco Settlement Legislation, and a Penn State Cancer Research Institute Grant of the Penn State College of Medicine. References Anttila, T., Saikku, P., Koskela, P., Bloigu, A., Dillner, J., Ikaheimo, I., Jellum, E., Lehtinen, M., Lenner, P., Hakulinen, T., Narvanen, A., Pukkala, E., Thoresen, S., Youngman, L., Paavonen, J., 2001. Serotypes of Chlamydia trachomatis and risk for development of cervical squamous cell carcinoma. JAMA 285 (1), 47–51. Bai, W., Oliveros-Saunders, B., Wang, Q., Acevedo-Duncan, M.E., Nicosia, S.V., 2000. Estrogen stimulation of ovarian surface epithelial cell proliferation. In vitro Cell Dev. Biol. Anim. 36 (10), 657– 666. Bedell, M., Hudson, J., Golub, T., Turyk, M., Hosken, M., Wilbanks, G., Laimins, L., 1991. Amplification of human papillomavirus genomes in vitro is dependent on epithelial differentiation. J. Virol. 65 (5), 2254 – 2260. Bejcek, B., Conley, A.J., 1986. A transforming plasmid from HSV-2 transformed cells contains rat DNA homologous to the HSV-1 and HSV-2 genomes. Virology 154 (1), 41–55. Blumel, J., Graper, S., Matz, B., 2000. Structure of simian virus 40 DNA replicated by herpes simplex virus type 1. Virology 276 (2), 445– 454. Brinton, L., 1991. Oral contraceptives and cervical neoplasia. Contraception 43 (6), 581–595. Brinton, L., Reeves, W., Brenes, M., Herrero, R., de Brillion, R., Gaitan, E., Tenorio, F., Garcia, M., Rawls, W., 1990. Oral contraceptive use and risk of invasive cervical cancer. Int. J. Epidemiol. 19 (1), 4 –11.

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