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Journal of Virology logoLink to Journal of Virology
. 2012 Jun;86(12):6668–6676. doi: 10.1128/JVI.06240-11

Cdk1 Inhibition Induces Mutually Inhibitory Apoptosis and Reactivation of Kaposi's Sarcoma-Associated Herpesvirus

Xudong Li 1,*, Shijia Chen 1,*, Ren Sun 1,✉
PMCID: PMC3393555  PMID: 22496227

Abstract

Primary effusion lymphoma (PEL) cells are predominantly infected by the latent form of Kaposi's sarcoma-associated herpesvirus (KSHV), with virus reactivation occurring in a small percentage of cells. Latency enables KSHV to persist in the host cell and promotes tumorigenesis through viral gene expression, thus presenting a major barrier to the elimination of KSHV and the treatment of PEL. Therefore, it is important to identify cellular genes that are essential for PEL cell survival or the maintenance of KSHV latency. Here we report that cyclin-dependent kinase 1 (Cdk1) inhibition can induce both apoptosis and KSHV reactivation in a population of PEL cells. Caspases, but not p53, are required for PEL cell apoptosis induced by Cdk1 inhibition. p38 kinase is activated by Cdk1 inhibition and mediates KSHV reactivation. Interestingly, upon Cdk1 inhibition, KSHV is reactivated predominantly in the nonapoptotic subpopulation of PEL cells. We provide evidence that this is due to mutual inhibition between apoptosis and KSHV reactivation. In addition, we found that KSHV reactivation activates protein kinase B (AKT/PKB), which promotes cell survival and facilitates KSHV reactivation. Our study thus establishes a key role for Cdk1 in PEL cell survival and the maintenance of KSHV latency and reveals a multifaceted relationship between KSHV reactivation and PEL cell apoptosis.

INTRODUCTION

Kaposi's sarcoma-associated herpesvirus (KSHV) is the etiological agent of three types of human tumors: Kaposi's sarcoma (KS), primary effusion lymphoma (PEL), and a plasmablastic type of multicentric Castleman disease (MCD) (10, 11, 16). PEL, also known as body cavity-based lymphoma (BCBL), is a deadly type of non-Hodgkin lymphoma with a short (<6 months) median survival time upon diagnosis due to a lack of sufficient potency and specificity of the current chemotherapy-based treatment regimens (2, 5, 50).

KSHV has two phases, latency and lytic replication, in its life cycle. PEL cells are predominantly infected with a latent form of KSHV (10), expressing only a few genes (9, 14, 37, 44, 45, 47, 52). Latency enables KSHV to persist in the host and constitutes a major obstacle to the elimination of KSHV from the host. In addition, latent KSHV does not passively exist in the host cell. Instead, the majority of KSHV latent proteins, including LANA, v-cyclin, vFLIP, and vIRF-3, can regulate cellular oncogenic pathways to promote cell proliferation and/or cell survival (3, 8, 12, 15, 19–22, 30, 35, 36, 39, 43, 49, 57). Some of these latent proteins and their cellular targets have been shown to be indispensable for PEL cell survival (13, 23, 26). Therefore, oncogenic pathways activated by KSHV latent proteins may be attractive targets for the treatment of PEL.

Latent KSHV can be reactivated to undergo lytic replication. The KSHV replication and transcription activator (RTA) is the key viral regulator of KSHV reactivation (38, 53). Cellular factors can regulate KSHV reactivation through modulating the level and/or activity of RTA (18, 32, 59–62). Thus, cellular factors essential for maintaining KSHV latency present another group of potential targets for disrupting latency and killing KSHV-associated tumor cells (1, 27, 58).

The proto-oncogene Myc is deregulated by two KSHV latent proteins, LANA and vIRF-3/LANA2 (8, 35, 36). LANA stabilizes and activates Myc (8, 35), whereas vIRF-3 stimulates Myc transcriptional activity (36), suggesting that Myc may play an important role in the manipulation of host cells by KSHV. Indeed, we recently showed that Myc knockdown in PEL cells not only results in cell cycle arrest and apoptosis but also disrupts latency, leading to KSHV reactivation (32). These results suggest that Myc-related cellular pathways may be potential targets for treating PEL. Although it is difficult to design drugs to directly target transcription factors such as Myc, elevated Myc expression may specifically sensitize cancer cells to p53-independent apoptosis induced by cyclin-dependent kinase 1 (Cdk1) inhibition (24), raising the possibility that PEL cells are also sensitive to Cdk1 inhibition.

We reasoned that in order for latent KSHV to avoid elimination when a host cell is about to undergo apoptosis, it has to be able to sense signals downstream of proapoptotic stimuli to induce reactivation. We therefore hypothesized that a cellular gene which is required for PEL cell survival may also be essential for the maintenance of KSHV latency. We recently showed that Myc is one such cellular gene (32). We undertook the current study to determine whether Cdk1 is also essential for both PEL cell survival and KSHV reactivation. More importantly, we explored the relationship between apoptosis and KSHV reactivation downstream of proapoptotic stimuli.

MATERIALS AND METHODS

Cell lines and reagents.

BC-3 and BCBL-1 cells were kindly provided by E. Cesarman (Cornell Medical College, NY). The construction of BC-3-G cells was previously described (62). Purvalanol A, Nutlin-3, and Z-VAD(OMe)-FMK were purchased from Calbiochem or Enzo Life Sciences. 12-O-tetradecanoylphorbol-13-acetate (TPA) was purchased from Sigma-Aldrich. The p53 (sc-6243) antibody was purchased from Santa Cruz Biotechnology. The Cdk1 (06-923) antibody was purchased from Millipore. The poly(ADP-ribose) polymerase (PARP), phospho-p38 mitogen-activated protein kinase (MAPK) (Thr180/Tyr182), phospho-Erk1/2 (Thr202/Tyr204), phospho-protein kinase B (AKT/PKB) (Ser473), and cleaved caspase 3 (catalog number 9661) antibodies were purchased from Cell Signaling Technology. The generation of anti-RTA rabbit serum was described previously (51).

shRNA vector-mediated knockdown of genes.

Short hairpin RNA (shRNA) plasmids were constructed by annealing 2 DNA oligonucleotides containing target sequences, followed by cloning the annealed DNA into the pLKO.1-TRC vector (Addgene). The target sequences are GGG GAT TCA GAA ATT GAT C (shCDK1) (34), GAC TCC AGT GGT AAT CTA C (shp53) (7), TGG CGC AAG ATG ACA AGG G (shKRTA) (32), GAT GAG GAA GAA ATC GAT G (shMyc) (32), and CAA CAA GAT GAA GAG CAC CAA (shCtrl) (32, 33). shRNA lentiviruses were produced according to the protocol provided by the manufacturer (Addgene). Briefly, shRNA vector was cotransfected with pCMV-dR8.2 dvpr (Addgene) and pCMV-VSV-G (Addgene) into 293T cells using the calcium phosphate transfection method. Lentiviruses were collected at 48 and 72 h posttransfection, filtered through 0.45-μm filters, and concentrated by centrifugation at 15,000 rpm using a Sorvall SA-600 rotor for 4 h at 2°C. Supernatant was removed, and virus pellets were resuspended with phosphate-buffered saline (PBS). To transduce cells, viruses were added to cells seeded in medium containing 4 μg/ml Polybrene. Lentiviral transduction was performed in the presence of 4 μg/ml Polybrene. When necessary, cells were subjected to a 3-μg/ml puromycin selection at 24 h posttransduction.

Cell viability assay.

Cells were seeded at a density of 5 × 104 or 1 × 105 cells per well of 48-well plates. Cells were then treated with chemicals or transduced with shRNA vectors. At different time points posttreatment or posttransduction, the numbers of live and dead cells were measured with the trypan blue (Sigma-Aldrich) exclusion assay using a hemocytometer. Cell viability was calculated as the percentage of live cells in the total cell population (live plus dead cells).

Quantitative reverse transcription-PCR (qRT-PCR) analysis.

Cells treated with chemicals were collected at 24 h posttreatment by centrifugation at 200 × g for 3 min. Cells were washed with PBS twice, followed by centrifugation. RNA was purified using a kit provided by Invitrogen. First-strand cDNA was synthesized using Superscript III polymerase (Invitrogen). Sybr green PCR was performed using probes specific for KSHV RTA and the human β-actin gene.

Apoptosis assays.

Apoptosis was measured with the annexin V-phycoerythrin (PE) apoptosis detection kit I (BD Biosciences Pharmingen) according to the manufacturer's protocol and analyzed by flow cytometry. Acquisition was performed on a FACScan flow cytometer (BD Biosciences).

Flow cytometry analysis of KSHV reactivation.

BC-3-G cells were treated with chemicals or transduced with shRNA vectors. At different time points posttreatment or posttransduction, cells were analyzed by flow cytometry for enhanced green fluorescent protein (EGFP) expression on a FACScan flow cytometer (BD Biosciences) or a FACSCanto II cytometer (BD Biosciences). In experiments that required simultaneous monitoring of apoptosis and KSHV reactivation, these readouts were simultaneously determined with annexin V-PE apoptosis detection kit I (BD Biosciences Pharmingen) for apoptosis and by EGFP expression for KSHV reactivation.

Quantitation of virion production.

Supernatants from cells were collected and cleared by centrifugation at 200 × g for 3 min, followed by a second centrifugation at 3,000 × g for 5 min. Cleared supernatants were then treated with DNase I (Invitrogen) at a concentration of 100 U/ml for 1 h to remove DNA not protected by virions. After DNase I was heat inactivated at 65°C for 30 min in the presence of 10 mM EDTA, supernatants were treated with proteinase K (Sigma-Aldrich) at 65°C for 2 h. Virion DNA was extracted with phenol-chloroform and precipitated with ethanol. Air-dried DNA was dissolved in 40 μl of Tris-EDTA (TE) buffer and measured with quantitative PCR (qPCR) using primers specific for the KSHV major capsid gene.

RESULTS

Cdk1 inhibition leads to PEL cell apoptosis.

Elevated Myc expression has been shown to increase cancer cell apoptosis induced by Cdk1 inhibition (24). Myc is deregulated in PEL cells (8, 35, 36). To evaluate whether Cdk1 is required for PEL cell survival, we treated two PEL cell lines, BC-3 and BCBL-1, or KSHV-negative DG75 cells with a 10 μM concentration of a specific Cdk1 inhibitor, purvalanol A (24, 25), or an equal concentration of its vehicle, dimethyl sulfoxide (DMSO). A trypan blue exclusion assay was used to determine cell viability at 3 days and 6 days posttreatment (Fig. 1A). Purvalanol A treatment reduced the viability of the BC-3 and BCBL-1 cells without reducing DG75 cell viability (Fig. 1A). To examine whether Myc plays a role in sensitizing PEL cells to Cdk1 inhibition, we first transduced BC-3 cells with a short hairpin RNA (shRNA) lentiviral vector targeting Myc, shMyc (32), or a control shRNA vector that does not target any human gene, shCtrl (32, 33). Cells were cultured for 48 h to allow sufficient time for shRNA vectors to function before being reseeded at an equal concentration and treated with 10 μM purvalanol A or DMSO as a vehicle control. At 48 h posttreatment, cell viability was measured with a trypan blue assay as relative numbers of viable cells compared to the control treatment (set as 100%) (Fig. 1B). Myc knockdown led to reduced numbers of viable cells, consistent with our previous results showing that Myc knockdown in PEL cells results in inhibition of both cell proliferation and apoptosis (32). Interestingly, reduction of cell viability by purvalanol A treatment was partially reversed in shMyc-transduced cells (Fig. 1B). We concluded that PEL cell sensitivity to Cdk1 inhibition is at least partially due to Myc expression.

Fig 1.

Fig 1

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Cdk1 inhibition leads to PEL cell apoptosis. (A) BC-3, BCBL-1, or DG75 cells were treated with 10 μM purvalanol A or DMSO as a vehicle control. The percentages of viable cells were determined with a trypan blue exclusion assay at day 3 and day 6 posttreatment. (B) Equal numbers of BC-3 cells transduced with the shCtrl or the shMyc lentiviral vector were seeded at 48 h posttransduction and treated with 10 μM purvalanol A or DMSO as a vehicle control. At 48 h posttreatment, cell viability was determined with a trypan blue exclusion assay as relative numbers of viable cells compared to the shCtrl and DMSO control. (C) BC-3, BCBL-1, or DG75 cells were treated with 10 μM purvalanol A or DMSO as a vehicle control for 24 h. Whole-cell lysates were analyzed with Western blotting for the expression of poly(ADP-ribose) polymerase (PARP) and with actin as a loading control. (D) BC-3 cells were pretreated with 100 μM Z-VAD or DMSO as a vehicle control for 1 h. Cells were then treated with 10 μM purvalanol A or DMSO in the presence of Z-VAD or DMSO. At 48 h posttreatment, cells were analyzed by flow cytometry for apoptosis. (E) BC-3 cells were transduced with shCtrl or shp53. At 72 h posttransduction, whole-cell lysates were analyzed with Western blotting for p53 expression. (F) Equal numbers of BC-3 cells transduced with shCtrl or shp53 were treated with 5 μM Nutlin-3, 5 μM purvalanol A, or DMSO as a vehicle control. At 48 h posttreatment, cell viability was determined by a trypan blue exclusion assay as relative numbers of viable cells compared to the DMSO control treatment. (G) BC-3 cells were transduced with shCtrl, shMyc, or shCDK1. At 120 h postransduction, whole-cell lysates were analyzed by Western blotting for cleaved caspase 3, PARP, and p53 expression. (H) BC-3 cells mock-transduced or transduced with shCtrl, shMyc, or shCDK1 were analyzed for apoptosis (DNA content of <2 N) with flow cytometry at 72 h posttransduction. (A, B, and F) Data are the means and standard deviations (SDs) from three independent experiments. (C, D, E, G, and H) Data are representative of three (C, E, and G) or two (D and H) independent experiments.

To determine whether Cdk1 inhibition induces PEL cell apoptosis, we first examined a commonly used marker for both caspase activation and apoptosis, the cleavage of poly(ADP-ribose) polymerase (PARP) by caspases. PARP cleavage was increased in BC-3 and BCBL-1 cells but not in DG75 cells treated with 10 μM purvalanol A (Fig. 1C), indicating that Cdk1 inhibition led to increased caspase activity in PEL cells. We also used flow cytometry to analyze apoptosis of BC-3 cells treated with purvalanol A or DMSO after they were pretreated with the pan-caspase inhibitor Z-VAD or DMSO as a vehicle control. In the absence of Z-VAD pretreatment, purvalanol A treatment increased the percentages of early apoptotic (annexin-positive/7-aminoactinomycin D (7-AAD)-negative) cells from 1.8% to 26% and late apoptotic/necrotic (annexin-positive/7-AAD-positive) cells from 3% to 25% at 48 h posttreatment (Fig. 1D). Z-VAD pretreatment reduced the percentages of early apoptotic cells from 26% to 4.8% and late apoptotic cells from 25% to 8.8% (Fig. 1D), indicating that caspase activation is required for PEL cell apoptosis induced by Cdk1 inhibition.

A previous study showed that apoptosis induced by Cdk1 inhibition in Myc-overexpressing cells is p53 independent (24). To determine whether PEL cell apoptosis induced by Cdk1 inhibition requires p53, we examined the effect of p53 knockdown (Fig. 1E) on the viability of BC-3 cells treated with either purvalanol A or Nutlin-3 (Fig. 1F), a murine double minute 2 (Mdm2) protein antagonist that has been shown to induce PEL cell apoptosis through p53 activation (46). Although p53 knockdown increased the viability of cells treated with Nutlin-3, it was not able to increase the viability of cells treated with purvalanol A (Fig. 1F), suggesting that PEL cell apoptosis induced by Cdk1 inhibition is p53 independent.

To further confirm that Cdk1 is required for PEL cell survival, we also examined the effect of Cdk1 knockdown on caspase activation and apoptosis. Transduction of BC-3 cells with shCDK1, an shRNA lentiviral vector targeting Cdk1 (34) (Fig. 2D), or with shMyc was able to increase the levels of both cleaved caspase 3 and cleaved PARP (Fig. 1G), suggesting that Cdk1 knockdown increases caspase activities in PEL cells. In addition, both shMyc and shCDK1 were able to increase the percentages of apoptotic cells as determined by flow cytometry (Fig. 1H), suggesting that similar to Cdk1 inhibition, Cdk1 knockdown also results in PEL cell apoptosis.

Fig 2.

Fig 2

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Cdk1 inhibition induces KSHV reactivation. (A) BC-3-G cells were treated with 10 μM purvalanol A or DMSO as a vehicle control. At 72 h posttreatment, cells were imaged with fluorescence microscopy to examine enhanced green fluorescent protein (EGFP) expression. Magnification, ×200. (B) BC-3-G cells were treated with 10 μM purvalanol A or DMSO as a vehicle control. At 48 h posttreatment, cells were analyzed with flow cytometry to determine the percentages of EGFP-positive cells. (C) BC-3-G cells were treated with 2.5 μM RO-3306 or DMSO as a vehicle control. At 48 h posttreatment, cells were analyzed with flow cytometry to determine the percentages of EGFP-positive cells. (D) BC-3 cells were transduced with shCtrl or shCDK1. At 96 h posttransduction, whole-cell lysates were analyzed with Western blotting for Cdk1 expression. Actin was analyzed as a loading control. (E) BC-3 or BCBL-1 cells were transduced with shCtrl or shCDK1, cultured for 24 h, and selected with 3 μg/ml of puromycin for 48 h. Whole-cell lysates were analyzed with Western blotting for the expression of KSHV lytic protein K8. Actin was analyzed as a loading control. (F) BC-3-G cells were cotransfected with shCDK1 or shCtrl and a red fluorescent protein (RFP) expression vector. At 72 h posttransfection, cells were analyzed with flow cytometry to determine the percentages of EGFP-positive cells among RFP-positive cells. (G) BCBL-1 cells were treated with 5 μM purvalanol A or DMSO as a vehicle control. At 96 h posttreatment, virion production was analyzed by measuring levels of virion-associated viral DNA with qPCR. (A, D, and E) Data are representative of three (A) or two (D and E) independent experiments. (B, C, F, and G) Data are the means and SDs from three independent experiments.

Cdk1 is required for the maintenance of KSHV latency in PEL cells.

We have shown that Myc knockdown can induce both KSHV reactivation and PEL cell apoptosis (32). We hypothesized that KSHV might have evolved the ability to sense proapoptotic signals to initiate reactivation as a survival mechanism. We therefore examined whether Cdk1 inhibition leads to KSHV reactivation in addition to inducing PEL cell apoptosis. To monitor KSHV reactivation, we used a reporter cell line, BC-3-G, which was derived from BC-3 cells. In BC-3-G cells, the expression of enhanced green fluorescent protein (EGFP) is driven by a minimal viral lytic promoter that responds specifically to RTA (62). When virus undergoes successful reactivation, RTA expression needs to be upregulated to a sufficient level for a sustained period of time in order for downstream viral lytic genes to be expressed at measurable levels to support virion production. Therefore, EGFP expression in BC-3-G cells can be used as an indicator of KSHV reactivation. Treatment of BC-3-G cells with 10 μM purvalanol A for 72 h increased the percentage of EGFP-positive cells compared with that for DMSO-treated cells (Fig. 2A), suggesting that Cdk1 inhibition induces KSHV reactivation. We also quantified the percentages of EGFP-positive cells with flow cytometry at 48 h posttreatment and found that there is an approximately 10-fold increase of the percentage of EGFP-positive cells (Fig. 2B). In addition, another highly specific inhibitor of Cdk1, RO-3306, which showed greater-than-15-fold selectivity against a diverse panel of eight human kinases (55), also increased the percentage of EGFP-positive BC-3-G cells (Fig. 2C).

To further confirm the essential role of Cdk1 in maintaining KSHV latency, we examined the effect of Cdk1 knockdown (Fig. 2D) on KSHV reactivation. Transduction of BC-3 or BCBL-1 cells with shCDK1 greatly increased levels of KSHV lytic protein K8 (Fig. 2E). In addition, cotransfection of shCDK1 and a red fluorescent protein (RFP) expression vector as a transfection marker increased the percentage of EGFP-positive cells among the RFP-positive transfected cell population compared to that for cells cotransfected with shCtrl and RFP (Fig. 2F), suggesting that Cdk1 knockdown induces KSHV reactivation. Furthermore, Cdk1 inhibition also increased virion production in BCBL-1 cells (Fig. 2G). Together, these results showed that Cdk1 is required for KSHV to maintain latency in PEL cells.

p38 is activated by Cdk1 inhibition and is required for KSHV reactivation.

Having shown that Cdk1 inhibition leads to KSHV reactivation, we asked which signaling pathway downstream of Cdk1 inhibition mediates KSHV reactivation. To this end, we examined the effect of Cdk1 inhibition on active phosphorylation of p38 and Erk1/2 mitogen-activated protein kinases (MAPKs) (Fig. 3A). MAPK pathways have been shown to play key roles in KSHV reactivation (60, 62). Treatment of BC-3 and BCBL-1 cells with purvalanol A results in increased active phosphorylation of p38 but not of Erk1/2 at 1 and 2 h posttreatment (Fig. 3A), indicating that Cdk1 inhibition activates p38 in PEL cells. We then examined whether p38 is required for KSHV reactivation induced by Cdk1 inhibition. Pretreatment of BC-3-G cells with one of two highly specific p38 inhibitors, p38-I-III (31) or p38-I-IV (41), abolished KSHV reactivation in cells treated with 10 μM purvalanol A (Fig. 3B). Therefore, Cdk1 inhibition in PEL cells leads to the activation of p38, which is required for KSHV reactivation.

Fig 3.

Fig 3

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p38 is required for KSHV reactivation induced by Cdk1 inhibition. (A) BC-3 or BCBL-1 cells were treated with 10 μM purvalanol A or DMSO as a vehicle control. At 0, 1, and 2 h posttreatment, whole-cell lysates were analyzed with Western blotting for p38 phosphorylation (Thr180/Tyr182) and Erk1/2 phosphorylation (Thr202/Tyr204). α-Tubulin was analyzed as a loading control. Data are representative of two independent experiments. (B) BC-3-G cells were pretreated with p38 inhibitor p38-I-III, p38 inhibitor p38-I-IV, or DMSO as a vehicle control for 1 h. Cells were then treated with 10 μM purvalanol A or DMSO. At 48 h posttreatment, cells were analyzed with flow cytometry to determine the percentages of EGFP-positive cells. Data are the means and SDs from three independent experiments.

Caspases inhibit KSHV reactivation upon Cdk1 inhibition.

Having established that Cdk1 inhibition leads to both apoptosis and KSHV reactivation in a population of PEL cells, we further explored the relationship between these two processes. To this end, we first used flow cytometry to evaluate whether KSHV reactivation preferentially occurs in the apoptotic or the nonapoptotic subpopulation of BC-3-G cells upon Cdk1 inhibition. We found that the majority of EGFP-positive cells belonged to the nonapoptotic subpopulation (Fig. 4A), indicative of potential antagonism between apoptosis and KSHV reactivation. The antagonism can result from inhibition of KSHV reactivation by apoptosis, inhibition of apoptosis by KSHV reactivation and lytic gene expression, or both. To determine whether apoptosis inhibits KSHV reactivation upon Cdk1 inhibition, we used the pan-caspase inhibitor Z-VAD to inhibit apoptosis induced by Cdk1 inhibition (Fig. 1D). Pretreatment of BC-3-G cells with Z-VAD increased the percentage of EGFP-positive cells approximately 2-fold (Fig. 4B), suggesting that caspases inhibit KSHV reactivation induced by Cdk1 inhibition. Similarly, Z-VAD also enhanced KSHV reactivation induced by another specific Cdk1 inhibitor, RO-3306 (55) (data not shown). Z-VAD treatment also increased KSHV reactivation induced by Cdk1 knockdown in BC-3-G cells (Fig. 4C). In addition, Z-VAD further increased virion production in BCBL-1 cells treated with purvalanol A (Fig. 4D). These results show that upon Cdk1 inhibition, caspases activated during apoptosis inhibit KSHV reactivation.

Fig 4.

Fig 4

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Caspase inhibition enhances KSHV reactivation induced by Cdk1 inhibition or Cdk1 knockdown. (A) BC-3-G cells treated with 10 μM purvalanol A for 72 h were stained with annexin V-PE and 7-AAD and analyzed with flow cytometry to examine EGFP expression and apoptosis. Percentages of EGFP-positive and EGFP-negative cells among the nonapoptotic, early apoptotic, and late apoptotic/necrotic subpopulations are shown. (B) BC-3-G cells were pretreated with 50 μM Z-VAD or DMSO as a vehicle control for 1 h. Cells were then treated with 10 μM purvalanol A or DMSO. At 48 h posttreatment, cells were analyzed with flow cytometry to determine the percentages of EGFP-positive cells. (C) BC-3-G cells were cotransfected with shCDK1 or shCtrl and an RFP expression vector. Cells were treated with 50 μM Z-VAD or DMSO as a vehicle control at 12 h posttransfection. At 48 h posttransfection, cells were analyzed with flow cytometry to determine the percentages of EGFP-positive cells among transfected (RFP-positive) cells. (D) BCBL-1 cells were pretreated with 50 μM Z-VAD or DMSO as a vehicle control for 1 h. Cells were then treated with 5 μM purvalanol A or DMSO. At 96 h posttreatment, virion production was analyzed by measuring levels of virion-associated viral DNA with qPCR. (A) Data are representative of two independent experiments. (B, C, and D) Data are the means and SDs from three independent experiments.

Caspases inhibit KSHV reactivation induced by p53 activation.

Because Cdk1 induces p53-independent KSHV reactivation in PEL cells (Fig. 1E), we asked whether caspases also inhibit KSHV reactivation in cells undergoing p53-dependent apoptosis. Activation of p53 with Nutlin-3 has been shown to induce apoptosis in PEL cells (46). We confirmed that Nutlin-3 can induce PEL cell apoptosis and also found that caspase inhibition with Z-VAD can prevent Nutlin-3-induced apoptosis (Fig. 5A). To examine the effect of Nutlin-3 on KSHV reactivation in the presence or absence of caspase inhibition, we treated BC-3-G cells with 5 μM Nutlin-3 or DMSO following pretreatment with 50 μM Z-VAD or DMSO. At 48 h posttreatment, Nutlin-3 alone slightly increased the percentage of EGFP-positive cells (Fig. 5B), suggesting that p53 activation by Nutlin-3 can induce KSHV reactivation to a small extent. Importantly, Z-VAD markedly enhanced the ability of Nutlin-3 to induce KSHV reactivation (Fig. 5B). Z-VAD was also able to increase the ability of Nutlin-3 to upregulate the mRNA level of RTA (Fig. 5C) and the protein level of K8 (Fig. 5D). These results indicate that caspases inhibit KSHV reactivation in PEL cells in which p53 is activated by Nutlin-3.

Fig 5.

Fig 5

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Caspase inhibition enhances KSHV reactivation induced by p53 activation. (A) BC-3-G cells were pretreated with 100 μM Z-VAD or DMSO as a vehicle control for 1 h before being treated with 10 μM Nutlin-3 or DMSO as a vehicle control. At 24 h and 48 h posttreatment, cells were analyzed with flow cytometry for apoptosis. (B) BC-3-G cells were pretreated with 50 μM Z-VAD or DMSO as a vehicle control for 1 h. Cells were then treated with 5 μM Nutlin-3 or DMSO as a vehicle control in the presence of Z-VAD or DMSO. At 48 h posttreatment, cells were analyzed with flow cytometry to determine the percentages of EGFP-positive cells. (C) BC-3 cells were pretreated with 100 μM Z-VAD or DMSO as a vehicle control for 1 h. Cells were then treated with 10 μM Nutlin-3 or DMSO as a vehicle control in the presence of Z-VAD or DMSO. At 24 h posttreatment, the levels of RTA mRNA were measured by qRT-PCR, normalized to the levels of actin mRNA, and presented as the fold induction over the levels of the DMSO control. (D) BC-3 cells were pretreated with 100 μM Z-VAD or DMSO as a vehicle control for 1 h. Cells were then treated with 10 μM Nutlin-3 or DMSO as a vehicle control in the presence of Z-VAD or DMSO. At 72 h posttreatment, whole-cell lysates were analyzed with Western blotting for K8 expression. Actin expression was analyzed as a loading control. (A and D) Data are representative of three independent experiments. (B and C) Data are the means and SDs from three independent experiments.

KSHV reactivation inhibits apoptosis induced by Cdk1 inhibition.

Having shown that inhibition of KSHV reactivation by caspases may contribute to the mutual antagonism between KSHV reactivation and apoptosis in PEL cells, we evaluated the possibility that KSHV reactivation may also inhibit apoptosis in PEL cells upon Cdk1 inhibition. We examined the effect of RTA knockdown on cell death induced by purvalanol A because of the essential role of RTA for KSHV reactivation (38, 53). We transduced BC-3 or BCBL-1 cells with shKRTA, an shRNA lentiviral vector that can effectively knock down KSHV RTA (32), or the control shRNA vector shCtrl. Cells were then treated with 10 μM purvalanol A or DMSO as a vehicle control for 96 h (Fig. 6). We found that shKRTA further reduced the viability of BC-3 (Fig. 6A) and BCBL-1 (Fig. 6B) cells treated with purvalanol A. Thus, KSHV reactivation can inhibit PEL cell death induced by Cdk1 inhibition.

Fig 6.

Fig 6

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KSHV reactivation inhibits apoptosis induced by Cdk1 inhibition. BC-3 (A) or BCBL-1 (B) cells transduced with shKRTA or shCtrl were treated with 10 μM purvalanol A or DMSO as a vehicle control. At 96 h posttreatment, the percentages of viable cells were measured with a trypan blue exclusion assay.

AKT promotes cell survival and facilitates KSHV reactivation induced by Cdk1 inhibition or p53 activation.

Having shown that KSHV reactivation inhibits apoptosis induced by Cdk1 inhibition (Fig. 6), we asked which signaling pathway activated during KSHV reactivation promotes cell survival. Previous studies showed that KSHV lytic proteins, such as K1 (54) and vGPCR (40), can activate the prosurvival AKT signaling pathway. We examined the effect of enforced RTA expression, which can initiate KSHV reactivation, on the activation status of AKT in BC-3 cells. RTA overexpression led to increased active phosphorylation of AKT at Ser473 (Fig. 7A), indicating that KSHV reactivation activates AKT, possibly through upregulating viral lytic proteins, such as K1 and vGPCR.

Fig 7.

Fig 7

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AKT promotes PEL cell survival and KSHV reactivation upon Cdk1 inhibition or p53 activation. (A) BC-3 cells were transfected with a plasmid expressing KSHV RTA or the empty backbone plasmid as a negative control (NC). At 24 h posttransfection, whole-cell lysates were analyzed with Western blotting for RTA expression and AKT phosphorylation (Ser473). α-Tubulin was analyzed as a loading control. (B) BC-3 and BCBL-1 cells were pretreated with 2.5 μM and 5 μM concentrations of the AKT inhibitor AKT-I-IV, respectively, or with DMSO as a vehicle control for 1 h. Cells were then treated with 10 μM purvalanol A, 5 μM Nutlin-3, or DMSO as a vehicle control. At 48 h posttreatment, cell viability was determined with a trypan blue exclusion assay as percentages of viable cells. (C) BC-3-G cells were pretreated with 2.5 μM AKT-I-IV or DMSO as a vehicle control for 1 h. Cells were then treated with 10 μM purvalanol A, 5 μM Nutlin-3, 20 ng/ml TPA, or DMSO as a vehicle control. At 48 h posttreatment, cells were analyzed with flow cytometry to determine the percentages of EGFP-positive cells. (D) BC-3-G cells were pretreated with either 50 μM Z-VAD or DMSO and either 2.5 μM AKT-I-IV or DMSO for 1 h. Cells were then treated with 10 μM purvalanol A, 5 μM Nutlin-3, or DMSO as a vehicle control. At 48 h posttreatment, cells were analyzed with flow cytometry to determine the percentages of EGFP-positive cells. (A) Data are representative of two independent experiments. (B, C, and D) Data are the means and SDs from three independent experiments.

We next asked whether increased AKT activity plays a role in inhibition of cell death by KSHV reactivation upon Cdk1 inhibition or p53 activation. Pretreatment of BC-3 or BCBL-1 cells with low concentrations of a specific AKT inhibitor, AKT-I-IV, drastically reduced the viability of PEL cells treated with either purvalanol A or Nutlin-3 while having a much smaller impact on cells treated with the vehicle control DMSO (Fig. 7B), suggesting that AKT activity is required for PEL cell survival upon Cdk1 inhibition or p53 activation.

Having shown that apoptosis inhibits KSHV reactivation upon Cdk1 inhibition (Fig. 4) or p53 activation (Fig. 5), we reasoned that increased cell death resulting from AKT inhibition would inhibit KSHV reactivation. Indeed, pretreatment of BC-3-G cells with AKT-I-IV abolished KSHV reactivation induced by either purvalanol A or Nutlin-3 (Fig. 7C). Interestingly, in contrast to the inhibitory effects of AKT inhibition on KSHV reactivation induced by Cdk1 inhibition or p53 activation, pretreatment of BC-3-G cells with AKT-I-IV further enhanced KSHV reactivation induced by 12-O-tetradecanoylphorbol-13-acetate (TPA), a well-known inducer of KSHV reactivation (Fig. 7C), consistent with results from a previous study (42). This may be due to the ability of TPA to promote cell survival independent of AKT activation (4), masking the effects of AKT inhibition. To determine whether the inhibition of KSHV reactivation by AKT results from enhanced cell death, we performed a set of experiments similar to those described in Fig. 6C with additional pretreatment of cells with Z-VAD or the vehicle control DMSO (Fig. 7D). Z-VAD pretreatment was able to rescue KSHV reactivation that would otherwise be abolished by AKT inhibition (Fig. 7D). Thus, abolishment of KSHV reactivation by AKT inhibition was probably due to increased caspase activities.

DISCUSSION

In this study, we showed that Cdk1 inhibition induces mutually inhibitory KSHV reactivation and apoptosis in PEL cells. Our current working model is summarized in Fig. 8. Cdk1 inhibition leads to both PEL cell apoptosis and KSHV reactivation, indicating that Cdk1 is required for both the maintenance of KSHV latency and PEL cell survival. Recently, we showed that Myc plays a similar role (32). Previous studies showed that nuclear factor kappa B (NF-κB) is another cellular factor playing a dual role of maintaining KSHV latency and promoting cell survival (6, 29). The ability of KSHV to sense proapoptotic signals to initiate reactivation may provide potential evolutionary advantages. Latency helps KSHV evade immune surveillance and elimination. However, when cells harboring latent KSHV are in danger of being eliminated by apoptosis, reactivation may enable the latent virus to escape from these cells. Therefore, the initiation of KSHV reactivation in response to various proapoptotic stimuli may help KSHV to evade elimination and to persist in the host.

Fig 8.

Fig 8

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Proposed model of antagonism between KSHV reactivation and apoptosis downstream of Cdk1 inhibition or p53 activation.

Although Cdk1 inhibition leads to both apoptosis and KSHV reactivation in a population of cells, we showed that KSHV reactivation occurs mostly in the nonapoptotic subpopulations of cells (Fig. 4A). We provided evidence that the antagonism between KSHV reactivation and apoptosis results from mutual inhibition between caspases activated during apoptosis and KSHV reactivation (Fig. 4, 5, and 6). Apoptosis can function as an innate cellular response to infection by depriving viruses of cellular machineries essential for virus replication (56). Apoptosis may also promote antigen presentation (28), enhancing host immune response to pathogens. Thus, it is also possible that KSHV may encode mechanisms to gauge the extent of apoptosis and to actively limit lytic gene expression when apoptosis advances to a stage that is difficult for the virus to control. Restriction of reactivation by caspases may help KSHV reduce exposure to the host immune system in the same vein as immune evasion provided by latency.

Our data also point to the coexistence of cellular signals with different roles in KSHV reactivation downstream of proapoptotic stimuli. In contrast to the inhibitory effects of caspases on KSHV reactivation (Fig. 4 and 5), p38, which is activated early during Cdk1 inhibition, promotes KSHV reactivation (Fig. 3). p38 is also required for KSHV reactivation induced by Myc knockdown (32) or p53 activation (data not shown), further supporting the notion that KSHV senses early signals downstream of proapoptotic stimuli to initiate reactivation, possibly to achieve kinetic advantage over the induction of apoptosis.

Lastly, we showed that RTA knockdown enhances cell death induced by Cdk1 inhibition (Fig. 6). Interestingly, RTA RNA interference (RNAi) with shKRTA led to decreased cell viability in the absence of purvalanol A treatment (Fig. 6). We observed a similar, albeit smaller, drop of cell viability with another RTA hairpin that was also less effective at inhibiting virus reactivation than shKRTA (data not shown), making the findings less likely to be merely a result of off-target effects of the hairpins we used. It is possible that RTA expressed at levels below the threshold for triggering virus reactivation may still regulate cellular survival pathways in a cell-autonomous manner. It is also possible that inhibition of spontaneous reactivation by these RTA hairpins suppresses the release of paracrine factors that may promote the survival of other cells harboring latent KSHV. In addition, we provided evidence that AKT activation during KSHV reactivation inhibits apoptosis and facilitates KSHV reactivation (Fig. 7). KSHV K1 (54) and vGPCR (40) may mediate the activation of AKT during KSHV reactivation. Other viral lytic proteins, such as vBcl-2 (48) and K7 (17), have been shown to have antiapoptotic functions and may contribute to the inhibition of apoptosis by KSHV reactivation.

Taken together, our results demonstrate that Cdk1 is required for both PEL cell survival and the maintenance of KSHV latency. More importantly, our data reveal a mutually inhibitory relationship between apoptosis and KSHV reactivation when PEL cells are subjected to a wide range of proapoptotic stimuli.

ACKNOWLEDGMENTS

We thank E. Cesarman and J. Jung for generously providing reagents, the Janis V. Giorgi Flow Cytometry Laboratory at UCLA for help on the FACS experiments, Victoria Bender for critical reading and editing, and all members of the Sun lab for helpful discussions.

This work was supported by a grant from the Burroughs Wellcome Fund and grants from the NIH (CA091791, CA127042, DE019085, and DE015612).

Footnotes

Published ahead of print 11 April 2012

REFERENCES

  • 1. Ambinder RF, Robertson KD, Moore SM, Yang J. 1996. Epstein-Barr virus as a therapeutic target in Hodgkin's disease and nasopharyngeal carcinoma. Semin. Cancer Biol. 7:217–226 [DOI] [PubMed] [Google Scholar]
  • 2. Aoki Y, Tosato G. 2004. Therapeutic options for human herpesvirus-8/Kaposi's sarcoma-associated herpesvirus-related disorders. Expert Rev. Anti Infect. Ther. 2:213–225 [DOI] [PubMed] [Google Scholar]
  • 3. Belanger C, et al. 2001. Human herpesvirus 8 viral FLICE-inhibitory protein inhibits Fas-mediated apoptosis through binding and prevention of procaspase-8 maturation. J. Hum. Virol. 4:62–73 [PubMed] [Google Scholar]
  • 4. Bertolotto C, Maulon L, Filippa N, Baier G, Auberger P. 2000. Protein kinase C theta and epsilon promote T-cell survival by a rsk-dependent phosphorylation and inactivation of BAD. J. Biol. Chem. 275:37246–37250 [DOI] [PubMed] [Google Scholar]
  • 5. Boulanger E, et al. 2005. Prognostic factors and outcome of human herpesvirus 8-associated primary effusion lymphoma in patients with AIDS. J. Clin. Oncol. 23:4372–4380 [DOI] [PubMed] [Google Scholar]
  • 6. Brown HJ, et al. 2003. NF-κB inhibits gammaherpesvirus lytic replication. J. Virol. 77:8532–8540 [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 7. Brummelkamp TR, Bernards R, Agami R. 2002. A system for stable expression of short interfering RNAs in mammalian cells. Science 296:550–553 [DOI] [PubMed] [Google Scholar]
  • 8. Bubman D, Guasparri I, Cesarman E. 2007. Deregulation of c-Myc in primary effusion lymphoma by Kaposi's sarcoma herpesvirus latency-associated nuclear antigen. Oncogene 26:4979–4986 [DOI] [PubMed] [Google Scholar]
  • 9. Burysek L, Yeow WS, Pitha PM. 1999. Unique properties of a second human herpesvirus 8-encoded interferon regulatory factor (vIRF-2). J. Hum. Virol. 2:19–32 [PubMed] [Google Scholar]
  • 10. Cesarman E, Chang Y, Moore PS, Said JW, Knowles DM. 1995. Kaposi's sarcoma-associated herpesvirus-like DNA sequences in AIDS-related body-cavity-based lymphomas. N. Engl. J. Med. 332:1186–1191 [DOI] [PubMed] [Google Scholar]
  • 11. Chang Y, et al. 1994. Identification of herpesvirus-like DNA sequences in AIDS-associated Kaposi's sarcoma. Science 266:1865–1869 [DOI] [PubMed] [Google Scholar]
  • 12. Chugh P, et al. 2005. Constitutive NF-κB activation, normal Fas-induced apoptosis, and increased incidence of lymphoma in human herpes virus 8 K13 transgenic mice. Proc. Natl. Acad. Sci. U. S. A. 102:12885–12890 [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 13. Curreli F, Friedman-Kien AE, Flore O. 2005. Glycyrrhizic acid alters Kaposi sarcoma-associated herpesvirus latency, triggering p53-mediated apoptosis in transformed B lymphocytes. J. Clin. Invest. 115:642–652 [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 14. Dittmer D, et al. 1998. A cluster of latently expressed genes in Kaposi's sarcoma-associated herpesvirus. J. Virol. 72:8309–8315 [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 15. Djerbi M, et al. 1999. The inhibitor of death receptor signaling, FLICE-inhibitory protein defines a new class of tumor progression factors. J. Exp. Med. 190:1025–1032 [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 16. Dupin N, et al. 2000. HHV-8 is associated with a plasmablastic variant of Castleman disease that is linked to HHV-8-positive plasmablastic lymphoma. Blood 95:1406–1412 [PubMed] [Google Scholar]
  • 17. Feng P, et al. 2002. Kaposi's sarcoma-associated herpesvirus mitochondrial K7 protein targets a cellular calcium-modulating cyclophilin ligand to modulate intracellular calcium concentration and inhibit apoptosis. J. Virol. 76:11491–11504 [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 18. Ford PW, et al. 2006. Raf/MEK/ERK signalling triggers reactivation of Kaposi's sarcoma-associated herpesvirus latency. J. Gen. Virol. 87:1139–1144 [DOI] [PubMed] [Google Scholar]
  • 19. Friborg J, Jr, Kong W, Hottiger MO, Nabel GJ. 1999. p53 inhibition by the LANA protein of KSHV protects against cell death. Nature 402:889–894 [DOI] [PubMed] [Google Scholar]
  • 20. Fujimuro M, Hayward SD. 2003. The latency-associated nuclear antigen of Kaposi's sarcoma-associated herpesvirus manipulates the activity of glycogen synthase kinase-3β. J. Virol. 77:8019–8030 [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 21. Fujimuro M, et al. 2003. A novel viral mechanism for dysregulation of beta-catenin in Kaposi's sarcoma-associated herpesvirus latency. Nat. Med. 9:300–306 [DOI] [PubMed] [Google Scholar]
  • 22. Godden-Kent D, et al. 1997. The cyclin encoded by Kaposi's sarcoma-associated herpesvirus stimulates cdk6 to phosphorylate the retinoblastoma protein and histone H1. J. Virol. 71:4193–4198 [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 23. Godfrey A, Anderson J, Papanastasiou A, Takeuchi Y, Boshoff C. 2005. Inhibiting primary effusion lymphoma by lentiviral vectors encoding short hairpin RNA. Blood 105:2510–2518 [DOI] [PubMed] [Google Scholar]
  • 24. Goga A, Yang D, Tward AD, Morgan DO, Bishop JM. 2007. Inhibition of CDK1 as a potential therapy for tumors over-expressing MYC. Nat. Med. 13:820–827 [DOI] [PubMed] [Google Scholar]
  • 25. Gray NS, et al. 1998. Exploiting chemical libraries, structure, and genomics in the search for kinase inhibitors. Science 281:533–538 [DOI] [PubMed] [Google Scholar]
  • 26. Guasparri I, Keller SA, Cesarman E. 2004. KSHV vFLIP is essential for the survival of infected lymphoma cells. J. Exp. Med. 199:993–1003 [DOI] [PMC free article] [PubMed] [Google Scholar] [Retracted]
  • 27. Gutierrez MI, Judde JG, Magrath IT, Bhatia KG. 1996. Switching viral latency to viral lysis: a novel therapeutic approach for Epstein-Barr virus-associated neoplasia. Cancer Res. 56:969–972 [PubMed] [Google Scholar]
  • 28. Inaba K, et al. 1998. Efficient presentation of phagocytosed cellular fragments on the major histocompatibility complex class II products of dendritic cells. J. Exp. Med. 188:2163–2173 [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 29. Keller SA, Schattner EJ, Cesarman E. 2000. Inhibition of NF-κB induces apoptosis of KSHV-infected primary effusion lymphoma cells. Blood 96:2537–2542 [PubMed] [Google Scholar]
  • 30. Knight JS, Cotter MA, II, Robertson ES. 2001. The latency-associated nuclear antigen of Kaposi's sarcoma-associated herpesvirus transactivates the telomerase reverse transcriptase promoter. J. Biol. Chem. 276:22971–22978 [DOI] [PubMed] [Google Scholar]
  • 31. Laufer SA, Wagner GK, Kotschenreuther DA, Albrecht W. 2003. Novel substituted pyridinyl imidazoles as potent anticytokine agents with low activity against hepatic cytochrome P450 enzymes. J. Med. Chem. 46:3230–3244 [DOI] [PubMed] [Google Scholar]
  • 32. Li X, Chen S, Feng J, Deng H, Sun R. 2010. Myc is required for the maintenance of Kaposi's sarcoma-associated herpesvirus latency. J. Virol. 84:8945–8948 [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 33. Li X, et al. 2010. Tpl2/AP-1 enhances murine gammaherpesvirus 68 lytic replication. J. Virol. 84:1881–1890 [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 34. Lindqvist A, van Zon W, Karlsson Rosenthal C, Wolthuis RM. 2007. Cyclin B1-Cdk1 activation continues after centrosome separation to control mitotic progression. PLoS Biol. 5:e123 doi:10.1371/journal.pbio.0050123 [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 35. Liu J, Martin HJ, Liao G, Hayward SD. 2007. The Kaposi's sarcoma-associated herpesvirus LANA protein stabilizes and activates c-Myc. J. Virol. 81:10451–10459 [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 36. Lubyova B, Kellum MJ, Frisancho JA, Pitha PM. 2007. Stimulation of c-Myc transcriptional activity by vIRF-3 of Kaposi sarcoma-associated herpesvirus. J. Biol. Chem. 282:31944–31953 [DOI] [PubMed] [Google Scholar]
  • 37. Lubyova B, Pitha PM. 2000. Characterization of a novel human herpesvirus 8-encoded protein, vIRF-3, that shows homology to viral and cellular interferon regulatory factors. J. Virol. 74:8194–8201 [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 38. Lukac DM, Renne R, Kirshner JR, Ganem D. 1998. Reactivation of Kaposi's sarcoma-associated herpesvirus infection from latency by expression of the ORF 50 transactivator, a homolog of the EBV R protein. Virology 252:304–312 [DOI] [PubMed] [Google Scholar]
  • 39. Mann DJ, Child ES, Swanton C, Laman H, Jones N. 1999. Modulation of p27(Kip1) levels by the cyclin encoded by Kaposi's sarcoma-associated herpesvirus. EMBO J. 18:654–663 [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 40. Montaner S, Sodhi A, Pece S, Mesri EA, Gutkind JS. 2001. The Kaposi's sarcoma-associated herpesvirus G protein-coupled receptor promotes endothelial cell survival through the activation of Akt/protein kinase B. Cancer Res. 61:2641–2648 [PubMed] [Google Scholar]
  • 41. Ottosen ER, et al. 2003. Synthesis and structure-activity relationship of aminobenzophenones. A novel class of p38 MAP kinase inhibitors with high antiinflammatory activity. J. Med. Chem. 46:5651–5662 [DOI] [PubMed] [Google Scholar]
  • 42. Peng L, et al. 2010. Inhibition of the phosphatidylinositol 3-kinase-Akt pathway enhances gamma-2 herpesvirus lytic replication and facilitates reactivation from latency. J. Gen. Virol. 91:463–469 [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 43. Radkov SA, Kellam P, Boshoff C. 2000. The latent nuclear antigen of Kaposi sarcoma-associated herpesvirus targets the retinoblastoma-E2F pathway and with the oncogene Hras transforms primary rat cells. Nat. Med. 6:1121–1127 [DOI] [PubMed] [Google Scholar]
  • 44. Rainbow L, et al. 1997. The 222- to 234-kilodalton latent nuclear protein (LNA) of Kaposi's sarcoma-associated herpesvirus (human herpesvirus 8) is encoded by orf73 and is a component of the latency-associated nuclear antigen. J. Virol. 71:5915–5921 [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 45. Rivas C, Thlick AE, Parravicini C, Moore PS, Chang Y. 2001. Kaposi's sarcoma-associated herpesvirus LANA2 is a B-cell-specific latent viral protein that inhibits p53. J. Virol. 75:429–438 [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 46. Sarek G, et al. 2007. Reactivation of the p53 pathway as a treatment modality for KSHV-induced lymphomas. J. Clin. Invest. 117:1019–1028 [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 47. Sarid R, Flore O, Bohenzky RA, Chang Y, Moore PS. 1998. Transcription mapping of the Kaposi's sarcoma-associated herpesvirus (human herpesvirus 8) genome in a body cavity-based lymphoma cell line (BC-1). J. Virol. 72:1005–1012 [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 48. Sarid R, Sato T, Bohenzky RA, Russo JJ, Chang Y. 1997. Kaposi's sarcoma-associated herpesvirus encodes a functional bcl-2 homologue. Nat. Med. 3:293–298 [DOI] [PubMed] [Google Scholar]
  • 49. Si H, Robertson ES. 2006. Kaposi's sarcoma-associated herpesvirus-encoded latency-associated nuclear antigen induces chromosomal instability through inhibition of p53 function. J. Virol. 80:697–709 [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 50. Simonelli C, et al. 2003. Clinical features and outcome of primary effusion lymphoma in HIV-infected patients: a single-institution study. J. Clin. Oncol. 21:3948–3954 [DOI] [PubMed] [Google Scholar]
  • 51. Song MJ, Li X, Brown HJ, Sun R. 2002. Characterization of interactions between RTA and the promoter of polyadenylated nuclear RNA in Kaposi's sarcoma-associated herpesvirus/human herpesvirus 8. J. Virol. 76:5000–5013 [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 52. Sturzl M, et al. 1999. Expression of K13/v-FLIP gene of human herpesvirus 8 and apoptosis in Kaposi's sarcoma spindle cells. J. Natl. Cancer Inst. 91:1725–1733 [DOI] [PubMed] [Google Scholar]
  • 53. Sun R, et al. 1998. A viral gene that activates lytic cycle expression of Kaposi's sarcoma-associated herpesvirus. Proc. Natl. Acad. Sci. U. S. A. 95:10866–10871 [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 54. Tomlinson CC, Damania B. 2004. The K1 protein of Kaposi's sarcoma-associated herpesvirus activates the Akt signaling pathway. J. Virol. 78:1918–1927 [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 55. Vassilev LT, et al. 2006. Selective small-molecule inhibitor reveals critical mitotic functions of human CDK1. Proc. Natl. Acad. Sci. U. S. A. 103:10660–10665 [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 56. Vaux DL, Haecker G, Strasser A. 1994. An evolutionary perspective on apoptosis. Cell 76:777–779 [DOI] [PubMed] [Google Scholar]
  • 57. Verschuren EW, Klefstrom J, Evan GI, Jones N. 2002. The oncogenic potential of Kaposi's sarcoma-associated herpesvirus cyclin is exposed by p53 loss in vitro and in vivo. Cancer Cell 2:229–241 [DOI] [PubMed] [Google Scholar]
  • 58. Westphal EM, et al. 1999. Induction of lytic Epstein-Barr virus (EBV) infection in EBV-associated malignancies using adenovirus vectors in vitro and in vivo. Cancer Res. 59:1485–1491 [PubMed] [Google Scholar]
  • 59. Wilson SJ, et al. 2007. X box binding protein XBP-1s transactivates the Kaposi's sarcoma-associated herpesvirus (KSHV) ORF50 promoter, linking plasma cell differentiation to KSHV reactivation from latency. J. Virol. 81:13578–13586 [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 60. Xie J, Ajibade AO, Ye F, Kuhne K, Gao SJ. 2008. Reactivation of Kaposi's sarcoma-associated herpesvirus from latency requires MEK/ERK, JNK and p38 multiple mitogen-activated protein kinase pathways. Virology 371:139–154 [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 61. Yu F, et al. 2007. B cell terminal differentiation factor XBP-1 induces reactivation of Kaposi's sarcoma-associated herpesvirus. FEBS Lett. 581:3485–3488 [DOI] [PubMed] [Google Scholar]
  • 62. Yu F, et al. 2007. Systematic identification of cellular signals reactivating Kaposi sarcoma-associated herpesvirus. PLoS Pathog. 3:e44 doi:10.1371/journal.ppat.0030044 [DOI] [PMC free article] [PubMed] [Google Scholar]

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