1, 25(OH)2 D3 Induces Reactivation and Death of Kaposi’s Sarcoma-Associated Herpesvirus of Primary Effusion Lymphoma cells

Kaposi’s sarcoma associated herpesvirus (KSHV) a gammaherpesvirus establishes perennial latency in the host with periodic reactivation. Occasionally change in the physiological condition like hypoxia, host cell differentiation can trigger the lytic switch and reactivation of the virus. The biologically active form of 1, 25(OH)2 D3 plays a critical role in the regulation of various physiological processes (e.g. regulation of mineral homeostasis and control of bone metabolism). Apart from its role in host physiology, 1, 25(OH)2 D3 has been implicated as a potential agent for the prevention and/or treatment of many a tumors. Here we show that 1, 25(OH)2 D3 induces both death of Kaposi sarcoma associated herpesvirus infected PEL cells and KSHV replication. 1, 25(OH)2 D3 mediated inhibition of proliferation was associated with apoptosis of the PEL cells, and virus reactivation. In addition, p38 signalling is required for KSHV reactivation. Furthermore, treatment of PEL cells with p38 inhibitor abrogated the expression of ORF57, thus blocking lytic switch. Furthermore, silencing of VDR resulted in reduced ORF57 expression compared to the control cells, signifying the potential role of 1, 25(OH)2 D3 in KSHV reactivation. Thus, our studies have revealed a novel role of 1, 25(OH)2 D3 in the regulation of KSHV reactivation and PEL cell death.

of angiogenesis, apoptosis, Immunomodulation, growth and differentiation of many cell types, including lymphoma cells [22][23][24][25][26] . VDR expression is reported in many cancers types including breast, prostrate, pancreas, colon, leukaemia's and lymphomas [27][28][29][30][31][32] . Exposure of these cells to 1, 25(OH)2 D3 induces apoptosis in cells. However, studies are lacking on the role of 1, 25(OH)2 D3 in viral pathogenesis, only very few studies have indicated that vitamin D3 deficiency may confer increased risk of influenza and respiratory tract infection 33,34 . In vitro studies have demonstrated the effect of 1, 25(OH)2 D3 in susceptibility and control of HIV infection 35 . Furthermore, pre-treatment of human monoblastoid U937 cell line and monocyte derived macrophages in cell culture model of HIV infection have demonstrated anti-viral effects 36 . However, the underlying mechanism or pathways involving these functions is unclear, due to varied activities and functions. In addition, it remains to be identified whether 1, 25(OH)2 D3 is protective or pathogenic in cases of viral infection.
Effect of 1, 25(OH)2 D3 on downregulation of NF-κB pathway in endothelial cells transformed by Kaposi sarcoma associated herpes virus G protein coupled receptor is known 37 . Further, it has been shown that 1, 25(OH)2 D3 also has anti-proliferative effect on KSHV GPCR transformed endothelial cells 38 . Gene expression profiling of PEL cells have demonstrated that VDR is highly expressed in PEL cells as compared to normal B and T cell lymphoma and their sensitivity to vitamin D analogue EB1089, implicates a role for VDR in KSHV pathogenesis 11 . In view of these facts, the current investigations were taken up to dissect the mechanism (s) of action of 1, 25(OH)2 D3 on PEL cells, in particular its effect on apoptosis and reactivation.

Material and Methods
Cells and Reagents. PEL cells (JSC-1 and HBL-6) were kindly provided by Erle Robertson (University of Pennsylvania). These cells were cultured in RPMI 1640 supplemented with 10% foetal bovine serum glutamine (300 mg/mL) and streptomycin (100 mg/mL) and penicillin (100 U/mL) under 5% CO2 at 37 °C. 1, 25(OH)2 D3 was purchased from Sigma-Aldrich and was reconstituted in 90% ethanol and stored at −80 °C in an inert atmosphere in the dark. In all experiments, equal amount of 90% ethanol were added to control cultures. Pan caspase inhibitor Z-VAD-FMK was purchased from R&D system. FITC annexin V apoptosis detection kit was purchased from BD Biosciences, SB203580 (p38 inhibitor) and PD98059 (ERK inhibitor) were purchased from InvivoGen, phorbol 12-myristate 13-acetate, sodium Butyrate and MTT reagent were purchased from Sigma-Aldrich.
Cell viability assay. All cells were plated in 96 well culture plate in complete medium at a density of 5 × 10 4 cells per well and treated with or without increasing concentration of 1, 25(OH)2 D3(10, 50, 100, 200 nM). The plates were incubated at 37 °C, 5% CO2, for 24, 48 and 72 hours, respectively. Then, MTT solution (10 μL) for a total volume of 100 μL was added in every well and incubated for 4 hours at 37 °C with 5% CO2. Subsequently, MTT-containing medium was removed gently and replaced with DMSO (100 μL per well) and absorbance was obtained at 570 nm on a microtiter plate reader.
shRNA mediated VDR knockdown. To knock down VDR expression, two validated lentiviral constructs expressing small hairpin RNA (shRNA) sequences to targeting 2 different regions of the human VDR transcript were used. The constructs were obtained from (Sigma-Aldrich). Details of the clones and target sequences are given in Table 1. Lentiviral particles were prepared using standard protocols, resuspended in serum-free media and used to transduce JSC-1 cells. After 48 h, stably transduced cells were selected for puromycin resistance (2.5 μg/mL) for 20 days.
Quantitative Real Time RT-PCR (qRT-PCR). Total RNA was extracted from cells using TRIzol reagent (Invitrogen, Life Technologies, USA) as per manufacturer's instruction, followed by treatment with DNase 1. One microgram total RNA was reverse transcribed using cDNA synthesis kit (Thermo Fischer, USA). Syber green R-5′-TTCCCGTTCTCAGCCTTGAC-3′ Table 2. List of qRT-PCR primers.
PCR was performed using primer specific for KSHV ORF57, RTA and the human GAPDH gene. Sequence of primers are mentioned in Table 2. Western blot analysis. Cells were lysed in modified RIPA buffer containing 150 mM NaCl, 1% NP-40, 50 mM Tris-HCl (pH 8), 0.5% deoxycholic acid, 0.1% SDS, 1% Triton X-100, protease and phosphatase inhibitors. Lysates were placed on ice for 45 minutes and then clarified by centrifugation. Supernatants were removed and total protein measured by Bradford assay. Forty microgram of protein lysate per lane was electrophoresed on 12% SDS-PAGE and transferred to nitrocellulose membranes. The membranes were blocked for 1 h in TBST blocking solution, containing 5% bovine serum albumin and then incubated with a primary antibody overnight at 4 °C. The membranes were washed at least 3 times with each wash for 10 min with washing solution (TBS and 0.1% Tween 20) and incubated for 45 min with appropriate horseradish peroxidase-conjugated secondary antibodies. The washed membranes were developed using ECL Blotting Substrate (Thermo Scientific). The β-actin,VDR, ORF57, K8α and ERK antibodies were purchased from Santa Cruz Biotechnology. The phospho-p38 mitogen-activated protein kinase (MAPK), LANA and caspase-3 antibodies were purchased from Imgenex.

Viral Load Assay.
For intracellular Viral load assay, DNA was isolated using Gene elute mammalian genomic DNA isolation kit according to manufacturer's instructions (Sigma-Aldrich) and KSHV replication was determined by qPCR using SYBR green PCR master mix (Agilent technology, USA). KSHV ORF57 gene expression were harvested and added to confluent monolayers of uninfected 293 cells in a 24 well dish. Polybrene (8 µg/ mL) was added to each well and the plate was spinoculated at 2500 rpm for 2 h at 26 °C as previously described 40 .
Ninety-six hours post-infection, intracellular viral loaded was determined by real time PCR. Furthermore, infection was validated by checking the expression of LANA by western blot.

Results
Antiproliferative effect of 1, 25(OH)2 D3 in PEL cell lines. The demonstration of VDR expression in diverse tumors and cancers has emphasized that the effect of 1, 25(OH)2 D3 is not limited to VDR expression only but also display a range of antiproliferative activities. JSC-1, HBL-6 and DG-75 cells were exposed to different 1, 25(OH)2 D3 concentrations, (0 to 200 nM), for 48 h and cell viability was tested. 1, 25(OH)2 D3 induced a dose-dependent loss of viability in JSC-1 and HBL-6 PEL cells as compared to control cells DG-75 (Fig. 1A).

1, 25(OH)2 D3 induces caspase-3 dependent cell apoptosis.
To determine whether the inhibitory effects of 1, 25(OH)2 D3 on viability was associated with the induction of apoptosis, we evaluated the percentage of apoptotic cells by annexinV/PI staining. 1, 25(OH)2 D3 treated JSC-1 cells showed significantly higher percentage of apoptotic cell after 48 h ( Fig. 2A). On the other hand, HBL-6 did not show any significant change in apoptosis ( Fig. 2A). Simultaneously we also evaluated the effect of 1, 25(OH)2 D3 on the expression of pro and anti-apoptotic proteins. Significant increase in the cleaved PARP and caspase-3 was observed in JSC-1 cells (Fig. 2B). However, only a modest change in the level of these proteins was found in HBL-6 cells (Fig. 2C) (Fig. 4A and B). We then tested whether the increase in the expression of lytic genes also correlated to progeny virus production. For this latently infected JSC-1 cells were used, wherein cell-free virus was isolated from JSC-1 supernatants 2 days post 1, 25(OH)2 D3 treatment,. Viral DNA was extracted, and viral genome copy number determined by qPCR(4D,E and F). Figure 4G clearly demonstrates that 1, 25(OH)2 D3 induced virus production when compared to the controls. However, KSHV reactivation was comparatively lower in 1, 25(OH) 2 D3 treatment than in the positive control, TPA (Fig. 4G). To validate further, a time course treatment of JSC-1 and HBL-6 cells with 1, 25(OH)2 D3 or a combination of both TPA and sodium butyrate (NaB) as a potent positive control for KSHV reactivation was performed [0,6,12,24,36 and 48 h] (Fig. 4C). Immunobloting, for expression of ORF57 and K8α showed lytic replication at the 24 h and 36 h with peak activation at 24 h (Fig. 4C). These results indicate that 1, 25(OH)2 D3 induces expression of lytic genes and progeny virus production.

1, 25(OH)2 D3 activates KSHV replication involving MAPK signalling pathway. To gain insights
into the mechanism underlying the role of 1, 25(OH)2 D3 in the induction of reactivation, we explored cellular signalling pathways that may mediate KSHV reactivation downstream of VDR signalling. Previous studies have shown that the mitogen-activated protein kinase (MAPK) signalling pathways play important roles in KSHV reactivation induced by phorbol esters, Ras and oxidative stress [41][42][43][44] . We therefore examined whether MAPK signalling is required for KSHV reactivation induced by 1, 25(OH)2 D3. PD 98059, a specific inhibitor of ERK, and SB 203580, a specific inhibitor of p38 MAPK, significantly inhibited KSHV reactivation induced by 1, 25(OH)2 D3, as indicated ORF57 protein levels (Fig. 5B). In contrast, the JNK inhibitor SP 600125 did not significantly affect KSHV reactivation induced by 1, 25(OH)2 D3 (Fig. 5B). Since 1, 25(OH)2 D3 upregulated RTA transcription ( Fig. 4A and B), we further tested whether ERK or p38 signaling is involved in this upregulation. Both PD 98059 and SB 203580 inhibited RTA upregulation but not SP 600125 (Fig. 5C). Thus, ERK and p38 signalling are involved in the upregulation of RTA and KSHV reactivation downstream of VDR signalling.

Effect of VDR knockdown on 1, 25(OH)2 D3 mediated KSHV Reactivation.
To further confirm the role of VDR in KSHV reactivation, we also used lentiviral shVDR, a plasmid that express short hairpin RNA (shRNA) targeting VDR, to examine the effect of VDR depletion on KSHV reactivation. Table 1 shows the 2 shRNA constructs targeting 2 discrete regions of the human VDR transcript. We selected transfected cells with puromycin resistance gene to obtain stably transduced cells JSC-1-VDRKO. VDR knock-down JSC-1 cells were used in 1, 25(OH)2 D3 reactivation assays. Viral reactivation was measured by western analysis for the lytic protein, ORF-57. The control cell line, pTRCJSC-1 showed significant reactivation with 1, 25(OH)2 D3 (Fig. 6D) as compared VDR knockdown cells. Notably, all knock-down cell lines were responsive to lytic reactivation by TPA, and showed significant reactivation (Fig. 6D). In summary, VDR knock-down lowers 1, 25(OH)2 D3 induced viral reactivation.

Discussion
In summary, we report here compelling evidence to KSHV reactivation through VDR signalling. VDR mediated reactivation from latency offers a paradigm for how KSHV may initiate lytic replication in vivo. The mechanisms controlling KSHV latncy and lytic replication are complex. Whether KSHV undergoes latent or lytic replication might depend on diverse factors including: the status of cellular signalling pathways, cell cycle, extracellular factors, cell types, stages of viral infection, and viral regulatory factors, and susceptibility to disease development [45][46][47][48][49][50][51] . Understanding the key cellular and molecular basis of KSHV latency and reactivation may provide newer control stategies. In this report, we demonstrate the involvement of VDR-dependent signal transduction in KSHV reactivation in latently infected cells. The present work dissects the action of 1, 25(OH)2 D3 in PEL cells. Vitamin D receptor belongs to the superfamily of steroid receptors, which act as ligand dependent transcription factor. It is reported that VDR is constitutively expressed in primary effusion lymphoma B cells at high levels 11,52 . It has been previously demonstrated that active form of 1, 25(OH)2 D3 promotes growth inhibition in lymphocytes and in a variety of human cancer cell lines [53][54][55][56][57] . PEL cell line used in this study responds robustly to 1, 25(OH)2 D3 during a 48 h treatment period (Fig. 1). We observed strong growth inhibition at early time point from 24 h. However, this response was lost with incubation beyond 72 hours in JSC-1 cells (Fig. 1A). In contrast, a delayed response was noted in HBL-6 cells and growth inhibition starts 72 h post treatment (Fig. 1B). On the other hand, DG-75 cells showed no inhibition. Difference in sensitivity of these (PEL) cells to ligand may be due to the difference in receptor expression (Fig. 1C).
Reduction in growth, proliferation and induction of apoptosis are likely to cause Herpes virus reactivation [58][59][60] . Previous studies have shown that two seemingly conflicting phenotypes of KSHV reactivation and the death of PEL cells occur simultaneously 43 . Notably, 1, 25(OH)2 D3 mediated inhibition of proliferation was associated with apoptosis of the PEL cells (Fig. 2). On the other hand, the reactivation of PEL cells observed (expression of lytic transcripts ORF57 and K8.1) was not affected by pan caspase inhibitor ( Fig. 3C and D) although it was able to suppress apoptosis (Fig. 3A), clearly indicating that these two actions are independent of each other. Even though the extent of reactivation by 1, 25(OH)2 D3 is comparatively lower than those caused by strong inducers, such as TPA. More importantly, unlike TPA and butyrate, 1, 25(OH)2 D3 is a natural product of cellular metabolism and plays a critical role in several physiological and pathological conditions. It is likely that 1, 25(OH)2 D3 may play a pivotal role in regulation and equilibrium between latent and lytic replication in PEL cells. Thus, our findings elucidate one of the possible mechanisms for the pathogenesis and reactivation associated with KSHV infection. To identify the mechanisms of reactivation of KSHV in PEL cells, several different signalling pathways have been investigated. Several authors have shown the involvement of MAPK pathways in 1, 25(OH)2 D3 treated cells. MAPK p38 has been shown to be involved skeletal and intestinal cells, thereby affecting cell cycle, growth and differentiation 61,62 . Previous reports have also shown the involvement of MAPK pathways in KSHV lytic replication during productive primary infection and reactivation from latency 63,64 . In this study, the p38 MAPK pathway however appears to be generally activated by 1, 25(OH)2 D3 in PEL cells as assessed by expression of p-p38 protein (Fig. 6A). Most importantly, this activation of p38 MAPK by 1, 25(OH)2 D3 led to reactivation of KSHV in PEL cells. Furthermore, the p38 kinase inhibitor SB203580 not only prevented p38 phosphorylation but also abrogated KSHV reactivation (Fig. 5B). On the other hand, ERK inhibitor, PD98059 only partially suppressed KSHV reactivation, while there was no change with JNK inhibitor (Fig. 5A and B) suggesting that MAPK pathways p38 and ERK may be involved in switching from latency to lytic phase in KSHV.
Treatment of 1, 25(OH)2 D3 in PEL cells caused p38, ERK expression and caspase activation indicating that signalling events bifurcate downstream of VDR in mediating these two processes, i.e., virus reactivation and cell death. KSHV reactivation from latency depends on the expression of RTA. Our finding mirrors this as RTA expression occurred following stimulation by 1, 25(OH)2 D3 (Fig. 4A and B). As RTA does not contain any VDRE, it is speculated that 1, 25(OH)2 D3 indirectly increases the RTA expression most likely via p38. Thus, increase in the RTA expression induced by 1, 25(OH)2 D3 may lead to a greater degree of KSHV reactivation, making RTA a sensitive regulator between latency and reactivation as also confirmed through infection of 293 cells (Fig. 4G).
Lastly, we determined the effects of VDR knockdown on PEL cell proliferation and KSHV reactivation. VDR Knockdown rendered JSC-1 cells significantly less susceptible to 1, 25(OH)2 D3 mediated KSHV reactivation, while virus reactivation by phorbol esters remained intact (Fig. 6D). The findings further suggest that 1, 25(OH)2 D3 may activate latent KSHV in vivo. Thus, our findings clearly establish a key role in which VDR signalling allows the virus to escape a cell that is destined to die and induces KSHV reactivation and lytic replication. These findings imply a cross talk between a host cell and a latent KSHV that determine the clinical consequences.