Valproic Acid, a Histone Deacetylase Inhibitor, Is an Antagonist for Oncolytic Adenoviral Gene Therapy

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doi:10.1016/j.ymthe.2006.07.009

Valproic Acid, a Histone Deacetylase Inhibitor, Is an Antagonist for Oncolytic Adenoviral Gene Therapy Naseruddin Ho¨ti,1 Wasim Chowdhury,1 Jer-Tsong Hsieh,2 Markus D. Sachs,3 Shawn E. Lupold,1 and Ronald Rodriguez1,* 1

Brady Urological Institute, Johns Hopkins University School of Medicine, Baltimore, MD 21287, USA 2 Department of Urology, University of Texas, Southwestern, Dallas, TX, USA 3 Department of Urology, Charite Medical School, Humboldt University Berlin, Schumannstrasse 20/21, 10117 Berlin, Germany *To whom correspondence and reprint requests should be addressed at the Brady Urological Institute, Marburg 205, Johns Hopkins Hospital, 600 North Wolfe Street, Baltimore, MD 21287, USA.

Available online 20 September 2006

Oncolytic adenoviruses preferentially replicate in and lyse tumor cells. However, their application to cancer gene therapy has been complicated by the low levels of coxsackie and adenovirus receptor (CAR) expressed in many solid tumors. Histone deacetylase inhibitors (HDACIs) significantly upregulate CAR expression in tumor cells and have additional antineoplastic activities. Therefore, there is a clear rationale for the combination of HDACIs and oncolytic adenoviral gene therapy. We present evidence that HDACI treatment significantly inhibits adenoviral replication, viral burst, and tumor cell kill. Valproic acid (VPA), a well-established HDACI, inhibits adenoviral replication late in the viral life cycle. We hypothesized that VPA induction of the cell-cycle-regulating protein p21WAF1/CIP1 may be partly responsible for this activity. We demonstrate that p21WAF1/CIP1 expression alone limits viral replication and decreases viral titers in different cancer cell models. We also demonstrate that VPA and replicating adenovirus mutually inhibit each otherTs ability to kill cells, independent of p21WAF1/CIP1 expression. These results not only identify the importance of p21WAF1/CIP1 in the biology of adenoviral replication, but also suggest that oncolytic adenoviral gene therapy will be inhibited rather than enhanced by VPA (HDACI) treatment. Key Words: adenovirus, VPA, fiber-IRES-GFP, p21WAF1/CIP1, E1A

INTRODUCTION Human adenovirus serotype 5 (Ad5) is a well-characterized gene therapy vector that has shown much promise in preclinical models and in limited clinical trials. These viruses can be modified to replicate preferentially in cancer cells, leading to tumor-selective cell kill or oncolysis [1–3]. However, many solid tumors are refractory to adenoviral infection because the cellular receptor, the coxsackie and adenovirus receptor (CAR), is lacking or significantly underexpressed [4–7]. Recently histone deacetylase inhibitors (HDACIs) have been shown to upregulate adenovirus receptors in certain cancer cells [4,8– 10] and to have independent anti-tumor benefits [11]. Therefore the combination of HDACIs with oncolytic adenoviruses is being explored in preclinical models of adenoviral cancer gene therapy [12]. Valproic acid (VPA) is a well-known class I HDACI that has been found to inhibit tumor cell proliferation and promote differentiation [13]. All known HDACI, including VPA, up-regulate p21WAF1/CIP1 (p21) expression [14,15], an important mediator of growth arrest and

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senescence in mammalian cells. Elevated p21 expression leads to growth arrest in both G1 and G2 phases of the cell cycle by inhibiting cyclin-dependent kinase (CDK) complexes [16]. Furthermore, p21 interacts with procaspase-3 in mitochondria, which prevents caspase-3 activation and apoptotic cell death [17]. Here we report for the first time that valproic acid inhibits adenoviral replication and viral spread. Cancer cells treated with both VPA and wild-type adenovirus 5 were found to be more resistant to induction of cell death than those treated with VPA or adenovirus alone. This effect was found to be VPA dose dependent. We provide evidence that the cell cycle inhibitor protein p21 plays a major role in rendering cells resistant to adenovirus replication, thus revealing a novel function for the CDK inhibitor.

RESULTS VPA Inhibits Adenovirus Replication To evaluate the potential benefit of combining VPA and replication-competent adenovirus on cancer cells, we

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used a reporter virus that quantifies adenoviral replication by linking green fluorescent protein (GFP) expression to viral major late gene expression through an internal ribosome entry site (IRES). The reporter virus, FFIG, is an E1-deleted, replication-incompetent, type 5 adenovirus in which Fiber-IRES-GFP replaces the normal fiber gene (Fig. 1A, schematic). Since the fiber gene is under the control of the major late promoter, its expression does not become evident until late in viral replication. Co-infection of FFIG with a replicative virus,

FIG. 1. VPA treatment significantly compromises adenovirus replication. (A) LNCaP cells treated with VPA at different concentrations (0, 1.2, 2.5, and 5.0 mM) were infected with CN702 adenovirus in the presence of a reporter Ad5Fiber-IRES-GFP virus. By following GFP expression, which is under the control of the replication-specific major late promoter (schematic), viral replication kinetics was quantified in real time. VPA treatment dose-dependently inhibits adenoviral replication. (B) Adenoviral- and VPA-induced cell kill in LNCaP cells was measured using the MTS assay. Untreated cells served as a mock for cells that had been infected with CN702 only or treated with 5 mM VPA for 72 h. VPA or CN702 alone significantly reduced viable cell numbers ( P b 0.05). The combination of VPA and CN702 had no significant effect on cell viability ( P N 0.05). This demonstrates that VPA-mediated inhibition of viral replication was not due to cell death. Error bars represent means F SE of quadruplicate experiments.

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CN702 (a wild-type Ad5 replication-competent E3deleted adenovirus [1]), or fully intact wild-type adenovirus (VR-1516) leads to replication of both viruses. Using this system, we were able to quantify adenovirus replication in real time in the face of drug stimuli. We plated LNCaP prostate cancer cells in a 96-well plate at a density of 2  104 cells per well. The next day we co-infected the cells with CN702 (m.o.i. 5) and FFIG reporter virus (m.o.i. 10) and incubated them with or without VPA (0-5 mM). As shown in Fig. 1A, adenoviruses were able to replicate more proficiently in the absence of VPA, compared to cells that were treated with VPA. Furthermore, the effects of VPA on adenoviral replication were dose dependent. We obtained similar results with DU145 and C4-2 prostate cancer cell lines (Supplementary Figs. 1A and 1B). This decrease in viral replication was not due to VPA- or CN702-induced cell death. The combination of CN702 and VPA (5 mM) demonstrated no significant change in cell viability compared with untreated cells (Fig. 1B), in which treatment with CN702 or VPA alone resulted in significant decreases in cell viability. Therefore the combination of VPA and viral gene therapy appears less effective at stimulating cell death. These data suggest that cell death induced by VPA and that induced by CN702 are mutually inhibitory. To demonstrate further that VPA inhibition of viral replication is independent of the E3 region that harbors viral death proteins we used fully intact wild-type adenovirus (VR-1516) and compared it with CN702 (E3deleted virus) in the presence and absence of VPA. LNCaP cells that had been treated with various concentrations of VPA in the presence of wild-type adenovirus showed a dose-dependent decrease in replication similar to that of CN702 virus compared to untreated VPA controls (Supplementary Figs. 1C and 1D). We obtained similar results with other HDACIs Supplementary Figs. 1E and 1F). Finally, we evaluated the combination of VPA and replication-competent adenovirus on T24 bladder cancer cells. This cell line expresses little to no CAR protein in the absence of HDACI treatment but can be induced to express CAR after being treated with various HDACIs (Supplementary Fig. 2A). Using T24 cells with different HDACIs, including VPA, dose dependently increased the infection (transduction of viral genome) by replicationincompetent Ad-TrackCMV-GFP virus (Supplementary Figs. 2B–2D). Despite the increases in viral infection, the replication-competent adenovirus CN702 showed little to no replication in T24 cells treated with various HDACIs (Supplementary Figs. 2E–2G). To demonstrate further that viral transduction into these cells was the consequence of the HDACI activity of these compounds, we used an analogue of VPA, valpromide (VPD), which lacks HDACI activity (Supplementary Fig. 3A), and found that VPD was unable to induce CAR expression or transduce viruses into T24 cells and thus was unable to facilitate CN702 replication in T24 cells (Supplementary

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expression, we treated PC-3 prostate cancer cells that had been stably transfected with CAR (PC-3 tailless) [24] with VPA in the presence or absence of wild-type and Ad-FFIG adenovirus. We observed a dose-dependent decrease in the GFP signal over time (Supplementary Fig. 4C); similarly the oncolysis of these cells by the wild-type adenovirus was significantly compromised when treated with both modalities at the same time (Supplementary Fig. 4E). All these data suggest that the inhibitory responses of HDACIs are independent of CAR expression.

FIG. 2. Lower viral replication in the presence of VPA is accompanied by lower CPE and viral output. LNCaP cells infected with CN702 (m.o.i. 5) and reporter FFIG virus (m.o.i. 10) were treated with or without VPA (0, 0.1, 2.4, 5 mM). 72 h posttreatment, cells were analyzed by phase-contrast and fluorescence microscopy. The combination of VPA and adenovirus resulted in a dosedependent reduction of cytopathic effect and viral spread. (B) Viral output of CN702 virus (m.o.i. 5) from LNCaP cells treated with or without VPA (at given concentrations) was titered and reported as output:input ratios from three individual experiments 72 h p.i. VPA dose-dependently reduced output titers. Error bars represent F SE from three different experiments. *Difference between the untreated and the treated LNCaP cells was statistically significant ( P b 0.05). (C) Total virus recovered from the in vivo xenograft models of LNCaP and DU145 cells after homogenization was subjected to viral titering on 293 cells. Data plots represent output-to-input ratio of CN702 virus for the VPA-treated and untreated animals. P values for LNCaP ( P = 0.031) and DU145 ( P = 0.0313) were statistically determined using Wilcoxon signed rank test and were significant between the treated and the untreated groups.

Figs. 3B and 3C). We also found that, unlike VPA, VPD was unable to rescue LNCaP cells from viral oncolysis (Supplementary Fig. 4A). Similarly, 0.1–2.4 mM VPD had no effect on viral replication in LNCaP cells (Supplementary Fig. 4B). Evaluation of viral replication in the presence of 5 mM VPD was hampered by significant cell death (Supplementary Figs. 4A and 4B). To clarify whether the inhibitory effects of HDACIs on viral replication and viral oncolysis were dependent on CAR

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VPA Inhibits Adenoviral Burst, Viral Spread, and Output Titers We applied fluorescence microscopy to follow viral infection and spread. We plated LNCaP cells on six-well plates at a density of 5  105 cells per well. We infected the cells on the following day using the same conditions as for Fig. 1 (5 pfu/cell of CN702, 10 pfu/cell of FFIG) in the presence or absence of VPA (0 to 5 mM). Differential interference contrast phase microscopy (i.e., Nomarski imaging) demonstrated an increase in cell survival when the cells were treated with both VPA and CN702 (Fig. 2A, Nomarski column). Cells treated with increasing amounts of VPA, when combined with the viral constructs, had an apparent dose-dependent increase in cell survival at 72 h posttreatment, consistent with our observation presented in Fig. 1B. While the surviving cell numbers increased with the combination of lytic adenoviruses and VPA, viral replication and spreading are markedly inhibited (Fig. 2A, GFP column). Similarly, viral replication is known to cause cell cytopathic effects (CPE), which include cell rounding and detachment. Cells that were infected with virus in the presence of VPA adhered to the surface, while infected cells in the absence of VPA demonstrated significant CPE. This VPA-induced inhibition of viral replication and spread was also reflected by decreasing viral titers (output:input), in a dose dependent fashion (Fig. 2B). To examine if this inhibitory effect of VPA on adenoviral replication remained in in vivo tumor xenograft models, we transplanted two different tumor cell lines (LNCaP and DU145 cells) subcutaneously into the dorsal region of nude Balb/c mice. The treatment group received 0.8% w/v VPA in drinking water 3 days prior to CN702 (1  108 pfu) intratumoral injections, while the control group received normal water as described under Materials and Methods. Similarly the VPA-treated group further received an ip VPA injection of 250 mg/kg/500 Al twice a day for 3 days starting from the day of viral inoculation. At the end of the experiment we resected and homogenized the tumors and titered the resulting homogenates on 293 cells. Titer data from the tumors were evaluated as input-to-output ratios. Animals treated with VPA had a significantly lower viral recovery ( P b 0.05) compared to untreated groups. These data are in good concordance with our in vitro data that VPA inhibits replication and decreases adenoviral inputto-output ratio.

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Induction of Early Adenovirus E1A and E1B Genes by VPA To dissect the molecular mechanisms underlying VPA inhibition of adenoviral replication, we evaluated adenoviral early and late protein expression to determine which phase of the viral life cycle is susceptible to treatment. To be consistent with the timing of other assays, we extracted cellular proteins 72 h postinfection. VPA treatment appreciably up-regulated adenoviral early protein expression (E1A and E1B) in a dose-dependent manner (Fig. 3A). However, the fiber protein, which is under the control of the major late promoter, was slightly down-regulated by VPA. These results suggest that the negative effects of VPA on viral replication do not occur in the early viral life cycle. The treatment of LNCaP cells with VPA dose-dependently increased p21 protein levels in the absence of adenovirus (Fig. 3B). This induction was independent of p53 expression. Interestingly, cells treated with high concentrations of VPA together with CN702 virus have reduced p21 expression levels (Fig. 2A, 2.4 and 5 mM VPA). These findings suggested a potential reciprocal relationship between p21 expression and viral replication. To elucidate further the mechanisms of VPA induction of viral early genes, we applied VPA to the 293HEK cell line, which constitutively expresses E1A and E1B. 293HEK cell treatment with various concentrations of VPA failed to induce E1A expression (Fig. 3C). These results suggest that VPA effects on viral biology may require other elements of the intact viral genome. Constitutive Expression of E1A Does Not Rescue Viral Replication in 293 Cells Treated with VPA Since VPA does not alter E1A or p21 expression in 293HEK cells (Fig. 3C), we reasoned that either the VPA effect on viral replication does not extend to this cell line or such an effect requires additional factors provided by the adenoviral genome. To test these competing hypotheses, we treated 293HEK cells with varying doses of VPA and followed viral spread of an E1-deleted viral vector that constitutively expresses GFP (Ad5-CMV-GFP). Hence, unlike with the FFIG replication reporter construct, GFP expression is constitutive and visible shortly after viral infection. Viral spread can be visualized microscopically by cell fluorescence. We infected cells with a low m.o.i. (0.1), so that viral burst and spread could be followed by microscopy. As shown in Fig. 4A, cells that were treated with VPA showed no viral spread, while the untreated cells actively packaged virus and it spread into the neighboring cells, forming green cometlike foci or plaques. We used a nonspreading, fiberdeleted adenovirus as a negative control for cell burst and spread (Supplementary Fig. 5A). We confirmed and further quantified these results by direct fluorescence measurements on a multiplate fluorometer (Supplementary Fig. 5B).

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FIG. 3. Early but not late gene expression of the adenovirus is induced by VPA. (A) VPA treatment of LNCaP cells at the given concentrations induces expression of E1A and E1B. The late viral capsid protein, fiber, was slightly down-regulated by VPA treatment. High levels of VPA in the presence of adenovirus also lead to decreased p21WAF1/CIP1 protein levels. (B) LNCaP cells treated with VPA in the absence of CN702 show an up-regulated expression of p21WAF1/CIP1, which is independent of p53. (C) 293HEK cells treated with increasing concentrations of VPA (0, 0.1, 0.6, 1.2, 2.4, and 5 mM) failed to show induced E1A expression. In all experiments, cells were harvested by 72 h postinfection/posttreatment.

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FIG. 4. Adenovirus burst assay in the presence and absence of VPA. 293HEK cells, which constitutively express E1A, were infected with E1A-deleted adenovirus Ad-5CMV-GFP at an m.o.i. of 0.1. Viral spread was monitored by fluorescence microscopy, for plaque formation (comet-like foci) and CPE, at the indicated concentrations of VPA for 72 h. Cells that were treated with VPA were resistant to adenoviral spread and CPE in a dose-dependent manner compared to the untreated cells.

Induced Expression of E1A Is a Consequence of p21WAF1/CIP1 Expression HDACIs are known to up-regulate p21 gene expression in a p53-independent manner. We were interested in determining if the observed VPA-induced expression of E1A was due to the up-regulation of p21. To answer this,

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we infected p21-proficient and p21-knockout HCT-116 cells with CN702 adenovirus (m.o.i. of 2) and evaluated E1A protein levels 36 h p.i. E1A levels were significantly higher in p21-proficient HCT116 cells compared to p21negative HCT-116 cells (Fig. 5A). In support of our previous data (Fig. 3A), the expression of p21 in HCT-

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FIG. 5. The CDK inhibitor p21WAF1/CIP1 regulates E1A expression. (A) HCT-116 p21 / and HCT-116 p21+/+ cells were infected with CN702 (m.o.i. 2), and 36 h p.i. total cellular protein was harvested and analyzed by Western blot. h-Actin was used to ensure equal amounts of protein loading across each wells. p21WAF1/CIP1 expression correlated with increased E1A expression. Similarly p21WAF1/CIP1 was down-regulated by viral infection in p21WAF1/CIP1 +/+ cells. (B) p21WAF1/CIP1 regulates the expression of E1A. HCT-116 p21 / cells were infected with CN702 adenovirus (m.o.i. 5) in the presence of increasing m.o.i. of a p21 adenoviral expression vector (Ad-CMV- p21WAF1/CIP1, m.o.i. 0, 2, 5, 10, 20, and 30); 36 h postinfection cell lysates were subjected to Western blot analysis.

116 p21+/+ cells was also drastically reduced after infection with CN702 wild-type adenovirus. To investigate further the role of p21 in E1A regulation, we co-infected HCT-116 p21 / cells with CN702 (m.o.i. 5) and increasing m.o.i. of a p21 adenoviral expression vector (AdCMV-p21) [25]. All cells received equal viral m.o.i. by normalization with a control adenovirus. E1A protein levels clearly correlated with p21 expression levels, in a dose-dependent fashion (Fig. 5B). These results suggest that p21-induced increases in E1A expression result in dysfunctional E1A or that p21 inhibits a downstream effector of E1A action. The CDK Inhibitor p21WAF1/CIP1 Is a Major Antagonist of Adenoviral Replication The p21 protein has been shown to interact directly with adenoviral E1A protein, resulting in inactivation of p21 CDK inhibitory activity [26]. We therefore hypothesized that p21 overexpression might be responsible for inhibiting adenoviral replication. To test this hypothesis, we applied the FFIG reporter virus to evaluate CN702 viral

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replication in cells treated with and without the p21expressing virus, Ad-CMV-p21, or a control virus. Results demonstrated that heterologous p21 expression significantly inhibits CN702 viral replication in the HCT-116 p21 / cell line (Fig. 6A, dark lines). Adenoviral replication was approximately threefold more efficient in cells lacking p21 expression than in the p21-positive HCT-116 p21+/+ reference cells (Figs. 6Aand 6B, No Tx). Furthermore, approximately fourfold more virus was produced by HCT-116 p21 / cells in comparison to p21+/+ cells (Supplementary Fig. 6). Adenoviral replication was also inhibited by additional heterologous p21 in the p21positive LNCaP prostate cancer cell line (Fig. 6C). AdCMV-p21 virus alone did not affect viral replication in HCT-116 parental cells, suggesting that in that cell line p21 effects are near saturable levels from autologous p21 expression (Fig. 6B). In all cell lines tested, the addition of 5 mM VPA resulted in a stronger viral inhibition than in untreated cells or an m.o.i. of 2 Ad-CMV-p21 alone (Figs. 6A–6C, gray lines). This effect was evident even in the presence of saturable p21 (Fig. 6B), suggesting non-p21-mediated antireplicative effects of VPA. The combination of VPA and Ad-CMV-p21 resulted in additional inhibition of viral replication in HCT-116 p21 / and LNCaP cells, indicating that the initial dosage of Ad-CMV-p21 (m.o.i. 2) was in the lower range of effect in these cell lines. Collectively, these results suggest that while p21 plays a major role in regulating adenoviral replication and spreading, it is not the only pathway by which VPA inhibits viral replication. Correspondingly, VPA treatment in p21-negative cells was shown to inhibit viral cell death in a dose-dependent manner (Fig. 6D), further supporting the role for a minor non-p21-mediated pathway for maximal inhibition of adenoviral replication by HDACIs.

DISCUSSION VPA is becoming increasingly appreciated as a potential cancer chemotherapeutic and has recently entered phase I/phase II clinical trials for solid tumors [27]. VPA has numerous anti-cancer activities, including inhibition of cell growth, induction of cellular differentiation, and inhibition of hypoxia-induced angiogenesis [28–31]. Additionally, VPA and other HDACIs have been shown to increase expression of the adenoviral receptor, CAR [4,8,9]. Cell surface CAR protein enhances cellular adhesion in tight junctions [32], which may explain the growth-inhibitory properties of CAR expression and the frequent down-regulation of CAR expression in many cancers [33]. The combination of CAR up-regulation and the antineoplastic effects of HDACIs predicts the obvious benefits of combining HDACIs and adenoviral gene therapy. This hypothesis has been well supported in the literature

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[8,10,12,34]. In this report, we demonstrate for the first time that the combination of replication-competent adenovirus with HDACIs will lead to reduced rather than increased therapeutic effect. VPA clearly inhibits adeno-

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viral replication, viral spread, and therefore any potential lytic therapeutic effect (Figs. 1 and 2). This inhibitory effect of VPA on the viral lytic cycle might be the consequence of HDACI activity of this compound, as no such effect was observed on the cells that were treated with valpromide (derivative of VPA that lacks HDACI activity) together with CN702 virus (Supplementary Fig. 4A). Using multiple cancer cell models, we were able to demonstrate that the VPA-induced p21 may be responsible for much of the inhibition of adenoviral replication. The cell-cycle-dependent kinase inhibitor, p21, has been linked to many other functions in addition to its activity to arrest the cells at G1 or G2. It is also known to interact with proliferating cell nuclear antigen (PCNA), with which it can inversely affect DNA replication [35]. Similarly p21 was found to interact with procaspase-3, inhibiting Fas-mediated cell death [17]. Using a p21 knockout and p21-proficient HCT-116 cells, we were able to demonstrate that viral replication was inversely correlated with the p21 status. Viruses replicate three times more efficiently in the absence of p21 (Fig. 6). Accordingly, p21-deficient cells produce far more virus per cell compared to cells that are p21 intact (Supplementary Fig. 6). A rational explanation for this might be that during adenovirus replication, cells are pushed into a pseudo-S phase of the cell cycle, which is a permissive phase for adenoviral DNA replication. It is possible that VPA abrogates this process by arresting the cells at either G1 or G2 phase [36], which could explain the deficiencies in adenoviral replication and spread despite the increases in adenoviral E1A expression. Furthermore, the interaction of p21 with PCNA may independently compromise viral DNA synthesis. The microenvironment within tumors is significantly different from that of normal tissues. A major difference is seen in the disorganized vasculature of tumors, which results in an unbalanced blood supply and significant perfusion heterogeneities. As a consequence, many regions within tumors are transiently or chronically hypoxic. It has been previously reported that hypoxic conditions lead to cell cycle arrest through p21 overexpression and induction of apoptosis that is independent of p53 [37,38]. Similarly there are data available that adenovirus replication is compromised in hypoxic tissue FIG. 6. p21WAF1/CIP1 and adenovirus replication. Cells were infected with a combination of either CN702 (m.o.i. 2), the control virus AdgvRxR (m.o.i. 2), and FFIG (m.o.i. 5) or CN702 (m.o.i. 2), Ad-CMV-p21 (m.o.i. 2), and FFIG (m.o.i. 5) (dark solid and dashed lines, respectively). Similarly, cells were treated with the same viral combinations in the presence or absence of 5 mM VPA (gray solid and dashed lines, respectively). Viral replication is indicated by GFP expression from the replication-deficient reporter virus FFIG. (A) HCT116 p21 / cells. (B) HCT-116 p21+/+ parental cells. (C) LNCaP cells (p21+/+). (D) Cell viability of HCT-116 p21WAF1/CIP1 / measured by MTS assay in cells infected with CN702 in the presence or absence of increasing concentrations of VPA. Data represent the fold survival of cells over that of untreated cells. Error bars indicate means F SE of the quadruplicate experiments. *P b 0.05.

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[39]. Our experimental finding might help explain these observations and suggests that any processes that result in hyperexpression of p21WAF-1 in these tissues might be counterproductive in oncolytic adenoviral therapy. To our knowledge this is the first report to demonstrate an HDACI as an antagonist to adenoviral replication. We also demonstrate that VPA treatment significantly up-regulates viral E1A expression (Fig. 3 and Table 1). However, the increased E1A appears to be dysfunctional, as the major late promoter is not induced by its presence. While p21 expression does appear to correlate well with the potential for adenoviral replication, it might not be the only mechanism of VPAinduced inhibition of replication. The combination of VPA with recombinant adenovirus Ad-CMV-p21 significantly lowered the viral replication more than p21 alone. Moreover, VPA treatment of p21-negative cells inhibits viral replication, though to a much lesser extent. Thus we predict that there might be pathways other than p21 in the cells that VPA activates to hinder adenovirus replication. In summary our results indicate the importance of p21 in the biology of adenovirus replication and further suggest that HDACIs, such as VPA, further inhibit adenoviral replication in a p21-independent pathway.

MATERIALS AND METHODS Reagents and antibodies. For Western blot analysis the following antibodies were used: pAb-E1A (1:1000; Santa Cruz Biotechnology, Santa Cruz, CA, USA), pAb-acetylated histone 3 (1:1000; Upstate, Charlottesville, VA, USA), mAb-p21 (1:1000; Upstate), mAb-p53 (1:1000; Upstate), mAb-E1B 55KD (1:200; Calbiochem, La Jolla, CA, USA), mAb h-actin (1:1000; Sigma, St. Louis, MO, USA), and anti-fiber mAb 4D2 (1:1000; AbCam, Inc., Cambridge, MA, USA). Anti-mouse IgG (HRP)-conjugated was from Sigma (1:20,000). RmcB monoclonal CAR antibody (1:1000) for FACS was from Chemicon (Temecula, CA, USA). Restriction enzymes were

TABLE 1: Densitometry data from Western blot analyses (Fig. 3) Tx

0 mM

0.1 mM

0.6 mM

1.2 mM

2.4 mM

5 mM

Fig. 3A E1A E1B Fiber p21

1.00 1.00 1.00 1.00

0.39 0.71 1.16 1.71

0.84 0.71 1.02 2.27

1.28 1.30 0.92 1.93

2.62 3.54 1.02 1.31

6.46 2.94 0.90 1.20

Fig. 3B p53 p21 H3Ac

1.00 1.00 1.00

1.27 1.21 1.40

1.15 1.26 0.36

1.03 1.31 1.57

1.21 1.32 4.38

0.94 1.45 9.79

Fig. 3C E1A 1.00 p21 1.00

1.14 1.04

1.04 1.02

1.21 1.07

1.09 0.86

1.12 0.98

Correlated densitometry data for Figs. 3A, 3B, and 3C demonstrating fold increase or decrease in the expression of the respective proteins after treatment with VPA at given concentrations.

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purchased from New England Biolabs (Beverly, MA, USA). Cell culture media were obtained from Cellgro (Herndon, VA, USA). Trypan blue was purchased from Invitrogen (Carlsbad, CA, USA). The majority of all other chemical reagents and compounds were ordered from Sigma, unless otherwise specified. Cell culture. LNCaP, 293HEK, and DU-145 cell lines were purchased from American Tissue Culture Collection (Manassas, VA, USA); HCT-116 p21+/+ parental and HCT-116 p21 / [18] were a kind gift from Dr. Bert Vogelstein (Johns Hopkins Oncology Center). LNCaP and DU145, HCT116 p21+/+ parental and HCT-116 p21 / , and 293HEK cells were respectively maintained in RPMI 1640 medium with l-glutamine (Cellgro), McCoy 5A (Cellgro), and DMEM (Cellgro), supplemented with heatinactivated fetal bovine serum 10% (GIBCO, Carlsbad, CA, USA), ciprofloxacin hydrochloride 5 Ag/ml (U.S. Biological, Swampscott, MA, USA), and gentamicin 50 Ag/ml (Quality Biological, Inc., Gaithersburg, MD, USA). Cells were allowed to grow to until 80–90% confluent and harvested with 0.05% trypsin/0.53 mM EDTA (Cellgro) before each subsequent passage. Western blot analysis. Cells were washed with 1 PBS and resuspended with 5 vol of cold lysis buffer (50 mM Tris–HCl, pH 7.5, 250 mM NaCl, 5 mM EDTA, 50 mM NaF, 0.5% NP-40) supplemented with protease inhibitor cocktail (Roche, Indianapolis, IN, USA). The cell lysate was incubated on ice for 30 min and was then centrifuged for 10 min at 48C. Equal amounts of proteins were separated by SDS–PAGE and the resolved proteins transferred to nitrocellulose membrane. After being blocked with 5% nonfat milk in TBST overnight at 48C, the blot was incubated with primary antibody for 1 h at room temperature. The membrane was then probed with HRP-conjugated secondary antibody for 1 h and developed with the ECL-Plus system (Amersham Pharmacia) using the manufacturerTs protocol. A reporter adenovirus (FFIG)-based system for measuring virus replication. We have applied a reporter system to quantify adenoviral replication in real time by linking GFP expression to the fiber gene through an IRES. This virus (FFIG) was a kind gift from Dr. Gary KetnerTs lab (Johns Hopkins Bloomberg School of Public Health). FFIG virus is an E1/E3-deleted first-generation replication-deficient adenovirus generated by brescueQ of fiber gene deletion by transfection of the shuttle vector containing the adenovirus type 5 Fiber-IRES-GFP flanked with wild-type adenovirus sequences into 293 cells infected with fiber-deleted adenovirus. Recombination in 293 cells leads to rescue of nonfunctional virus, such that only fiber recombinants are capable replicating and spreading (while the manuscript for this article was being prepared, this system was independently reported by Lie et al. [19]). Since fiber mRNA is part of the major late transcription unit, expression of GFP occurs only late in viral replication and therefore is a surrogate for the terminal phases of viral replication. The schematic representation of this design is provided at the top of each figure for which this reporter (FFIG) was used. For reporter experiments, cells were plated into 96-well plates at 1  104/well overnight and co-infected the next day with an E3-deleted wild-type replicationcompetent type 5 adenovirus (CN702) [1] at an m.o.i. of 5–10 pfu/cell and the reporter virus FFIG at an m.o.i. of 10–20 pfu/cell. The GFP fluorescence signals were measured at the indicated time points using a multiplate fluorometer (Cytofluor II; PerSpective Biosystems, Inc.), utilizing an excitation wavelength of 485 F 20 nm and an emission wavelength of 530 F 25 nm. Background fluorescence was measured in cells that were mock infected with FFIG. GFP data were plotted as fold GFP induction relative to the mock-infected cells. All samples were performed in either triplicate or quadruplicate and results plotted with error bars representing the standard error of the mean. Construction of Ad5-CMV-GFP and a pseudotyped Ad-Fiber-Fib1 virus. Both Ad5-CMV-GFP and Ad-Fiber-Fib1 were generated by the AdEasy vector system [23]. Briefly, the E1 shuttle vector, pAdTrack (which contains the CMV promoter driving GFP in the E1 region), was linearized by PmeI digestion and recombined with the respective AdEasy parent vectors in stably transformed BJ5183 Escherichia coli, as previously described [40]. Resulting recombinants constitutively express the GFP

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reporter from the viral E1 region and are therefore replication defective. The parent vector AdEasy-1 (Ad5, DE1, DE3) was applied for Ad5-CMVGFP virus and the parent vector pFex-Fib1 (AdEasy-1 with nonexpressing fiber) for the Ad-Fiber-Fib1 virus. Ad-Fiber-Fib1 contains a frameshift mutation, which completely interrupts fiber production. This lack of fiber completely impedes viral infection and spread (Supplementary Fig. 2). Resulting viral plasmids were linearized by PacI digestion and transfected for viral packaging. Ad5-CMV-GFP virus was produced in 293HEK cells. Ad-Fiber-Fib1 viruses were produced in 911 cells, which constitutively express wild-type Ad5 fiber. This complementation, known as pseudotyping, packages the Ad-Fiber-Fib1 genome with wild-type fiber on the capsids only. Following infection, only the mutant Fib1 fiber is expressed; therefore virus is unable to spread on standard E1-containing packaging lines. Both virus were amplified in their respective cell lines, purified using the AdenoPure adenovirus purification kit (PURESYN, Inc., Malvern, PA, USA), and titered using the Adeno-X Rapid Titer Kit (Clontech Laboratories, Inc., Mountain View, CA, USA). 3-(4,5-Dimethylthiazol-2-yl)-5-(3-carboxy-methoxyphenyl)-2-(4-sulfophenyl)-2H-tetrazolium (MTS) assay. The metabolic viability of the cells was monitored using a MTS assay kit (CellTiter 96) from Promega (Madison, WI, USA) [20]. The assay is based on the older, widely used 3(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide (MTT) assay, which measures the colored formazan produced in a reduction reaction driven by nutrient metabolism, which can be quantified spectrophotometrically. The MTS assay has a major advantage over the MTT assay, as the formazan formed during the MTS assay is soluble and exits the cells [21], while the formazan produced by MTT is crystalline and must be solubilized (which requires disrupting the cells) [22].Cells were seeded onto 96-well plates and cultured in the presence of test agents for the indicated time intervals. A mixture of MTS and phenazine methosulfate (an electron-coupling reagent; final concentrations, 333 Ag/ml and 25 AM, respectively) was added, and the cells were incubated for 30 min at 378C. Formazan formed from the reduction of MTS was quantified by measurement of absorbance of the medium at 490 nm using a microplate reader (all data have been normalized to the background signals). Flow cytometry for CAR expression. T24 cells were treated with VPA at different concentrations (0 to 5 mM for 72 h) or SAHA (5 AM). Similarly 293 cells were included as a positive control for CAR expression. Cells were harvested at a density of 106 cells/ml and incubated with a 1:1000 dilution of the RmcB monoclonal CAR antibody (Chemicon) in a singlecell suspension for 60 min at 48C. Cells were then washed with PBS containing 2% FBS and treated with R-phycoerythrin-conjugated rat antimouse IgG1 monoclonal antibody (BD PharMingen, San Diego, CA, USA) for 30 min at 48C. Finally, the cell suspension was washed and fixed with 1% paraformaldehyde. Fluorescence analysis was performed by fluorescence-activated cell analysis (FACSCalibur; Becton–Dickinson, San Jose, CA, USA) at the Johns Hopkins Flow Cytometry Core Facility. Determination of the positive cell population was carried out by gating the righthand tail of the negative control (secondary antibody only) for each cell line at 1% and applying this setting to each sample acquisition. Viral burst assay. Viral replication kinetics analysis was applied to 293HEK monolayers seeded in 24-well plates at 2  105 cells per well. The following day, cells were infected with 0.1 pfu/cell of Ad-CMV-GFP or control virus Ad-Fiber-Fib1. After infection, these cells were treated with different concentrations of VPA from 0.1 to 5 mM. The cultures were incubated in 1 ml medium for 3 days and every 24 h cells were observed by fluorescent microscopy for any GFP-expressing comet-like foci (viral burst) [23]. Total well fluorescence was also quantified by multiplate fluorometer (Cytofluor II; PerSpective Biosystems, Inc.) utilizing an excitation wavelength of 485 F 20 nm and an emission wavelength of 530 F 25 nm. Viral amplification: output-to-input assay. Viral output-to-input assays were performed using the Adeno-X Rapid Titer Kit (Clontech Laboratories, Inc.). Briefly cells were infected with CN702 adenovirus (m.o.i. 5) in six-well plates in the presence or absence of VPA. Seventy-two hours postinfection cells were harvested in the same medium and subjected to

MOLECULAR THERAPY Vol. 14, No. 6, December 2006 Copyright C The American Society of Gene Therapy

three rounds of freeze–thaw cycles. Total infectious viruses were measured by titering them on 293HEK cells according to the recommendation of the Adeno-X Rapid Titer Kit protocol. The bamplification ratioQ of a virus produced from an infected cell (output) to the amount originally used to infect the cells in the first place (input) was determined and plotted as output-to-input ratio. In vivo virus replication assay. Tumors xenografts were established by inoculating 5  106 cells into the dorsal region of athymic nude mice. Three weeks from the initial transplants animals were put on either 0.8% VPA w/v in their drinking water (VPA-treated group) or normal water (untreated group) for 3 days before viral injection. Each tumor was injected with 1  108 pfu of CN702 virus in 50 Al of PBS. Similarly the VPAtreated group further received an ip injection of 250 mg/kg/500 Al twice a day for 3 days. By the end of 72 h total tumor mass was homogenized in 2 ml of PBS buffer (OMNI International, GA, USA). After three rounds of freeze–thaw cycles at 378C total tissue lysates were centrifuged at 3000g for 20 min. Total virus recovery was quantitated by plating serial dilutions of the homogenate supernatants on 293HEK cells. Virus titers were determined by hexon staining using the Adeno-X Rapid Titer Kit (Clontech Laboratories). The amplification ratio of a virus produced inside the inoculated tumor (output) to the amount originally used to inject the tumor in the first place (input) was determined and plotted as output–input ratio. Statistical analysis. All experiments were done in triplicate or quadruplicate and plotted with standard errors of the mean. All statistical analyses were performed using Prism 4.0 (GraphPad, Inc.) or Excel running on an IBM PC-compatible computer using the Windows XP operating system. Statistical comparisons for paired data were analyzed by the Student t test for the in vitro assays, while Wilcoxon signed rank test was used to analyze the statistical significance for in vivo xenograft models. Statistical significance was defined as P b 0.05.

ACKNOWLEDGMENTS We are thankful to Dr. Gary Ketner (Johns Hopkins University School of Medicine) for constructing and providing us with FFIG adenovirus. We are also thankful to Dr. Bert Vogelstein (Johns Hopkins School of Medicine) for providing us with p21 wild-type and knockout HCT-116 cell lines. Furthermore we are thankful to Drs. Elizabeth Platz and Siobhan Sutcliffe (Johns Hopkins School of Public Health) for helping us with statistical analysis and Maggie Liu for her technical support throughout this study. This study was supported in part by grants from the NIH, 2T32DK07552 and 2P50CA58236-09A1; by the Maren Foundation; and by the Department of Defense Prostate Cancer Research Program, under Award DAMD17-03-2-0033, which is managed by the U.S. Army, Medical Research and Material. RECEIVED FOR PUBLICATION DECEMBER 16, 2005; REVISED JULY 31 2006; ACCEPTED JULY 31, 2006.

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