A Noncanonical Function of Polycomb Repressive Complexes Promotes Human Cytomegalovirus Lytic DNA Replication and Serves as a Novel Cellular Target for Antiviral Intervention

Polycomb group (PcG) proteins are primarily known as transcriptional repressors that modify chromatin and contribute to the establishment and maintenance of cell fates. Furthermore, emerging evidence indicates that overexpression of PcG proteins in various types of cancers contributes to the dysregulation of cellular proliferation. Consequently, several inhibitors targeting PcG proteins are presently undergoing preclinical and clinical evaluation. Here, we show that infection with human cytomegalovirus also induces a strong upregulation of several PcG proteins. Our data suggest that viral DNA replication depends on a noncanonical function of polycomb repressor complexes which is independent of the so-far-described enzymatic activities of individual PcG factors. Importantly, we observe that a subclass of inhibitory drugs that affect the abundance of PcG proteins strongly interferes with viral replication. This principle may serve as a novel promising target for antiviral treatment.

ABSTRACT

Chromatin-based modifications of herpesviral genomes play a crucial role in dictating the outcome of infection. Consistent with this, host cell multiprotein complexes, such as polycomb repressive complexes (PRCs), were proposed to act as epigenetic regulators of herpesviral latency. In particular, PRC2 has recently been shown to contribute to the silencing of human cytomegalovirus (HCMV) genomes. Here, we identify a novel proviral role of PRC1 and PRC2, the two main polycomb repressive complexes, during productive HCMV infection. Western blot analyses revealed strong HCMV-mediated upregulation of RING finger protein 1B (RING1B) and B lymphoma Moloney murine leukemia virus insertion region 1 homolog (BMI1) as well as of enhancer of zeste homolog 2 (EZH2), suppressor of zeste 12 (SUZ12), and embryonic ectoderm development (EED), which constitute the core components of PRC1 and PRC2, respectively. Furthermore, we observed a relocalization of PRC components to viral replication compartments, whereas histone modifications conferred by the respective PRCs were specifically excluded from these sites. Depletion of individual PRC1/PRC2 proteins by RNA interference resulted in a significant reduction of newly synthesized viral genomes and, in consequence, a decreased release of viral particles. Furthermore, accelerated native isolation of protein on nascent DNA (aniPOND) revealed a physical association of EZH2 and BMI1 with nascent HCMV DNA, suggesting a direct contribution of PRC proteins to viral DNA replication. Strikingly, substances solely inhibiting the enzymatic activity of PRC1/2 did not exert antiviral effects, while drugs affecting the abundance of PRC core components strongly compromised HCMV genome synthesis and particle release. Taken together, our data reveal an enzymatically independent, noncanonical function of both PRC1 and PRC2 during HCMV DNA replication, which may serve as a novel cellular target for antiviral therapy.

IMPORTANCE Polycomb group (PcG) proteins are primarily known as transcriptional repressors that modify chromatin and contribute to the establishment and maintenance of cell fates. Furthermore, emerging evidence indicates that overexpression of PcG proteins in various types of cancers contributes to the dysregulation of cellular proliferation. Consequently, several inhibitors targeting PcG proteins are presently undergoing preclinical and clinical evaluation. Here, we show that infection with human cytomegalovirus also induces a strong upregulation of several PcG proteins. Our data suggest that viral DNA replication depends on a noncanonical function of polycomb repressor complexes which is independent of the so-far-described enzymatic activities of individual PcG factors. Importantly, we observe that a subclass of inhibitory drugs that affect the abundance of PcG proteins strongly interferes with viral replication. This principle may serve as a novel promising target for antiviral treatment.

INTRODUCTION

Human cytomegalovirus (HCMV) is a double-stranded DNA virus which belongs to the subfamily of betaherpesviruses. This human pathogen has a high seroprevalence in the population and still poses a serious health risk. Like all herpesviruses, it induces lifelong persistence by the establishment of latency. While HCMV infection usually proceeds subclinically in immunocompetent individuals, primary infections as well as phases of reactivation can cause severe disease in patients with a compromised immune system, such as AIDS patients or transplant recipients. Furthermore, congenital HCMV infection is the leading virus-associated cause of fetal malformations and birth defects (1). The currently licensed anti-HCMV drugs are limited by toxic side effects and the development of antiviral resistance (2). Therefore, there is an ongoing need for new antiviral strategies.

During productive infection, viral gene expression advances in a well-regulated cascade-like manner comprising three phases (3). The expression of the immediate early (IE) genes, driven by the potent major immediate early enhancer-promoter (MIEP), is the prerequisite for a successful replication cycle. During latency, the synthesis of IE proteins and, thereby, the onset of lytic infection are blocked, and the viral genome is maintained as an extrachromosomal circular episome (4, 5). It is well established that epigenetic-based mechanisms play a crucial role in dictating the outcome of HCMV infection. Immediately after nuclear entry, the viral genome associates with histones, and the chromatin state of the viral genome decides whether the infection proceeds in a lytic or latent manner (6). Transcriptionally permissive euchromatin predominates during lytic replication, while latency is characterized by the formation of transcriptionally inactive heterochromatin. Consistently, the heterochromatin-associated complex polycomb repressive complex 2 (PRC2) has been reported to associate with the MIEP region of the HCMV genome to contribute to the silencing of gene expression in latently infected cells (7, 8). In contrast, there is evidence that PRC2 may promote productive replication, since it was shown that depletion or pharmacological inhibition of this complex impairs efficient HCMV gene expression (9, 10).

PRC2 is composed of three core components. Enhancer of zeste homolog 2 (EZH2) harbors the enzymatic activity of this complex, while embryonic ectoderm development (EED) and suppressor of zeste 12 (SUZ12) have important regulatory functions and are essential for the functionality of the complex (11). Through its methyltransferase activity, EZH2 catalyzes the di- and trimethylation of histone H3 at lysine 27, which leads to the condensation of the respective chromatin region. The latter histone modification is a hallmark of facultative heterochromatin, which, in contrast to constitutive heterochromatin, retains the potential to revert to euchromatin again. By this regulation of transcriptional programs, PRC2 is a key player in several cellular processes, such as cell differentiation, stem cell self-renewal, and cell cycle progression (11, 12). In its role as a transcriptional repressor, PRC2 works in concert with a second polycomb repressor complex termed PRC1. Its subunits RING finger protein 1B (RING1B) and B lymphoma Moloney murine leukemia virus insertion region 1 homolog (BMI1) form a heterodimeric E3 ubiquitin ligase, which catalyzes the ubiquitination of histone H2A (H2Aub) at lysine 119. Most studies demonstrated that PRC2-conferred histone methylation is the prerequisite for PRC1 recruitment and for subsequent processes of chromatin condensation, but recent reports indicate that histone ubiquitination by PRC1 can also create binding sites for PRC2 and thereby initiate heterochromatin formation (13–16). Strikingly, the PRC2 core protein EED is also able to physically interact with PRC1, functioning as an active part of it (17). This underlines the close interregulation between these complexes and suggests that EED is an epigenetic exchange factor for the recruitment and function of both PRC1 and PRC2.

Besides their well-established roles as transcriptional repressors, PRCs were recently linked to other cellular processes. Upon ionizing radiation, both polycomb group (PcG) proteins were found to accumulate at sites of double-strand breaks (DSBs) and to promote an efficient DNA damage response (DDR) (18, 19). In this context, both PRC1 and PRC2 seem to promote transcription inhibition at sites of DNA damage but were also suggested to participate in the choice between the two main DSB repair pathways, homologous recombination and nonhomologous end joining (18, 20). Furthermore, emerging studies link PcG proteins to the field of DNA replication (21–24). Several components of both PRC1 and PRC2 localize at active replication forks, and depletion of these factors is associated with slower progression, altered symmetry, as well as stalled replication forks (23, 24).

In this report, we focus on the role of both PRC1 and PRC2 during lytic HCMV replication. We observed a strong upregulation of all major PRC components and identified a proviral function of both complexes during viral DNA replication, which was independent of the catalytic activities of PRC1/2 but could be inhibited by drugs affecting the abundance of PRC core components.

RESULTS

Polycomb group protein expression is upregulated during productive HCMV infection.

Polycomb repressive complex 2 (PRC2) has previously been proposed to play important roles during HCMV infection (7–9). Since several studies indicate that PRC2 commonly works in concert with the related multiprotein complex PRC1, we set out to investigate the relevance of polycomb group (PcG) proteins of both complexes during productive HCMV infection in further detail. For this, we infected permissive human foreskin fibroblasts (HFFs) with HCMV strain AD169 at a multiplicity of infection (MOI) of 3 and analyzed PcG protein levels throughout the viral replication cycle (Fig. 1A). This revealed a strong upregulation of all major PRC1 components (RING1B and BMI1) and PRC2 components (EZH2, EED, and SUZ12) starting at 24 to 48 h postinfection (hpi). In particular, upregulation was pronounced for EED, EZH2, and RING1B, since these proteins were expressed at very low levels in uninfected fibroblasts. The histone modifications conferred by PRC1 and PRC2, ubiquitinylated histone H2A (H2Aub) and trimethylated histone H3 (H3K27me3), respectively, were increased in parallel. Furthermore, we ascertained that the increased expression of PcG proteins was virus strain independent, since we also observed strong upregulation following infection with the clinical isolate TB40/E (Fig. 1B). To answer the question of whether this upregulation is due to enhanced transcription of the respective genes, total RNA was isolated from AD169-infected HFF cells at the indicated times postinfection, and PcG-specific mRNA levels were measured by reverse transcription-quantitative PCR (qRT-PCR) (Fig. 1C). This revealed increased mRNA levels of representatives of both PRC1 (RING1B) and PRC2 (EED, EZH2, and SUZ12) corresponding to the times of enhanced protein expression. Next, we were interested to investigate whether viral gene expression was responsible for PcG protein upregulation. HFF cells were treated with cycloheximide (CHX) in parallel with infection with HCMV strain AD169 (Fig. 1D), thereby preventing de novo viral protein expression, as confirmed by the detection of the immediate early protein IE1. In addition, an infection experiment using UV-inactivated AD169 was performed (Fig. 1E). Western blot (WB) analysis revealed a prevention of PcG protein upregulation upon CHX treatment as well as after infection with UV-inactivated virus, which implies that viral gene expression is necessary for enhanced PcG protein expression following HCMV infection. Taken together, our data demonstrate an HCMV-induced upregulation of the major PRC1 and PRC2 core factors on the mRNA as well as protein levels during the course of infection.

FIG 1.

HCMV infection leads to an upregulation of polycomb group proteins. (A and B) HFF cells were infected with the laboratory HCMV strain AD169 (A) and the clinical strain TB40/E (B) at an MOI of 3. Samples were harvested at the indicated time points postinfection, and whole-cell extracts were subjected to subsequent SDS-PAGE and Western blot analysis. Cellular PcG proteins were detected by the indicated antibodies. The detection of viral immediate early (IE1), early (pUL44), and late (pp28, MCP) proteins was used to monitor progression of the replication cycle. β-Actin served as a loading control. (C) HFF cells were infected with AD169 at an MOI of 3 and harvested at the indicated time points postinfection. RNA was isolated using TRIzol and the Direct-zol RNA miniprep kit (Zymo Research) and synthesized into cDNA by RT-PCR. Transcript levels were assessed by SYBR green PCR, and relative mRNA levels were calculated by normalization against the value for the housekeeping gene GAPDH. Values are derived from biological triplicates and represent mean values ± standard deviations (SD). Statistical analysis was performed by ordinary one-way analysis of variance (ANOVA). n.s., not significant; *, P ≤ 0.05; **, P ≤ 0.01; ***, P ≤ 0.001; ****, P ≤ 0.0001. (D) HFF cells were infected with AD169 at an MOI of 3 and treated with cycloheximide (CHX) (150 μg/ml) in parallel. After the indicated time points postinfection, whole-cell lysates were prepared and subjected to Western blot analysis. Cellular and viral proteins were detected by the indicated antibodies. IE proteins (IE1 or IE2) served as controls for blocked de novo protein synthesis, while pp65 ensured the income of tegument proteins. β-Actin served as a loading control. (E) HFF cells were infected with either AD169 or UV-inactivated AD169 at an MOI of 1. Whole-cell extracts were generated at the indicated time points and subjected to Western blot analysis. Detection of proteins was equivalent to that for panel D.

PcG proteins relocalize to viral replication compartments following HCMV infection.

To further analyze the fate of polycomb group complexes during HCMV infection, we assessed their subcellular localization by indirect immunofluorescence analysis. Intriguingly, while PRC1 and -2 components exhibited an evenly distributed nuclear localization in noninfected cells (Fig. 2A and B, left), they relocalized into specific intranuclear inclusions during infection, which we determined to be viral replication compartments (VRCs) by costaining of the viral processivity factor pUL44, a marker for VRCs (25) (Fig. 2A and B, right). Next, we investigated the localization of the PRC-mediated histone modifications H3K27me3 and H2Aub as well as total histone H3 in mock-infected and infected cells. In uninfected cells, a diffuse distribution throughout the nucleus was detected (Fig. 2C, left). Interestingly, the repressive histone marks instituted by PRC1/2 turned out to be specifically excluded from VRCs in infected cells (Fig. 2C, right). In contrast, the localization of histone H3 was unchanged after infection (Fig. 2C, bottom right). Thus, these findings suggest a role of PRC1 and -2 during viral DNA synthesis, which, however, seems to be distinct from their enzymatic activity.

FIG 2.

Polycomb group proteins localize to HCMV replication compartments. HFF cells were infected with AD169 at an MOI of 1. At 72 hpi, cells were fixed with 4% paraformaldehyde and permeabilized with 1% SDS. Samples were stained with specific primary antibodies against the indicated PcG proteins (PRC2 core subunit factors [A] and PRC1 core subunit factors [B]) and corresponding histone modifications (PRC2 H3K27me3, PRC1 H2Aub, and total H3 [C]). The viral protein pUL44 served as a marker for viral replication compartments. The nucleus was counterstained with DAPI. Samples were analyzed by confocal microscopy.

PcG proteins play a proviral role for HCMV replication and spread.

Next, we wanted to clarify whether the upregulation of PcG proteins acts pro- or antivirally during infection. In order to address this, we generated HFF knockdown cells for single PRC1 (HFF/shRING1 and HFF/shBMI1) and PRC2 (HFF/shEZH2 and HFF/shEED) components by lentiviral transduction of the respective short hairpin RNAs (shRNAs). The knockdown efficiency was tested by Western blot analysis of mock- and AD169-infected knockdown HFFs (Fig. 3A). In noninfected cells, expression of the shRNAs resulted in a complete depletion of the respective PcG proteins (Fig. 3A, compare lanes 1 and 3 to lanes 5, 7, 9, and 11). However, upon infection, the HCMV-induced upregulation of PcG proteins could not entirely be inhibited by shRNA expression, with the notable exception of shBMI1 (Fig. 3A, compare lanes 5 with 6, 7 with 8, 9 with 10, and 11 with 12). Nonetheless, PRC protein levels were considerably reduced compared to those in infected cells that were transduced with the control shRNA shNone (empty vector) or shC (scrambled shRNA) (Fig. 3A, compare lanes 6, 8, 10, and 12 to lanes 2 and 4). Thus, in the context of HCMV infection, PcG protein expression could clearly be diminished by the expression of shRNAs. Additionally, knockdown of specific PRC components simultaneously reduced the protein levels of other core proteins, which is in accordance with current literature describing a high dependence of PcG protein stability on complex integrity and the interaction of all core components (26). In particular, EED and RING1B (Fig. 3A, lanes 7 to 10) influenced each other’s abundance. This is of particular interest since EED has been described as a shared component of both PRC1 and PRC2, thereby acting as an epigenetic exchange factor (17). This further supports the notion that both PRC complexes are functionally interconnected. Next, we used HFF knockdown cells in a multiround replication experiment in order to investigate the role of PcG proteins for HCMV infection (Fig. 3B). Knockdown and control cells were infected with the reporter virus AD169-GFP at a low MOI of 0.02, and the green fluorescent protein (GFP) signal was quantified at 7 days postinfection (dpi). We observed that knockdown of all PcG proteins reduced the viral replication efficacy significantly, indicating a proviral function of PRCs. Interestingly, the effect was most pronounced for the EED and RING1B knockdown cells. In summary, our findings suggest that PcG proteins play an important role during lytic HCMV infection, as they appear to be required for efficient viral replication.

FIG 3.

Knockdown of PcG proteins negatively affects HCMV replication. (A) Control cells (HFF/shNone and HFF/shC) and knockdown cells (HFF/shEZH2, HFF/shEED, HFF/shRING1B, and HFF/shBMI1) were infected with AD169 at an MOI of 0.1 and harvested together with mock-infected cells at 72 hpi. Whole-cell lysates were subjected to Western blot analysis. Cellular proteins were detected by the indicated antibodies. β-Actin served as a loading control. (B) PcG knockdown and control cells were infected with AD169-GFP at an MOI of 0.002 and lysed at 7 days postinfection to perform quantitative GFP fluorometry. Values are derived from biological quadruplicates and represent mean values ± SD. Statistical analysis was performed by ordinary one-way ANOVA. n.s., not significant; ****, P ≤ 0.0001.

HCMV depends on PcG functions at different steps of the viral life cycle.

Next, we aimed to specify the stage of infection at which PcG proteins exert their proviral function in order to obtain information about their mode of action. For this, we infected knockdown and control HFFs with AD169 at an MOI of 0.1 and harvested the cells at 72 hpi. By Western blot analysis, we detected viral proteins expressed during the IE (IE1p72 and IE2p86), early (E) (pUL84 and pUL44), and late (L) (IE2p60, IE2p40, monocyte chemoattractant protein [MCP], and pp28) phases of infection (Fig. 4A). Here, we concentrated on HFF/shRING1B and HFF/shEED cells as representatives of loss of PRC1 and PRC2 function, respectively, since these cell lines showed the most pronounced effect on viral replication (Fig. 3B). Knockdown of EED and RING1B led to a slight decrease of IE protein levels as well as a stronger reduction of E and especially L proteins (Fig. 4A, compare lanes 6 and 8 to lanes 2 and 4). A previous study had already indicated that depletion of EZH2 has a negative effect on the initiation of the IE phase of HCMV infection (9). In that study, PRC2 was demonstrated to epigenetically repress the expression of the cellular protein Gfi-1 (growth factor independence 1), which itself acts as a transcriptional repressor of the viral major immediate early enhancer-promoter (MIEP). However, considering the observed strong upregulation of PcG proteins and localization to VRCs as infection progresses, we assumed an additional function during later replication stages. Interestingly, closer analysis revealed that the requirement of PRC1/2 activity at IE times of infection is MOI dependent. As evident from Fig. 4B, only infection of EED and RING1B knockdown cells under low-MOI conditions (MOI of 0.1) (Fig. 4B, top) negatively affected IE1 protein abundance, while infection at an MOI of 0.5 or 1 (Fig. 4B, middle and bottom) resulted in equal IE1 expression levels. Thus, in order to circumvent the effect of PcG proteins on viral IE gene expression, we infected EED and RING1B knockdown cells with AD169 at an MOI of 1 and harvested the cells at 72 hpi (Fig. 4C). Western blot analysis revealed that while IE1 protein expression was not affected by EED or RING1B depletion, loss of PRC1 and -2 negatively affected viral E and especially L gene expression (Fig. 4C, compare lanes 6 and 8 to lanes 2 and 4). To investigate whether viral E genes are downregulated in the absence of RING1B, cells were harvested at either 24 hpi or 72 hpi in the presence of the DNA polymerase inhibitor foscarnet (phosphonoformic acid [PFA]). As shown in Fig. 4D, no effect on the expression of the viral early-late protein pUL44, pUL84, or pp65 could be detected under these conditions (Fig. 4D). In summary, our results reveal a novel, MOI-independent function of PRC1 and -2 components during the L phase of infection, which can be discriminated from effects of PRC components at the IE stage.

FIG 4.

PcG proteins function at different stages of HCMV replication depending on the MOI. (A) PRC knockdown cells and control cells were infected with AD169 at an MOI of 0.1 and harvested together with mock-infected cells at 72 hpi. Whole-cell lysates were subjected to Western blot analysis, and immediate early (IE1p72 and IE2p86), early (pUL44, pUL84, and pp65), and late (IE2p60, IE2p40, pp28, and MCP) viral proteins were detected, as indicated. β-Actin served as a loading control. (B) Knockdown cells and control cells were infected with AD169 at MOIs of 0.1, 0.5, and 1.0 and harvested at 24 hpi. Whole-cell lysates were prepared, and IE1 levels were assessed by Western blot analysis. (C) Knockdown and control cells were infected with AD169 at an MOI of 1 and harvested together with mock-treated cells at 72 hpi. Whole-cell lysates were subjected to Western blot analysis, and viral proteins were detected by the indicated antibodies as described above for panel A. β-Actin served as a loading control. (D) RING1B knockdown cells and control cells were infected with AD169 at an MOI of 1. In parallel with infection, 250 μM phosphonoformic acid (PFA) was added to the indicated samples. At 24 and 72 hpi, the cells were harvested and subjected to Western blot analysis for detection of viral protein levels as described above for panel A.

PcG proteins promote efficient HCMV DNA replication.

Since we observed an attenuation of late gene expression upon PRC knockdown, which depends on efficient viral DNA replication, as well as an accumulation of PcG proteins in VRCs (Fig. 2A and Fig. 4C), we hypothesized that PRCs may play a role during viral DNA replication. To investigate this, we infected HFF/shEED and HFF/shRING1B cells with AD169 at a high MOI (MOI of 0.5) (Fig. 4B, middle). At 10 hpi and 72 hpi, total DNA was extracted and submitted to quantitative TaqMan PCR for assessing viral genome copy numbers (Fig. 5A). In order to calculate the number of HCMV genomes per cell, the obtained values were correlated with the value for the cellular housekeeping gene albumin. Analysis of the parental viral genomes at 10 hpi showed no difference between control and knockdown cells, demonstrating that depletion of EED and RING1B did not affect the entry of viral particles (Fig. 5A, left). However, quantification of viral progeny DNA at 72 hpi showed a significant decrease of HCMV genomes in HFF/shEED and HFF/RING1B cells compared to control cells, indicating a positive role of PcG proteins for viral DNA replication (Fig. 5A, right). Since we assumed that reduced DNA synthesis would also affect the release of viral particles, we quantified viral genome copies in the supernatants of infected knockdown and control cells by TaqMan PCR (Fig. 5B). The drastic reduction of nascent viral particles released from EED and RING1B knockdown HFFs compared to control cells confirmed our hypothesis (Fig. 5B). In summary, these results highlight an important role of both PRC1 and PRC2 proteins during the process of HCMV genome amplification.

FIG 5.

PcG knockdown negatively affects viral DNA replication. (A) Knockdown and control cells were infected with AD169 at an MOI of 0.5, and total DNA was extracted at 10 hpi and 72 hpi using the DNeasy blood and tissue kit (Qiagen). Viral genomes were quantified by TaqMan real-time PCR specific for IE1, and genome copy numbers were calculated as indicated in the graph. Values are derived from triplicate experiments and represent mean values ± SD. (B) Knockdown and control cells were infected with AD169 at an MOI of 0.5, and supernatants were harvested at 72 hpi. Viral genomes released into the supernatant were quantified by TaqMan real-time PCR. HFF/shNone cells were set to 100%. Values are derived from biological triplicates and represent mean values ± SD. Statistical analysis was performed by ordinary one-way ANOVA. n.s., not significant; **, P ≤ 0.01; ****, P ≤ 0.0001.

PcG proteins associate with replicating HCMV genomes.

Our observation that PcG proteins are required for efficient HCMV DNA amplification (Fig. 5A) is in accordance with recent reports in the literature which directly implicated PcG factors in the regulation of cellular DNA replication (23, 24). In several studies, a direct association of PcG proteins with replicating host cell DNA could be demonstrated (21–24). To test whether PcG proteins also bind to replicating HCMV DNA, we performed accelerated native isolation of protein on nascent DNA (aniPOND) (22). This technique is based on the labeling of newly synthesized DNA with the nucleoside analog 5-ethynyl-2′-deoxyuridine (EdU) and biotinylating the EdU-labeled DNA by click chemistry, followed by affinity purification of the biotinylated DNA, and the subsequent analysis of the proteins associated with it. By indirect immunofluorescence analysis, we first confirmed that incorporation of the thymidine analog EdU preferentially occurred in viral DNA (Fig. 6A). HFF cells were grown to confluence prior to infection in order to achieve a growth arrest in G₀. This not only allowed an efficient start of HCMV gene expression and replication but additionally guaranteed a block of cellular DNA synthesis (27–31). At 72 hpi, we labeled mock- and AD169-infected HFF cells for 4 h with 10 μM EdU. After fixation, we performed antibody staining of pUL44 as a marker for VRCs and a click reaction with fluorophore-coupled azide for visualization of newly synthesized DNA (Fig. 6A). In noninfected cells, which were all pUL44 negative (Fig. 6A, top), only very few cells were labeled by EdU incorporation (1.33%) (see quantification in Fig. 6B). On the contrary, in infected samples (Fig. 6A, bottom), the majority of cells were EdU as well as pUL44 positive (see quantification in Fig. 6B). Moreover, the confocal microscopy images of infected cells in Fig. 6A (bottom) demonstrated a clear colocalization of EdU and pUL44, indicating an exclusive DNA replication activity in VRCs. In contrast to that in uninfected cells, EdU was evenly distributed throughout the nucleus when integrated into cellular DNA (Fig. 6A, top). After having demonstrated the preferential labeling of viral DNA, we used the same experimental setup as the one described above for aniPOND (Fig. 6C). After precipitation of the EdU-labeled DNA, we analyzed DNA-bound proteins. While the no-click control was negative for captured proteins, we were able to observe a coprecipitation of viral genomes with EZH2 and BMI1, indicating that these PRC1 and -2 core factors associate with newly synthesized viral DNA. In summary, these experiments suggest that PRC components such as EZH2 and BMI1 are physically present at nascent viral DNA, which supports the hypothesis of PcG proteins favoring viral replication.

FIG 6.

PcG proteins bind to replicating HCMV genomes. (A) HFF cells were infected with AD169 at an MOI of 1.5. At 72 hpi, newly synthesized DNA was labeled with EdU (10 μM) for 4 h, and Alexa 488 azide was conjugated by click chemistry. The viral protein pUL44 was visualized by antibody staining. The samples were analyzed by confocal microscopy. (B) Graph representing the percentage of cells positive for EdU only or for both EdU and pUL44 (mock, n = 150; infected, n = 150). (C) HFF cells were infected with AD169 at an MOI of 1.5 and used for accelerated native isolation of proteins on nascent DNA (aniPOND). At 72 hpi, cells were labeled for 4 h and biotinylated by click chemistry for affinity purification. No click reaction served as the negative control.

PcG proteins contribute to efficient viral DNA replication in a catalytically independent manner.

Since PRC1 and -2 have emerged as promising drug targets in cancer therapy, we were interested to test the impact of diverse PRC1/2-inhibitory substances on HCMV DNA replication (Fig. 7A). The inhibitors 3-deazaneplanocin A (DZNep), UNC1999, and GSK126 target the enzymatic activity of PRC2. Wedelolactone (WDL) disturbs the interaction between EED and EZH2, thereby disrupting the functional PRC2, whereas A395 inactivates PRC2 by inhibiting the interaction of EED with trimethylated histone H3 (H3K27me3) (26, 32–35). PTC-209 blocks the expression of BMI1, a core component of PRC1, and PRT4165 targets the enzymatic activity of the above-mentioned complex (36, 37). To avoid any effects on the IE phase of infection (Fig. 4B), we added the substances at 24 hpi and isolated total DNA at 96 hpi to assess viral genome copy numbers per cell (Fig. 7A). This revealed that only the PRC2 inhibitors DZNep and WDL as well as the PRC1 inhibitor PTC-209 were able to compromise HCMV DNA replication comparably to the positive control ganciclovir (GCV), while all other substances had no significant effect (Fig. 7A). In accordance with this finding, only DZNep, WDL, and PTC-209 were able to also negatively affect progression of viral gene expression when added after IE times of infection (Fig. 7B, compare lanes 4, 5, and 10 to lanes 2 and 3) and to decrease the release of viral particles equivalently to the effect of the positive control GCV (Fig. 7C). Interestingly, upon treatment with WDL and PTC-209, the decrease of nascent HCMV particles was even more pronounced compared to the reduction of viral DNA replication (WDL, 3.9% to 41.6%; PTC-209, 0.4% to 8.6%) (compare Fig. 7A and C). This suggests that inhibition of PcG complexes not only leads to inefficient DNA replication but also may result in defective viral DNA, which can be quantified by TaqMan PCR (Fig. 7A) but is not packaged into viral particles (Fig. 7C). To elucidate differences in the modes of action between the antivirally active and nonactive substances, we next analyzed their impacts on PcG protein levels and the conferred histone modifications H3K27me3 and H2Aub (Fig. 7D). Interestingly, while each substance was able to restrain the enzymatic activity of the targeted PRC, DZNep, WDL, and PTC-209 additionally had the capacity to affect complex integrity by reducing the protein levels of essential PRC core components (Fig. 7D, compare lanes 4, 5, and 10 to lanes 2 and 3). Thus, as summarized in Fig. 7E, only substances that negatively affected complex stability, leading to the downregulation of certain PRC core factors, had the capacity to compromise HCMV genome synthesis, while inhibition of PRC1/2’s enzymatic activity alone was insufficient. Taken together, these results lead to the overall conclusion that PRCs exert a noncanonical, enzymatically independent function during HCMV DNA replication.

FIG 7.

Inhibitors depleting PcG proteins impair efficient viral DNA synthesis. (A) HFF cells were infected with AD169 at an MOI of 0.1. At 24 hpi, either DMSO or one of the following substances was added: DZNep (10 μM), wedelolactone (WDL) (20 μM), UNC1999 (300 nM), GSK126 (2.5 μM), A395 (300 nM), PTC-209 (1 μM), and PRT4165 (20 μM). At 96 hpi, total DNA was extracted using the DNeasy blood and tissue kit (Qiagen). Viral genomes were quantified by TaqMan real-time PCR specific for IE1; quantification of the cellular gene albumin was used to calculate HCMV genomes per cell. Values are derived from biological triplicates and represent mean values ± SD. (B) HFF cells were infected with AD169 at an MOI of 0.1 and treated with PRC inhibitors listed above for panel A at 24 hpi. In addition, A395N (300 nM) was used as a negative control for the inhibitor A395. At 96 hpi, samples were harvested and subjected to Western blot analysis. Viral proteins were detected by the indicated antibodies. β-Actin served as a loading control. (C) HFF cells were infected with AD169 at an MOI of 0.1 and treated with the PRC inhibitors listed above for panel A at 24 hpi. Supernatants were harvested at 96 hpi, and contained viral genomes were quantified by HCMV IE1-specific TaqMan real-time PCR. Untreated HFF cells were set to 100%. Values are derived from biological triplicates and represent mean values ± SD. (D) Samples were obtained as described above for panel B. Cellular proteins were detected by the indicated antibodies. β-Actin served as a loading control. (E) Table showing PRC inhibitors utilized and the respective targeted complex. Moreover, the table summarizes the effects of the inhibitors on PRC enzymatic activity and complex stability as well as HCMV DNA replication. For panels A and C, statistical analysis was performed by ordinary one-way ANOVA. n.s., not significant; ***, P ≤ 0.001; ****, P ≤ 0.0001.

PRC inhibitors as potential novel anti-HCMV agents.

Given our observation that HCMV depends on PRC1 and -2 at different steps of the viral life cycle (Fig. 4), we were interested in addressing the impact of PRC inhibitors on HCMV infection in general. Therefore, we applied the PRC2 inhibitors DZNep, WDL, UNC1999, GSK126, and A395 as well as the PRC1 inhibitors PTC-209 and PRT4165 in parallel with infection and investigated the effect on viral spread throughout several HCMV replication cycles by performing a GFP assay and quantifying the GFP signal at 7 dpi (Fig. 8A). The result of this experiment confirmed our previous observation that only the complex-destabilizing inhibitors DZNep, WDL, and PTC-209 were able to inhibit viral spread to an extent equal to that of GCV. In this experimental setting, the inhibitory effect of WDL was weak compared to what is shown in Fig. 7C, which may be an indication of a short half-life of this substance in cell culture medium. Additionally, since the inhibitors solely repressing the enzymatic activity of PRCs did not exhibit antiviral activity (UNC1999, GSK126, A395, and PRT4165) (Fig. 8A), this suggests that the role of PRCs during IE times of infection (Fig. 4) is also enzymatically independent. In order to ensure that the effects that we observed were specific, potential toxicity of the substances used had to be excluded. Therefore, noninfected HFF cells were treated with the PRC1 and PRC2 inhibitors for 24 h, before the cell culture supernatants were analyzed for lactate dehydrogenase (LDH) release. As shown in Fig. 8B, no increased LDH release could be observed, indicating that the inhibitors used were not toxic. Moreover, the influence of the antivirally active substances DZNep, WDL, and PTC-209 on cellular viability was tested over a prolonged incubation period. Noninfected HFF cells were treated with increasing concentrations of the inhibitors. Supernatants were analyzed for LDH release every 24 h for 4 days (Fig. 8C). As shown in Fig. 8C, no substance exceeded a threshold of 20% lysis even when considerably higher concentrations were used (Fig. 8C). In summary, we demonstrate that only inhibitors affecting the complex integrity of PRCs, but not inhibitors of their enzymatic activity alone, are antivirally active over several HCMV replication cycles. The extent of this antiviral activity was comparable to that of the approved HCMV drug ganciclovir, which corroborates the importance of the noncanonical function of PRCs for lytic HCMV replication and demonstrates a possible novel strategy for antiviral treatment.

FIG 8.

PRC inhibitors effectively impair HCMV replication. (A) HFF cells were infected with AD169-GFP at an MOI of 0.002 and treated in parallel with the PRC inhibitors DZNep (10 μM), WDL (30 μM), UNC1999 (300 nM), GSK126 (2.5 μM), A395 (300 nM), PTC-209 (1 μM), and PRT4165 (30 μM). Samples were lysed at 7 days postinfection to perform quantitative GFP fluorometry. Values are derived from biological quadruplicates and represent mean values ± SD. Statistical analysis was performed by ordinary one-way ANOVA. n.s., not significant; *, P ≤ 0.05; ****, P ≤ 0.0001. (B) HFF cells were seeded and treated with the PRC inhibitors listed above for panel A. After 24 h, toxicity was measured using the CytoTox 96 nonradioactive cytotoxicity assay (Promega). Cell lysis was set to 100%. Staurosporine (STP) (5 μM) served as a positive control. Values are derived from biological triplicates and represent mean values ± SD. (C) HFF cells were seeded and treated with different concentrations of GCV, DZNep, WDL, and PTC-209. Supernatants were harvested every 24 h and subjected to the CytoTox 96 nonradioactive cytotoxicity assay (Promega) as described above for panel B.

DISCUSSION

Polycomb repressive complexes (PRCs) are crucial cellular regulators and are therefore modulated by a multitude of viruses for both productive as well as latent infections (8, 38–41). In this study, we dissected the role of the two main complexes formed by polycomb group proteins, PRC1 and PRC2, during lytic HCMV replication. We demonstrated that the core components of both complexes were strongly upregulated, probably resulting from enhanced transcription, which is reflected by increased mRNA as well as protein levels (Fig. 1A and C). Nevertheless, an additional stabilizing effect during infection cannot be excluded. The upregulation was observed at about 24 hpi, independent of the virus strain but dependent on viral gene expression (Fig. 1B and D). The identification of the mechanisms and viral factors responsible for the enhanced PcG gene expression is still pending, but we propose a role of immediate early protein 2, since EZH2 and EED expressions are controlled by the E2F pathway, which is reportedly activated by IE2 (42, 43). Consistent with this assumption, a DNA microarray analysis performed previously identified the PRC2 component EZH2 to be upregulated upon IE2 expression (43). Hence, a similar mechanism for other PcG proteins is conceivable.

In order to elucidate the role of PRC components for lytic HCMV replication, we generated primary human fibroblasts harboring knockdowns of single PRC1 and -2 proteins and found that this resulted in an inhibition of viral replication (Fig. 3). In contrast to the PRC2-mediated repression during HCMV latency, this suggested a proviral function of PcG proteins promoting lytic replication. Strikingly, downregulation of EED and RING1B had the most pronounced effect on viral replication. This may be attributed to the codepletion of the respective proteins upon knockdown of the individual factors (Fig. 3A), which supports the prevailing model of complex integrity being a prerequisite for the stability of polycomb proteins (26). The fact that RING1B as a PRC1 component depends on a PRC2 factor is consistent with the recent finding that EED is a shared component of PRC1 and -2 acting as an exchange factor for recruitment and function of both complexes (17). A similar role for EED may be assumed during the course of HCMV infection.

We were able to identify an active function of PRCs during viral DNA replication since depletion of PcG proteins resulted in a reduced number of nascent viral genomes (Fig. 5A) and decreased delayed early as well as late protein levels, whose expression depends on viral genome amplification (Fig. 4C and D). Consistent with this, all PRC1/2 core components exhibited a changed localization from an even distribution in mock samples to an accumulation in VRCs upon infection (Fig. 2A and B). This is in accordance with a recent study where a similar PcG protein relocalization to VRCs was observed upon murine cytomegalovirus (MCMV) infection, emphasizing a possible conservation of PRC functions for different CMV species (44). Finally, we used aniPOND, a technique of coprecipitation of bound proteins with labeled DNA, which has previously been applied to study the dynamics of cellular factors during herpes simplex virus 1 (HSV-1) replication (45). This revealed a direct physical association of the PRC1 and -2 core components BMI1 and EZH2 with nascent HCMV genomes. In accordance with our findings, several studies recently suggested a potential role of PRCs in the process of cellular DNA replication (23, 24). It was shown that the loss of both complexes led to slower-progressing, asymmetric, or stalled replication forks. While PcG protein binding was detected at cellular replication forks by aniPOND, the underlying mechanisms of how PcG factors contribute to cellular DNA replication are still poorly defined (22–24). While some studies describe PRCs to transfer epigenetic marks to nascent histones for the inheritance of the epigenome, others propose an active contribution to the progression of replication forks (23, 24, 46–48).

In this study, we provide evidence that the promoting function of PcG factors on viral DNA replication occurs in a catalytically independent manner. By investigating the effects of several inhibitory substances on HCMV infection, which target solely the catalytic activity of the respective complexes or in addition target the PcG protein abundance, we show that only PTC-209, DZNep, and WDL, described previously to alter PRC1 and PRC2 protein levels (26, 34, 36), were able to resemble the results obtained in knockdown cells, i.e., reduction of nascent viral genomes and particles as well as inhibition of viral replication (Fig. 7A and C and Fig. 8A). In contrast, substances only reducing the conferred histone modifications had no effect on HCMV infection. In accordance with this, our localization studies demonstrated that H3K27me3 and H2Aub were specifically excluded from VRCs, thereby exhibiting a different localization than all analyzed PRC components (Fig. 2C). In this context, it is noteworthy that while the knockdown of EED and RING1B had a more pronounced effect on viral replication than the depletion of EZH2 and BMI1, the respective histone modifications were less affected (Fig. 3A). Hence, our results highlight a noncanonical, enzymatically independent mode of action of both PRC1 and PRC2 during HCMV infection. These noncatalytic functions have been reported previously in the field of oncology, especially for EZH2 but also for EED, RING1B, and BMI1 (49–55).

The proviral role of PRCs is consistent with previous studies describing that PRC2 facilitates lytic HCMV infection by inhibiting the expression of antiviral factors, thereby creating a favorable cellular environment for viral replication (9, 10). We not only were able to reproduce the impact of PRC2 knockdown on the IE phase of infection but also could expand this finding to the closely related complex PRC1. Furthermore, we demonstrate that a lack of PcG proteins during this phase of infection could be compensated for by higher infectious doses (Fig. 4B), while the DNA replication-related role of PRCs is MOI independent. In contrast to previous studies, we observed that the function of PRC2 during IE times of HCMV infection did not depend on its enzymatic activity (10). This is supported by the observation that substances that solely inhibited the enzymatic activity of PRC components did not affect viral spread when analyzed during several rounds of HCMV replication (Fig. 8A). Also, preincubation of cells with these compounds before infection did not reveal an antiviral effect (data not shown).

In the light of our results, PRCs represent attractive anti-HCMV targets. As summarized in Fig. 9 (right), in the context of latency, PRC2 associates with the MIEP to silence viral lytic gene expression through its methyltransferase activity (7, 8). During lytic HCMV infection (Fig. 9, left), PcG proteins of both PRC1 as well as PRC2 are upregulated and relocalize to viral replication compartments, where they contribute to efficient viral DNA synthesis in a catalytically independent manner. Hence, a pharmacological disruption of PRC2 in latently infected cells, which has been shown to result in a derepression of silenced viral genomes, should subsequently also lead to an abortion of the productive replication cycle of HCMV. This may form the basis for a strategy to target the latent viral reservoir. Taken together, our results significantly broaden the current knowledge about the relevance of PcG proteins for HCMV infection and identify a novel promising target for antiviral treatment.

FIG 9.

Differential roles of PRC1 and -2 during lytic and latent HCMV infection. PRCs play an important role in lytic as well as latent HCMV infection. (Left) During productive HCMV replication, the core components of PRC1 and -2 are heavily upregulated, translocate to replicating viral DNA inside viral replication compartments (VRCs), and promote the efficient synthesis of HCMV genomes. In this regard, both PRCs function in a noncanonical manner independent of their enzymatic activity, as they can be targeted only by compounds that inhibit complex stability. (Right) On the contrary, in cells which support the establishment of HCMV latency, PRC2 is recruited to the viral DNA to silence viral lytic gene expression. Through its methyltransferase activity, PRC2 induces heterochromatinization of HCMV genomes, which can be antagonized by substances that inhibit the enzymatic activity of the complex. Ac, acetylation; M, methylation.

MATERIALS AND METHODS

Oligonucleotides and plasmid constructs.

The oligonucleotide primers used for this study were purchased from Biomers GmbH (Ulm, Germany) or Biomol GmbH (Hamburg, Germany) and are listed in Table 1. In order to construct lentiviral vectors encoding PRC-specific short hairpin RNAs (shRNAs) or a control shRNA, the respective sequences were amplified using the primers 5′shRNA_EZH2 and 3′shRNA_EZH2 for pLVX-shEZH2, 5′shRNA_EED and 3′shRNA_ EED for pLVX-shEED, 5′shRNA_RING1B and 3′shRNA_ RING1B for pLVX-shRING1B, 5′shRNA_BMI1 and 3′shRNA_BMI1 for pLVX-shBMI1, as well as 5′shRNA_Control and 3′shRNA_Control for pLVX-shControl (shC) and inserted into the lentiviral vector pLVX-shRNA1 via BamHI and EcoRI.

TABLE 1.

Oligonucleotides used for plasmid constructions and PCRs

Cells and viruses.

Primary human foreskin fibroblasts (HFFs) were prepared from human foreskin tissue as described previously (56) and cultivated in Eagle’s minimal essential medium (MEM) (Gibco, Thermo Fisher Scientific, Waltham, MA, USA) supplemented with 7.5% fetal calf serum (FCS) (Sigma-Aldrich, St. Louis, MO, USA). HFFs stably expressing small interfering RNAs directed against different PRC components and the respective control cells were maintained in Eagle’s minimal essential medium supplemented with 10% fetal calf serum and 2 μg/ml puromycin. Infection experiments were performed with either the laboratory HCMV strain AD169, the recombinant reporter virus AD169-GFP (57), or the clinical isolate HCMV TB40/E (58). UV-inactivated AD169 was generated by the exposure of wild-type (wt) AD169 to UV light (0.12 J/cm²). All virus stocks were titrated via IE1p72 fluorescence (59). For this purpose, HFF cells were infected with various dilutions of virus stocks and incubated for 24 h. Subsequently, the cells were fixed and stained with the antibody p63-27 directed against IE1p72. IE1-expressing cells were detected, and viral titers were calculated as IE protein-forming units (IEU) per milliliter. Additionally, for AD169-GFP, the 25% tissue culture infection dose (TCID₂₅) was determined as described previously (57), which was equivalent to a multiplicity of infection (MOI) of 0.002.

Generation of stable HFF cells by lentivirus transduction.

For the generation of HFF cells stably expressing shRNAs targeting EZH2, EED, RING1B, and BMI1 as well as the respective control cells expressing shNone (shN) and shControl (shC), the Lenti-X shRNA expression system based on the vector pLVX-shRNA1 (Clontech, Palo Alto, CA, USA) was used. For generation of replication-deficient lentiviruses, HEK293T cells were seeded into 10-cm dishes at a density of 5 × 10⁶ cells/dish and cotransfected with either the empty pLVX-shRNA1 vector (shNone) or vectors containing control shRNA or shRNA against polycomb factors together with packaging plasmids encoding pLP1, pLP2, and pLP/VSV-G using the Lipofectamine 2000 reagent (Invitrogen, Karlsruhe, Germany). Viral supernatants were harvested at 48 h posttransfection, cleared by centrifugation, filtered, and stored at −80°C. HFFs of low passage numbers were incubated for 24 h with lentiviral supernatants in the presence of 7.5 μg/ml Polybrene (Sigma-Aldrich, St. Louis, MO, USA), followed by selection of stably transduced HFF cell populations by the addition of 5 μg/ml puromycin.

Inhibitory substances.

Cycloheximide and phosphonoformic acid (PFA) were purchased from Sigma-Aldrich (St. Louis, MO, USA) and added to the cell culture supernatant in parallel with infection at a concentration of 150 μg/ml or 250 μM, respectively. Ganciclovir from Sigma-Aldrich (St. Louis, MO, USA) was used at 20 μM. DZNep and PRT4165 were ordered from Tocris Bioscience (Bristol, UK) and were applied at 10 μM and 20 μM, respectively. GSK126 was obtained from Cayman Chemical (Ann Arbor, MI, USA) and used at a concentration of 2.5 μM. Wedelolactone, A395, A395N, PTC-209, and UNC1999 were purchased from Sigma-Aldrich (St. Louis, MO, USA) and applied at 20 μM, 300 nM, 1 μM, and 300 nM, respectively. DZNep, A395, A395N, and PFA were dissolved in H₂O, while all other stocks were prepared in dimethyl sulfoxide (DMSO) (Sigma-Aldrich, St. Louis, MO, USA).

Western blot analysis, immunofluorescence, and antibodies.

For Western blot analysis, 3 × 10⁵ wt or stably transduced HFF cells were grown in 6-well plates, harvested and lysed in Roti-Load Laemmli buffer (Roth GmbH, Karlsruhe, Germany) at the respective times postinfection, and boiled at 95°C for 10 min. Proteins were separated on sodium dodecyl sulfate-containing 8% or 12.5% polyacrylamide gels and transferred on nitrocellulose membranes (GE Healthcare, Munich, Germany). Subsequent chemiluminescence detection was performed according to the manufacturer’s instructions (ECL Western blotting detection kit; Amersham Pharmacia Europe, Freiburg, Germany). For immunofluorescence analysis, 1.5 × 10⁵ HFF cells were seeded on coverslips, fixed at 3 days postinfection with 4% paraformaldehyde for 10 min at room temperature, and permeabilized with 1% sodium dodecyl sulfate (SDS) in phosphate-buffered saline without calcium and magnesium (PBSo) for 5 min at room temperature. The samples were incubated with blocking solution containing 2 mg/ml γ-globulin (Cohn fraction II; Sigma-Aldrich, St. Louis, MO, USA) for 30 min, primary antibodies against cellular antigens for 1 h, primary antibody against pUL44 for 30 min, and secondary antibodies for 30 min at 37°C. Finally, the cells were mounted using Vectashield mounting medium containing 4′,6-diamidino-2-phenylindole (DAPI) (Vector Laboratories, Burlingame, CA, USA). The samples were analyzed using a Leica TCS SP5 confocal microscope with 488-nm and 543-nm laser lines, where each channel was scanned separately under image capture conditions, thereby eliminating channel overlap. The images were then exported and processed with Adobe Photoshop CS5. The following monoclonal antibodies were applied for Western blotting and immunofluorescence analysis: anti-UL44 BS510 (kindly provided by B. Plachter, Mainz, Germany), anti-pp65 65-33 (kindly provided by W. Britt, Birmingham, AL, USA), anti-MCP 28-4 (60), anti-pp28 41-18 (61), anti-EZH2 (clone D2C9, catalog number 5246), anti-SUZ12 (clone D39F6, catalog number 3737), anti-RING1B (clone D22F2, catalog number 5694), anti-BMI1 (clone D20B7, catalog number 6964), anti-trimethyl-histone H3 (Lys27) (H3K27me3) (clone C36B11, catalog number 9733), anti-ubiquityl-histone H2A (Lys119) (H2Aub) (clone D27C4, catalog number 8240), anti-H3 (clone D2B12, catalog number 4620) (all purchased from Cell Signaling Technology, Danvers, MA, USA), and β-actin AC-15 (Sigma-Aldrich, St. Louis, MO, USA). The following polyclonal antibodies were applied for Western blotting and immunofluorescence analysis: anti-EED (clone H-300, catalog number sc-28701; Santa Cruz, CA, USA), anti-IE86 (directed against IE2-p86) (62), and anti-pUL84 (62). Horseradish peroxidase-conjugated anti-mouse and anti-rabbit secondary antibodies for Western blot analysis were obtained from Dianova (Hamburg, Germany), and Alexa Fluor 488- and Alexa Fluor 555-conjugated secondary antibodies for indirect immunofluorescence experiments were purchased from Molecular Probes (Karlsruhe, Germany).

RNA isolation and SYBR green reverse transcription-quantitative PCR.

RNAs from 6 × 10⁵ mock-infected or infected HFF cells were isolated in triplicate using TRIzol reagent (Invitrogen, Karlsruhe, Germany) and a Direct-zol RNA miniprep kit (Zymo Research, Freiburg, Germany) according to the manufacturers’ instructions. First-strand cDNA synthesis was performed with 1 μg of total RNA using a Maxima first-strand cDNA synthesis kit (Thermo Fisher Scientific, Waltham, MA, USA). The cDNAs were then used for quantitative real-time SYBR green PCR as described previously (59). For amplification of EED transcripts, the primers EED-fw and EED-rev were used. EZH2 transcripts were amplified using the primers EZH2-fw and EZH2-rev, and SUZ12 transcripts were amplified with the primers SUZ12-fw and SUZ12-rev. For RING1B amplification, RING1B-fw and RING1B-rev were used. For normalization of the quantitative PCR values, the housekeeping gene GAPDH (glyceraldehyde-3-phosphate dehydrogenase) was amplified using the primers huGAPDH_TM_for and huGAPDH_tm-rev.

Quantitative TaqMan real-time PCR.

A total of 3 × 10⁵ infected HFF cells were used in triplicates for TaqMan real-time PCR. For quantification of intracellular viral genomes, total DNA was extracted using the DNeasy blood and tissue kit according to the manufacturer’s instructions (Qiagen, Hilden, Germany). For assessing viral genome copy numbers released from infected HFFs, the cell culture supernatants were collected, centrifuged at 1,500 × g, and treated with proteinase K (Sigma-Aldrich, St. Louis, MO, USA) for 1 h at 56°C, followed by an inactivation step for 5 min at 95°C. Next, 5 μl from each sample was utilized for quantitative real-time PCR (TaqMan PCR), which was conducted as described previously (59). To quantify viral DNA, a sequence within the major immediate early gene region was amplified using the primers 5′CMV and 3′CMV as well as the labeled probe CMV MIE FAM/TAMRA. For analysis of intracellular genomes, quantification of cellular albumin genes was performed in parallel using the primers 5′ Alb and 3′ Alb along with the labeled probe Alb FAM/TAMRA.

GFP reporter assay.

To investigate viral growth and spread, a GFP reporter infection assay was performed as described previously (57). Briefly, 1.8 × 10⁵ wild-type HFF cells or stably transduced HFFs were grown in 12-well plates and infected with the reporter virus HCMV AD169-GFP at an MOI of 0.002. In the respective experiments, compounds were added in parallel with infection. After 7 days, the cells were lysed, followed by automated quantitative GFP fluorometry.

Click chemistry and aniPOND.

For the visualization of replicating DNA, 3 × 10⁵ HFF cells were seeded on coverslips and grown to confluence. At the indicated time points, they were labeled with 10 μM 5-ethynyl-2′-deoxyuridine (EdU) for 4 h, followed by fixation with 4% paraformaldehyde for 10 min and a quenching step using 50 mM glycine and 50 mM NH₄Cl in PBSo. The cells were permeabilized with PBSo containing 0.2% Triton X-100 (Sigma-Aldrich, St. Louis, MO, USA) on ice for 20 min before antibody labeling was conducted as described above. The conjugation of Alexa Fluor 488 azide to labeled DNA was performed by click chemistry using 10 mM sodium ascorbate and 1 mM CuSO₄ for 2 h at room temperature. Further immunofluorescence analysis was performed as described above. For accelerated native isolation of proteins on nascent DNA (aniPOND), four 175-cm² tissue culture dishes of confluent HFF cells (∼1.2 × 10⁷ cells per sample) were infected with AD169. At 3 days postinfection, replicating DNA was labeled with 10 μM EdU for 4 h. Subsequent lysis, biotinylation, and precipitation of labeled DNA were performed as described previously (22). Briefly, after the incubation period, cell nuclei were harvested using an NP-40-containing nucleus extraction buffer, which combined the termination of EdU labeling and the lysis of cells. After washing, the click reaction was performed by incubation of the nuclei together with 25 μM biotin azide, 10 mM sodium ascorbate, and 2 mM CuSO₄ at 4°C for 1 h, followed by another washing step. Replacement of biotin azide with DMSO served as a negative control. Lysis of the nuclei was achieved by two short and one extensive sonication step. Input controls were taken before biotinylated DNA was captured by incubation with Dynabeads MyOne streptavidin T1 (Thermo Fisher Scientific, Waltham, MA, USA) for 16 h. After extensive washing, the bound DNA was resolved with Roti-Load Laemmli buffer (Roth GmbH, Karlsruhe, Germany) by boiling at 95°C for 15 min. The samples were subjected to Western blot analysis together with the input controls.

Cytotoxicity assay.

The determination of cytotoxic effects was performed using the CytoTox 96 nonradioactive cytotoxicity assay (Promega, Madison, WI, USA) according to the manufacturer’s instructions. Therefore, 1.35 × 10⁴ HFF cells were seeded in 96-well plates and treated with the indicated substances for 24 h. The cell culture supernatant was analyzed for the release of lactate dehydrogenase (LDH). Alternatively, 8.1 × 10⁴ HFF cells were seeded in 24-well plates, and the indicated substances were added. Cell culture supernatants were collected and analyzed every 24 h. Total cell lysates served as positive controls for maximal LDH release.