The Human Cytomegalovirus Lytic Cycle Is Induced by 1,25-Dihydroxyvitamin D3 in Peripheral Blood Monocytes and in the THP-1 Monocytic Cell Line

Abstract

Human cytomegalovirus (HCMV) resides in a latent form in hematopoietic progenitors and undifferentiated cells within the myeloid lineage. Maturation and differentiation along the myeloid lineage triggers lytic replication. Here, we used peripheral blood monocytes and the monocytic cell line THP-1 to investigate the effects of 1,25-dihydroxyvitamin D3 on HCMV replication. Interestingly, 1,25-dihydroxyvitamin D3 induces lytic replication marked by upregulation of HCMV gene expression and production of infectious virus. Moreover, we demonstrate that the effects of 1,25-dihydroxyvitamin D3 correlate with maturation/differentiation of the monocytes and not by directly stimulating the MIEP. These results are somewhat surprising as 1,25-dihydroxyvitamin D3 typically boosts immunity to bacteria and viruses rather than driving the infectious life cycle as it does for HCMV. Defining the signaling pathways kindled by 1,25-dihydroxyvitamin D3 will lead to a better understanding of the underlying molecular mechanisms that determine the fate of HCMV once it infects cells in the myeloid lineage.

Introduction

Human cytomegalovirus (HCMV) is a β-herpesvirus that spreads broadly throughout the human population (Sinzger, Digel, and Jahn, 2008). In general, about 50-70% of people are serologically positive for HCMV worldwide (Bate, Dollard, and Cannon, 2010). Although in immunocompetent individuals HCMV infection is typically asymptomatic, in the case of congenital infection, the virus can cause severe neurological sequelae such as deafness and developmental defects following infection of the fetus (Grosse, Ross, and Dollard, 2008; Johnson and Anderson, 2014). In immunocompromised individuals, including those with HIV/AIDS or those receiving organ transplants, HCMV can cause devastating morbidity and mortality including pneumonia, retinitis, and transplant rejection (Ljungman, Hakki, and Boeckh, 2011; Paya et al., 2004; Yen et al., 2015). Moreover, many studies have shown that HCMV can be associated with chronic diseases such as atherosclerosis and hypertension, cancer, and autoimmune disease (Dziurzynski et al., 2012; Li et al., 2011; Streblow, Orloff, and Nelson, 2001; Varani et al., 2009). Therefore, understanding the biology of HCMV infection is both clinically relevant and intensively studied with regards to potential pharmacological intervention. Like other herpesviruses, HCMV can establish latency in the human body, thus making the eradication of the virus from infected individual a difficult task (Grinde, 2013; White, Suzanne Beard, and Barton, 2012). The cellular reservoirs for HCMV latency include hematopoietic stem cells, common myeloid progenitor cells, and monocytes (Bego and St Jeor, 2006; Goodrum et al., 2002; Sinclair, 2008; Taylor-Wiedeman et al., 1991). Deciphering the mechanisms that regulate the latent/lytic switch in HCMV infected cells could lead to the identification of novel therapeutics that could be used to regulate latency. Previous studies have indicated that both viral and cellular factors are involved in the control of latent and lytic cycles in myeloid progenitors and monocytes (Chan, Nogalski, and Yurochko, 2012; Goodrum et al., 2007; Kew et al., 2014; Keyes et al., 2012b; O'Connor, Vanicek, and Murphy, 2014; Smith et al., 2004; Stevenson et al., 2014); however, the molecular mechanisms remain unresolved, and it is highly probable that there are numerous cellular and viral regulatory factors that have yet to be identified. In light of this, further investigation of the mechanisms and factors that influence the switch between HCMV latency and lytic replication in clinically relevant myeloid cell types is needed.

It is known that the developmental maturation of monocytes into macrophages and dendritic cells can reactivate HCMV from latency leading to the production of new infectious virus (Chan et al., 2008; Reeves and Sinclair, 2013; Smith et al., 2004; Soderberg-Naucler, Fish, and Nelson, 1997; Soderberg-Naucler et al., 2001; Stevenson et al., 2014). In addition, there are various extracellular stimuli (i.e. PMA) that can trigger monocyte to macrophage differentiation (Greenberger, Newburger, and Sakakeeny, 1980; Hemmi and Breitman, 1985; Huber et al., 2014; Naito, 2008; Nakamura et al., 1986; Netea et al., 2008) and some of these stimuli have also been shown to directly induce the HCMV immediate-early gene promoter, which is essential for induction of the HCMV lytic cycle (Ghazal et al., 1992; Kline et al., 1998; Stein et al., 1993). Since the activities of these stimuli appear to be multi-factorial, it is difficult to determine if the major influence of these stimuli on lytic replication is induction of the IE promoter, promotion of cellular maturation/differentiation or a combination of both activities. THP-1 cells are a monocytic cell line that is commonly used in combination with primary blood derived monocytes to study the interaction between HCMV and myeloid cells and gain insight into the latent/lytic switch (Saffert, Penkert, and Kalejta, 2010; Van Damme et al., 2014). It is well known that HCMV enters latency or a quiescent state in undifferentiated THP-1, and the virus typically enters into the lytic cycle after it infects phorbol 12-myristate 13-acetate (PMA) treated THP-1 cells (Qin, Penkert, and Kalejta, 2013; Weinshenker, Wilton, and Rice, 1988). As a consequence, PMA is a reagent of choice used to promote myeloid differentiation in studies aimed at inducing lytic replication in in vitro systems. However, PMA is a synthetic compound resembling diacylglycerol (DAG) that is capable of activating a broad range of cell signaling pathways (Castagna et al., 1982; Niedel, Kuhn, and Vandenbark, 1983; Swindle, Hunt, and Coleman, 2002). In this research we sought to identify additional physiologically relevant compounds that could trigger both monocyte differentiation and HCMV lytic infection. Vitamin D3 is a hormone that is produced by the human body and acquired in a supplemental fashion through diet (Baeke et al., 2010; Holick, 2003; Lamberg-Allardt, 2006). The most well-known effects of vitamin D3 and its active metabolite 1,25-dihydroxyvitamin D3 are to regulate homeostasis of calcium and phosphorus and promote bone development through interaction with the vitamin D receptor (VDR), a member of the nuclear receptor family of transcription factors (Goltzman, Hendy, and White, 2014; Kannan and Lim, 2014). Interestingly, blood leukocytes robustly express the VDR and results of studies performed in vitro in human myeloid cell lines and ex vivo in murine bone marrow cells have demonstrated that 1,25-dihydroxyvitamin D3 has the ability to induce monocyte-macrophage differentiation (Gemelli et al., 2008; Hmama et al., 1999; Lagishetty, Liu, and Hewison, 2011; Liu et al., 2006; O'Kelly et al., 2002, Bhalla, 1983 #83; Provvedini et al., 1983). It is therefore not surprising that 1,25-dihydroxyvitamin D3 has been demonstrated to exhibit antibacterial and antiviral effects (Korf, Decallonne, and Mathieu, 2014; Luong and Nguyen, 2011; Maxwell, Carbone, and Wood, 2012; Spector, 2011). The importance of 1,25-dihydroxyvitamin D3 in regulation of immune system function has been further highlighted by studies which suggest that 1,25-dihydroxyvitamin D3 or synthetic analogues of 1,25-dihydroxyvitamin D3 could be used as potent candidates for the treatment for autoimmune diseases, infectious diseases and anticancer therapies (Salomon et al., 2014; Yuzefpolskiy et al., 2014; Zhang, Wan, and Liu, 2013). Nonetheless, the effect of 1,25-dihydroxyvitamin D3 on HCMV replication in monocytes and macrophages remains unknown. Therefore, we explored the possibility that peripheral blood monocytes and THP-1 cells could be used to determine the effect of 1,25-dihydroxyvitamin D3 on HCMV replication in myeloid cells. According to the results of previous studies, 1,25-dihydroxyvitamin D3 treatment induces THP-1 cells to differentiate into mature monocytes, with high CD14 expression (Daigneault et al., 2010; Hmama et al., 1999; Schwende et al., 1996) and therefore we also hypothesized that we also could use this model to study HCMV replication in 1,25-dihydroxyvitamin D3 treated cells that are in the transition from the promonocytic to macrophage stages.

Interestingly, we found that the HCMV lytic phase can be induced in 1,25-dihydroxyvitamin D3 treated primary monocytes and in THP-1 cells with infectious virus being produced by these cells. In contrast to PMA treated cells, 1,25-dihydroxyvitamin D3 does not have a direct effect on the HCMV immediate-early gene promoter in reporter gene assays suggesting that the predominant effect of 1,25-dihydroxyvitamin D3 is to drive differentiation and not necessarily to directly stimulate IE promoter activity. When 1,25-dihydroxyvitamin D3 is combined with PMA to differentiate THP-1 cells, no additive effect on HCMV replication is observed. These results demonstrate that 1,25-dihydroxyvitamin D3 induces a set of differentiation related signaling pathways that creates a favorable cellular milieu for HCMV lytic infection. Moreover, our results suggest that clinical/dietary supplementation with vitamin D3 could be problematic in patients susceptible to reactivation-based HCMV disease.

Materials and Methods

General Reagents

1,25-dihydroxyvitamin D3 and phorbol 12-myristate 13-acetate (PMA) were purchased from Sigma-Aldrich. APC conjugated anti-human CD14, anti-human CD11b, anti-human CD54, anti-human CD36 antibodies and PE conjugated anti-mouse IgG1 antibodies were obtained from eBioscience. Anti-CMV IE1/IE2 antibody (mAb810) and an Alexa Fluor® 488 conjugated version of mAB810 were purchased from Millipore. Anti-CMV UL44 antibody (mAb 25G11, IgG1 isotype) was a kind gift of John Shanley, and anti-CMV pp65 antibody was obtained from Virusys Corporation.

Cell culture and differentiation of THP-1 cells

THP-1 cells were maintained in RPMI-1640 (Roswell Park Memorial Institute Institute-1640) supplemented with 10% FBS, 100 IU/ml penicillin and 100 μg/ml streptomycin at 37°C in 5% CO₂. THP-1 cells were passaged every 3 days to maintain the cell density between 0.2×10⁶ and 1×10⁶ cells/ml. Human foreskin fibroblasts (HFFs) were maintained in DMEM (Dulbecco Modified Eagle's Medium) supplemented with 10% Fetal Clone III serum, 100 IU/ml penicillin and 100μg/ml streptomycin at 37°C in 5% CO₂. THP-1 cells were treated with 80nM PMA or 100nM 1,25-dihydroxyvitamin D3 for three days to induce cellular maturation/differentiation.

Propagation and purification of virus

The HCMV TB40E-mCherry(3×FLAGUS28) virus was generously provided by Dr. Christine O' Connor from the Cleveland Clinic (Miller et al., 2012; O'Connor and Shenk, 2011). This virus was characterized and demonstrated to grow with similar kinetics and to similar titers as does HCMV TB40E. To propagate virus, HFFs were infected with TB40E viruses at an m.o.i. of 0.04. Viral supernatant was harvested at days 9, 11, and 13 post-infection. Cell Culture supernatant containing virus was centrifuged at 1800×g for 3 minutes at 21°C to remove cellular debris. The clarified supernatant was overlayed on a 20% D-sorbitol/1mM MgCl₂ cushion and subjected to ultracentrifugation at 24,000 rpm for 1hr at 21°C. Supernatant was decanted, and the viral pellet was resuspended in RPMI-1640 culture media. Viral supernatant was aliquoted and stored at 80°C.

Isolation of monocytes from peripheral blood of normal donors

Blood was diluted 2 fold with Dulbecco's Phosphate-Buffered Saline (DPBS) containing 2 mM EDTA. Diluted blood was carefully layered over 15 ml of Ficoll-paque^® PLUS in a 50 ml conical tube. Conical tubes were centrifuged at 400×g for 30-40 minutes at 20°C in a swinging bucket rotor without brake. After centrifuge, the upper layer was aspirated, leaving buffy coat containing the mononuclear cell layer undisturbed at interphase. The buffy coat was transfer to clean tube, and fresh DPBS with 2 mM EDTA was added to fill the tubes. The cells were centrifuged at 300xg for 10 minutes at 20°C. Then supernatant was carefully removed. In order to remove platelets, cells were resuspended in 50 ml of DPBS with 2 mM EDTA, and centrifuged at 200×g for 10-15 minutes at 20°C. The supernatant was removed afterward. The wash step was repeated once to further deplete platelets. Cells were resuspended in 80 μl of buffer (DPBS containing 0.5% BSA and 2 mM EDTA) per 10⁷ cells. 20 μl of CD14 MicroBeads (Miltenyi Biotec) was added per 10⁷ cells, and cells were incubated in the cold room (2-8°C) for 30 minutes. After incubation, cells were washed by adding 1-2 ml of buffer (DPBS containing 0.5% BSA and 2 mM EDTA) per 10⁷ cells, and centrifuged at 300×g for 10 minutes. 10⁸ cells were resuspended in 500 μl of buffer, and passed through LS Column (Miltenyi Biotec). The column was washed 3 times with buffer. The column was removed from MACS Separator (Miltenyi Biotec), and cells were flushed out by firmly pushing the plunger into the column.

Culture and infection of primary peripheral blood derived monocytes

CD14⁺ monocytes isolated as described above were resuspended in RPMI-1640 supplemented with 10% FBS, 100 IU/ml penicillin and 100 μg/ml streptomycin. Cells were then cultured in 1,25-dihydroxyvitamin D3 (100 nM), PMA (80 nM) or the appropriate vehicle control (EtOH or DMSO) for 2 days at 37°C in 5% CO₂. After 2 days of culture in 1,25-dihydroxyvitamin D3, PMA or vehicle, viral supernatant was added at a MOI of 10, and cells were incubated overnight. After overnight incubation with virus, cells treated with vehicle or 1,25-dihydroxyvitamin D3 were pelleted by centrifugation, viral inoculums were removed, and cells were treated with 1× trypsin for 5 minutes to remove attached but un-internalized virions (O'Connor and Murphy, 2012). Cells were then resuspended in fresh RPMI supplemented as described above and cultured for 4-6 days. In the case of the PMA treated cells that were adhered to the culture plates, viral inoculums were aspirated, cells were washed thoroughly with DPBS, fed with fresh RPMI and supplements and cultured for 4-6 days.

HCMV infection of THP-1 cells

THP-1 cells were infected with HCMV TB40E-mCherry (3×FLAGUS28) at MOIs as indicated in the figure legends. After the viral supernatant was added, cells were centrifuged at 21°C and 1000×g for 30 minutes to enhance infectivity. After overnight culture, cells were spun down and the inoculums were removed. Cells were incubated in 1× trypsin for 5 minutes to remove attached but un-internalized virions (O'Connor and Murphy, 2012). The trypsin reactions were neutralized by adding equal volumes of fresh culture media. The supernatant is aspirated, and cells were resuspended in fresh culture media. Because the PMA-differentiated cells firmly adhere to the plates, the centrifuge step is omitted from infection protocol. After overnight culture, culture media were removed and cells were washed with 1× DPBS, and fresh media were added to the cells.

Western blot analysis of HCMV gene expression

1×10⁶ infected THP-1 cells were lysed in NP-40 cell lysis buffer (50 mM HEPES pH7.4, 0.5% NP-40, 250 mM NaCl, 20% glycerol, 2 mM EDTA, 100 μM Sodium Orthovanadate, 1 mM Sodium Fluoride, 1× complete Mini protease inhibitor). Cell lysates were sonicated for 20 seconds on level 1 using a Sonic Dismembrator Model 100 (Fisher Scientific). Protein concentrations for each lysate sample were determined using the Bio-Rad protein assay reagent. Lysates were mixed with Laemmli sample buffer, and heated at 100° for 10 minutes. 30 μg of protein was loaded into each lane for electrophoresis. Proteins were transferred to nitrocellulose membranes using a semi-dry transfer apparatus (Continental Lab Products). Membranes were blocked with 5% non-fat dried milk for 1 hour and membranes were incubated with primary antibody at 4° overnight. Membranes were washed 3 times with Tris buffered saline with Tween-20 (TBST) and then incubated with secondary antibody for 2 hours. Membranes were washed with TBST 3 times and subjected to antibody detection using the SuperSignal West Pico chemiluminescent substrate (Thermo Scientific). Luminescence emitted from the membranes was detected by classic blue autoradiography film BX. Films were developed by Kodak min-R mammography processor.

MIEP reporter gene construction

The HCMV immediate-early gene promoter (bp 52586-53162 in GenBank entry AC146907.1) was PCR amplified from FIX-BAC bacmid DNA using the following primers. HCMV MIEP promoter forward primer: 5′-TAACCCGGGTAGTAATCAATTACGGGG-3′, HCMV MIEP promoter reverse primer: 5′-TCGAGATCTCTGACGGTTCACTAAACG-3′. PCR amplification was performed for 30 cycles, consisting of denaturation at 94°C for 30 seconds, annealing at 55°C for 30 seconds, and extension at 72°C for 30 seconds. The HCMV MIEP promoter fragment was cloned into the XmaI and BglII sites of pGL3 and sequenced to confirm the identity of the MIEP fragment (Genewiz, Inc).

Luciferase assays

2×10⁵ THP-1 cells were plated per well in 24-well plates and cultured overnight. For each transfection, 410 ng pcDNA3, 60 ng pGL3-HCMV MIEP, 30 ng phRGTK-renilla and 1.5 μl TransIT-2020 was diluted into 50 μl RPMI-1640 and incubated for 15 minutes. The incubated transfection reagent was then added into each well. 4 hours after transfection, vehicle (ethanol or DMSO), 1,25-dihydroxyvitamin D3 (100 nM), or PMA (80 nM) was added to the designated wells and the cells were incubated for a further 24 hours. Cells were lysed in 200 μl of 1× passive lysis buffer and 10 μl of the cell lysate was used in luciferase assay reactions. 50 μl of Firefly-luciferase substrate was added to cell lysate and luciferase activity was measured on a Glomax 20/20 luminometer (Promega). 50 μl Stop & Glow solution was added to each reaction, and luminescence of the control reporter Renilla-Luciferase was measured. The Firefly-luciferase reading of vehicle control (ethanol) was divided by the Renilla-Luciferase reading of vehicle control and that value was defined as 1. The fold changes were then determined by dividing the luciferase to renilla ratios of the experimental conditions to the ratio of vehicle control.

Infectious center assays

1×10⁵ HFF cells were plated into wells of 12-well plates and cultured overnight. 1×10⁴ THP-1 cells were harvested at six days post-infection and co-cultured with HFFs for 2 days. Culture media was then removed and the plates were washed twice with 1× PBS. Cell monolayers were covered with overlay media (a 1:1 mixture of 1.5% carboxymethyl cellulose (Sigma), and 2× MEM supplemented with 20% FCIII serum, nonessential amino acids, and penicillin-streptomycin) and incubated for another 8 days to allow for plaque development. Cells were fixed with methanol, stained with 10% Geimsa (Sigma) and plaques were counted using a dissecting microscope.

Plaque assays

After infection, culture media from each cell sample was harvested on the indicated days, and 10 μl of culture media was added to HFF monolayers plated in 12-well plates the previous day. Virus was adsorbed to HFF monolayers for 3 hours, culture media were removed, and covered with overlay media (a 1:1 mixture of 1.5% carboxymethyl cellulose (Sigma), and 2× MEM supplemented with 20% FCIII serum, nonessential amino acids, and penicillin-streptomycin) and incubated for another 8 days to allow for plaque development. Cells were fixed with methanol, stained with 10% Geimsa (Sigma) and plaques were counted using a dissecting microscope.

Flow cytometry

For cell surface marker analyses, cells were harvested and resuspended in 50μl of a 0.5% BSA/DPBS solution containing 1:200 dilution of the appropriate APC-conjugated antibody (CD11b, CD14, etc). Cells were incubated at room temperature for 1 hour. Cells were washed with 0.5% BSA/DPBS solution, resuspended in fresh DPBS and analyzed by flow cytometry on a BD FACSCalibur. For HCMV IE protein staining, cells were harvested, resuspended in 100 μl DPBS and an equal volume of 70% ice cold EtOH was added to fix the cells. Fixed cells were washed and permeabilized with 0.5% BSA/DPBS solution containing 0.5% tween-20. Cells were then resuspended in 0.5% BSA/DPBS solution containing 1:200 dilution of Alexa488 conjugated anti-HCMV IE antibody, and incubated at room temperature for 1 hour. Stained cells were washed with 500μl 0.5% BSA/DPBS solution containing 0.5% tween-20. For UL44 and IE co-staining, cells were fixed and permeabilized as above and then resuspended in 0.5% BSA/DPBS solution containing 1:10 dilution anti-HCMV UL44 antibody. Cells were incubated for 1 hour at room temperature, washed as described above and then incubated for 1 hour in 0.5% BSA/DPBS solution containing 1:250 dilution of PE-conjugated anti-mouse IgG1 to label the UL44 primary antibody. UL44 stained cells were then washed and stained for IE proteins as described above. After staining, cells were resuspended in DPBS and analyzed by flow cytometry.

Semi-quantitative PCR for viral DNA copy number

2×10⁵ THP-1 cells were harvested on days 1 and 6 post-infection, and lysed in 100 μl DNA lysis buffer (Kondo, Kaneshima, and Mocarski, 1994) containing 20 μg of Proteinase K at 55°C overnight. Proteinase K activity wasstopped by incubating DNA lysates at 100°C for 15 minutes and DNAs were used for semi-quantitative PCR. Primers for HCMV IE amplification: HCMV IE forward 5′-ATGGAGTCCTCTGCCAAGAGAAAGATGGAC-3′, HCMV IE reverse 5′-CAATACACTTCATCTCCTCGAAAGG -3′ (Bego et al., 2005). Primers used for GAPDHamplification: GAPDH forward 5′-GAGCCAAAAGGGTCATC-3′, GAPDH reverse primer 5′-GTGGTCATGAGTCCTTC-3′ (Juckem et al., 2008). PCR amplification was performed for 30cycles, consisting of denaturation at 94°C for 30 seconds, annealing at 55°C for 30 seconds,and extension at 72° for 30 seconds. Band intensities were measured by NIH Image software and calculated as a ratio of HCMV IE DNA over cellular GAPDH DNA.

Chromatin Immunoprecipitation

5×10⁶ cells were harvested at 2 days post-infection and fixed in PBS containing 1% formaldehyde for 10 minutes at room temperature. 2.5 M glycine was added to stop the fixation reaction, and cells were washed in ice cold PBS. Cells were lysed in cell lysis buffer (0.5 mM PIPES Ph8, 85 mM KCl, 0.5% NP40, 1× protein inhibitor cocktail). Nuclei were pelleted by centrifuge and resuspended in 500 μl nuclear lysis buffer (50 mM Tris-HCl Ph8, 10 mM EDTA,1% SDS,1× protein inhibitor cocktail). The lysates were sonicated for six cycles (30 seconds on/ 30 seconds off) using program 3 on a Sonic Dismembrator Model 100 (Fisher Scientific). Lysates were then diluted 5 fold in IP dilution buffer (0.01% SDS, 1.1% Triton-X 100, 1.2 mM EDTA, 16.7 mM Tris-HCl pH8, 167 mM NaCl, 1× protein inhibitor cocktail) and pre-cleared with 30μl sepharose beads for 2 hours at 4°C. 3 μg of anti-tri-methylated histone 3 antibody (Millipore) was added into each reaction and rotated overnight at 4 °. Protein A/G beads were added to each reaction and incubated at 4°for 3 hours to capture primary antibody. Protein A/G beads was pelleted by centrifugation and washed with 1× dialysis buffer (2 mM EDTA, 50 mM Tris-HCl pH8, 0.2% Sarkosyl, 1× protein inhibitor cocktail) 3 times. Bound chromatin fragments were eluted using elution buffer (50 mM NaHCO₃, 1% SDS). After elution, RNase A and NaCl were added to make final concentration of 0.083 mg/ml and 0.2 M respectively and the solution was incubated at 65°C overnight to reverse cross-links. 34 μg of Proteinase K was added to digest proteins at 45°C for 2 hours. Primers for amplification of MIEP enhancer: Forward primer: 5′-TTGGGCATACGCGATATCTG-3′. Reverse primer: 5′-GCCTCATATCGTCTGTCACC-3′ (Abraham and Kulesza, 2013). The DNA fragments were recovered using a Fermentus gel extraction kit and 20 ng of immunoprecipitated DNA was used for PCR amplification. PCR reaction conditions are the same as mentioned above except were performed for 36 cycles. The signal from chromatin immunoprecipitation samples were normalized to signal from respective input samples.

Results

1,25-dihydroxyvitamin D3 promotes HCMV replication in primary monocytes and THP-1 cells

Vitamin D3 is a natural hormone that is produced by human body and typically supplemented through diet (Baeke et al., 2010; Holick, 2003; Lamberg-Allardt, 2006). In addition to the regulation of the homeostasis of calcium and phosphorus (Garabedian and Ulmann, 1979; Goltzman, Hendy, and White, 2014), vitamin D3 has been shown to play multiple roles in immune responses including modulating T cell and B cell activity (Terrier et al., 2012), promoting monocyte-macrophage differentiation (Pan et al., 1997; Takahashi, Nakamura, and Iho, 1997), stimulating the anti-bacterial and anti-viral effects of macrophages (Campbell and Spector, 2012; Verway et al., 2013), and driving lineage commitment of hematopoietic progenitor cells (Bunce, Brown, and Hewison, 1997). Vitamin D3 like other endocrine hormones is carried by the circulatory system to various tissues (Baeke et al., 2010), and research has also shown that many cells within the immune system have the enzyme that can convert Vitamin D3 into its active form, 1,25-dihydroxyvitamin D3 (Ooi et al., 2014; Shahijanian et al., 2014; Stoffels et al., 2006). Therefore, vitamin D3 can execute its effects on a wide variety of cells in either an endocrine or paracrine fashion (Hewison, 2012). HCMV is a β-herpesvirus which can infect myeloid cells and establish latent and/or lytic infections within cells of this lineage (Sinclair, 2010). Although studies have shown that allogeneic stimulation or stimulation with cytokines like TNF-α can stimulate HCMV IE promoter activity and drive lytic replication in myeloid cells (Prosch et al., 1995; Soderberg-Naucler, Fish, and Nelson, 1997; Stein et al., 1993), the effect of 1,25-dihydroxyvitamin D3 on HCMV replication in myeloid cells remains unexplored. To determine if 1,25-dihydroxyvitamin D3 may influence HCMV lytic replication in myeloid cells, we examined the effect of 1,25-dihydroxyvitamin D3 on the ability of CD14 positive peripheral blood monocytes to support lytic replication. Peripheral blood mononuclear cells (PBMCs) from healthy anonymous donors were isolated by Ficoll-paque and CD14 positive monocytes were subsequently isolated from PBMCs using CD14 magnetic beads (Miltenyi). Monocytes were treated with 1,25-dihydroxyvitamin D3 (100 nM) for 2 days and infected with HCMV TB40E at an MOI of 10. On days 4 and 6 post-infection, cells were harvested and subsequently co-cultured with human foreskin fibroblasts (infectious center assays) to determine whether the infected monocytes were producing infectious virus. Interestingly, monocytes treated with 1,25-dihydroxyvitamin D3 exhibited a 5-8 fold increase in infectious centers over cells treated with the vehicle control ethanol (Figure 1A). These data indicate that 1,25-dihydroxyvitamin D3 treatment can create a milieu in blood monocytes that more efficiently supports HCMV virus production. HCMV replication in primary monocytes treated with the phorbol ester phorbal 12-myristate 13-acetate (PMA) was similarly examined (Figure 1B). PMA is a well-established inducer of HCMV replication in a number of systems (Qin, Penkert, and Kalejta, 2013; Weinshenker, Wilton, and Rice, 1988). PMA treated cells exhibited a 15-30 fold increase in infectious centers over cells treated with the vehicle control DMSO (Figure 1B). Thus, for comparison, while it is readily apparent that 1,25-dihydroxyvitamin D3 is a robust inducer of HCMV replication in monocytes, the level of replication achieved is not as strong as that achieved by the phorbol ester, consistent with what might be expected for a natural compound like 1,25-dihydroxyvitamin D3.

Figure 1. 1,25-dihydroxyvitamin D3 promotes HCMV replication in primary peripheral blood derived monocytes.

Monocytes were treated with 100 nM 1,25-dihydroxyvitamin D3 (A) or 80 nM PMA (B) for 2 days and then infected with HCMV TB40E at a MOI of 10. On 4 days and 6 days post-infection, cells were harvested and co-cultured with HFF fibroblasts for 2 days. After 2 days of co-culture, fibroblast monolayers were overlayed with CMC/MEM and incubated for 8 days to allow for plaque development. The data represent 4-8 independent experiments performed in duplicate. VitD3, 1,25-dihydoxyvitamin D3. * p<0.05, ** p<0.01.

We then sought a model system that could be utilized to provide a more mechanistic explanation for this finding. THP-1, an established monocytic cell line and model system frequently used in HCMV studies (Keyes et al., 2012a; Saffert, Penkert, and Kalejta, 2010), was then deployed to further explore the effects of 1,25-dihydroxyvitamin D3 on HCMV lytic replication. Since it is well established that PMA can drive HCMV production in THP-1 cells, we again used PMA as a control in these experiments (Weinshenker, Wilton, and Rice, 1988). THP-1 cells were treated with vehicle (ethanol), 1,25-dihydroxyvitamin D3 (100 nM) or PMA (80 nM) for 3 days before infection. Cells were then infected with HCMV TB40E at an MOI of 10. On day 6 post-infection, THP-1 cells were co-cultured with HFFs in infectious center assays. In THP-1 cells treated with the ethanol control, only a very low number of plaques were detected in infectious center assays, which indicate that THP-1 cells rarely support lytic phase replication after HCMV infection (Figure 2). In PMA treated THP-1 cells, there was a 40- fold increase in the number of plaques arising in infectious center assays supporting early studies that reported the induction of lytic phase replication by PMA treatment. Importantly, 1,25-dihydroxyvitamin D3 treatment of THP-1 cells resulted in a 10-fold increase in the number of plaques arising in infectious center assays. We repeated this experiment and examined infectious center production at days 6 and 8 post-infection and obtained similar results, indicating that the difference in lytic replication observed between 1,25-dihydroxyvitamin D3 and PMA treated cells is not simply the result of a delay in virus replication in the 1,25-dihydroxyvitamin D3 treated cells (data not shown). Therefore, since 1,25-dihydroxyvitamin D3 can promote lytic virus production in both peripheral blood monocytes and in the monocytic cell line THP-1, we conclude that THP-1 cells would provide a viable model to recapitulate and further explore the effects of 1,25-dihydroxyvitamin D3 on HCMV replication in myeloid cells.

Figure 2. 1,25-dihydroxyvitamin D3 promotes HCMV replication in the THP-1 monocytic cell line.

THP-1 monocytes were treated with 1,25-dihydroxyvitamin D3 (100 nM) or PMA (80 nM) for 3 days and then infected with HCMV TB40E at a MOI of 10. On day 6 post-infection, cells were harvested and co-cultured with HFF fibroblasts for 2 days. After 2 days of co-culture, fibroblast monolayers were overlayed with CMC/MEM and incubated for 8 days to allow for plaque development. The data represent five independent experiments performed in duplicate. VitD3, 1,25-dihydoxyvitamin D3. ** p<0.01, *** p<0.001.

1,25-dihydroxyvitamin D3 treatment does not influence the ability of HCMV to establish an initial infection in monocytes

Since we found that 1,25-dihydroxyvitamin D3 treatment can dramatically increase the number of plaques that arise from THP-1 cells in co-culture infectious center assays, we wanted to determine whether this difference could be the result of increased infectivity or entry of HCMV virions into 1,25-dihydroxyvitamin D3 treated cells. We used semi-quantitative PCR to examine viral genome copy number in cells at various time points post-infection (Figure 3). If 1,25-dihydroxyvitamin D3 leads to increased infectivity of the monocytes, we would expect to see increased viral DNA levels at early time points post-infection. However, at day 1 post-infection, cells treated with 1,25-dihydroxyvitamin D3 or PMA exhibited similar viral copy numbers to that of vehicle control cells. The results are depicted in Figure 3A and quantitative results from six independent experiments are shown in Figure 3B. This result suggested that HCMV infects control and 1,25-dihydroxyvitamin D3 treated cells with equivalent efficiency. At day 6 post-infection, the PCR signal for viral genomes in control THP-1 cells declined while the signal from 1,25-dihydroxyvitamin D3- and PMA-treated cells was maintained or increased, consistent with the conclusion that 1,25-dihydroxyvitamin D3 and PMA treated cells are supporting lytic HCMV replication.

Figure 3. 1,25-dihydroxyvitamin D3 treatment does not influence the ability of HCMV to establish an initial infection.

(A) THP-1 monocytes were treated 1,25-dihydroxyvitamin D3 (100 nM) or PMA (80 nM) for 3 days and then infected with HCMV TB40E at a MOI of 10. On days 1 and 6 post-infection, DNA from THP-1 cells subjected to the indicated treatments was amplified by PCR for HCMV genomes (IE region) or cellular genomes (GAPDH). PCR products were visualized by agarose gel electrophoresis. (B) The PCR signals of viral DNA were normalized to the signals of GAPDH. Data shown are the means +/- SEM of six independent experiments. VitD3, 1,25-dihydroxyvitamin D3. n.s. non-significant, *p<0.05, **p<0.01, d.p.i. day post-infection.

1,25-dihydroxyvitamin D3 treated THP-1 cells are more likely to exhibit IE gene expression following infection

While an equivalent amount of HCMV DNA is initially present following infection of control or 1,25-dihydroxyvitamin D3 treated cells, it is clear that the 1,25-dihydroxyvitamin D3 treated cells support a robust increase in productive HCMV replication. Therefore, we next chose to examine HCMV gene expression profiles in 1,25-dihydroxyvitamin D3 and PMA treated cells. Immediate early (IE) gene expression is typically repressed in cells that fail to undergo lytic phase induction, but is expressed rapidly after infection in cells capable of supporting lytic replication (Keyes et al., 2012a; Meier, 2001; Turtinen and Seufzer, 1994). We examined IE gene expression in cells treated with vehicle control, 1,25-dihydroxyvitamin D3 or PMA (Figure 4). Flow cytometric staining with anti-IE-Alex488 antibodies was performed as this enabled us to assess not only the frequency with which IE positive cells arise but also the relative level of IE antigens per cell. Infected cells were harvested and examined at day 1 post-infection. Compared to vehicle control, 1,25-dihydroxyvitamin D3 treatment resulted in a significantly higher percentage of IE positive cells by day 1 post-infection (Figures 4A and B). Interestingly, while the number of IE positive cells is significantly increased with 1,25-dihydroxyvitamin D3 treatment, there is no difference in the relative IE expression per cell as the mean fluorescent intensities are similar when comparing vehicle and 1,25-dihydroxyvitamin D3 treated cells (Figure 4C). PMA, in contrast, caused an increase in both the percentage of IE positive cells and relative IE expression per cell, suggesting that the mechanisms utilized by 1,25-dihydroxyvitamin D3 and PMA to promote lytic replication may be distinct. We did not detect any differences in the subcellular localization of IE1/2 when comparing control, 1,25-dihydroxyvitamin D3, and PMA treated cells indicating that changes in the compartmentalization of IE proteins is unlikely to account for the mechanism of 1,25-dihydroxyvitamin D3 induced HCMV replication (data not shown).

Figure 4. 1,25-dihydroxyvitamin D3 treatment increases the percentage of cells supporting HCMV IE gene expression.

(A) THP-1 monocytes were treated with 1,25-dihydroxyvitamin D3 (100 nM) or PMA (80 nM) for 3 days and then infected with HCMV TB40E at a MOI of 10. At 1 day post-infection, cells were fixed, permeabilized and stained with anti-HCMV IE antibody mAB 810-Alexa488. Cells were analyzed by flow cytometry. (B) The percentage of IE positive cells at day 1 post-infection is presented graphically. The data are derived from four independent experiments including the one depicted in panel A. (C) The mean fluorescence intensity of IE positive cells at day 1 post-infection is presented graphically. The data are derived from four independent experiments including the one depicted in panel A. VitD3, 1,25-dihydroxyvitamin D3. ** p<0.01,*** p<0.001.

Taken together, while viral genome copy numbers are initially equivalent, the 1,25-dihydroxyvitamin D3 treated cells are more highly likely to be capable of initiating IE protein production consistent with their ability to progress to the lytic phase (Figures 2 and 4). Moreover, while 20-40% of THP-1 cells treated with 1,25-dihydroxyvitamin D3 or PMA express IE antigens, it is evident that not all cells that progress through the IE phase go on to produce infectious virus based on infectious center assays, indicating that there are additional blocks subsequent to IE expression that control the progression to the lytic phase in HCMV infected myeloid cells.

HCMV early and late genes are expressed in 1,25-dihydroxyvitamin D3 stimulated cells

Although IE expression is important for initiation of lytic infection, the expression of early and late genes are needed to complete the lytic phase (McDonough and Spector, 1983; Wathen and Stinski, 1982). Therefore, the expression of early and late HCMV genes was examined by western blot in vehicle control, 1,25-dihydroxyvitamin D3 and PMA treated cells (Figures 5A and 5B). For these experiments, we analyzed UL44, a processivity factor associated with the viral DNA polymerase (Sinigalia et al., 2008), which is expressed with delayed-early kinetics (Hwang et al., 2000) and pp65, a tegument protein that is expressed with late kinetics (Kalejta, 2008). Vehicle control infected cells exhibited undetectable levels of either the delayed early UL44 or late pp65 proteins while cells treated with 1,25-dihydroxyvitamin D3 or PMA prior to infection showed dramatic upregulation of both UL44 and pp65. Representative western blots are depicted in Figure 5A and the results are depicted graphically in Figure 5B in which the blots were quantitated and viral protein levels are shown relative to the cellular actin protein as a control. To further investigate viral gene expression patterns in these cells and determine what percentage of IE positive cells progress to the early phase as evidenced by UL44 expression, cells at days 1 and 4 post-infection were co-stained for IE1/2 and UL44 expression and analyzed by flow cytometry (Figure 5C). In vehicle treated cultures, about 7.5% of the cells were IE positive at day 1 post-infection, while the percentage of IE positive cells dropped to 4.4% at day 4 post-infection. Of the 4.4% IE positive cells at day 4 post-infection only 11% of cells were UL44 positive (0.5% of the total cellular population). In 1,25-dihydroxyvitamin D3 treated cultures, approximately 33% of the cells were IE positive at day 1 post-infection, while the percentage of IE positive cells declined to 19% at day 4 post-infection. However, of the 19% IE positive cells, 33% were UL44 also positive (6.5% of the total cellular population). In PMA treated cultures, 40% of the cells were IE positive at day 1 post-infection and the percentage decreased slightly to 38% at day 4 post-infection. Of the 38% IE positive cells, 48% were also UL44 positive (18% of the total cellular population). The results of these early and late gene expression profiling experiments are also in line with the results of infectious center assay and are all consistent with the conclusion that 1,25-dihydroxyvitamin D3 promotes HCMV lytic replication in myeloid cells.

Figure 5. 1,25-dihydroxyvitamin D3 promotes HCMV early and late gene expression.

(A) THP-1 monocytes were treated with 1,25-dihydroxyvitamin D3 (100 nM) or PMA (80 nM) for 3 days and then infected with HCMV TB40E at a MOI of 10. At the indicated times post-infection, cells extracts were analyzed by western blot for UL44 (early) and pp65 (late) gene expression. Western blot analyses demonstrated that UL44 and pp65 are robustly expressed in 1,25-dihydroxyvitamin D3 and PMA treated cells. Cell extracts were also analyzed for actin expression as an internal control. (B) The results of four independent experiments are shown graphically as the ratio of UL44 or pp65 to actin. (C) THP-1 cells treated as described in panel A were stained for IE and UL44 proteins at days 1 and 4 post-infection and analyzed by flow cytometry. IE⁺ cells were gated (left panel in each pair) and then plotted as a function of UL44 expression (right panel in each pair) to determine the fraction of IE⁺ cells that have entered into the early phase as assessed by UL44 expression. The results shown are representative of 4 independent experiments. VitD3, 1,25-dihydroxyvitamin D3., *p<0.05, **p<0.01.

1,25-dihydroxyvitamin D3 uses a mechanism distinct from that of PMA to promote lytic replication

Due to the fact that both 1,25-dihydroxyvitamin D3 and PMA can prime THP-1 cells to support lytic infection, it would be intriguing to determine if these two reagents deploy the same mechanism or if 1,25-dihydroxyvitamin D3 functions in a manner distinct from that of PMA. Moreover, since it is clear that IE protein expression is critical for the onset of lytic replication, we wished to investigate the effects of 1,25-dihydroxyvitamin D3 and PMA on IE gene expression in a more detailed manner. Based on published reports (Abraham and Kulesza, 2013), it has been demonstrated that the HCMV IE enhancer region in THP-1 cells after infection is marked by histone 3 lysine 27 trimethylation (H3K27me3), and that the H3K27me3 mark at the IE enhancer is significantly decreased after PMA treatment. H3K27me3 is associated with a closed chromatin conformation and silenced gene expression (Fu et al., 2014), therefore it appears that decreased H3K27me3 in the IE region correlates with an open chromatin conformation and increased MIEP activity. We wanted to determine whether H3K27me3 associated with the IE enhancer is also decreased in 1,25-dihydroxyvitamin D3 treated cells. Using chromatin immunoprecipitation (CHIP) followed by PCR for the IE enhancer region, we find that the IE enhancer region in control cells is in fact modified by H3K27me3 as reported by others (Figure 6A) (Abraham and Kulesza, 2013; Rossetto, Tarrant-Elorza, and Pari, 2013). However, in both 1,25-dihydroxyvitamin D3 and PMA treated cells the CHIP-PCR signal is 4 to 10 fold weaker indicative of decreased H3K27me3 at the IE enhancer. Thus, these results are consistent with our analyses of IE1 protein expression and suggest that the transition of the MIEP enhancer region into an open conformation in 1,25-dihydroxyvitamin D3 treated cells is an important prerequisite for the transition to the lytic phase.

Figure 6. 1,25-dihydroxyvitamin D3 promotes K27 demethylation of histone H3 associated with the HCMV IE enhancer region, but does not directly stimulate IE promoter activity.

(A) THP-1 monocytes were treated with 1,25-dihydroxyvitamin D3 (100 nM) or PMA (80 nM) for 3 days and then infected with HCMV TB40E at a MOI of 10. At 2 days post-infection, chromatin immunoprecipitation was used to examine the K27 methylation status of histone H3 associated with the HCMV IE enhancer region. The results presented are derived from 3 independent experiments. (B) MIEP-luciferase activity was assessed in transient reporter gene assays in the presence of 1,25-dihydroxyvitamin D3 or PMA. MIEP-luciferase activity was normalized to the internal control renilla luciferase. The results presented are derived from 3-5 independent experiments performed in duplicate. VitD3, 1,25-dihydroxyvitamin D3. *p<0.05, **p<0.01.

The MIEP contains binding sites for a number of transcription factors that are responsive to PMA such as NF-κB and CREB (Liu et al., 2010), but it is unknown if the MIEP would be directly responsive to 1,25-dihydroxyvitamin D3. We cloned the HMCV IE promoter from the HCMV-FIX strain into the luciferase reporter pGL3 so that we could test whether 1,25-dihydroxyvitamin D3, like PMA would be able to directly stimulate the MIEP-luciferase reporter gene. We transfected THP-1 cells with pGL3-MIEP, stimulated cells with either 1,25-dihydroxyvitamin D3 or PMA, and measured luciferase activity. Reporter luciferase was internally controlled by comparison to constitutively expressed Rennila-luciferase. Interestingly, the Luciferase assay results demonstrate that while PMA can directly stimulate MIEP promoter activity, 1,25-dihydroxyvitamin D3 cannot (Figure 6B). These data are consistent with our flow cytometry data in Figure 4C, which indicated that 1,25-dihydroxyvitamin D3 treated cells are more likely to support IE protein expression but do not exhibit increased levels of the IE protein on a per cell basis (Figure 4C). Thus, taken together, while both 1,25-dihydroxyvitamin D3 and PMA can promote an open conformation of the MIEP followed by IE protein production and onset of the lytic phase, the mechanisms used by the two inducers are not identical as the effects of PMA can at least be partially explained by a direct effect on the MIEP promoter.

Since the two inducers appeared to not utilize totally overlapping mechanisms we investigated whether the effects of 1,25-dihydroxyvitamin D3 would be additive regarding the onset of lytic phase. To test this hypothesis, THP-1 cells were treated with PMA or PMA + 1,25-dihydroxyvitamin D3 prior to infection. Culture media was harvested at multiple time points post-infection and viral titers were assessed by plaque assay. Compared to PMA treated cells, 1,25-dihydroxyvitamin D3 plus PMA did not show an additive effect as the kinetics and magnitude of the viral growth is the same (Figure 7). Thus, while the mechanisms are not totally overlapping, the convergence of the two compounds on creating an environment more suitable for IE protein expression seems to be a necessary prerequisite leading to the onset of the lytic viral lifecycle.

Figure 7. 1,25-dihydroxyvitamin D3 and PMA do not function in a cooperative manner to increase lytic replication.

THP-1 cells were differentiated with PMA (80 nM) or PMA (80 nM) and 1,25-dihydroxyvitamin D3 (100nM) for one day, and then infected with HCMV at an MOI of 7. Cell supernatants were harvested at the indicated times post-infection and viral titers were determined by plaque assay on human foreskin fibroblasts. The results indicate that 1,25-dihydroxyvitamin D3 and PMA do not function cooperatively to enhance HCMV replication in THP-1 cells. The results are derived from 4 independent experiments. VitD3, 1,25-dihydroxyvitamin D3.

The differentiation of THP-1 cells triggered by 1,25-dihydroxyvitamin D3 plays an important role in releasing the restriction on HCMV IE expression

Based on published studies it is clear that 1,25-dihydroxyvitamin D3, like PMA, is an inducer of monocyte differentiation and maturation (Greenberger, Newburger, and Sakakeeny, 1980; Hmama et al., 1999; Schwende et al., 1996). However, while PMA induces the THP-1 cells to mature into a more highly differentiated macrophage-like state marked by strong adherence of the cells to plastic, 1,25-dihydroxyvitamin D3 induces differentiation into a mature monocyte marked by a non-adherent phenotype (Schwende et al., 1996). Therefore, we wished to investigate the influence of monocyte differentiation properties on 1,25-dihydroxyvitamin D3 induced HCMV replication in the THP-1 model. Consistent with previous studies (Daigneault et al., 2010; Schwende et al., 1996), we observed that PMA but not 1,25-dihydroxyvitamin D3 induced the appearance of a macrophage phenotype with flattened cells tightly adhered to the flask (Figure 8A). Immunostaining for the cell differentiation markers CD14, CD11b, CD36, CD54, and CD68 was also performed and the results demonstrate that 1,25-dihydroxyvitamin D3 consistently induces strong CD14 and moderate CD11b expression while having little to no effect on CD36, CD54, and CD68 expression (Figure 8B and Supplemental Figure 1). In contrast, PMA induced moderate levels of CD14, while inducing robust levels of CD11b, CD36, CD54, and CD68 (Figure 8B and Supplemental Figure 1). These results are entirely consistent with those of published studies and are further supportive of the conclusion that 1,25-dihydroxyvitamin D3 induces an intermediate differentiation phenotype typical of a mature monocyte, while PMA causes a terminally differentiated phenotype typical of a macrophage (Kremlev and Phelps, 1997; Kunisch et al., 2004; Munoz-Pacheco et al., 2012; Schwende et al., 1996). Interestingly, while PMA induces terminal differentiation and halts cellular proliferation, 1,25-dihydroxyvitamin D3 treated cells continued to proliferate in agreement with previous studies (data not shown) (Schwende et al., 1996).

Figure 8. 1,25-dihydroxyvitamin D3 and PMA induced THP-1 differentiation is phenotypically distinct.

(A) THP-1 cells were treated with 1,25-dihydroxyvitamin D3 (100 nM) or PMA (80 nM) for 3 days and photographed with a Olympus Q Color 5 camera equipped with QCapture Pro Software. (B) Cells treated as in panel A were stained with the indicated antibodies and analyzed by flow cytometry. Data from four independent experiments is depicted graphically. VitD3, 1,25-dihydroxyvitamin D3. ** p<0.01.

We then asked whether there was a correlation between 1,25-dihydroxyvitamin D3 promoted differentiation and onset of lytic replication. Since 1,25-dihydroxyvitamin D3 appeared to not directly stimulate MIEP promoter activity, we hypothesized that the effects of 1,25-dihydroxyvitamin D3 on IE expression might be delayed and only occur as the cells differentiated into mature monocytes. To test this postulate, THP-1 cells were treated with 1,25-dihydroxyvitamin D3 for different times (6 hours to 3 days) prior to infection with HCMV (experimental conditions are depicted graphically in Figure 9C). Expression of CD14 was monitored throughout the time course to assess cellular differentiation and IE positivity was assessed as a measure of the relative ability of the cells to promote lytic replication. IE expression was measured at 24 hours post-infection in all cases. This experimental system enables us to assess the expression of the differentiation marker CD14 and the HCMV IE protein in a series of cells in different stages of maturation. CD14 expression as measured by flow cytometry peaked at between 1 and 2 days post-stimulation with 1,25-dihydroxyvitamin D3 (Figure 9A). Interestingly, we did not observe significant effects of 1,25-dihydroxyvitamin D3 on IE positivity until 2 days post-stimulation with 1,25-dihydroxyvitamin D3 (Figure 9B). This finding is consistent with the conclusion that 1,25-dihydroxyvitamin D3-induction of lytic HCMV replication correlates with 1,25-dihydroxyvitamin D3 induced cellular differentiation. Moreover, the lack of a significant effect of 1,25-dihydroxyvitamin D3 at early time points argues against a direct effect of 1,25-dihydroxyvitamin D3 on the MIEP similar to what we observed in reporter assays in Figure 6.

Figure 9. The timing and magnitude of 1,25-dihydroxyvitamin D3 induced differentiation of THP-1 monocytes is important for supporting HCMV lytic infection.

(A) THP-1 cells were treated with 1,25-dihydroxyvitamin D3 (100 nM) for indicated times and CD14 expression was analyzed by flow cytometry. (B) Cells were treated with 1,25-dihydroxyvitamin D3 for the indicated times prior to infection with HCMV TB40E. At 24 hours post-infection cells were fixed, permeabilized and stained with HCMV IE antibody mAb810-Alexa488. The results represent flow cytometric analysis of IE positive cells and are derived from three independent experiments. (C) Experimental set up for the experiments depicted here. Cells were treated for variable lengths prior to infection with HCMV TB40E. In all cases IE gene expression was analyzed at 24 hours post-infection. VitD3, 1,25-dihydroxyvitamin D3. **p < 0.01, ***p < 0.001.

Discussion

In this study, we found that 1,25-dihydroxyvitamin D3, a hormone present in the circulatory system and in many tissues (Prietl et al., 2013), can promote HCMV replication in primary peripheral blood monocytes and in THP-1 cells. Our data are consistent with a mechanism whereby 1,25-dihydroxyvitamin D3 induced HCMV replication involves the induction of monocyte differentiation. Based on our data, we propose that monocyte maturation/differentiation induced by 1,25-dihydroxyvitamin D3 leads to a modification of histone 3 K27 methylation in the HCMV IE enhancer region, which results in a conversion to an open chromatin conformation and induction of IE gene expression. Moreover, since 1,25-dihydroxyvitamin D3 does not promote cell cycle arrest and terminal differentiation of monocyte cell lines in vitro (Schwende et al., 1996), this system represents an interesting paradigm that could be utilized to study reactivation of virus in model systems. In particular, while previous studies have shown that PMA differentiated THP-1 macrophages can support permissive infection, in this study we found that monocytes in the transition between mature monocytes and macrophage stages, like the state induced by 1,25-dihydroxyvitamin D3, can also support permissive infection.

Our work is the first to explore the effect of 1,25-dihydroxyvitamin D3 on HCMV in myeloid cells. Although the concentration of 1,25-dihydroxyvitamin D3 used in our study is somewhat higher than the concentrations found in vivo (Grande et al., 2002), published studies have demonstrated that in hematopoietic progenitor cells derived from umbilical cord blood, a similar dose of 1,25-dihydroxyvitamin D3 added once a week has the same effects on monocyte-macrophage differentiation as does physiological concentrations of 1,25-dihydroxyvitamin D3 supplemented daily (Grande et al., 2002). In this case the difference between the two conditions is that high dose treatment causes more rapid differentiation and lineage commitment than does physiological concentrations and thus the high doses like those used in our study simply facilitate experiments performed in an in vitro setting.

Although in this research, our main focus is to define the effect of 1,25-dihydroxyvitamin D3 on HCMV lytic replication in peripheral blood monocytes, an equally attractive question to be addressed in the future is to ask what effect 1,25-dihydroxyvitamin D3 has on HCMV replication in hematopoietic progenitor cells (HPCs), a cell type well-known as a reservoir for HCMV latency (Goodrum et al., 2002; Maciejewski and St Jeor, 1999). In particular does 1,25-dihydroxyvitamin D3 treatment of HPCs affect the ability of HCMV to choose a latent or lytic path and would the presence of high concentrations of 1,25-dihydroxyvitamin D3 promote lytic reactivation? It is known in the HCMV field that macrophages and mature dendritic cells can support lytic infection but hematopoietic progenitor cells, myeloid progenitor cells and monocytes are cell types known to typically establish a latent infection (Sinclair, 2008; Sinclair, 2010). THP-1 cells treated with 1,25-dihydroxyvitamin D3 do not show a phenotype characteristic of mature macrophages and therefore, 1,25-dihydroxyvitamin D3 appears to induce a differentiation state in between monocytes and macrophages. Previously, studies of HCMV in myeloid cells have focused on hematopoietic progenitor cells (Goodrum et al., 2007; Maciejewski et al., 1992), myeloid progenitor cells (Cheung et al., 2006; Kondo, Kaneshima, and Mocarski, 1994), monocytes (Chan, Nogalski, and Yurochko, 2012; Keyes et al., 2012b; Stevenson et al., 2014) or macrophages (Sanchez et al., 2012), and thus our research offers some additional insight into HCMV replication in cells that are transitioning between the monocyte and macrophage stages. Although previous studies suggest that HCMV typically enters the lytic phase in mature macrophages (Smith et al., 2004; Turtinen and Seufzer, 1994; Weinshenker, Wilton, and Rice, 1988) our results indicated that cells transitioning between the monocyte and macrophage stages can also support lytic infection. HCMV infection itself has been shown to promote monocyte differentiation but the differentiation patterns triggered by infection alone are not typically capable of efficiently driving lytic infection (Chan, Nogalski, and Yurochko, 2012; Stevenson et al., 2014). It is highly possible that there is a differentiation threshold that is needed to be passed in order to appropriately kindle a lytic infection. 1,25-dihydroxyvitamin D3 treatment may prime the cells in the differentiation process, and when infection provides the appropriate additional differentiation signals, the threshold is surpassed.

In these studies we examined whether 1,25-dihydroxyvitamin D3 differentiated monocytes can support lytic infection, but as mentioned above an important question to ask in the future is whether 1,25-dihydroxyvitamin D3 may also be involved in the reactivation of HCMV from latency. Recently, glucocorticoids have been shown to trigger reactivation of HCMV in primary monocytes through direct activation of the IE promoter (Van Damme et al., 2014). Since there are studies showing crosstalk between glucocorticoids and 1,25-dihydroxyvitamin D3 (Hidalgo et al., 2011), and since 1,25-dihydroxyvitamin D3 can enhance glucocorticoid action in human monocytes (Zhang, Leung, and Goleva, 2013; Zhang, Leung, and Goleva, 2014), it reasonable to speculate that 1,25-dihydroxyvitamin D3 may also be involved in the regulation of HCMV reactivation or work in concert with glucocorticoids in this process. Perhaps since the mechanisms of actions appear to be disparate (glucocorticoids directly on the MIEP and 1,25-dihydroxyvitamin D3 on cellular differentiation) these two hormones may work synergistically to affect HCMV replication and/or reactivation.

Differentiation of monocytes by 1,25-dihydroxyvitamin D3 appears to robustly induce the HCMV lytic phase, but the precise molecular mechanism(s) that regulates this activity remain unknown. Based on our studies, we postulate that signaling activity and gene expression patterns typically triggered by 1,25-dihydroxyvitamin D3 to drive monocyte maturation/differentiation are required for HCMV lytic infection to proceed. Global comparison of downstream gene expression induced by 1,25-dihydroxyvitamin D3, PMA and/or glucocorticoids in monocytes by microarray or RNA-seq will be helpful to narrow down the list of candidates, and increase the likelihood of identifying the essential molecules that regulate this switch.

From a clinical perspective, there are many ongoing studies showing that 1,25-dihydroxyvitamin D3 or its synthetic analogs can have anti-cancer effects, and can have beneficial effects on cardiovascular or autoimmune disease (Delvin et al., 2014; James et al., 1998; Menezes et al., 2014). Since there is some evidence supporting a role for HCMV infection in the progression of cancers (Michaelis et al., 2011; Soroceanu and Cobbs, 2011), cardiovascular and autoimmune diseases (Caposio, Orloff, and Streblow, 2011; Varani et al., 2009), our study could prompt questions regarding whether or not 1,25-dihydroxyvitamin D3 and its synthetic analogs should be used therapeutically. In addition, the effect of 1,25-dihydroxyvitamin D3 on HCMV replication in other cell types including cancer cells is still unknown. Moreover, while 1,25-dihydroxyvitamin D3 can have genomic and non-genomic effects (Norman et al., 1992), whether those synthetic analogs have the same effect of 1,25-dihydroxyvitamin D3 on HCMV replication is another intriguing question to answer and which may provide an important tool to tease out the specific pathways involved in lytic induction.

Supplementary Material

supplement

Supplemental figure 1. 1,25-dihydroxyvitamin D3 and PMA induced THP-1 differentiation is phenotypically distinct: THP-1 cells were treated with ethanol (left column), 1,25-dihydroxyvitamin D3 (100 nM) (middle column) or PMA (80 nM) (right column) for 3 days, stained with the indicated antibodies and analyzed by flow cytometry. The area depicted by the black line represents unstained cells, and the gray shade area shows the cells stained with the indicated antibody. The histograms presented are representative of 4 independent experiments.

• Vitamin D3 induces HCMV replication in blood monocytes and in THP-1 cells

• Vitamin D3 induced HCMV replication is driven by monocyte differentiation

• The Vitamin D3 effect is independent of a direct activity on the MIEP

• Defining the Vitamin D3 induced pathways will give insight into lytic switching