Abstract
The accessory protein Vpr of Human Immunodeficiency Virus type 1 (HIV-1) enhances replication of the virus in macrophage but less so in cycling T cells. Virus particle packaged Vpr is released in target cells shortly after entry, suggesting it is required early in infection. A commonly observed activity of Vpr is the induction of cell cycle arrest in T cells but why this is so is unknown. Here we observe, by co-immunoprecipitation assay, an interaction between Vpr and endogenous REAF (RNA-associated Early-stage Antiviral Factor, RPRD2), a protein previously shown to potently restrict HIV infection. After HIV-1 infects macrophages, within 30 minutes of viral entry, Vpr induces the degradation of REAF. Subsequently, as replication continues, REAF expression is upregulated – a response curtailed by Vpr. Using VLPs we show that Vpr alone is sufficient to down modulate REAF in MDMs. REAF is more highly expressed in differentiated macrophages than in cycling T cells. Expression in cycling cells is cell cycle dependent and knockdown induces cell cycle perturbation. Therefore, our results support the hypothesis that Vpr induces the degradation of a factor involved in the cell cycle that impedes HIV infection in macrophages.
Introduction
Human Immunodeficiency Virus type 1 (HIV-1) infects CD4+ T cells and macrophages in vivo and causes Acquired Immunodeficiency Syndrome (AIDS). HIV-1 has four non-structural accessory genes nef, vif, vpu and vpr. Nef, vif and vpu mitigate host innate immunity. A function for Vpr has been elusive, but it is required for replication in macrophages and for pathogenesis in vivo (1, 2). A widely acknowledged but poorly understood Vpr-mediated phenotype is that an infected population of cycling cells accumulates in the G2/M phase of the cell cycle. The observed phenomenon is achieved by manipulation of the cullin4A-DBB1 (DCAF1) E3 ubiquitin ligase and the recruitment of an unknown substrate for proteasomal degradation. A large number of Vpr substrates have been proposed (3–11). Yan et al. (2019) show that helicase-like transcription factor (HLTF) restricts replication of HIV-1 in T cells shown previously by Lahouassa et al. 2016 to be down modulated by Vpr about six hours post infection (12, 13). Furthermore, Greenwood et al. 2019 report on the effect that Vpr has on large scale remodelling of nearly 2000 cellular proteins including those that bind nucleic acids and those involved with the cell cycle (14).
Substantial but finite quantities of Vpr are incorporated into viral particles and released from the major capsid protein (CA) after entry into the cell (15, 16). Concurrently, reverse transcription transcribes the RNA genome into DNA, which integrates into the host cell DNA. The early release of Vpr from the CA (17) suggests it has an early function prior to integration. When considering the role of Vpr in cell tropism and pathogenesis it is important to focus on proteins that have a direct effect on viral replication. Here we show that within 30 minutes of cellular entry, Vpr containing virus induces the degradation of RNA-associated Early-stage Antiviral Factor (REAF, also known as RPRD2). REAF has been shown to limit the completion of pro-viral DNA synthesis and integration (18). Here we show that it restricts replication of HIV-1 in macrophages, a restriction overcome by Vpr.
Results
HIV-1 Vpr interacts with REAF and overcomes restriction
HeLa-CD4, knocked down for REAF (HeLa-CD4 shRNA-REAF, Figure 1A) were challenged with two HIV-1 primary macrophage tropic isolates 89.6 and 2044 (19). Figure 1B showns both viruses could replicate more efficiently in the absence of REAF (between 20 and 50 fold). To determine if a viral accessory gene could overcome this REAF effect we tested infectivity of HIV-1 89.6WT and mutants deleted for vpr (89.6Δvpr), vif (89.6Δvif) or vpu (89.6Δvpu). Figure 1C shows that despite a standard virus input (50 FFU/ml, as measured on HeLa-CD4), the prevention of REAF expression using short hairpin RNA (shRNA) alleviates the need for Vpr. There is significantly greater rescue of HIV-1 89.6Δvpr (>40 fold, p < 0.0001) compared to HIV-1 89.6WT or other mutant viruses with an intact vpr gene but lacking vpu or vif. Thus, vpr potently overcomes the restriction imposed by REAF. We further confirmed that the mutant HIV-1 89.6Δvpris restricted in HeLa-CD4 compared to HIV-1 89.6WT. Figure 1D shows that despite equal inputs of p24, significantly fewer foci of infection (FFU) result from challenge with HIV-1 89.6Δvpr compared to HIV-1 89.6WT. Further support of a role for Vpr is supported in Figures 1E and F. When PM1 cells are challenged with a HIV-1 89.6WT, which has an intact vpr, REAF protein is down modulated (Figure 1E). The observed down modulation is dependent on the presence of Vpr as HIV-1 89.6∆vpr is incapable of degrading REAF (Figure 1F). Moreover, Figure 1G shows that Vpr and REAF interact with each other, either directly or as part of a complex, as they are co-immunoprecipitated.
Macrophages are a target for HIV infection (20) and numerous studies support the notion that vpr is crucial for their efficient infection. We therefore focussed on REAF effects in this primary cell type. Using Western blotting we followed REAF protein during the maturation of blood derived monocytes (day 0) to macrophages (Figure 3H). We observed that REAF levels increase over time up to 7 days of maturation. To investigate the ability of Vpr to deplete REAF protein in MDMs we used imaging flow cytometry to quantify the overall REAF mean fluorescence intensity (MFI) in large populations (5000) of target cells after infection. Previously we reported that REAF affects the production of reverse transcripts early after infection and that at this critical time point REAF is transiently down modulated in HeLa-CD4 (21). Here we confirmed that the loss of REAF from MDMs also occurs early, where the Vpr dependent loss of REAF is observed from 30 minutes after infection (Figure 1I) with recovery around 2 hours. In contrast, without Vpr (HIV-1 89.6Δvpr) the opposite occurs, REAF levels increase after infection. The increase in REAF levels is most potent after 30 mins of infection with HIV-1 89.6Δvpr.
Previously it was reported that Vpr is released from the capsid and enters the nucleus shortly after infection (11). We generated a virus with a mutated form of Vpr (F34I) described as lacking the ability to localise to the nuclear membrane or to interact with the nuclear transport proteins importin-α and nucleoporins (22). Interestingly, HIV-1 89.6F34I, similar to HIV-1 89.6Δvpr, can no longer deplete REAF in MDMs (Figure 1H). We confirmed that HIV-1 89.6Δvpr has restricted replication in MDMs compared with the wild type virus expressing Vpr (HIV-1 89.6WT) (Figure 1I). Like HIV-1 89.6Δvpr, the mutant virus (HIV-1 89.6F34I) replicates less efficiently in MDMs.
Other targets of Vpr have been proposed. It recruits SLX4-SLX1/MUS81-EME1 endonucleases to DCAF1, activating MUS81 degradation and triggering arrest in G2/M (23). It also degrades helicase-like transcription factor (HLTF) (13) and has recently been shown to enhance infection of HIV-1 in T-cells lines (Yan et al. 2019). We show here both HLTF (Figure 1J) and MUS81 (Figure 1K) are depleted by virus concurrently with REAF within 60 minutes of infection. In contrast to REAF, HLTF and MUS81 levels do not recover.
Fluctuation of subcellular REAF expression after HIV-1 infection is Vpr dependent
In view of the above, using imaging flow cytometry, we sought to determine the relative subcellular localization of REAF. We found that REAF is mostly present in the nuclear compartment compared to the cytoplasm (Figure 2A) of both Hela-CD4 (left) and particularly so in primary macrophages (right). We looked more carefully at the cellular distribution of REAF at the early time points following HIV-1 infection. Here, HeLa-CD4 infected with HIV-1 89.6WT or HIV-1 89.6Δvpr were quantified for REAF nuclear or cytoplasmic protein over the first three hours following infection by imaging flow cytometry. Following challenge with HIV-1 89.6Δvpr, REAF levels increase in both the nucleus (~25%, Figure 2B, left) and cytoplasm (~10%, Figure 2B, right) within 0.5 hours and nuclear levels remain high for 3 hours. However, in the presence of Vpr (HIV-1 89.6WT) the observed increase in REAF is curtailed, with a steady decline from 0.5-2 hours. The decline is most marked in the nucleus with ~20% reduction by 1 hour and ~30% at 2 hours. By 3 hours, levels of REAF protein recover. The virus carries finite quantities of Vpr (17), which potentially explains why there is a pause in the down modulation of REAF. Imaging flow cytometry software determined the ‘nuclear enrichment score’ over time after infection with HIV-1 89.6WT or HIV-1 89.6∆vpr (Figure 2C). The lower the score the less REAF in the nucleus relative to in the cell overall. By 1-2 hours, a significant segregation emerges and so the presence of Vpr, relative nuclear levels of REAF are suppressed. Lower levels of REAF were also observed in the cytoplasm over time but to a much lesser extent (Figure 2B, right).
Nuclear levels of REAF were compared in a number of primary cell types using imaging flow cytometry (Figure 2D). When compared to either monocytes or resting/activated T cells, both MDMs and dendritic cells (DCs) highly express nuclear REAF. Antiviral factors are often upregulated in response to pathogen associated molecular patterns. Polyriboinosinic:polyribocytidylic acid (poly(I:C)) is a double-stranded RNA, used to stimulate viral infection associated molecular pattern recognition pathways. Figure 2E shows poly(I:C) induction of REAF in THP-1, a macrophage cell line.
We sought to determine if the above fluctuations of REAF observed in HeLa-CD4 cells also occur in MDMs. The subcellular fluctuation of REAF levels from two donors after challenge was determined using imaging flow cytometry. With vpr (HIV-1 89.6WT), nuclear REAF decreases within 2 hours of viral infection of macrophages from two donors (Figure 2F), similar to that observed in HeLa-CD4 (Figure 2B). In contrast, also in both donors, nuclear REAF rapidly increases from as early as 0.5 hours when the virus does not contain Vpr (HIV-1 89.6Δvpr) (Figure 2F). For the cytoplasmic compartment, a similar picture emerges for REAF fluctuation. In both donors, when Vpr is absent, REAF levels increase rapidly within 0.5 hours of infection (Figure 2F). This cytoplasmic increase is curtailed in donor 1 when Vpr is present. In donor 2 the loss of Nuclear REAF after HIV-1 89.6WT infection is paralleled by an increase in cytoplasmic REAF. Similar kinetics of total REAF protein fluctuation were observed, in the presence or absence of Vpr, in two further donors. This was measured by Western blotting and is presented in Expanded View Figure 1.
We sought to determine if knockdown of REAF in primary macrophages resulted in an increased susceptibility to HIV-1 infection. In Figure 2G primary MDMs were treated with siRNA targeting REAF (siREAF) or a control protein (siCB). Cells lacking REAF were found to be significantly more susceptible to infection with HIV-1 89.6.
In Figure 2H, MDMs were treated with virus-like particles (VLPs) containing Vpr and Western blotting confirmed that Vpr alone is sufficient to induce the down modulation of REAF. MDMs, pre-treated with VLPs (Figure 2H) were then subsequently infected with either HIV-1 89.6WT or 89.6Δvpr. Cells with VLP-induced down modulation of REAF were found to be significantly more susceptible to infection with either virus but particularly virus lacking Vpr of its own.
Depletion of REAF results in G2/M accumulation and mitotic cells are more susceptible to HIV-1 infection
After knockdown of REAF in THP-1 (THP-1 shRNA-REAF, Figure 3A), there is a clear increase in mitosis, indicated by the phosphorylation of histone H3 (Ser10/Thr11). We further confirmed G2/M arrest after knockdown of REAF in a T cell line, PM1 (PM1-shRNA-REAF, Figure 3B-D). Reduction of REAF mRNA (Figure 3B) and protein (Figure 3C) was confirmed. An accumulation of cells at G2/M, illustrated by the significant increase in G2/G1 ratio (Figure 3D, insert), was seen comparable to previous reports of Vpr induced cell cycle arrest (Figure 3D)(14).
We defined the cell cycle phases (G0/1, S and G2/M) of primary human monocytes and analysed REAF expression. Levels are lowest in G0/1, increase through S phase, and peak in G2/M (Figure 3E). Using imaging flow cytometry, we further analysed the subcellular localization of REAF during mitosis (Figure 3F). An asynchronous population had a nuclear enrichment score of 0.92 (solid grey, left). Nocodazole-treated cells diverged into two populations (0.13, 1.53, Figure 3F, black outline, left). To confirm the divergence in the nuclear enrichment scores corresponded to cell cycle, the mitotic cell marker (histone H3 pSer28) was used to identify such cells. Mitotic cells were shown to have a lower nuclear enrichment score (0.17), and thus lower levels of REAF in the nucleus relative to the cell overall. The treatment of cells with nocodazole resulted in an increase in the population with a low nuclear enrichment score (Figure 3F, right). Representative images are shown in Figure 3G. Using confocal microscopy, REAF is observed in both the cytoplasm and nucleus through interphase, prophase and prometaphase but excluded from chromatin during metaphase, anaphase and telophase (Figure 3H).
Vpr has been shown, to varying degrees, to be more beneficial for replication in macrophages than in cycling T cells (22, 24–27). We compared the susceptibility of a mitotic enriched population of HeLa-CD4 to an asynchronous population (Figure 3I-K). A thymidine/nocodazole treated population was confirmed to be mitotic enriched using flow cytometry (Figure 3I) and Western blotting (Figure 3J). The mitotic enriched population was more susceptible to infection with HIV-1 89.6WT (approximately 7-11 fold, Figure 3K). A further experiment using HIV-1 89.6 (VSV-G) with a GFP reporter as a challenge virus confirmed the increased susceptibility of mitotic cells to infection (Expanded View Figure 2).
IFNα induces many HIV restriction factors (28, 29). We used RNA microarray analysis to determine if IFNα upregulated REAF mRNA in MDMs. Figure 4A shows IFNα induced upregulation of antiviral genes, including HIV restriction factors APOBEC3G, IFITM1-3, MX2, Tetherin and Viperin (30) but with little or no upregulation of REAF mRNA. Nor was REAF mRNA upregulated in MDMs or CD4+ T cells in response to IFNα (Figure 4B).
Restriction factors are often under evolutionary positive selection at sites that interact with virus. We compared REAF DNA sequences from 15 extant primate species using PAML package for signatures of positive natural selection. We found no evidence of positive selection of REAF in the primate lineage (Figure 4C).
Discussion
The deletion of vpr in HIV-1 leads to significant impairment of replication in both HeLa-CD4 and primary macrophages. A number of experiments presented here point to a role for Vpr in the counter-restriction of the antiviral protein REAF. First, HIV-1 replication is significantly enhanced by knockdown of REAF in either HeLa-CD4 or primary macrophages and this phenotype is even more pronounced for viruses lacking vpr. Second, REAF is down modulated early after infection in a manner dependent on both the presence of Vpr and, as demonstrated by a vpr single point mutation, its localisation to the nuclear envelope. Third, using VLPs we show that Vpr alone is sufficient to down modulate REAF in MDMs. Finally, we show, by co-immunoprecipitation, that REAF and Vpr physically interact with each other either directly or as part of a complex. Taken together, our results highlight the importance of the relationship between REAF and the HIV-1 accessory protein Vpr.
Others have shown a specific requirement for Vpr in infection of non-dividing cells and only reduced replication rates in cycling T cells (20). Indeed, REAF is more highly expressed MDMs compared to cycling T cells. The requirement for Vpr in macrophage infection is substantiated here where reduced viral particle production is observed after infection of MDMs with either HIV-1 89.6∆vpr or HIV-1 89.6F34I compared to HIV-1 89.6WT. This is the first demonstration of a vpr-alleviated impairment of HIV-1 replication in primary macrophages. Recently however Yan et al. (2019) show that HLTF, a proposed target of Vpr, restricts replication of HIV-1 in T cells while Lahouassa et al. (2016) also reported a Vpr dependent loss of HLTF at six hours post infection (8). Our results also support a role for HLTF in HIV replication. We show that HLTF down modulation occurs concomitantly with REAF as early as 0.5 hours post infection. Interestingly, HLTF and REAF were identified in the same screen for proteins that interact with single-stranded DNA (31).
The transient nature and timing of REAF depletion is consistent with its ability to impede the production of reverse transcripts early in infection (21). After an initial down modulation of REAF following infection REAF depletion is paused one hour later, perhaps attributable to the finite quantities of Vpr carried in the virus particle (17). In contrast however, down modulation of HLTF and MUS81 (another proposed target of Vpr) is maintained, perhaps suggesting alternate or additional roles for these factors later in infection (27, 22). Recently, Greenwood et al., using a whole cell proteomics screen for factors up or down modulated by Vpr in T cells, identified HLTF among almost 2000 proteins affected, underlining the promiscuous activity of Vpr (14). It is now critical, in light of these findings, that attention is directed to those proposed Vpr targets that affect replication of HIV-1 in primary cells. The recovery of REAF expression prior to 6 hours after infection most likely explains why it was not identified in the data collected by Greenwood et al. (14).
Our model is that Vpr is brought into the cell by HIV-1, in limited, but sufficient quantities to down modulate REAF in the timeframe required for reverse transcription to proceed unhindered. Nuclear localisation of Vpr is required for the down modulation of REAF perhaps similar to the Vpx mediated depletion of the reverse transcription inhibitor SAMHD1, for which degradation is initiated in the nucleus(32). We propose that REAF is linked to the innate immune response as treatment of primary macrophages with poly(I:C) induces its expression. Furthermore, HIV-1 without vpr, induces the expression of REAF to high levels in primary macrophages within 0.5 hours of infection.
A poorly understood phenomenon is that Vpr induces cell cycle arrest in the G2/M phase after infection. We report here that the loss of REAF from cycling cells contributes to an accumulation of the population in G2/M. Other studies suggest that vpr is less dispensable for infection of non-dividing cells (20). In fact, REAF is more highly expressed MDMs compared to cycling T cells supporting this premise.
REAF is unlike the evolving HIV restriction factors like APOBEC3G, SAMHD1, TRIM5 or BST2/Tetherin and is more similar to SERINC3 and 5 which are not under positive selection (33, 34). We propose that REAF is a multi-functional or ‘moonlighting’ protein with at least two cellular roles (35). REAF has many properties of restriction factors (30, 36). It interacts with HIV-1 reverse transcripts, impeding reverse transcription (21). It is germline encoded, constitutively expressed in cells, regulated by the proteasome system, suppressed by Vpr and upregulated by poly(I:C). Our results support the current model for Vpr activity which is that it induces the degradation of a protein involved in an unknown restriction of HIV-1.
Materials and Methods
Ethics Statement
Leucocyte cones from blood donors, from which PBMCs were isolated, were obtained from the NHS Blood Transfusion service, St. George’s Hospital, London. Donors were anonymous and thus patient consent was not required. The local ethical approval reference number is 06/Q0603/59.
Cell Lines
HEK-293T (ATCC), PM1, THP-1, C8166, HeLa-CD4 parental (all NIBSC AIDS Reagents) and HeLa-CD4 shRNA-REAF (previously described) were maintained at 37°C in 5% CO2 (18). Cells were cultured in Dulbecco’s Modified Eagle Medium (DMEM, ThermoFisher) supplemented with fetal bovine serum (FBS) 5-10% from Thermo Fisher and appropriate antibiotics. HeLa-CD4-shRNA-REAF were selected for resistance to puromycin in media supplemented with 10µg/ml puromycin.
The isopropyl β-D-1-thiogalactopyranoside (IPTG)-inducible vector pLKO-IPTG-3xLacO (Sigma) was used to express short hairpin RNAs (shRNAs) targeted against REAF (Sigma Mission TRCN0000141116). Additionally, a non-target (scramble) control was prepared. Viral particles for cell line transductions were prepared by co-transfecting HEK-293T cells with pLKO-IPTG-3xLacO, the Gag/Pol packaging vector pLP1, a Rev expression vector pLP2, and the vesicular stomatitis virus G protein (VSV-G) expression vector pVPack-VSV-G (Stratagene). After 72 hours, virus was clarified by low-speed centrifugation and passed through a 0.45-m-pore-size filter. THP-1 and PM1 cells were transduced by culturing viral particles in the presence of 8g/ml Polybrene for 72 hours, after which resistant colonies were selected and maintained with 2µg/ml puromycin. Culturing cells in the presence of 1mM IPTG for 72 hours induced expression of shRNAs.
Transfections and Virus/VLP Production
The HIV-1 primary isolate 2044 has been previously described (19). The infectious molecular clone for HIV-1 89.6 was obtained from the Centre for AIDS Research (NIBSC, UK). Infectious full-length and chimeric HIV clones were prepared by linear polyethylenimine 25K (Polysciences), Lipofectamine 2000 (Invitrogen) or Lipofectamine 3000 (Invitrogen) transfection of HEK-293T. Virus-like particles (VLPs) were produced by linear polyethylenimine 25K (Polysciences) transfection of HEK-293T. The VLP packaging vector was a gift from N. Landau and production is described in reference (20). HIV-1 89.6 (VSV-G) was generated by combining the transfer vector pCSGW with the envelope pMDG VSV-G and the core construct p8.91-89.6gag in HEK-293T as has been previously described (21).
Plasmid constructs HIV-1 89.6Δvif, HIV-1 89.6Δvpr and HIV-1 89.6Δvpu were generated from the HIV-1 89.6 molecular clone, using overlap extension PCR (29). Clones were confirmed by plasmid sequencing (Source BioScience). Primer sequences are available upon request. HIV-1 p89.6 vpr mutant F341 was made by site directed mutagenesis (Agilent) of the p89.6 plasmid. HEK-293T were plated at 2×104/cm2 in 8-well chamber slides (for confocal microscopy), or 10cm dishes (for virus and VLP production) 48 hours prior to transfection. For virus/VLP production, supernatant was harvested 72 hours post transfection and cleared of cell debris by centrifugation at 500 x g for 5 minutes. Stocks were stored at −80°C. All viruses were amplified by C8166 for 48 hours before harvest.
Titration of Replication Competent Virus
HeLa-CD4 were seeded at 1.5×104 cells/well in 48-well plates to form an adherent monolayer of cells. Cell monolayers were challenged with serial 1/5 dilutions of virus and titre was assessed after 48 hours by in situ intracellular staining of HIV-1 p24 to identify individual foci of viral replication (FFU), as described previously (21). For infection time course experiments, 400-500μl of 1×105 FFU/ml (HeLa-CD4) or 3×103 FFU/ml (MDMs) virus was added per well to cells cultured in 6-well trays for 24 hours (HeLa-CD4) or 7 days (for MDMs). For Figure 3I, cells were challenged with 50ng p24 in 6-well plates with 2×106 MDMs per well. Supernatants were harvested on days 0, 2, 8, 21 and 28 post challenge and p24 concentration analysed by ELISA.
p24 ELISA
ELISA plates were pre-coated with 5μg/ml sheep anti-HIV-1 p24 antibody (Aalto Bio Reagents) at 4°C overnight. Viral supernatants were treated with 1% Empigen® BB for 30 minutes at 56°C, then plated at 1:10 dilution in Tris-bufered saline (TBS) on anti-p24-coated plates and incubated for 3 hours at room temperature. Alkaline phosphatase-conjugated mouse anti-HIV-1 p24 monoclonal antibody (Aalto Bio Reagents) diluted in 20% sheep serum, 0.05% v/v Tween-20, TBS was then added and incubated for 1 hour at room temperature. Plates were washed 4 times with 0.01% v/v Tween-20 in PBS and twice with ELISA Light washing buffer (ThermoFisher). CSPD substrate with Sapphire II enhancer (ThermoFisher) was added and incubated for 30 minutes at room temperature before a plate reader was used to detect chemiluminiscence.
cDNA Synthesis and qPCR
Total RNA was extracted from PM1 cells using the ReliaPrep RNA kit (Promega) and in one-step reverse transcription qPCR (Quantbio) using TaqMan probes for the detection of amplified transcripts. Data acquisition was performed on an Agilent Mx3000 and analyzed on MxPro software.
From MDMs, total RNA was extracted using an RNeasy Plant Mini Kit (QIAGEN), and cDNA was synthesized with SuperScript™ III First-Strand Synthesis System (Invitrogen), according to manufacturer’s instructions. cDNA was subjected to real-time quantitative PCR (qPCR) using REAF, OAS1 and β-actin primer pairs with SYBR® Green detection of amplified transcripts (QuantiTect SYBR Green PCR Kit, QIAGEN). Data acquisition and analysis were performed using the ABI PRISM™ 7500 SDS software. Primer sequences are available upon request.
Gene Expression RNA Microarray
Prior to microarray analysis, RNA from MDMs was prepared using the Illumina™ TotalPrep™ RNA Amplification Kit (Ambion), according to manufacturer’s instructions. The probes were hybridized on an Illumina™ HT12v3 bead array following the manufacturer’s standard hybridization and scanning protocols. Raw measurements were processed by GenomeStudio software (Illumina), and quantile normalized. All microarray data are publicly available in the Gene Expression Omnibus (GEO) database with accession number GSE54455.
IFN, Poly(I:C) and Treatment
MDMs, CD4+ T cells and THP-1 were treated with IFN (100-500IU/ml, specified) for 24 or 48 hours (specified) before harvest for RNA extraction; analysis by Western blotting or imaging flow cytometry. THP-1 were treated with poly(I:C) (25μg/ml, HMW/LyoVec™, Invitrogen) for 48 hours before analysis by Western blotting or imaging flow cytometry. Prior to IFN or poly(I:C) treatment, THP-1 were treated with phorbol 12-myristate 13-acetate (PMA, 62ng/ml) for 3 days and then PMA-free DMEM for 2 days to allow differentiation and recovery. For Figure 4A and Figure S6C-D, recombinant IFNα was purchased from Sigma (Interferon-αA/D human Cat. No. I4401-100KU) and is a combination of human subtypes 1 and 2. For Figure S6A-B and E, recombinant human IFNs are from Peprotech.
Western blotting
Cells were harvested and lysed in 30-50μl of radioimmunoprecipitation assay (RIPA) buffer supplemented with NaF (5µM), Na2VO3 (5µM), β-glycerophosphate (5µM) and 1x Protease Inhibitor Cocktail (Cytoskeleton). The protein concentration of each sample was determined using BCA Protein Assay Kit (Pierce). 12.5-70µg of total protein was separated by SDS-PAGE (4-12% Bis-Tris Gels, Invitrogen), at 120V for 1 hour 45 minutes in MOPS SDS Running Buffer (Invitrogen). Separated proteins were transferred onto nitrocellulose membrane (0.45μm pore size, GE Healthcare) at 45V for 2 hours, in ice-cold 20% (v/v) Methanol NuPAGE™ Transfer Buffer (ThermoFisher). Membranes were blocked for 1 hour at room temperature in 5% (w/v) non-fat milk powder in TBS-T buffer. Specific proteins were detected with primary antibodies by incubation with membranes overnight at 4°C and with secondary antibodies for 1 hour at room temperature. All antibodies were diluted in blocking buffer. Proteins were visualized using ECL Prime Western Blotting Detection Reagent (GE Healthcare) and imaged using either ChemiDoc Gel Imaging System (Bio-Rad) or exposed to CL-XPosure films (ThermoScientific) and developed.
Antibodies
Primary rabbit polyclonal antibody to REAF (RbpAb-RPRD2) has been previously described (21). For imaging flow cytometry and confocal microscopy, RbpAb-RPRD2 was detected using goat anti-rabbit IgG conjugated with Alexa Fluor 647 (Invitrogen). FITC-labelled anti-phospho-histone H3 (Ser28) Alexa 488 was used (BD Bioscience) for imaging flow cytometry and confocal microscopy. MsmAb-IFITM1 (clone 5B5E2, Proteintech), was detected by goat anti-mouse IgG Alexa Fluor 555 (ThermoFisher) for imaging flow cytometry, and by anti-mouse IgG antibody conjugated to HRP (GE Healthcare) for Western blotting, as were MsmAb-Mus81 and MsmAb-GFP (both Abcam). Also for Western blotting, RbpAb-RPRD2, RbmAb-IFITM3 (EPR5242, Insight Biotechnology), RbpAb-GAPDH, RbpAb-βActin, RbmAb-phospho-histone H3 (Ser10/Thr11) and RbpAb-HLTF (all Abcam) were detected with secondary antibody: donkey anti-rabbit IgG conjugated to HRP (GE Healthcare).
Immunoprecipitation
HEK-293T, transfected with either VPR-GFP or GFP control expression vector, were lysed 72hrs post transfection in RIPA buffer supplemented with NaF (5µM), Na2VO3 (5µM), β-glycerophosphate (5µM) and 1x Protease Inhibitor Cocktail (Cytoskeleton). Total protein concentration was determined using BCA Protein Assay Kit (Pierce). GFP-TRAP® magnetic agarose beads were equilibrated in ice cold dilution buffer (10 mM Tris/Cl pH 7.5; 150 mM NaCl; 0.5 mM EDTA) according to manufacturer’s instructions (Chromotek). Cell lysates containing 100µg of total protein were incubated with 10µl of equilibrated beads for 2 hours at 4°C with gentle agitation. Beads were washed three times with PBST buffer before analysis by Western blotting.
Magnetic Separation of Primary Human Lymphocytes
Peripheral blood mononuclear cells (PBMCs) were isolated from leukocyte cones (NHS Blood Transfusion service, St. George’s Hospital, London) by density gradient centrifugation with Lymphoprep™ density gradient medium (STEMCELL™ Technologies). Peripheral monocytes were isolated from PBMCs, using the human CD14+ magnetic beads (Miltenyi Biotech) according to manufacturer’s instructions. CD4+ T cells were isolated from the flow-through, using the human CD4+ T cell isolation kit (Miltenyi Biotech). CD14+ monocytes, and CD4+ T cells were either differentiated, or fixed directly after isolation for intracellular staining. To obtain M1 and M2 macrophages (M1/M2 MDMs), monocytes were treated with either granulocyte-macrophage colony stimulating factor (GM-CSF, 100ng/ml, Peprotech) or macrophage colony stimulating factor (M-CSF, 100ng/ml) for 7 days, with medium replenished on day 4. To obtain dendritic cells (DC), monocytes were treated with GM-CSF (50ng/ml) and IL-4 (50ng/ml) for 7 days, with medium replenished on day 4. Activated CD4+ T cells were obtained by stimulating freshly isolated CD4+ T cells at 1×106/ml with T cell activator CD3/CD28 Dynabeads (ThermoFisher), at a bead-cell-ratio of 1, for 7 days. Magnetic beads were removed prior to intracellular staining and flow cytometry.
Immunofluorescence
Transfected cells were washed with PBS and fixed in 2% paraformaldehyde/PBS for 10 minutes, at room temperature. Fixed cells were then permeabilized in 0.2% Triton™-X100/PBS for 20 minutes, at room temperature. Cells were incubated with primary antibodies in PBS containing 0.1% Triton-X100 and 2% BSA overnight at 4°C. After 3 washes in PBS, cells were then labeled with secondary antibodies in the same buffer for 1 hour, at room temperature, and washed 3 times with PBS. For confocal microscopy, nuclei were counterstained with Hoechst 33342 (2µM, ThermoFisher) for 5 minutes, at room temperature. Labeled cells were mounted with ProLong™ Diamond Antifade Mountant (ThermoFisher) and analyzed on a laser scanning confocal microscope LSM 710 (Carl Zeiss). Images were acquired with ZEN software and analyzed with ImageJ.
Imaging Flow Cytometry
Cells were fixed in FIX&PERM® Solution A (Nordic MUbio) for 30 minutes, and permeabilized with 0.2% Triton™-X 100/PBS. MDMs were blocked with human serum (1%). The staining buffer used was: 0.1% Triton™-X 100 0.5% FBS. Nuclei were stained with DAPI (1µg/ml) for two hours. Imaging flow cytometry was performed using the Amnis ImageStream®x Mark II Flow Cytometer (Merck) and INSPIRE® software (Amnis). A minimum of 10,000 events were collected for each sample, gating strategy is shown in Figure S7. IDEAS® software (Amnis) was used for analysis and to determine the ‘nuclear enrichment score’. The nuclear enrichment score is a comparison of the intensity of REAF fluorescence inside the nucleus to the total fluorescence intensity of the entire cell. A lower nuclear enrichment score indicates a lower proportion of overall REAF is located within the nucleus.
Statistics
Statistical significance in all experiments was calculated by Student’s t-test (two tailed) or ANOVA (indicated). Data are represented as mean ± standard deviation (error bars). GraphPad Prism and Excel were used for calculation and illustration of graphs.
Cells Synchronization
HeLa-CD4 were synchronized at the G2/M border by nocodazole (200ng/ml) for 16 hours. Where synchronized cells were infected with virus, an initial S phase block with thymidine (4mM) was induced for 24 hours followed by a PBS wash and a treatment with nocodazole (100ng/ml) for a further 16 hours. Collecting only those cells that were in suspension, as well as those that detached easily with a manual “shake-off”, enriched the population of mitotic cells.
Cell Cycle Analysis
Cell cycle phase distribution was determined by analysis of DNA content via either flow cytometry (BD FACS Canto™ II) or imaging flow cytometry. Cells were fixed in ice-cold ethanol (70%), treated with ribonuclease A (100µg/ml) and stained with propidium iodide (PI, 50µg/ml) or fixed in FIX&PERM® Solution A (Nordic MUbio) and stained with DAPI (1µg/ml). Mitotic cells were also identified by flow cytometry using the anti-phospho-histone H3 (Ser28) antibody. Cell lysates were assessed by Western blotting using the anti-phospho-histone H3 (Ser10/Thr11) antibody as an additional mitotic marker. Chromatin morphology and anti-phospho-histone H3 (Ser28) were used to determine the cells in indicated phases of the cell cycle and mitosis in confocal microscopy experiments. Cell cycle status of PM1 cells was determined via propidium iodide (PI) staining using FxCycle PI/RNse solution (ThermoFisher). Stained cells were analyzed on an NxT flow cytometer (ThermoFisher).
Evolutionary Analysis
To ascertain the evolutionary trajectory of REAF, we analyzed DNA sequence alignments of REAF from 15 species of extant primates using codeml (as implemented by PAML 4.2) (37). The evolution of REAF was compared to several NSsites models of selection, M1, M7 and M8a (neutral models with site classes of dN/dS <1 or ≥1) and M2, M8 (positive selection models allowing an additional site class with dN/dS >1). Two models of codon frequencies (F61 and F3×4) and two different seed values for dN/dS (ω) were used in the maximum likelihood simulations. Likelihood ratio tests were performed to evaluate which model of evolution the data fit significantly better. The p-value indicates the confidence with which the null model (M1, M7, M8a) can be rejected in favor of the model of positive selection (M2, M8). The alignment of REAF was analyzed by GARD to confirm the lack recombination during REAF evolution (38). Neither positively selected sites nor signatures of episodic diversifying selection were detected within REAF by additional evolutionary analysis by REL and FEL or MEME (39).
Data Availability
All RNA microarray data is available in the gene expression omnibus (GEO) database with accession number GSE54455.
Acknowledgements
This work was supported partly by an MRC Senior Non-Clinical Fellowship awarded to AM (G117/547) and PhD studentships awarded by QMUL Life Sciences Institute (LSI) (CEJ) and The Rosetrees Trust (JMG and CEJ, M665 and M275). RDS was supported by the Wellcome Trust-University of Edinburgh Institutional Strategic Support Fund. The monoclonal antibodies to HIV-1 p24 (EVA365 and 366) were provided by the EU Programme EVA Centre for AIDS Reagents, NIBSC, UK (AVIP Contract Number LSHP-CT-2004-503487). The Wellcome Trust (101604/Z/13/Z) funded the purchase of Amnis ImageStream™ imaging flow cytometer. We wish to thank N. Landau for the kind gift of VPL constructs.