|Home | About | Journals | Submit | Contact Us | Français|
The human SAMHD1 protein is a novel retroviral restriction factor expressed in myeloid cells. Previous work has correlated the deoxynucleotide triphosphohydrolase activity of SAMHD1 with its ability to block HIV-1 and SIVmac infection. SAMHD1 is comprised of the sterile alpha motif (SAM) and histidine–aspartic (HD) domains; however the contribution of these domains to retroviral restriction is not understood. Mutagenesis and deletion studies revealed that expression of the sole HD domain of SAMHD1 is sufficient to achieve potent restriction of HIV-1 and SIVmac. We demonstrated that the HD domain of SAMHD1 is essential for the ability of SAMHD1 to oligomerize by using a biochemical assay. In agreement with previous observations, we mapped the RNA-binding ability of SAMHD1 to the HD domain. We also demonstrated a direct interaction of SAMHD1 with RNA by using enzymatically-active purified SAMHD1 protein from insect cells. Interestingly, we showed that double-stranded RNA inhibits the enzymatic activity of SAMHD1 in vitro suggesting the possibility that RNA from a pathogen might modulate the enzymatic activity of SAMHD1 in cells. By contrast, we found that the SAM domain is dispensable for retroviral restriction, oligomerization and RNA binding. Finally we tested the ability of SAMHD1 to block the infection of retroviruses other than HIV-1 and SIVmac. These results showed that SAMHD1 blocks infection of HIV-2, feline immunodeficiency virus (FIV), bovine immunodeficiency virus (BIV), Equine infectious anemia virus (EIAV), N-tropic murine leukemia virus (N-MLV), and B-tropic murine leukemia virus (B-MLV).
Infection of primary macrophages and dendritic cells by Simian Immunodeficiency virus (SIVmac) requires the accessory protein Vpx, which is encoded in the SIV genome (Ayinde et al., 2010). SIVmac particles without Vpx (SIVΔVpx) are unable to infect primary macrophages. Vpx is essential for both SIV infection of primary macrophages and viral pathogenesis in vivo (Belshan et al., 2006; Fletcher et al., 1996; Gibbs et al., 1995; Hirsch et al., 1998). Vpx is incorporated into viral particles suggesting that it might be acting immediately after viral fusion in the target cells (Jin et al., 2001; Kappes et al., 1993; Park and Sodroski, 1995; Selig et al., 1999). Viral reverse transcription is prevented in primary macrophages when cells are infected with either Vpx-deficient SIVmac or HIV-2 (Bergamaschi et al., 2009; Fujita et al., 2008; Goujon et al., 2007; Kaushik et al., 2009; Srivastava et al., 2008). These experiments suggest that when Vpx is incorporated into viral particles, the virus overcomes a pre-reverse transcription block in macrophages and dendritic cells. Remarkably, Vpx also increases the ability of HIV-1 to infect macrophages and dendritic cells when Vpx is incorporated into HIV-1 particles or supplied in trans (Goujon et al., 2008; Sunseri et al., 2011). This suggests that the block imposed by macrophages to SIVΔVpx is similar to the one imposed by macrophages to HIV-1.
Recent work identified SAMHD1 as the protein that blocks infection of SIVΔVpx, HIV-2ΔVpx and HIV-1 before reverse transcription in macrophages and dendritic cells (Hrecka et al., 2011; Laguette et al., 2011). However, SIV, HIV-2 and HIV-1 expressing Vpx overcome the SAMHD1-dependent restriction (Hrecka et al., 2011; Laguette et al., 2011). Mechanistic studies have suggested that Vpx induces the proteasomal degradation of SAMHD1 (Berger et al., 2011; Hrecka et al., 2011; Laguette et al., 2011). Interestingly, the C-terminal region of SAMHD1 contains a Vpx binding motif, which is important for the ability of Vpx to degrade SAMHD1 (Laguette et al., 2012; Lim et al., 2012; Zhang et al., 2012). In addition, findings by Lim et al. (2012) have suggested that the SAM domain of SAMHD1 is important for the ability of Vpx to induce SAMHD1 degradation.
SAMHD1 comprises a sterile alpha motif (SAM) and a histidine–aspartic (HD) domain. SAM domains are protein interaction modules that mediate interaction with other SAM domains or non-SAM domain-containing proteins (Rice et al., 2009). Moreover, SAM domains in other proteins bind a specific sequence of DNA acting as either transcription activators or repressors (Qiao and Bowie, 2005). The HD domain is found in a super family of enzymes with a predicted phosphohydrolase activity (Zimmerman et al., 2008). In agreement, recent work has demonstrated that SAMHD1 is a dGTP-regulated deoxynucleotide triphosphohydrolase that might be involved in decreasing the overall cellular levels of triphosphodeoxynucleotides (Goldstone et al., 2011; Kim et al., 2012; Lahouassa et al., 2012b; Powell et al., 2011). This evidence agrees with the hypothesis that an overall decrease in the level of dNTPs is responsible for the block imposed to lentiviral infection.
The SAM and HD domains have been extensively studied on other proteins; however, the contribution of the different domains of SAMHD1 to HIV-1 restriction has not been investigated. This work explores the contribution of the different domains of SAMHD1 to HIV-1 restriction. By performing a series of deletion constructs, we found that the expression of the sole HD domain of SAMHD1 in U937 cells is sufficient to potently block HIV-1 infection. By contrast we showed that the SAM domain seems to be dispensable for restriction. Using biochemical assays, we demonstrated that the HD domain of SAMHD1 is the basic requirement for the ability of SAMHD1 to oligomerize and bind RNA. Furthermore, we explored the ability of SAMHD1 and the HD domain to block infection by different retroviruses.
The recently discovered restriction factor SAMHD1 comprises of a SAM and HD domain. Although the enzymatic activity of the HD domain is required for the ability of SAMHD1 to block HIV-1 and SIVmac infection (Goldstone et al., 2011; Lahouassa et al., 2012b; Powell et al., 2011), the contribution of the different regions of the SAMHD1 protein to restriction is not understood. To understand the contribution of the SAM and HD domain to lentiviral restriction, we generated a series of deletion constructs to find the minimal requirements for potent restriction (Fig. 1A). The different SAMHD1 variants were stably expressed in the human monocytic cell line U937 by using the LPCX vector system (Fig. 1B), as previously described (Brandariz-Nunez et al., 2012). U937 cells stably expressing SAMHD1 variants were induced to differentiation by PMA treatment (Schwende et al., 1996). Differentiated U937 cells were challenged with increasing amounts of HIV-1 containing the green fluorescent protein as a reporter (HIV-1-GFP) (Fig. 1C and Table 1). Overall these studies showed that expression of the SAMHD1 variant 112–582, which contains only the HD domain, is sufficient to block HIV-1 infection. In agreement, SAMHD1 variants Δ164–319, 1–328 and HD206AA that have either a defective or deleted HD domain completely lost their ability to block HIV-1 infection (Fig. 1C and Table 1). Because deletion of the SAM domain in the variant 112–626 did not affect HIV-1 restriction, this result suggested that the SAM domain is dispensable for restriction. In agreement with our previous findings that SAMHD1 nuclear localization is not necessary for restriction, the SAMHD1 variant 15–626, which is missing the nuclear localization signal (NLS), potently restricts HIV-1. Because SAMHD1 decreases the cellular levels of dNTPs in PMA-differentiated U937 cells, we next measured dNTP levels in differentiated U937 cells expressing the different SAMHD1 variants (Fig. 1D and Table 1). In agreement with the notion that the HD domain is required for enzymatic activity, we found that constructs with an intact HD domain decreased the cellular levels of dNTPs (Fig. 1D and Table 1).
Overall these mapping studies suggested that the HD domain of SAMHD1 is sufficient to achieve potent restriction of HIV-1. Moreover, these studies suggested that the SAM domain is dispensable for restriction.
Structural studies revealed that the recombinant HD domain protein of SAMHD1 produced in bacteria is dimeric (Goldstone et al., 2011). We wanted to know whether SAMHD1 is oligomerized in mammalian cells. Next we evaluated the biochemical ability of SAMHD1 containing a FLAG tag (SAMHD1-FLAG) to interact with SAMHD1 containing an HA tag (SAMHD1-HA). For this purpose, we cotransfected similar amounts of both SAMHD1-HA and SAMHD1-FLAG. Cells were lysed, and SAMHD1-FLAG was immunoprecipitated by using anti-FLAG beads. Proteins eluted by the FLAG peptide were separated by SDS-PAGE gels and analyzed by Western blotting using anti-HA and anti-FLAG antibodies (Fig. 2). In agreement with the notion that SAMHD1 forms oligomers, SAMHD1-FLAG interacts with SAMHD1-HA (Fig. 2). These results suggested that SAMHD1 forms oligomers when expressed in mammalian cells.
We next wanted to analyze the contribution of oligomerization to HIV-1 restriction by SAMHD1. Moreover, the ability of SAMHD1 to form oligomers is used as a surrogate assay to evaluate proper folding of the protein. For this purpose, we studied the ability of the different SAMHD1-FLAG variants to interact with a wild type SAMHD1-HA (Fig. 3 and Table 1). Interestingly, we observed that only SAMHD1-FLAG variants that contain an intact HD domain such as 15–626, Δ45–110, 112–626 and 112–582 showed the ability to interact with SAMHD1-HA (Fig. 3 and Table 1). By contrast, deletion constructs where the HD domain was affected were deficient for oligomerization (Fig. 3 and Table 1). These results suggested that the HD domain is the main determinant for the oligomerization ability of SAMHD1. Surprisingly, the SAMHD1 variant HD206AA, which is the mutation on the active site of SAMHD1, showed a reproducible decrease on oligomerization (Fig. 3 and Table 1). These results suggested that the HD domain of SAMHD1 is the main oligomerization determinant of the protein and is required for restriction.
Recent experiments demonstrated the ability of SAMHD1 to interact with nucleic acids (Goncalves et al., 2012). This work showed that SAMHD1 preferentially binds RNA by testing the ability of SAMHD1 from total mammalian or bacterial extracts to bind the interferon-stimulatory DNA sequence containing a phosphorothioate backbone (ISD-PS), which is an RNA analog. These experiments raised the possibility that SAMHD1 is indirectly interacting with RNA. Here we wanted to know whether purified SAMHD1 directly interacts with RNA. For this purpose, we tested the ability of full-length SAMHD1 purified from baculovirus-infected insect cells to bind RNA (Fig. S1A and Fig. 4A). As shown in Fig. 4A, purified SAMHD1 binds directly ISD-PS in a dose dependent manner. To control for the bona fide nature of the purified SAMHD1 protein, we tested the triphosphodeoxynucleotidase (dNTPase) activity of the protein. In agreement with the previous findings, our purified full-length recombinant SAMHD1 protein exhibits dGTP-dependent dTTPase activity (Fig. 4B). The recombinant SAMHD1 protein hydrolyzes 0.004 ng of dTTP/µg of SAMHD1/min. These results indicated a direct interaction between SAMHD1 and nucleic acids such as RNA.
Because SAMHD1 binds RNA, we tested the ability of RNA to modulate the dNTPase activity of SAMHD1. For this purpose, we measured the ability of purified recombinant SAMHD1 to hydrolyze α32P –TTP in the presence of the RNA analog ISD-PS (Fig. 4C). Interestingly, the presence of double-stranded RNA in the reaction inhibited the enzymatic activity of recombinant SAMHD1. These results suggested that the enzymatic activity of SAMHD1 is modulated by double-stranded RNA.
Next, we tested the ability of SAMHD1 variants expressed in human 293 cells to bind RNA. To control for the bona fide nature of the immunoprecipitated SAMHD1 protein from human cells, we tested its enzymatic activity (Fig. S1B). To assay the ability of SAMHD1 variants to bind RNA, we tested the ability of SAMHD1 variants produced in human 293T cells to bind ISD-PS (Fig. 4D and Table 1). Analysis of binding to RNA revealed that each and every SAMHD1 variant containing a complete HD domain interacts with RNA (Fig. 4D and Table 1). Moreover, we found that SAMHD1 variants containing the specific N-terminal (residues 112–328) or C-terminal region (residues 329–582) of the HD domain interact with RNA (Fig. 4D and Table 1). By contrast, the SAMHD1 variant 151–328 that exhibits a shorter N-terminal region of the HD domain no longer binds RNA. These results indicated that the N-terminal and C-terminal region of the HD domain are the minimal protein segments with the ability to bind RNA. However, it is also possible that the SAMHD1 variant 151–328 is a misfolded protein.
Previous work has demonstrated that SAMHD1 is a nuclearly localized protein (Brandariz-Nunez et al., 2012; Rice et al., 2009). Next we investigated the subcellular localization of the SAMHD1 variants studied in this work in the human HeLa cells (Fig. 5 and Table 1). In agreement with our findings suggesting that the NLS of SAMHD1 is located on the first 15 amino acids (11KRPR14) of the protein (Brandariz-Nunez et al., 2012), all SAMHD1 variants containing the NLS localized to the nucleus of HeLa cells (Fig. 5 and Table 1). By contrast, SAMHD1 variants missing the NLS were localized to either the cytoplasm or the cytoplasm and the nucleus (Fig. 5 and Table 1). In agreement with previous findings (Brandariz-Nunez et al., 2012), these results demonstrated that SAMHD1 blocks HIV-1 infection independent of its cellular localization.
Because the HD domain of SAMHD1 potently blocks HIV-1 infection, we tested the ability of SAMHD1 and the HD domain to block different retroviruses. For this purpose, we challenged PMA-treated U937 cells expressing either SAMHD1 or the HD domain (112–582) with increasing amounts of the indicated retrovirus containing the green fluorescent protein (GFP) as a reporter of infection (Fig. 6A). As control, we performed similar infections in PMA-treated U937 cells containing the empty vector LPCX. As expected, wild type SAMHD1 blocked HIV-1 infection; however, the sole HD domain was more effective in blocking HIV-1. Similarly, the HD domain of SAMHD1 more potently blocked infection when compared to wild type SAMHD1 when the following retroviruses were tested (Fig. 6A): HIV-2, SIVmac, SIVmacΔVpx, bovine immunodeficiency virus (BIV), feline immunodeficiency virus (FIV), N-tropic murine leukemia virus (N-MLV) and B-tropic murine leukemia virus (B-MLV). Interestingly, equine infectious anemia virus (EIAV) seems to partially overcome SAMHD1-mediated restriction suggesting that EIAV has a mechanism to overcome SAMHD1. To confirm these observations using cells that endogenously express SAMHD1, we challenged THP-1 cells either treated with PMA or DMSO with increasing amounts of the indicated retroviruses (Fig. 6B). PMA-treated THP-1 cells potently block infection by HIV-1, HIV-2, SIVmacΔVpx, BIV, FIV, EIAV, N-MLV and B-MLV. By contrast, SIVmac infection, which carries Vpx, was not blocked by PMA-treated THP-1 cells when compared to SIVmacΔVpx. These results demonstrated the wide range of retroviruses that can be inhibited by either expression of SAMHD1 or the sole HD domain.
Here we studied the contribution of the different domains of SAMHD1 to its antiviral activity. From these studies, we have learned the following: (1) the SAM domain of SAMHD1 seems to be dispensable for retroviral restriction, (2) the sole HD domain is sufficient for potent restriction of HIV-1 and other retroviruses, (3) we have established a method to measure the ability of SAMHD1 to oligomerize in mammalian cells, (4) an intact HD domain is required for SAMHD1 oligomerization, (5) the N-terminal (residues 112–328) and C-terminal (residues 329–582) regions of the HD domain are the minimal protein sequences with the ability to bind RNA, (6) cytosolic SAMHD1 variants are potent restrictors, (7) the HD domain restricts retroviral infection more potently when compared to the full-length SAMHD1, and (8) SAMHD1 and the HD domain of SAMHD1 potently block infection of HIV-1, HIV-2, SIVmacΔVpx, FIV, BIV, N-MLV and B-MLV.
SAM domains can potentially interact with each other, with other domains and with RNA (Qiao and Bowie, 2005; Rice et al., 2009). However, the SAM domain of SAMHD1 seems to be dispensable for the ability of SAMHD1 to block retroviral infection or to bind RNA. Our oligomerization assays revealed that full-length SAMHD1 does not interact with the SAMHD1 variant (1–150) that contains the SAM domain, suggesting that the SAM domain of SAMHD1 might not be interacting with itself in the context of a dimeric SAMHD1; however, the SAMHD1 variant 1–150 that contains the SAM domain might be a misfolded protein. This work did not eliminate the possibility that the SAM domain is involved in the regulation of SAMHD1’s enzymatic activity.
The fact that expression of the sole HD domain of SAMHD1 is sufficient to achieve potent retroviral restriction agrees with the notion that dNTPs depletion is the only necessary step required for restriction (Goldstone et al., 2011; Kim et al., 2012; Lahouassa et al., 2012b; Powell et al., 2011). Interestingly, cycling U937 cells expressing the HD domain of SAMHD1, which were not induced to differentiate by PMA, were permissive to HIV-1 infection (data not shown). These results suggested that in cycling cells the sole HD domain undergoes a regulation that inhibits its enzymatic activity. An alternative possibility is that the levels of dNTPs in cycling U937 cells are high enough that having an enzymatically active HD domain does not influence the overall levels of dNTPs. This alternative possibility could be tested by studying the sensitivity of HIV-1 infection in cycling U937 cells expressing the HD domain to hydroxyurea, which is an inhibitor of ribonucleotide reductase (RNR). The enzyme RNR is responsible for the synthesis of deoxynucleotides, which provide the substrate for synthesis of dNTPs. If the HD domain of SAMHD1 expressed in cycling U937 cells is an enzymatically active HD domain, infection of HIV-1 might be more sensitive to hydoxyurea when compared to control U937 cells.
In agreement with the dimeric structure of SAMHD1 (Goldstone et al., 2011), we demonstrated that SAMHD1 expressed in mammalian cells is an oligomer by an assay that measures SAMHD1 oligomerization. However, our oligomerization assay has the limitation that it does not provide information regarding the stoichiometry of the oligomer (i.e. dimer, trimer, etc). Besides this drawback, this assay could be used as a surrogate to evaluate protein folding. Using this assay, we demonstrated that an intact HD domain is required for oligomerization. Interestingly, we observed that the SAMHD1 variant HD206AA, which is a mutation on the active site of SAMHD1, is defective for oligomerization (Fig. 3). This observation raises the possibility that this mutant is enzymatically defective because of a defect on oligomerization, suggesting that oligomerization contributes to the enzymatic activity of SAMHD1. It will be important to determine whether dimerization of the HD domain is required for enzymatic activity and retroviral restriction.
Previous work demonstrated the ability of SAMHD1 to interact with RNA (Goncalves et al., 2012). However, these experiments used unpurified SAMHD1 protein in mammalian or bacterial extracts raising the possibility that the observed interaction between SAMHD1 and RNA is mediated by a second protein present in the extract. Here we used purified SAMHD1 protein from insect cells, and showed that enzymatically active SAMHD1 has the ability to directly interact with RNA. Furthermore, our mapping studies revealed that the HD domain is the basic motif required for the ability of SAMHD1 to bind RNA, as previously suggested (Goncalves et al., 2012). Additionally, we found SAMHD1 variants that are able to bind RNA but unable to oligomerize, suggesting that oligomerization is not required for RNA binding. Surprisingly, RNA binding to SAMHD1 dramatically affected its enzymatic activity bringing up the possibility that when SAMHD1 interacts with the RNA of a pathogen it allows the recovery of cellular dNTPs, which might be beneficial for the host response.
In agreement with our previous observations (Brandariz-Nunez et al., 2012), the subcellular localization of SAMHD1 variants did not influence SAMHD1’s ability to block retroviral infection. These results suggested that dNTPs are depleted from either the nucleus or cytoplasm.
Our findings showed that the sole HD domain restricts more potently than the full-length SAMHD1. These results suggest that the HD domain alone is more active as an enzyme, which indirectly implies that the HD domain (SAMHD1 variant 112–582) is missing regulatory parts of the protein that control enzymatic activity. Future experiments will address the possibility that other regions of the SAMHD1 protein regulate the enzymatic activity of the HD domain.
Human THP-1(ATCC#TIB-202) and U937 (ATCC#CRL-1593) cells were grown in RPMI supplemented with 10% (v/v) fetal bovine serum and 1% (v/v) penicilin/streptomycin. Human HeLa cells (ATCC# CCL-2) were grown on DMEM supplemented with 10% fetal bovine serum and and 1% (w/v) penicilin/streptomycin. LPCX-SAMHD1-HA and LPCX-SAMHD1-FLAG plasmids expressing the codon optimized SAMHD1 fused to either HA or FLAG epitope were previously described (Brandariz-Nunez et al., 2012). The plasmids expressing SAMHD1 deletion mutants were created using pLPCX-SAMHD1-FLAG as template and specific primers. PCR products were digested and cloned into the EcoRI and ClaI sites of pLPCX. Orientation of the inserts was confirmed by sequencing and restriction analysis.
Retroviral vectors encoding wild type or mutant SAMHD1 proteins fused to FLAG were created using the LPCX vector (Clontech). Recombinant viruses were produced in 293T cells by co-transfecting the LPCX plasmids with the pVPack-GP and pVPack-VSV-G packaging plasmids (Stratagene). The pVPack-VSV-G plasmid encodes the vesicular stomatitis virus G envelope glycoprotein, which allows efficient entry into a wide range of vertebrate cells (Yee et al., 1994). Transduced human monocytic U937 cells were selected in 0.4 µg/ml puromycin (Sigma).
Cellular proteins were extracted with radioimmunoprecipitation assay (RIPA) as previously described (Lienlaf et al., 2011). Detection of proteins by Western blotting was performed using anti-FLAG (Sigma), anti-SAMHD1 (Abnova), anti-GAPDH (Ambion) or anti-HA (Sigma). Secondary antibodies against rabbit and mouse conjugated to Alexa Fluor 680 were obtained from Li-Cor. Bands were detected by scanning blots using the Li-Cor Odyssey Imaging System in the 700 nm channel.
Recombinant retroviruses expressing GFP, pseudotyped with the VSV-G glycoprotein, were prepared as described (Diaz-Griffero et al., 2008). For infections, 6 × 104 cells seeded in 24-well plates were either treated with 10 ng/ml phorbol-12-myristate-3-acetate (PMA) or DMSO for 16 h. PMA stock solution was prepared in DMSO at 250 µg/ml. Subsequently, cells were incubated with the indicated retrovirus for 48 h at 37 °C. The percentage of GFP-positive cells was determined by flow cytometry (Becton Dickinson). Viral stocks were titrated by serial dilution on dog Cf2Th cells.
Approximately 1.0 × 107 human 293T cells were cotransfected with plasmids encoding FLAG-tagged and HA-tagged mutant and wild type SAMHD1 proteins. After 24 h, cells were lysed in 0.5 ml of whole-cell extract (WCE) buffer (50 mM Tris [pH 8.0], 280 mM NaCl, 0.5% IGEPAL, 10% glycerol, 5 mM MgCl2, 50 µg/ml ethidium bromide, 50 U/ml benzonase tail [Roche]). Lysates were centrifuged at 14,000 rpm for 1 h at 4 °C. Post-spin lysates were then pre-cleared using protein A-agarose (Sigma) for 1 h at 4 °C; a small aliquot of each of these lysates was stored as input. Pre-cleared lysates containing the tagged proteins were incubated with anti-FLAG-agarose beads (Sigma) for 2 h at 4 °C. Anti-FLAG-agarose beads were washed three times in WCE buffer, and immune complexes were eluted using 200 µg of FLAG tripeptide/ml in WCE buffer. The eluted samples were separated by SDS-PAGE and analyzed by Western blotting using either anti-HA or anti-FLAG antibodies.
Nucleic-acid binding assay was performed as previously described (Goncalves et al., 2012). In brief, the synthetic DNA phosphorothioate-containing interferon-stimulatory DNA (ISD-PS), which is an RNA analog, was synthesized with a 5′-biotin tag using the following primers:
Sense and antisense primers were incubated at 65 °C for 20 min, and primers were allowed to anneal by cooling down to room temperature. Annealed primers were immobilized on an Ultralink Immobilized Streptavidin Plus Gel (Pierce). Cells were lysed using TAP lysis buffer (50 mM Tris pH 7.5, 100 mM NaCl, 5% glycerol, 0.2% NP-40, 1.5 mM MgCl2, 25 mM NaF, 1 mM Na3VO4, protease inhibitors) and lysates were cleared by centrifugation. Cleared lysates (Input) were incubated with immobilized nucleic acids at 4 °C on a rotary wheel for 2 h in the presence of 10 µg/ml of Calf-thymus DNA (Sigma) as a competitor. Unbound proteins were removed by three consecutive washes in TAP lysis buffer. Bound proteins to nucleic acids (Bound) were eluted by boiling samples in SDS sample buffer (63 mM Tris–HCl, 10% Glycerol 2% SDS, 0.0025% Bromophenol Blue) and analyzed by Western blotting using anti-FLAG antibodies (Sigma).
2 × 106 to 3 × 106 cells were collected for each cell type. Cells were washed twice with 1× PBS, pelleted and resuspended in ice cold 65% methanol. Cells were vortexed for 2 min and incubated at 95 °C for 3 min. Cells were centrifuged at 14000 rpm for 3 min and the supernatant was transferred to a new tube for the complete drying of the methanol in a speed vac. The dried samples were resuspended in molecular grade dH2O. An 18-nucleotide primer labeled at the 5′ end with 32 P (5′-GTCCCTGTTCGGGCGCCA-3–) was annealed at a 1:2 ratio to four different 19-nucleotide templates (5′-NTGGCGCCCGAACAGGGAC-3′), where ‘N’ represents the nucleotide variation at the 5′ end. Reaction condition contains 200 fmoles of template primer, 2 µl of 0.5 mM dNTP mix for positive control or dNTP cell extract, 4 µl of excess HIV-1 RT, 25mM Tris–HCl, pH 8.0, 2 mM dithiothreitol, 100 mM KCl, 5 mM MgCl2, and 10 µM oligo(dT) to a final volume of 20 µL. The reaction was incubated at 37 °C for 5 min before being quenched with 10 µL of 40 mM EDTA and 99% (vol/vol) formamide at 95 °C for 5 min. The extended primer products were resolved on a 14% urea–PAGE gel and analyzed using a phosphoimager. The extended products were quantified using QuantityOne software to quantify percent volume of saturation. The quantified dNTP content of each sample was accounted for based on its dilution factor, so that each sample volume was adjusted to obtain a signal within the linear range of the assay (Kim et al., 2012; Lahouassa et al., 2012a).
5.8 µM SAMHD1 was incubated with 1 mM dTTP and 100 µM dGTP in reaction buffer (50 mM Tris–HCl pH 8, 50 mM KCl, 5 mM MgCl2, 0.1% Triton-X100) for 2 h at 37 °C. Reactions were terminated by incubation at 75 °C for 10 min and diluted 20-fold into 12.5% acetonitrile containing 58 µM dCMP as a reference control. HPLC: Dionex DNAPac PA100 column (4 × 50 mm2) was equilibrated with running buffer (25 mM Tris–HCl pH 8, 0.5% acetonitrile) for 10 min, 99 µl sample was injected and eluted with a linear gradient of 240 mM NH4Cl for 12 min, run at an isocratic gradient with 240 mM NH4Cl for 5 min, and column was again equilibrated with running buffer (Beckman Coulter System Gold 126 Solvent Module). Absorbance was measured with a Beckman Coulter System Gold 166 Detector at 254 nm and dTTP abundance was determined by integrating the area under each peak using 32 Karat 8.0 Software. dTTP levels were normalized by the amount of dCMP detected in each diluted sample.
Recombinant SAMHD1 (5 µM) was incubated with or without 100 µM dGTP, with or without 6 µM dsRNA analog, 500 µM dTTP and 0.25 µl α32P-dTTP (PerkinElmer) in SAMHD1 reaction buffer (50 mM Tris–HCl pH 8, 50 mM KCl, 5 mM MgCl2, 0.1% Triton-X 100) in a 17.5 µl final volume. Reactions were initiated by addition of SAMHD1, incubated for 1 h at 37 °C, and terminated by incubation for 10 min at 70 °C. The no enzyme control reaction and the antarctic phosphatase reaction contained both dGTP and dsRNA. The antarctic phosphatase reaction (2 µl, New England BioLabs) was used to show the mobility of monophosphates on the plate as a comparison to triphosphate mobility. Reactions were spotted (0.5 µl) on a TLC PEI Cellulose F plate (EMD Chemicals) and separated in a 0.8 M LiCl solvent. Product formation was analyzed on a Bio-Rad Personal Molecular Imager.
Transfections of cell monolayers were performed using Lipofectamine Plus reagent (Invitrogen), according to the manufacturer’s instructions. Transfections were incubated at 37 °C for 24h. Indirect immunofluorescence microscopy was perfomed as previously described (Diaz-Griffero et al., 2002). Transfected monolayers grown on coverslips were washed twice with PBS1X (137 mM NaCl, KCl 2.7 mM, Na2HPO4 · 2H2O 10mM, KH2PO4 mM) and fixed for 15 min in 3.9% paraformaldehyde in PBS1X. Fixed cells were washed twice in PBS1X, permeabilized for 4 min in permeabilizing buffer (0.5% Triton X-100 in PBS), and then blocked in PBS1X containing 2% bovine serum albumin (blocking buffer) for 1 h at room temperature. Cells were then incubated for 1 h at room temperature with primary antibodies diluted in blocking buffer. After three washes with PBS, cells were incubated for 30 min in secondary antibodies and 1 µg of DAPI (49, 69-diamidino-2-phenylindole)/ml. Samples were mounted for fluorescence microscopy by using the ProLong Antifade Kit (Molecular Probes, Eugene, OR). Images were obtained with a Zeiss Observer Z1 microscope using a 63× objective, and deconvolution was performed using the software AxioVision V184.108.40.206 (Carl Zeiss Imaging Solutions).
Recombinant baculovirus was generated using the Bac-to-Bac system (Invitrogen) following the manufacturer’s instructions. For construction of the transfer vector, full-length SAMHD1 was amplified by PCR using specific primers and cloned into the EcoRI and XbaI sites of the pFastBac-1 vector containing an OSF epitope (One-Strep and Flag) previously cloned into BamHI and EcoRI sites of the plasmid. The correctness of the construct was confirmed by sequencing. The pFastBac1-OSF-SAMHD1 vector was recombined into the bacmid by transformation of DH10 Bac cells (Invitogen). The recombinant bacmid was obtained according to the supplier’s protocol. Single colonies were grown to stationary phase in 2 ml of LB medium supplemented with the necessary antibiotics (50 µg/ml kanamycin, 7 µg/ml gentamicin, and 10 µg/ml tetracycline shaking at 300 rpm) for up 24 h at 37 °C. Bacterial pellets were resuspended in 0.3 ml of Solution I [15 mM Tris–HCl (pH 8.0), 10 mM EDTA, 100 µg/ml RNase A] and 0.3 ml of Solution II (0.2 N NaOH, 1% SDS) and incubated at room temperature for 5 min. Samples were treated with 0.3 ml of 3 M potassium acetate (pH 5.5) on ice for 10 min and cebtrifugated for 10 min at 14,000g. The supernatant was transferred to a tube containing isopropanol and incubated on ice for 10 min. The DNA was pelleted by centrifugation for 15 min at 14,000g at room temperature. The pellets were washed with 0.5 ml of 70% ethanol. The pellet was air dried at room temperature. DNA samples were resuspended in 40 µl milli-Q water. The presence of the SAMHD1 gene in the bacmids was confirmed by PCR using the pUC/M13 forward and reverse primers described by Invitrogen. The PCR products were sequenced to confirm the presence of the insert containing the OSF-SAMHD1 construct. The bacmid was then used to generate the corresponding recombinant baculovirus according to the supplier’s protocol as described (Brandariz-Nunez et al., 2010a, b). Briefly, the bacmid was transfected into Spodoptera frugiperda (SF9) cells using Lipofectamine Plus reagent (Invitrogen). Cells were incubated at 28 °C in SF-900 II serum-free insect cell medium (Invitrogen) for 3 days, and recombinant viruses expressing full-length SAMHD1 were havested (P1 viral stock). For amplification of P1 viral stock, a monolayer culture (2 × 106 cells/ml) was infected with 0.5 ml of a P1 viral stock. The P2 viral stock was harvested 3 days post-infection. Expression of OSF-SAMHD1 was analyzed in SF9 cells infected with P2 viral sotcks by coomassie stain and Western blotting using anti-FLAG and anti-SAMHD1 antibodies.
7.5 × 108 Sf9 insect cells growing in suspension were infected with 5 PFU/cell of the recombinant baculovirus and incubated at 28 °C for 72 h. All subsequent steps were carried out at 4 °C. Cells were collected by centrifugation at 500g for 8 min, resuspended in 60 ml of lysis buffer (250 mM NaCl, 50 mM Tris [pH 8.0], 1.5% Triton X-100, 1 mM TCEP, and mammalian protease inhibitor cocktail [Sigma]). The lysate was clarified by centrifugation (20,000g, 40 min), filtered (0.45 mm), and incubated in 1.5 ml StrepTactin Superflow affinity resin (Qiagen). The bound protein was washed three three-times with 15 ml of buffer (50 mM NaCl, 50 mM Tris [pH 8.0], 1 mM TCEP) and eluted with washing buffer supplemented with 2.5 mM d-desthiobiotin. The eluate was analyzed by SDS-PAGE and Western blotting using anti-FLAG or anti-SAMHD1 antibodies.
This work was funded by the NIH No. R01 AI087390 to F.D.-G, and a K99/R00 Pathway to Independence Award to F.D.-G. from the National Institutes of Health No. 4R00MH086162-02.
Appendix A. Supporting information
Supplementary data associated with this article can be found in the online version at http://dx.doi.org/10.1016/j.virol.2012.10.029.