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J Virol. 2008 December; 82(23): 11889–11901. Published online 2008 September 25. doi: 10.1128/JVI.01537-08. | PMCID: PMC2583693 |
Copyright © 2008, American Society for Microbiology Equine Infectious Anemia Virus Resists the Antiretroviral Activity of Equine APOBEC3 Proteins through a Packaging-Independent Mechanism ![[down-pointing small open triangle]](/corehtml/pmc/pmcents/x25BF.gif) Hal P. Bogerd,1 Rebecca L. Tallmadge,2 J. Lindsay Oaks,2 Susan Carpenter,2 and Bryan R. Cullen1* Department of Molecular Genetics and Microbiology and Center for Virology, Duke University Medical Center, Durham, North Carolina 27710,1 Department of Veterinary Microbiology and Pathology, Washington State University, Pullman, Washington 991642 Received July 21, 2008; Accepted September 19, 2008. Equine infectious anemia virus (EIAV), uniquely among lentiviruses, does not encode a vif gene product. Other lentiviruses, including human immunodeficiency virus type 1 (HIV-1), use Vif to neutralize members of the APOBEC3 (A3) family of intrinsic immunity factors that would otherwise inhibit viral infectivity. This suggests either that equine cells infected by EIAV in vivo do not express active A3 proteins or that EIAV has developed a novel mechanism to avoid inhibition by equine A3 (eA3). Here, we demonstrate that horses encode six distinct A3 proteins, four of which contain a single copy of the cytidine deaminase (CDA) consensus active site and two of which contain two CDA motifs. This represents a level of complexity previously seen only in primates. Phylogenetic analysis of equine single-CDA A3 proteins revealed two proteins related to human A3A (hA3A), one related to hA3C, and one related to hA3H. Both equine double-CDA proteins are similar to hA3F and were named eA3F1 and eA3F2. Analysis of eA3F1 and eA3F2 expression in vivo shows that the mRNAs encoding these proteins are widely expressed, including in cells that are natural EIAV targets. Both eA3F1 and eA3F2 inhibit retrotransposon mobility, while eA3F1 is a potent inhibitor of a Vif-deficient HIV-1 mutant and induces extensive editing of HIV-1 reverse transcripts. However, both eA3F1 and eA3F2 are weak inhibitors of EIAV. Surprisingly, eA3F1 and eA3F2 were packaged into EIAV and HIV-1 virions as effectively as hA3G, although only the latter inhibited EIAV infectivity. Moreover, all three proteins bound both the HIV-1 and EIAV nucleocapsid protein specifically in vitro. It therefore appears that EIAV has evolved a novel mechanism to specifically neutralize the biological activities of the cognate eA3F1 and eA3F2 proteins at a step subsequent to virion incorporation. Equine infectious anemia virus (EIAV) is a macrophage tropic lentivirus that causes a lifelong persistent infection in horses and other equids ( 62, 63). Experimental infection of horses induces a reproducible clinical disease course and provides a useful model to examine mechanisms of immune control and virus persistence during long-term lentivirus infection ( 37, 45). Upon infection, most horses suffer an early episode of acute disease, including fever, viremia, and thrombocytopenia associated with high levels of virus replication in tissue macrophages ( 31, 51, 63). Resolution of acute disease is concurrent with the appearance of virus-specific cytotoxic T lymphocyte and neutralizing antibody and a decrease in plasma viremia ( 29, 46, 66). Recurrent episodes of high-titer plasma viremia and associated disease often occur within the first year after initial infection but generally abate in frequency and severity with time. Thereafter, most horses enter a lifelong clinically unapparent stage with no evident compromise to their long-term health. Notably, these horses do not eliminate the virus but become lifelong carriers. EIAV is similar to other lentiviruses, such as human immunodeficiency virus type 1 (HIV-1), in its overall genome organization; however, EIAV is genetically the simplest lentivirus and contains only three regulatory/accessory genes: tat, rev, and S2. The Tat and Rev proteins of EIAV are functionally homologous to HIV-1 Tat and Rev but differ somewhat in the organization of functional domains and the RNA structures that regulate viral gene expression ( 3, 10, 11, 24, 35, 36). Of particular interest, EIAV is unique among lentiviruses in that it lacks a vif gene. The APOBEC3 (A3) family of cytidine deaminases (CDAs) was first identified in humans, where seven distinct A3 genes are found in close proximity on human chromosome 22 ( 15, 32). The ability of A3 proteins to function as potent inhibitors of retroviral infectivity was first defined for human A3G (hA3G), which inhibits the infectivity of HIV-1 variants lacking an intact vif gene (HIV-1ΔVif) ( 16, 64). Conversely, wild-type HIV-1 is largely unaffected by hA3G expression. Subsequent research has demonstrated that the HIV-1 Vif protein directly binds to hA3G and induces its degradation via the proteasome ( 14, 42, 65, 72). In the absence of Vif, hA3G interacts with the nucleocapsid (NC) domain of the HIV-1 Gag polyprotein and is specifically packaged into progeny virion particles ( 2, 12, 60, 73). Upon subsequent infection of a susceptible cell, hA3G interferes with retroviral reverse transcription, at least in part by inducing the deamination of dC residues on the proviral DNA minus strand, resulting in dU residues that then template the introduction of A residues, instead of G residues, on the proviral plus strand ( 30, 40, 71, 74). While massive mutagenesis of reverse transcripts by hA3G is clearly an important part of this protein's inhibitory activity ( 47, 61), evidence also exists that hA3G can inhibit retroviral infectivity in the absence of detectable editing ( 50). In addition to hA3G, humans also encode six other A3 family members, named hA3A, hA3B, hA3C, hA3D/E, hA3F, and hA3H ( 15, 32, 52). These proteins can be subdivided into three proteins that contain a single-CDA active site consensus sequence (hA3A, hA3C, and hA3H) and four that contain two tandem CDA domains (hA3B, hA3D/E, hA3F, and hA3G). At least three hA3 proteins can function as potent inhibitors of a range of retroviruses, i.e., hA3B, hA3F, and hA3G ( 4, 20, 38, 70), and these are all double-CDA-domain proteins. On the other hand, single-CDA-domain proteins, such as hA3A, can function as efficient inhibitors of retrotransposon mobility ( 7, 13). In the case of hA3G, mutational analysis has revealed that the amino-terminal CDA is enzymatically inactive and functions to recruit hA3G into retroviral virion particles, while the enzymatically active carboxy-terminal CDA is required for inhibition of infectivity ( 28, 49). Recent data suggest that the segregation of A3 proteins into a virion-packaging domain and an inhibitory domain may greatly facilitate their antiretroviral activities ( 27). There is considerable interest in understanding the selective forces that shape the evolution and activity of A3 genes. A number of A3 genes have been identified in nonprimate species, and several of these have been characterized for antiviral activity against a range of viruses and/or retroelements. At least one double-CDA-domain A3 protein with antiviral activity has been identified in mice, cows, sheep, and pigs ( 33, 34, 41, 70). The mouse genome contains one A3 gene ( 15, 69); however, genomic characterization of the A3 genes in cows, sheep, and pigs is incomplete. In cats, where the A3 locus has been examined in more detail, there are four distinct A3 genes encoding five A3 proteins. Three of the A3 proteins were shown to be active against feline foamy virus, whereas the other two proteins were active against feline immunodeficiency virus and feline leukemia virus ( 39, 48). Sequences of equine and canine A3 genes have been included in phylogenetic studies ( 15, 52), but no functional data are available. While HIV-1 can block the inhibitory activities of hA3G and hA3F by using Vif to induce their degradation ( 16), other retroviruses have evolved other mechanisms to permit their replication to occur unimpeded by host cell A3 proteins. In the case of foamy viruses, the viral Bet protein has been found to directly bind to hA3G and to sequester this protein away from progeny virions without inducing its degradation ( 39, 58). Human T-cell leukemia virus type I and Mason-Pfizer monkey virus have both been reported to selectively exclude A3 proteins encoded by their healthy host species from virion particles ( 17, 19), although Mason-Pfizer monkey virus packages, and is inhibited by, murine A3 (mA3). Finally, murine leukemia virus (MLV) has also been reported to discriminate against its cognate A3 protein, mA3, but not against the heterologous hA3G protein in virion incorporation ( 1, 20, 34). MLV has also been reported to be inhibited less effectively by virion-incorporated mA3 than by hA3G, even when virions containing equivalent levels of each protein were analyzed ( 9, 57). In this paper, we characterize the equine A3 gene locus and ask how the lentivirus EIAV, which uniquely lacks a vif gene, is able to grow in the presence of these equine A3 gene products. We report that horses express six distinct A3 genes, four of which encode single-CDA domains and two of which, equine A3F1 (eA3F1) and eA3F2, contain double-CDA domain proteins. While eA3F1 is a potent inhibitor of several heterologous retroviruses, including HIV-1ΔVif, both eA3F1 and eA3F2 are weak inhibitors of EIAV. Inhibition of HIV-1 infectivity by eA3F1 is associated with high levels of editing of HIV-1 reverse transcripts. Both eA3F1 and eA3F2 are enzymatically active CDAs, and both can function as effective inhibitors of retrotransposon mobility. Moreover, mRNAs encoding both eA3F1 and eA3F2 are expressed at readily detectable levels in vivo in tissues that support EIAV replication, including macrophages. Surprisingly, we demonstrate that eA3F1 and eA3F2 are incorporated into EIAV virions as effectively as hA3G, although only the heterologous hA3G protein can inhibit EIAV infectivity. Characterization of the equine A3 gene locus. The human double-CDA domain A3 genes with known antiviral activities (hA3B, hA3F, and hA3G) were used to perform BLASTN and/or TBLASTN searches of the NCBI equine sequence databases. A total of 59 expressed sequence tag (EST) clones were identified from peripheral blood lymphocyte libraries (Table 1). Representative clones were obtained from the respective libraries, sequenced in full, and used to BLAST the horse genome sequence through NCBI (EquCab1; accession number NW_001799702). This identified four distinct genes on chromosome 28, each containing a single-CDA domain. Two additional equine A3 genes containing double-CDA domains, EcA3F1 and EcA3F2, were identified following reverse transcription-PCR (RT-PCR) amplification of total RNA isolated from both fetal equine kidney (FEK) cells and equine peripheral blood mononuclear cells (PBMCs). Subsequently, partial cDNAs of EcA3F1 and EcA3F2 were also obtained by RT-PCR amplification using RNA samples derived from additional unrelated horses. | TABLE 1.GenBank accession numbers of equine APOBEC sequences |
To determine phylogenetic relationships among the A3 family members, the equine deaminase domains were aligned with CDA domains of several species, including hA3A ( NM_145699), hA3B ( NM_004900), hA3C ( NM_014508), hA3D/E ( NM_152426), hA3F ( NM_145298), hA3G ( NM_021822), hA3H ( NM_181773), cow A3F ( DQ974646), sheep A3F ( DQ974645), pig A3F ( DQ974647), mA3 ( NM_030255), rat A3 ( NM_001033703), cat A3Ca ( EU109281), cat A3Cb ( EU109281), cat A3Cc ( EU109281), cat A3H ( EU109281), hAID ( NM_020661), hAPOBEC1 ( NM_001644), and hAPOBEC2 ( NM_006789). Phylogenetic trees were constructed by the neighbor-joining method using a p-distance model, and the reliability of branching orders was assessed by bootstrap analysis using 1,000 replicates with MEGA4 software ( 68). Phylogenetic analyses were also performed with equine APOBEC family members AID, APOBEC1, and APOBEC2 to ensure the sequences identified were indeed A3 family members. Analysis of A3 mRNA expression patterns. Tissues (brain, kidney, liver, bone marrow, lung, lymph node, and spleen) were collected postmortem from a clinically healthy pony, snap-frozen in liquid nitrogen, and stored at −80°C until use. PBMCs were obtained from a clinically healthy Arabian horse and isolated by density gradient centrifugation (Histopaque; Sigma). Monocytes and monocyte-derived macrophages (MDMs) were derived from the whole blood of a clinically healthy Arabian horse using previously described differential adherence methods ( 56). Equine dermal (ED) cells were obtained from the ATCC (CCL57), and FEK cells were isolated and cultured from primary cells as previously described ( 55). Total RNA was extracted using TRIzol reagent (Invitrogen). RNA pellets were resuspended in 80 μl of water and 2 μl of RNase inhibitor (RNaseOUT; Invitrogen) and treated with DNase (Turbo DNase-free kit; Applied Biosystems). cDNA was produced from 1 μg of total RNA by random hexamer priming and 200 U of MLV reverse transcriptase (Invitrogen) for 1 h at 37°C. Reactions were terminated by being boiled for 5 min, and the cDNA was stored at −20°C until used. Reverse transcription efficiency was verified in the cDNA preparations by quantitative real-time PCR for 18S rRNA ( 23). For analyses of mRNA expression, 1 μl of cDNA (approximately 40 ng) was amplified with eA3F1- or eA3F2-specific primers designed to exclusively amplify double-CDA-containing transcripts. The sequences of the eA3F1 primers were 5′-CTGGCCGTGATGTTGCG-3′ and 5′-GCAGTCTCTGAAATCCCA-3′, amplifying a 579-bp product. The sequences of the eA3F2 gene primers were 5′-CATGGTCTTCAGGGATTTCAG-3′ and 5′-GAAGCGCTCACTTGAGAATC-3′, resulting in a 683-bp product. Expression plasmids encoding eA3F1 or eA3F2 were included as controls for specificity. The β-actin primers were 5′-GCTCGTCGTCGACAACGGCT-3′ and 5′-CAAACATGATCTGGGTCATCTTCTC-3′. Amplification reactions consisted of 20 mM Tris-HCl (pH 8.4), 50 mM KCl, 1.5 mM MgCl2, 0.2 mM deoxynucleoside triphosphates, 0.2 μM of each primer, and 2.5 U of Taq DNA polymerase (Invitrogen, Carlsbad, CA). Cycling conditions were as follows: initial denaturation at 95°C for 2 min and 40 cycles of 95°C for 30 s, 60°C for 10 s, and 72°C for 1 min, followed by a final extension of 72°C for 10 min. One-fifth of the reaction mixture volume was run on 1% agarose gel and stained with ethidium bromide for visualization. Construction of molecular clones. Total RNA was isolated from FEK cells, and equine PBMCs were stimulated with phorbol 12-myristate 13-acetate using TRI reagent (Sigma). First-strand cDNAs were synthesized using total RNA from both FEK cells and equine PBMCs, using an oligo(dT) primer and SuperScript II reverse transcriptase (Invitrogen). Primer pairs were designed to amplify the predicted A3 cDNAs identified in the NCBI's Horse Genome Resource. The primer pairs for the predicted equine gene XM_001499905.1 (sense primer, 5′-gcgcGGTACCaccatggagaagttggatcct-3′; antisense primer, 5′-gcgcCAATTGcttgagaatctcctcaagg-3′) and the XM_001499905.1 sense primer when paired with an antisense primer based on equine gene sequence for XM_001499895.1 (5′-gcgcCAATTGcttgagaaggtcctcaagctttctggccaggagat-3′) produced PCR products of ~1.1 kb. Capital letters indicate introduced Asp718 and MfeI restriction enzyme sites. The PCR products were digested with Asp718 and MfeI and cloned in frame into pcDNA3-HA (digested with Asp718/EcoRI) to generate the expression plasmids peA3F1-HA and peA3F2-HA. The eA3F1 and eA3F2 cDNA sequences were then determined. The eA3 expression cassettes, including the influenza hemagglutinin (HA) tag, were also transferred as HindIII/XhoI fragments into the pK vector, which does not contain a neo selection cassette, to generate pK/eA3F1-HA and pK/eA3F2-HA. The following mammalian expression plasmids have been previously described: pEV53B ( 53); pUNC-SIN6.1CLW-1 ( 54); pHIT/G ( 60); pNL4-3ΔVifΔEnv, phA3G-HA, and pNL-Luc-HXBΔVif ( 6); pK/hA3G-HA and pK/hA3A-HA ( 7); phA3A-HA ( 70); pSIV-AGM-LucΔVif ( 41); pNCS ( 25); pDJ33/440N1neo TNF ( 18); and pCMVMus-6Dneo TNF ( 22). pFB-Luc was obtained from Stratagene. A bacterial expression plasmid, pGEX4T-EIAV NC, expressing glutathione S-transferase (GST) fused to 62 amino acids of EIAV Gag, spanning the two zinc fingers of the NC domain (GGPLKAAQT CYN CGKP HLSSQ CRAPKV CFK CKQPG HFSKQ CRSVPKNGKQGAQGRPQKQTF; critical zinc binding residues are underlined), was constructed by amplification of the relevant region of the EIAV gag gene. This EIAV sequence was cloned in frame into the BamHI/XhoI sites of pGEX4T. The pGEX4T-HIV NC plasmid, which encodes HIV-1 NC fused to GST, has been previously described ( 5). Cell culture. HeLa and 293T cells were cultured in Dulbecco's modified Eagle's medium supplemented with 5% fetal bovine serum and gentamicin (Gibco). Viral production and luciferase assay. A total of 3 × 105 293T cells were transfected with 62 ng, 125 ng, or 250 ng of an A3 expression plasmid, with pcDNA3 filler plasmid added to a total of 1 μg. In addition, cells were also cotransfected with pNL-Luc-HXBΔVif (2 μg), pSIV-AGM-LucΔVif (2 μg), pNCS(1 μg), and pFB-LUC (1 μg) or pEV53B (1 μg) and pUNC-SIN6.1CLW-1 (1 μg). All viruses were pseudotyped by the addition of 200 ng of the vesicular stomatitis virus G (VSV-G) expression plasmid pHIT/G to the transfection cocktail. Forty-four hours posttransfection, virus-containing supernatants were collected, passed through 0.45-μm-pore-sized filters and used to infect naïve 293T cells. At this time, viral producer cells were lysed in sodium dodecyl sulfate (SDS)-acrylamide gel loading buffer containing β-mercaptoethanol, and this lysate was then analyzed by Western blot analysis for A3 and HIV-1 or EIAV Gag protein levels. Twenty-four hours postinfection, the infected 293T cells were lysed in passive lysis buffer (Promega) and luciferase activity was determined as previously described, using Promega's luciferase assay system. Protein purification and binding assays. Escherichia coli strain BL21 was transformed with pGEX4T, pGEX4T-HIV NC, or pGEX4T-EIAV NC. Protein expression was induced by the addition of 1 mM isopropyl-β- d-thiogalactopyranoside (IPTG) (Invitrogen). Protein purification and A3:Gag binding assays were performed as previously described ( 5). All A3 proteins were produced by transfecting 10 μg of expression plasmid into 1.5 × 10 6 293T cells using the calcium phosphate method. At 44 h posttransfection, cells were lysed in 4 ml of 150 mM NaCl, 50 mM Tris (pH 7.5), and 0.5% NP-40 (binding buffer) and centrifuged to remove insoluble matter. One milliliter of the A3-containing clarified supernatant was incubated with ~250 ng of partially purified recombinant GST, GST-HIV NC, or GST-EIAV NC, and 50 μl of washed glutathione-agarose beads at 4°C for 60 min. The agarose beads/protein complexes were then washed four times with binding buffer, and bound proteins were eluted with 100 μl of SDS-acrylamide gel loading buffer containing β-mercaptoethanol. Input and bound fractions were then analyzed by Western blot analysis, as previously described ( 5, 6). Packaging of hA3G, eA3F2, and eA3F1 into HIV-1 and EIAV virions. A total of 1.5 × 106 293T cells in a 10-cm dish were transfected by the calcium phosphate method with 1.5 μg of an A3 expression plasmid, 4.5 μg pcDNA3 filler, and either 12 μg of the pNL4-3ΔVifΔEnv HIV-1 proviral clone or 6 μg of pEV53B and 6 μg of pUNC-SIN6.1CLW-1. At 48 h posttransfection, virus-containing supernatants were collected, filtered, and layered over binding buffer supplemented with 20% sucrose. Virus-producing cells were lysed in SDS-acrylamide gel loading buffer containing β-mercaptoethanol and used for Western blot analysis of the levels of A3 and viral Gag expression. Sucrose gradients were spun at 40,000 rpm for 90 min. Supernatants were discarded, and the pelleted virus was resuspended in SDS-acrylamide gel loading buffer containing β-mercaptoethanol. The lysate and purified virions were then analyzed by Western blot analysis. Editing of HIV-1 proviral DNA. HIV-1 virions were produced in transfected 293T cells as described above and used to infect naïve 293T cells. Twenty-four hours postinfection, duplicate samples were either analyzed for luciferase activity to confirm inhibition of infectivity (data not shown) or lysed and total DNA was isolated using a DNeasy kit (Qiagen). Purified DNA was then digested with DpnI to cleave any contaminating plasmid DNA. The 3′ end of the virally encoded luciferase reporter gene was amplified by PCR, cloned, and sequenced. Western blot analyses. All Western blots were developed using the Lumi-Light Western blotting substrate (Roche), as previously described. Proteins were detected using the following reagents: HA-tagged proteins detected with a mouse monoclonal anti-HA antibody (Covance), followed by a secondary goat anti-mouse horseradish peroxidase (HRP)-conjugated antibody (Amersham). HIV-1 Gag was detected using a rabbit polyclonal anti-p24 antiserum (Division of AIDS, NIAID, NIH; produced by BioMolecular Technologies), followed by a secondary goat anti-rabbit HRP-conjugated antibody (Amersham). EIAV Gag was detected using serum from an EIAV-infected horse, followed by a secondary goat anti-horse HRP-conjugated antiserum (GeneTex, Inc.) Bacterial mutator assay. A3 cDNAs, including the C-terminal HA tag, were excised from the relevant pK-based plasmid by cleavage with Asp718 and XhoI and subcloned into Asp718 and SalI sites present in the bacterial expression plasmid pTrc99A (AP Biosciences). The uracil DNA glycosylase-deficient E. coli strain BW310 ( 28) was transformed with the pTrc99A parental plasmid and vectors encoding the various A3 cDNAs. Transformed bacteria were then selected overnight on LB plates containing ampicillin. Twenty colonies were pooled into 2 ml of LB medium plus ampicillin plus 1 mM IPTG and cultures grown overnight at 37°C. One hundred microliters of the saturated culture was then plated on LB plates containing 100 μg/ml of rifampin, and the total number of rifampin-resistant (Rif r) colonies per plate was counted 24 h later. To verify protein expression, 100 μl of the saturated IPTG-induced culture was lysed and analyzed by Western blot analysis as described above. Retrotransposition assays. The retrotransposition assays used have been previously described ( 7, 22). Briefly, 3 × 10 5 HeLa cells were seeded into 35-mm culture dishes and then transfected with 2 μg of reporter plasmid (the intracisternal A particle [IAP] retrotransposition indicator plasmid pDJ33/440N1neo TNF, the MusD retrotransposition indicator plasmid pCMVMus-6Dneo TNF, or the control neo expression plasmid pcDNA3) and 2 μg of the control pK parental plasmid, pK/hA3A-HA, pK/hA3G-HA, pK/eA3F2-HA, or pK/eA3F1-HA. At 72 h posttransfection, the cells were transferred to a 10-cm dish and subjected to selection with 700 μg/ml G418 (Geneticin) for an additional 12 days. Neomycin-resistant (Neo r) colonies were then stained with crystal violet and counted. Subcellular localization of the eA3F2 and eA3F1 proteins. HeLa cells were transfected with 2 μg of either peA3F1-HA or peA3F2-HA. At 44 h posttransfection, the cells were permeabilized ( 8), and eA3F1 and eA3F2 were visualized by staining with an anti-HA mouse monoclonal antibody, followed by goat anti-mouse antiserum conjugated to fluorescein isothiocyanate. Nuclei were identified by staining with Hoechst stain, as previously described ( 8). Identifying the equine A3 genome locus and expressed sequences. The number of A3 genes in mammalian species is quite variable, ranging from a single gene in mice to seven genes in humans ( 15, 32, 52). Up to four A3 genes have been reported in other nonprimate species, including dogs, cats, cows, pigs, and sheep ( 15, 33, 48, 52). The recent availability of the horse genome sequence allowed a detailed physical characterization of the equine A3 gene locus. Searches of equine EST and genome databases identified a number of genes homologous to the hA3G gene. RT-PCR and sequencing were used to delineate gene boundaries and verify gene identity and expression in equine lymphoid cells. Collectively, these analyses identified six equine A3 genes that map in a cluster on equine chromosome 28. The six genes are arranged in a head-to-tail orientation and span 83,126 bp (Fig. ). With the exception of a single exon in the opposite orientation (Fig. ), no other A3-like genes were found in this locus, and no A3 sequences were identified outside of this region. The A3 gene locus on equine chromosome 28 is flanked by NPTXR and CBX7, which also flank the human A3 gene locus on human chromosome 22. The CBX6 gene, which is located between human NPTXR and hA3A, appears to be present in the same location in horses; however, equine CBX6 is not yet annotated. Overall, the conservation of gene order between horse chromosome 28 and human chromosome 22 extends from C22orf28 through ALG12, spanning approximately 15 million bases of the horse chromosome. Therefore, the equine A3 gene locus contains six expressed genes and is syntenic to the A3 segment of human chromosome 22. Four of the six equine genes encode a single-CDA domain, and two genes encode double-CDA domains. Phylogenetic analyses indicated the eight equine CDA domains grouped with A3 sequences and segregated by Z1a, Z1b, or Z2 CDA domain designation ( 15) (Fig. ). Within the “Z” groups, the CDA domains group by species rather than by gene designation. Using the phylogenetic relationships and the number of CDA domains present per gene, we could group the six genes into two hA3A-like genes, two hA3F-like genes, one hA3C-like gene, and one hA3H-like gene. Although we recognize that it is difficult to assign nomenclature without functional characterization, we have designated these genes, listed in the order they occur in the locus, EcA3A1, EcA3A2, EcA3F1, EcA3F2, EcA3C, and EcA3H (Fig. ). The two EcA3A genes were previously identified as A3 by Conticello et al. ( 15). Each gene encodes one CDA domain and clusters with human Z1b CDA sequences, characteristics that are shared by hA3A. The first gene in the locus, EcA3A1, contains an atypical CDA sequence (DXEX 27PCX 2C rather than HXEX 27PCX 2C). The aspartate mutation is present in 29 equine EST clones as well as the annotated genome. It is not known how this mutation affects activity, but EcA3A1 was expressed in PBMCs from several horses (data not shown). The EcA3F1 and EcA3F2 genes each contain two Z1a CDA domains and cluster with A3F sequences from multiple species. Moreover, the predicted eA3F1 and eA3F2 proteins show extensive sequence homology to hA3F ( HsA3F) at the amino acid level (Fig. ). The transcript for EcA3F1 was amplified from PBMCs stimulated with phorbol 12-myristate 13-acetate, while several single- and double-CDA-domain transcripts for EcA3F2 were amplified from both FEK and equine PBMCs (data not shown). Single-domain EcA3F2 transcripts may represent alternately spliced transcripts and/or PCR artifacts. The EcA3C gene contains a single Z1a CDA domain, while EcA3H contains a single Z2 CDA domain and is thus most similar to hA3C and hA3H, respectively (Fig. ). Interestingly, EcA3C and EcA3H are computationally predicted to form a single, seven-exon, double Z1a/Z2 CDA gene ( XM_001501833). This predicted genomic organization and complement of CDA domains is most similar to the A3F gene described in artiodactyl species ( 33). However, despite repeated attempts, we failed to detect any double-CDA transcripts by RT-PCR (data not shown). Rather, we consistently detected the independent expression of either the 5′ or the 3′ half of this predicted double-CDA gene, which was also consistent with the EST clones (Table 1). Therefore, we conclude that EcA3C and EcA3H are in fact two separate genes, each with a single-CDA domain. Finally, comparison of the nucleotide sequence identity and amino acid similarity within and among the equine A3 genes confirmed the presence of six distinct genes (Table 2). The EcA3A1 and EcA3A2 genes were closely related, as were EcA3F1 and EcA3F2; however, the percent identity among the alleles of each gene was higher and did not overlap the percent identity between the different genes. In summary, we identified six expressed genes in the eA3 locus, which is more than currently described for any other nonprimate species. Strikingly, the repertoire and organization of the eight equine CDA domains (five Z1a in single- and double-domain genes, two Z1b in single-domain genes, and one Z2 single-domain gene) are most similar to the human CDA organization rather than that of artiodactyl species, cats, or mice ( 15, 33, 48). | TABLE 2.Nucleotide identity and amino acid similarity among the equine APOBEC3 genes and allelesa |
Tissue distribution of e3AF1 and e3AF2. The number and repertoire of the equine A3 genes suggest they may have evolved to play an important role in antiviral defense. Previous work has revealed that A3 proteins that show strong antiretroviral activity, such as hA3G, generally contain two CDA domains ( 16), and it was therefore of particular interest to analyze the two equine A3 double-CDA genes for breadth of expression and antiviral properties, especially against EIAV. To determine whether EcA3F1 and EcA3F2 transcripts (henceforth termed eA3F1 and eA3F2) were expressed in cells relevant to EIAV replication, a cDNA panel was assembled from tissues and cells that differ in permissiveness for EIAV replication in vivo and in vitro. The panel included seven somatic and lymphoid tissues (brain, kidney, liver, lung, bone marrow, lymph node, and spleen), circulating lymphoid cells (PBMCs and monocytes), and in vitro-cultured MDMs, ED cells, and FEK cells. Overall, eA3F1 and eA3F2 showed similar, variable patterns of mRNA expression in equine cells and tissues (Fig. ). Abundant levels of each transcript were detected in the liver, lung, and lymphoid tissues, including PBMCs and MDMs. Low but detectable levels of mRNA were also present in brain, kidney, and FEK cells and monocytes; however, no amplified products were detected in ED cells. The abundant mRNA expression in spleen cells and MDMs are of particular interest because EIAV is a macrophage-tropic lentivirus, and the main site of infection and replication in the acute and chronic stages of disease are the spleen and tissue macrophages of the lung, liver, kidney, lymph node, and bone marrow tissues ( 44, 51, 63). In contrast, very low mRNA levels were detected in circulating monocytes, which are not permissive for EIAV replication ( 43, 63). Field isolates of EIAV do not readily replicate in vitro, but certain isolates have been adapted to fibroblast cell lines, such as ED and FEK. Interestingly, low to no levels of eA3F1 and eA3F2 mRNA were present in FEK and ED cells. Collectively, these results demonstrate that transcripts encoding the equine double-CDA-domain proteins eA3F1 and eA3F2 are expressed in cells and tissues that are permissive for EIAV replication in vivo and, furthermore, that expression of these genes is not correlated with restriction of EIAV replication in vivo or in vitro. Functional analyses of e3AF1 and e3AF2. An important attribute of A3 proteins is their ability to edit dC residues to dU on single-stranded DNA templates ( 71). To test whether eA3F1 and eA3F2 showed this enzymatic activity, we used a previously described ( 8, 28) DNA mutation assay in bacteria. This assay measures the ability of a protein to mutate the E. coli RNA polymerase B gene ( rpoB). Mutations in rpoB are then detected by screening for the frequency of Rif r colonies. Expression of hA3A and hA3G has previously been shown to greatly or modestly enhance, respectively, the frequency of rpoB mutations ( 8, 28). As shown in Fig. , hA3A increased the incidence of Rifr colonies by ~70-fold when expressed in bacteria, while hA3G increased their incidence by a more modest ~5-fold. Similarly, expression of eA3F2 increased the number of Rifr colonies by ~14-fold, while eA3F1 had an ~3-fold enhancing effect. DNA sequence analysis of the rpoB gene in the Rifr colonies derived from the eA3F1- or eA3F2-expressing bacteria showed almost exclusively C-to-T mutations, as predicted (data not shown). We therefore conclude that both eA3F1 and eA3F2 are functional CDAs. We next asked whether eA3F1 or eA3F2 would be able to inhibit the infectivity of HIV-1, simian immunodeficiency virus (SIV), EIAV, or MLV by producing infectious retroviral vector particles based on each of these viruses in the presence of eA3F1 or eA3F2. hA3G, a potent inhibitor of many different retroviruses, and hA3A, a relatively weak inhibitor, were used as controls. For this purpose, we cotransfected 293T cells with a VSV-G expression plasmid (pHIT/G) and a plasmid encoding one of these four A3 proteins, together with a plasmid which encodes an HIV-1-based lentiviral reporter virus encoding luciferase but lacking an intact vif gene (pNL-Luc-HXBΔVif), or a similar plasmid encoding a ΔVif version of an African green monkey SIV provirus (pSIV-AGM-LucΔVif). Alternatively, the A3 expression plasmids were cotransfected with a plasmid encoding an MLV-based retroviral vector expressing luciferase (pFB-luc) and an MLV packaging plasmid (pNCS) or a plasmid encoding an EIAV-based lentiviral vector expressing luciferase (pUNC-SIN6.1CLW-1) and an EIAV packaging plasmid (pEV53B). Of note, pEV53B expresses every known EIAV protein except Env and Tat ( 53). As shown in Fig. , hA3G acted as a potent inhibitor of the infectivity of HIV-1ΔVif, SIVagmΔVif, EIAV, and MLV, while hA3A had little effect on viral infectivity, with the exception of SIVagm, which was inhibited by approximately threefold. eA3F1 inhibited HIV-1ΔVif infectivity by ~25-fold, SIVagmΔVif infectivity by ~10-fold, and MLV infectivity by ~3-fold but inhibited EIAV infectivity by <2-fold. Finally, eA3F2 inhibited SIVagmΔVif infectivity by ~10-fold but reduced HIV-1ΔVif, MLV, and EIAV infectivity by ≤2-fold (Fig. ). The data presented in Fig. suggest that eA3F1 and eA3F2 are not effective inhibitors of EIAV infectivity but that they are capable of selectively inhibiting other retroviruses. To further confirm this result, we performed a limited dose-response analysis looking at the abilities of eA3F1, eA3F2, hA3G, and hA3A to inhibit EIAV or HIV-1ΔVif infectivity (Fig. ). As may be observed, these data confirmed the data reported in Fig. showing that eA3F1 is a potent inhibitor of HIV-1ΔVif infectivity, while hA3G is a potent inhibitor of EIAV infectivity. In contrast, neither eA3F1 nor eA3F2 exerted a strong inhibitory effect on EIAV. eA3F1 can edit HIV-1 reverse transcripts. Inhibition of retroviral infectivity by A3 proteins is usually, but not invariably, correlated with a significant level of editing of retroviral reverse transcripts ( 30, 40, 50, 74). Specifically, hA3G and other inhibitory A3 proteins can edit dC residues on the proviral minus strand to dU residues, resulting in misincorporation of A in place of G during copying of the proviral DNA plus strand by the reverse transcriptase ( 71). As eA3F2 did not exert a strong inhibitory effect on any retrovirus tested (Fig. ), we instead asked whether eA3F1 was capable of editing HIV-1 reverse transcripts made by HIV-1ΔVif virions generated in its presence. As shown in Fig. , eA3F1 in fact induced a level of G-to-A editing that was at least equal to that seen upon coexpression of hA3G with HIV-1ΔVif. We also noted a significant increase in C-to-T mutations, which suggests that hA3G and eA3F1 are also able to edit the HIV-1 proviral plus strand, albeit clearly less effectively than the minus strand. In contrast, preliminary experiments have not detected any editing of EIAV reverse transcripts produced from EIAV virions derived from cells expressing eA3F1, although editing of EIAV reverse transcripts by hA3G was readily detected (data not shown). Previous work from several groups has shown that hA3G prefers to edit C residues located 3′ to another C, while hA3F prefers to edit C residues located 3′ to T residues ( 4, 38, 70). Consistent with the sequence similarity between hA3F and eA3F1 (Fig. ), we observed that eA3F1 also preferred to edit TC, rather than CC, targets (Fig. ). Virion packaging of eA3F1 and eA3F2. In order to inhibit retroviral infectivity, A3 proteins have to be specifically packaged into retroviral virion particles and then exert their inhibitory effect during the reverse transcription process in newly infected target cells ( 16). It has been reported that some retroviruses that lack vif are able to selectively exclude specific A3 proteins from virion particles and thereby achieve resistance to their inhibitory effects ( 17, 19). To address whether the poor inhibition of EIAV infectivity exerted by eA3F1 and eA3F2 (Fig. and ) results from a similar exclusion phenomenon, we produced HIV-1ΔVif and EIAV virions in the presence of hA3A, hA3G, eA3F1, or eA3F2 and then examined whether these proteins were packaged effectively. We have previously reported that hA3A is not effectively packaged into HIV-1ΔVif virions ( 27), so this A3 protein is used here as a control for packaging specificity. As shown in Fig. , and as expected, hA3A was not packaged into HIV-1ΔVif virions, while packaging of hA3G was efficient. Interestingly, both eA3F2, which is a weak inhibitor of HIV-1ΔVif infectivity, and eA3F1, a strong inhibitor (Fig. and ), appeared to be packaged into HIV-1 virions as effectively as hA3G (Fig. ). Analysis of A3 protein packaging into EIAV virions gave a closely similar result. Again hA3A failed to package, while the incorporation of hA3G, eA3F1, and eA3F2 was efficient and closely comparable. We therefore conclude that the inability of eA3F1 or eA3F2 to inhibit EIAV infectivity effectively, compared to hA3G, does not reflect a difference in virion incorporation efficiency. To begin to address whether eA3F1 and eA3F2 were being packaged into retroviral virion cores, the experiments shown in Fig. were repeated in the presence of the detergent Triton X-100, which has been reported to strip away proteins that copurify with HIV-1 virions but that are not located within the virion core ( 26). However, treatment of virions with this detergent did not reduce the association of hA3G, eA3F1, or eA3F2 with either HIV-1 or EIAV virion particles (data not shown), thus suggesting that these proteins are indeed core associated. The resistance of EIAV to inhibition by eA3F1 and eA3F2 observed in Fig. and must therefore occur via a mechanism that acts subsequent to their incorporation into virions. One possible mechanism of resistance after packaging, first suggested by Abudu et al. ( 1) in the case of MLV resistance to inhibition by mA3, is that the packaged eA3F proteins might be cleaved by the viral protease after viral budding. At least in the case of eA3F1, we did in fact occasionally see an eA3F1 cleavage product in EIAV virions that was not present in the producer cells (Fig. , lane 6). However, as cleavage was both far from complete and observed at similar levels in HIV-1 virions (Fig. , lane 6), which are highly susceptible to inhibition by eA3F1 (Fig. and ), protease cleavage seems unlikely to explain the insensitivity of EIAV to eA3F1 inhibition. Previously, we and others have reported that the selective incorporation of A3 proteins into retroviral virions is due to a specific interaction between the packaged A3 protein and the NC domain of the Gag polyprotein ( 2, 12, 60, 73). As eA3F1 and eA3F2 are apparently able to specifically package into both HIV-1ΔVif and EIAV virion cores (Fig. ), we therefore predicted that packaging should correlate with the ability to interact with the EIAV and/or HIV-1 NC protein in vitro. To perform this analysis, we incubated recombinant bacterial GST-HIV NC and GST-EIAV NC fusion proteins, or unfused GST, with recombinant mammalian HA-tagged hA3A, hA3G, eA3F1, or eA3F2. As may be observed in Fig. , hA3A, which fails to incorporate into either HIV-1 or EIAV virion particles (Fig. ), also fails to specifically bind either the HIV-1 or EIAV NC protein. In contrast, hA3G, eA3F1, and eA3F2, which are selectively incorporated into both HIV-1 and EIAV virions (Fig. ), also specifically interact with both GST-HIV NC and GST-EIAV NC but not with unfused GST (Fig. ). These data therefore suggest that incorporation of eA3F1 and eA3F2 into EIAV virions, like incorporation of hA3G into HIV-1 virions, is mediated by a specific interaction with the NC domain of each Gag polyprotein. Inhibition of retrotransposon mobility by eA3F1 and eA3F2. While the A3 proteins first became of interest due to their ability to inhibit the infectivity of exogenous retroviruses, it is now clear that many A3 proteins are also capable of inhibiting retrotransposon mobility ( 7, 8, 13, 21, 22, 67). To test whether eA3F1 or eA3F2 also shared this activity, we examined their ability to inhibit the mobility of two murine long-terminal-repeat retrotransposons, MusD and IAP, using previously described retrotransposition indicator constructs ( 18, 22). These constructs contain a neo gene, inserted into the MusD or IAP retrotransposon in the antisense orientation, disrupted by an intron in the sense orientation. Therefore, Neo r can only occur if the indicator construct is transcribed and spliced and then reverse transcribed and integrated. To perform this analysis, we cotransfected HeLa cells with the MusD or IAP retrotransposition indicator constructs together with vectors encoding hA3A, hA3G, eA3F1, or eA3F2. As a control for nonspecific toxicity, we also cotransfected each of these A3 expression plasmids with pcDNA3, which contains an intact neo gene that can confer Neo r without requiring a reverse transcription step. As shown in Fig. , none of these A3 proteins exerted any nonspecific inhibitory activity on Neo r conferred by pcDNA3. In contrast, and as previously described, hA3A acts as a potent inhibitor of IAP mobility, reducing retrotransposition of IAP by ~20-fold, while hA3G exerted a modest ~2- to 3-fold inhibitory effect on IAP mobility ( 7, 22). Similarly, eA3F2 was observed to inhibit IAP mobility by ~10-fold, while eA3F1 exerted an ~5-fold inhibitory effect (Fig. ). In the case of the MusD retrotransposon, hA3G, again as previously reported, inhibited retrotransposition by ~10-fold, while hA3A had only a modest inhibitory effect ( 22). In the case of MusD, eA3F2 proved to be an ineffective inhibitor, while eA3F1 inhibited MusD mobility by ~10-fold (Fig. ). We therefore conclude that while eA3F1 and eA3F2 can both function as potent inhibitors of retrotransposon mobility, these two eA3 proteins clearly have different abilities to affect the mobility of distinct retrotransposon species. The molecular bases for these differences are currently unclear. eA3F1 and eA3F2 show distinct subcellular localizations. While the majority of the two-CDA-domain A3 proteins, all of which are too large to passively diffuse into the nucleus, are found localized in the cytoplasm, at least one hA3 protein, hA3B, is localized to nuclei and in fact functions as a nucleocytoplasmic shuttle protein ( 8, 67). Analysis of the subcellular localization of the two equine double-CDA-domain proteins revealed that eA3F1, like hA3G and hA3F, localizes to the cytoplasm, while eA3F2, like hA3B, is found predominantly in the nucleus (Fig. ). While the functional relevance of this difference in subcellular localization is currently unclear, it is interesting to note that both eA3F1 and eA3F2 are able to package into HIV-1 and EIAV virions with comparable efficiency (Fig. ), even though these are thought to be assembled exclusively in the cytoplasm. Species-specific expansion and divergence of mammalian A3 genes have occurred in response to evolutionary pressure from exogenous and endogenous retroviruses and/or retroelements. The presence of six A3 genes in the equine genome (Fig. ), including two Z1a/Z1a CDA genes (Fig. ), is greater than that reported in any other nonprimate species and is nearly equal to that found in the human genome. The variability in the number of A3 genes among mammalian species, ranging from one to seven, results from gene expansion by duplication within a syntenic region ( 15, 59). An evolutionary history of gene duplication is also likely for the equine A3 genes, since eA3A1 and eA3A2 are remarkably similar to each other, as are eA3F1 and eA3F2 (Fig. and ; Table 2). It is not known whether the six equine A3 genes have redundant or separate functions. However, the observed differences between eA3F1 and eA3F2 with respect to antiviral activity, subcellular localization, and retrotransposon specificity (Fig. , , , and ) indicate that at least some of the equine genes are functionally distinct. Phylogenetic segregation of CDA domains into Z1a, Z1b, or Z2 types has facilitated comparative analyses of the repertoire of A3 genes within and among different species ( 15, 33, 52). Based on the complement of Z domains, it is evident that the usage and organization of the equine CDA domains are most similar to that seen in humans (Fig. ). Like hA3F, the equine double-CDA-domain proteins eA3F1 and eA3F2 comprise two Z1a domains. In contrast, a Z1a/Z2 configuration is found in all of the double-CDA-domain proteins of nonprimate species characterized to date, including artiodactyls (A3F), cats (A3CH), and mice (A3) ( 15, 33, 48). Horses are not evolutionarily closer to humans than these other species; therefore, the similar complement of double-CDA domains may have arisen from similar selective pressures during the evolution of A3 genes. The studies reported here demonstrate that eA3F1 has potent antiviral activity against HIV-1 and MLV but only modest activity against EIAV (Fig. and ). This suggests that EIAV has evolved a mechanism to resist or inhibit the intrinsic antiviral activity of eA3F1. Retroviruses that lack vif use various strategies to resist the antiviral activity of A3 proteins, and most, if not all, inhibit incorporation of cognate A3 proteins into virions ( 16). A possible exception to this generalization has recently been reported for MLV, which appears to package the cognate mA3 protein into MLV virion particles yet is apparently less susceptible to inhibition by virion-incorporated mA3 than by equivalent levels of virion-incorporated hA3G ( 9, 57). However, other groups have suggested that MLV virions also package hA3G somewhat more efficiently than mA3 ( 1, 20, 34). In this paper, we demonstrate that EIAV does not block the expression of eA3F1 or eA3F2 in transfected cells (Fig. and ) or prevent EIAV NC binding or packaging of eA3F1 or eA3F2 into virions (Fig. and ). These data suggest that EIAV may employ a novel mechanism to escape the antiviral activities of eA3F1 and eA3F2 that act at a step after virion incorporation ( 57). This resistance does not result from any lack of biological activity on the part of the eA3 proteins, as eA3F1 at least is clearly a potent inhibitor of HIV-1 and SIV infectivity and effectively edits HIV-1 reverse transcripts after virion incorporation (Fig. , , and ). Moreover, both eA3F1 and eA3F2 are active CDAs (Fig. ). The ability of EIAV to avoid inhibition by eA3F1 and eA3F2 at an undefined step after virion incorporation presumably explains the ability of EIAV to replicate in vivo in tissues that, as measured by mRNA expression levels, appear to produce substantial levels of these equine intrinsic immunity factors (Fig. ). R.L.T. was supported by NIH grant T32 AI007025. Work at WSU was partially supported by grants from the Schindler Equine Research Funds and Washington State Equine Research Funds. Work in the laboratory of B.R.C. was supported by NIH grant R01-AI065301. We thank Steven Goff (Columbia University), Nathaniel Landau (NYU), Thierry Heidmann (Institute Gustave Roussy), Fred Fuller (NCSU), Reuben Harris (University of Minnesota), and John C. Olsen (UNC) for reagents used in this research. The rabbit polyclonal HIV-1SF2 p24 antiserum reagent was obtained through the NIH AIDS Research and Reference Reagent Program, Division of AIDS, NIAID, NIH. We thank Sue Pritchard and Yousuf Jafarey for excellent technical assistance. 1. Abudu, A., A. Takaori-Kondo, T. Izumi, K. Shirakawa, M. Kobayashi, A. Sasada, K. Fukunaga, and T. Uchiyama. 2006. Murine retrovirus escapes from murine APOBEC3 via two distinct novel mechanisms. Curr. Biol. 161565-1570. [PubMed] 2. Alce, T. M., and W. Popik. 2004. APOBEC3G is incorporated into virus-like particles by a direct interaction with HIV-1 Gag nucleocapsid protein. J. Biol. Chem. 2793408 51 -34086. [PubMed] 3. Belshan, M., G. S. Park, P. Bilodeau, C. M. Stoltzfus, and S. Carpenter. 2000. Binding of equine infectious anemia virus rev to an exon splicing enhancer mediates alternative splicing and nuclear export of viral mRNAs. Mol. Cell. Biol. 203550-3557. [PubMed] 4. Bishop, K. N., R. K. Holmes, A. M. Sheehy, N. O. Davidson, S. J. Cho, and M. H. Malim. 2004. Cytidine deamination of retroviral DNA by diverse APOBEC proteins. Curr. Biol. 141392-1396. [PubMed] 5. Bogerd, H. P., and B. R. Cullen. 2008. Single-stranded RNA facilitates nucleocapsid: APOBEC3G complex formation. RNA 141228-1236. [PubMed] 6. Bogerd, H. P., B. P. Doehle, H. L. Wiegand, and B. R. Cullen. 2004. A single amino acid difference in the host APOBEC3G protein controls the primate species specificity of HIV type 1 virion infectivity factor. Proc. Natl. Acad. Sci. USA 1013770-3774. [PubMed] 7. Bogerd, H. P., H. L. Wiegand, B. P. Doehle, K. K. Lueders, and B. R. Cullen. 2006. APOBEC3A and APOBEC3B are potent inhibitors of LTR-retrotransposon function in human cells. Nucleic Acids Res. 3489-95. [PubMed] 8. Bogerd, H. P., H. L. Wiegand, A. E. Hulme, J. L. Garcia-Perez, K. S. O'Shea, J. V. Moran, and B. R. Cullen. 2006. Cellular inhibitors of long interspersed element 1 and Alu retrotransposition. Proc. Natl. Acad. Sci. USA 1038780-8785. [PubMed] 9. Browne, E. P., and D. R. Littman. 2008. Species-specific restriction of Apobec3-mediated hypermutation. J. Virol. 821305-1313. [PubMed] 10. Carroll, R., L. Martarano, and D. Derse. 1991. Identification of lentivirus Tat functional domains through generation of equine infectious anemia virus/human immunodeficiency virus type 1 tat gene chimeras. J. Virol. 653460-3467. [PubMed] 11. Carvalho, M., and D. Derse. 1991. Mutational analysis of the equine infectious anemia virus Tat-responsive element. J. Virol. 653468-3474. [PubMed] 12. Cen, S., F. Guo, M. Niu, J. Saadatmand, J. Deflassieux, and L. Kleiman. 2004. The interaction between HIV-1 Gag and APOBEC3G. J. Biol. Chem. 27933177-33184. [PubMed] 13. Chen, H., C. E. Lilley, Q. Yu, D. V. Lee, J. Chou, I. Narvaiza, N. R. Landau, and M. D. Weitzman. 2006. APOBEC3A is a potent inhibitor of adeno-associated virus and retrotransposons. Curr. Biol. 16480-485. [PubMed] 14. Conticello, S. G., R. S. Harris, and M. S. Neuberger. 2003. The Vif protein of HIV triggers degradation of the human antiretroviral DNA deaminase APOBEC3G. Curr. Biol. 132009-2013. [PubMed] 15. Conticello, S. G., C. J. Thomas, S. K. Petersen-Mahrt, and M. S. Neuberger. 2005. Evolution of the AID/APOBEC family of polynucleotide (deoxy)cytidine deaminases. Mol. Biol. Evol. 22367-377. [PubMed] 16. Cullen, B. R. 2006. Role and mechanism of action of the APOBEC3 family of antiretroviral resistance factors. J. Virol. 801067-1076. [PubMed] 17. Derse, D., S. A. Hill, G. Princler, P. Lloyd, and G. Heidecker. 2007. Resistance of human T cell leukemia virus type 1 to APOBEC3G restriction is mediated by elements in nucleocapsid. Proc. Natl. Acad. Sci. USA 1042915-2920. [PubMed] 18. Dewannieux, M., A. Dupressoir, F. Harper, G. Pierron, and T. Heidmann. 2004. Identification of autonomous IAP LTR retrotransposons mobile in mammalian cells. Nat. Genet. 36534-539. [PubMed] 19. Doehle, B. P., H. P. Bogerd, H. L. Wiegand, N. Jouvenet, P. D. Bieniasz, E. Hunter, and B. R. Cullen. 2006. The betaretrovirus Mason-Pfizer monkey virus selectively excludes simian APOBEC3G from virion particles. J. Virol. 8012102-12108. [PubMed] 20. Doehle, B. P., A. Schafer, H. L. Wiegand, H. P. Bogerd, and B. R. Cullen. 2005. Differential sensitivity of murine leukemia virus to APOBEC3-mediated inhibition is governed by virion exclusion. J. Virol. 798201-8207. [PubMed] 21. Dutko, J. A., A. Schafer, A. E. Kenny, B. R. Cullen, and M. J. Curcio. 2005. Inhibition of a yeast LTR retrotransposon by human APOBEC3 cytidine deaminases. Curr. Biol. 15661-666. [PubMed] 22. Esnault, C., O. Heidmann, F. Delebecque, M. Dewannieux, D. Ribet, A. J. Hance, T. Heidmann, and O. Schwartz. 2005. APOBEC3G cytidine deaminase inhibits retrotransposition of endogenous retroviruses. Nature 433430-433. [PubMed] 23. Flaminio, M. J., A. Yen, and D. F. Antczak. 2004. The proliferation inhibitory proteins p27Kip1 and retinoblastoma are involved in the control of equine lymphocyte proliferation. Vet. Immunol. Immunopathol. 102363-377. [PubMed] 24. Fridell, R. A., K. M. Partin, S. Carpenter, and B. R. Cullen. 1993. Identification of the activation domain of equine infectious anemia virus Rev. J. Virol. 677317-7323. [PubMed] 25. Gao, G., and S. P. Goff. 1998. Replication defect of moloney murine leukemia virus with a mutant reverse transcriptase that can incorporate ribonucleotides and deoxyribonucleotides. J. Virol. 725905-5911. [PubMed] 26. Goila-Gaur, R., M. A. Khan, E. Miyagi, S. Kao, and K. Strebel. 2007. Targeting APOBEC3A to the viral nucleoprotein complex confers antiviral activity. Retrovirology 461. [PubMed] 27. Gooch, B. D., and B. R. Cullen. 2008. Functional domain organization of human APOBEC3G. Virology 379118-124. [PubMed] 28. Haché, G., M. T. Liddament, and R. S. Harris. 2005. The retroviral hypermutation specificity of APOBEC3F and APOBEC3G is governed by the C-terminal DNA cytosine deaminase domain. J. Biol. Chem. 28010920-10924. [PubMed] 29. Hammond, S. A., S. J. Cook, D. L. Lichtenstein, C. J. Issel, and R. C. Montelaro. 1997. Maturation of the cellular and humoral immune responses to persistent infection in horses by equine infectious anemia virus is a complex and lengthy process. J. Virol. 713840-3852. [PubMed] 30. Harris, R. S., K. N. Bishop, A. M. Sheehy, H. M. Craig, S. K. Petersen-Mahrt, I. N. Watt, M. S. Neuberger, and M. H. Malim. 2003. DNA deamination mediates innate immunity to retroviral infection. Cell 113803-809. [PubMed] 31. Harrold, S. M., S. J. Cook, R. F. Cook, K. E. Rushlow, C. J. Issel, and R. C. Montelaro. 2000. Tissue sites of persistent infection and active replication of equine infectious anemia virus during acute disease and asymptomatic infection in experimentally infected equids. J. Virol. 743112-3121. [PubMed] 32. Jarmuz, A., A. Chester, J. Bayliss, J. Gisbourne, I. Dunham, J. Scott, and N. Navaratnam. 2002. An anthropoid-specific locus of orphan C to U RNA-editing enzymes on chromosome 22. Genomics 79285-296. [PubMed] 33. Jónsson, S. R., G. Hache, M. D. Stenglein, S. C. Fahrenkrug, V. Andresdottir, and R. S. Harris. 2006. Evolutionarily conserved and non-conserved retrovirus restriction activities of artiodactyl APOBEC3F proteins. Nucleic Acids Res. 345683-5694. [PubMed] 34. Kobayashi, M., A. Takaori-Kondo, K. Shindo, A. Abudu, K. Fukunaga, and T. Uchiyama. 2004. APOBEC3G targets specific virus species. J. Virol. 788238-8244. [PubMed] 35. Lee, J. H., G. Culver, S. Carpenter, and D. Dobbs. 2008. Analysis of the EIAV Rev-responsive element (RRE) reveals a conserved RNA motif required for high affinity Rev binding in both HIV-1 and EIAV. PLoS ONE 3e2272. [PubMed] 36. Lee, J. H., S. C. Murphy, M. Belshan, W. O. Sparks, Y. Wannemuehler, S. Liu, T. J. Hope, D. Dobbs, and S. Carpenter. 2006. Characterization of functional domains of equine infectious anemia virus Rev suggests a bipartite RNA-binding domain. J. Virol. 803844-3852. [PubMed] 37. Leroux, C., J. L. Cadore, and R. C. Montelaro. 2004. Equine infectious anemia virus (EIAV): what has HIV's country cousin got to tell us? Vet. Res. 35485-512. [PubMed] 38. Liddament, M. T., W. L. Brown, A. J. Schumacher, and R. S. Harris. 2004. APOBEC3F properties and hypermutation preferences indicate activity against HIV-1 in vivo. Curr. Biol. 141385-1391. [PubMed] 39. Löchelt, M., F. Romen, P. Bastone, H. Muckenfuss, N. Kirchner, Y. B. Kim, U. Truyen, U. Rosler, M. Battenberg, A. Saib, E. Flory, K. Cichutek, and C. Munk. 2005. The antiretroviral activity of APOBEC3 is inhibited by the foamy virus accessory Bet protein. Proc. Natl. Acad. Sci. USA 1027982-7987. [PubMed] 40. Mangeat, B., P. Turelli, G. Caron, M. Friedli, L. Perrin, and D. Trono. 2003. Broad antiretroviral defence by human APOBEC3G through lethal editing of nascent reverse transcripts. Nature 42499-103. [PubMed] 41. Mariani, R., D. Chen, B. Schrofelbauer, F. Navarro, R. Konig, B. Bollman, C. Munk, H. Nymark-McMahon, and N. R. Landau. 2003. Species-specific exclusion of APOBEC3G from HIV-1 virions by Vif. Cell 11421-31. [PubMed] 42. Marin, M., K. M. Rose, S. L. Kozak, and D. Kabat. 2003. HIV-1 Vif protein binds the editing enzyme APOBEC3G and induces its degradation. Nat. Med. 91398-1403. [PubMed] 43. Maury, W. 1994. Monocyte maturation controls expression of equine infectious anemia virus. J. Virol. 686270-6279. [PubMed] 44. McGuire, T. C., T. B. Crawford, and J. B. Henson. 1971. Immunofluorescent localization of equine infectious anemia virus in tissue. Am. J. Pathol. 62283-294. [PubMed] 45. McGuire, T. C., D. G. Fraser, and R. H. Mealey. 2002. Cytotoxic T lymphocytes and neutralizing antibody in the control of equine infectious anemia virus. Viral Immunol. 15521-531. [PubMed] 46. McGuire, T. C., D. B. Tumas, K. M. Byrne, M. T. Hines, S. R. Leib, A. L. Brassfield, K. I. O'Rourke, and L. E. Perryman. 1994. Major histocompatibility complex-restricted CD8+ cytotoxic T lymphocytes from horses with equine infectious anemia virus recognize Env and Gag/PR proteins. J. Virol. 681459-1467. [PubMed] 47. Miyagi, E., S. Opi, H. Takeuchi, M. Khan, R. Goila-Gaur, S. Kao, and K. Strebel. 2007. Enzymatically active APOBEC3G is required for efficient inhibition of human immunodeficiency virus type 1. J. Virol. 8113346-13353. [PubMed] 48. Münk, C., T. Beck, J. Zielonka, A. Hotz-Wagenblatt, S. Chareza, M. Battenberg, J. Thielebein, K. Cichutek, I. G. Bravo, S. J. O'Brien, M. Lochelt, and N. Yuhki. 2008. Functions, structure, and read-through alternative splicing of feline APOBEC3 genes. Genome Biol. 9R48. [PubMed] 49. Navarro, F., B. Bollman, H. Chen, R. Konig, Q. Yu, K. Chiles, and N. R. Landau. 2005. Complementary function of the two catalytic domains of APOBEC3G. Virology 333374-386. [PubMed] 50. Newman, E. N., R. K. Holmes, H. M. Craig, K. C. Klein, J. R. Lingappa, M. H. Malim, and A. M. Sheehy. 2005. Antiviral function of APOBEC3G can be dissociated from cytidine deaminase activity. Curr. Biol. 15166-170. [PubMed] 51. Oaks, J. L., T. C. McGuire, C. Ulibarri, and T. B. Crawford. 1998. Equine infectious anemia virus is found in tissue macrophages during subclinical infection. J. Virol. 727263-7269. [PubMed] 52. OhAinle, M., J. A. Kerns, H. S. Malik, and M. Emerman. 2006. Adaptive evolution and antiviral activity of the conserved mammalian cytidine deaminase APOBEC3H. J. Virol. 803853-3862. [PubMed] 53. Olsen, J. C. 1998. Gene transfer vectors derived from equine infectious anemia virus. Gene Ther. 51481-1487. [PubMed] 54. O'Rourke, J. P., H. Hiraragi, K. Urban, M. Patel, J. C. Olsen, and B. A. Bunnell. 2003. Analysis of gene transfer and expression in skeletal muscle using enhanced EIAV lentivirus vectors. Mol. Ther. 7632-639. [PubMed] 55. O'Rourke, K., L. E. Perryman, and T. C. McGuire. 1988. Antiviral, anti-glycoprotein and neutralizing antibodies in foals with equine infectious anaemia virus. J. Gen. Virol. 69667-674. [PubMed] 56. Raabe, M. R., C. J. Issel, and R. C. Montelaro. 1998. Equine monocyte-derived macrophage cultures and their applications for infectivity and neutralization studies of equine infectious anemia virus. J. Virol. Methods 7187-104. [PubMed] 57. Rulli, S. J., Jr., J. Mirro, S. A. Hill, P. Lloyd, R. J. Gorelick, J. M. Coffin, D. Derse, and A. Rein. 2008. Interactions of murine APOBEC3 and human APOBEC3G with murine leukemia viruses. J. Virol. 826566-6575. [PubMed] 58. Russell, R. A., H. L. Wiegand, M. D. Moore, A. Schafer, M. O. McClure, and B. R. Cullen. 2005. Foamy virus Bet proteins function as novel inhibitors of the APOBEC3 family of innate antiretroviral defense factors. J. Virol. 798724-8731. [PubMed] 59. Sawyer, S. L., M. Emerman, and H. S. Malik. 2004. Ancient adaptive evolution of the primate antiviral DNA-editing enzyme APOBEC3G. PLoS Biol. 2e275. [PubMed] 60. Schäfer, A., H. P. Bogerd, and B. R. Cullen. 2004. Specific packaging of APOBEC3G into HIV-1 virions is mediated by the nucleocapsid domain of the gag polyprotein precursor. Virology 328163-168. [PubMed] 61. Schumacher, A. J., G. Hache, D. A. Macduff, W. L. Brown, and R. S. Harris. 2008. The DNA deaminase activity of human APOBEC3G is required for Ty1, MusD, and human immunodeficiency virus type 1 restriction. J. Virol. 822652-2660. [PubMed] 62. Sellon, D. C., F. J. Fuller, and T. C. McGuire. 1994. The immunopathogenesis of equine infectious anemia virus. Virus Res. 32111-138. [PubMed] 63. Sellon, D. C., S. T. Perry, L. Coggins, and F. J. Fuller. 1992. Wild-type equine infectious anemia virus replicates in vivo predominantly in tissue macrophages, not in peripheral blood monocytes. J. Virol. 665906-5913. [PubMed] 64. Sheehy, A. M., N. C. Gaddis, J. D. Choi, and M. H. Malim. 2002. Isolation of a human gene that inhibits HIV-1 infection and is suppressed by the viral Vif protein. Nature 418646-650. [PubMed] 65. Sheehy, A. M., N. C. Gaddis, and M. H. Malim. 2003. The antiretroviral enzyme APOBEC3G is degraded by the proteasome in response to HIV-1 Vif. Nat. Med. 91404-1407. [PubMed] 66. Sponseller, B. A., W. O. Sparks, Y. Wannemuehler, Y. Li, A. K. Antons, J. L. Oaks, and S. Carpenter. 2007. Immune selection of equine infectious anemia virus env variants during the long-term inapparent stage of disease. Virology 363156-165. [PubMed] 67. Stenglein, M. D., and R. S. Harris. 2006. APOBEC3B and APOBEC3F inhibit L1 retrotransposition by a DNA deamination-independent mechanism. J. Biol. Chem. 28116837-16841. [PubMed] 68. Tamura, K., J. Dudley, M. Nei, and S. Kumar. 2007. MEGA4: Molecular Evolutionary Genetics Analysis (MEGA) software version 4.0. Mol. Biol. Evol. 241596-1599. [PubMed] 69. Wedekind, J. E., G. S. Dance, M. P. Sowden, and H. C. Smith. 2003. Messenger RNA editing in mammals: new members of the APOBEC family seeking roles in the family business. Trends Genet. 19207-216. [PubMed] 70. Wiegand, H. L., B. P. Doehle, H. P. Bogerd, and B. R. Cullen. 2004. A second human antiretroviral factor, APOBEC3F, is suppressed by the HIV-1 and HIV-2 Vif proteins. EMBO J. 232451-2458. [PubMed] 71. Yu, Q., R. Konig, S. Pillai, K. Chiles, M. Kearney, S. Palmer, D. Richman, J. M. Coffin, and N. R. Landau. 2004. Single-strand specificity of APOBEC3G accounts for minus-strand deamination of the HIV genome. Nat. Struct. Mol. Biol. 11435-442. [PubMed] 72. Yu, X., Y. Yu, B. Liu, K. Luo, W. Kong, P. Mao, and X. F. Yu. 2003. Induction of APOBEC3G ubiquitination and degradation by an HIV-1 Vif-Cul5-SCF complex. Science 3021056-1060. [PubMed] 73. Zennou, V., D. Perez-Caballero, H. Gottlinger, and P. D. Bieniasz. 2004. APOBEC3G incorporation into human immunodeficiency virus type 1 particles. J. Virol. 7812058-12061. [PubMed] 74. Zhang, H., B. Yang, R. J. Pomerantz, C. Zhang, S. C. Arunachalam, and L. Gao. 2003. The cytidine deaminase CEM15 induces hypermutation in newly synthesized HIV-1 DNA. Nature 42494-98. [PubMed]
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