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Nucleic Acids Res. 2012 January; 40(1): 345–359.
Published online 2011 September 8. doi:  10.1093/nar/gkr694
PMCID: PMC3245923

Biochemical, inhibition and inhibitor resistance studies of xenotropic murine leukemia virus-related virus reverse transcriptase


We report key mechanistic differences between the reverse transcriptases (RT) of human immunodeficiency virus type-1 (HIV-1) and of xenotropic murine leukemia virus-related virus (XMRV), a gammaretrovirus that can infect human cells. Steady and pre-steady state kinetics demonstrated that XMRV RT is significantly less efficient in DNA synthesis and in unblocking chain-terminated primers. Surface plasmon resonance experiments showed that the gammaretroviral enzyme has a remarkably higher dissociation rate (koff) from DNA, which also results in lower processivity than HIV-1 RT. Transient kinetics of mismatch incorporation revealed that XMRV RT has higher fidelity than HIV-1 RT. We identified RNA aptamers that potently inhibit XMRV, but not HIV-1 RT. XMRV RT is highly susceptible to some nucleoside RT inhibitors, including Translocation Deficient RT inhibitors, but not to non-nucleoside RT inhibitors. We demonstrated that XMRV RT mutants K103R and Q190M, which are equivalent to HIV-1 mutants that are resistant to tenofovir (K65R) and AZT (Q151M), are also resistant to the respective drugs, suggesting that XMRV can acquire resistance to these compounds through the decreased incorporation mechanism reported in HIV-1.


Xenotropic murine leukemia virus-related virus (XMRV) is a gammaretrovirus that was first identified in some prostate cancer tissues (1,2) While some subsequent reports confirmed the presence of XMRV in prostate cancer samples (3–6), several others found little or no evidence of the virus in patient samples (7–9). XMRV DNA was also reported in 67% of patients with chronic fatigue syndrome (CFS) (10), but several subsequent studies in Europe and the USA failed to identify XMRV DNA in CFS patients or healthy controls (11–15). Hence, the relevance of XMRV to human disease remains unclear (16) and have been challenged (17). Most recently, it has been reported that XMRV has been generated through recombination of two separate proviruses suggesting that the association of XMRV with human disease is due to contamination of human samples with virus originating from this recombination event (18). Nonetheless, as a retrovirus that can infect human cells, XMRV can be very helpful in advancing our understanding of the mechanisms of retroviral reverse transcription, inhibition and drug resistance. XMRV RT is similar to the Moloney murine leukemia virus (MoMLV) RT, which has been the subject of structural and biochemical studies (19–24). Most of the differences between these gammaretroviral enzymes are at the RNase H domain (Supplementary Figure S1). Comparisons of human immunodeficiency virus type-1 (HIV) RT with MoMLV RT have revealed structural and sequence differences (21). For example, HIV-1 RT is a heterodimer composed of two related subunits (25,26) [reviewed in (27,28)]. Its larger p66 subunit (~66 kDa) contains both the polymerase and RNase H domains; the smaller p51 subunit, (~51 kDa), is derived from the p66 subunit by proteolytic cleavage and its role is to provide structural support and optimize RT’s biochemical functions (29). In contrast, structural studies have demonstrated that MoMLV RT is a monomer of about 74 kDa, although one study reported that it may form a homodimer during DNA synthesis (30). So far, there are no published biochemical or structural studies on XMRV RT. Hence, the present study on this enzyme and its comparison to related enzymes provides an excellent opportunity to advance our biochemical understanding of the mechanism of reverse transcription, its inhibition and drug resistance.


Expression and purification of XMRV, HIV-1 and MoMLV RTs

The plasmid pBSK-XMRV containing the coding sequence of XMRV RT from the VP62 clone (GenBank: DQ399707.1) was chemically synthesized and optimized for bacterial expression by Epoch Biolabs Inc (Missouri City, Texas, USA). The 2013 bp XMRV RT sequence was amplified from pBSK-XMRVRT by PCR, using the forward and reverse primers 1 (all primer sequences are shown in Supplementary Table S1), resulting in NdeI and HindIII restriction sites. Drug resistant XMRV RT mutants Q190M and K103R (equivalent to HIV-1 Q151M RT and K65R) were generated by site-directed mutagenesis using forward and reverse primers 2 and 3. The digested amplicons were ligated into pET-28a (Novagen), resulting into a construct that expresses an N-terminal hexa-histidine tag. pET-28a-MRT encoding full-length wild-type MoMLV RT was provided by Dr M. Modak (New Jersey Medical School, Newark NJ, USA).

Expression and purification of MoMLV and XMRV RTs were carried out similarly to our previously published protocols (23,24). Briefly, RTs were expressed in BL21-pLysS Escherichia coli (Invitrogen) grown at 37°C and induced with 150 µM IPTG at OD600 0.8, followed by 16 h growth at 17°C. A cell pellet from a 3 l culture was incubated with 40 ml lysis buffer (50 mM Tris–HCl, pH 7.8, 500 mM NaCl, 1 mM PMSF, 0.1% NP-40, 1% sucrose and 2 mg/ml lysozyme), then sonicated and centrifuged at 15,000 g for 30 min. The supernatant was diluted 2-fold in Buffer A (50 mM Tri–HCl pH 7.8, 1 mM PMSF, 4% streptomycin sulfate and 10% sucrose), stirred on ice for 30 min and centrifuged. The supernatant was loaded on a Ni-NTA column and bound proteins were washed with 20 ml Buffer B (20 mM Tris–HCl pH 7.5, 500 mM NaCl) and 5 mM imidazole, followed by 20 ml Buffer B with 75 mM imidazole. RT was eluted in 2 ml fractions with 20 ml buffer B containing 300 mM imidazole. Fractions with RT were pooled and further purified by size exclusion chromatography (Superdex 75; GE Healthcare). RTs (>95% pure) were stored in 50 mM Tris–HCl pH 7.0, 100 mM NaCl, 1 mM DTT, 0.1% NP-40 and 30% glycerol in 10 μl aliquots at −20°C. Protein concentrations were determined by measuring UV280 (molar extinction coefficients of 106 and 103 M−1 cm−1 for XMRV and MoMLV RT).

HIV-1 RT was cloned in a pETduo vector and purified as described previously (29,31,32). Oligonucleotide sequences (IDT-Coralville, IA, USA) of DNA/RNA substrates are shown in Supplementary Table S1. Nucleotides were purchased from Fermentas (Glen Burnie, MD, USA). They were treated with inorganic pyrophosphatase (Roche Diagnostics, Mannheim, Germany) as described previously (33) to remove PPi that might interfere with excision assays.

Steady state kinetics

Steady state parameters Km and kcat for dATP incorporation were determined using single nucleotide incorporation gel-based assays. XMRV RT and MoMLV RT reactions were carried out in 50 mM Tris–HCl pH 7.8, 60 mM KCl, 0.1 mM DTT, 0.01% NP-40 and 0.01% bovine serum albumin (BSA) (Reaction Buffer) with 6 mM MgCl2 or 1.5 mM MnCl2, 0.5 mM EDTA, 200 nM or 100 nM Td26/5′-Cy3-Pd18b, 20 nM or 5 nM RT for XMRV and MoMLV RTs, respectively and varying concentrations of dNTP in a final volume of 10 µl. The reactions for HIV-1 RT were carried out in Reaction Buffer with 100 nM Td26/5′-Cy3-Pd18b, 10 nM HIV-1 RT and 6 mM MgCl2 in a 20 µl reaction. All the concentrations mentioned here and in subsequent assays reflect final concentration of reactants otherwise mentioned reactions were stopped after 15 min for XMRV, 4 min for MoMLV RT, and 2.5 min for HIV-1. The products were resolved on 15% polyacrylamide–7M urea gels. The gels were scanned with a Fuji Fla-5000 PhosphorImager (Stamford, CT, USA) and the bands were quantified using MultiGauge. Results were plotted using GraphPad Prism 4. Km and kcat were determined graphically using Michaelis–Menten equation.

Gel mobility shift assays

Formation of RT-DNA binary complex: 20 nM Td31/5′-Cy3-Pd18a (Supplementary Table S1) was incubated for 10 minutes with increasing amounts of MoMLV or XMRV RT in 50 mM Tris–HCl pH 7.8, 0.01% BSA, 5 mM MgCl2 and 10% (v/v) sucrose. The complexes were resolved on native 6% polyacrylamide 50 mM Tris–borate gel and visualized as described above.

Active site titration and determination of KD.DNA

Active site concentrations and kinetic constants of DNA binding for XMRV, HIV-1 and MoMLV RTs were determined using pre-steady state experiments. Reactions with XMRV and MoMLV RTs were carried out in the reaction buffers listed above. For XMRV RT 100 nM protein was pre-incubated with increasing concentrations of Td31/5′-Cy3-Pd18a, followed by rapid mixing with a reaction mixture containing 5 mM MgCl2 and 100 µM next incoming nucleotide (dATP). The reactions were quenched at various times (5 ms to 4 s) by adding EDTA to a final concentration of 50 mM. The amounts of 19-mer product were quantified and plotted against time. The data were fit to the following burst equation:

equation image

where A is the amplitude of the burst phase that represents the RT–DNA complex at the start of the reaction, kobs is the observed burst rate constant for dNTP incorporation, kss is the steady state rate constant and t is the reaction time. The rate constant of the linear phase (kcat) was estimated by dividing the slope of the linear phase by the enzyme concentration. The active site concentration and T/P binding affinity (KD.DNA) were determined by plotting the amplitude (A) against the concentration of T/P. Data were fit to the quadratic equation (Equation 2) using non-linear regression:

equation image

where KD is the dissociation constant for the RT–DNA complex, and [RT] is the concentration of active polymerase. HIV-1 RT’s DNA binding affinity was determined as previously described (29).

Surface plasmon resonance assay

We used surface plasmon resonance (SPR) to measure the binding constants of XMRV and HIV-1 RTs to double-stranded DNA. Experiments were carried out using a Biacore T100 (GE Healthcare). To prepare the sensor chip surface we used the 5′-biotin-Td37/Pd25 oligonucleotide (Supplementary Table S1). One hundred and twenty RUs of this DNA duplex were bound in channel 2 of a streptavidin-coated sensor chip [Series S Sensor Chip SA (certified)] by flowing a solution of 0.1 µM DNA at a flow rate of 10 µl/min in a buffer containing 50 mM Tris pH 7.8, 50 mM NaCl. The binding constants were determined as follows: RT binding was observed by flowing solutions containing increasing concentrations of the enzyme (0.2, 0.5, 1, 2, 5, 10, 20, 50, 100 and 200 nM) in 50 mM Tris pH 7.8, 60 mM KCl, 1 mM DTT, 0.01% NP40 and 10 mM MgCl2 in channels 1 (background) and 2 (test sample) at 30 µl/min. The trace obtained in channel 1 was subtracted from the trace in channel 2 to obtain the binding signal of RT. This signal was analyzed using the Biacore T100 Evaluation software to determine KD.DNA, kon and koff.

Pre-steady state kinetics of dNTP incorporation

The optimal nucleotide incorporation rates (kpol) were obtained by pre-steady state kinetics analysis using single nucleotide incorporation assays. A solution containing XMRV RT (150 nM final concentration) and Td31/5′-Cy3-Pd18a (40 nM) was rapidly mixed with a solution of MgCl2 (5 mM) and varying dATP (5–200 μM) for 0.1 to 6 s before quenching with EDTA (50 mM) (all concentrations in parentheses are final, unless otherwise stated). Products were resolved and quantified as described above. Burst phase incorporation rates and substrate affinities were obtained from fitting the data to Equation 1. Turnover rates (kpol), dNTP binding to the RT-DNA complex (Kd.dATP), and observed burst rates (kobs) were fit to the hyperbolic equation:

equation image

HIV-1 RT’s DNA binding affinity was determined as previously described (29).

Fidelity of DNA synthesis

The fidelity (error-proneness) of XMRV RT was determined and compared with that of MoMLV RT and HIV-1 RT by primer extension assays using 10 nM heteropolymeric Td100/5′-Cy3-Pd18a. Reactions (10 µl) were carried out in Reaction Buffer containing all four dNTPs (100 μM each) or only three dNTPs (missing either dATP, dGTP or dTTP) at 100 μM each. Incubations of the XMRV and MoMLV (50 nM) reactions were at 37°C for 45 min and 30 min for HIV-1 RT (20 nM). Reactions were initiated by adding dNTPs, stopped with equal volume of formamide-bromophenol blue, and an aliquot was run on a 16% polyacrylamide–7M urea gel.

Kinetics of mismatch incorporation

For these experiments, instead of including the next correct nucleotide (dATP) in the polymerase reactions, we used dTTP as the mismatched incoming nucleotide. Hence, 50 nM XMRV RT was pre-incubated with 35 nM Td31/5′-Cy3-Pd18a in reaction mixture. Reactions were initiated by adding dTTP (5–750 µM) and 5 mM MgCl2, followed by incubation (37°C) for 5 min, due to the decreased mismatch incorporation rate of XMRV. For MoMLV RT, 30 nM RT and 20 nM DNA used and the reactions were carried out for 2.5 minutes. For HIV-1, 30 nM RT, 20 nM DNA and 0–200 µM nucleotide were used and the reactions were carried out for 2.5 min. The amount of extended primer was quantified and plotted against the concentration of dTTP. The data were used to derive the Kd.dNTP of incorrect nucleotide binding, the rate kpol (using Equations 1 and 3) and the efficiency of the misincorporation reaction (kpol/Kd.dTTP).

Determination of in vivo fidelity

ANGIE P cells, which contain a retroviral vector (GA-1) that encodes a bacterial β-galactosidase gene (lacZ) and a neomycin phosphotransferase gene, were plated (5 × 106 cells/100 mm dish) and after 24 h were transfected using the calcium phosphate precipitation method with a plasmid expressing either XMRV or amphotropic MLV (AM-MLV) (three independent transfections per vector). After 48 h, the culture medium with XMRV or (AM-MLV) was harvested, serially diluted and used to infect D17 target cells (2 × 105 cells/60 mm dish) in the presence of polybrene. The infected D17 cells were selected for resistance to G418 (400 µg/ml) in the presence of 1 µM AZT to suppress reinfection, and characterized by staining with 5-bromo-4-chloro-3-indoyl-β-d-galacto-pyranoside (X-Gal) ~2 weeks after G418 selection. The frequencies of inactivating mutations in lacZ quantified as described before (blue versus white colonies) (34).

Processivity of DNA synthesis—trap assay

Processivity reactions were carried out in Reaction Buffer containing 20 nM Td100/Pd18, 100 μM of each dNTP, 30 nM HIV-1 RT, 50 nM MoMLV RT or 100 nM XMRV RT and 1μg/μl unlabeled calf thymus DNA trap in 50 μL. Enzymes were pre-incubated with Td100/Pd18 for 1 min before adding dNTPs (100 µM each) together with the calf thymus DNA trap. Reactions were incubated at 37°C, and 10 μl aliquots were taken out at 3, 7.5 and 15 min for HIV-1 RT or at 7.5, 15 and 30 min for XMRV RT and MoMLV RT, and mixed with equal volume of loading dye. The effectiveness of the trap was determined by pre-incubating the enzyme with the trap before adding Td100/Pd18. Control DNA synthesis was measured in absence of trap under the same conditions. Reaction products were resolved as above.

Single turnover processivity assays

Thirty nanomolar Td31/5′-Cy3-Pd18a was pre-incubated for 10 min with 100 nM XMRV or 50 nM MoMLV RT in Reaction Buffer, then rapidly mixed with 100 µM dNTPs, 5 mM MgCl2 for varying times (0.1–45 s) before quenching with EDTA (50 mM final). Single turnover processivity of HIV-1 RT was assayed with 40 nM enzyme, 20 nM DNA and 50 μM of each nucleotide were used. The reaction products were resolved and quantified as described above. The data were fit to a one-phase exponential decay equation for the elongation of the 18-mer primer. The rates of appearance and extension of products from subsequent nucleotide incorporations (19- and 27-mer) were obtained by fitting the intensities of corresponding bands to double exponential (Equation 4):

equation image

where A is the amplitude, P is the amount of 19-mer, 20-mer or higher length products, k1 is the rate of product generation, k2 the rate of subsequent elongation and C a constant (29,35).

Assays for reverse transcriptase inhibition

DNA synthesis by 50 nM XMRV RT or MoMLV RT was carried out in Reaction Buffer using 20 nM Td100/5′-Cy3-Pd18a, 2.5 µM dNTP, 5 mM MgCl2 and varying amounts of NRTI (0–100 µM). Reactions were quenched with 95% formamide after 1 h incubation at 37°C (38). In experiments with aptamers 10 nM XMRV RT, 20 nM Td31/5′-Cy3-Pd18a and 50 µM dNTPs were used in the presence of varying amounts of aptamer for 30 min (0–500 nM for m.1.3; 0–25 nM for m.1.4 and m.1.1FL). The inhibition of DNA polymerization was monitored by resolving the products on 15% polyacrylamide–7 M urea gels and visualized as described above. Bands corresponding to full extension products were quantified using MultiGauge Software and IC50s were obtained from dose–response curves using GraphPad Prism.

PPi- and ATP-dependent excision and rescue of T/PAZT-MP or T/PEFdA-MP

The ability of enzymes to use PPi or ATP to unblock template-primers that had AZT-MP (T/PAZT-MP) or EFdA-MP (T/PEFdA-MP) at their 3′ primer ends was measured as follows: 20 nM of T/PAZT-MP or T/PEFdA-MP were prepared as described before (32). They were incubated at 37°C with either 60 nM HIV-1 RT or 200 nM XMRV RT in the presence of 0.15 mM PPi or 3.5 mM ATP for PPi- or ATP-dependent rescue reactions, respectively. Reactions were initiated by the addition of MgCl2 (6 mM). Aliquots were removed at different times (0–90 min) and analyzed as above. Rescue assays were performed in the presence of 100 µM dATP to prevent EFdA-MP reincorporation, 0.5 µM dTTP, 10 µM ddGTP and 10 mM MgCl2.

Molecular modeling

The sequence of XMRV RT from the VP62 clone was aligned with that of MoMLV RT (PDB: 1RW3) (21,22) using ClustalW. To generate the homology model of XMRV RT, we used the Prime protocol of the Schrödinger software suite (Schrödinger Inc. NY). The resulting molecular model was further energy minimized by OPLS2005 force field using the Impact option of Schrödinger. The final model was validated with PROCHECK v.3.5.4.


Comparison of RT sequences

The XMRV and MoMLV enzymes are closely related (~95% sequence identity) with most of the differences between them being in the RNase H domain (Supplementary Figure S1). While XMRV and MoMLV differ significantly from HIV-1 RT, the known polymerase motifs (A–F) are well conserved in all three enzymes (Supplementary Figure S1). Specifically, the active site aspartates in Motifs A and C (Figure 9) (D150, D224, D225 in XMRV RT; D150, D224, D225 in MoMLV RT; D110, D185, D186 in HIV-1 RT) are conserved in all three RTs. Also, the three enzymes are similar in Motif B, which is involved in dNTP binding and multidrug resistance (AZT and dideoxy-nucleoside drugs) through the decreased incorporation mechanism (27,39–41). Specifically, all three enzymes have a glutamine at the start of this motif (Q151 in HIV-1 RT, Q190 in XMRV RT and Q190 in MoMLV RT). Motif D includes HIV-1 RT residues L210 and T215, which when mutated they enhance excision of AZT from the AZT-terminated primer terminus. This motif is mostly different in XMRV and MoMLV RTs, where the corresponding residues are N226 and A231 (Supplementary Figure S1). K219 of HIV-1 RT Motif D is proximal to the dNTP-binding pocket and is also conserved in the other enzymes (K235). The DNA primer grip (Motif E) (36,42) in HIV-1 RT (M230G231Y232) is slightly different in the gammaretroviral enzymes (L245G246Y247). Motif F at the fingers subdomain of all enzymes has two conserved lysines that bind the triphosphate of the dNTP (K65 and K72 in HIV-1 RT; K103 and K110 in XMRV and MoMLV RTs).

Figure 9.
Molecular model of XMRV RT. Ribbons diagram of XMRV RT with the conserved polymerase Motifs color-coded: Motif A (green), B (brown), C (purple), D (red), E (orange) and F (blue). The residues that differ from MoMLV’s polymerase domain are shown ...

Several HIV-1 residues involved in NRTI resistance have the resistance mutations in XMRV and MoMLV RTs (Table 1). Hence, XMRV and MoMLV RTs have a Val as the X residue (codon 223) of the conserved YXDD sequence of Motif C. An M184V mutation at this position in HIV-1 RT causes strong, steric hindrance-based, resistance to 3TC and FTC (43–45), and to a lesser extent to ddI, ABC [reviewed in (46)], and translocation defective RT inhibitors (TDRTIs) (43) (Table 1). Similarly, the M41L mutation, which causes excision-based AZT resistance in HIV is already present in XMRV and MoMLV RT (L81, Table 1). The gammaretroviral enzymes differ from HIV-1 RT in several other HIV drug resistance sites (HIV residues 62, 67, 69, 70, 75, 77, 115, 210, 215) (Table 1). Finally, there are also differences in residues that are essential for NNRTI binding in HIV-1 RT: W229 changes to Y268 in XMRV RT, Y181 to L220, Y188 to L227 and G190 to A229 (Table 1) (27,28,47–49).

Table 1.
HIV-1 RT drug resistance mutations with wild-type XMRV RT and MoMLV RT residues

Preparation of MoMLV and XMRV RTs

The sequence coding for full-length XMRV RT from the VP-62 clone (NCBI RefSeq: NC_007815) (1) was optimized for expression in bacteria, synthesized by Epoch Biolabs and cloned as described in ‘Materials and Methods’ section. Both XMRV RT and MoMLV RT were tagged with a hexahistidine sequence at the N-terminus and expressed with a yield of ~2 mg/l of culture. Purified enzymes (>95% pure, Supplementary Figure S2) were stored at −20°C. The presence of NP-40 or glycerol was critical for enzyme stability.

Steady state kinetics of nucleotide incorporation

Initial polymerase activity assays using Td31/5′-Cy3-Pd18a displayed overall slower polymerase activity of XMRV RT compared to HIV-1 and MoMLV RTs. This observation led us to investigate the steady state nucleotide incorporation properties of XMRV RT using single nucleotide incorporation assays. The estimated values for kcat (19.9 min−1 for HIV-1 RT (32), 3.3 min−1 for MoMLV RT, 0.6 min−1 for XMRV RT) and Km.dNTP (0.07 µM for HIV-1 RT (32), 3.3 μM for MoMLV RT, 3.0 µM for XMRV RT) show that XMRV RT has a drastically reduced efficacy (kcat/Km.dNTP) at nucleotide incorporation, compared to both MoMLV and HIV-1 RTs.

DNA binding affinity

To assess if the efficiency of XMRV RT was also affected by a lower DNA binding affinity we measured the DNA binding affinity of the enzymes using three methods: gel-mobility shift assays, pre-steady state kinetics and SPR. Gel-mobility shift assays showed that the KD.DNA for XMRV RT was marginally higher than that for HIV-1 RT and MoMLV RT (data not shown) (50) suggesting weaker binding to DNA.

DNA binding affinity using pre-steady state kinetics

Pre-steady state kinetics allows estimation of the fraction of active polymerase sites as well as the KD.DNA value for the enzyme. The amplitudes of DNA extensions using XMRV RT and/or MoMLV RT at varying DNA concentrations were plotted against the DNA concentration and the data were fit to the quadratic equation (Equation 2), yielding a KD.DNA of 33 nM for XMRV RT, 19 nM for MoMLV RT (Table 2) and 12.5 nM for HIV-1 RT (32). These values did not change significantly when tested with DNA of different lengths (data not shown). Hence, the transient kinetic experiments confirmed the findings of the gel-mobility shift assays showing XMRV RT to have lower DNA binding affinity than HIV-1 RT.

Table 2.
Kinetic parameters of DNA binding and synthesis by HIV-1 and XMRV RTs

Binding kinetics of XMRV and HIV-1 RT to double-stranded DNA

Measurements of KD.DNA using gel-mobility shift assays and pre-steady state kinetic methods do not offer insights regarding the kinetics of binding and release of nucleic acid from the viral polymerases. Hence, we used SPR to measure directly DNA binding and the DNA dissociation components of the KD.DNA. We attached on the SPR chip a nucleic acid biotinylated at the 5′ template end and immobilized it on a streptavidin sensor chip. Various concentrations of either XMRV or HIV-1 RT were flowed over the chip to measure the association (kon) and dissociation (koff) rates of the enzymes in real time (Figure 1). HIV-1 RT had considerably slower dissociation rates than XMRV RT, and longer dissociation phases were needed to obtain reliable values.

Figure 1.
Assessment of KD.DNA, kon and koff using surface plasmon resonance. SPR was used to measure the binding affinity of RTs to a nucleic acid substrate. Increasing concentrations of each RT (0.2, 0.5, 1, 2, 5, 10, 20, 50, 100 and 200 nM) were injected ...

Several methods were tested to best fit our data. The ‘heterogeneous ligand’ method gave the best fit for both XMRV and HIV-1 RT. In this model the x2 values for DNA binding to XMRV and HIV-1 RT were 9.3 RU2 and 48.1 RU2, respectively, compared to 15.1 RU2 and 152 RU2 when we tried fitting the data in a ‘homogeneous ligand’ model. The former model assumes that RT binds DNA in two different modes and provides two association (kon) and two dissociation constants (koff).

Our data show that XMRV RT has a slightly faster rate of association (kon) than HIV-1 RT. We measured two kon values of 7.3 × 106 M−1s−1 and 8.2 × 104 M−1s−1 for XMRV RT versus 7.6 × 105 M−1s−1 and 1.2 × 106 M−1s−1 for HIV-1 RT. Interestingly, the dissociation rate of XMRV RT was significantly faster than that of HIV-1 RT (0.28 s−1 and 0.0045 s−1 for XMRV RT and 7.8 × 10−4 s−1and 0.0076 s−1 for HIV-1 RT) (Table 3). This difference in dissociation rate resulted in a KD.DNA at least 1 order of magnitude higher for XMRV RT compared to HIV-1 RT (38 and 54 nM versus 1.0 and 6.1 nM for XMRV and HIV-1 RT, respectively) (Table 3).

Table 3.
DNA binding constants for HIV-1 and XMRV RTs from surface plasmon resonance

Nucleotide binding affinity and optimal incorporation efficiency

A transient-state kinetics approach was used to estimate the dNTP binding affinity (Kd.dNTP) and maximum nucleotide incorporation rate (kpol) (55). The rates at varying concentrations of next incoming nucleotide (dATP) were determined by plotting the amount of extended primer as a function of time. The rates were then plotted against dATP concentration. The data were fit to a hyperbola (Equation 3). The Kd.dNTP for XMRV RT is 26.6 μM and the kpol is 8.9 s−1 (Figure 2) (Table 2). Under similar conditions the Kd.dNTP and kpol were 1.3 µM and 24.4 s−1 for HIV-1 RT (29) and 25 µM and 14.1 s−1 for MoMLV RT.

Figure 2.
Pre-steady state kinetics of nucleotide incorporation by XMRV RT. 150 nM XMRV RT was pre-incubated with 40 nM Td31/5′-Cy3-Pd18a rapidly mixed with a solution containing MgCl2 (5 mM) and varying concentrations of dATP: 25 µM ...

Fidelity of nucleotide incorporation

To assess whether XMRV RT displays high nucleotide incorporation fidelity we monitored the incorporation of three dNTPs by XMRV RT and compared with HIV-1 RT (52). The results of fidelity assay are shown in Figure 3. The lanes marked ‘4dNTPs’ for all enzymes represent the DNA synthesis using a Td100/5′-Cy3-Pd18a template-primer in the presence of all four dNTPs. The subsequent lanes, marked ‘-dNTP’, correspond to the synthesis of DNA in the absence of that specific deoxynucleotide triphosphate. The comparison of the DNA synthesis in the absence of one nucleotide by HIV-1 RT, MoMLV RT and XMRV RT shows that HIV-1 and MoMLV RTs were able to misincorporate and extend the primer beyond the missing nucleotide more efficiently than XMRV RT, suggesting that the latter is a less error prone DNA polymerase. It should be noted that the higher fidelity of XMRV is not the result of measuring a smaller number of errors because of the decreased replication rate, as the assay conditions were optimized to allow production of the same amount of full length product in the presence of all four dNTPs for and MoMLV RTs. To further investigate the fidelity of DNA synthesis by XMRV RT, the kinetics of mismatch nucleotide incorporation were carried out in a quantitative manner by monitoring the incorporation of single mismatched nucleotide under pre-steady state conditions. The estimated KD.dTTP (mismatch) and kpol values show that XMRV RT has a lower affinity for a mismatched nucleotide but comparable turnover number than MoMLV RT, suggesting that the observed higher fidelity over MoMLV RT is due to differences during the nucleotide-binding step (Table 4). However, compared to HIV-1 RT, XMRV RT has decreased both affinity and incorporation rate, suggesting that its higher fidelity is the result of both decreased binding of mismatched nucleotides and slow rate of incorporation.

Figure 3.
Comparison of in vitro fidelity of HIV-1, MoMLV and XMRV RTs. Extension of 10 nM Td100/5′-Cy3-Pd18a by HIV-1 RT, MoMLV RT or XMRV RT (20, 50 and 50 nM, respectively) in the presence of 150 µM each of three out of ...
Table 4.
Kinetics of mismatch incorporation for HIV-1, MoMLV and XMRV RTs

Intracellular fidelity by measuring LacZ mutation frequency

The ANGIE P cells used for this assay are a D17-based encapsidating cell line and contain an MLV-based retroviral vector (GA-1), which encodes a bacterial β-galactosidase gene (lacZ) and a neomycin phosphotransferase gene (neo). Replication fidelity is a measure of the frequency of lacZ inactivation and was determined by measuring lacZ non-expressing white colonies. The results show that the number of white colonies was not statistically different in the case of XMRV as compared to AM-MLV, suggesting that under these conditions the fidelity of XMRV is not significantly different than that of AM-MLV (Figure 4).

Figure 4.
Comparison of in vivo fidelity of XMRV with amphotropic MLV. The ANGIE P cells used for this assay contain a retroviral vector (GA-1), which encodes a bacterial β-galactosidase gene (lacZ) and a neomycin phosphotransferase gene. Replication fidelity ...

Processivity of DNA synthesis

Processivity is the probability of translocation of a polymerase along a template and predicts the number of cycles of nucleotide incorporation during one productive enzyme–DNA binding event. We assessed XMRV RT’s processivity of DNA synthesis in comparison to HIV and MoMLV RTs using both a gel-based trap assay and a quantitative pre-steady state assay. In the gel-based assay, the enzymes were pre-incubated with template-primer, then the reaction was initiated by the addition of all four nucleotides together with calf thymus DNA, which was used as a trap to bind free enzyme dissociated from the substrate during the course of the reaction (38). The length of the DNA product is an inverse measure of termination probability, as previously described. As a control, we used lanes where no trap was present; establishing that the same amount of total polymerase activity (processive and non-processive) is provided in all cases. The results indicate that XMRV RT is less processive than HIV-1 and MoMLV RTs with shorter DNA product after 30 min of reaction in the presence of trap (Figure 5).

Figure 5.
Processivity (trap assay) of HIV-RT, MoMLV RT and XMRV RT. DNA synthesis was monitored in the presence of calf thymus DNA as an enzyme trap. Each enzyme (30 nM HIV RT, 100 nM MoMLV RT or 100 nM XMRV RT) was pre-incubated with 40 nM ...

To measure processivity quantitatively we applied a single turnover processivity assay developed by Patel et al. (35) (Figure 6). In this assay, the rates of consecutive nucleotide incorporations under single turnover conditions are monitored. The rate of elongation incorporation (k1) and the rate of processive DNA synthesis (k2) (Equation 4) were calculated at several template positions for each enzyme. The ratio of the rate of processive DNA synthesis to the rate of nucleotide incorporation (k2/k1) is referred to as the processivity index (35). The absolute values of these constants for HIV-1 RT, XMRV and MoMLV RT and their ratios are collected in Table 5. XMRV RT is clearly the least processive for each extension product. The difference in processivity varies significantly depending on sequence or sequence context (decrease in processivity from 3-fold up to 10-fold). While the current data do not allow generalization of rules for pausing at specific sites, this clearly shows consistently that XMRV is not as efficient as MoMLV RT in polymerizing processively through ‘difficult spots’.

Figure 6.
Single-turnover processivity assays. 30 nM Td31/Cy3-Pd18a was combined with 100 nM XMRV RT or 50 nM MoMLV RT in RT buffer before rapidly mixing with all four dNTPs (100 µM each) and 5 mM MgCl2 for varying ...
Table 5.
Single turnover processivity parameters of HIV-1, MoMLV and XMRV RTs

Susceptibility of XMRV RT to NRTIs, TDRTIs and NNRTIs

Previous studies have shown that XMRV is inhibited by some antivirals (53–56). However, the susceptibility of XMRV RT has not been tested against a wide variety of nucleoside RT inhibitors (NRTIs) that block replication by chain-terminating the primer, or by preventing translocation after their incorporation into the nascent DNA chain (TDRTIs) (32,57,58). In addition, the susceptibility of XMRV RT to non-nucleoside RT inhibitors (NNRTIs) or RNA aptamers that can be selected to block reverse transcriptases (59–63) has not been established.

Hence, we performed gel-based primer extension assays in the presence of various inhibitors. As shown in Table 6, most of the HIV-1 RT inhibitors also block XMRV RT with significantly varying IC50s. The most potent inhibitors tested were ENdA (4′-ethynyl-2-amino-2′-deoxyadenosine) followed by EFdA. EFdA was also potent at inhibiting wild-type XMRV replication in cell culture with an EC50 of 40 nM from three independent experiments (standard error was 10 nM).

Table 6.
Inhibition of XMRV and MoMLV RTs

Unlike HIV-1 RT, XMRV RT and MoMLV RT lack the two tyrosine residues (Y181 andY188 in HIV-1 RT) (Supplementary Figure S1) that are known to contribute to NNRTI binding. Hence, the gammaretroviral enzymes were not inhibited by the NNRTIs tested (TMC-125 and efavirenz) (Supplementary Figure S3).

Susceptibility of XMRV RT to RNA aptamers

We also tested XMRV RT’s susceptibility to three independent RNA aptamers that had been previously selected against MoMLV RT (60). The aptamers inhibited XMRV RT to varying extents with IC50s ranging from 2 to 52 nM (Figure 7). Most notable was the m.1.1FL aptamer which gave IC50s of 2 and 4 nM for XMRV RT (Figure 7) and MoMLV RT respectively, without inhibiting HIV-1 RT (data not shown). These inhibition assays utilized truncated forms of aptamers m.1.3 and m.1.4 lacking the original primer-binding segments of the aptamers, demonstrating that these 5′ and 3′ segments are not required.

Figure 7.
Inhibition of XMRV RT by RNA aptamers. 10 nM XMRV RT was incubated with increasing amounts of RNA aptamer in Reaction Buffer for 5 min at 37°C followed by addition of 20 nM Td31/Cy3-Pd18a and 50 µM of each ...

PPi-mediated excision activity of XMRV RT

A key mechanism of NRTI resistance in HIV-1 RT is based on inhibitor excision from the primer end, using a pyrophospholytic reaction (64,65). The pyrophosphate donor in vivo is likely to be ATP, although PPi can efficiently unblock NRTI-terminated primers. This excision activity is present in wild-type HIV-1 RT, and is enhanced in the presence of AZT-resistance mutations. We measured the ability of wild-type XMRV to unblock primers terminated with AZT or EFdA in the presence of PPi. We found that unlike HIV-1 RT that excised AZT-MP efficiently under these conditions, XMRV RT had considerably lower excision activity (Figure 8). Similar excision experiments where ATP was used instead of PPi showed that XMRV is very inefficient in ATP-based excision as compared to HIV-1 RT (data not shown).

Figure 8.
PPi-mediated unblocking of AZT-(A) and EFdA-(B) terminated DNA. About 20 nM of (A) AZT- or (B) EFdA-terminated Td31/Cy3-Pd18c (T/PAZT-MP or T/PEFdA-MP) was incubated with HIV-1 RT (60 nM) or XMRV RT (200 nM) in the presence of ...

Susceptibility of mutant XMRV RTs to AZT-TP and tenofovir-DP

The HIV-1 RT mutation Q151M confers resistance to AZT by enhancing discrimination of the nucleotide analog leading to its reduced incorporation (37,66–68). Another HIV-1 RT mutation, K65R, decreases susceptibility to tenofovir (69,70). Since AZT and tenofovir are potent inhibitors of XMRV (Table 6) (54–56), we wanted to investigate whether the XMRV RT mutant equivalents of HIV Q151M and K65R (XMRV Q190M and K103R) would confer XMRV RT resistance to AZT and tenofovir. We constructed these mutant clones and tested their susceptibility to AZT and tenofovir in the same manner as wild-type XMRV RT. Interestingly, Q190M XMRV RT has a decreased susceptibility to AZT (approximately 5-fold increase in the IC50). Similarly, the K103R XMRV RT mutant enzyme was less susceptible to tenofovir, increasing the IC50 by at least 2-fold.

Molecular model of XMRV RT

Given the significant sequence similarity between XMRV and MoMLV RTs, the resulting homology model of XMRV RT is highly similar to MoMLV RT (>1.5Å rms) and of excellent quality. Since the input structure of MoMLV RT did not contain the RNase H domain of the enzyme, the XMRV RT model is also missing this domain. The molecular model of the polymerase domain of XMRV RT is shown in Figure 9. An alignment of the MoMLV RT crystal structure (22) with the XMRV RT homology model highlights the few changes in the polymerase domain of XMRV RT. These are L29 (P in MoMLV), Q234 (L in MoMLV), R238 (Q in MoMLV) and N422 (D in MoMLV). From these, residue 422 is located in the nucleic acid binding cleft and may contribute to differences in the interactions with nucleic acid substrate. However, most of the differences between the gammaretroviral enzymes are in their RNase H domains and also in the first 30 N-terminal residues of the polymerase domain, for which we do not have structural information since they were not included in the original crystal structure of MoMLV RT. The differences between XMRV RT and HIV-1 RT are very significant. Unlike the HIV enzyme, XMRV RT appears to be a monomer in solution. Moreover, alignment of the HIV-1 RT–DNA complex with XMRV RT based on their active sites at the palm subdomains shows that the thumb subdomain of XMRV RT would have to be repositioned to be able to accommodate nucleic acid.


Early studies reported the presence of XMRV in stromal cells from prostate cancer patient samples and also in CFS clinical samples. Some of the subsequent studies confirmed these findings whereas several others failed to identify XMRV in prostate cancer or in CFS patients, even when same samples were used (71). It was recently reported that human sample contamination with mouse DNA can occur frequently (17,72–74). Moreover, two coauthors from this study have recently demonstrated that XMRV is the product of recombination events between two MLV proviruses, suggesting that XMRV may not be relevant to human disease (18). Nonetheless, XMRV is still an important human retrovirus and comparisons with HIV can provide valuable insights into the fundamental mechanisms of DNA polymerization, RT inhibition and drug resistance. (75).

There is high degree of sequence similarity between the XMRV and MoMLV RTs (95% amino acid identity), and much less so with HIV-1 RT. Based on gel filtration experiments we conclude that unlike HIV-1 RT, but similar to MoMLV RT, XMRV RT exists in solution primarily as a monomer. We also included comparisons with HIV-1 RT in this study as it has been extensively studied and provides an excellent frame of reference.

We report here that there are significant differences in the DNA polymerization efficiency of the three enzymes. Although the polymerase active sites of the XMRV and MoMLV enzymes are almost identical, there is a considerable decrease in the efficiency of nucleotide incorporation by XMRV RT. Most differences in sequence are at the RNase H domain and are likely to affect polymerization by changing the positioning of DNA at the polymerase active site.

We have recently solved the crystal structure of the XMRV RNase H at high resolution (1.5Å) (pdb 3P1G) (Kirby, K.A. et al., submitted for publication). We observed major differences in affinity for nucleic acid that we determined with gel-mobility shift assays and with pre-steady state kinetics. SPR experiments dissected in more detail the specific defect of XMRV RT in binding DNA. Surprisingly, XMRV RT can associate very rapidly with DNA, even more so than HIV-1 RT (Figure 1 and Table 3). However, it dissociates from DNA much faster than the HIV enzyme, resulting in an overall reduced binding affinity. A possible reason for the fast association and dissociation rates of XMRV RT may be the apparent monomeric state, which might offer facile access to the nucleic acid binding cleft, although with less contacts and lower affinity than HIV-1 RT, which is a heterodimer (76,77). This high rate of XMRV RT dissociation from DNA likely contributes to the decreased processivity observed in our study, and may have consequences in the recombination rates of this virus.

Previous sequences of XMRV from prostate cancer tumors showed low variability, suggesting that the virus may have a high fidelity of replication (1,10). Our study demonstrated that HIV-1 RT and MoMLV RT incorporated mismatched nucleotides and extended past the mismatches more efficiently than XMRV RT. Pre-steady state kinetics established that the higher overall fidelity of XMRV RT over MoMLV RT is due to a lower affinity for mismatched nucleotides. When compared to HIV-1 RT, however, XMRV RT has differs in both the nucleotide binding and incorporation steps. Nonetheless, XMRV did not have higher fidelity than a related amphotropic MLV virus or HIV-1 in a cell-based assay. It is possible that the high dNTP concentration in dividing cells can suppress mismatching events. We have previously shown (39) that as nucleotide concentrations vary in different cell lines, this can affect viral susceptibility to NRTIs, and possibly in this case also incorporation of mismatched nucleotides. Additional cell-based studies using multiple cell lines and a large panel of viruses should provide a better understanding of the relation between in vivo and in vitro fidelity.

Early studies have reported susceptibility of XMRV to some antiretrovirals that have been used in the treatment of HIV infection (53–56). In those studies the compounds were tested at the virus level. To better understand the interactions of inhibitors at their RT target level we tested here the ability of these and several more compounds to block the polymerase activity of XMRV RT. We found that two TDRTIs, EFdA-TP and ENdA-TP were very potent RT inhibitors (IC50s: 0.43 µM and 0.14 µM, respectively). Unlike other NRTIs, these compounds have a 3′ OH group and are known to efficiently inhibit HIV replication by blocking translocation (32,58,78). Preliminary experiments demonstrated that they also block XMRV RT by the same mechanism (data not shown).

In HIV, moderate resistance to EFdA is conferred by the emergence of the M184V mutation at the conserved X position of the conserved YXDD motif of the polymerase active site. Interestingly, XMRV and MoMLV RTs already have a valine (V223) at this position. This difference is likely to contribute to the better potency of EFdA against HIV-1 RT than XMRV RT or MoMLV RT (57,58). It may also contribute to the decreased ability of XMRV RT to unblock chain-terminated primers, as was also reported for M184V HIV-1 RT (79) and to the enhanced fidelity reported here for XMRV RT, which is also reminiscent of the previously reported high fidelity of M184V HIV-1 RT (80,81). Nonetheless, despite the presence of a Val in the YMDD motif of XMRV RT we found EFdA to inhibit very efficiently replication-competent or pseudotyped XMRV, with submicromolar EC50s (40 and 110 nM, respectively).

Previously, highly potent aptamers were selected to inhibit MoMLV RT (60). We demonstrate here that the three aptamers we tested have varying potency against XMRV RT. Aptamer m.1.1FL was the most potent inhibitor of XMRV RT and MoMLV RT in in vitro assays (IC50 = 2 and 4 nM, respectively). The fact that XMRV and MoMLV RTs are inhibited by the same aptamers at comparable efficiencies suggests that the RT residues that are different in the two enzymes are not critical to the binding of the aptamer. In contrast, heterodimeric HIV-1 RT has a very different binding cleft and is not inhibited by these aptamers.

Tenofovir is an essential component of HIV therapies and is also a potent inhibitor of XMRV RT. HIV resistance to tenofovir is conferred by a single codon mutation (K65R). HIV-1 RT residue 65 is known to interact with the incoming dNTP or the activated tenofovir analog (tenofovir diphosphate) (82). K65R causes resistance to tenofovir by lowering the kpol for the incorporation of the inhibitor into the nascent DNA. We prepared XMRV RT with the equivalent mutation, K103R, and determined that it has decreased susceptibility to tenofovir. Hence, it is possible for XMRV to develop tenofovir resistance through the same mechanism as HIV-1 RT. HIV resistance to AZT can occur by either decreased binding/incorporation or increased excision of the chain-terminating NRTI (33,83). HIV-1 RTs containing the M41L, D67N, K70R, T215Y/F, K219E/Q mutations show enhanced removal of AZT. Our experiments show that unlike wild-type HIV-1 RT, XMRV RT is not able to excise NRTI-terminated primers. Similarly, it was previously shown that MoMLV RT is not capable of unblocking chain-terminated primers (33).

In HIV, decreased binding of AZT is conferred initially in the presence of the primary Q151M mutation, followed by secondary mutations F77L, A62V, V75I and F116Y (27,47,84). XMRV RT already differs from wild-type HIV-1 RT in the first three of these residues (P104, Q113 and L115 versus A62, V75 and F77) (Table 1). We demonstrated that introducing the primary Q→M mutation at the equivalent XMRV RT site (Q190M) resulted in an enzyme with decreased susceptibility to AZT. Hence, it appears that these residues can confer AZT resistance to XMRV by reduced incorporation of nucleotide analogs, as is the case in HIV-2 (41). At this point we do not know if introduction of as yet unknown mutations could endow XMRV RT with the ability to unblock chain-terminated nucleic acids. The details of the molecular mechanism of XMRV resistance to tenofovir and AZT are under investigation.

In conclusion, our study provides detailed biochemical analysis of the mechanisms of polymerization, inhibition, fidelity, processivity and drug resistance of XMRV RT and how it compares with the closely related enzyme MoMLV RT and the more distantly related HIV-1 RT. The findings enhance our understanding of the basic mechanisms of reverse transcription.


Supplementary Data are available at NAR Online.


NIH grants (AI076119, AI079801, and AI094715, to S.G.S.), (AI074389, to D.H.B.), (AI079801 to M.A.P.); NIH Bench-to-Bedside Award and the Intramural Research Program of the NIH, National Cancer Institute, Center for Cancer Research (to V.K.P); Ministry of Knowledge and Economy, Bilateral International Collaborative R&D Program, Republic of Korea; Canadian Institutes of Health Research (CIHR) and University of Missouri (to S-L.L.); amfAR Mathilde Krim Fellowship and a CIHR Fellowship (to B.M.). Funding for open access charge: NIH grants (AI076119, AI094715, AI074389, AI079801).

Conflict of interest statement. None declared.

Supplementary Material

Supplementary Data:


The content of this publication does not necessarily reflect the views or policies of the Department of Health Human Services, nor does mention of trade names, commercial products or organizations imply endorsement by the U.S. Government.


1. Urisman A, Molinaro RJ, Fischer N, Plummer SJ, Casey G, Klein EA, Malathi K, Magi-Galluzzi C, Tubbs RR, Ganem D, et al. Identification of a novel gammaretrovirus in prostate tumors of patients homozygous for R462Q RNASEL variant. PLoS Pathog. 2006;2:e25. [PMC free article] [PubMed]
2. Malathi K, Dong B, Gale M, Jr, Silverman RH. Small self-RNA generated by RNase L amplifies antiviral innate immunity. Nature. 2007;448:816–819. [PubMed]
3. Arnold RS, Makarova NV, Osunkoya AO, Suppiah S, Scott TA, Johnson NA, Bhosle SM, Liotta D, Hunter E, Marshall FF, et al. XMRV infection in patients with prostate cancer: novel serologic assay and correlation with PCR and FISH. Urology. 75:755–761. [PubMed]
4. Dong B, Kim S, Hong S, Das Gupta J, Malathi K, Klein EA, Ganem D, Derisi JL, Chow SA, Silverman RH. An infectious retrovirus susceptible to an IFN antiviral pathway from human prostate tumors. Proc Natl Acad Sci. USA. 2007;104:1655–1660. [PubMed]
5. Knouf EC, Metzger MJ, Mitchell PS, Arroyo JD, Chevillet JR, Tewari M, Miller AD. Multiple integrated copies and high-level production of the human retrovirus XMRV (xenotropic murine leukemia virus-related virus) from 22Rv1 prostate carcinoma cells. J. Virol. 2009;83:7353–7356. [PMC free article] [PubMed]
6. Schlaberg R, Choe DJ, Brown KR, Thaker HM, Singh IR. XMRV is present in malignant prostatic epithelium and is associated with prostate cancer, especially high-grade tumors. Proc. Natl Acad. Sci. USA. 2009;106:16351–16356. [PubMed]
7. Sabunciyan S, Mandelberg N, Rabkin CS, Yolken R, Viscidi R. No difference in antibody titers against xenotropic MLV related virus in prostate cancer cases and cancer-free controls. Mol. Cell. Probes. 25:134–136. [PMC free article] [PubMed]
8. Verhaegh GW, de Jong AS, Smit FP, Jannink SA, Melchers WJ, Schalken JA. Prevalence of human xenotropic murine leukemia virus-related gammaretrovirus (XMRV) in Dutch prostate cancer patients. Prostate. 2011;71:415–420. [PubMed]
9. Hohn O, Krause H, Barbarotto P, Niederstadt L, Beimforde N, Denner J, Miller K, Kurth R, Bannert N. Lack of evidence for xenotropic murine leukemia virus-related virus(XMRV) in German prostate cancer patients. Retrovirology. 2009;6:92. [PMC free article] [PubMed]
10. Lombardi VC, Ruscetti FW, Das Gupta J, Pfost MA, Hagen KS, Peterson DL, Ruscetti SK, Bagni RK, Petrow-Sadowski C, Gold B, et al. Detection of an infectious retrovirus, XMRV, in blood cells of patients with chronic fatigue syndrome. Science. 2009;326:585–589. [PubMed]
11. Henrich TJ, Li JZ, Felsenstein D, Kotton CN, Plenge RM, Pereyra F, Marty FM, Lin NH, Grazioso P, Crochiere DM, et al. Xenotropic murine leukemia virus-related virus prevalence in patients with chronic fatigue syndrome or chronic immunomodulatory conditions. J. Infect. Dis. 2010;202:1478–1481. [PMC free article] [PubMed]
12. Erlwein O, Kaye S, McClure MO, Weber J, Wills G, Collier D, Wessely S, Cleare A. Failure to detect the novel retrovirus XMRV in chronic fatigue syndrome. PLoS ONE. 2010;5:e8519. [PMC free article] [PubMed]
13. Groom HC, Boucherit VC, Makinson K, Randal E, Baptista S, Hagan S, Gow JW, Mattes FM, Breuer J, Kerr JR, et al. Absence of xenotropic murine leukaemia virus-related virus in UK patients with chronic fatigue syndrome. Retrovirology. 2010;7:10. [PMC free article] [PubMed]
14. Switzer WM, Jia H, Hohn O, Zheng H, Tang S, Shankar A, Bannert N, Simmons G, Hendry RM, Falkenberg VR, et al. Absence of evidence of xenotropic murine leukemia virus-related virus infection in persons with chronic fatigue syndrome and healthy controls in the United States. Retrovirology. 2010;7:57. [PMC free article] [PubMed]
15. Satterfield BC, Garcia RA, Jia H, Tang S, Zheng H, Switzer WM. Serologic and PCR testing of persons with chronic fatigue syndrome in the United States shows no association with xenotropic or polytropic murine leukemia virus-related viruses. Retrovirology. 2011;8:12. [PMC free article] [PubMed]
16. Menendez-Arias L. Evidence and controversies on the role of XMRV in prostate cancer and chronic fatigue syndrome. Rev. Med. Virol. 21:3–17. [PubMed]
17. Hue S, Gray ER, Gall A, Katzourakis A, Tan CP, Houldcroft CJ, McLaren S, Pillay D, Futreal A, Garson JA, et al. Disease-associated XMRV sequences are consistent with laboratory contamination. Retrovirology. 2010;7:111. [PMC free article] [PubMed]
18. Paprotka T, Delviks-Frankenberry KA, Cingoz O, Martinez A, Kung HJ, Tepper CG, Hu WS, Fivash MJ, Jr, Coffin JM, Pathak VK. Recombinant origin of the retrovirus XMRV. Science. 2011;333:97–101. [PMC free article] [PubMed]
19. Singh K, Kaushik N, Jin J, Madhusudanan M, Modak MJ. Role of Q190 of MuLV RT in ddNTP resistance and fidelity of DNA synthesis: a molecular model of interactions with substrates. Protein Eng. 2000;13:635–643. [PubMed]
20. Telesnitsky A, Goff SP. Two defective forms of reverse transcriptase can complement to restore retroviral infectivity. EMBO J. 1993;12:4433–4438. [PubMed]
21. Georgiadis MM, Jessen SM, Ogata CM, Telesnitsky A, Goff SP, Hendrickson WA. Mechanistic implications from the structure of a catalytic fragment of Moloney murine leukemia virus reverse transcriptase. Structure. 1995;3:879–892. [PubMed]
22. Das D, Georgiadis MM. The crystal structure of the monomeric reverse transcriptase from Moloney murine leukemia virus. Structure. 2004;12:819–829. [PubMed]
23. Chowdhury K, Kaushik N, Pandey VN, Modak MJ. Elucidation of the role of Arg 110 of murine leukemia virus reverse transcriptase in the catalytic mechanism: biochemical characterization of its mutant enzymes. Biochemistry. 1996;35:16610–16620. [PubMed]
24. Kaushik N, Chowdhury K, Pandey VN, Modak MJ. Valine of the YVDD motif of moloney murine leukemia virus reverse transcriptase: role in the fidelity of DNA synthesis. Biochemistry. 2000;39:5155–5165. [PubMed]
25. Jacobo-Molina A, Ding J, Nanni RG, Clark AD, Jr, Lu X, Tantillo C, Williams RL, Kamer G, Ferris AL, Clark P, et al. Crystal structure of human immunodeficiency virus type 1 reverse transcriptase complexed with double-stranded DNA at 3.0 A resolution shows bent DNA. Proc. Natl Acad. Sci. USA. 1993;90:6320–6324. [PubMed]
26. Kohlstaedt LA, Wang J, Friedman JM, Rice PA, Steitz TA. Crystal structure at 3.5 A resolution of HIV-1 reverse transcriptase complexed with an inhibitor. Science. 1992;256:1783–1790. [PubMed]
27. Sarafianos SG, Marchand B, Das K, Himmel DM, Parniak MA, Hughes SH, Arnold E. Structure and function of HIV-1 reverse transcriptase: molecular mechanisms of polymerization and inhibition. J. Mol. Biol. 2009;385:693–713. [PMC free article] [PubMed]
28. Singh K, Marchand B, Kirby KA, Michailidis E, Sarafianos SG. Structural aspects of drug resistance and inhibition of HIV-1 reverse transcriptase. Viruses. 2010;2:606–638. [PMC free article] [PubMed]
29. Schuckmann MM, Marchand B, Hachiya A, Kodama EN, Kirby KA, Singh K, Sarafianos SG. The N348I mutation at the connection subdomain of HIV-1 reverse transcriptase decreases binding to nevirapine. J. Biol. Chem. 2010;285:38700–38709. [PMC free article] [PubMed]
30. Telesnitsky A, Goff SP. RNase H domain mutations affect the interaction between Moloney murine leukemia virus reverse transcriptase and its primer-template. Proc. Natl Acad. Sci. USA. 1993;90:1276–1280. [PubMed]
31. Bauman JD, Das K, Ho WC, Baweja M, Himmel DM, Clark AD, Jr, Oren DA, Boyer PL, Hughes SH, Shatkin AJ, et al. Crystal engineering of HIV-1 reverse transcriptase for structure-based drug design. Nucleic Acids Res. 2008;36:5083–5092. [PMC free article] [PubMed]
32. Michailidis E, Marchand B, Kodama EN, Singh K, Matsuoka M, Kirby KA, Ryan EM, Sawani AM, Nagy E, Ashida N, et al. Mechanism of inhibition of HIV-1 reverse transcriptase by 4′-Ethynyl-2-fluoro-2′-deoxyadenosine triphosphate, a translocation-defective reverse transcriptase inhibitor. J. Biol. Chem. 2009;284:35681–35691. [PMC free article] [PubMed]
33. Meyer PR, Matsuura SE, So AG, Scott WA. Unblocking of chain-terminated primer by HIV-1 reverse transcriptase through a nucleotide-dependent mechanism. Proc. Natl Acad. Sci. USA. 1998;95:13471–13476. [PubMed]
34. Halvas EK, Svarovskaia ES, Pathak VK. Development of an in vivo assay to identify structural determinants in murine leukemia virus reverse transcriptase important for fidelity. J Virol. 2000;74:312–319. [PMC free article] [PubMed]
35. Patel SS, Wong I, Johnson KA. Pre-steady-state kinetic analysis of processive DNA replication including complete characterization of an exonuclease-deficient mutant. Biochemistry. 1991;30:511–525. [PubMed]
36. Sarafianos SG, Clark AD, Jr, Das K, Tuske S, Birktoft JJ, Ilankumaran P, Ramesha AR, Sayer JM, Jerina DM, Boyer PL, et al. Structures of HIV-1 reverse transcriptase with pre- and post-translocation AZTMP-terminated DNA. EMBO J. 2002;21:6614–6624. [PubMed]
37. Tuske S, Sarafianos SG, Clark AD, Jr, Ding J, Naeger LK, White KL, Miller MD, Gibbs CS, Boyer PL, Clark P, et al. Structures of HIV-1 RT-DNA complexes before and after incorporation of the anti-AIDS drug tenofovir. Nat. Struct. Mol. Biol. 2004;11:469–474. [PubMed]
38. Sarafianos SG, Pandey VN, Kaushik N, Modak MJ. Site-directed mutagenesis of arginine 72 of HIV-1 reverse transcriptase. Catalytic role and inhibitor sensitivity. J. Biol. Chem. 1995;270:19729–19735. [PubMed]
39. Hachiya A, Kodama EN, Schuckmann MM, Kirby KA, Michailidis E, Sakagami Y, Oka S, Singh K, Sarafianos SG. K70Q adds high-level tenofovir resistance to “Q151M complex” HIV reverse transcriptase through the enhanced discrimination mechanism. PLoS One. 2011;6:e16242. [PMC free article] [PubMed]
40. Sarafianos SG, Das K, Ding J, Boyer PL, Hughes SH, Arnold E. Touching the heart of HIV-1 drug resistance: the fingers close down on the dNTP at the polymerase active site. Chem. Biol. 1999;6:R137–R146. [PubMed]
41. Boyer PL, Sarafianos SG, Clark PK, Arnold E, Hughes SH. Why do HIV-1 and HIV-2 use different pathways to develop AZT resistance? PLoS Pathog. 2006;2:e10. [PMC free article] [PubMed]
42. Powell MD, Ghosh M, Jacques PS, Howard KJ, Le Grice SF, Levin JG. Alanine-scanning mutations in the “primer grip” of p66 HIV-1 reverse transcriptase result in selective loss of RNA priming activity. J. Biol. Chem. 1997;272:13262–13269. [PubMed]
43. Sarafianos SG, Das K, Clark AD, Jr, Ding J, Boyer PL, Hughes SH, Arnold E. Lamivudine (3TC) resistance in HIV-1 reverse transcriptase involves steric hindrance with beta-branched amino acids. Proc. Natl Acad. Sci. USA. 1999;96:10027–10032. [PubMed]
44. Boucher CA, Cammack N, Schipper P, Schuurman R, Rouse P, Wainberg MA, Cameron JM. High-level resistance to (-) enantiomeric 2′-deoxy-3′-thiacytidine in vitro is due to one amino acid substitution in the catalytic site of human immunodeficiency virus type 1 reverse transcriptase. Antimicrob. Agents Chemother. 1993;37:2231–2234. [PMC free article] [PubMed]
45. Tisdale M, Kemp SD, Parry NR, Larder BA. Rapid in vitro selection of human immunodeficiency virus type 1 resistant to 3′-thiacytidine inhibitors due to a mutation in the YMDD region of reverse transcriptase. Proc. Natl Acad. Sci. USA. 1993;90:5653–5656. [PubMed]
46. Menendez-Arias L. Molecular basis of human immunodeficiency virus drug resistance: an update. Antiviral Res. 2010;85:210–231. [PubMed]
47. Sarafianos SG, Das K, Hughes SH, Arnold E. Taking aim at a moving target: designing drugs to inhibit drug-resistant HIV-1 reverse transcriptases. Curr. Opin. Struct. Biol. 2004;14:716–730. [PubMed]
48. Menendez-Arias L. Molecular basis of human immunodeficiency virus drug resistance: an update. Antiviral Res. 2010;85:210–231. [PubMed]
49. Menendez-Arias L, Berkhout B. Retroviral reverse transcription. Virus Res. 2008;134:1–3. [PubMed]
50. Shi Q, Singh K, Srivastava A, Kaushik N, Modak MJ. Lysine 152 of MuLV reverse transcriptase is required for the integrity of the active site. Biochemistry. 2002;41:14831–14842. [PubMed]
51. Johnson KA. Conformational coupling in DNA polymerase fidelity. Annu. Rev. Biochem. 1993;62:685–713. [PubMed]
52. Rezende LF, Prasad VR. Nucleoside-analog resistance mutations in HIV-1 reverse transcriptase and their influence on polymerase fidelity and viral mutation rates. Int. J. Biochem. Cell Biol. 2004;36:1716–1734. [PubMed]
53. Paprotka T, Venkatachari NJ, Chaipan C, Burdick R, Delviks-Frankenberry KA, Hu WS, Pathak VK. Inhibition of xenotropic murine leukemia virus-related virus by APOBEC3 proteins and antiviral drugs. J. Virol. 84:5719–5729. [PMC free article] [PubMed]
54. Sakuma R, Sakuma T, Ohmine S, Silverman RH, Ikeda Y. Xenotropic murine leukemia virus-related virus is susceptible to AZT. Virology. 2010;397:1–6. [PMC free article] [PubMed]
55. Singh IR, Gorzynski JE, Drobysheva D, Bassit L, Schinazi RF. Raltegravir is a potent inhibitor of XMRV, a virus implicated in prostate cancer and chronic fatigue syndrome. PLoS One. 2010;5:e9948. [PMC free article] [PubMed]
56. Smith RA, Gottlieb GS, Miller AD. Susceptibility of the human retrovirus XMRV to antiretroviral inhibitors. Retrovirology. 2010;7:70. [PMC free article] [PubMed]
57. Kawamoto A, Kodama E, Sarafianos SG, Sakagami Y, Kohgo S, Kitano K, Ashida N, Iwai Y, Hayakawa H, Nakata H, et al. 2′-deoxy-4′-C-ethynyl-2-halo-adenosines active against drug-resistant human immunodeficiency virus type 1 variants. Int. J. Biochem. Cell Biol. 2008;40:2410–2420. [PubMed]
58. Kodama EI, Kohgo S, Kitano K, Machida H, Gatanaga H, Shigeta S, Matsuoka M, Ohrui H, Mitsuya H. 4′-Ethynyl nucleoside analogs: potent inhibitors of multidrug-resistant human immunodeficiency virus variants in vitro. Antimicrob. Agents Chemother. 2001;45:1539–1546. [PMC free article] [PubMed]
59. Kissel JD, Held DM, Hardy RW, Burke DH. Single-stranded DNA aptamer RT1t49 inhibits RT polymerase and RNase H functions of HIV type 1, HIV type 2, and SIVCPZ RTs. AIDS Res. Hum. Retroviruses. 2007;23:699–708. [PubMed]
60. Chen H, Gold L. Selection of high-affinity RNA ligands to reverse transcriptase: inhibition of cDNA synthesis and RNase H activity. Biochemistry. 1994;33:8746–8756. [PubMed]
61. Joshi PJ, Fisher TS, Prasad VR. Anti-HIV inhibitors based on nucleic acids: emergence of aptamers as potent antivirals. Curr. Drug Targets Infect. Disord. 2003;3:383–400. [PubMed]
62. DeStefano JJ, Nair GR. Novel aptamer inhibitors of human immunodeficiency virus reverse transcriptase. Oligonucleotides. 2008;18:133–144. [PMC free article] [PubMed]
63. DeStefano JJ, Cristofaro JV. Selection of primer-template sequences that bind human immunodeficiency virus reverse transcriptase with high affinity. Nucleic Acids Res. 2006;34:130–139. [PMC free article] [PubMed]
64. Arion D, Kaushik N, McCormick S, Borkow G, Parniak MA. Phenotypic mechanism of HIV-1 resistance to 3′-azido-3′-deoxythymidine (AZT): increased polymerization processivity and enhanced sensitivity to pyrophosphate of the mutant viral reverse transcriptase. Biochemistry. 1998;37:15908–15917. [PubMed]
65. Sarafianos SG, Hughes SH, Arnold E. Designing anti-AIDS drugs targeting the major mechanism of HIV-1 RT resistance to nucleoside analog drugs. Int. J. Biochem. Cell Biol. 2004;36:1706–1715. [PubMed]
66. Shafer RW, Kozal MJ, Winters MA, Iversen AK, Katzenstein DA, Ragni MV, Meyer WA, 3rd, Gupta P, Rasheed S, Coombs R, et al. Combination therapy with zidovudine and didanosine selects for drug-resistant human immunodeficiency virus type 1 strains with unique patterns of pol gene mutations. J. Infect. Dis. 1994;169:722–729. [PubMed]
67. Shirasaka T, Kavlick MF, Ueno T, Gao WY, Kojima E, Alcaide ML, Chokekijchai S, Roy BM, Arnold E, Yarchoan R, et al. Emergence of human immunodeficiency virus type 1 variants with resistance to multiple dideoxynucleosides in patients receiving therapy with dideoxynucleosides. Proc. Natl Acad. Sci. USA. 1995;92:2398–2402. [PubMed]
68. Iversen AK, Shafer RW, Wehrly K, Winters MA, Mullins JI, Chesebro B, Merigan TC. Multidrug-resistant human immunodeficiency virus type 1 strains resulting from combination antiretroviral therapy. J. Virol. 1996;70:1086–1090. [PMC free article] [PubMed]
69. Winters MA, Shafer RW, Jellinger RA, Mamtora G, Gingeras T, Merigan TC. Human immunodeficiency virus type 1 reverse transcriptase genotype and drug susceptibility changes in infected individuals receiving dideoxyinosine monotherapy for 1 to 2 years. Antimicrob. Agents Chemother. 1997;41:757–762. [PMC free article] [PubMed]
70. Gu Z, Gao Q, Fang H, Salomon H, Parniak MA, Goldberg E, Cameron J, Wainberg MA. Identification of a mutation at codon 65 in the IKKK motif of reverse transcriptase that encodes human immunodeficiency virus resistance to 2′,3′-dideoxycytidine and 2′,3′-dideoxy-3′-thiacytidine. Antimicrob. Agents Chemother. 1994;38:275–281. [PMC free article] [PubMed]
71. Knox K, Carrigan D, Simmons G, Teque F, Zhou Y, Hackett J, Jr, Qiu X, Luk KC, Schochetman G, Knox A, et al. No evidence of murine-like gammaretroviruses in CFS patients previously identified as XMRV-infected. Science. 2011 [PubMed]
72. Oakes B, Tai AK, Cingoz O, Henefield MH, Levine S, Coffin JM, Huber BT. Contamination of human DNA samples with mouse DNA can lead to false detection of XMRV-like sequences. Retrovirology. 2010;7:109. [PMC free article] [PubMed]
73. Robinson MJ, Erlwein OW, Kaye S, Weber J, Cingoz O, Patel A, Walker MM, Kim WJ, Uiprasertkul M, Coffin JM, et al. Mouse DNA contamination in human tissue tested for XMRV. Retrovirology. 2010;7:108. [PMC free article] [PubMed]
74. Sato E, Furuta RA, Miyazawa T. An endogenous murine leukemia viral genome contaminant in a commercial RT-PCR kit is amplified using standard primers for XMRV. Retrovirology. 2010;7:110. [PMC free article] [PubMed]
75. Coffin JM, Stoye JP. Virology. A new virus for old diseases? Science. 2009;326:530–531. [PMC free article] [PubMed]
76. Huang H, Chopra R, Verdine GL, Harrison SC. Structure of a covalently trapped catalytic complex of HIV-1 reverse transcriptase: implications for drug resistance. Science. 1998;282:1669–1675. [PubMed]
77. Sarafianos SG, Das K, Tantillo C, Clark AD, Jr, Ding J, Whitcomb JM, Boyer PL, Hughes SH, Arnold E. Crystal structure of HIV-1 reverse transcriptase in complex with a polypurine tract RNA:DNA. EMBO J. 2001;20:1449–1461. [PubMed]
78. Kirby KA, Singh K, Michailidis E, Marchand B, Kodama EN, Ashida N, Mitsuya H, Parniak MA, Sarafianos SG. The sugar ring conformation of 4′-ethynyl-2-fluoro-2′-deoxyadenosine and its recognition by the polymerase active site of hiv reverse transcriptase. Cell Mol. Biol. 57:40–46. [PMC free article] [PubMed]
79. Gotte M, Arion D, Parniak MA, Wainberg MA. The M184V mutation in the reverse transcriptase of human immunodeficiency virus type 1 impairs rescue of chain-terminated DNA synthesis. J. Virol. 2000;74:3579–3585. [PMC free article] [PubMed]
80. Wainberg MA, Drosopoulos WC, Salomon H, Hsu M, Borkow G, Parniak M, Gu Z, Song Q, Manne J, Islam S, et al. Enhanced fidelity of 3TC-selected mutant HIV-1 reverse transcriptase. Science. 1996;271:1282–1285. [PubMed]
81. Pandey VN, Kaushik N, Rege N, Sarafianos SG, Yadav PN, Modak MJ. Role of methionine 184 of human immunodeficiency virus type-1 reverse transcriptase in the polymerase function and fidelity of DNA synthesis. Biochemistry. 1996;35:2168–2179. [PubMed]
82. Das K, Bandwar RP, White KL, Feng JY, Sarafianos SG, Tuske S, Tu X, Clark AD, Jr, Boyer PL, Hou X, et al. Structural basis for the role of the K65R mutation in HIV-1 reverse transcriptase polymerization, excision antagonism, and tenofovir resistance. J. Biol. Chem. 2009;284:35092–35100. [PMC free article] [PubMed]
83. Meyer PR, Matsuura SE, Mian AM, So AG, Scott WA. A mechanism of AZT resistance: an increase in nucleotide-dependent primer unblocking by mutant HIV-1 reverse transcriptase. Mol. Cell. 1999;4:35–43. [PubMed]
84. Ueno T, Shirasaka T, Mitsuya H. Enzymatic characterization of human immunodeficiency virus type 1 reverse transcriptase resistant to multiple 2′,3′-dideoxynucleoside 5′-triphosphates. J. Biol. Chem. 1995;270:23605–23611. [PubMed]
85. Rhee SY, Gonzales MJ, Kantor R, Betts BJ, Ravela J, Shafer RW. Human immunodeficiency virus reverse transcriptase and protease sequence database. Nucleic Acids Res. 2003;31:298–303. [PMC free article] [PubMed]

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