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Integrase (IN), the HIV-1 enzyme responsible for the integration of the viral genome into the chromosomes of infected cells, is the target of the recently approved antiviral raltegravir (RAL). Despite this drug's activity against viruses resistant to other antiretrovirals, failures of raltegravir therapy were observed, in association with the emergence of resistance due to mutations in the integrase coding region. Two pathways involving primary mutations on residues N155 and Q148 have been characterized. It was suggested that mutations at residue Y143 might constitute a third primary pathway for resistance. The aims of this study were to investigate the susceptibility of HIV-1 Y143R/C mutants to raltegravir and to determine the effects of these mutations on the IN-mediated reactions. Our observations demonstrate that Y143R/C mutants are strongly impaired for both of these activities in vitro. However, Y143R/C activity can be kinetically restored, thereby reproducing the effect of the secondary G140S mutation that rescues the defect associated with the Q148R/H mutants. A molecular modeling study confirmed that Y143R/C mutations play a role similar to that determined for Q148R/H mutations. In the viral replicative context, this defect leads to a partial block of integration responsible for a weak replicative capacity. Nevertheless, the Y143 mutant presented a high level of resistance to raltegravir. Furthermore, the 50% effective concentration (EC50) determined for Y143R/C mutants was significantly higher than that obtained with G140S/Q148R mutants. Altogether our results not only show that the mutation at position Y143 is one of the mechanisms conferring resistance to RAL but also explain the delayed emergence of this mutation.
Integration of the HIV-1 viral genome into that of the host cell is a key step of viral replication, since the integrated genome, the provirus, is the genomic DNA form that allows the expression of the viral gene and the subsequent formation of progeny viruses. This step is catalyzed by the viral protein integrase (IN), encoded by the pol gene and responsible for the integration of double-stranded DNA generated by reverse transcription of the RNA genome (for a review, see reference 11). To complete this process, IN catalyzes two successive reactions. The first is the 3′-processing (3′-P) reaction, during which terminal GpT dinucleotides are cleaved from each 3′ end of long terminal repeats (LTR), producing CpA 3′-hydroxyl ends. This reaction takes place within a nucleoprotein complex known as the preintegration complex (PIC), which translocates through the nuclear pore into the nucleus, where the second reaction or strand transfer (ST) occurs. During this second step, IN transfers both newly exposed 3′ extremities of viral DNA into the target genome in a concerted manner by one-step transesterification reactions, resulting in full-site integration (2). Due to this key function in the viral cycle, IN is an attractive target for antiretroviral drugs (ARVs). IN inhibitors constitute a new class of antiretroviral agents blocking HIV-1 activity (19). To date, only integrase strand transfer inhibitors (INSTIs) have shown potent antiviral activity in vivo. Raltegravir (RAL), which was approved in October 2007, is the first drug of the INSTI group to reach therapeutic use after having demonstrated a rapid, potent, and sustained antiretroviral effect in patients with advanced HIV-1 infection (17, 30). RAL is well tolerated and, due to its mechanism of action, is active against viruses resistant to other classes of antiretroviral drugs, such as nucleoside reverse transcriptase inhibitors (NRTI), nonnucleoside reverse transcriptase inhibitors (NNRTI), protease inhibitors (PI), and entry inhibitors (17). However, as observed for other ARVs, specific resistance mutations located in the IN coding region of replicating viruses, associated with a reduced susceptibility to RAL, have been identified in patients failing to respond to treatment with RAL (28). Although resistance to RAL in vivo has been linked to 14 mutations (35), each to a varying degree, virological failure is clearly linked to two main independent genetic pathways involving primary mutations on residues N155 (N155H) and Q148 (Q148K/R/H). Secondary mutations that increase the fitness of the resistant viruses have been identified in both pathways. In particular, the secondary G140S mutation, observed in tandem with the Q148H mutation, rescues a replicative defect due to the presence of the primary mutation Q148H (12). Several other mutations at other positions have also been described, such as L68I/V, D232N, G163K/R, E138A, E157Q, L74M, and V151I (For a review, see reference 5). Nevertheless, with the hindsight of several months of treatment, only a few mutations that might constitute another pathway of resistance have emerged. These mutations involve residues E92, E157, and Y143. While the first two mutations are subject to controversy as primary mutations for RAL resistance (35), the third one seems to significantly decrease susceptibility to the inhibitor. Interestingly, Y143R/C/H occurs less frequently and appears to occur later than the first two pathways (9). We identified Y143R/C mutations in the IN mutation patterns associated with patients that failed to respond to RAL treatment. We investigated the impact of the substitution at the Y143 residue in their sensitivity to RAL, the overall catalytic activity of the recombinant protein, and the replication efficiency of viruses with the Y143R/C mutation. Y143R/C mutations conferred a high resistance to RAL in vitro as well as in vivo. The overall catalytic activity of IN in vitro and in vivo was highly impaired, and this occurred mainly at the integration step.
Four patients who failed to respond to RAL treatment were retrospectively studied. The genotypic analysis of IN resistance and the follow-up of HIV-1 viral load were carried out following French national guidelines (39). All patients received at least one NRTI with one boosted PI with or without enfuvirtide in their optimized regimen. The optimized regimen associated with RAL was selected according to previous antiretroviral exposure and genotypic resistance testing interpreted using the last French ANRS algorithm (www.hivfrenchresistance.org). RNA was extracted from 500 μl of plasma, and a fragment encompassing the entire IN coding region of the pol gene was amplified and sequenced as described previously (27).
MT4 cells were cultured in RPMI 1640 containing 10% fetal calf serum. 293T and HeLa-P4 cells were cultured in Dulbecco's modified Eagle medium supplemented with 10% fetal calf serum, 100 U penicillin/ml (Invitrogen), and 100 μg streptomycin/ml (Invitrogen). HIV-1 IN (Y143R/C) mutants were generated by site-directed mutagenesis as previously described (12). Briefly, the fragment encoding IN of the replication-competent pNL4.3 virus was digested with AgeI and EcoRI and inserted into the Bluescript vector, and IN mutants were obtained by mutagenesis (QuikChange mutagenesis kit; Stratagene). The constructs were checked by sequencing, and the fragment was then inserted into pNL4.3. HIV-1 virus stocks of all mutants were prepared by transfecting 293T cells. Transfection assays were carried out by the calcium phosphate method. Forty-eight hours posttransfection, viral supernatants were filtered through a 0.45-μm-pore-size filter and immediately frozen at −80°C. The HIV-1 p24gag antigen content in viral supernatants was determined by enzyme-linked immunosorbent assay (Perkin-Elmer Life Sciences).
The viral titer was determined in HeLa-P4 cells. HeLa-P4 cells are HeLa CD4 LTR-LacZ cells in which lacZ expression is induced by the HIV transactivator protein Tat, allowing the precise quantification of HIV-1 infectivity. Cells were infected in triplicate in 96-well plates with wild-type (WT) or Y143R/C viruses (equivalent of 3 ng of p24gag antigen or with higher viral concentrations, as described in Results). The viral inoculum was left for 48 h. The viral titer was determined 48 h after infection by quantifying β-galactosidase activity in HeLa-P4 lysates in a colorimetric assay based on the cleavage of chlorophenol red-β-d-galactopyranoside (CPRG) by β-galactosidase as described previously (12). For 50% effective concentration (EC50) determination, cells were infected with viruses and grown in the presence of increasing concentrations of RAL. The EC50 was defined as the drug concentration resulting in β-galactosidase levels that are 50% lower than those in untreated infected cells. Cell survival was also estimated with a standard MTT (3-[4,5-dimethylthiazol-2-yl]-2,5-diphenyltetrazolium bromide) assay (12).
MT4 cells were concentrated to 2.106/ml and infected with viruses (50 ng of p24 antigen par 106 cells). Viral inoculum remains throughout the course of the experiment. If required, cells were treated with 100 nM RAL inhibitor (Merck & Co.) 1 h before infection. RAL was maintained during the course of the experiment. At various time points after infection, 1 million to 2 million cells were harvested and dry cell pellets were frozen at −80°C until use.
Total cell DNA was extracted with a QIAamp blood DNA minikit (Qiagen, Courtaboeuf, France). Quantifications of total HIV-1 DNA, 2-LTR circles, and integrated HIV-1 DNA were performed by real-time PCR on a LightCycler instrument (Roche Diagnostics) using the fit point method provided by the LightCycler quantification software, version 3.5 (Roche Diagnostics) as previously described (3). Cell equivalents were calculated based on amplification of the β-globin gene with commercially available materials (control kit DNA; Roche Diagnostics). 2-LTR circles and total and integrated HIV-1 DNA levels were determined and expressed as copy numbers per 106 cells.
The Y143R and Y143C mutations were obtained by site-directed mutagenesis of pET-15b containing the WT sequence. The wild-type and mutant HIV-1 INs used for DNA-binding and 3′-P assays were produced in Escherichia coli BL21(RIL) and purified under nondenaturing conditions as previously described (25). Activity assays were carried out as described in reference 25. Oligonucleotides (ODNs) were radiolabeled with T4 polynucleotide kinase (Biolabs) and [γ-32P]ATP (3,000 Ci/mmol) (Amersham) and purified on a Sephadex G-10 column (GE Healthcare). Double-stranded ODNs were obtained by mixing equimolar amounts of complementary strands in the presence of 100 mM NaCl. 3′-P and strand transfer assays were carried out at 37°C in a buffer containing 10 mM HEPES (pH 7.2)-1 mM dithiothreitol (DTT) (7.5 mM) in the presence of 3.25 nM and 12.5 nM DNA substrates, respectively. Products were separated by electrophoresis in denaturing 18% acrylamide/urea gels. Gels were analyzed with a Molecular Dynamics Storm phosphorimager (GE Healthcare) and quantified using the Image Quant 4.1 software program. Fifty percent inhibitory concentration (IC50) calculation and t tests were performed using Prism 5.0 software (GraphPad Software, San Diego, CA).
Steady-state fluorescence anisotropy values (r) were recorded on a Beacon 2000 instrument (Panvera, Madison, WI) in a cell maintained at 25°C or 37°C under thermostatic control. The principle underlying the anisotropy-based assay has been published elsewhere for DNA binding (1, 36) and 3′-P (10, 18), respectively. Briefly, IN binding to fluorescein-labeled DNA (double-stranded 21-mer oligonucleotide [ODN] mimicking the U5 viral DNA end) increases the r value, making it possible to calculate the fractional saturation function: [DNA*IN]/[DNA]0. The DNA binding step was recorded at 25°C, using ODNs fluorescein labeled at the 3′-terminal GT nucleotide. The percentage of complexes was then calculated as
where rfree and rsat are the anisotropy values characterizing the free and bound oligonucleotides, respectively. Following DNA binding, the sample was then incubated at 37°C to record 3′-P activity. As the fluorophore is linked to the released dinucleotide, 3′-P activity significantly decreases the r value with respect to that for nonprocessed DNA. Activity can be calculated in fixed-time experiments by disrupting all the IN*DNA complexes with SDS (0.25%, final). The fraction of dinucleotides released is given by
where rNP and rdinu are the anisotropy values for pure solutions of nonprocessed double-stranded ODN and dinucleotide, respectively. The formation of IN/DNA complexes and the subsequent 3′-processing reaction were carried out by incubating fluorescein-labeled ODNs (4 nM) with IN in 20 mM HEPES (pH 7.2), 1 mM dithiothreitol, 30 mM NaCl, and 10 mM MgCl2. Standard 3′-P and ST activity tests based on gel electrophoresis were performed as previously described (25).
All calculations were carried out on a PC running RedHat Enterprise Linux 5 software. Modeling, analysis, and graphic generation were performed with SYBYL (version 8.0) software (Tripos Inc., 2008).
Three-dimensional (3D) models of raltegravir resistance mutants were built by individual amino acid substitutions, using previously generated models of the catalytic core of wild-type IN, and were optimized as previously described (31). The stereochemical quality of the models was assessed with the ProTable Procheck software program (24), which showed that more than 97% of the nonglycine residues in all models had dihedral angles in the most favored and allowed regions of the Ramachandran plot, consistent with high model quality.
the overall hydrophobicity (lipophilicity) of a molecule was calculated/predicted by its partition coefficient (logP), represented as fragmental increments, fi:
On the basis of the atomic partial lipophilic values, fi, a distance-dependent function has been defined for the lipophilicity potential (LP) of proteins (20):
where di = distance of a certain point in space from atom i with
LPHM was implemented in MOLCAD with two of fi sets based on Crippen atomic partial lipophilicities (16) used for characterizing hydrophobicity of the wild type and the mutants.
Hydrogen-bonding (HB) sites on the molecular surface were localized by the original MOLCAD method.
Sequence analysis of clinical isolates obtained during RAL treatment has led to the identification of various mutations at specific positions only observed in the IN coding region (28). Indeed, two main genetic pathways have been characterized involving residues N155 (mutation N155H) and Q148 (mutations Q148H/K/R) (23). Recent data suggest the existence of a third pathway, involving the Y143 residue, correlated with virological resistance to RAL (35). Here, four HIV-1-infected patients failing to respond to 400 mg RAL, administered twice daily, and for whom a mutation in position 143 was found in the viral IN sequence at the time of failure were studied retrospectively. These patients with isolates with the Y143 mutation represented an approximate ratio of 4% among patients failing to respond to RAL, while clinical isolates displaying the N155H or G140S/Q148R/H mutations represented, respectively, 61 and 35% of the resistant profiles observed in our clinical center.
Isolates from patients 1 to 3 had a Y143R mutation, and virus from patient 4 had two mutations, Y143Y/C and N155N/H (Tables (Tables11 and and2).2). Their baseline characteristics are given in Table Table1.1. At the start of RAL therapy, the median CD4 count was 164.5 cells/mm3 (range, 3 to 383 cells/mm3) and the median plasma HIV-1 RNA level was 4.7 log10 copies/ml (range, 3.7 to 4.8 log10 copies/ml). All patients harbored highly mutated viruses with resistance to NRTI, NNRTI, and PI; their genotypic sensitivity score (number of active ARV in the background regimen associated with RAL) was between 1 and 2. The complete nucleotide sequences of IN were determined for clinical isolates from each patient at various time points before and during RAL treatment. We found no IN resistance mutations in the bulk of the PCR products amplified from plasma HIV-1 RNA before the introduction of RAL for any of the patients. At the time of failure (median, 41 weeks; range, 16 to 48 weeks), the only change in sequence analyzed was a mutation in position 143 in the IN sequence. These findings confirmed that a specific mutation at Y143 was correlated with a failure to respond to RAL treatment. Mutation of this residue may constitute a third mechanism of resistance against RAL. Thus, we investigated Y143 and its role in resistance to RAL by producing viruses that have the Y143R or Y143C mutation.
To determine whether the Y143R and Y43C mutations conferred resistance to RAL, these mutations were introduced into a WT pNL4.3 viral backbone. First, mutated viruses were produced following transfection of these genomic clones into 293T cells. Levels of p24 similar to those obtained with the WT virus were observed for each mutant, showing that none of these mutations significantly impaired late viral replication steps (Fig. (Fig.1A).1A). Then, the impact of these mutations on the early replication steps, i.e., from entry to integration, was determined at 48 h postinfection on HeLa-P4 reporter cells (Fig. (Fig.1B).1B). Cells were infected with similar amounts of viral particles (3 ng of p24), and β-galactosidase levels were evaluated 48 h postinfection. Under these conditions, about 40% of the cells were infected with the wild-type virus. To rule out a possible bias due to the colorimetric measurement method, the background corresponding to the CPRG signal obtained for an abortive viral infection in the presence of 50 μM zidovudine (ZDV) was systematically determined. Both the Y143R and Y143C mutations significantly decreased viral infectivity (15% and 10% of the WT level for Y143C and Y143R, respectively), suggesting that mutant IN activity was affected (Fig. (Fig.1B).1B). We then quantified RAL resistance in Y143R/C mutants following infection with higher virus concentrations (30 ng p24 antigen). To avoid a bias due to a possible correlation between the amount of infectious viruses present and the effectiveness of RAL, we determined first whether a change in the infectious virus concentration affects the apparent EC50. In agreement with previous studies (33), no significant change was observed, with an EC50 equal to 7 nM after various infectious virus concentrations to more than 1 order of magnitude, thereby confirming the high potency of RAL as an inhibitor of HIV-1 integration (Fig. (Fig.1C).1C). No cytotoxicity due either to RAL or to viral replication was observed in this model by a standard MTT assay up to 72 h postinfection as already described (12). For both Y143R and Y143C viruses, the EC50 was not reached within the range of concentrations tested (EC50 > 100 nM), thus demonstrating a strong resistance to RAL (Fig. (Fig.1C1C).
Although Y143R/C viruses are highly resistant to RAL, their infectivity was deeply altered, suggesting a defect during the early replicative steps. To determine which replicative step was impaired, MT4 cells were infected with either WT or resistant viruses in the presence or absence of 100 nM RAL, and the various viral DNA species were quantified by real-time PCR 24 h postinfection. The amounts of total viral DNA were similar for all viruses, indicating that the mutations did not affect the viral entry and reverse transcription steps (data not shown). Moreover, the amount of viral DNA was not decreased by the addition of 100 nM RAL, confirming that this ARV does not affect the reverse transcription step.
Efficiency of viral integration was evaluated by Alu-LTR real-time nested PCR amplification of integrated proviruses (3). For the wild-type virus, 10% of the total DNA was integrated, in agreement with previous reports (3). In contrast, for the Y143R and Y143C mutants, the amount of integrated DNA was equal to 2.1% and 1.1%, respectively. Thus, the integration for the Y143 mutants was much less than that for WT viruses, thereby indicating that a block in integration may account for the replication defect (Fig. (Fig.2A).2A). In the presence of 100 nM RAL, only 27% of the total DNA of the WT virus was integrated in comparison with results for the control without RAL, thereby confirming the efficacy of INSTI in blocking the integration step (Fig. (Fig.2A).2A). This decrease in the amount of integrated viral DNA was not observed for the Y143R/C mutants, since the ratio between integrated DNA and unintegrated DNA was not affected by the addition of RAL despite the weaker integration efficiency. The ratio of integrated DNA in the absence of RAL over integrated DNA in the presence of RAL was not significantly different for the Y143R and Y143C mutants, whereas it was equal to 3.6 for the WT. Thus, the inhibitor was not efficient at blocking viral integration of resistant viruses.
A defect in viral integration affects the formation of episomic 2-LTR circles, for which accumulation is a hallmark of integration impairment. This impairment is observed if ST inhibitors are added during infection or in the case of infection by IN catalytic mutants (19, 32, 37). Infection with WT virus in the presence of RAL led to an increase in 2-LTR circles, as expected (Fig. (Fig.2B).2B). The number of 2-LTR circles resulting from resistant viruses in the absence of RAL was also slightly higher than that observed for WT viruses, thus confirming a partial block of integration for the mutants (Fig. (Fig.2B).2B). Finally, no significant difference in the amount of 2-LTR circles was observed for resistant Y143R viruses in the presence or absence of RAL, thus confirming that the inhibitor was not able to block viral integration of this mutant. Intriguingly, a slight increase of 2-LTR circles was observed for the Y143C mutant in the presence of RAL. This may hint that this mutant remains partially sensitive to the inhibitor. However, this hypothesis is not supported by the absence of integration inhibition (Fig. (Fig.2A).2A). Nevertheless, the increase is small compared to that observed with the wild-type virus. Together, these findings demonstrated that (i) the Y143R/C mutation affects viral integration efficiency and (ii) this mutation provides the virus with potent resistance to RAL originating in its inability to block viral integration.
Although greater resistance may explain the emergence of the Y143 mutants, their weak replicative capacity is an obstacle to it. To address the possibility that Y143 mutants have a significant advantage over the other resistant viruses, we compared the viral replication of Y143R/C and G140S/Q148H resistant viruses in the single-round infection assay in the presence of increasing concentration of RAL. As shown in Fig. Fig.3,3, Y143R/C viruses remained poorly susceptible to RAL for concentrations up to 5 μM, demonstrating a dramatic resistance to the drug. In contrast, G140S/Q148H mutants were affected for concentrations above 100 nM, with an apparent EC50 of the drug reached at 2 μM. Furthermore, Y143R/C viruses were significantly more resistant to RAL than the other single mutants, Q148H and N155H viruses (12). In conclusion, Y143R/C mutations provide additional resistance compared to that of Q148 mutants.
The defect in the integration process observed in the viral context could be due to a defect in the 3′-P or ST reaction. To address this question, we investigated the impact of Y143 mutations on the two catalytic activities of the enzyme (3′-P and ST reaction). We first used a steady-state fluorescence anisotropy assay. The determination of the anisotropy of the fluorescence emitted by a fluorophore covalently linked to an ODN mimicking the viral DNA allows monitoring of both the binding of IN to the ODN substrate and the subsequent 3′-P reaction (18). The binding of IN to the ODN increases the steady-state anisotropy value (r), allowing the calculation of the fractional saturation function (Fig. (Fig.4A)4A) . When the fluorophore is linked to the 3′ extremity, the release of the dinucleotide product resulting from the 3′ processing significantly decreases r. First, we used this assay to check that Y143R/C mutants did not affect IN-DNA complex formation (Fig. (Fig.4A).4A). We then monitored 3′ processing as a function of IN concentration in the steady-state anisotropy assay, allowing the reaction to be followed from an initial precisely quantified amount of IN/DNA complexes (18). 3′-P activity as a function of the IN concentration gave a characteristic bell-shaped curve, with activity reaching a plateau at a concentration of about 200 nM similarly to that of the wild-type protein (Fig. (Fig.4B).4B). However, the 3′-P efficiency of the Y143R and Y143C mutants was significantly reduced in comparison with that of the WT, since the overall yields were only 30% and 22% that of WT enzyme for Y143C and Y143R mutants, respectively. Moreover, increasing the mutant concentration did not allow the activity of the wild-type protein to be reached. This result is consistent with previous results showing that mutant protein DNA binding properties were not affected but rather displayed a catalysis-related defect.
Taken together, these data indicate that the Y143R and Y143C mutations affect 3′-P activities of the proteins; these effects occur mostly at the catalytic step without affecting the overall affinity for the viral substrate, a situation reminiscent of that with the Q148 mutation (12). We have previously reported that the RAL-resistant G140S/Q148H double mutant is a catalytic mutant, the activity of which is recovered by allowing the reaction to proceed for an extended time at 37°C. The yield from the 3′-P reaction in Y143R/C mutants also continuously increased over time, reaching up to 80% of the yield obtained with the WT (Fig. (Fig.4C).4C). Thus, Y143R/C mutants may behave similarly to the previously described G140S/Q148H mutant.
To determine whether viral resistance to RAL was related to how it affects IN, we quantified the efficacy of RAL against the enzyme in vitro. To quantify both activities, we used a gel assay that allows the monitoring of both 3′-P and ST (4, 25). We first quantified the 3′-P activity in the presence of increasing RAL concentrations (Fig. (Fig.5A).5A). Results obtained for the 3′-P reaction with radiolabeled ODNs were consistent with those obtained by steady-state anisotropy (reduction of the 3′-P activity of the Y143C and Y143R mutants in comparison with that of WT IN) (Fig. (Fig.5A,5A, right panel). RAL did not significantly inhibit the 3′-P reaction in vitro, as expected for an INSTI compound (14, 21, 29). This assay, which used short blunt ODNs, also revealed that the ST reaction was impaired for both mutants. To allow a precise quantification of RAL inhibition of the ST reaction, the experiment was performed with precleaved ODNs mimicking the processed substrate, which have been reported to enhance ST activity (26). Again, we found that the ST reaction in Y143R/C mutants was less efficient than that in the WT, with the Y143C mutant being the most affected (Fig. (Fig.5B).5B). The ST reaction yield was 60% and 21% of that of the WT for the Y143R and Y143C mutants, respectively (Fig. (Fig.5B,5B, right panel). Despite this weak activity, the quantification of RAL inhibition demonstrated that both mutants were highly resistant to RAL (IC50 above 300 nM) in comparison with the WT (Fig. (Fig.66).
These findings demonstrated that the mutation at the Y143 position was detrimental to overall IN activity, explaining the integration defect observed in cell culture. The nature of the amino acid at the Y143 position (Y143R or Y143C) observed in our clinical study slightly influenced the 3′-P reaction (the difference was not statistically significant in the gel assay). However, the substitution of a C residue (instead of R) was more detrimental to the ST reaction. Nevertheless, even though Y143 mutants are less active than the WT, the mutations observed in patients and studied in this report led to strong RAL resistance.
To gain insight into the origin of the resistance induced by the Y143R/C mutations and for comparison against results with the Q148R mutation, we analyzed the structural and molecular effects induced by these alternative RAL resistance mutations. First we observed that the 3D models of the Y143R/C and Q148R mutants were structurally equivalent and could be superimposed on the WT protein perfectly. In particular, the Y143R/C and Q148R mutations result in the conservation of the catalytic loop, particularly its Ω-shaped hairpin. Y143 and Q148 are adjacent residues in the catalytic loop structure at the neck of its hairpin. This proximity in an invariant structure, as well as the high mobility of the loop, suggests comparable roles of these mutations in resistance to RAL.
We previously demonstrated that the Q148R/H/K mutation alters the specificity of DNA recognition by IN (31). We thus compared the hydrophobicity and hydrogen-bonding patterns of the WT and mutant proteins, both factors playing key roles in the binding of substrates or inhibitors. To compare the contributions of residues Y143R/C and Q148R in modulating the target binding properties and their roles in the complex formation with substrate (DNA) or RAL, we generated the molecular Connolly and MOLCAD surfaces for the WT and mutant INs and analyzed the lipophilic potential (LP) and hydrogen bonding (HB) properties. Lipophilicities of the Y143R/C and Q148R mutants were similar, whereas LP on the IN surface in mutants was considerably lower than that in the WT enzyme (Fig. (Fig.7,7, top). Therefore, a reduced capability to stabilize inhibitors through nonspecific hydrophobic interaction in the vicinity of these residues is expected. The analysis of hydrogen bonding sites on the IN surface also highlighted significant changes to the donor/acceptor properties of the mutants in comparison with those of the WT enzyme (Fig. (Fig.7,7, bottom). In particular, both mutations Q148R and Y143R/C contributed to form a pure hydrogen donor site instead of the mixed donor/acceptor binding sites observed in the context of the wild-type enzyme (Fig. (Fig.8A8A).
Next, two theoretical models of the tetrameric HIV-1 IN with bound LTR DNAs were used to analyze the interactions between the LTR terminal nucleotides and the integrase residues involved in resistance to raltegravir (6, 38). According to the theoretical model suggested by Chen et al. (6), the unpaired 5′-AC base pair of the viral DNA is precisely positioned between these two residues, allowing 3′-processing contacts between Q148/Y143 and the 5′-AC overhang (Fig. (Fig.8B,8B, left). In a model suggested by Wielens et al. (38), the 3′-processed viral DNA is situated in a position allowing short-range contacts between both the Q148 and Y143 residues and the A-T base pair in the third position on the viral LTR (Fig. (Fig.8B,8B, right). Each model corresponds to one of the two possibilities predicted for the way wild-type residues recognize the single adenine at position −2 (6) or the A-T base pairs at position 1 (38). Thus, Y143R could be considered to be a functional and structural alternative to the Q148R mutation.
To date, two different resistance pathways are associated with failure to respond to RAL therapy; these involve the primary mutation N155H or Q148R/H/K (8, 23, 28, 35). A retrospective study of four patients who failed to respond to RAL therapy revealed Y143R/C mutations, supporting the suggestion that these mutations are also responsible for primary resistance to RAL. Here we studied the impact of Y143R/C mutations on viral replication and on the activity of recombinant INs.
Unlike what we previously found for the G140S/Q148H mutant virus, which replicate efficiently in the single-round replication assay (12), the replicative capacity of Y143 mutant virus evaluated in a similar assay was dramatically lower than that of the wild-type virus. This result is consistent with previous observations that show that Y143N mutant replication is significantly delayed in comparison with that of the wild-type virus owing to a specific defect in early replication (34). Quantification of viral DNA species clearly indicated that the Y143 mutations led to a significant replicative defect, resulting in low fitness in the absence of the integrase inhibitor. In contrast, the integration process of both Y143R/C mutants was not affected by RAL, highlighting strong resistance conferred by Y143 substitutions; this may thus confer a higher fitness to the mutant virus in the presence of the inhibitor. In the presence of 5 μM RAL, Y143 mutants were still capable of replicating whereas G140S/Q148R resistant virus was fully susceptible to RAL. This comparison shows that Y143 mutants have a replicative advantage in the presence of the drug over the double mutant, possibly explaining the eventual emergence of this mutant despite its poor replicative capacity.
In vitro, we observed a significant catalytic defect for both 3′ processing and strand transfer activities of Y143R/C proteins. This confirms that integration impairment observed when these mutations were introduced into the viral background originated from a defect in IN activity. If assayed in vitro, mutated INs showed a reduced susceptibility to RAL, which accounts for the resistance of the virus. Thus, Y143R/C appears to be sufficient at conferring high resistance to the enzyme both in vitro and in the viral context.
Interestingly, 3′-P activity was slightly more affected for Y143R mutants, but not ST activity, which was more efficient for this mutant than for the Y143C mutant. This observation could provide a possible explanation for the prevalence of this specific mutant, since both mutants displayed similar resistances to RAL in vitro and in vivo. From this viewpoint, it is interesting to note that the Y143C change requires only a single nucleotide mutation (UAU to UGU or UAC to UGC) whereas Y143R involves a minima two successive mutations (UAU to CGU or UAC to CGC), and thus, Y143C may be a transient form of the Y143R pathway.
Y143 mutants do compare, both in vitro and in vivo, with the Q148H mutant that emerges only in the context of the double mutation G140S/Q148H (12, 35). We previously demonstrated that the Q148H mutant is a thermodynamic mutant, whereas the G140S/Q148H mutant is a kinetics mutant (12). In contrast to the Q148H mutant, which was highly impaired in its overall activity, the G140S/Q148H double mutant was capable of reaching wild-type levels of activity provided that incubation time was increased, indicating a functional rescue of the Q148H defect by the G140S substitution (12). Increasing the incubation time of the reaction in Y143 mutants also restored the enzyme activity to levels observed for the wild-type enzyme, albeit less rapidly than with the G140S/Q148H mutant. This thus suggests that Y143 mutants are functionally equivalent to the G140S/Q148H mutants.
The modeling analysis of the molecular effects induced by these alternative RAL resistance mutations supported this hypothesis in that both the G140S/Q148H and Y143R mutations might prompt alternative substrate recognition, particularly the post-3′-processing contact with the 5′-AC overhang. Although this hypothesis remains a theoretical model owing to the lack of structural data, it is supported by biochemical results showing that both the Y143 and Q148 residues are involved in specifically binding the extremity of viral DNA (13, 15, 22). The resistance of recombinant IN to RAL in vitro and molecular modeling study suggested that the presence of R or H at position 148 prevents RAL binding while mutated IN is still capable of interacting with DNA (12, 31). Thus, the replacement of neighboring Y by R may provide a similar effect, thereby providing an alternative pathway for RAL resistance.
In conclusion, this study shows that mutations at position Y143 constitute one pathway conferring high-level resistance to RAL both in vivo and in vitro. Despite impairment of IN activities observed in vitro, these mutated viruses are able to replicate efficiently in vivo as was observed before for the two other pathways of resistance to RAL (positions 155 and 148). From this viewpoint, it is possible that the polymorphisms present at the origin in the sequences of patients have a positive effect on the activity of the integrase containing the Y143 mutations, which went unnoticed for the single mutations in an HXB2 background that we studied. However, first the genetic backgrounds of the four patients were quite different, and second, several of these polymorphisms (T125A, H51D, N232D, and T206S) are commonly found, in particular T125A and T206S, associated with the CRF02_AG strains, and so far the studies of these strains have failed to show any difference in the resistance patterns induced by RAL (5). Thus, the mechanism underlying this in vivo replicative efficiency of a virus encoding a catalytically crippled integrase remains to be elucidated.
The research leading to these results received funding from Sidaction, Agence Nationale de Recherches sur le SIDA (ANRS), and the European Community's Seventh Framework Programme (FP7/2007-2013) under the project Collaborative HIV and Anti-HIV Drug Resistance Network (CHAIN), grant agreement no. 223131. S. Thierry is the recipient of a fellowship from ANRS.
Published ahead of print on 9 November 2009.