Second-site suppressors of HIV-1 capsid mutations: restoration of intracellular activities without correction of intrinsic capsid stability defects
© Yang et al; licensee BioMed Central Ltd. 2012
Received: 12 January 2012
Accepted: 19 April 2012
Published: 19 April 2012
Disassembly of the viral capsid following penetration into the cytoplasm, or uncoating, is a poorly understood stage of retrovirus infection. Based on previous studies of HIV-1 CA mutants exhibiting altered capsid stability, we concluded that formation of a capsid of optimal intrinsic stability is crucial for HIV-1 infection.
To further examine the connection between HIV-1 capsid stability and infectivity, we isolated second-site suppressors of HIV-1 mutants exhibiting unstable (P38A) or hyperstable (E45A) capsids. We identified the respective suppressor mutations, T216I and R132T, which restored virus replication in a human T cell line and markedly enhanced the fitness of the original mutants as revealed in single-cycle infection assays. Analysis of the corresponding purified N-terminal domain CA proteins by NMR spectroscopy demonstrated that the E45A and R132T mutations induced structural changes that are localized to the regions of the mutations, while the P38A mutation resulted in changes extending to neighboring regions in space. Unexpectedly, neither suppressor mutation corrected the intrinsic viral capsid stability defect associated with the respective original mutation. Nonetheless, the R132T mutation rescued the selective infectivity impairment exhibited by the E45A mutant in aphidicolin-arrested cells, and the double mutant regained sensitivity to the small molecule inhibitor PF74. The T216I mutation rescued the impaired ability of the P38A mutant virus to abrogate restriction by TRIMCyp and TRIM5α.
The second-site suppressor mutations in CA that we have identified rescue virus infection without correcting the intrinsic capsid stability defects associated with the P38A and E45A mutations. The suppressors also restored wild type virus function in several cell-based assays. We propose that while proper HIV-1 uncoating in target cells is dependent on the intrinsic stability of the viral capsid, the effects of stability-altering mutations can be mitigated by additional mutations that affect interactions with host factors in target cells or the consequences of these interactions. The ability of mutations at other CA surfaces to compensate for effects at the NTD-NTD interface further indicates that uncoating in target cells is controlled by multiple intersubunit interfaces in the viral capsid.
KeywordsHIV-1 Capsid Uncoating Suppressor mutations Inhibitor
During retrovirus maturation, the viral capsid protein (CA) assembles into a shell, referred to as the capsid, surrounding the viral genomic ribonucleoprotein (RNP) complex. HIV-1 CA consists of 231 residues that fold into two distinct domains connected by a flexible linker. Various surfaces of CA are involved in HIV-1 capsid formation. Interactions between the N-terminal domains (NTDs) form hexamers through an intersubunit NTD-NTD interface, while the C-terminal domains (CTDs) form dimers that connect adjacent hexamers through a CTD-CTD interface [1–5]. The existence of an NTD-CTD interface in the retroviral capsid was originally inferred from studies of the Rous sarcoma virus (RSV), in which two lethal MHR mutations in the CTD were rescued by compensatory mutations in the NTD . In HIV-1, NTD-CTD contacts were detected by hydrogen-deuterium exchange  and chemical cross-linking , and in structures of CA hexamers and pentamers [9, 10].
Following cell entry, the HIV-1 particle releases its core into the host cytoplasm. Subsequently, the core undergoes an uncoating process, which we define as disassembly or dissociation of the viral capsid from the internal RNP complex . The details of HIV-1 uncoating, including the timing, location, and mechanism, are poorly understood. HIV-1 preintegration complexes (PICs) isolated from the cytoplasm several hours after virus entry contain only low levels of CA protein, suggesting that viral uncoating occurs in the cytoplasm prior to nuclear entry [12, 13]. Reverse transcription complexes have been isolated at earlier time points, with some of these complexes retaining low levels of CA . Recent studies employing imaging and pharmacologic approaches have suggested that uncoating takes place within a few hours following cell entry, and may be linked to reverse transcription [15, 16].
In a previous study, we observed that mutations in CA that result in altering the intrinsic stability of the HIV-1 capsid are associated with impaired infectivity . These capsid stability mutants were competent for viral particle assembly and release and exhibited normal core morphology by electron microscopy, but most exhibited defects in reverse transcription in target cells. One of the mutants, Q63A/Q67A, resulting in unstable capsids in vitro, was competent for reverse transcription but impaired for nuclear import [17, 18]. Interestingly, PICs recovered from this mutant contained elevated levels of CA and were impaired for integration in vitro. Collectively, these studies suggest that uncoating occurs progressively in the cytoplasm and is required for productive reverse transcription, nuclear import, and integration.
Viral determinants besides CA may also be involved in HIV-1 uncoating. A triple-stranded viral DNA structure created during reverse transcription was implicated in uncoating at the nuclear pore. This so-called "DNA flap" structure was shown to be required for nuclear import, but not reverse transcription . Mutations in the HIV-1 matrix protein (MA) can also result in early postentry defects, suggesting that MA also regulates uncoating [20–22].
HIV-1 uncoating may also be modulated by host cell factors. Cellular proteins that may affect uncoating include cyclophilin A and restrictive alleles of TRIM5α [23, 24]. However, whether these proteins play a direct role in uncoating remains unknown, owing to a lack of sensitive and validated cell-based assays to study HIV-1 uncoating in target cells and the difficulty in distinguishing functional from nonfunctional particles . A recent study also reported that lysates from activated CD4+ lymphocytes, but not quiescent CD4+ lymphocytes, induce disassembly of isolated HIV-1 cores in vitro, suggesting that HIV-1 uncoating may differ in activated vs. resting T cells .
Amino acids known to affect HIV-1 capsid stability are located on both domains of the CA protein in varying locations, suggesting that multiple surfaces of the CA protein are involved in stabilizing intersubunit contacts. In the present study, we identified second-site substitutions in CA that restore the ability of HIV-1 mutants containing unstable (P38A) or hyperstable (E45A) capsids to replicate in T cells. Surprisingly, the suppressor mutations rescue HIV-1 infectivity without correcting the intrinsic capsid stability defects associated with the original mutations.
Second-site suppressor mutations restore replication to the P38A and E45A mutants
Second-site suppressor mutations enhance the infectivity of the P38A and E45A mutants in a single-cycle assay
The T216I mutations partially relieves the reverse transcription impairment exhibited by P38A
Second-site suppressor mutations do not restore normal capsid stability in vitro
Suppressor mutations do not restore normal CA assembly kinetics in vitro
Structural assessment of mutant CA151 proteins by NMR
The R132T mutation relieves the cell-cycle dependence of the E45A mutant
Addition of the R132T substitution restores sensitivity of the E45A mutant to the small molecule inhibitor PF74
The T216I mutation restores the ability of P38A particles to saturate TRIM5 host restrictions in monkey cells
The host tripartite motif proteins TRIMCyp and TRIM5α act as barriers to lentivirus infection by targeting the viral capsid in a species-specific manner [34–39]. Restriction of HIV-1 by TRIM5 proteins in simian cells can be saturated by adding virus-like particles (VLPs) in trans. In previous work, we showed that the ability of HIV-1 VLPs to overcome restriction by TRIM5α and TRIMCyp is reduced by CA mutations that destabilize the viral capsid [40, 41], but not by mutations that render the capsid hyperstable, leading us to conclude that efficient trans-abrogation of TRIM5 restriction requires particles with a stable capsid. The ability to abrogate restriction in trans does not depend on the infectivity of the decoy particles, as poorly infectious mutants containing hyperstable capsids (e.g. E45A) were competent in the assay. Our previous results from the assay also supported the conclusion that HIV-1 CA mutants containing unstable capsids undergo accelerated uncoating in target cells. Therefore, the dependence of trans-abrogation activity on capsid stability provides an informative probe of capsid disassembly in target cells.
Uncoating, defined as disassembly of the viral capsid in target cells, is a critical yet poorly understood stage of retroviral infection. In the present work, we observed that the R132T and T216I mutations can partially correct the infectivity defects associated with E45A and P38A, respectively, without correcting the altered capsid stability perturbations created by the original mutations. Similar observations have been reported for RSV, where second site suppressors of CA mutants failed to correct aberrant capsid stability . To elucidate the mechanisms by which the suppressor mutations rescue the original mutant phenotypes, we examined the kinetics of CA protein assembly in vitro, and performed target cell-based assays for viral functions that depend on HIV-1 capsid stability. CA assembly assays did not uncover any correlation between capsid stability and the mutant phenotypes. The cell-based assays, however, were more revealing. The T216I mutation reversed the impaired ability of the P38A mutant to abrogate TRIMCyp- and TRIM5α-mediated restriction of HIV-1. We previously demonstrated a correlation between efficiency of saturation of restriction and intrinsic capsid stability [40, 41]; therefore, the restored ability of P38A/T216I to saturate CA-specific host restriction likely results from stabilization of the viral capsid in the target cells. Furthermore, the R132T mutation partially rescued the impaired ability of the E45A mutant to infect nondividing cells, possibly via reversal of the uncoating defect caused by the E45A mutation. Similarly, the R132T suppressor restored sensitivity to PF74, a compound that inhibits HIV-1 by destabilizing the viral capsid. Collectively, these data strongly suggest that the E45A uncoating defect is rescued by the R132T mutation. Apparently, the putative uncoating defect associated with the P38A and E45A mutations can be rescued in target cells without restoring normal intrinsic capsid stability.
Although uncoating has not been definitively linked to the ability of lentiviruses such as HIV-1 to infect nondividing cells, recent studies demonstrate a role of the CA protein [25, 32], and completion of uncoating may be a requirement for mitosis-independent infection. In accordance with this hypothesis, impaired infection of nondividing cells is associated with CA mutants E45A and Q63A/Q67A, which exhibit hyperstable capsids and/or slower uncoating in target cells. Our observation that R132T rescues the impaired ability of E45A mutant particles to infect arrested cells confirms that the ability of lentiviruses to infect nondividing cells is dependent on a function of the viral capsid, and suggests that this is linked to the mechanism or timing of uncoating.
Our results with the P38A/T216I mutant are reminiscent of another study of CA-NTD mutants in which a second-site substitution at position 208 in the CTD restored replication to mutants containing substitutions at His62 in the NTD, which were associated with structural impairments in the capsid . A recent cryoEM structural model of the assembled capsid may help explain how T216I, which resides in the C-terminal domain of CA, modulates the effects of the P38A substitution in the N-terminal domain. In the model, T216 resides at the CTD-CTD interhexamer interface. Substitution of Cys at positions 207 and 216 resulted in intersubunit disulfide crosslinking within HIV-1 particles, supporting the model . Thus, the T216I substitution may stabilize the CTD-CTD interhexamer interface, thereby offsetting the structural effects of the P38A substitution. Moreover, T216 is also near the NTD-CTD intersubunit interface in the hexamer (Figure 10B), where it may also modulate the stability of the viral capsid.
In our previous studies, we observed a strong dependence of HIV-1 infection on the proper stability of the viral capsid. Specifically, ten CA mutants exhibiting altered capsid stability were impaired for infectivity, while five other mutants with normal infectivity exhibited capsid stability similar to the wild type . This begs the question: how do the CA suppressor mutations rescue HIV-1 infectivity without restoring normal capsid stability in vitro? While the simplest interpretation is that capsid stability is irrelevant for infection, we think this improbable because of the strong correlation we previously observed. One possible alternative explanation is that the biochemical assays we have employed are limited in their ability to detect subtle changes in capsid stability. HIV-1 capsid stability is altered by changes in pH and ionic strength in vitro , and the conditions in which uncoating takes place in target cells are unknown. A second possibility, which we favor, is that the suppressor mutations alter interactions with host factors that participate in HIV-1 uncoating in target cells. Based on the current and previous in vitro and cell-based studies, we propose that HIV-1 infection is dependent on the proper HIV-1 capsid stability, but that the effects of changes in intrinsic capsid stability can also be mitigated by secondary mutations that alter interactions with host cell molecules, or their consequences. In support of this hypothesis, we have observed that an HIV-1 CA mutant with five substitutions in CA, remains infectious despite exhibiting a remarkably stable capsid [33, 44]. Furthermore, the observation that HIV-1 can escape requirement for expression of host factor point mutations in CA confirms that CA exhibits functional plasticity . Additional studies of the determinants of HIV-1 capsid stability and its interactions with host factors should lead to an improved understanding of HIV-1 uncoating and facilitate the discovery and development of pharmacologic inhibitors targeting this critical stage of the virus life cycle.
We have identified second-site suppressors of mutations that alter the stability of the viral capsid. The suppressors restore the ability of the mutant viruses to replicate in T cells, and revert the mutant phenotypes in several cell-based functional assays. However, the biochemical defects in capsid stability produced by the original mutations were not corrected by the suppressor mutations. We propose that proper uncoating in target cells depends on the intrinsic stability of the capsid and on interactions between the viral capsid and host cell factors.
Cells and viruses
293 T, HeLa-CD4/LTR-lacZ (Hela-P4) cells were cultured in Dulbecco's modified Eagle's medium (Cellgro) supplemented with 10% fetal bovine serum, penicillin (50 IU/ml), and streptomycin (50 μg/ml) at 37°C and 5% CO2. CEM cells were cultured in RPMI 1640 medium with the same supplements. The wild-type HIV-1 proviral DNA construct R9 , encoding full-length open reading frames for all HIV-1 structural and accessory genes, was used for these studies. R9 clones encoding mutations E45A and P38A in CA were previously described  and were the generous gift of Dr. Wes Sundquist. HIV-GFP, an env-defective proviral clone encoding GFP in place of nef , was used to generate pseudotyped reporter particles. Viruses were produced by calcium phosphate transfection of 293 T cells (20 μg of plasmid DNA per 2 × 106 cells) as previously described . Five μg of pHCMV-G  were included in the transfection mix for producing VSV-G-pseudotyped viruses with env-defective variants of the respective proviral clones. One day after transfection, culture supernatants were harvested and clarified by passage through 0.45-μm-pore-size filters, and aliquots were frozen and stored at -80°C. The CA concentration of each virus stock was quantified by p24 enzyme-linked immunosorbent assay (ELISA), as previously described . Reverse transcriptase activity was quantified in viral stocks by polymerization of 3H-TTP, as previously described .
HIV-1 replication assay
Viral replication was determined by inoculation of cultures of CEM cells (2 × 105 cells in 0.2 ml) with quantities of virus corresponding to 2 ng of p24 antigen. Every two days, samples of culture supernatants (100 μl) were withdrawn and replaced with an equal volume of fresh medium. Reverse transcriptase activity in culture supernatants was quantified as previously described .
HIV-1 adaptation studies
Virus stocks, normalized for RT activity, were used to inoculate CEM cell cultures. Culture supernatants were harvested at peak of RT activity, and normalized quantities of viruses were re-inoculated in to new CEM cultures. The mutant viruses emerged after approximately one month of culture. Two additional passages of infection in fresh CEM cells were performed until accelerated growth (emergence in 15-20 days) was observed. Cells were harvested immediately following the peak of RT activity from the third passage. Proviral DNA was purified with the DNeasy kit (Qiagen), and regions of Gag were PCR amplified using pfu DNA polymerase (Stratagene) to a 800-bp BssHII-SpeI fragment spanning MA and part of CA coding region and a 500-bp SpeI-ApaI fragment spanning part of CA and NC coding region. The BssHII-SpeI fragment was amplified using sense primer 5'-GGAGATCTCTCGACGCAG-3' and anti-sense primer 5'- TTTAATCCCAGGATTATCCAT-3'. The SpeI-ApaI fragment was amplified using sense primer 5'-GCATGCAGGGCCTATTGC -3' and anti-sense primer 5'- CCTGTCTCTCAGTACAATC-3'. Following identification of additional mutations by DNA sequencing of the PCR products, the corresponding SpeI-ApaI fragments were transferred into R9.E45A or R9.P38A proviral clones. The individual single R132T and T216I mutants were constructed by transferring the mutant SpeI-ApaI segments into the wild-type R9 plasmid. Mutant viral constructs were verified by sequencing of the corresponding regions.
Assay of HIV-1 infectivity
The HeLa-P4 cell line, a HeLa cell clone engineered to express CD4 and an integrated long terminal repeat (LTR)-lacZ reporter cassette , was used to quantify HIV-1 infectivity, as previously described . Infected cells were identified by staining with X-gal, and the specific infectivity of each virus was defined as the number of infected cells per ng of p24 antigen in the inoculum. PF74 was synthesized by the Vanderbilt Institute of Chemical Biology Synthesis Core, Vanderbilt University, Nashville, TN 37212. Assays of PF74 inhibition were performed with a fixed concentration of each virus, with the inhibitor present at the indicated concentrations in the inocula.
Assay of HIV-1 uncoating in vitro
The uncoating of purified HIV-1 cores was assayed as described previously [17, 55]. Purified cores were diluted in STE buffer and incubated at 37°C. Following incubation, the particles were subjected to ultracentrifugation to separate free CA from intact cores. Supernatants and pellets were analyzed for CA content by ELISA to determine the percentage of total CA released from the cores during the incubation.
VSV-pseudotyped HIV-GFP reporter particles were titrated onto FRhK-4 and OMK cell monolayers to determine the appropriate dose for use in the assays. The abrogation-of-restriction assay was performed as previously described [34, 40]. Cultures (20,000 cells per well in 24-well plates) were inoculated with fixed (subsaturating) quantity of an HIV-GFP reporter virus together with various concentrations of VSV-G-pseudotyped non-reporter viruses in the presence of polybrene (8 μg/ml) in a total volume of 300 μl. One day later, complete medium (1 ml) was added. Two days after infection, cells were detached using trypsin and were fixed by the addition of an equal volume of phosphate-buffered saline containing 4% paraformaldehyde. Cellular GFP expression was quantified by flow cytometry by using a FACSCalibur instrument (Becton Dickinson), and the percentage of GFP-expressing cells was determined with Cellquest software. A minimum quantity of 5,000 cells was analyzed for each sample.
CA protein expression and purification
pET21a-based plasmids were constructed for full-length CA protein expression in E. coli BL21-DE3. An NdeI-XhoI fragment was amplified from WT and mutant CA cDNA using sense primer 5'- GGAATCCCATATGCCTATAGTGCAGAACCTCCAGGGG-3' and anti-sense primer 5'- CCCCTCGAGTCACAAAACTCTTGCTTTATGGCCGGG-3' and inserted into the NdeI-XhoI site of pET21a and verified by DNA sequencing. CA proteins were expressed and purified as previously described , except where noted. Proteins were eluted from a UnosphereQ (Bio-Rad) column at NaCl concentrations ranging from 70 to 90 mM. Most of the E45A protein was present in the flow-through. The protein solutions were dialyzed into 50 mM sodium phosphate buffer, pH 8.0, concentrated with Centriprep centrifugal filters (Millipore) to a concentration of ~300-500 μM and stored frozen at -80°C until needed. The proteins were > 99% pure by SDS-PAGE.
To produce isotopically-labeled CA proteins for NMR studies, cells were grown in modified minimal medium at 23°C using 15NH4Cl and/or 13 C glucose as a sole nitrogen and carbon source in E. coli Rosetta 2 (DE3) and induced for 16 h with 0.4 mM IPTG. Soluble CA-NTD proteins were obtained by sonication in lysis buffer containing 25 mM sodium phosphate, pH 7.0, 1 mM DTT, and 0.02% sodium azide followed by ultracentrifugation at 127,000 × g. Proteins were purified by using 10 mL Hi-Trap QP (GE Healthcare, Piscataway, NJ) at pH 7.0 and pH 8.5 with 1 M NaCl gradient. The fractions containing CA-NTD were further purified using Hi-Load Superdex75 26/60 (GE Healthcare, Piscataway, NJ) equilibrated with 25 mM sodium phosphate, pH 6.5, 100 mM NaCl, 1 mM DTT, and 0.02% sodium azide. The molecular mass of each CA-NTD protein was confirmed by LC-ESI-TOF mass spectrometry (Bruker Daltonics, Billerica, MA) and isotope labeling efficiency was estimated to be greater than 99%.
Assays of CA assembly in vitro
Purified full-length CA proteins were assembled by rapid dilution into concentrated NaCl solutions at 23°C to yield a final concentration of 2.25 M NaCl and protein of 50 μM, and the course of the reaction was monitored by turbidity. For kinetic analysis, the reaction was rapidly mixed and placed into a 1-mm quartz cuvette. Approximately 20 s elapsed between the time of the addition of salt and the first time point measured. The increase in optical density was monitored at 350 nm for 30 min as previously described .
with Δδ HN and Δδ N representing the 1HN and 15 N chemical shift differences, respectively.
Cell cycle arrest
To generate cell cycle-arrested target cells for infectivity assay, HeLa-P4 cells (4 × 104 cells/well) were plated in 48-well plates. Aphidicolin (Sigma-Aldrich) was added to a final concentration of 2 μg/ml upon seeding [25, 57]. The next day, cells were inoculated with viruses overnight; aphidicolin was also present during this period. Cultures were replenished with 0.5 ml fresh media and cultured for another 24 hours prior to X-gal staining to reveal infected cells.
Assays of HIV-1 proviral DNA in target cells
HIV-1 stocks were treated with DNaseI, and quantities corresponding to 100 ng p24 were used to inoculate cultures of HeLa-P4 cells (100,000) in triplicate, in the presence or absence of the reverse transcriptase inhibitor Efavirenz (1 μM). Total DNA was harvested at 8 hours post-inoculation and was purified using the DNeasy kit (Qiagen). DNA was eluted in 100 μl of water. Five microliter aliquots were assayed for 2nd-strand transfer reverse transcription products by quantitative PCR, as previously described .
Molecular graphics images in Figure 6 were produced with MOLMOL , and those in Figure 9 were produced using the UCSF Chimera package from the Resource for Biocomputing, Visualization, and Informatics at the University of California, San Francisco (supported by NIH P41 RR-01081). The images are based on the 1.9A resolution structure of the CA hexamer (Protein Data Bank entry 3H47.pdb).
Human immunodeficiency virus type 1
Nuclear magnetic resonance
Rous sarcoma virus
Vesicular stomatitis virus glycoprotein.
We thank Wes Sundquist for the P38A and E45A mutant proviruses. The following reagents were obtained through the NIH AIDS Research and Reference Reagent Program, Division of AIDS, NIAID, NIH: HIV-Ig from NABI and NHLBI; HIV-1 p24 hybridoma (183-H12-5 C) from Dr. Bruce Chesebro, and Efavirenz. Compound PF74 was provided by the Vanderbilt Institute of Chemical Biology Synthesis Core. This work was supported by NIH grants AI076121 and GM082251.
- Gamble TR, Vajdos FF, Yoo S, Worthylake DK, Houseweart M, Sundquist WI, Hill CP: Crystal structure of human cyclophilin A bound to the amino-terminal domain of HIV-1 capsid. Cell. 1996, 87: 1285-1294. 10.1016/S0092-8674(00)81823-1.View ArticlePubMedGoogle Scholar
- Gamble TR, Yoo S, Vajdos FF, von Schwedler UK, Worthylake DK, Wang H, McCutcheon JP, Sundquist WI, Hill CP: Structure of the carboxyl-terminal dimerization domain of the HIV-1 capsid protein. Science. 1997, 278: 849-853. 10.1126/science.278.5339.849.View ArticlePubMedGoogle Scholar
- Gitti RK, Lee BM, Walker J, Summers MF, Yoo S, Sundquist WI: Structure of the amino-terminal core domain of the HIV-1 capsid protein. Science. 1996, 273: 231-235. 10.1126/science.273.5272.231.View ArticlePubMedGoogle Scholar
- Li S, Hill CP, Sundquist WI, Finch JT: Image reconstructions of helical assemblies of the HIV-1 CA protein. Nature. 2000, 407: 409-413. 10.1038/35030177.View ArticlePubMedGoogle Scholar
- Momany C, Kovari LC, Prongay AJ, Keller W, Gitti RK, Lee BM, Gorbalenya AE, Tong L, McClure J, Ehrlich LS, et al: Crystal structure of dimeric HIV-1 capsid protein. Nat Struct Biol. 1996, 3: 763-770. 10.1038/nsb0996-763.View ArticlePubMedGoogle Scholar
- Bowzard JB, Wills JW, Craven RC: Second-site suppressors of Rous sarcoma virus Ca mutations: evidence for interdomain interactions. J Virol. 2001, 75: 6850-6856. 10.1128/JVI.75.15.6850-6856.2001.PubMed CentralView ArticlePubMedGoogle Scholar
- Lanman J, Lam TT, Emmett MR, Marshall AG, Sakalian M, Prevelige PE: Key interactions in HIV-1 maturation identified by hydrogen-deuterium exchange. Nat Struct Mol Biol. 2004, 11: 676-677. 10.1038/nsmb790.View ArticlePubMedGoogle Scholar
- Lanman J, Lam TT, Barnes S, Sakalian M, Emmett MR, Marshall AG, Prevelige PE: Identification of novel interactions in HIV-1 capsid protein assembly by high-resolution mass spectrometry. J Mol Biol. 2003, 325: 759-772. 10.1016/S0022-2836(02)01245-7.View ArticlePubMedGoogle Scholar
- Ganser-Pornillos BK, Cheng A, Yeager M: Structure of full-length HIV-1 CA: a model for the mature capsid lattice. Cell. 2007, 131: 70-79. 10.1016/j.cell.2007.08.018.View ArticlePubMedGoogle Scholar
- Pornillos O, Ganser-Pornillos BK, Kelly BN, Hua Y, Whitby FG, Stout CD, Sundquist WI, Hill CP, Yeager M: X-ray structures of the hexameric building block of the HIV capsid. Cell. 2009, 137: 1282-1292. 10.1016/j.cell.2009.04.063.PubMed CentralView ArticlePubMedGoogle Scholar
- Aiken C: Viral and cellular factors that regulate HIV-1 uncoating. Curr Opin HIV AIDS. 2006, 1: 194-199. 10.1097/01.COH.0000221591.11294.c1.View ArticlePubMedGoogle Scholar
- Miller MD, Farnet CM, Bushman FD: Human immunodeficiency virus type 1 preintegration complexes: studies of organization and composition. J Virol. 1997, 71: 5382-5390.PubMed CentralPubMedGoogle Scholar
- Farnet CM, Haseltine WA: Determination of viral proteins present in the human immunodeficiency virus type 1 preintegration complex. J Virol. 1991, 65: 1910-1915.PubMed CentralPubMedGoogle Scholar
- Fassati A, Goff SP: Characterization of intracellular reverse transcription complexes of human immunodeficiency virus type 1. J Virol. 2001, 75: 3626-3635. 10.1128/JVI.75.8.3626-3635.2001.PubMed CentralView ArticlePubMedGoogle Scholar
- Arfi V, Lienard J, Nguyen XN, Berger G, Rigal D, Darlix JL, Cimarelli A: Characterization of the behavior of functional viral genomes during the early steps of human immunodeficiency virus type 1 infection. J Virol. 2009, 83: 7524-7535. 10.1128/JVI.00429-09.PubMed CentralView ArticlePubMedGoogle Scholar
- Hulme AE, Perez O, Hope TJ: Complementary assays reveal a relationship between HIV-1 uncoating and reverse transcription. Proc Natl Acad Sci USA. 2011, 108: 9975-9980. 10.1073/pnas.1014522108.PubMed CentralView ArticlePubMedGoogle Scholar
- Forshey BM, von Schwedler U, Sundquist WI, Aiken C: Formation of a human immunodeficiency virus type 1 core of optimal stability is crucial for viral replication. J Virol. 2002, 76: 5667-5677. 10.1128/JVI.76.11.5667-5677.2002.PubMed CentralView ArticlePubMedGoogle Scholar
- Dismuke DJ, Aiken C: Evidence for a functional link between uncoating of the human immunodeficiency virus type 1 core and nuclear import of the viral preintegration complex. J Virol. 2006, 80: 3712-3720. 10.1128/JVI.80.8.3712-3720.2006.PubMed CentralView ArticlePubMedGoogle Scholar
- Arhel NJ, Souquere-Besse S, Munier S, Souque P, Guadagnini S, Rutherford S, Prevost MC, Allen TD, Charneau P: HIV-1 DNA Flap formation promotes uncoating of the pre-integration complex at the nuclear pore. EMBO J. 2007, 26: 3025-3037. 10.1038/sj.emboj.7601740.PubMed CentralView ArticlePubMedGoogle Scholar
- Casella CR, Raffini LJ, Panganiban AT: Pleiotropic mutations in the HIV-1 matrix protein that affect diverse steps in replication. Virology. 1997, 228: 294-306. 10.1006/viro.1996.8355.View ArticlePubMedGoogle Scholar
- Kiernan RE, Ono A, Englund G, Freed EO: Role of matrix in an early postentry step in the human immunodeficiency virus type 1 life cycle. J Virol. 1998, 72: 4116-4126.PubMed CentralPubMedGoogle Scholar
- Davis MR, Jiang J, Zhou J, Freed EO, Aiken C: A mutation in the human immunodeficiency virus type 1 Gag protein destabilizes the interaction of the envelope protein subunits gp120 and gp41. J Virol. 2006, 80: 2405-2417. 10.1128/JVI.80.5.2405-2417.2006.PubMed CentralView ArticlePubMedGoogle Scholar
- Stremlau M, Perron M, Lee M, Li Y, Song B, Javanbakht H, Diaz-Griffero F, Anderson DJ, Sundquist WI, Sodroski J: Specific recognition and accelerated uncoating of retroviral capsids by the TRIM5alpha restriction factor. Proc Natl Acad Sci USA. 2006, 103: 5514-5519. 10.1073/pnas.0509996103.PubMed CentralView ArticlePubMedGoogle Scholar
- Luban J, Bossolt KL, Franke EK, Kalpana GV, Goff SP: Human immunodeficiency virus type 1 Gag protein binds to cyclophilins A and B. Cell. 1993, 73: 1067-1078. 10.1016/0092-8674(93)90637-6.View ArticlePubMedGoogle Scholar
- Yamashita M, Perez O, Hope TJ, Emerman M: Evidence for direct involvement of the capsid protein in HIV infection of nondividing cells. PLoS Pathog. 2007, 3: 1502-1510.View ArticlePubMedGoogle Scholar
- Auewarakul P, Wacharapornin P, Srichatrapimuk S, Chutipongtanate S, Puthavathana P: Uncoating of HIV-1 requires cellular activation. Virology. 2005, 337: 93-101. 10.1016/j.virol.2005.02.028.View ArticlePubMedGoogle Scholar
- Campbell S, Vogt VM: Self-assembly in vitro of purified CA-NC proteins from Rous sarcoma virus and human immunodeficiency virus type 1. J Virol. 1995, 69: 6487-6497.PubMed CentralPubMedGoogle Scholar
- Ehrlich LS, Agresta BE, Carter CA: Assembly of recombinant human immunodeficiency virus type 1 capsid protein in vitro. J Virol. 1992, 66: 4874-4883.PubMed CentralPubMedGoogle Scholar
- Gross I, Hohenberg H, Krausslich HG: In vitro assembly properties of purified bacterially expressed capsid proteins of human immunodeficiency virus. Eur J Biochem. 1997, 249: 592-600. 10.1111/j.1432-1033.1997.t01-1-00592.x.View ArticlePubMedGoogle Scholar
- von Schwedler UK, Stemmler TL, Klishko VY, Li S, Albertine KH, Davis DR, Sundquist WI: Proteolytic refolding of the HIV-1 capsid protein amino-terminus facilitates viral core assembly. EMBO J. 1998, 17: 1555-1568. 10.1093/emboj/17.6.1555.PubMed CentralView ArticlePubMedGoogle Scholar
- Douglas CC, Thomas D, Lanman J, Prevelige PE: Investigation of N-terminal domain charged residues on the assembly and stability of HIV-1 CA. Biochemistry. 2004, 43: 10435-10441. 10.1021/bi049359g.View ArticlePubMedGoogle Scholar
- Yamashita M, Emerman M: Capsid is a dominant determinant of retrovirus infectivity in nondividing cells. J Virol. 2004, 78: 5670-5678. 10.1128/JVI.78.11.5670-5678.2004.PubMed CentralView ArticlePubMedGoogle Scholar
- Shi J, Zhou J, Shah VB, Aiken C, Whitby K: Small-molecule inhibition of human immunodeficiency virus type 1 infection by virus capsid destabilization. J Virol. 2011, 85: 542-549. 10.1128/JVI.01406-10.PubMed CentralView ArticlePubMedGoogle Scholar
- Cowan S, Hatziioannou T, Cunningham T, Muesing MA, Gottlinger HG, Bieniasz PD: Cellular inhibitors with Fv1-like activity restrict human and simian immunodeficiency virus tropism. Proc Natl Acad Sci USA. 2002, 99: 11914-11919. 10.1073/pnas.162299499.PubMed CentralView ArticlePubMedGoogle Scholar
- Hatziioannou T, Perez-Caballero D, Yang A, Cowan S, Bieniasz PD: Retrovirus resistance factors Ref1 and Lv1 are species-specific variants of TRIM5alpha. Proc Natl Acad Sci USA. 2004, 101: 10774-10779. 10.1073/pnas.0402361101.PubMed CentralView ArticlePubMedGoogle Scholar
- Keckesova Z, Ylinen LM, Towers GJ: The human and African green monkey TRIM5alpha genes encode Ref1 and Lv1 retroviral restriction factor activities. Proc Natl Acad Sci USA. 2004, 101: 10780-10785. 10.1073/pnas.0402474101.PubMed CentralView ArticlePubMedGoogle Scholar
- Yap MW, Nisole S, Lynch C, Stoye JP: Trim5alpha protein restricts both HIV-1 and murine leukemia virus. Proc Natl Acad Sci USA. 2004, 101: 10786-10791. 10.1073/pnas.0402876101.PubMed CentralView ArticlePubMedGoogle Scholar
- Nisole S, Lynch C, Stoye JP, Yap MW: A Trim5-cyclophilin A fusion protein found in owl monkey kidney cells can restrict HIV-1. Proc Natl Acad Sci USA. 2004, 101: 13324-13328. 10.1073/pnas.0404640101.PubMed CentralView ArticlePubMedGoogle Scholar
- Sayah DM, Sokolskaja E, Berthoux L, Luban J: Cyclophilin A retrotransposition into TRIM5 explains owl monkey resistance to HIV-1. Nature. 2004, 430: 569-573. 10.1038/nature02777.View ArticlePubMedGoogle Scholar
- Forshey BM, Shi J, Aiken C: Structural requirements for recognition of the human immunodeficiency virus type 1 core during host restriction in owl monkey cells. J Virol. 2005, 79: 869-875. 10.1128/JVI.79.2.869-875.2005.PubMed CentralView ArticlePubMedGoogle Scholar
- Shi J, Aiken C: Saturation of TRIM5alpha-mediated restriction of HIV-1 infection depends on the stability of the incoming viral capsid. Virology. 2006, 350: 493-500. 10.1016/j.virol.2006.03.013.View ArticlePubMedGoogle Scholar
- Noviello CM, Lopez CS, Kukull B, McNett H, Still A, Eccles J, Sloan R, Barklis E: Second-site compensatory mutations of HIV-1 capsid mutations. J Virol. 2011, 85: 4730-4738. 10.1128/JVI.00099-11.PubMed CentralView ArticlePubMedGoogle Scholar
- Byeon IJ, Meng X, Jung J, Zhao G, Yang R, Ahn J, Shi J, Concel J, Aiken C, Zhang P, Gronenborn AM: Structural convergence between Cryo-EM and NMR reveals intersubunit interactions critical for HIV-1 capsid function. Cell. 2009, 139: 780-790. 10.1016/j.cell.2009.10.010.PubMed CentralView ArticlePubMedGoogle Scholar
- Blair WS, Pickford C, Irving SL, Brown DG, Anderson M, Bazin R, Cao J, Ciaramella G, Isaacson J, Jackson L, et al: HIV capsid is a tractable target for small molecule therapeutic intervention. PLoS Pathog. 2010, 6: e1001220-10.1371/journal.ppat.1001220.PubMed CentralView ArticlePubMedGoogle Scholar
- Lee K, Ambrose Z, Martin TD, Oztop I, Mulky A, Julias JG, Vandegraaff N, Baumann JG, Wang R, Yuen W, et al: Flexible use of nuclear import pathways by HIV-1. Cell Host Microbe. 2010, 7: 221-233. 10.1016/j.chom.2010.02.007.PubMed CentralView ArticlePubMedGoogle Scholar
- Swingler S, Gallay P, Camaur D, Song J, Abo A, Trono D: The Nef protein of human immunodeficiency virus type 1 enhances serine phosphorylation of the viral matrix. J Virol. 1997, 71: 4372-4377.PubMed CentralPubMedGoogle Scholar
- von Schwedler UK, Stray KM, Garrus JE, Sundquist WI: Functional surfaces of the human immunodeficiency virus type 1 capsid protein. J Virol. 2003, 77: 5439-5450. 10.1128/JVI.77.9.5439-5450.2003.PubMed CentralView ArticlePubMedGoogle Scholar
- He J, Chen Y, Farzan M, Choe H, Ohagen A, Gartner S, Busciglio J, Yang X, Hofmann W, Newman W, et al: CCR3 and CCR5 are co-receptors for HIV-1 infection of microglia. Nature. 1997, 385: 645-649. 10.1038/385645a0.View ArticlePubMedGoogle Scholar
- Chen C, Okayama H: High-efficiency transformation of mammalian cells by plasmid DNA. Mol Cell Biol. 1987, 7: 2745-2752.PubMed CentralView ArticlePubMedGoogle Scholar
- Yee JK, Friedmann T, Burns JC: Generation of high-titer pseudotyped retroviral with very broad host range. Methods Cell Biol. 1994, 43: 99-112.View ArticlePubMedGoogle Scholar
- Wehrly K, Chesebro B: p24 antigen capture assay for quantification of human immunodeficiency virus using readily available inexpensive reagents. Methods. 1997, 12: 288-293. 10.1006/meth.1997.0481.View ArticlePubMedGoogle Scholar
- Aiken C, Trono D: Nef stimulates human immunodeficiency virus type 1 proviral DNA synthesis. J Virol. 1995, 69: 5048-5056.PubMed CentralPubMedGoogle Scholar
- Yang R, Aiken C: A mutation in alpha helix 3 of CA renders human immunodeficiency virus type 1 cyclosporin A resistant and dependent: rescue by a second-site substitution in a distal region of CA. J Virol. 2007, 81: 3749-3756. 10.1128/JVI.02634-06.PubMed CentralView ArticlePubMedGoogle Scholar
- Charneau P, Alizon M, Clavel F: A second origin of DNA plus-strand synthesis is required for optimal human immunodeficiency virus replication. J Virol. 1992, 66: 2814-2820.PubMed CentralPubMedGoogle Scholar
- Shah VB, Aiken C: In vitro Uncoating of HIV-1 Cores. J Vis Exp. 2011, 57: e3384-Google Scholar
- Ganser BK, Li S, Klishko VY, Finch JT, Sundquist WI: Assembly and analysis of conical models for the HIV-1 core. Science. 1999, 283: 80-83. 10.1126/science.283.5398.80.View ArticlePubMedGoogle Scholar
- Qi M, Yang R, Aiken C: Cyclophilin A-dependent restriction of human immunodeficiency virus type 1 capsid mutants for infection of nondividing cells. J Virol. 2008, 82: 12001-12008. 10.1128/JVI.01518-08.PubMed CentralView ArticlePubMedGoogle Scholar
- Koradi R, Billeter M, Wuthrich K: MOLMOL: a program for display and analysis of macromolecular structures. J Mol Graph. 1996, 14: 51-55. 10.1016/0263-7855(96)00009-4. 29-32View ArticlePubMedGoogle Scholar
- Pettersen EF, Goddard TD, Huang CC, Couch GS, Greenblatt DM, Meng EC, Ferrin TE: UCSF Chimera-a visualization system for exploratory research and analysis. J Comput Chem. 2004, 25: 1605-1612. 10.1002/jcc.20084.View ArticlePubMedGoogle Scholar
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