- Open Access
Analysis of the mechanical properties of wild type and hyperstable mutants of the HIV-1 capsid
© Ramalho et al. 2016
- Received: 8 February 2016
- Accepted: 8 March 2016
- Published: 15 March 2016
The human immunodeficiency virus (HIV-1) capsid is a self-assembled protein shell that contains the viral genome. During the stages between viral entry into a host cell and nuclear import of the viral DNA, the capsid dissociates in a process known as uncoating, which leads to the release of the viral genetic material. Mutations that alter the stability of the capsid affect the uncoating rate and impair HIV-1 infectivity.
To gain further insight into the role of capsid stability during uncoating, we used atomic force spectroscopy to quantify the stiffness of the capsid. Empty in vitro assemblies of wild type (WT) and mutant recombinant HIV-1 capsid protein (CA) as well as isolated WT and mutant HIV-1 cores (i.e., filled capsids) were analyzed. We find that hyperstable CA mutant assemblies (A204C, A14C/E45C, E45A and E45A/R132T) are significantly stiffer than WT assemblies. However, the hardening effect of disulfide crosslinking (A204C and A14C/E45C) is lower than that of hydrophobic interactions (E45A and E45A/R132T).
Our results demonstrate that mutations that increase the intrinsic stability of the HIV-1 capsid have an increased stiffness of their lattice.
- Atomic force microscopy
The human immunodeficiency virus (HIV) is an enveloped retrovirus whose genetic material is initially in single-stranded RNA form. In mature viral particles, the viral genomic RNA is encapsidated within a cone-shaped capsid. The HIV capsid is a thin conical shell 100–120 nm in length that is formed during viral maturation by the assembly of about 1500 capsid protein (CA) molecules, which are organized into about 250 hexamers and 12 pentamers. It is widely accepted that the presence of pentamers induces the curvature necessary to form the cone shape of the capsid [1–3].
During infection, the viral RNA is reverse transcribed into double-stranded DNA and is then integrated into a host chromosome. Lentiviruses, such as HIV, infect non-dividing cells by traversing the nuclear pore as a nucleoproteic pre-integration complex that contains the viral DNA and integrase. Since the intact capsid is too large to cross the nuclear pore, capsid dissociation (uncoating) and HIV genome release are thought to occur prior to or during nuclear import. The state of the capsid during these steps is unclear, however, the capsid was demonstrated to play major roles, including evasion of innate immunity, motor protein recruitment for transport towards the nucleus , and mediation of nuclear import .
While the mechanism and timing of uncoating are presently unclear (see  and  for reviews), several studies suggest a correlation between capsid stability, uncoating, and viral infectivity. The cellular proteins transportin 3 (TNPO3) and cyclophilin A (CypA) were shown to interact with the viral capsid and to affect infection by modulating uncoating. In vitro data show that transportin 3 promotes uncoating whereas cyclophilin A can inhibit it [7, 8]. Several studies have shown that mutations in the CA protein affect capsid stability and, consequently, the uncoating rate [3, 9–12]. The mutated virus particles have significantly reduced infectivity and exhibit impairments in reverse transcription and virion trafficking. Interestingly, applying inhibitors to partially block reverse transcription gives rise to delayed uncoating [13, 14]. However, Xu et al.  suggested that the early stage of uncoating, which permits entry of a dye molecule into the capsid, is independent of reverse transcription. Nonetheless, the authors postulated a coupling between capsid uncoating and reverse transcription. Recent theoretical models  utilized the conversion of ssRNA into a dsDNA, which has a larger persistence length, as the mechanical driving force for sundering the capsid.
To assess the feasibility of any mechanical model of capsid uncoating, knowledge of capsid stiffness is essential. Previously, we have measured the mechanical properties of HIV-1 particles using nano-indentation measurements with an atomic force microscope (AFM) [17, 18]. Here, we apply the same nano-indentation method to measure the stiffness of empty capsids independently from the viral envelope. These empty capsids self-assemble from purified recombinant CA under high salt conditions in vitro . In addition to measuring the stiffness of self-assembled wild-type capsids, we measured the stiffness values of assemblies of four hyperstable CA mutants. Comparing the wild type with these hyperstable mutants provides insight into the stabilizing effect of disulfide bonds (A204C and A14C/E45C) compared with hydrophobic interactions (E45A and E45A/R132T). Finally, we measured the stiffness of wild-type and E45A viral cores (i.e., of filled capsids) purified from virus particles. Our data demonstrate that mutations that delay HIV-1 uncoating also increase the physical stiffness of the HIV-1 capsid.
CA assemblies form tubes and capsid-like cones
While the hyperstable mutant assemblies maintain their structure for 5–10 days without any detectable damage, WT CA assemblies were the least stable, being visibly disassembled after 48 h. Therefore, all measurements were performed on freshly assembled samples.
In order to compare the in vitro CA assembly with native viral capsids, HIV-1 cores were purified and their structures were visualized by AFM. Cores are, as described, conical in shape, with lengths of 100–120 nm, widths of ~90 nm, and heights of 20–40 nm. No difference in shape was apparent between WT (Fig. 1e) and E45A (not shown) cores. A cross-sectional analysis of representative WT and E45A cores is shown in Fig. 1f. Cores appear to be wider than expected due to convolution between the sample and the AFM probe.
Using sharp AFM tips, we were able to resolve the capsid honeycomb lattice on the surface of the CA assemblies (Fig. 2a). To visualize individual CA hexamers, we zoomed onto the top area of the capsid (Fig. 2b) where the curvature is minimal. In this relatively flat region, the features of the individual hexamers became evident. Measured hexamer diameter was approximately 10 nm, which is consistent with dimensions obtained by other methods .
Hyperstable CA mutations increase capsid stiffness
The stiffness value of the empty WT HIV-1 capsid shell (assembled from recombinant CA) is significantly lower than that of a filled WT capsid (isolated viral core) or an enveloped virion [17, 18, 20]. The stiffness of the empty capsid is comparable to that of the empty capsid of a non-enveloped ssRNA virus, such as the norovirus . In contrast, empty capsids of icosahedral DNA viruses, such as the enveloped herpes simplex virus 1  and the non-enveloped mouse minute virus , are six- to ten-fold stiffer than empty capsids of WT HIV-1. Double-strand DNA is a much stiffer polymer than single-strand DNA or RNA, and is thought to exert high pressures on the capsid wall [16, 24, 25]. This suggests that capsid mechanical properties are highly adapted to the nature of their contents, as previously suggested .
The mechanical properties of retroviral capsids are also modulated by their contents, as we observed by comparing WT HIV-1 cores with recombinant WT CA assemblies. The RNA–protein complexes within the core are not thought to occupy the entire volume of the capsid , which rules out a potential contribution of internal pressure to capsid stiffness. Instead, we hypothesize that the capsid contents and the capsid interact in a manner that reinforces shell stiffness. Similar stabilizing interactions have been described in DNA viruses . In contrast, this reinforcement was not observed in the case of the E45A mutant: the stiffness value for E45A CA assemblies was nearly identical to that of its isolated native core. This is probably because the empty capsid is relatively stiff, such that the capsid’s contents make no apparent contribution.
AFM analysis requires adsorption of the sample to a substrate which often deforms its structure. It is therefore possible, that the deformation of WT HIV-1 core increases the packing density of its contents, which leads to elevated core stiffness values compared with its WT CA counterpart. To address this possibility, we estimated the extent of deformation of the adsorbed core. The maximal thickness of an HIV-1 core is approximately 50 nm . In our AFM analysis, the measured height of isolated cores is 35–50 nm, indicating that adsorption deforms the cores to only a relatively small extent. In addition, the contents of HIV-1 core are thought to occupy only about 20 % of their volume . Taken together, the small deformation and low content occupancy suggest that the apparent reinforcement of WT cores is unlikely to be due to an increase in the packing density of their content.
Stiffness can be defined as the force needed to elastically deform a structure. In multimeric assemblies, such as capsids, this is intrinsically connected to the strength of the interactions between the building blocks and between the monomers within each building block. To explore the effect of inter- and intra-hexamer interactions on the stiffness of the capsid, we studied a series of four capsid-stabilized CA mutants. Mutations E45A and E45A/R132T increase hydrophobic intra-hexamer interactions, the double A14C/E45C mutation introduces covalent intra-hexamer crosslinking via a disulfide bond, while A204C introduces inter-hexamer disulfide crosslinking. We found that all four mutations elevate capsid stiffness in comparison with that of the assembled WT capsid. Interestingly, increasing intra-hexamer interactions resulted in a larger increase in capsid stiffness than from increasing inter-hexamer interactions (Fig. 4). In addition, we observed that increasing the hydrophobic interactions between CA monomers has a significantly larger effect on capsid stiffness than increasing covalent crosslinking (compare Fig. 4 A14C/E45C to E45A and E45A/R132T). To explain this result one has to consider the orientation of the applied force compared to the above monomers interactions. If the force is being applied along the bond axis then covalent bond is expected to be stronger than hydrophobic interactions. However, to measure the stiffness of the capsids we apply a force which is perpendicular to the monomer–monomer interactions axis. Such a perpendicular force does not act on the bond itself, but rather on the interactions between monomers’ interfaces which are mostly hydrophobic. Hence, our results imply that the applied force is acting on interactions between these hydrophobic interfaces and therefore stronger hydrophobic interactions lead to greater stiffness.
Capsid stability has been previously assessed biochemically by assembly efficiency and by comparing dissociation over time [3, 11, 28, 29]. According to those analyses, the capsid disulfide crosslinking mutants, A204C and A14C/E45C, are the most stable of the four mutants analyzed in our study. This apparent discrepancy between the above result and our findings indicates that biochemical stability does not strictly correlate with mechanical stiffness, as has been previously shown .
Finally, the E45A/R132T double mutant has previously been shown to partially restore viral infectivity lost following the original E45A mutation while maintaining the latter’s stability in vitro . The authors suggested that the mutation instead restored the capsid’s interactions with cellular factors, or the consequences thereof. In support of their hypothesis, we find that the stiffness of E45A/R132T mutant CA assemblies is similar to that of the E45A mutant.
We present an AFM analysis of the morphology and mechanical properties of the HIV-1 capsid. The topographic AFM images of CA assemblies and cores are in good agreement with TEM-resolved structures.
Using self-assembled capsid protein structures allowed us to measure capsid stiffness in isolation from its contents, which revealed that the empty WT capsid is extremely soft and that its core contents have a reinforcing effect on it. This finding of an extremely soft capsid fits well with uncoating models  in which DNA synthesis mechanically destabilizes the capsid up to a breaking point.
As our results show, capsid-stabilizing mutations significantly stiffen the capsid structure, which, as Forshey et al.  showed, results in decreased infectivity in several of these mutations. The higher stiffness of the native WT HIV-1 cores compared to the corresponding recombinant CA assemblies suggests that the contents of the viral core buttresses the capsid walls. This suggests, in turn, that the rigidification of the viral genome that occurs during reverse transcription may provide a trigger for uncoating, as suggested from previous uncoating studies in cells [13, 14, 31]. Nevertheless, capsid stiffness is not strictly correlated with infectivity, as shown by the phenotype of the E45A/R132T double mutant. Future work will test the effects of reverse transcription and cellular factors on capsid stiffness and uncoating in vitro.
Purified recombinant WT and mutant HIV-1 capsid proteins (E45A, E45A/R132T, A204C and A14C/E45C) were restored from lyophilized form by resuspension in storage buffer (20 mM Tris–HCl at pH 8.0, 40 mM NaCl, 60 mM β-mercaptoethanol) to a final concentration of 160 μM and self-assembled using a method previously described . Briefly, the resuspended CA was dialyzed against CAB (capsid assembly buffer: 100 mM Tris–HCl at pH 8.0, 200 mM NaCl) overnight at 4 °C in mini dialysis cups (Slyde-A-Lyzer MINI, Thermo Scientific). The resulting CA assemblies were then characterized using AFM and cryo-transmission electron microscopy (cryo-TEM).
Viral core purification
Pseudovirion particles were produced by transfection of human embryonic kidney (HEK) 293T cells with the ΔEnv HIV-1 genome vector containing either the WT or E45A CA mutant sequence, using a method previously described . Briefly, cells were transfected using polyethylenimine (branched, MW ~25,000, Aldrich), the medium was changed after 20 h and the supernatant containing pseudoviral particles was collected 26 h post-transfection. The supernatant was centrifuged for 10 min at 1000×g, filtered through a 0.45 μm pore filter, and centrifuged over an OptiMEM (Sigma) cushion at 100,000×g for 2 h at 4 °C. Viral pellet was resuspended in TNE (100 mM NaCl, 0.1 mM EDTA, 50 mM Tris–HCl pH 7.4) and concentrated using a Vivaspin 20 column (100,000 MWCO, Sartorius). Viral cores were purified by mixing an aliquot of purified and concentrated HIV-1 pseudoviral particles with an equal amount of 1 % Triton X-100 in 3-(N-morpholino)propanesulfonic acid (MOPS) buffer (200 mM NaCl, 100 mM MOPS, pH 7.0). The mixture was incubated for 2 min at 4 °C and cores were spun down at 13,800×g for 8 min. Supernatant was gently removed and pelleted cores were washed twice by adding 80 µL of MOPS buffer and centrifuging at 13,800×g for 8 min. The pellet was resuspended in 10 µL MOPS by pipetting and the resulting cores were characterized using AFM.
Atomic force microscopy (AFM)
For preparation of AFM samples, 10–20 μL of solution containing capsid assemblies or cores was deposited on hexamethyldisilazane- (HMDS-) coated microscope glass slides, incubated for 15–30 min at room temperature, rinsed, and measured in buffer (CAB for capsid assemblies and MOPS for cores).
Measurements were carried out with a JPK Nanowizard ULTRA Speed AFM (JPK Instruments, Berlin, Germany) mounted on an inverted optical microscope (Axio Observer, Carl Zeiss, Heidelberg, Germany). Silicon nitride probes (mean cantilever spring constant, kcant = 0.1 N/m, DNP, Bruker) were used for stiffness measurements, and sharp silicon probes (mean kcant = 0.07 N/m, MSNL, Bruker) were used for high-resolution imaging. Topographic imaging was performed in quantitative imaging (QI) mode, which is a force-curve based imaging mode.
Capsid stiffness was determined based on indentation type experiments as previously described . Briefly, 100 force–distance (F–D) curves were obtained for each point stiffness measurement at a rate of 20 Hz. Each individual F–D curve was acquired by elastically indenting the sample to a maximum of 4 nm (corresponding to a maximum force of 0.2–1.5 nN). Prior to analysis, each curve within a set was shifted to set the deflection in the non-contact section to zero. The set of F–D curves was then averaged. Stiffness was derived mathematically from the slope of the force distance curve. A linear function was fitted to a region of the loading part of the force-distance curve bounded by 3 and 4 nm indentation depths. Averaged force-distance curves were converted from deflection units (V) to a loading force (N) by multiplying the former by the deflection sensitivity (in nm/V, derived from a force-distance curve performed on glass) and the spring constant (N/m) of the cantilever. The measured stiffness comprises the stiffness constants of both the capsid (kCA) and the cantilever (kcant). The stiffness of the capsid was computed according to Hook’s law on the assumption that our experimental system can be modeled as two springs arranged in series. To reduce error in the calculated point stiffness, we chose cantilevers such that the measured point stiffness was less than 70 % of the cantilever spring constant. Statistical differences between stiffness means were tested using parametric (ANOVA) and nonparametric (Kruskal–Wallis) statistics at a level of ≤0.0001. Pairwise comparisons were done using the nonparametric Mann–Whitney U statistical test.
RR and SR performed research and analyzed data, JZ contributed reagents, CA and IR designed research and wrote the manuscript. All authors read and approved the final manuscript.
We thank Einat Nativ-Roth and Alexander Upcher of the Ilse Katz Institute for Nanoscale Science and Technology for high-resolution cryo-TEM measurements. This work was supported by the Israel Science Foundation (Grant 1115/13). R.R. and S.R. were supported by fellowships from the Kreitman school of Advanced Studies (Ben-Gurion University of the Negev). J.Z. and C.A. received support from the National Institutes of Health (P50 GM082251).
The authors declare that they have no competing interests.
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- Ganser BK, Li S, Klishko VY, Finch JT, Sundquist WI. Assembly and analysis of conical models for the HIV-1 core. Science. 1999;283:80–3.View ArticlePubMedGoogle Scholar
- Sundquist WI, Krausslich HG. HIV-1 assembly, budding, and maturation. Cold Spring Harb Perspect Med. 2012;2:a006924.View ArticlePubMedPubMed CentralGoogle Scholar
- Zhao G, Perilla JR, Yufenyuy EL, Meng X, Chen B, Ning J, Ahn J, Gronenborn AM, Schulten K, Aiken C, Zhang P. Mature HIV-1 capsid structure by cryo-electron microscopy and all-atom molecular dynamics. Nature. 2013;497:643–6.View ArticlePubMedPubMed CentralGoogle Scholar
- Malikov V, da Silva ES, Jovasevic V, Bennett G, de Souza Aranha Vieira DA, Schulte B, Diaz-Griffero F, Walsh D, Naghavi MH. HIV-1 capsids bind and exploit the kinesin-1 adaptor FEZ1 for inward movement to the nucleus. Nat Commun. 2015;6:6660.View ArticlePubMedPubMed CentralGoogle Scholar
- Ambrose Z, Aiken C. HIV-1 uncoating: connection to nuclear entry and regulation by host proteins. Virology. 2014;454–455:371–9.View ArticlePubMedGoogle Scholar
- Arhel N, Kirchhoff F. Host proteins involved in HIV infection: new therapeutic targets. Biochim Biophys Acta. 2010;1802:313–21.View ArticlePubMedGoogle Scholar
- Li Y, Kar AK, Sodroski J. Target cell type-dependent modulation of human immunodeficiency virus type 1 capsid disassembly by cyclophilin A. J Virol. 2009;83:10951–62.View ArticlePubMedPubMed CentralGoogle Scholar
- Shah VB, Shi J, Hout DR, Oztop I, Krishnan L, Ahn J, Shotwell MS, Engelman A, Aiken C. The host proteins transportin SR2/TNPO3 and cyclophilin A exert opposing effects on HIV-1 uncoating. J Virol. 2013;87:422–32.View ArticlePubMedPubMed CentralGoogle Scholar
- Ambrose Z, Lee K, Ndjomou J, Xu H, Oztop I, Matous J, Takemura T, Unutmaz D, Engelman A, Hughes SH, KewalRamani VN. Human immunodeficiency virus type 1 capsid mutation N74D alters cyclophilin A dependence and impairs macrophage infection. J Virol. 2012;86:4708–14.View ArticlePubMedPubMed CentralGoogle Scholar
- Byeon IJL, Meng X, Jung JW, Zhao GP, Yang RF, Ahn JW, Shi J, Concel J, Aiken C, Zhang PJ, Gronenborn AM. Structural convergence between cryo-EM and NMR reveals intersubunit interactions critical for HIV-1 capsid function. Cell. 2009;139:780–90.View ArticlePubMedPubMed CentralGoogle 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;76:5667–77.View ArticlePubMedPubMed CentralGoogle Scholar
- Yufenyuy EL, Aiken C. The NTD-CTD intersubunit interface plays a critical role in assembly and stabilization of the HIV-1 capsid. Retrovirology 2013;10:29.View ArticlePubMedPubMed CentralGoogle 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;108:9975–80.View ArticlePubMedPubMed CentralGoogle Scholar
- Yang Y, Fricke T, Diaz-Griffero F. Inhibition of reverse transcriptase activity increases stability of the HIV-1 core. J Virol. 2013;87:683–7.View ArticlePubMedPubMed CentralGoogle Scholar
- Xu H, Franks T, Gibson G, Huber K, Rahm N, Strambio De Castillia C, Luban J, Aiken C, Watkins S, Sluis-Cremer N, Ambrose Z. Evidence for biphasic uncoating during HIV-1 infection from a novel imaging assay. Retrovirology. 2013;10:70.View ArticlePubMedPubMed CentralGoogle Scholar
- Rouzina I, Bruinsma R. DNA confinement drives uncoating of the HIV Virus. Eur Phys J Spec Top. 2014;223:1745–54.View ArticleGoogle Scholar
- Kol N, Shi Y, Tsvitov M, Barlam D, Shneck RZ, Kay MS, Rousso I. A stiffness switch in human immunodeficiency virus. Biophys J. 2007;92:1777–83.View ArticlePubMedPubMed CentralGoogle Scholar
- Pang H-B, Hevroni L, Kol N, Eckert DM, Tsvitov M, Kay MS, Rousso I. Virion stiffness regulates immature HIV-1 entry. Retrovirology. 2013;10:4.View ArticlePubMedPubMed CentralGoogle Scholar
- Ehrlich LS, Agresta BE, Carter CA. Assembly of recombinant human immunodeficiency virus type 1 capsid protein in vitro. J Virol. 1992;66:4874–83.PubMedPubMed CentralGoogle Scholar
- Kol N, Gladnikoff M, Barlam D, Shneck RZ, Rein A, Rousso I. Mechanical properties of murine leukemia virus particles: effect of maturation. Biophys J. 2006;91:767–74.View ArticlePubMedPubMed CentralGoogle Scholar
- Cuellar JL, Meinhoevel F, Hoehne M, Donath E. Size and mechanical stability of norovirus capsids depend on pH: a nanoindentation study. J Gen Virol. 2010;91:2449–56.View ArticlePubMedGoogle Scholar
- Roos WH, Radtke K, Kniesmeijer E, Geertsema H, Sodeik B, Wuite GJ. Scaffold expulsion and genome packaging trigger stabilization of herpes simplex virus capsids. Proc Natl Acad Sci USA. 2009;106:9673–8.View ArticlePubMedPubMed CentralGoogle Scholar
- Carrasco C, Carreira A, Schaap IA, Serena PA, Gomez-Herrero J, Mateu MG, de Pablo PJ. DNA-mediated anisotropic mechanical reinforcement of a virus. Proc Natl Acad Sci USA. 2006;103:13706–11.View ArticlePubMedPubMed CentralGoogle Scholar
- Evilevitch A, Lavelle L, Knobler CM, Raspaud E, Gelbart WM. Osmotic pressure inhibition of DNA ejection from phage. Proc Natl Acad Sci USA. 2003;100:9292–5.View ArticlePubMedPubMed CentralGoogle Scholar
- Ivanovska I, Wuite G, Jonsson B, Evilevitch A. Internal DNA pressure modifies stability of WT phage. Proc Natl Acad Sci USA. 2007;104:9603–8.View ArticlePubMedPubMed CentralGoogle Scholar
- Mateu MG. Mechanical properties of viruses analyzed by atomic force microscopy: a virological perspective. Virus Res. 2012;168:1–22.View ArticlePubMedGoogle Scholar
- Carrasco C, Castellanos M, de Pablo PJ, Mateu MG. Manipulation of the mechanical properties of a virus by protein engineering. Proc Natl Acad Sci USA. 2008;105:4150–5.View ArticlePubMedPubMed CentralGoogle Scholar
- Pornillos O, Ganser-Pornillos BK, Banumathi S, Hua Y, Yeager M. Disulfide bond stabilization of the hexameric capsomer of human immunodeficiency virus. J Mol Biol. 2010;401:985–95.View ArticlePubMedPubMed CentralGoogle Scholar
- Yang R, Shi J, Byeon I-JL, Ahn J, Sheehan JH, Meiler J, Gronenborn AM, Aiken C. Second-site suppressors of HIV-1 capsid mutations: restoration of intracellular activities without correction of intrinsic capsid stability defects. Retrovirology. 2012;9:30.View ArticlePubMedPubMed CentralGoogle Scholar
- Roos WH, Wuite GL. Nanoindentation studies reveal material properties of viruses. Adv Mater. 2009;21:1187–92.View ArticleGoogle 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;26:3025–37.View ArticlePubMedPubMed CentralGoogle Scholar
- Kol N, Tsvitov M, Hevroni L, Wolf SG, Pang HB, Kay MS, Rousso I. The effect of purification method on the completeness of the immature HIV-1 Gag shell. J Virol Methods. 2010;169:244–7.View ArticlePubMedGoogle Scholar