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J. Biol. Chem., Vol. 280, Issue 8, 6792-6801, February 25, 2005

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Structural and Dynamics Studies of the D54A Mutant of Human T Cell Leukemia Virus-1 Capsid Protein^*

From the ^aHoward Hughes Medical Institute, New York, New York, the ^dLaboratory of Biophysical Chemistry, NHLBI, National Institutes of Health, Bethesda, Maryland 20892, the ^fDepartment of Biochemistry and Molecular Biophysics and Department of Microbiology, Columbia University, College of Physicians and Surgeons, and the ⁱDepartment of Molecular Genetics and Microbiology, Stony Brook University, Stony Brook, New York 11794-5222

Received for publication, July 19, 2004 , and in revised form, November 23, 2004.

	ABSTRACT

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
The human T cell leukemia virus and the human immunodeficiency virus share a highly conserved, predominantly helical two-domain mature capsid (CA) protein structure with an N-terminal -hairpin. Despite overall structural similarity, differences exist in the backbone dynamic properties of the CA N-terminal domain. Since studies with other retroviruses suggest that the -hairpin is critical for formation of a CA-CA interface, we investigated the functional role of the human T cell leukemia virus -hairpin by disrupting the salt bridge between Pro¹ and Asp⁵⁴ that stabilizes the -hairpin. NMR ¹⁵N relaxation data were used to characterize the backbone dynamics of the D54A mutant in the context of the N-terminal domains, compared with the wild-type counterpart. Moreover, the effect of the mutation on proteolytic processing and release of virus-like particles (VLPs) from human cells in culture was determined. Conformational and dynamic changes resulting from the mutation were detected by NMR spectroscopy. The mutation also altered the conformation of mature CA in cells and VLPs, as reflected by differential antibody recognition of the wild-type and mutated CA proteins. In contrast, the mutation did not detectably affect antibody recognition of the CA protein precursor or release of VLPs assembled by the precursor, consistent with the fact that the hairpin cannot form in the precursor molecule. The particle morphology and size were not detectably affected. The results indicate that the -hairpin contributes to the overall structure of the mature CA protein and suggest that differences in the backbone dynamics of the -hairpin contribute to mature CA structure, possibly introducing flexibility into interface formation during proteolytic maturation.

	INTRODUCTION

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
All retroviruses encode a precursor polyprotein designated the group-specific antigen (Gag)¹ that is sufficient to direct the formation and release of immature viral particles (1). In the presence of the virus-encoded protease, the Gag precursor is cleaved as the immature virion buds, generating three products: matrix (MA), capsid (CA), and nucleocapsid (NC), which refold and organize spatially to form an infectious mature virion. The CA protein condenses around the NC·RNA complex at the center of the virion to form a core structure within a shell composed of MA proteins. Whereas the noninfectious immature particles of all retroviruses are generally spherical, the mature infectious particles are morphologically distinct and are characterized by the shape of the CA core structure. The mature virion is conical for human immunodeficiency virus type 1 (HIV-1) and other lentiviruses and spherical or irregularly polyhedral for human T cell leukemia virus type 1 (HTLV-1) and other members of the retrovirus group, including Rous sarcoma virus and Moloney murine leukemia virus. The morphology of the core and viral infectivity are closely correlated; mutations that result in loss of the characteristic core structure also result in loss of infectivity (2, 3).

The CA protein that forms the core consists of two independently folded domains joined by a flexible linker peptide (4). The structure is highly conserved among retroviral CA proteins that have been examined (5–10); mutations in the C-terminal domain of HIV-1 block CA protein dimerization and particle assembly (3, 11–13). Mutations in the N-terminal domain (NTD) of HIV-1 impair CA protein assembly to a significantly lesser extent, but the assembled particles are noninfectious (3, 11, 13, 14). In contrast, mutations in the C-terminal domain of the HTLV-1 CA protein had little effect on viral particle assembly and release, whereas mutations throughout the NTD of the protein had much greater impact (15). NTD insertion mutations completely blocked particle release of assembled particles; NTD substitution mutations reduced release of mature particles by 5–50%. In particular, a substitution in helix 3 (Asp⁵⁴-Leu⁵⁵ to Arg-Ser) produced a 10-fold greater defect than substitutions elsewhere in the NTD and blocked particle assembly almost completely (15). These observations suggest that the NTD and C-terminal domain in the HIV-1 and HTLV-1 CA proteins do not function in the same manner.

The N-terminal residues of the HTLV-1 (8), HIV-1 (4), and Rous sarcoma virus (6) CA proteins form a -hairpin structure. It has been proposed that upon proteolysis from the Gag precursor, the liberated amino-terminal end of the CA protein refolds into a -hairpin structure that is stabilized by the formation of a salt bridge between the highly conserved Pro¹ and an Asp/Glu residue from helix 3. The refolded amino terminus is hypothesized to create a new CA-CA interface that is essential for building the condensed conical core that will initiate replication in the next round of infection (2). The fact that the spumavirus subgroup lacks the conserved Pro and Asp residues and does not undergo proteolytic maturation supports the hypothesis that the sequence conservation is linked to the refolding function and core assembly. Interestingly, a 2-amino acid insertion next to the conserved Asp⁵⁴–Leu⁵⁵ residues in helix 3 of the HTLV-1 CA protein significantly reduced formation of viral particles (15). In contrast, an insertion next to the conserved Asp⁵¹–Leu⁵² residues in the NTD of the HIV-1 CA protein had no detectable effect on viral particle formation (13). This difference suggests that structurally equivalent regions can form functionally distinct assembly interfaces during capsid assembly, despite conservation of the secondary and tertiary structures. However, it seems unlikely that the insertion perturbed the -hairpin structure, since a point mutation in the HIV-1 CA helix 3 contact residue, Asp⁵¹ to Ala, reduced viral particle assembly to 20–100% of WT efficiency, reduced viral infectivity correspondingly, and blocked formation of the typical cone-shaped core structure (2). The fact that much of the Gag precursor incorporated into the viral particles was processed aberrantly to truncated CA protein suggests that the mutation exposed the N terminus to inappropriate proteolysis and possibly accounts for the reduced assembly efficiency.

These several observations suggest that the outcome of perturbing the CA protein assembly process can be significantly different for HTLV-1 and HIV-1. Our previous studies (8) indicated that, although the -hairpins of both CAs make contact with the helical core through the conserved salt bridge, they adopt significantly different orientations relative to it. Subsequent NMR transverse relaxation T₁ and steady-state hetero-nuclear nuclear Overhauser effect (NOE) measurements (16) also showed the rigidity of this structural element with values virtually identical to the rest of the well structured protein backbone. In contrast, the -hairpin in HIV-1 CA exhibited a pattern of significant local mobility (on a fast time scale) in the same type of experiments (17). Thus, the same structural element can exhibit distinct dynamics and orientations in two highly related proteins.

In this study, we adopted an approach that combines structural, biochemical, and functional techniques to study the effect of disrupting the -hairpin loop on the structure and dynamics of HTLV-1 CA in solution and on the ability to assemble virus-like particles (VLPs) in mammalian cells. We examined the effect of disrupting the HTLV-1 CA Pro¹–Asp⁵⁴ salt bridge by mutating Asp⁵⁴ to Ala. Consistent with the notion that the HTLV-1 NTD is a less flexible structure (16), we found that the D54A mutation completely destabilized the -hairpin in the HTLV-1 NTD and caused localized conformational and dynamical changes in the region near the mutation. We observed significantly smaller changes (<0.2-ppm amide proton shifts) in helices 1, 2, 3, and 6, which are proximal to the -hairpin in the three-dimensional structure. These findings contrast with the greater perturbation of the equivalent regions that result from the D51A mutation in the HIV-1 CA NTD (2). The conformational alterations detected by NMR in CA₁₃₄ were maintained in the Gag protein expressed in 293T cells, as indicated by differential monoclonal antibody recognition of the WT and D54A mutated CA proteins following proteolytic maturation. The mutated CA protein was assembled and released into VLPs formed by the Gag and Gag-Pro proteins. Moreover, processing of D54A Gag polyprotein, in contrast to HIV-1 D51A Gag, was not affected. Thus, our data indicate that differences in the backbone dynamics of the -hairpin of retroviral capsids correlate with differences in the course of mature CA assembly, processing, and release.

	MATERIALS AND METHODS

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
Protein Expression, Purification, and Preparation of NMR Samples— The DNA fragment encoding the D54A mutant was cloned into the NdeI/SapI sites of the pTYB1 plasmid vector (New England BioLabs), as described previously (8, 16). The recombinant plasmid was transformed into Escherichia coli ER2566 strain for protein expression, and the purified D54A mutant was obtained by the method described previously (8, 16). The uniformly ¹⁵N- and ¹⁵N/¹³C-labeled NMR samples (0.8–1.0 mM protein concentration) were prepared in 250 µl of 10 mM Tris acetate, 0.1 mM NaN₃, pH 6.0, containing 92.5% H₂O and 7.5% ²H₂O.

NMR Spectroscopy—NMR experiments were carried out either on a Bruker AVANCE 600, operating at the ¹H frequency of 600 MHz, or on a Bruker AVANCE 800, operating at the ¹H frequency of 800 MHz, both equipped with a 5-mm shielded triple gradient triple resonance probe head. The NMR experiments were performed at 27 °C. The spectra were processed using the NMRPipe package (18) and analyzed with PIPP/STAPP (19). Chemical shifts were referenced to the internal water signal (4.735822 ppm at 27 °C).

Chemical Shift Assignments—The sequence backbone NMR resonance assignments were derived from the three-dimensional HNCO (20), CBCA(CO)NH (21), and HNCACB (22) experiments. The side chain ¹³C and ¹H assignments were obtained from analysis of the three-dimensional HBHACONH (23), CBCA(CO)NH, and HNCACB spectra.

Backbone ¹⁵N Relaxation Measurements—Longitudinal (T₁) and transversal (T₁) relaxation times for the backbone ¹⁵N nuclei were measured using conventional pulse sequences (24), adapted to include the Watergate scheme (25), pulsed field gradient (26), and a semiconstant time evolution period in t₁ (23). The T₁ experiment used continuous ¹⁵N spin-lock field strength of 2.5 kHz (27).

800-MHz Data—T₁, T₁, and NOE data were acquired as 768*(t₂) x 128*(t₁) data sets with 16, 32, and 64 scans per t₁ point, respectively. Eight spectra were recorded for the T₁ experiments using relaxation delays of 12, 84, 324, 524, 724, 884, 1084, and 1284 ms. For T₁ values, eight spectra were recorded in an interleaved manner (to minimize any effects from spectrometer field drifting and sample heating) using the following ¹⁵N delays: 4.89, 14.49, 28.89, 43.29, 55.29, 74.49, 93.69, and 117.69 ms. ¹⁵N-{¹H} NOE values were measured using a relaxation delay of 3.76 s with the water flip-back NOE pulse sequence described by (28), yielding data sets of 768*(t₂) x 128*(t₁) after accumulation of 64 scans per t₁ point.

600-MHz Data—All experiments were acquired as 512*(t₂) x 128*(t₁) data sets with 32, 64, and 128 scans per t₁ point for T₁, T₁, and NOE, respectively. The following ¹⁵N delays were used for the T₁ experiments: 11, 83, 123, 323, 403, 563, 803, and 1003 ms. The T₁ relaxation decay was sampled at eight different time points: 4.84, 14.44, 26.44, 38.44, 60.04, 88.84, 112.84, and 139.24 ms. ¹⁵N-{¹H} NOE values were measured using a relaxation delay of 3.76 s with the water flip-back NOE pulse sequence.

Relaxation times were calculated by nonlinear fittings of the delay-dependent peak intensities to an exponential decay function with a peak intensity base line of zero. ¹⁵N-{¹H} NOE values were calculated as the intensity ratios of the ¹⁵N-¹H correlation peaks from pairs of spectra acquired with and without ¹H presaturation during the recycle time. The saturated and unsaturated experiments were interleaved; the absence of saturation was obtained by shifting the proton frequency offresonance by 3 MHz during the recycle time. ¹⁵N-{¹H} NOE values were corrected as described previously to compensate for the effects of incomplete ¹H magnetization recovery (28).

Construction of Mutated HTLV-1 gag and gag-pro—HTLV-1 Gag and Gag-Pro mammalian cell expression plasmids were constructed as described previously (15). Briefly, the fragment containing the entire HTLV-1 gag or gag and pro genes was cut from p5'X13 plasmid (a gift from Therese Astier-Gin, University of Bordeaux, France) using AvrII and ApaI sites, and the cDNA fragment was inserted into the plasmid pSV between XbaI and SmaI sites. The plasmids obtained were named pSVgag and pSVgag-pro. ptax, prex, and pX plasmids that express the HTLV-1 X region accessory proteins Tax and Rex were gifts from Dr. David Derse (NCI-Frederick). Mutants in which alanine was substituted for the aspartic acid residue in position 54 (D54A) and alanine or isoleucine was substituted for the glycine residue in position 7 (G7A and G7I, respectively) were constructed using QuikChange site-directed mutagenesis kit (Stratagene) according to the manufacturer's recommendations.

Cell Culture and Transfection—Human 293T cells were maintained at 37 °C and 5% CO₂. in Dulbecco's modified Eagle's medium containing 10% fetal bovine serum. Transfections were performed in 10-cm dishes containing 2 x 10⁶ cells by using either calcium phosphate or Fugene 6 transfection reagent (Roche Applied Science) following the manufacturer's recommendations.

Preparation of Cytoplasmic Extracts, Isolation of VLPs, and Protein Analysis—Cells were lysed on ice in 300–500 µl of radioimmune precipitation buffer (50 mM Tris, pH 8, 150 mM NaCl, 1% Nonidet P-40, 0.5% deoxycholic acid, and 0.1% SDS), sonicated, and centrifuged for 20 min at 4 °C at 13,000 rpm. HTLV-1 Gag proteins were immunoprecipitated with rabbit anti-HTLV-1 capsid antibody (Advanced Biosciences Laboratories) and collected on protein A-Sepharose beads. Proteins were analyzed on 12.5% SDS-polyacrylamide gels and transferred to nitrocellulose membranes (Millipore Corp.). Gag was subsequently detected by Western blot analysis using mouse anti-CA or goat anti-CA antibodies. To detect HTLV-1 Gag proteins released from transfected cells, the culture medium was collected 48 h post-transfection, filtered through a 0.45-µm filter, and spun through a 20% sucrose cushion for 1 h at 25,000 rpm. The pellets were resuspended in phosphate-buffered saline and further analyzed by SDS-PAGE as described above, negative staining and electron microscopy, or rate zonal centrifugation. HTLV-1 Gag proteins were then visualized by Western blot analysis using mouse anti-CA, goat anti-CA, or mouse anti-MA antibodies.

	RESULTS

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
Experimental Approach—Initial studies were conducted to determine whether the substitution to A of the Asp⁵⁴ residue in the NTD of the HTLV-1 CA 134 fragment changed the structure of the protein. The [¹H-¹⁵N] HSQC spectrum, a "fingerprint" of the protein that correlates backbone amide ¹H and ¹⁵N resonances, is a very good indicator of whether the protein undergoes a substantial or minute change(s). This is due to the sensitivity of the NMR resonances to the local chemical environment. A substantial change in the HSQC spectrum can mean global or local structural changes close to the mutation site. This can further be evaluated by looking at the full backbone (amide H and N as well as C, C, and C') resonances and comparing them to the wild type structure. The areas in the structure associated with the largest chemical shift changes are the ones experiencing the largest perturbation due to the mutation. Furthermore, since the backbone chemical shifts are indicative of the secondary structure, they will also provide a measure of possible changes in the secondary structure caused by the mutation. However, backbone resonance data do not differentiate a local geometric change (such as reorientation of a secondary structure element) from local unfolding. Thus, we also obtained NMR backbone relaxation data to complement the above data sets and to address whether the observed changes were associated with folding or unfolding of parts of the protein structure.

Backbone Resonance Assignments for the CA₁₃₄ D54A Mutant—The effect of mutating HTLV-1 CA₁₃₄ residue Asp⁵⁴ to Ala on the -hairpin structure was determined. The [¹H-¹⁵N] HSQC spectrum of the D54A mutant was obtained (Fig. 1B) and is compared with that of wild type (WT) CA₁₃₄ in Fig. 1A. 116 of 121 possible ¹H^N and ¹⁵N resonances (134 residues minus 13 prolines) were assigned (97.5%). Sequential connectivities for the ¹H^N, ¹⁵N, ¹³C, and ¹³C nuclei were derived from HNCACB and CBCA(CO)NH spectra. The ¹³C' frequencies were identified from three-dimensional HNCO analysis. ¹H, ¹H assignments were obtained from HBHACONH analysis. The assignments of ¹³C/¹³C and ¹H/¹H were checked using the [¹³C-¹H] CT-HSQC analysis. In total, 94% of the backbone assignments as well as 90% of the ¹³C and ¹H were obtained. Interestingly, two residues downstream of Asp⁵⁴, Gln⁵⁶ and Asp⁵⁷, could not be assigned due to exchange. This is in contrast to the WT protein, in which these residues do not exhibit exchange (8). Leu⁵⁵ did not exhibit exchange in either protein. The fact that a comparable percentage of backbone assignments could be made for the WT and mutated proteins indicates that the substitution of Ala for Asp⁵⁴ did not cause gross aggregation or unfolding.

View larger version (26K): [in this window] [in a new window]   FIG. 1. HSQC of the wild type HTLV-1 (A) and D54A mutant (B). The corresponding secondary structures are given in C and D for the wild type and D54A mutant HTLV-1, respectively.

Chemical Shift Changes—Backbone ¹H^N, ¹⁵N, ¹³C, and ¹³C chemical shift differences between the WT and the mutant are presented in Fig. 2. A total of 44 of 121 residues exhibited displacements larger than 0.2 ppm in ¹H^N and 0.3 ppm in ¹⁵N chemical shifts. The changes in chemical shifts of backbone ¹³C, ¹³C, ¹H^N, and ¹⁵N reflect the difference in local chemical environment due to the D54A mutation. As might be expected, the largest changes were observed for Asp⁵⁴. In addition, substantial changes also were detected at the N-terminal residues (Pro¹–Met¹⁷) that form the -hairpin in the WT protein. Smaller differences occurred throughout the -1 region (residues Asp¹⁹–Ser²⁸), at the carboxyl terminus of -2 (residues Ile⁴¹–Asp⁴⁹), at -5 (residues Gln¹⁰⁰–Asn¹⁰²) and at the beginning of -6 (residues Gln¹⁰⁷–Tyr¹¹³). A worm representation of the WT protein structure (colored red) with the locations of residues exhibiting significant ¹H and ¹⁵N chemical shift changes (colored green and blue) is shown in Fig. 3. Changes due to D54A mutation were mainly localized to the immediate environment of the -hairpin.

View larger version (23K): [in this window] [in a new window]   FIG. 2. C, C, HN, and 15N chemical shift differences between the wild type and the D54A mutant HTLV-1 versus residue number. The changes in chemical shift at the mutation site are colored in red. The secondary structure elements of the D54A mutant protein are shown in the top panel.

View larger version (66K): [in this window] [in a new window]   FIG. 3. 15N, HN chemical shift differences in the wild type "worm" representation. Residues coded in blue have chemical shift differences larger than 0.2 ppm for HN or 0.3 ppm for 15N, green for the D54A mutation site, yellow for residues that are not assigned (Gln56 and Asp57), and red for residues with small chemical shift differences.

1

1

1

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View larger version (27K): [in this window] [in a new window]   FIG. 4. Experimental T1, T1, and NOE values at 600 MHz (filled circles), 800 MHz (open circles) for the D54A mutant, and 600 MHz (asterisks) for the wild type HTLV-1 are plotted against residue number.

The rotational motion of a nonspherical protein like CA₁₃₄ is described through two independent rotation axes. One slow and one fast diffusion axis lie along the short part and long part of the protein, respectively. Since the relaxation of a ¹⁵N nucleus on the backbone of a protein is dominated by the dipolar interaction along the NH bond, an NH bond oriented parallel to the long axis of the protein will not experience the fast rotation around that axis but will be very sensitive to the slow rotation around the short axis. The reverse will be true for NH bonds oriented along the short axis of the protein. This differentiation will govern how the relaxation rates (i.e. the T₁, T₂, or their ratio) will be distributed throughout the protein.

The ¹⁵N T₁/T₂ ratio depends on the orientation of the NH bond vector relative to the rotational diffusion tensor; this has been called the molecular frame (31). The ¹⁵N T₁/T₂ ratio is, to a first approximation, independent of the rapid internal motions and magnitude of the chemical shift anisotropy. Since the presence of very rapid internal motions causes almost the same fractional increase in T₁ and T₂, T₁/T₂ ratios can be used to derive the rotational diffusion tensor. Residues with a significant contribution from internal motion to the observed T₁/T₂ ratios must be eliminated. Residues with NOE < 0.65 and residues that exhibit conformational exchange on a microsecond/millisecond time scale must be excluded. We found 17 residues with NOE values lower than 0.65 (Met³, His⁴, His⁶, Gly⁷, Ala⁸, Asn¹¹, Arg¹³, Gln²⁹, Gln⁴⁶, Leu⁵⁵, Ile⁸⁷, Gly¹²⁶, Ala¹²⁸, Lys¹²⁹, Ser¹³², Trp¹³³, and Ala¹³⁴). The same criteria used to identify the WT residues with slower internal motions (16) were used in evaluation of the mutant.

The residues that do not fulfill these criteria exhibit additional line broadening, commonly described by the exchange term R_ex (32). The R_ex contribution increases proportionally with the magnetic field. Thus, for the 800-MHz data, the following residues were excluded: Ile⁴¹, Arg⁴², Thr⁸⁸, Asn⁹¹, Arg⁹⁸, Ala¹⁰¹, Arg¹¹⁰, and Gln¹¹⁶. The remaining 82 (or 87; based on 112 residues at 800 MHz) residues at 600 MHz (800 MHz) were used to obtain the rotational diffusion tensor.

Since the orientation of the NH bond defines the relative relaxation rates with respect to the average for the whole protein, one can choose a helix in which typically all of the NH bond vectors are nearly parallel to the helix axis (thus, their T₁/T₂ ratios are uniform) to provide a qualitative measure of how this helix should be oriented with respect to the long axes of the molecule. Different helices exhibit different average T₁/T₂ values corresponding to the orientation relative to the rotational diffusion tensor or the frame of the molecule; the helix with highest average T₁/T₂ value is nearly collinear with the principal component of the diffusion tensor (D||), whereas the helix with the lowest average T₁/T₂ value is nearly perpendicular to it (Fig. 5). The observed average T₁/T₂ values for the different helices in the mutant are as follows: ₁ = 9.29, ₂ = 9.47, ₃ = 8.65, ₄ = 8.88, ₅ = 9.12, and ₆ = 9.29 at 600 MHz and ₁ = 13.69, ₂ = 14.33, ₃ = 13.02, ₄ = 13.49, ₅ = 14.29, and ₆ = 14.19 at 800 MHz. The average values for these helices do not vary as much as they do for the WT protein. This qualitatively suggests that the anisotropy (D||/D) value for the mutant will be smaller than the WT protein (i.e. the mutant has a more compact structure).

View larger version (18K): [in this window] [in a new window]   FIG. 5. T1/T2 ratios measured at 600 MHz (filled circles) and 800 MHz (open circles) versus residue number.

The model free internal diffusion parameters are shown in Fig. 6. The square of generalized order parameter (S²) had average values of 0.87 for the whole molecule, neglecting the high flexibility regions (i.e. the N and C termini). Within the cone model of Brainard and Szabo (34), this average value would correspond to motion within a semiangle of 18°. The N-terminal 13 residues and C-terminal 7 residues of D54A had significantly lower S² values. In contrast, only the C-terminal region of the WT protein possessed low S² values. Two residues in the homologue CyP A binding loop (Gly⁸⁶ and Gly⁹⁵) as well as Leu⁵⁵ showed S² values that are slightly less than average for the well structured region of the mutant protein, whereas in the WT only Gly⁹⁵ exhibited lower than average S² value.

View larger version (16K): [in this window] [in a new window]   FIG. 6. The square of the generalized order parameter (S2) and the effective correlation time (e) versus residue number.

Differential Antibody Recognition of WT and Mutated CA₁₃₄—The chemical shift analysis indicated that disruption of the salt bridge by mutation of Asp⁵⁴ affected regions of ₂, ₃, ₄, ₅, and ₆, regions that are distal to the mutation. The mutation apparently affected an antigenic site in the NTD, since the mutated CA₁₃₄ protein (Fig. 7A, lane 2) was no longer recognized in Western analysis by a monoclonal antibody that detected the WT protein (Fig. 7A, lane 1). In contrast, a polyclonal antibody detected both the WT (Fig. 7B, lane 1) and the mutated (Fig. 7B, lane 2) proteins, as revealed by stripping the blot and reprobing with a rabbit polyclonal anti-CA antibody. The results suggested that disruption of the salt bridge resulted in global changes in the protein that, directly or indirectly, affected the overall protein surface.

View larger version (31K): [in this window] [in a new window]   FIG. 7. Western analysis of CA134. WT CA134 (lane 1) and mutant D54A CA134 (lane 2) proteins were expressed in E. coli and analyzed by SDS-PAGE and Western blotting, using a monoclonal anti-HTLV-1 CA antibody (A) and a polyclonal anti-HTLV-1 CA antibody (B).

Gag and Gag-Pro precursor polyproteins of both the WT and the D54A mutant were blotted and then probed with either a mouse monoclonal antibody (Fig. 8A) or a goat polyclonal antibody (Fig. 8B). As expected, no specific signals were detected in mock-transfected cells (lanes 1 and 4). In contrast, cells transfected with DNA encoding the WT (lanes 2 and 5) or the mutant (lanes 3 and 6) expressed the HTLV-1 Gag precursor protein (p53Gag). The selectivity of the monoclonal antibody for WT over mutated CA was maintained in the natural Gag and Gag-Pro context, as indicated by the differential recognition of the WT CA protein (lanes 2 and 5) but not the mutated D54A CA protein (lanes 3 and 6). This defect was directly related to disruption of the salt bridge between Pro¹ and Asp⁵⁴, because, whereas the monoclonal antibody recognized the WT and the mutant D54A Gag protein (i.e. the unprocessed precursors) almost comparably (Fig. 8A, lanes 2 and 3), the antibody recognized the mature mutant D54A CA protein in the cell lysate and in VLPs poorly (compare lane 2 with lane 3 and lane 5 with lane 6, respectively). In contrast, the polyclonal antibody recognized the WT and the mutant D54A p53 Gag precursor protein as well as the WT and the mutant D54A mature CA proteins to comparable extents (Fig. 8B, compare lane 2 with lane 3 and lane 5 with lane 6), as was the case for CA₁₃₄. Moreover, although the mutated mature D54A CA protein was not efficiently recognized by the monoclonal antibody (Fig. 8A, lanes 5 and 6), comparable amounts of the WT and mutated CA proteins were detected by the polyclonal antibody (Fig. 8B, lanes 5 and 6). These results indicate that destabilization of the hairpin did not prevent release of VLPs at WT levels.

View larger version (45K): [in this window] [in a new window]   FIG. 8. Analysis of the effect of Asp54 mutation on proteolytic processing and VLP release. 293 T cells, mock-transfected (lanes 1 and 4) or transfected with the WT pSV gag (lanes 2 and 5) or with the pSV D54A gag mutant (lanes 3 and 6) construct, were harvested 48 h post-transfection. The Gag-related proteins in the cytoplasm (lanes 1–3) or released as VLPs in the culture media (lanes 4–6) were analyzed by SDS-PAGE and Western blotting using either an anti-HTLV-1 CA monoclonal antibody (A) or a polyclonal antibody (B).

View larger version (42K): [in this window] [in a new window]   FIG. 9. Comparison of the effect of Gly7 and Asp54 mutations on proteolytic processing and VLP release. 293 T cells, mock-transfected (lane 1) or transfected with WT pSV gag (lane 2), pSVgag D54A, G7I, or G7A mutant constructs (lanes 3–5) or with pSVgag-pro (lane 6), pSV gag-pro D54A, G7I, or G7A mutant constructs (lanes 7–9), were harvested 48 h post-transfection. Gag-related protein expression in the cytoplasm (A) or VLPs in the culture media (B and C) were analyzed by SDS-PAGE followed by Western blotting using either an anti-HTLV-1 CA monoclonal antibody (A and B) or a monoclonal anti-MA antibody (C).

View larger version (106K): [in this window] [in a new window]   FIG. 10. Comparison of WT and D54A VLP morphology and size by electron microscopy. VLPs isolated from the culture media from cells transfected with WT pSVgag-pro (A) or pSV gag-pro D54A(B) were stained with uranyl acetate and examined by electron microscopy.

View larger version (31K): [in this window] [in a new window]   FIG. 11. Analysis of WT and D54A VLP on sucrose gradients. VLPs assembled by WT Gag-Pro (A) or the D54A Gag-Pro mutant (B) polyproteins and isolated as described under "Materials and Methods" were gently resuspended in phosphate-buffered saline and layered on a 10–55% sucrose gradient. After centrifugation for 24 h, 25 fractions were collected and analyzed by SDS-PAGE and Western blotting using goat anti-HTLV-I CA antibody. The migration position of the CA-related proteins in the VLPs is shown on the left; the density of the samples () is shown on the right.

	DISCUSSION

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

In the current study, the amino-terminal residues (Pro¹–Arg¹³) of the D54A mutant exhibited relaxation times and NOE values that are characteristic of highly disordered regions in contrast to the WT protein, where that region is well defined. The C and C chemical shifts for these residues are close to their random coil values. In addition, long range NOEs that are characteristics for -hairpin are absent in the D54A mutant. Therefore, all of these data indicate that the amino-terminal region of the mutated CA protein does not form a -hairpin, and, in the context of the Gag precursor, it did not refold into a -hairpin following proteolytic processing. Although the D54A mutation is located near the amino terminus of -3, we observed chemical shift changes (i.e. structural modifications) throughout the -hairpin and its neighboring helices. For the -hairpin region, the chemical shifts were displaced in general toward random coil values, indicating that the disruption of the salt bridge favored the unfolding of the amino terminus of the CA protein and completely destabilized the formation of the -hairpin. Changes in the neighboring helices ₂, ₃, and ₄ reflect differences in packing within them due to the unfolding of the -hairpin. In addition, we also observed secondary effects in ₅ and ₆ due to changes in the first three helices.

Several observations can be made by comparing the 600-MHz relaxation data (T₁, T₁, and NOE) obtained for the mutant and the native proteins. The average T₁ value for the well structured region (residues Lys¹⁸ to Ala¹²³) was higher in the mutant than in the WT protein (754 and 708 ms for the mutant and wild type, respectively); the average T₁ value for the mutant was lower than the WT value (85 versus 92 ms, respectively); the pattern of relaxation data for the CyP A binding loop homologue (residues Gly⁸⁶–Pro⁹⁶) was identical; and the region near the mutation (residues Lys⁵³ and Ala⁵⁴) exhibited higher mobility than its counterpart in the WT. These observations suggest that disruption of the salt bridge does not affect the central loop. Curiously, the differences in average T₁ and T₁ values of the two proteins suggest that a small amount of the mutant protein might be aggregated. Considering the average T₁ and T₁ values at the high protein concentration at which the experiments were carried out as well as the 2–3% experimental errors, the estimated maximum dimer population is 5% (36). A determination of the concentration dependence of protein oligomerization not be helpful in this case, because the experimental error, which is already close to the relative variation of the T₁ and T₁ averages (T₁/T₁ 6.5%; T₁/T₁ 7.6%), will be higher at lower concentrations. Alternatively, the differences in average T₁ and T₁ values might reflect significant differences in the hydrodynamic properties of the two proteins. On the other hand, it is important to point out that the principal axis of diffusion is the same for both WT and mutant proteins (within 5°). Nevertheless, the anisotropy for the mutant is significantly smaller. This can be attributed to the lack (i.e. disruption) of structured -hairpin at the N-terminal end of the mutated protein. Furthermore, the residual in the diffusion tensor fitting to the measured relaxation rates for the mutant is not as good as the WT, as measured by the ². This suggests that there is some local structural difference between the two proteins. This is also supported by long _e values for Gln²⁹ and Leu⁵⁵, suggesting the presence of slow motion that was absent in the WT protein as well as changes in chemical shifts of residues away from the -hairpin and the mutation site. Thus, although the axis of diffusion is the same for the two proteins, the overall shape or topology and local structural as well as dynamic details are not. This result is consistent with the differential antibody recognition between the WT and the D54A mutant proteins observed using Western analysis (Figs. 7, 8, 9). As noted above, there may be sufficient renaturation of the protein during the Western analysis procedure to permit these differences to be distinguished by the antibody probes.

Our study shows that destabilization of the -hairpin by the Asp⁵⁴ to Ala mutation alone did not affect Gag processing, caused no detectable reduction in VLP assembly or release, and did not detectably alter particle size or morphology. Asp⁵⁴ and other residues within -3 may be involved in formation of the maturation interface without participation of the -hairpin. Interestingly, Rayne et al. (15) found that mutation of Asp⁵⁴-Leu⁵⁵ to RS significantly impaired VLP morphogenesis and release from 293T cells. As already noted, mutations in the homologous site in HIV-1, Asp⁵¹-Leu⁵²-Asn⁵³, had much smaller effects compared with effects observed by Rayne et al. (13). The observed differences in the flexibility of the NTD structures may explain the more drastic effect of the mutation on HIV-1 particle production. Conformational alterations in helices 3 and 6 have been associated with the morphological switch from tubes to spheres that occurs upon proteolytic maturation of the HIV-1 Gag precursor. The observation that these regions were only minimally perturbed by disruption of the salt bridge in HTLV-1 is consistent with the fact that both the immature and mature capsids of this virus are spherical (i.e. no apparent morphological transition occurs). Thus, we hypothesize that the function of the -hairpin may be to control formation of interfaces during the proteolytic maturation process to ensure proper core formation. It seems likely that understanding the role of the highly conserved -hairpin and the neighboring helices could provide additional insights into how the retroviral CA proteins form structurally and functionally distinct assemblages.

	FOOTNOTES

 
^* This work was supported in part by National Institutes of Health (NIH) Grant GM 48294 (to C. A. C.) and facilitated by the infrastructure and resources provided by the NIH-funded center for AIDS research program P30 AI 42848. The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

The ¹H^N, ¹⁵N, ¹³C, ¹H, ¹³C', ¹³C, and ¹H chemical shifts for the D54A mutant have been deposited in the BioMagnetic Resonance Bank (www.bmrb.wisc.edu) under BMRB accession code 5334 [BMRB] .

^b A Howard Hughes Medical Institute (HHMI) associate.

^c These authors contributed equally to this work.

^e Present address: Center for Eukaryotic Structural Genomics, Dept. of Biochemistry, University of Wisconsin, Madison, WI 53706-1544.

^g An HHMI principal investigator.

^h To whom correspondence may be addressed. Tel.: 301-402-3029; Fax: 301-402-3405; E-mail: nico{at}helix.nih.gov. j To whom correspondence may be addressed. Tel.: 631-632-8801; Fax: 631-632-9797; E-mail: ccarter{at}ms.cc.sunysb.edu.

¹ The abbreviations used are: Gag, group-specific antigen; MA, matrix; CA, capsid; NC, nucleocapsid; HIV-1, human immunodeficiency virus type 1; HTLV-1, human T cell leukemia virus type 1; NTD, N-terminal domain; VLP, virus-like particle.

	ACKNOWLEDGMENTS

 
We thank M. Auerbach for assistance with electron microscopy and Drs. T. Astier-Gin and D. Derse for reagents.

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TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 

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