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Logo of nihpaAbout Author manuscriptsSubmit a manuscriptHHS Public Access; Author Manuscript; Accepted for publication in peer reviewed journal;
Structure. Author manuscript; available in PMC 2010 August 1.
Published in final edited form as:
PMCID: PMC2913127

Crystal structure of the APOBEC3G catalytic domain reveals potential oligomerization interfaces


APOBEC3G is a DNA cytidine deaminase that has anti-viral activity against HIV-1 and other pathogenic viruses. In this study the crystal structure of the catalytically active C-terminal domain was determined to 2.25 Å. This structure corroborates features previously observed in NMR studies, a bulge in the second β-strand and a lengthening of the second α-helix. Oligomerization is postulated to be critical for the function of APOBEC3G. In this structure, four extensive intermolecular interfaces are observed suggesting potential models for APOBEC3G oligomerization. The structural and functional significance of these interfaces was probed by solution NMR and disruptive variants were designed and tested for DNA deaminase and anti-HIV activities. The variant designed to disrupt the most extensive interface lost both activities. NMR solution data provides evidence that another interface, which coordinates a novel zinc site, also exists. Thus, the observed crystallographic interfaces of APOBEC3G may be important for both oligomerization and function.


APOBEC3G (A3G), a DNA cytidine deaminase, belongs to the larger APOBEC family of proteins. Members include activation induced deaminase (AID), APOBEC1, APOBEC2, APOBEC4, and, in addition to A3G, six APOBEC3s (A3s), (Jarmuz et al., 2002; Sawyer et al., 2004; Zhang and Webb, 2004; Conticello et al., 2005) [reviews by: (Goila-Gaur and Strebel, 2008; Chiu and Greene, 2008, 2009; Malim, 2009)]. These proteins have diverse biological functions that include editing mRNA (APOBEC1), diversifying antibody gene DNA (AID), and restricting the mobilization of retroviruses and retrotransposons (A3s).

All the A3s (A, B, C, DE, F, G, and H) are single-strand DNA cytidine deaminases known to inhibit multiple retroelement substrates. The deaminases possess either one or two conserved zinc-coordinating (Z) motifs, with the consensus amino acid signature, Hx1Ex24–28PCx2–4C (Conticello, 2008; Jarmuz et al., 2002; LaRue et al., 2008; LaRue et al., 2009). Zinc coordination is mediated by a histidine and two cysteines. The hydroxide that subsequently converts cytidine to uridine (C-to-U deamination) is generated when the active site glutamate removes hydrogen from water.

HIV-1, the retrovirus that causes AIDS, is the most prominent pathogen restricted by A3G (Harris et al., 2003; Zhang et al., 2003; Lecossier et al., 2003; Mangeat et al., 2003; Sheehy et al., 2002). A3G suppresses HIV-1 infectivity by entering viral particles and deaminating the viral cDNA cytidines to uridines during reverse transcription. The uridines introduce adenosines in the complementary genomic strand which are often detected as the hallmark G-to-A hypermutations. These mutations produce stop codons and amino acid changes that can ultimately inactivate the virus. However, HIV-1 efficiently counteracts restriction by A3G, with the viral Vif protein binding and targeting the cytidine deaminase for proteasomal degradation [reviews by: (Goila-Gaur and Strebel, 2008; Chiu and Greene, 2008, 2009; Malim, 2009)].

The life-or-death interaction between human A3G and HIV-1 Vif has made A3G the prototype member of the APOBEC family for biochemical and structural investigations. A3G has two consecutive Z-motifs giving the sequence of the enzyme an internal pseudo-symmetry. In the past two years, significant advances have been made in elucidating the structure of the C-terminal catalytic domain of A3G (A3G-CTD) but the conformations of several important functional regions differ between three NMR structures and a single X-ray crystal structure (Chen et al., 2008; Furukawa et al., 2009; Harjes et al., 2009; Holden et al., 2008). The present study aims to resolve some of the controversy by determining a new, higher-resolution crystal structure of A3G-CTD. This new structure differs from the recently reported crystal structure (PDB ID: 3E1U with R-factor 25.2%, R-free 26.7%, Resolution 2.3 Å) (Holden et al., 2008) and strongly supports the structural integrity of the NMR structures (PDB IDs: 2JYW, 2KBO, 2KEM) (Chen et al., 2008; Furukawa et al., 2009; Harjes et al., 2009). Most importantly, this new crystal structure reveals four extensive interfaces, of which one or more may be important for A3G oligomerization and biological activity.


The crystal structure of the human APOBEC3G catalytic domain, A3G191-384-2K3A—where 2K3A is L234K, C243A, F310K, C321A and C356A (Harjes et al., 2009)—was determined at 2.25 Å resolution. The A3G191-384-2K3A crystal structure was solved by molecular replacement, using a highly truncated model based on (PDB ID: 3E1U) (Holden et al., 2008). The crystal structure contains two molecules of A3G191-384-2K3A in the asymmetric unit. The final refinement statistics are: Rfactor (Rfree) 16.57% (20.82%) (Table 1).

Table 1
Crystallographic statistics for the A3G191-384-2K3A crystal structure.

The crystal structure of A3G191-384-2K3A has a core α-β-α fold consistent with other cytidine deaminases (Chen et al., 2008; Holden et al., 2008; Prochnow et al., 2007). As seen with the other A3G-CTD structures (Harjes et al., 2009; Holden et al., 2008), this structure has a five-stranded β-sheet surrounded on both sides by six α-helices (Fig. 1). Secondary structural elements are numbered after the A3G191-384-2K3A NMR structure (PDB ID: 2KEM) (Harjes et al., 2009) so that consistent comparisons can be made between the structures. The second β-strand is discontinuous, as also observed in the wild-type A3G-CTD (Furukawa et al., 2009) and A3G-CTD-2K3A (Chen et al., 2008; Harjes et al., 2009) NMR structures. As previously observed, the catalytic zinc is coordinated directly by H257, C288, and C291, and indirectly by the catalytic residue E259 via a water molecule. Thus, in terms of the overall fold, this structure is similar to the previously published structures but many key specific differences exist.

Figure 1
Crystal structure of A3G191-384-2K3A with the α-helices highlighted in red and β-sheets highlighted in yellow, labeled as in (Harjes et al., 2009). The catalytic zinc ion is shown in cyan and the intermolecular zinc ion is depicted in ...

Crystal structure comparison

Comparison of A3G191-384-2K3A with the 3E1U crystal structure reveals numerous differences (highlighted in green and blue in Fig. 2A). Some of these differences are in loop regions but other variations include large regions of secondary structure, notably the β1-β2 region (M227-Q237), the β2′-α2 region (H248-G255), and the α2-β3 region (P267-D274). In the A3G191-384-2K3A crystal structure, the second β-strand is discontinuous, with residues L235-R239 forming a prominent “bulge”. This has been observed in both mutant and wild-type A3G-CTD NMR structures (Furukawa et al., 2009; Harjes et al., 2009; Chen et al., 2008).

Figure 2
(A) Major regions of structural differences between A3G191-384-2K3A and 3E1U highlighted in blue and green. Residues F268-D272, which were mis-traced in the 3E1U structure are highlighted in red. (B) NOE violations of more than 6 Å (red lines ...

Crystallography and NMR spectroscopy are complementary techniques with well-established methods for verifying the structural integrity of protein structures. PDB_REDO recently organized some of the verification of crystal structures in the Protein Data Bank (Joosten et al., 2009b; Joosten et al., 2009a; Sanderson, 2009). Unfortunately, the only other A3G-CTD crystal structure in the database was flagged as having discrepancies in the PDB_REDO database (PDB ID: 3E1U with R-factor 25.2%, R-free 26.7%, Resolution 2.3 Å). Many of the regions in the 3E1U structure (Holden et al., 2008) that differ from the NMR structures, are regions where the 3E1U structure’s experimental data are the weakest (α-carbon B-factors shown in Fig. 2C). The regions differing between these structures include β1-β2, β2′-α2 and α2-β3.

Evidence for ambiguity in the 3E1U structure is apparent in four ways: (1) 36 NOE restraints are violated in the 3E1U structure, by more than 6 Å, with respect to NMR structural data; including in the very well-ordered α-helical and β-sheet regions that are well-established by NMR (Fig. 2B and Experimental Procedures). (2) In addition to being flagged in PDB-REDO, analyses of the structure and experimental structure-factors of the 3E1U structure, downloaded from the Protein Data Bank (, revealed seven residues that were in unfavorable Ramachandran space (3 in disallowed and 4 in generously allowed regions). Most significantly, residues F268-D272 are completely mis-traced in the electron density, burying K270 away from the surface (Fig. 2D). These residues, located near the end of α2-helix, may be essential for the stability of the A3G-CTD active site because mutating W269 or L271 abrogates deaminase activity (Chen et al., 2008). When we re-fit this region in the 3E1U data, the final turn of α2-helix fit better within the electron density (Fig. 2D). Thus, the structure in this region converged towards the conformation observed in the NMR structures not the one originally modeled in the 3E1U structure. The electron density in two other regions, β1-β2 and β2′-α2, remained ambiguous, making refitting unattainable. (3) In contrast to these regions in the 3E1U structure, our crystal structure of A3G-191-384-2K3A (R-factor 16.57%, R-free 20.82%, Resolution 2.25 Å) is well-ordered. Our structure of A3G191-384-2K3A has only 9 NOE restraint violations to the NMR data; the α2-helix is well ordered; and the structure has lower α-carbon B-factors (Table 1) (Fig. 2). (4) Mass spectrometry analysis validates that the β1-β2 structure is conserved in full-length A3G. In contrast to the 3E1U structure, all NMR structures and the A3G191-384-2K3A crystal structure show that residues M227 and W232 are adjacent to each other in the two neighboring beta-strands (β1-β2). This was verified by introducing two cysteine mutations: M227C in β1 and W232C in β2, into both full-length A3G-2K3A (Fig. S1) and A3G (data not shown). Mass spectrometry determined that these two residues, M227C and W232C, form a disulfide bond and no peptides containing free M227C or W232C are detected in either construct. In the 3E1U structure, M227 and W232 are not adjacent, rather R226 and L235 (part of the putative “continuous β2” strand) face each other. Applying similar mass spectrometry analysis to full-length A3G, with mutations R226C and L235C, did not result in a detectable peptide containing a disulfide bond (data not shown).

Authors of the 3E1U structure write: “Therefore, an intact full-length β2 strand and the five-stranded β-sheet core is probably the feature of wild-type APOBEC3G-CD2 and all other APOBEC proteins.” (Holden et al., 2008). However, the A3G-191-384-2K3A crystal structure and the additional experimental data described above do not support this statement. These data suggest that the “bulge” observed between the β2-β2′ strands in NMR experiments and the A3G-191-384-2K3A crystal structure, is not merely an experimental artifact but an intrinsic feature of A3G-CTD structure.

Analysis of A3G191-384-2K3A crystal packing interfaces

The two molecules in the asymmetric unit in the crystal structure of A3G191-384-2K3A pack in such a way as to produce four major interfaces, all of which are the result of non-crystallographic symmetry, with surface areas of: 901 Å2, 604 Å2, 427 Å2 and 246 Å2, respectively (Figs. 36). Multiple sequence alignment of the A3G-CTD to ten other homologs: A3G-NTD, A3A, A3B-CTD/NTD, A3C, A3DE-NTD/CTD, A3F- NTD/CTD and A3H reveals that many of the residues contributing more than 30 Å2 to these interfaces are unique to A3G-CTD (Fig. 7).

Figure 3
Interface 1 between two A3G-CTD molecules in the asymmetric unit (A) Surface representation and (B) details of the largest interaction interface. Molecule A is shown in dark green, molecule B is shown in light green.
Figure 6
Interface 4 between two A3G-CTD molecules in the asymmetric unit (A) Surface representation and (B) detailed interactions. Same coloring scheme as in Figure 3.
Figure 7
Multiple sequence alignment of all human APOBEC3 protein sequences (Refseq) by their respective NTDs and CTDs aligned to the A3G-CTD sequence. Residues contributing more than 30 Å2 to the buried surface area on an inter-molecular interface are ...

The largest interaction interface (901 Å2) displays excellent shape complementarity as observed from the surface representation of interfacing molecules (Fig. 3A). Extensive surface contacts are observed, primarily between identical residues at the α1-loop-β1 from both molecules in the asymmetric unit. Twelve residues in each molecule contribute at least 30 Å2 to the interface, however, residues W211, R213 and Q318 together contribute over 450 Å2 of the interfacial area (Table S1) (Fig. 3B). None of these three residues are conserved across the other APOBEC3’s (Fig. 7). Three direct hydrogen bonds, nine water-mediated hydrogen bonds and one ionic interaction, occur across the entire interface (Table S2). All of these contacts verify the intimacy of this extensive interface. As the residues forming this extensive interface are not conserved among the other APOBEC3’s or the NTD of A3G (Fig. 7), this interface may be unique to A3G-CTD.

To assess the functional significance of this interface, a variant of A3G was made with the three amino acid substitutions W211A, R213A and R374E designed to profoundly disrupt the observed packing. This variant shows near undetectable levels (background) of DNA deaminase activity in vitro and abrogated anti-viral activity in the Vif-deficient HIV-1 reporter virus assay (Fig. 8A, 8B). In contrast, the active site A3G-E259Q variant is catalytically dead and unable to inhibit Vif-deficient HIV-1, in agreement with prior studies (Schumacher et al., 2008; Hache et al., 2008) (Fig. 8B). All of the variants studied have near normal cellular expression levels and incorporate into viral particles (Fig. 8C). Thus the residues that are integral to the first interface in the A3G191-384-2K3A crystal structure are essential to both A3G deaminase and anti-viral activity.

Figure 8
DNA deaminase activity and HIV-1 restriction data. (A) A graph showing the results of a DNA oligonucleotide-based deamination assay with the indicated A3G-GFP constructs (MOCK: GFP only; WT: A3G-GFP; E259Q: A3G-E259Q-GFP; INT1: interface 1 A3G-GFP mutant ...

The second largest interface (604 Å2) involves the β2′-loop-α2 residues 247–254 in each of the two molecules of the asymmetric unit (Fig. 4A, 4B). At this interface, eight residues of each molecule bury extensive surface area, with the largest surface area being buried by residues F252 and F268 (Table S1). This loop also coordinates an intermolecular zinc-binding site. H248 and H250 of one molecule in the asymmetric unit and the second molecule’s C261 and D264 (through a water-mediated hydrogen bond) coordinate an intermolecular zinc ion (Table S2). The combination of these four residues occurring simultaneously is unique to A3G-CTD amongst the APOBEC3’s (Fig. 7). An additional four hydrogen/water mediated bonds are formed within this interface. This zinc-coordinating interface may provide insights into how a metal mediated switch could specifically modulate A3G oligomerization.

Figure 4
Interface 2 between two A3G-CTD molecules in the asymmetric unit (A) Surface representation and (B) detailed interactions. Same coloring scheme as in Figure 3. (C) Interface 2 overview with coloring based on data from HSQC spectra. Blue residues: signal ...

NMR experiments support Zn2+ mediated oligomerization at this interface. Through titration of Zn2+ into isotope labeled A3G191-384-2K3A, loss of signal intensities and chemical shift perturbations were observed at 1mM Zn2+ (labeled blue, Fig. S2). The residues most affected are located in the β2′-loop-α2 and α2 regions. In fact, these residues in the A3G191-384-2K3A crystal structure (blue, Fig. 4C) map predominantly to the zinc-coordinating second interface, described above, indicating that this interface may contribute to zinc mediated oligomerization in solution.

To test the ability of zinc to modulate oligomerization a further series of NMR experiments was performed. A reference HSQC spectrum was taken at 50μM of Zn2+ concentration (Fig. 9A) then, Zn2+ concentration was increased to 1.25mM causing the disappearance of amide proton NMR signals (Fig. 9B). Next, 0.4 mM ethylenediaminetetraacetic acid (EDTA) was added to chelate the free Zn2+. This caused the signals within the NMR spectra to reappear (Fig. 9C). The similarity of chemical shifts, signal intensities and lineshapes of HSQC signals in Figs. 9A and 9C combine to suggest that zinc mediates the equilibrium of A3G-CTD between oligomeric (Fig. 9B) and monomeric states.

Figure 9
Zn2+ dependent A3G191-384-2K3A aggregation is reversible. HSQC spectra of A3G191-384-2K3A with (A) 50μM Zn2+, (B)1.25mM Zn2+, and (C)1.25mM Zn2+ / 0.4mM EDTA. Protein concentration was 300μM for all spectra

A variant designed to disrupt this zinc binding site in the second interface, H248A, H250A and C261A, shows normal expression levels and anti-viral activity (Fig. 8B, 8C). These observations demonstrate that zinc-mediated oligomerization may not be essential for A3G’s HIV-1 restriction activity. This variant is partly defective for DNA deaminase activity (Fig. 8A). This alteration in DNA deaminase activity, may be attributable to C261A alone (Chen et al., 2008), and was not significant enough to compromise A3G anti-viral activity. Thus the residues that are integral to the second interface in the A3G191-384-2K3A crystal structure appear to impact A3G deaminase activity.

The third interface (427 Å2) involves residues at N-terminal (β1-β2 strands) and C-terminal ends of A3G191-384-2K3A (Fig. 5A, 5B) (Table S1). Nine residues contribute extensively to the interface with Q354 burying the largest surface area at this interface followed by Q237, H228 and G355. In six of the eleven aligned APOBEC3 sequences (Q or E354)/G355 are conserved (Fig. 7) but not H228 or Q237. Four direct and three water mediated hydrogen bonds are formed at this interface. Specifically, Q237 and R238 at the β2-β2′ “bulge” of one molecule (molecule A), form intermolecular hydrogen bonds to N-terminal residues R194 (side chain disordered) and S196 with the other molecule (molecule B). In addition, Q354 in the α5–α6 loop (molecule B) forms a hydrogen bond with R226 in β1 (molecule A), this interaction is potentially conserved in APOBEC3A and APOBEC3B-CTD. This interface would correspond closest to the β2-β2 interface observed in APOBEC2, but is clearly not formed and would entail extensive rearrangement of the α5 and α6 helices and their connecting loop. Of the four major interfaces, the packing of the third interface is least complementary and mutations at this interface (H228A, V233R and L235R) do not disrupt A3G’s DNA deaminase or antiviral activities (Fig. 8).

Figure 5
Interface 3 between two A3G-CTD molecules in the asymmetric unit (A) Surface representation and (B) detailed interactions. Same coloring scheme as in Figure 3.

The final of the four major interfaces buries a total of 246 Å2 and involves substantial burial of six residues from each molecule and the formation of two hydrogen bonds and a salt bridge (Fig. 6A, 6B) (Tables S1, S2). This interface is primarily made up of the tops of the α5 and α6 helices. Interestingly, residues in α5 and α6 are also affected by the addition of Zn2+. These residues showed small chemical shift changes in HSQC spectra, suggesting that this region is involved in intermolecular interactions under a fast exchange (labeled yellow, Fig. 4C, Fig. S2). At this interface, residues Y340 and D365 contribute most to the buried interface area. These two residues are well conserved among the various APOBEC3’s (Fig. 7) Y340 is conserved in all but one and (D/E/Q)365 is conserved in eight of the eleven APOBEC3 sequences. Both of these residues and the less conserved S341 are involved in hydrogen bonding; a salt-bridge is observed between the moderately conserved K344 and D365. Although extensive sequence conservation is present, this interface is not as large as the others and a variant of A3G with mutations Y340A, S341A, K344A and D365A has normal DNA deaminase and HIV-1 restriction activities (Fig. 8).


In this study, the crystal structure of the catalytically active C-terminal domain of APOBEC3G was determined to 2.25 Å. This structure differs significantly from the previously published crystal structure of this domain (PDB ID: 3E1U) (Holden et al., 2008). Here we have also demonstrated that the 3E1U structure’s deposited experimental data do not unambiguously describe the structure deposited in the PDB. In contrast, our well-resolved crystal structure of A3G191-384-2K3A validates the published mutant and wild-type NMR structures (Chen et al., 2008; Furukawa et al., 2009; Harjes et al., 2009), both in respect to the β2-β2′ “bulge” and a longer α2-helix with the residues F268-D272 properly placed. Thus, this structure represents the first clearly resolved crystal structure of the catalytic domain of APOBEC3G.

In addition mass spectrometry data support the observed β1-β2 conformation and presence of a bulge between the β2-β2′ strands of the A3G catalytic domain. We verified the register of the β1-β2 secondary structure elements by probing the proximity of key residues in this region through carefully engineered disulfide linkages. These results thereby confirm the structural integrity of the β1-β2 conformation observed in all NMR structures and the A3G191-384-2K3A crystal structure described here (Chen et al., 2008; Furukawa et al., 2009; Harjes et al., 2009).

The unique interfaces observed between the asymmetric units of this crystal structure also lend insights into the potential role of oligomerization in APOBEC3G function. Full-length APOBEC3G sequence displays pseudo-symmetry (Fig. 7), yet none of the largest interfaces observed in the crystal structure are conserved between the N- and C- terminal halves of the protein sequence nor are do they position the termini in a manner that could represent the full length enzyme. This is also consistent with the recent model proposed for the full-length A3G holoenzyme (Harjes et al., 2009).

Beyond the full-length A3G enzyme, oligomerization —often dimerization or tetramerization—is a trend among the known deaminases. Escherichia coli cytidine deaminase and Staphylococcus aureus TadA form dimers; Bacillus subtilis cytidine deaminase, yeast CDD1, and human cytidine deaminase form tetramers; and human ADAR1 and ADAR2 are homodimers (Betts et al., 1994; Carlow et al., 1999; Chung et al., 2005; Johansson et al., 2002; Losey et al., 2006; Navaratnam and Sarwar, 2006; Valente and Nishikura, 2007; Xie et al., 2004). Multiple studies have shown that A3G can exist in states ranging from monomer to megadalton complexes (Bennett et al., 2008; Chiu et al., 2006; 1994; Jarmuz et al., 2002; Wedekind et al., 2006). The importance of A3G oligomerization to HIV-1 suppression is controversial; some studies conclude that oligomerization is dispensable (Opi et al., 2006) while others claim that oligomerization is essential (Huthoff et al., 2009). The present study adds new insights into the potential roles of these interfaces for anti-viral and deaminase activities.

The surface(s) of A3G that forms these oligomers still remain unclear. SAXS, co-immunoprecipitation and FRET studies have implicated the C-terminal half of A3G in oligomerization (Bennett et al., 2008; Wedekind et al., 2006). This work potentially contrasts with more recent co-immunoprecipitation studies demonstrating that N-terminal residues are required for RNA-mediated dimerization (Huthoff et al., 2009; Friew et al., 2009). Possibly both sets of studies have some merit and two distinct surfaces of A3G contribute to oligomerization.

Based on our analyses and data, we hypothesize that there are a variety of potential surfaces for oligomerization. The interactions observed at the two most extensive interfaces, the α1-loop-β1 (Fig. 3) and the β2′-loop-α2 (Fig. 4) which coordinates a zinc ion, represent unique A3G homo-oligomerization interfaces, while the tops of the α5 and α6 helices (Fig. 6) could modulate both homo- and hetero- oligomerization of the different APOBEC3 family members.

A combination of solution NMR and the impact of variants designed to disrupt the interfaces support the possibility that three of these interfaces may have functional significance. Mutations introduced to disrupt the first interface reduce both DNA deaminase activity and anti-viral activity of A3G (Fig. 8). However whether this is change in activity is due to a change in oligomerization or direct disruption in nucleic acid binding affinity remains to be determined. While mutations to disrupt the other interfaces do not alter the anti-viral activity of A3G, these interfaces still may impact oligomerization. The solution NMR data demonstrates a clear relationship between zinc-mediated EDTA-reversible oligomerization and specific chemical shifts of residues most strongly in the second interface that binds the intermolecular zinc but secondarily to the fourth interface (Fig. 9 and Fig. S2). The functional significance of the zinc mediated oligomerization remains to be elucidated. The interfaces observed in this new A3G-CTD crystal structure may therefore have important functional implications in the role of oligomerization for the activity of A3G and will guide future studies.

Experimental Procedures

Protein expression and purification

BL21(DE3)-CodonPlus(RIL) (Stratagene, La Jolla, CA) bacterial cells, transformed with the pGEX6P1-A3G191-384-2K3A vector (Chen et al., 2007; Harjes et al., 2009), were grown to an O.D.600 of 0.6 [TB media, 100 μg/ml Ampicillin and 34 μg/ml Chloramphenicol]. They were induced overnight [0.5 mM IPTG, 18°C] to express GST-A3G191-384-2K3A. The cells were then harvested by centrifugation [5000 rpm, 15 min, 4°C] and re-suspended in lysis buffer [50 mM Na2HPO4/NaH2PO4 (pH 7.4) and protease inhibitor (Roche, Basel, Switzerland)]. Resuspension was followed by lysis with a cell disruptor. The lysate was centrifuged [43,000×g, 20 min, 4°C] and the supernatant incubated overnight with Glutathione-Sepharose (GE Healthcare, Piscataway, NJ) beads. The beads were washed with lysis buffer and incubated with PreScission Protease (GE Healthcare, Piscataway, NJ) in cleavage buffer [50 mM Na2HPO4/NaH2PO4 (pH 7.4), 0.005% Tween20, and 1 mM dithiothreitol]. A3G191-384-2K3A in the supernatant was further purified by FPLC (Pharmacia/GE Healthcare, Uppsala, Sweden) using a Superdex-75 size-exclusion column.

Protein crystallization

Purified A3G191-384-2K3A protein was exchanged into crystallization buffer [50 mM HEPES pH 7.5, 1 mM tris(2-carboxyethyl)phosphine, 50 μM zinc acetate, and 150 mM NaCl] with a PD-10 column (GE Healthcare, Piscataway, NJ). The protein was concentrated to 10 mg/ml using Amicon centrifugal filters (Millipore, Billerica, MA). Crystals were grown by the hanging-drop/vapor-diffusion method on greased VDX plates (Hampton Research, Aliso Viejo, CA) at 4°C with the reservoir solution [100 mM HEPES (pH 7.5), 10% Polyethylene glycol 4000, and 100 mM magnesium chloride].

Data collection and processing

Diffraction data for the A3G191-384-2K3A protein crystals were collected by synchrotron radiation at Argonne National Laboratory (Advanced Photon Source, Chicago, IL) BioCARS 14-BMC under cryogenic conditions. Diffraction images were indexed and scaled using the program HKL2000 (Otwinowski and Minor, 1997); the structure was solved in P212121 space group at 2.25 Å.

Structure solution and refinement

The molecular replacement solution, calculated by PHASER (McCoy et al., 2007) with the final translation function Z-score of 41.1, contained two molecules in the asymmetric unit. Phases were subsequently improved by building solvent molecules using ARP/wARP (Langer et al., 2008). Refinement was carried out with REFMAC5 (Murshudov et al., 1997) in the CCP4 suite (Collaborative-Computational-Project, 1994), with cycles of rigid-body and restrained refinement; Non-Crystallographic Symmetry (NCS) restraints were applied in the first two cycles and TLS parameters (Painter and Merritt, 2006) were used in subsequent refinement cycles. A free R-value with 5% of the data was used to limit the possibility of over-refinement. Electron density was viewed by the program COOT (Emsley and Cowtan, 2004) and was used for model optimization.

Structure comparison and analysis

Graphical visualization and analysis was achieved using PyMOL (DeLano Scientific LLC, Palo Alto CA) (DeLano, 2002). The interaction interfaces formed between molecules in the A3G191-384-2K3A crystal were analyzed by the Protein interfaces, surfaces and assemblies service PISA at European Bioinformatics Institute (Krissinel and Henrick, 2007). The NOE violations were calculated using accept.inp protocol of CNS (Brünger et al., 1998). The A3G-CTD crystal structure (PDB ID: 3E1U) was used as the input coordinate file and the NOE files used to calculate the A3G191-384-2K3A NMR structure (PDB ID: 2KEM), were set as NOE distance restraints. The algorithm PRANK, as implemented in the software PRANKSTER, performed the multiple sequence alignments, with the +F option and default values (Loytynoja and Goldman, 2005). All human APOBEC3 sequences were downloaded from the NCBI Refseq database (APOBEC3A: NP_663745.1; APOBEC3B: NP_004891.3; APOBEC3C: NP_055323.2; APOBEC3DE: NP_689639.2; APOBEC3F: NP_660341.2; APOBEC3G: NP_068594.1; APOBEC3H: NP_861438.1) (Pruitt et al., 2007).

DNA deaminase activity assays

The A3G-GFP and A3G-E259Q-GFP expression constructs have been described (Stenglein et al., 2008; Schumacher et al., 2008). All derivatives were constructed by combinatorial or sequential site-directed mutagenesis (Stratagene), with primer sequences available on request. The indicated A3G-GFP or interface mutant expression constructs were transfected into HEK-293 cells using Fugene (Roche). Cell lysates were prepared and DNA deaminase activities were measured as described (Thielen et al., 2007). The sequence of the single-strand DNA substrate was: 5′-(6-FAM)-AAA-TTC-TAA-TAG-ATA-ATG-TGA-(TAMRA).

HIV-1 infectivity studies and immunoblots

These were performed as described (Harjes et al., 2009). HIV-GFP reporter viruses were produced by Fugene-mediated transfection (Roche) of 293T cells with a five plasmid cocktail (Liddament et al., 2004). The HIV-GFP proviral plasmid CS-CG, the Gag-Pol expression plasmid, the Rev expression plasmid, and the VSV-G envelope expression plasmid constituted 0.9 μg of the cocktail, and the vector control or the A3G expression plasmid constituted the remaining 0.1 μg. Viruses containing supernatants were harvested 48 h post transfection by filtering cell-free supernatants (0.22-μm PVDF; Millipore) and pelleting by centrifugation (20,000 g, 2 hrs). The resulting viral pellet was resuspended directly in SDS gel loading buffer, fractionated by SDS-PAGE, transferred to a PVDF membrane (Millipore), and probed with an anti-GFP antibody JL-8 (Invitrogen) to detect the A3G-GFP constructs. An anti-p24 monoclonal antibody (Simon et al., 1997) provided by M. Malim through the National Institutes of Health AIDS Research and Reference Reagent Program was used as loading control. Both monoclonal antibodies were detected using a horseradish-peroxidase-conjugated goat anti-mouse IgG serum (Bio-Rad), followed by chemiluminescent imaging (Roche). After harvesting the viral supernatants, A3G levels in virus-producing cells were monitored by extracting soluble proteins with RIPA buffer (1 h, 4 °C, gentle rotation), removing particulates by centrifugation (10 min, 20,000 g), and immunoblotting, as described above. An anti-tubulin monoclonal antibody (Covance) was used as cellular lysate loading control.

NMR Zn2+ titration experiments

15N-labeled A3G191-384-2K3A protein was expressed and purified (in 1mM DTT, 200mM NaCl, 0.005% Tween 20 (Fisher scientific) and 50mM tris(hydroxymethyl)aminomethane buffer, pH 8.0) as described in (Harjes et al., 2009). ZnCl2 was titrated into 300 μM 15N-labeled A3G191-384-2K3A at concentrations of 50μM, 200 μM, 400 μM, 600 μM, 800 μM, 1mM and 1.25mM. A heteronuclear single quantum coherence (HSQC) spectrum was recorded at each Zn2+ concentration, which enabled specific amino acid chemical shift perturbations to be detected. Bruker 700MHz equipped with a cryoprobe was used to collect all NMR spectra.

Mass spectrometry of cysteine mutants

Cysteine substitution variants were constructed using QuikChange protocol (Stratagene) and verified by DNA sequencing. A3G191-384-2K3A, A3G191-384-2K3A-M227C-W232C, A3G1-384-M227C-W232C and A3G1-384-R226C-L235C were produced using the same procedure as described in the protein expression and purification. Proteins were isolated from sodium dodecyl sulfate (SDS) polyacrylamide gel after Coomassie Brilliant Blue (CBB) staining. Isolated protein bands were digested using trypsin and subjected to mass spectrometry (MS) at the Taplin Biological Mass Spectrometry Facility (Harvard Medical School). Both high resolution full MS scans and high resolution MS/MS scans were performed using LTQ-FT and LTQ-Orbitrap (Thermo Electron) to verify the peptide.

Supplementary Material


The authors would like to thank K. Romano, S. Mittal, and M. Kolli for assistance with data collection; W. Royer and R. Bandaranayake for assistance with initial data processing, and Dr. Ross Tomaino for assistance with Mass spectrometry. Use of the Advanced Photon Source was supported by the U.S. Department of Energy, Basic Energy Sciences, Office of Science, under Contract No. DE-AC02-06CH11357. Use of the BioCARS Sector 14 was supported by the National Institutes of Health, National Center for Research Resources, under grant number RR007707.

This work was supported by grants from the National Institutes of Health (A1067021 and GM65347 to C.A.S; AI073167 to H.M. and AI064046 to R.S.H.). The University of Minnesota Supercomputing institute and NMR Core (NSF BIR-961477) provided key instrumentation. All Mass spectrometry analyses were performed at the Taplin Biological Mass Spectrometry Facility of Harvard Medical School.


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