Title: Structural analysis of membrane-bound retrovirus capsid proteins
Abstract: Article15 March 1997free access Structural analysis of membrane-bound retrovirus capsid proteins Eric Barklis Corresponding Author Eric Barklis Vollum Institute and Department of Microbiology, Portland, OR, 97201 USAE.Barklis, J.McDermott, S.Wilkens and D.Thompson contributed equally to this work Search for more papers by this author Jason McDermott Jason McDermott Vollum Institute and Department of Microbiology, Portland, OR, 97201 USAE.Barklis, J.McDermott, S.Wilkens and D.Thompson contributed equally to this work Search for more papers by this author Stephan Wilkens Stephan Wilkens Institute of Molecular Biology, Eugene, OR, 97403 USA Search for more papers by this author Eric Schabtach Eric Schabtach Institute of Neuroscience, University of Oregon, Eugene, OR, 97403 USAE.Barklis, J.McDermott, S.Wilkens and D.Thompson contributed equally to this work Search for more papers by this author M.F. Schmid M.F. Schmid Verna and Marrs McLean Department of Biochemistry and W.M.Keck Center for Computational Biology, Baylor College of Medicine, Houston, TX, 77030 USA Search for more papers by this author Stephen Fuller Stephen Fuller Structural Biology Programme, European Molecular Biology Laboratory, D-69012 Heidelberg, Germany Search for more papers by this author Sonya Karanjia Sonya Karanjia Vollum Institute and Department of Microbiology, Portland, OR, 97201 USA Search for more papers by this author Zachary Love Zachary Love Vollum Institute and Department of Microbiology, Portland, OR, 97201 USA Search for more papers by this author Russell Jones Russell Jones Department of Pathology, Oregon Health Sciences University, Portland, OR, 97201 USA Search for more papers by this author Yuanjui Rui Yuanjui Rui Department of Chemistry, Purdue University, West Lafayette, IN, 47907 USA Search for more papers by this author Xiumin Zhao Xiumin Zhao Department of Chemistry, Purdue University, West Lafayette, IN, 47907 USA Search for more papers by this author David Thompson David Thompson Department of Chemistry, Purdue University, West Lafayette, IN, 47907 USAE.Barklis, J.McDermott, S.Wilkens and D.Thompson contributed equally to this work Search for more papers by this author Eric Barklis Corresponding Author Eric Barklis Vollum Institute and Department of Microbiology, Portland, OR, 97201 USAE.Barklis, J.McDermott, S.Wilkens and D.Thompson contributed equally to this work Search for more papers by this author Jason McDermott Jason McDermott Vollum Institute and Department of Microbiology, Portland, OR, 97201 USAE.Barklis, J.McDermott, S.Wilkens and D.Thompson contributed equally to this work Search for more papers by this author Stephan Wilkens Stephan Wilkens Institute of Molecular Biology, Eugene, OR, 97403 USA Search for more papers by this author Eric Schabtach Eric Schabtach Institute of Neuroscience, University of Oregon, Eugene, OR, 97403 USAE.Barklis, J.McDermott, S.Wilkens and D.Thompson contributed equally to this work Search for more papers by this author M.F. Schmid M.F. Schmid Verna and Marrs McLean Department of Biochemistry and W.M.Keck Center for Computational Biology, Baylor College of Medicine, Houston, TX, 77030 USA Search for more papers by this author Stephen Fuller Stephen Fuller Structural Biology Programme, European Molecular Biology Laboratory, D-69012 Heidelberg, Germany Search for more papers by this author Sonya Karanjia Sonya Karanjia Vollum Institute and Department of Microbiology, Portland, OR, 97201 USA Search for more papers by this author Zachary Love Zachary Love Vollum Institute and Department of Microbiology, Portland, OR, 97201 USA Search for more papers by this author Russell Jones Russell Jones Department of Pathology, Oregon Health Sciences University, Portland, OR, 97201 USA Search for more papers by this author Yuanjui Rui Yuanjui Rui Department of Chemistry, Purdue University, West Lafayette, IN, 47907 USA Search for more papers by this author Xiumin Zhao Xiumin Zhao Department of Chemistry, Purdue University, West Lafayette, IN, 47907 USA Search for more papers by this author David Thompson David Thompson Department of Chemistry, Purdue University, West Lafayette, IN, 47907 USAE.Barklis, J.McDermott, S.Wilkens and D.Thompson contributed equally to this work Search for more papers by this author Author Information Eric Barklis 1, Jason McDermott1, Stephan Wilkens2, Eric Schabtach3, M.F. Schmid4, Stephen Fuller5, Sonya Karanjia1, Zachary Love1, Russell Jones6, Yuanjui Rui7, Xiumin Zhao7 and David Thompson7 1Vollum Institute and Department of Microbiology, Portland, OR, 97201 USA 2Institute of Molecular Biology, Eugene, OR, 97403 USA 3Institute of Neuroscience, University of Oregon, Eugene, OR, 97403 USA 4Verna and Marrs McLean Department of Biochemistry and W.M.Keck Center for Computational Biology, Baylor College of Medicine, Houston, TX, 77030 USA 5Structural Biology Programme, European Molecular Biology Laboratory, D-69012 Heidelberg, Germany 6Department of Pathology, Oregon Health Sciences University, Portland, OR, 97201 USA 7Department of Chemistry, Purdue University, West Lafayette, IN, 47907 USA The EMBO Journal (1997)16:1199-1213https://doi.org/10.1093/emboj/16.6.1199 PDFDownload PDF of article text and main figures. ToolsAdd to favoritesDownload CitationsTrack CitationsPermissions ShareFacebookTwitterLinked InMendeleyWechatReddit Figures & Info We have developed a system for analysis of histidine-tagged (His-tagged) retrovirus core (Gag) proteins, assembled in vitro on lipid monolayers consisting of egg phosphatidylcholine (PC) plus the novel lipid DHGN. DHGN was shown to chelate nickel by atomic absorption spectrometry, and DHGN-containing monolayers specifically bound gold conjugates of His-tagged proteins. Using PC+DHGN monolayers, we examined membrane-bound arrays of an N-terminal His-tagged Moloney murine leukemia virus (M-MuLV) capsid (CA) protein, His-MoCA, and in vivo studies suggest that in vitro-derived His-MoCA arrays reflect some of the Gag protein interactions which occur in assembling virus particles. The His-MoCA proteins formed extensive two-dimensional (2D) protein crystals, with reflections out to 9.5 Å resolution. The image-analyzed 2D projection of His-MoCA arrays revealed a distinct cage-like network. The asymmetry of the individual building blocks of the network led to the formation of two types of hexamer rings, surrounding protein-free cage holes. These results predict that Gag hexamers constitute a retrovirus core substructure, and that cage hole sizes define an exclusion limit for entry of retrovirus envelope proteins, or other plasma membrane proteins, into virus particles. We believe that the 2D crystallization method will permit the detailed analysis of retroviral Gag proteins and other His-tagged proteins. Introduction A number of cellular and viral components are present in all infectious mammalian retroviruses. Such viruses are surrounded by host-derived lipid membrane envelopes, which carry receptor-binding/fusion proteins, the surface (SU) and transmembrane (TM) proteins, that are encoded by the retroviral env genes (Weiss et al., 1984; Leis et al., 1988). Within each mammalian retrovirus particle are two copies of the viral RNA genome with associated, cellularly derived tRNAs, which serve as primers during reverse transcription. Virus interiors also contain 1000–5000 copies of the viral Gag (group-specific antigen) proteins, and 10–100 copies of the viral pol gene products including the viral protease (PR), reverse transcriptase (RT), RNaseH and integrase (IN) (Weiss et al., 1984). Components of C-type retroviruses, such as Moloney murine leukemia virus (M-MuLV), and lentiviruses, such as the human immunodeficiency virus (HIV), are delivered for assembly at the plasma membranes of infected cells (Weiss et al., 1984), although assembly can occur at other intracellular membrane locations (Hansen et al., 1990; Faecke et al., 1993; Wang et al., 1993). The one viral component that has been shown to be necessary and sufficient for C-type and lentivirus particle assembly is the Gag protein, which is synthesized as a precursor polyprotein (Prgag), and normally cleaved into mature processed Gag proteins by the viral PR during or after budding. The proteolytic processing of Prgag is required for infectivity, and results in a major change in the virus core, in which electron-dense material adjacent to the periphery of immature particles reorganizes into central spherical, cylindrical or cone-shaped structures (Weiss et al., 1984; Stewart et al., 1990). Three mature Gag proteins, matrix (MA), nucleocapsid (NC) and capsid (CA), are present in both C-type viruses and lentiviruses. During biosynthesis, MA is myristylated at its amino-terminus, and this fatty acid modification is necessary for membrane association of Prgag proteins (Rein et al., 1986). Genetic experiments have demonstrated that the HIV-1 matrix protein also interacts with the HIV Env protein complex (Yu et al., 1992; Faecke et al., 1993; Wang et al., 1993). However, the central 80–90% of HIV MA can be deleted without blocking assembly of virus particles (Faecke et al., 1993) that are conditionally infectious (Wang et al., 1993). At or near the C-termini of lentivirus or C-type retrovirus, Prgag proteins are Cys–His, zinc finger-containing NC domains. The NC domains have RNA-binding capabilities, but it is not clear whether NC completely accounts for the specificity of RNA encapsidation into virus particles (Zhang and Barklis, 1995). While MA interacts with viral membranes and Env proteins, and NC interacts with the viral RNA, the Gag CA domains appear to establish interprotein contacts that are essential to the oligomerization of Prgag proteins, and assembly of Gag–Pol and Gag fusion proteins into virions (Hansen et al., 1990; Jones et al., 1990; Chazal et al., 1994; Mammano et al., 1994; Wang et al., 1994; Craven et al., 1995; Hansen and Barklis, 1995; Srinivasakumar et al., 1995; McDermott et al., 1996). Despite the apparent conservation of retrovirus capsid protein function, only a 20–30 residue section in the C-terminal half of CA, called the major homology region (MHR), is well conserved at the primary sequence level (Mammano et al., 1994; Craven et al., 1995; Hansen and Barklis, 1995; Clish et al., 1996; McDermott et al., 1996). In addition to the three conserved mature Gag proteins (MA, CA and NC), M-MuLV encodes a p12 protein between the MA and CA domains, and HIV encodes a p6 domain at the C-terminus of its Prgag: while these domains are essential to the infectivities of their respective viruses, they are not absolutely required for virus particle assembly (Crawford and Goff, 1984; Spearman et al., 1994; Hansen and Barklis, 1995). Three-dimensional (3D) structures of HIV MA (Matthews et al., 1994; Hill et al., 1996) and NC (Morrelet et al., 1992) are available, as are partial structures of HIV CA (Clish et al., 1996; Gitti et al., 1996; Momany et al., 1996). However, despite observations of preferred interprotein contacts in 3D crystals (Hill et al., 1996), it is unclear how the individual Gag proteins fit into the immature or mature virus particle structures. Ordinarily, retrovirus structure analysis might proceed by X-ray diffraction analysis of 3D crystals of virus, or by electron microscopy (EM) and image analysis of homogeneous preparations of virus particles. Unfortunately, these approaches have proven difficult, since the virions do not crystallize, preparations do not appear homogeneous, and it is unclear whether mature or immature virus cores have helical or icosohedral symmetry, which would be a great help in image reconstruction (Choi et al., 1991; Fuller et al., 1995, 1996). In the absence of homogeneous, in vivo-derived virus preparations, several groups have undertaken the analysis of virus-like particles formed in vitro from Prgag proteins (Nermut et al., 1994; Klikova et al., 1995) or their derivatives (Ehrlich et al., 1992; Campbell and Vogt, 1995; J.McDermott and E.Barklis, unpublished observations). Such studies have shown that Gag-derived monomer proteins can assemble to form higher order structures, but, for the most part, resolution has been such that the nature of Gag interprotein contacts remains unclear. Nevertheless one analysis of HIV Prgag proteins assembled at the plasma membranes of baculovirus vector-infected cells provided sufficient resolution to suggest that these proteins form a fullerene cage-like network at the membrane (Nermut et al., 1994). Because of the difficulties with traditional X-ray and EM approaches, and the current limited resolution available via analysis of standard in vitro assembly products, we have adapted a lipid monolayer approach (Uzgiris and Kornberg, 1983; Darst et al., 1991a,b; Celia et al., 1994; Olofsson et al., 1994; Avila-Sakar and Chiu, 1996) to determine how Gag proteins may associate with each other at a membrane. Our approach (Zhao et al., 1994), outlined in Figure 1, has been to employ a nickel-chelating lipid to facilitate the two dimensional (2D) crystallization of N–terminal histidine-tagged (His-tagged) Gag proteins at a lipid monolayer. This approach recently has been used, with modifications, to examine a variety of interfacial lipid–protein interactions (Schmitt et al., 1994; Zhao et al., 1994; Dietrich et al., 1995, 1996; Kubalek et al., 1995; Ng et al., 1995; Frey et al., 1996), and can be used to obtain protein 3D structures, using image analysis resources developed over the past two decades (Unwin and Henderson, 1975; Fuller et al., 1979; Baldwin et al., 1988; Frank et al., 1988; Henderson et al., 1990; Schmid et al., 1993; Kuhlbrandt et al., 1994). The monolayer technique described in Figure 1 appears ideally suited for the analysis of retroviral Gag proteins, since their natural function is to oligomerize at the face of a membrane. Furthermore, at least for HIV, only the membrane-anchoring activity of the amino-terminal Gag matrix domain is required for virus particle assembly (Facke et al., 1993; Wang et al., 1993). In our current study, we have substituted a His-tag for the membrane-anchoring myristate moiety of the Gag protein, and have examined crystalline arrays of a His-tagged M-MuLV capsid protein (His-MoCA) formed on a monolayer of egg phosphatidyl choline (PC) and a novel nickel-chelating lipid, 1,2-di-O-hexadecyl-sn-glycero-3-[1′-(2″-R-hydroxy-3′-N-(5-amino-1-carboxypentyl)-iminodiacetic acid) propyl ether] (DHGN). The His-MoCA protein forms 2D crystals with reflections at 9.5 Å resolution, and can be classified as either orthorhombic (a= 79.6 Å, b= 137.5 Å, γ = 90°) or hexagonal (a= b= 79.7 Å, γ = 60°). Image reconstruction shows a cage-like array of proteins, similar to that seen for HIV-1 Prgag proteins (Nermut et al., 1994). In 2D projections, the putative His-MoCA monomers form two different types of hexagonal units surrounding two different protein-free cage holes: a circular hole, with a diameter of 19.2 Å; and a triangular cage hole (length 28.0 Å, width 23.2 Å). Our results permit specific predictions concerning retrovirus particle structure and capsid protein interactions with MA, NC and Env proteins. Figure 1.Model for forming histidine-tagged protein arrays on a monolayer of nickel-chelating lipids at the air–buffer interface. Shown is a model, based on previous lipid monolayer studies (Uzgiris and Kornberg, 1983; Darst et al., 1991a,b; Celia et al., 1994; Olofsson et al., 1994), in which His-tagged proteins bind to nickel-chelating lipids at the air–buffer interface. For our purposes, the proteins are N-terminally His-tagged retrovirus Gag proteins and the nickel-chelating lipid is DHGN, a dialkylglycerol derivative carrying an NTA (Hochuli et al., 1987) headgroup. Analysis of 2D protein arrays, following previously established methods (Unwin and Henderson, 1975; Fuller et al., 1979; Schmid et al., 1993), involves adhering arrays to EM grids; photography of stained or unstained protein crystals at 20 000–100 000× magnification; evaluation of 2D arrays by optical diffraction of EM negatives; digitization of negatives; and image processing to obtain structural details. In many circumstances, resolution is in the 10–30 Å range, sufficient for observation of protein–protein interactions, and in some cases atomic resolution has been achieved (Baldwin et al., 1988; Henderson et al., 1990; Kuhlbrandt et al., 1994; Avila-Sakar and Chiu, 1996). Download figure Download PowerPoint Results Characterization of the nickel-chelating lipid, DHGN To foster the development of a lipid monolayer approach for the analysis of retrovirus Gag protein interactions, we designed a nickel-chelating lipid to serve as a ligand to His-tagged Gag proteins. Our objective was to combine the nickel-chelating nitrilotriacetic acid (NTA; Hochuli et al., 1987) group with an activated diacyl glycerol derivative (Thompson et al., 1994). For this purpose, a convergent synthesis scheme was developed (Figure 2; Materials and methods), where compounds 3 and 5 (Figure 2) served as immediate precursors to DHGN (Figure 2, compound 6; Zhao et al., 1994). The crude DHGN product was purified by two rounds of silica gel chromatography, producing DHGN, whose structure was confirmed by IR and NMR spectroscopy (Materials and methods). Figure 2.Synthesis of the nickel-chelating lipid, DHGN. DHGN [compound 6; 1,2-di-O-hexadecyl-sn-glycero-3-(1′-(2″-R-hydroxy-3′-N-(5-amino-1-carboxypentyl)-iminodiacetic acid) propyl ether] was prepared from starting materials Nϵ-benzyloxycarbonyl-L-lysine (compound 1) and 1,2-di-O-hexadecyl-sn-glycerol (compound 4) in a multistep synthesis. Compound 4 was converted to 1,2-di-O-hexadecyl-sn-glyceryl-3-R-glycidyl ether (compound 5) by alkylation with R-glycidyl-3-nitrobenzenesulfonate. Compound 3 [N-(5-amino-1-carboxypentyl)-iminodiacetic acid] was prepared from compound 1 as described previously (Hochuli et al., 1987) by acetylation with bromoacetic acid to give N-(5-benzyloxycarbonylamino-1-carboxypentyl)-iminodiacetic acid (compound 2), which was hydrogenated in the presence of Pd-C catalyst to yield the compound 3 intermediate. DHGN was generated by epoxide ring opening of compound 5 via nucleophilic attack with compound 3. The crude DHGN product was purified by silica gel chromatography and the structure confirmed by IR and NMR spectroscopy. Download figure Download PowerPoint While the presence of the NTA headgroup on DHGN suggested that the lipid should chelate nickel, it was necessary to test this assumption directly. To do so without excessive consumption of DHGN, we resorted to atomic absorption spectrometry using an instrument equipped with a graphite furnace, capable of detecting micromolar nickel concentrations of 20 μl samples (Materials and methods). For analysis of nickel binding, a simple extraction assay was employed. As shown in Figure 3A, limiting amounts of nickel sulfate in water were mixed with lipids in chloroform and, after phase separation, nickel levels were measured in each phase. When this protocol was used with control lipids OG (octyl glucoside) and DHP (dihexadecyl phosphate), Ni2+ remained in the aqueous phase (Figure 3B). In contrast, DHGN chelated nickel ions, resulting in their extraction to the organic phase. As expected, EDTA, which blocks His tag protein binding to NTA-resin, effectively competed with DHGN for Ni2+, inhibiting DHGN-mediated extraction of nickel. Figure 3.Atomic absorption analysis of nickel chelation to lipids. (A) Nickel was measured with a Perkin Elmer 603 atomic absorption spectrometer equipped with an HGA-400 programmer and a graphite furnace. For analysis of nickel binding to lipids, 250 μM lipid solutions in 0.1 ml of chloroform were overlaid with an equal volume of 20 μM nickel sulfate in distilled water. Samples were rocked for 1–18 h at 25°C, after which phases were separated by low speed centrifugation. Triplicate 20 μl samples from aqueous (top) and organic (bottom) phases were measured and compared with nickel sulfate standards in distilled water to determine the nickel concentrations in each phase. (B) Nickel concentrations in aqueous (white) and organic (black) phases are as shown. In one experiment, EDTA was added to the aqueous phase to a final concentration of 50 mM. Neither EDTA nor any of the stock lipid solutions showed any nickel content above backround prior to the phase fractionation experiments. Download figure Download PowerPoint The above experiments indicated that DHGN chelates Ni2+, but did not permit an assessment of whether DHGN, charged with nickel, was capable of binding His-tagged proteins. To test this, we modified the approach illustrated in Figure 1 by using His-tagged or untagged proteins conjugated to gold particles, rather than using free proteins in the buffer subphase. By this method, we reasoned it would be possible to quantitate protein binding to monolayers via gold particle counts. Thus, using standard procedures (Geoghegan and Ackerman, 1977; Horisberger and Clerc, 1985; see Materials and methods), gold conjugates were prepared to His-tagged activating transcription factor (His-ATF–gold) and His-tagged cAMP-responsive element modulator (His-CREM–gold). As a control, we conjugated bovine serum albumin (BSA–gold) and, to examine potential non-specific binding of a basic protein, cytochrome c (cytoC–gold; pI = 10.4) also was conjugated. Total gold particle conjugate concentrations were determined as described in Materials and methods, while binding to lipid monolayers was assessed by EM of glucose-embedded monolayers after binding incubations and washes (see Materials and methods). An example of our results is shown in Figure 4, where His-CREM–gold was incubated beneath a monolayer of PC only (Figure 4A) or PC plus DHGN (Figure 4B). As illustrated, on PC-only monolayers, the His-CREM–gold conjugate washed to the edges of the lacy carbon support. In contrast, when lipid monolayers consisted of PC plus DHGN, gold particles were observed over the surface of the monolayers, suggesting a specific binding of the His-tagged moiety to the DHGN headgroup. This interpretation was substantiated when gold particle counts were performed (Table I). All conjugates showed total gold counts of 100–300 particles/nl, and PC monolayer binding of 3–32 particles/1000 nm2. However, binding of gold-labeled, His-tagged proteins to DHGN-containing monolayers was enhanced at least 10-fold, while binding of control conjugates was unimproved. These results support the notion that nickel-bound DHGN serves as a ligand for His-tagged proteins. Figure 4.Protein–gold conjugate binding to lipid monolayers. Histidine-tagged CREM proteins (His-CREM), conjugated to 20 nm gold particles as described in Materials and methods, were incubated under standard conditions beneath lipid monolayers composed of PC only (A) or PC plus nickel-charged DHGN (B). After incubations, monolayers were lifted onto lacy carbon grids, rinsed for 30 s in 1% glucose, dried, and viewed and photographed on the Philips 301 TEM. Carbon supports are clearly visible as the broad dark lines; monolayers are visible with occasional holes caused by beam damage, while gold particles appear as black dots. Note that with either monolayer, gold particles adhere or are washed to the carbon support regions; however, gold-conjugated His-CREM particles are present on the monolayer only when DHGN is present (B). The black bar in (A) indicates 10 nm. Download figure Download PowerPoint Table 1. Binding of untagged and His-tagged protein–gold conjugates to control and DHGN-containing lipid monolayers Protein Total particles per 10 nl Particles bound per 1000 nm2 PC only PC+DHGN BSA 198 32 22 CytoC 163 6 1 His-CREM 228 19 369 His-ATF 117 12 117 Cytochrome C (cytoC), bovine serum albumin (BSA) and histidine-tagged proteins His-CREM, and His-ATF were conjugated to 20 nm gold particles, purified and used to assess binding to lipid monolayers. Total gold particle numbers were quantitated by drying 2 μl of 1:10 dilutions of gold conjugate stocks onto carbon–formvar grids, and counting all gold particles in 200 EM fields at 250 000× magnification (1.06 μm2) on a Philips 301 TEM to calculate gold particles per 10 nl of undiluted stocks. To assess binding to monolayers, particles were incubated beneath lipid monolayers containing phosphatidylcholine (PC) only, or PC plus nickel-charged DHGN (PC+DHGN), as described in Materials and methods. After incubations, monolayers were lifted onto lacy carbon grids, rinsed for 30 s in 1% glucose, dried, viewed and photographed. Binding was quantitated by counting particles on hole-free, carbon support-free monolayer areas in 20 fields at 91 000× magnification to obtain particles bound per 1000 nm2. The success of the above experiments afforded us an avenue to evaluate the effects of incubation conditions on DHGN binding to His tags. Thus, lipid, salt, pH, ion and reducing agents were examined for their effects in gold conjugate binding studies (Table II). As expected, PC-only monolayers (Table II; 0% DHGN) and monolayers of PC plus DHGN that had not been charged with Ni2+ (20% DHGN no nickel) showed no appreciable His-tagged gold binding. We also observed that reduction of DHGN to PC ratios down to 1% DHGN reduced binding only slightly, whereas increasing the percentage to 50% reduced binding levels to 16% of the standard conditions. We interpret the results with 1% DHGN to indicate that this amount of DHGN (6×108 excess over gold particles) is still functionally saturating in our incubations, while we hypothesize that high levels of DHGN may disrupt PC monolayers in some as yet undetermined fashion. While high levels of DHGN reduced binding, variation of subphase salt and pH conditions showed little effect on binding (Table II). However, certain divalent cations seemed to interfere with the DHGN–His tag interaction. In particular, CaCl2 and FeCl2 reduced binding levels 4- to 5-fold, and 5 mM NiCl2 in the subphase eliminated binding in a more dramatic fashion than even EDTA. Finally, as might be expected from their effects on metal chelate chromatography (Hochuli et al., 1987), dithiothreitol (DTT) and β-mercaptoethanol (β-Me) also inhibited His-tagged gold conjugate binding to DHGN-containing monolayers, which suggests that reducing agents must be used sparingly in studies with this lipid. Table 2. Variables affecting His-tagged gold particle binding to lipid monolayers Variations Binding as % of standard conditions Lipid/salt 0% DHGN 0 1% DHGN 68 20% DHGN 100 50% DHGN 16 20% DHGN (no nickel) 7 0 mM NaCl 58 250 mM NaCla 100 500 mM NaCl 119 pH/buffer pH 9.0, phosphate 90 pH 8.5, phosphate 142 pH 7.8, phosphatea 100 pH 7.4, phosphate 112 pH 7.0, phosphate 76 pH 6.5, phosphate 96 pH 6.0, phosphate 135 pH 5.5, phosphate 133 pH 5.0, phosphate 130 pH 7.8, phosphatea 100 pH 7.8, Tris–HCl 118 pH 7.8, HEPES 70 Ion/miscellaneous No additionsa 100 0.1 mM ZnCl2 112 5 mM MgCl2 81 1 mM CaCl2 25 1 mM FeCl2 22 5 mM NiCl2 0 10 mM EDTA 7 5 mM DTT 0 1 mM DTT 9 5 mM β-Me 0 1 mM β-Me 9 Gold particle conjugates to His-ATF (His-ATF–gold) were incubated under lipid monolayers using a variety of different conditions to ascertain how different variables affect His tag binding to DHGN-containing monolayers. Incubations were performed as described in Materials and methods, and were compared with results from standard condition incubations (incubations marked with a) performed in parallel. Lipid variations involved using different percentages of nickel-charged DHGN in incubations, or employed DHGN which had not been pre-bound with Ni2+. Salt variations were performed using different concentrations of NaCl in the subphase; pH and buffer variations used 25 mM buffer substitutions in place of the standard 25 mM phosphate, pH 7.8 buffer; and ion, EDTA and reducing agent incubations involved addition of these agents to the subphase solutions of standard incubations. Results are listed as percentages of gold particle counts per unit area obtained in the parallel standard incubations. Analysis of M-MuLV capsid protein arrays Provided with DHGN, which can bind His-tagged proteins, it was possible to examine whether His-tagged M-MuLV Gag proteins could form 2D crystals on DHGN-containing monolayers. Initially, we have chosen to test an N-terminally His-tagged M-MuLV capsid protein (His-MoCA) for several reasons: capsid–capsid interactions have been shown to be major determinants in retrovirus particle assembly (Hansen et al., 1990, 1993; Jones et al., 1990; Wang et al., 1993, 1994; Mammano et al., 1994; Craven et al., 1995; Hansen and Barklis, 1995; McDermott et al., 1996); tethering of His-MoCA to our model monolayer is analogous to myristate-mediated anchoring of Gag proteins in vivo; His-MoCA is not subject to interdomain proteolytic processing in bacteria; and His-MoCA may serve as a foundation for comparison with larger Gag-derived fragments in the future. Our hypothesis is that His-MoCA interactions reflect interactions between membrane-tethered capsid domains in assembling or immature M-MuLV particles, and in vivo experiments concerning the relevance of this interpretation are described below. As produced from the pET15B-MoCA plasmid in Escherichia coli, His-MoCA is a 35 kDa protein, composed of the 30 kDa M-MuLV ca