Identification of Complement Regulatory Domains in
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病菌学杂志 2005年第19期
National Centre for Cell Science, Pune University Campus, Ganeshkhind, Pune 411007, India
ABSTRACT
Vaccinia virus encodes a homolog of the human complement regulators named vaccinia virus complement control protein (VCP). It is composed of four contiguous complement control protein (CCP) domains. Previously, VCP has been shown to bind to C3b and C4b and to inactivate the classical and alternative pathway C3 convertases by accelerating the decay of the classical pathway C3 convertase and (to a limited extent) the alternative pathway C3 convertase, as well as by supporting the factor I-mediated inactivation of C3b and C4b (the subunits of C3 convertases). In this study, we have mapped the CCP domains of VCP important for its cofactor activities, decay-accelerating activities, and binding to the target proteins by utilizing a series of deletion mutants. Our data indicate the following. (i) CCPs 1 to 3 are essential for cofactor activity for C3b and C4b; however, CCP 4 also contributes to the optimal activity. (ii) CCPs 1 to 2 are enough to mediate the classical pathway decay-accelerating activity but show very minimal activity, and all the four CCPs are necessary for its efficient activity. (iii) CCPs 2 to 4 mediate the alternative pathway decay-accelerating activity. (iv) CCPs 1 to 3 are required for binding to C3b and C4b, but the presence of CCP 4 enhances the affinity for both the target proteins. These results together demonstrate that the entire length of the protein is required for VCP's various functional activities and suggests why the four-domain structure of viral CCP is conserved in poxviruses.
INTRODUCTION
The complement system is an ancient and one of the principal innate immune defense mechanisms against all invading pathogens, including viruses (7, 34). It has a unique ability to recognize and label pathogens as non-self, with and without the help of pattern recognition molecules (7, 27). Apart from its direct effect on pathogens, it also plays an additional role in initiation and enhancement of the pathogen-specific adaptive immune responses (6, 34, 42). A significant body of knowledge indicates that both acute and latent viruses are susceptible to neutralization by the complement system (3, 7, 25). Thus, complement exerts a strong selective pressure on viruses during infection. To survive as successful pathogens, viruses have evolved different strategies to subvert the complement system. Efficient complement evasion mechanisms evolved by viruses include direct complement inactivation by encoding structural and/or functional homologs of host complement regulatory proteins or by capturing host membrane regulatory proteins and using host receptors for cellular entry (3, 25). Important examples include viruses belonging to the family of poxviruses, herpesviruses, retroviruses, paramyxoviruses, and picornaviruses (1, 10, 28, 30, 33, 51, 56).
Vaccinia virus (VV), a prototype of the family Poxviridae and genus Orthopoxvirus, is a cytoplasmic large DNA virus (18, 32). VV embodies an illustrious example of complement evasion. It is known to have developed two efficient mechanisms to evade the complement system. (i) It encodes within its genome a 27-kDa protein, homologous to human complement regulators, known as VV complement control protein (VCP), which is a potent soluble complement inhibitor (22, 29, 46). (ii) It incorporates host complement proteins (membrane cofactor protein [MCP], CD46; decay-accelerating factor [DAF], CD55; and CD59) while budding, rendering itself resistant to complement attack (57).
VCP, encoded by the C21L gene, is one of the first viral proteins identified to have complement-binding activity (23). The primary structure of VCP consists of 244 amino acids with a signal peptide of 19 amino acids. It comprises four CCP domains that are typically present in human regulators of complement activation (RCA) proteins and exhibits considerable sequence similarity to factor H (26%), C4b-binding protein (37%), MCP (35%), DAF (38%), and complement receptor 1 (CR1) (37%). The nuclear magnetic resonance structure of VCP modules (CCPs 2 to 3 and CCPs 3 to 4) revealed that each CCP folds into a compact six-?-strand structure similar to CCP 16 of factor H (13). Recent elucidation of the crystal structure of VCP depicted it as a highly extended molecule with heparin-binding site at its C-terminal region (9, 36). An interesting feature revealed by the crystal structure was the charge distribution of VCP domains: CCP domains 1 and 4 are surrounded by a positive field, while the middle two domains (CCPs 2 and 3) are surrounded by a predominantly negative field.
Functional studies revealed that VCP protects vaccinia virions from antibody-dependent complement-mediated neutralization (15); vaccinia virus mutants that do not express VCP are attenuated in vivo (15). Initial studies of the mechanism of complement inhibition performed using both partially purified VCP and culture medium containing secreted VCP have shown that it inhibits the classical pathway-mediated lysis of sheep erythrocytes, binds to C3b and C4b, and accelerates the decay of the classical and (to a limited extent) the alternative pathway C3 convertases (29). A later study performed using purified recombinant VCP demonstrated that it possesses factor I cofactor activity for C3b and C4b and is a poor inhibitor of the alternative pathway, in comparison to the classical pathway (46). Recently, the binding mechanism of VCP with C3b and C4b has been characterized using surface plasmon resonance (SPR). The data suggested that binding of VCP to C3b and C4b follows a simple 1:1 binding model and does not involve multiple-site interactions, as is observed for factor H and CR1 (2). Further, binding to C3b and C4b involves ionic interactions, and the binding site of VCP on C3b and C4b is located within the C3dg and C4c regions, respectively (2).
Although significant efforts were made to establish VCP as a complement inhibitor (3, 17), no efforts were made to shed light on the identification of its domains important for its complement regulatory activities: factor I cofactor activity for C3b and C4b, and decay-accelerating activity for the classical and alternative pathway C3 convertases. In this paper, we have identified which of the four CCPs of VCP are required to confer cofactor activities, decay-accelerating activities, and binding to target proteins C3b and C4b. Our results suggest that a minimum of three CCP domains are required for its cofactor activities, binding to C3b and C4b, and its limited alternative pathway decay-accelerating activity and that two domains are enough to mediate the classical pathway decay-accelerating activity.
MATERIALS AND METHODS
Reagents and buffers. Antibody-coated sheep erythrocytes (designated EA) were prepared by incubating the sheep erythrocytes with anti-sheep erythrocyte antibodies (ICN Biomedical, Inc., Irvine, Calif.). Antiserum recognizing VCP was raised in rabbit by immunizing with purified VCP. Buffers used were VBS (5 mM barbital and 145 mM NaCl, pH 7.4), GVB (VBS containing 0.1% gelatin), DGVB (2.5 mM barbital and 73 mM NaCl, 0.1% gelatin, and 2.5% dextrose), DGVB++ (DGVB containing 0.5 mM MgCl2 and 0.15 mM CaCl2), and phosphate-buffered saline (PBS) (10 mM sodium phosphate and 145 mM NaCl, pH 7.4). Mg EGTA contained 0.1 M MgCl2 and 0.1 M EGTA.
Purified complement proteins. The human complement factors B, D, H, and I were kindly provided by Michael K. Pangburn (University of Texas Health Center, Tyler, Tex.) and the recombinant human soluble CR1 (sCR1) was a generous gift of Henry Marsh (AVANT Immunotherapeutics, Inc., Needham, Mass.). Human C1, C4, and C2 were purchased from Calbiochem (La Jolla, Calif.). Human C3 was purified according to Hammer et al. (12) with minor modifications. Twenty parts of human plasma were mixed with 1 part of inhibitor solution (1 M KH2PO4, 0.2 M Na4 EDTA, 0.2 M benzamidine, and 1 mM phenylmethylsulfonyl fluoride) and precipitated first with 4.5% polyethylene glycol and then with 12% polyethylene glycol at 0°C. The 12% pellet was then dissolved in 20 mM sodium phosphate, pH 7.4, and loaded onto a Source Q column (1.2 by 9.5 cm; Amersham Pharmacia Biotech, Uppsala, Sweden). The bound proteins were eluted with a linear gradient of 0 to 0.5 M NaCl. Fractions containing C3 were identified by sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) and immunodiffusion, pooled, and loaded onto a Mono Q 10/10 column (Amersham Pharmacia Biotech) in 10 mM sodium phosphate, pH 7.9. Bound proteins were eluted with a linear salt gradient of 0 to 0.5 M and subjected to SDS-PAGE analysis. C3-containing fractions were pooled, concentrated by ultrafiltration, and loaded onto Superose 12 (two columns connected in a series; Amersham Pharmacia Biotech) in PBS. Homogeneous C3 fractions as analyzed by SDS-PAGE were pooled. The purified C3 contained >90% native C3, as determined by analyzing the protein sample on a Mono S column (Amersham Pharmacia Biotech) (39). C3b was generated by limited trypsin cleavage and purified on a Mono Q 5/5 column (46). Human C4b used was purified as previously described (2) or purchased from Calbiochem (La Jolla, Calif.).
Construction of deletion mutants. The deletion constructs comprising CCPs 1 to 3, 2 to 4, 1 to 2, 2 to 3, and 3 to 4 of VCP were constructed from full-length VCP (46) by PCR amplification and cloned into the yeast expression vector pPICZ (Invitrogen, Carlsbad, Calif.), downstream of the AOX1 methanol-inducible promoter by the standard protocol. For amplification of the respective VCP deletion mutants, the following sequence-specific primers were used: for CCP 1-3, AS-1 (5'-GGAATTCTGCTGTACTATTCCGTCACGACCC-3') and AS-5 (5'-GCTCTAGATTATTTAACAATCTGACACGTGGGTGG-3'); for CCP 2-4, AS-2 (5'-GGAATTCTGCCCATCGCCTCGAGATATCG-3') and AS-4 (5'-GCTCTAGATTAGCGTACACATTTTGGAAGTTCCG-3'); for CCP 1-2, AS-1 and AS-6 (5'-GCTCTAGATTATTTAACAGATTCACAAATAGGTGCC-3'); for CCP 2-3, AS-2 and AS-5; and for CCP 3-4, AS-3 (5'-GGAATTCTGCCAATCCCCTCCATCTATATCCAACGG-3') and AS-4. The primers incorporated restriction sites EcoRI and XbaI (underlined) in the forward and the reverse orientations, respectively, and a stop codon (in boldface type) in the reverse orientation primers. The amplification was carried out using Titanium Taq DNA polymerase (B. D. Biosciences Clonetech, Palo Alto, CA), with an initial cycle of denaturation at 94°C for 2 min, followed by 25 cycles of 1 min at 94°C, 1 min at 60°C or 65°C according to the annealing temperature of the primer, 1 min at 72°C, and a final extension for 10 min at 72°C. All the PCR products were digested with EcoRI and XbaI and cloned into pPICZ EcoRI and XbaI cut vector. The validity of all the constructs was confirmed by automated DNA sequencing. Five micrograms of each of the SacI-digested pPICZ VCP deletion constructs was then integrated into Pichia pastoris as per the manufacturer's protocol. The integration of the deletion mutants in Pichia were confirmed by amplifying the respective genomic DNA using the 5'AOX1 and 3'AOX1 primers and with a combination of gene-specific primers. Further, the validated integrants were used for expression of the recombinant proteins.
Expression and purification of vaccinia virus complement control protein and its deletion mutants. The vaccinia virus complement control protein cloned in Pichia pastoris was expressed and purified as previously described (2). Expression of VCP mutants was performed as described below. A single colony of recombinant Pichia pastoris expressing the respective VCP mutant was inoculated in 10 ml of BMGY medium (100 mM potassium phosphate, pH 6.0, 10 g/liter yeast extract, 20 g/liter peptone, 13.4 g/liter yeast nitrogen base, 0.4 mg/liter biotin, and 1% glycerol) and incubated overnight at 30°C in a shaking incubator. This inoculum was then added to 400 ml of BMGY and incubated for 48 h at 30°C with shaking. The cells were centrifuged, resuspended in 400 ml of BMMY (BMGY containing 0.5% methanol, but without 1% glycerol), and incubated at 30°C for an additional 96 h with vigorous shaking; 0.5% methanol was added after every 24 h. After incubation, cells were pelleted and the supernatant containing the VCP mutant was collected for purification.
For purification, the supernatants containing VCP mutants were concentrated by ultrafiltration and precipitated with 80% ammonium sulfate at 0°C. The pellets were then suspended and dialyzed in PBS. For further purification, CCP 1-3 and CCP 1-2 mutants were passed through a PD-10 column (Amersham Pharmacia Biotech) in 10 mM sodium phosphate (pH 7.4), loaded onto a heparin-agarose column in the same buffer, and eluted with 500 mM NaCl. The eluted proteins were then exchanged into 20 mM Tris (pH 7.9), loaded onto a Mono Q column, and eluted with a linear gradient of 0 to 500 mM NaCl. For purification of CCP 2-4 and CCP 2-3 mutants, the samples were passed through DEAE-Sephacel (Sigma, St. Louis, Mo.) preequilibrated with 10 mM sodium phosphate, pH 7.4, containing 150 mM NaCl. The flowthrough was collected, passed through a PD-10 column in 20 mM Tris (pH 7.9), and loaded onto a Mono Q column. The bound proteins were eluted with a linear gradient of 0 to 500 mM NaCl. CCP 3-4 mutant was purified by being loaded onto DEAE-Sephacel in 10 mM sodium phosphate, pH 7.4, containing 150 mM NaCl. The flowthrough containing CCP 3-4 mutant was collected and buffer exchange was performed using a PD-10 column preequilibrated with 5 mM sodium acetate, pH 4.0. The sample was then loaded onto a Mono S column, and the bound proteins were eluted with a linear salt gradient of 0 to 500 mM NaCl. Eluted fractions in all the above purifications were analyzed by SDS-PAGE and Western blotting using anti-VCP antibodies; fractions containing pure proteins were pooled, dialyzed into PBS, and concentrated by ultrafiltration.
Mass analysis and protein sequencing. The molecular mass determination and sequencing of the VCP mutants were performed at the Biomolecular Research Facility (University of Virginia, Charlottesville, VA) as described below. The molecular weights of the VCP mutants were determined by matrix-assisted laser desorption ionization-time of flight mass spectrometry (MS) using an Applied Biosystems Voyager DE-Pro. The samples were dissolved in 70% acetonitrile-30% water, mixed with sinapinic acid matrix, and spotted on the matrix-assisted laser desorption ionization plate. Approximately 200 spectra were averaged for each sample.
The sequencing of VCP mutants was performed as described below. The samples were run on a 12% SDS-PAGE gel and stained with Coomassie blue. The protein bands were cut, transferred to a siliconized tube, washed, and destained overnight in 200 μl of 50% methanol. The gel pieces were then dehydrated in acetonitrile, rehydrated in 30 μl of 10 mM dithiolthreitol (DTT) in 0.1 M ammonium bicarbonate, and reduced at room temperature for 30 min. The DTT solution was then removed, and the samples were alkylated in 30 μl of 50 mM iodoacetamide in 0.1 M ammonium bicarbonate at room temperature for 30 min. The reagent was removed, and the gel pieces were dehydrated in 100 μl of acetonitrile. The gel pieces were subjected to two cycles of dehydration and rehydration in acetonitrile and 0.1 M ammonium bicarbonate and then dried by vacuum centrifugation. The gel pieces were rehydrated in 20 ng/μl trypsin (second band for each sample in chymotrypsin) in 50 mM ammonium bicarbonate on ice for 10 min, excess enzyme solution was removed, and 20 μl of 50 mM ammonium bicarbonate was added. The samples were digested overnight at 37°C, and the peptides formed were extracted from the polyacrylamide in two 30-μl aliquots of 50% acetonitrile-5% formic acid. These extracts were combined and evaporated to 25 μl for MS analysis, which was done using a liquid chromatography-MS system consisting of a Finnigan LCQ ion trap mass spectrometer with a Protana nanospray ion source interfaced to a self-packed 8 cm by 75 μm (inner diameter) Phenomenex Jupiter 10-μm C18 reversed-phase capillary column. The digest was analyzed using the double-play capability of the instrument acquiring one full scan mass spectrum to determine peptide molecular weights, followed by four product ion spectra to determine amino acid sequences in sequential scans. The data were analyzed by database searching using the Sequest algorithm and by manual interpretation.
SPR measurements. The kinetics of binding of VCP and VCP mutants to human C3b and C4b were determined by using the SPR-based biosensor BIACORE 2000 (Biacore AB, Uppsala, Sweden). Binding experiments were performed using physiologic ionic strength buffer (PBS-T, 10 mM sodium phosphate, 145 mM NaCl, pH 7.4, containing 0.05% Tween-20) at 25°C. Addition of 0.05% Tween-20 blocked the nonspecific adsorption of VCP to the sensor chips (50). In this assay, both C3b and C4b were oriented in their physiological orientation on a streptavidin chip (Sensor Chip SA, Biacore AB) by labeling their free -SH group with biotin (2). In brief, 914 response units (RUs) of C3b and 916 RUs of C4b were immobilized on FC-2 and FC-3, respectively, and FC-1 served as a control flow cell. Binding was measured at 50 μl/min to avoid the mass transport effect, and it was measured for 120 s. Dissociation was measured for 180 s. The sensor chip surfaces were regenerated with 30-s pulses of 0.2 M sodium carbonate, pH 9.5. Sensograms obtained for the control flow cell (Fc-1) were subtracted from the data for the flow cell immobilized with either C3b or C4b, and the SPR data were analyzed by BIAevaluation software version 4.1 using global fitting.
Measurement of factor I cofactor activity. Both qualitative and quantitative analyses of factor I cofactor activity of VCP and its mutants for C3b and C4b were determined using 10 mM phosphate buffer, pH 7.4, containing 145 mM NaCl. In these assays, 2.7 μg of C3b or 2.9 μg of C4b was mixed with 100 ng of factor I and VCP (as indicated in the figure legends) or its mutants and incubated at 37°C for 4 h in a total volume of 15 μl. The reactions were stopped by the addition of sample buffer containing DTT and electrophoresed on a 9% SDS-PAGE gel for determining C3b cleavages and a 10% SDS-PAGE for C4b cleavages. The cleavage products were visualized by staining the gel with Coomassie blue. For quantitative analysis, gels were scanned for densitometric analysis by using the ChemiDoc XRS system (Bio-Rad, Segrate, Italy) to calculate the percentage of '-chain. The data obtained were normalized by considering 100% '-chain to be equal to the '-chain intensity obtained in the absence of factor I (control). The binding data were fit using nonlinear regression analysis (GraFit; Erithacus Software, London, United Kingdom) and a four-parameter logistic analysis was performed to identify the best-fit 50% inhibitory concentration value (the concentration of VCP causing 50% '-chain cleavage).
Measurement of C3 convertase decay-accelerating activity. The classical pathway decay-accelerating activity of VCP and its mutants was determined by forming EAC142 (37). In brief, the classical pathway C3 convertase EAC142 was made by incubating 100 μl of EA (4 x 109/ml) in DGVB++ with 4 μg C1 for 20 min at 30°C in a total volume of 120 μl. The cells were then washed with ice-cold DGVB++, resuspended in 100 μl DGVB++, and incubated for 20 min at 30°C with 8 μg of C4. After the incubation, they were mixed with 4 μg of C2 and further incubated for 4 min at 30°C to form EAC142. The convertase formation was then stopped by adding EDTA to a final concentration of 10 mM. To define the effect of VCP and its mutants on the decay of EAC142, 10 μl of EAC142 (109/ml) was mixed with various concentrations of VCP or its mutants in a total volume of 25 μl for 5 min at 22°C. The reaction mixture was then mixed with 125 μl of DGVB and 100 μl of guinea pig serum diluted 1:100 in DGVB containing 40 mM EDTA, incubated 30 min at 37°C, and centrifuged. The percentage of lysis was determined by measuring the absorbance at 405 nM.
The alternative pathway C3 convertase decay was measured by forming C3b,Bb on rabbit (ER) or sheep (ES) erythrocytes. ERC3b and ESC3b were generated by incubating ER or ES with C3, factor B, and factor D in the presence of NiCl2 as previously described (43, 47). C3b,Bb on ER or ES was made by mixing 150 μl of C3b-coated rabbit or sheep erythrocytes (109/ml) with 45 μg of factor B and 0.76 μg of factor D in 210 μl of GVB containing 3.5 mM NiCl2. The reaction mixture was then incubated for 5 min at 30°C and stopped by the addition of EDTA to a final concentration of 10 mM. Immediately thereafter, 10 μl of ERC3b,Bb or ESC3b,Bb (109/ml) was mixed with various concentrations of VCP or its mutants in a total volume of 50 μl and incubated for 10 min at 37°C. The reaction mixture was then mixed with 50 μl of human serum diluted 1:5 in GVB containing 20 mM EDTA and incubated at 37°C for 20 min. After incubation, the reaction mixture was mixed with 200 μl of GVBE and centrifuged. The percentage of lysis was determined as above. The data generated were normalized by setting 100% lysis equal to the lysis that occurred in the absence of an inhibitor.
RESULTS
Design, expression, purification, and characterization of VCP deletion mutants. Three aspects were taken into consideration while designing the mutants. First, since VCP is a soluble protein, mutants were expressed as soluble proteins and not as membrane-bound fusion proteins as described in an earlier study, where attempts were made to localize the C3b-binding site (45). Second, it has previously been shown for the human complement regulators that apart from the CCP domains, the interdomain linker regions may also play a role in the ligand binding (14); therefore, each construct started with the first Cys of the starting domain and ended with the last residue of the interdomain linker. This design kept the entire linker region at the C-terminal side of each mutant (see the amino acid sequences listed in Table 1). Third, several previous studies on the human regulators have shown that two or three contiguous domains (5, 14, 19), as well as their native configuration (i.e., distance between the successive domains) (4, 24), is essential for their complement regulatory activities; therefore, mutants designed contained either two or three successive CCP domains.
Earlier, we and others have shown that the Pichia expression system produces a high yield of properly folded VCP (36, 46). Thus, we utilized the same expression system for expressing VCP mutants. The culture supernatants containing the expressed mutants were purified by using a series of chromatographic procedures as described in Materials and Methods. The purified mutants were homogeneous and >95% pure as determined by SDS-PAGE analysis (Fig. 1). To confirm the identity of the mutants, they were subjected to sequencing by mass spectrometry. The amino acid sequences of the expressed mutants were consistent with the predicted sequences confirming the identity of each mutant (Table 1). The addition of EAEAEF in the beginning of each mutant was the result of cloning into the Pichia vector. The molecular masses of the expressed mutants were analyzed by SDS-PAGE and mass spectrometry. In SDS-PAGE, all the proteins migrated higher than their molecular weights; however, this was not due to differences in glycosylation because the primary sequence of VCP does not contain any glycosylation site (23). The data obtained using mass spectrometry confirmed that the molecular masses of the mutants were similar to the calculated masses (Table 1).
Characterization of factor I cofactor activity of VCP mutants. A fluid-phase assay was utilized to characterize the factor I cofactor activity of VCP mutants for C3b and C4b. In this assay, C3b or C4b was incubated with the mutant proteins and factor I, and cleavage of the '-chain was assessed by running the samples on SDS-PAGE gels. VCP and the CCP 1-3 mutant supported cleavage of the '-chain of C3b (Fig. 2), as well as C4b (Fig. 3), but none of the other mutants showed any detectable activity. These data indicate that CCPs 1 to 3 comprise the minimum region required for the factor I cofactor activity for both C3b and C4b. Incubation of the CCP 1-3 mutant with C3b and factor I did not result in complete cleavage of the '-chain of C3b (Fig. 2). We therefore sought to analyze the relative factor I cofactor activity of VCP and the CCP 1-3 mutant for C3b and C4b. In this assay, C3b or C4b was incubated with factor I and increasing concentrations of either VCP or CCP 1-3 mutant. Figure 4 shows that higher concentrations of the CCP 1-3 mutant than that of VCP were required to cleave the '-chains of C3b and C4b. The concentrations of VCP required to cleave 50% of the '-chain of C3b and C4b were 0.12 μM and 0.038 μM, respectively, while the concentrations of the CCP 1-3 mutant required to cleave 50% of the '-chain of C3b and C4b were 4.9 μM and 0.58 μM, respectively. These data indicate that in comparison to VCP, the CCP 1-3 mutant has 41- and 15-fold-lower factor I cofactor activity for C3b and C4b, respectively.
Characterization of decay-accelerating activity of VCP mutants. A previous study had demonstrated that apart from factor I cofactor activity, VCP also possesses significant decay-accelerating activity for the classical and limited decay-accelerating activity for the alternative pathway C3 convertases (29). To identify the domains required for the decay-accelerating activity, we measured the activity of VCP mutants using hemolytic assays. For measuring the decay-accelerating activity for the classical pathway C3 convertase C4b,2a, the enzyme was formed on the sensitized sheep erythrocytes and allowed to incubate with the mutant proteins. The remaining C3 convertase activity was quantitated by measuring hemolysis after adding EDTA sera (a source of C3 to C9). VCP showed significant decay-accelerating activity even at a 100 nM concentration. On a molar basis, VCP was only 1.8-fold-less active than sCR1. Among the mutants, CCP 1-3 and CCP 1-2 mutants showed decay-accelerating activity, while the other mutants did not show any activity (Fig. 5), indicating that CCPs 1 to 2 comprise the minimum region required for the classical pathway. It should be pointed out here that the concentration of CCP 1-2 mutant required to inhibit the decay-accelerating activity was 5,600-fold higher than the concentration of VCP (Fig. 5), suggesting that in addition to CCPs 1 to 2, CCPs 3 and 4 also contribute significantly to the classical pathway decay-accelerating activity.
For measuring the alternative pathway decay-accelerating activity of VCP and its mutants, the alternative pathway C3 convertase C3b,Bb was formed on rabbit erythrocytes. The enzyme was incubated with VCP or the mutant proteins, and the remaining enzyme activity was detected by adding EDTA sera and measuring hemolysis. Surprisingly, neither VCP nor its mutants showed any alternative pathway decay-accelerating activity even at a 50 μM concentration. In a parallel assay, factor H and sCR1 showed significant inhibition at 100 nM and 2.5 nM concentrations, respectively (Fig. 6A). Since the previous study which reported limited alternative pathway decay-accelerating activity of VCP utilized sheep erythrocytes for forming C3b,Bb (29) and not the rabbit erythrocytes, we tested if the lack of inhibition in our assays was due to the differences in the cells used. Thus, we formed C3b,Bb on sheep erythrocytes and measured the decay-accelerating activity of VCP and its mutants. On sheep cells, VCP accelerated the decay of alternative pathway C3 convertase, though the concentration of VCP required to decay 50% alternative pathway C3 convertase was 64 μM, which was 2,400-fold higher than that required to decay the classical pathway C3 convertase (Fig. 6B). Among VCP mutants, only the CCP 2-4 mutant showed inhibition, although like VCP at a similarly high concentration, suggesting that the limited AP decay-accelerating activity is imparted by CCPs 2 to 4.
Effect of membrane sialic acid on the decay-accelerating activity of VCP. From the data presented in Fig. 6, it is clear that VCP shows limited decay-accelerating activity against the alternative pathway C3 convertase present on sheep erythrocytes but not against the convertase present on rabbit erythrocytes. Previously, differences in the decay-accelerating activity against the alternative pathway C3 convertase present on sheep versus rabbit erythrocytes have been reported for human complement regulator factor H. It has been shown that factor H displays enhanced decay-accelerating activity towards the alternative pathway C3 convertase present on sheep erythrocytes; this has been attributed to the interaction of its heparin-polyanion-binding site with sialic acid present on the erythrocyte membrane (rabbit erythrocytes contain approximately 13% of the sialic acid present on sheep erythrocytes) (8, 40). Since VCP contains a heparin-binding site on CCP 4 (9), we sought to analyze if the differences in its decay activity on sheep versus rabbit erythrocytes were due to interaction of its heparin-polyanion-binding site with sialic acid present on sheep erythrocytes. For this purpose, we treated sheep erythrocytes with neuraminidase and compared the decay-accelerating activity of VCP against the C3 convertase present on untreated and treated erythrocytes; factor H was included as a positive control. As expected, factor H showed reduced alternative pathway C3 convertase decay-accelerating activity on neuraminidase-treated cells compared to untreated cells (Fig. 7); the decay activity was 7.4-fold less on the treated than on the untreated sheep cells. However, VCP did not show any difference in the decay activity, thereby indicating that like factor H, the heparin-binding site on VCP does not help in enhancing the decay of the alternative pathway C3 convertase on sheep cells. Thus, the observed differences in the decay activities of VCP on sheep and rabbit erythrocytes were not due to the differences in the membrane sialic acid content.
Binding of VCP mutants to C3b and C4b. From the data presented above, it is clear that CCPs 1 to 2 and 2 to 4 are the minimum regions required for the decay-accelerating activities and that CCPs 1 to 3 are required for cofactor activities. To determine whether reduced decay-accelerating activities and cofactor activities of the VCP mutants correlate with binding to C3b and C4b, we performed a quantitative analysis of binding of VCP and its mutants to C3b and C4b using a surface plasmon resonance assay. In this assay, C3b and C4b were oriented on a streptavidin chip by labeling their free -SH groups with biotin (2). This setup mimicked the physiological orientation of these proteins and also allowed accurate determination of the binding constants (2). When VCP or its deletion mutants were allowed to flow over the sensor chip, only VCP and the CCP 1-3 mutant bound to C3b and C4b (Fig. 8). Further analysis showed that the CCP 1-3 mutant had 36- and 228-fold-lower affinity for C3b and C4b, respectively, than VCP (Fig. 9 and Table 2). This decrease was primarily due to reduction in the on-rates of the mutant (Table 2). These data suggest that poor factor I cofactor activities of the CCP 1-3 mutant could be due to its lower affinity for C3b and C4b. It is, however, interesting that the CCP 1-2 and CCP 2-4 mutants, which showed the classical and alternative pathway decay-accelerating activities, respectively, showed no detectable binding to C4b and C3b.
DISCUSSION
In the present study, we utilized a series of VCP deletion mutants to address the contribution of each CCP domain to its cofactor activities, decay-accelerating activities, and binding to C3b and C4b. Our data on the localization of domains required for factor I cofactor activity for C3b and C4b indicate that a minimum of three CCPs (CCPs 1 to 3) are essential for these activities, because further deletion of either CCP 1 or CCP 3 completely abolished the cofactor activities (Fig. 2 and 3). These data are consistent with the previous observations on mapping of the human RCA proteins (factor H, C4bp, MCP, and CR1) for cofactor activities, which indicated that a minimum of three CCPs are required for cofactor activities (4, 11, 14, 16, 19, 20, 44, 52). There is now a consensus that factor I-mediated inactivation of C3b and C4b involves two steps: (i) recognition of C3b or C4b by the cofactor and (ii) interaction of factor I with C3b or C4b and the cofactor (48, 54). Thus, it is conceivable that the recognition sites for C3b or C4b and factor I on the complement regulators are spatially conserved and require three CCPs. This, however, does not seem to be a general phenomenon as our recent data on kaposica, the complement regulator of the Kaposi's sarcoma-associated herpesvirus, indicated that only two CCPs are enough to impart cofactor activities in kaposica (35). Since CCPs 1 to 3 are required for both C3b and C4b cofactor activities, it can be envisaged that the recognition site for C3b and C4b on VCP is formed by similar residues. This possibility, however, appears to be less likely for VCP as it binds to different regions on C3b and C4b; its binding site on C3b is located on the C3dg region, while its binding site on C4b is located on the C4c region (2).
VCP has previously been shown to possess efficient decay-accelerating activity for the classical pathway and weak decay activity for the alternative pathway C3 convertases (29). Our data show that CCPs 1 to 2 of VCP form the minimum region essential for its classical pathway decay-accelerating activity, but CCPs 3 and 4 are required for its complete activity (Fig. 5). Earlier studies of DAF showed that CCPs 2 to 3 are important for its classical pathway decay-accelerating activity (5). It is pertinent to point out here that CCPs 1 to 2 of VCP show maximum sequence similarity to CCPs 2 to 3 of DAF, which possesses classical pathway decay-accelerating activity (5). Our data show that VCP possesses limited decay-accelerating activity only against the alternative pathway C3 convertase present on the sheep erythrocytes and not against the C3 convertase present on rabbit erythrocytes (Fig. 6). Since VCP contains a heparin-binding site like factor H (9), it is likely that similar to factor H (31, 49), the heparin-binding site on VCP helps it to enhance its decay-accelerating activity against the alternative pathway C3 convertase present on the sheep erythrocytes, which contains a high surface density of sialic acid. However, this premise did not hold true, as the decay rate of VCP did not reduce on neuraminidase-treated sheep erythrocytes (Fig. 7). The weak decay activity was also observed with the CCP 2-4 mutant, suggesting that CCPs 2 to 4 are important for this activity. Like VCP, the Kaposi's sarcoma-associated herpesvirus complement control protein kaposica also contains very weak decay-accelerating activity against alternative pathway C3 convertase (35, 55). Thus, it seems that viral homologs of complement control proteins primarily contain three out of four C3 convertase regulatory activities of the host complement regulators.
Our data on binding of VCP to target proteins measured using SPR identified CCPs 1 to 3 as the smallest structural unit capable of binding to C3b and C4b (Fig. 8). The presence of CCP 4, however, significantly enhanced the affinities for C3b and C4b (Fig. 9 and Table 2). These findings are consistent with the previous data on binding of the human RCA proteins to C3b and C4b, which essentially suggested that C3b or C4b binding primarily resides within three or four CCPs (4, 14, 19, 20, 52). Previously, two studies attempted to localize the C3b-binding site on VCP. Both these studies suggested that all the four CCPs are required for C3b binding. In the first study, various domains of VCP were expressed as membrane-bound CR2 fusion proteins (45). The lack of binding of the CCP 1-3 mutant to C3b in this study (as opposed to our present study) could have been due to lower sensitivity of the assay used. Alternatively, it is possible that the fusion of CCP 3 to CR2 could have sterically hindered the access to residues present on CCP 3, which are critical for binding to C3b. Unlike the first study, the second study utilized soluble proteins and SPR technology for detecting binding but failed to test binding of the CCP 1-3 mutant to C3b due to lack of this mutant (53).
Even though ligand binding is a prerequisite for cofactor and decay activities, a large body of evidence indicates that ligand binding does not always correlate well with cofactor and decay activities (26, 41). Our data support this premise. For example, the CCP 1-3 mutant showed a 36-fold decrease in affinity for C3b and a 228-fold decrease in affinity for C4b compared to VCP (Table 2). Although the decrease in C3b binding of the CCP 1-3 mutant correlated well with a 41-fold reduction in its C3b cofactor activity, the dramatic reduction in its affinity for C4b did not correlate with the 15-fold decrease in C4b cofactor activity. Similarly, CCP 1-2 and CCP 2-4 mutants which exhibited the classical and alternative pathway decay-accelerating activities, respectively, did not show detectable binding to C4b and C3b. It is likely that like DAF, these mutants might have better affinity for the C3 convertases than for their subunits C4b and C3b; DAF has been shown to bind to C4b,2a with 1,000-fold higher affinity than to C4b alone (38).
The data discussed above point out that although only two or three CCPs are enough to mediate the cofactor and decay activities of VCP, the entire length of the protein is essential for its full functional activity. It is likely that the residues scattered on two or three CCPs together form a binding site for C3 convertases or their subunits (C3b or C4b) and that the other CCPs contribute to the conformational stability of the complexes. Alternatively, a larger surface spanning all the four domains of VCP contributes to forming the recognition site for the convertases and C3b or C4b. These possibilities assume that functional domains are arranged in a linear fashion. This, however, may not be true; it is also likely that the recognition sites are discontinuous and that the middle domains or regions serve as essential spacers. These data, along with our recent observation on kaposica, which also indicated that the entire length of the protein is important for its complete function (35), explain why the four-CCP structure of viral CCP is highly conserved in viruses.
The important question in vaccinia virus biology is how VCP helps to protect the vaccinia virus from complement-mediated neutralization in vivo. The complement system is known to neutralize viruses in the presence and the absence of specific antibodies. Previously, it has been shown that extracellular enveloped virus, which is important for the virus dissemination, is protected from the alternative pathway due to the presence of host complement regulators on its envelope (57). However, this form remains susceptible to classical pathway-mediated neutralization (57). Thus, vaccinia virus must encode an effective regulator against the classical pathway to overcome the classical pathway-mediated neutralization. Earlier studies have shown VCP to be a better regulator of the classical pathway than even the host complement regulators factor H and C4-binding protein (21, 46); in the present manuscript, we have shown that VCP is only twofold-less effective than sCR1 in decaying the classical pathway C3 convertase (Fig. 5). The question is whether this inhibition of the classical pathway is also seen in vivo? In an earlier in vivo study, it was shown that the lesions formed by the VCP deletion mutant and the wild-type virus were similar for the first few days, but the lesion size was significantly reduced after 5 days in the mutant virus (15). Further, the appearance of antibody capable of mediating complement-enhanced neutralization correlated well with this timing (15), suggesting that VCP effectively inhibits classical pathway-mediated neutralization in vivo. In the present manuscript, we have shown which of the domains of VCP are required for each of its complement regulatory activities. Further delineation of the vital control points in VCP important for inhibition of the classical pathway decay-accelerating activity and C4b cofactor activity and targeting these control points would allow evaluation of the relative importance of these activities in immune evasion and may provide an alternative approach to manage vaccinia virus vaccine-related complications.
ACKNOWLEDGMENTS
We thank John D. Lambris (Department of Pathology and Laboratory Medicine, University of Pennsylvania, Philadelphia, PA) for his continuous support; Michael K. Pangburn (Department of Biochemistry, University of Texas Health Science Center, Tyler, Texas) for his support and the generous gift of complement proteins factors B, D, H, and I; Henry Marsh (AVANT Immunotherapeutics, Inc., Needham, Mass.) for providing sCR1; Nicholas E. Sherman (Biomolecular Research Facility, University of Virginia, Charlottsville, Va.) for sequencing VCP mutants; and Gabriela Canziani (Protein Interaction Facility, University of Utah, Salt Lake City, Utah) for advice on maintenance of Biacore.
This work was supported by a Wellcome Trust Senior Research Fellowship in Biomedical Science in India to A.S. The authors also acknowledge financial assistance to A.K.S. by the Council of Scientific and Industrial Research of India.
J.M. and J.B. contributed equally to this work.
REFERENCES
Alcami, A., and U. H. Koszinowski. 2000. Viral mechanisms of immune evasion. Trends Microbiol. 8:410-418.
Bernet, J., J. Mullick, Y. Panse, P. B. Parab, and A. Sahu. 2004. Kinetic analysis of the interactions between vaccinia virus complement control protein and human complement proteins C3b and C4b. J. Virol. 78:9446-9457.
Bernet, J., J. Mullick, A. K. Singh, and A. Sahu. 2003. Viral mimicry of the complement system. J. Biosci. 28:249-264.
Blom, A. M., L. Kask, and B. Dahlback. 2001. Structural requirements for the complement regulatory activities of C4BP. J. Biol. Chem. 276:27136-27144.
Brodbeck, W. G., D. Liu, J. Sperry, C. Mold, and M. E. Medof. 1996. Localization of classical and alternative pathway regulatory activity within the decay-accelerating factor. J. Immunol. 156:2528-2533.
Carroll, M. C. 2004. The complement system in regulation of adaptive immunity. Nat. Immunol. 5:981-986.
Cooper, N. R. 1998. Complement and viruses, p. 393-407. In J. E. Volanakis and M. M. Frank (ed.), The human complement system in health and disease. Marcel Dekker, Inc., New York, N.Y.
Fearon, D. T. 1978. Regulation of membrane sialic acid of ?1H-dependent decay-dissociation of amplification C3 convertase of the alternative complement pathway. Proc. Natl. Acad. Sci. USA 75:1971-1978.
Ganesh, V. K., S. A. Smith, G. J. Kotwal, and K. H. Murthy. 2004. Structure of vaccinia complement protein in complex with heparin and potential implications for complement regulation. Proc. Natl. Acad. Sci. USA 101:8924-8929.
Gewurz, B. E., R. Gaudet, D. Tortorella, E. W. Wang, and H. L. Ploegh. 2001. Virus subversion of immunity: a structural perspective. Curr. Opin. Immunol. 13:442-450.
Gordon, D. L., R. M. Kaufman, T. K. Blackmore, J. Kwong, and D. M. Lublin. 1995. Identification of complement regulatory domains in human factor H. J. Immunol. 155:348-356.
Hammer, C. H., G. H. Wirtz, L. Renfer, H. D. Gresham, and B. F. Tack. 1981. Large scale isolation of functionally active components of the human complement system. J. Biol. Chem. 256:3995-4006.
Henderson, C. E., K. Bromek, N. P. Mullin, B. O. Smith, D. Uhrin, and P. N. Barlow. 2001. Solution structure and dynamics of the central CCP module pair of a poxvirus complement control protein. J. Mol. Biol. 307:323-339.
Hourcade, D., M. K. Liszewski, M. Krych-Goldberg, and J. P. Atkinson. 2000. Functional domains, structural variations and pathogen interactions of MCP, DAF and CR1. Immunopharmacology 49:103-116.
Isaacs, S. N., G. J. Kotwal, and B. Moss. 1992. Vaccinia virus complement-control protein prevents antibody-dependent complement-enhanced neutralization of infectivity and contributes to virulence. Proc. Natl. Acad. Sci. USA 89:628-632.
Iwata, K., T. Seya, Y. Yanagi, J. M. Pesando, P. M. Johnson, M. Okabe, S. Ueda, H. Ariga, and S. Nagasawa. 1995. Diversity of sites for measles virus binding and for inactivation of complement C3b and C4b on membrane cofactor protein CD46. J. Biol. Chem. 270:15148-15152.
Jha, P., and G. J. Kotwal. 2003. Vaccinia complement control protein: multi-functional protein and a potential wonder drug. J. Biosci. 28:265-271.
Johnston, J. B., and G. McFadden. 2003. Poxvirus immunomodulatory strategies: current perspectives. J. Virol. 77:6093-6100.
Jokiranta, T. S., J. Hellwage, V. Koistinen, P. F. Zipfel, and S. Meri. 2000. Each of the three binding sites on complement factor H interacts with a distinct site on C3b. J. Biol. Chem. 275:27657-27662.
Kalli, K. R., P. H. Hsu, T. J. Bartow, J. M. Ahearn, A. K. Matsumoto, L. B. Klickstein, and D. T. Fearon. 1992. Mapping of the C3b-binding site of CR1 and construction of a (CR1)2-F(ab')2 chimeric complement inhibitor. J. Exp. Med. 174:1451-1460.
Kotwal, G. J. 1994. Purification of virokines using ultrafiltration. Am. Biotechnol. Lab. 12:76-77.
Kotwal, G. J., S. N. Isaacs, R. Mckenzie, M. M. Frank, and B. Moss. 1990. Inhibition of the complement cascade by the major secretory protein of vaccinia virus. Science 250:827-830.
Kotwal, G. J., and B. Moss. 1988. Vaccinia virus encodes a secretory polypeptide structurally related to complement control proteins. Nature 335:176-178.
Kuhn, S., C. Skerka, and P. F. Zipfel. 1995. Mapping of the complement regulatory domains in the human factor H-like protein 1 and in factor H. J. Immunol. 155:5663-5670.
Lachmann, P. J. 2002. Microbial subversion of the immune response. Proc. Natl. Acad. Sci. USA 99:8461-8462.
Lambris, J. D., A. Sahu, and R. Wetsel. 1998. The chemistry and biology of C3, C4, and C5, p. 83-118. In J. E. Volanakis and M. Frank (ed.), The human complement system in health and disease. Marcel Dekker, Inc., New York, N.Y.
Lu, J., C. Teh, U. Kishore, and K. B. Reid. 2002. Collectins and ficolins: sugar pattern recognition molecules of the mammalian innate immune system. Biochim. Biophys. Acta 1572:387-400.
Lubinski, J., T. Nagashunmugam, and H. M. Friedman. 1998. Viral interference with antibody and complement. Semin. Cell Dev. Biol. 9:329-337.
Mckenzie, R., G. J. Kotwal, B. Moss, C. H. Hammer, and M. M. Frank. 1992. Regulation of complement activity by vaccinia virus complement-control protein. J. Infect. Dis. 166:1245-1250.
Means, R. E., S. M. Lang, Y. H. Chung, and J. U. Jung. 2002. Kaposi's sarcoma associated herpesvirus immune evasion strategies. Front. Biosci. 7:e185-e203.
Meri, S., and M. K. Pangburn. 1990. Discrimination between activators and nonactivators of the alternative pathway of complement: regulation via a sialic acid/polyanion binding site on factor H. Proc. Natl. Acad. Sci. USA 87:3982-3986.
Moss, B. 2001. Poxviridae: the viruses and their replication, p. 2849-2883. In D. M. Knipe, P. M. Howley, D. E. Griffin, R. A. Lamb, M. A. Martin, B. Roizman, and S. E. Straus (ed.), Fields virology, 4th ed. Lippincott Williams and Wilkins, Philadelphia, Pa.
Mullick, J., J. Bernet, A. K. Singh, J. D. Lambris, and A. Sahu. 2003. Kaposi's sarcoma-associated herpesvirus (human herpesvirus-8) open reading frame 4 protein (kaposica) is a functional homolog of complement control proteins. J. Virol. 77:3878-3881.
Mullick, J., A. Kadam, and A. Sahu. 2003. Herpes and pox viral complement control proteins: ‘the mask of self.’ Trends Immunol. 24:500-507.
Mullick, J., A. K. Singh, Y. Panse, V. Yadav, J. Bernet, and A. Sahu. 2005. Identification of functional domains in kaposica, the complement control protein homolog of Kaposi's sarcoma-associated herpesvirus (human herpesvirus-8). J. Virol. 79:5850-5856.
Murthy, K. H., S. A. Smith, V. K. Ganesh, K. W. Judge, N. Mullin, P. N. Barlow, C. M. Ogata, and G. J. Kotwal. 2001. Crystal structure of a complement control protein that regulates both pathways of complement activation and binds heparan sulfate proteoglycans. Cell 104:301-311.
Pan, Q., R. O. Ebanks, and D. E. Isenman. 2000. Two clusters of acidic amino acids near the NH2 terminus of complement component C4 alpha'-chain are important for C2 binding. J. Immunol. 165:2518-2527.
Pangburn, M. K. 1986. Differences between the binding sites of the complement regulatory proteins DAF, CR1, and factor H on C3 convertases. J. Immunol. 136:2216-2221.
Pangburn, M. K. 1987. A fluorimetric assay for native C3. The hemolytically active form of the third component of human complement. J. Immunol. Methods 102:7-14.
Pangburn, M. K., and H. J. Müller-Eberhard. 1978. Complement C3 convertase: cell surface restriction of ?1H control and generation of restriction on neuraminidase-treated cells. Proc. Natl. Acad. Sci. USA 75:2416-2420.
Pangburn, M. K., K. L. Pangburn, V. Koistinen, S. Meri, and A. K. Sharma. 2000. Molecular mechanisms of target recognition in an innate immune system: interactions among factor H, C3b, and target in the alternative pathway of human complement. J. Immunol. 164:4742-4751.
Ploegh, H. L. 1998. Viral strategies of immune evasion. Science 280:248-253.
Pryzdial, E. L., and D. E. Isenman. 1986. A reexamination of the role of magnesium in the human alternative pathway of complement. Mol. Immunol. 23:87-96.
Reilly, B. D., S. C. Makrides, P. J. Ford, H. C. Marsh, and C. Mold. 1994. Quantitative analysis of C4b dimer binding to distinct sites on the C3b/C4b receptor (CR1). J. Biol. Chem. 269:7696-7701.
Rosengard, A. M., L. C. Alonso, L. C. Korb, W. M. Baldwin III, F. Sanfilippo, L. A. Turka, and J. M. Ahearn. 1999. Functional characterization of soluble and membrane-bound forms of vaccinia virus complement control protein (VCP). Mol. Immunol. 36:685-697.
Sahu, A., S. N. Isaacs, A. M. Soulika, and J. D. Lambris. 1998. Interaction of vaccinia virus complement control protein with human complement proteins: factor I-mediated degradation of C3b to iC3b1 inactivates the alternative complement pathway. J. Immunol. 160:5596-5604.
Sahu, A., T. R. Kozel, and M. K. Pangburn. 1994. Specificity of the thioester-containing reactive site of human C3 and its significance to complement activation. Biochem. J. 302:429-436.
Sahu, A., and J. D. Lambris. 2001. Structure and biology of complement protein C3, a connecting link between innate and acquired immunity. Immunol. Rev. 180:35-48.
Sahu, A., and M. K. Pangburn. 1993. Identification of multiple sites of interaction between heparin and the complement system. Mol. Immunol. 30:679-684.
Sahu, A., A. M. Soulika, D. Morikis, L. Spruce, W. T. Moore, and J. D. Lambris. 2000. Binding kinetics, structure-activity relationship, and biotransformation of the complement inhibitor compstatin. J. Immunol. 165:2491-2499.
Seet, B. T., J. B. Johnston, C. R. Brunetti, J. W. Barrett, H. Everett, C. Cameron, J. Sypula, S. H. Nazarian, A. Lucas, and G. McFadden. 2003. Poxviruses and immune evasion. Annu. Rev. Immunol. 21:377-423.
Sharma, A. K., and M. K. Pangburn. 1996. Identification of three physically and functionally distinct binding sites for C3b in human complement factor H by deletion mutagenesis. Proc. Natl. Acad. Sci. USA 93:10996-11001.
Smith, S. A., R. Sreenivasan, G. Krishnasamy, K. W. Judge, K. H. Murthy, S. J. Arjunwadkar, D. R. Pugh, and G. J. Kotwal. 2003. Mapping of regions within the vaccinia virus complement control protein involved in dose-dependent binding to key complement components and heparin using surface plasmon resonance. Biochim. Biophys. Acta 1650:30-39.
Soames, C. J., and R. B. Sim. 1997. Interactions between human complement components factor H, factor I and C3b. Biochem. J. 326:553-561.
Spiller, O. B., D. J. Blackbourn, L. Mark, D. G. Proctor, and A. M. Blom. 2003. Functional activity of the complement regulator encoded by Kaposi's sarcoma-associated herpesvirus. J. Biol. Chem. 278:9283-9289.
Stoiber, H., C. Speth, and M. P. Dierich. 2003. Role of complement in the control of HIV dynamics and pathogenesis. Vaccine 21(Suppl. 2):S77-S82.
Vanderplasschen, A., E. Mathew, M. Hollinshead, R. B. Sim, and G. L. Smith. 1998. Extracellular enveloped vaccinia virus is resistant to complement because of incorporation of host complement control proteins into its envelope. Proc. Natl. Acad. Sci. USA 95:7544-7549.
Warren, L. 1959. The thiobarbituric acid assay of sialic acids. J. Biol. Chem. 234:1971-1975.(Jayati Mullick, John Bern)
ABSTRACT
Vaccinia virus encodes a homolog of the human complement regulators named vaccinia virus complement control protein (VCP). It is composed of four contiguous complement control protein (CCP) domains. Previously, VCP has been shown to bind to C3b and C4b and to inactivate the classical and alternative pathway C3 convertases by accelerating the decay of the classical pathway C3 convertase and (to a limited extent) the alternative pathway C3 convertase, as well as by supporting the factor I-mediated inactivation of C3b and C4b (the subunits of C3 convertases). In this study, we have mapped the CCP domains of VCP important for its cofactor activities, decay-accelerating activities, and binding to the target proteins by utilizing a series of deletion mutants. Our data indicate the following. (i) CCPs 1 to 3 are essential for cofactor activity for C3b and C4b; however, CCP 4 also contributes to the optimal activity. (ii) CCPs 1 to 2 are enough to mediate the classical pathway decay-accelerating activity but show very minimal activity, and all the four CCPs are necessary for its efficient activity. (iii) CCPs 2 to 4 mediate the alternative pathway decay-accelerating activity. (iv) CCPs 1 to 3 are required for binding to C3b and C4b, but the presence of CCP 4 enhances the affinity for both the target proteins. These results together demonstrate that the entire length of the protein is required for VCP's various functional activities and suggests why the four-domain structure of viral CCP is conserved in poxviruses.
INTRODUCTION
The complement system is an ancient and one of the principal innate immune defense mechanisms against all invading pathogens, including viruses (7, 34). It has a unique ability to recognize and label pathogens as non-self, with and without the help of pattern recognition molecules (7, 27). Apart from its direct effect on pathogens, it also plays an additional role in initiation and enhancement of the pathogen-specific adaptive immune responses (6, 34, 42). A significant body of knowledge indicates that both acute and latent viruses are susceptible to neutralization by the complement system (3, 7, 25). Thus, complement exerts a strong selective pressure on viruses during infection. To survive as successful pathogens, viruses have evolved different strategies to subvert the complement system. Efficient complement evasion mechanisms evolved by viruses include direct complement inactivation by encoding structural and/or functional homologs of host complement regulatory proteins or by capturing host membrane regulatory proteins and using host receptors for cellular entry (3, 25). Important examples include viruses belonging to the family of poxviruses, herpesviruses, retroviruses, paramyxoviruses, and picornaviruses (1, 10, 28, 30, 33, 51, 56).
Vaccinia virus (VV), a prototype of the family Poxviridae and genus Orthopoxvirus, is a cytoplasmic large DNA virus (18, 32). VV embodies an illustrious example of complement evasion. It is known to have developed two efficient mechanisms to evade the complement system. (i) It encodes within its genome a 27-kDa protein, homologous to human complement regulators, known as VV complement control protein (VCP), which is a potent soluble complement inhibitor (22, 29, 46). (ii) It incorporates host complement proteins (membrane cofactor protein [MCP], CD46; decay-accelerating factor [DAF], CD55; and CD59) while budding, rendering itself resistant to complement attack (57).
VCP, encoded by the C21L gene, is one of the first viral proteins identified to have complement-binding activity (23). The primary structure of VCP consists of 244 amino acids with a signal peptide of 19 amino acids. It comprises four CCP domains that are typically present in human regulators of complement activation (RCA) proteins and exhibits considerable sequence similarity to factor H (26%), C4b-binding protein (37%), MCP (35%), DAF (38%), and complement receptor 1 (CR1) (37%). The nuclear magnetic resonance structure of VCP modules (CCPs 2 to 3 and CCPs 3 to 4) revealed that each CCP folds into a compact six-?-strand structure similar to CCP 16 of factor H (13). Recent elucidation of the crystal structure of VCP depicted it as a highly extended molecule with heparin-binding site at its C-terminal region (9, 36). An interesting feature revealed by the crystal structure was the charge distribution of VCP domains: CCP domains 1 and 4 are surrounded by a positive field, while the middle two domains (CCPs 2 and 3) are surrounded by a predominantly negative field.
Functional studies revealed that VCP protects vaccinia virions from antibody-dependent complement-mediated neutralization (15); vaccinia virus mutants that do not express VCP are attenuated in vivo (15). Initial studies of the mechanism of complement inhibition performed using both partially purified VCP and culture medium containing secreted VCP have shown that it inhibits the classical pathway-mediated lysis of sheep erythrocytes, binds to C3b and C4b, and accelerates the decay of the classical and (to a limited extent) the alternative pathway C3 convertases (29). A later study performed using purified recombinant VCP demonstrated that it possesses factor I cofactor activity for C3b and C4b and is a poor inhibitor of the alternative pathway, in comparison to the classical pathway (46). Recently, the binding mechanism of VCP with C3b and C4b has been characterized using surface plasmon resonance (SPR). The data suggested that binding of VCP to C3b and C4b follows a simple 1:1 binding model and does not involve multiple-site interactions, as is observed for factor H and CR1 (2). Further, binding to C3b and C4b involves ionic interactions, and the binding site of VCP on C3b and C4b is located within the C3dg and C4c regions, respectively (2).
Although significant efforts were made to establish VCP as a complement inhibitor (3, 17), no efforts were made to shed light on the identification of its domains important for its complement regulatory activities: factor I cofactor activity for C3b and C4b, and decay-accelerating activity for the classical and alternative pathway C3 convertases. In this paper, we have identified which of the four CCPs of VCP are required to confer cofactor activities, decay-accelerating activities, and binding to target proteins C3b and C4b. Our results suggest that a minimum of three CCP domains are required for its cofactor activities, binding to C3b and C4b, and its limited alternative pathway decay-accelerating activity and that two domains are enough to mediate the classical pathway decay-accelerating activity.
MATERIALS AND METHODS
Reagents and buffers. Antibody-coated sheep erythrocytes (designated EA) were prepared by incubating the sheep erythrocytes with anti-sheep erythrocyte antibodies (ICN Biomedical, Inc., Irvine, Calif.). Antiserum recognizing VCP was raised in rabbit by immunizing with purified VCP. Buffers used were VBS (5 mM barbital and 145 mM NaCl, pH 7.4), GVB (VBS containing 0.1% gelatin), DGVB (2.5 mM barbital and 73 mM NaCl, 0.1% gelatin, and 2.5% dextrose), DGVB++ (DGVB containing 0.5 mM MgCl2 and 0.15 mM CaCl2), and phosphate-buffered saline (PBS) (10 mM sodium phosphate and 145 mM NaCl, pH 7.4). Mg EGTA contained 0.1 M MgCl2 and 0.1 M EGTA.
Purified complement proteins. The human complement factors B, D, H, and I were kindly provided by Michael K. Pangburn (University of Texas Health Center, Tyler, Tex.) and the recombinant human soluble CR1 (sCR1) was a generous gift of Henry Marsh (AVANT Immunotherapeutics, Inc., Needham, Mass.). Human C1, C4, and C2 were purchased from Calbiochem (La Jolla, Calif.). Human C3 was purified according to Hammer et al. (12) with minor modifications. Twenty parts of human plasma were mixed with 1 part of inhibitor solution (1 M KH2PO4, 0.2 M Na4 EDTA, 0.2 M benzamidine, and 1 mM phenylmethylsulfonyl fluoride) and precipitated first with 4.5% polyethylene glycol and then with 12% polyethylene glycol at 0°C. The 12% pellet was then dissolved in 20 mM sodium phosphate, pH 7.4, and loaded onto a Source Q column (1.2 by 9.5 cm; Amersham Pharmacia Biotech, Uppsala, Sweden). The bound proteins were eluted with a linear gradient of 0 to 0.5 M NaCl. Fractions containing C3 were identified by sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) and immunodiffusion, pooled, and loaded onto a Mono Q 10/10 column (Amersham Pharmacia Biotech) in 10 mM sodium phosphate, pH 7.9. Bound proteins were eluted with a linear salt gradient of 0 to 0.5 M and subjected to SDS-PAGE analysis. C3-containing fractions were pooled, concentrated by ultrafiltration, and loaded onto Superose 12 (two columns connected in a series; Amersham Pharmacia Biotech) in PBS. Homogeneous C3 fractions as analyzed by SDS-PAGE were pooled. The purified C3 contained >90% native C3, as determined by analyzing the protein sample on a Mono S column (Amersham Pharmacia Biotech) (39). C3b was generated by limited trypsin cleavage and purified on a Mono Q 5/5 column (46). Human C4b used was purified as previously described (2) or purchased from Calbiochem (La Jolla, Calif.).
Construction of deletion mutants. The deletion constructs comprising CCPs 1 to 3, 2 to 4, 1 to 2, 2 to 3, and 3 to 4 of VCP were constructed from full-length VCP (46) by PCR amplification and cloned into the yeast expression vector pPICZ (Invitrogen, Carlsbad, Calif.), downstream of the AOX1 methanol-inducible promoter by the standard protocol. For amplification of the respective VCP deletion mutants, the following sequence-specific primers were used: for CCP 1-3, AS-1 (5'-GGAATTCTGCTGTACTATTCCGTCACGACCC-3') and AS-5 (5'-GCTCTAGATTATTTAACAATCTGACACGTGGGTGG-3'); for CCP 2-4, AS-2 (5'-GGAATTCTGCCCATCGCCTCGAGATATCG-3') and AS-4 (5'-GCTCTAGATTAGCGTACACATTTTGGAAGTTCCG-3'); for CCP 1-2, AS-1 and AS-6 (5'-GCTCTAGATTATTTAACAGATTCACAAATAGGTGCC-3'); for CCP 2-3, AS-2 and AS-5; and for CCP 3-4, AS-3 (5'-GGAATTCTGCCAATCCCCTCCATCTATATCCAACGG-3') and AS-4. The primers incorporated restriction sites EcoRI and XbaI (underlined) in the forward and the reverse orientations, respectively, and a stop codon (in boldface type) in the reverse orientation primers. The amplification was carried out using Titanium Taq DNA polymerase (B. D. Biosciences Clonetech, Palo Alto, CA), with an initial cycle of denaturation at 94°C for 2 min, followed by 25 cycles of 1 min at 94°C, 1 min at 60°C or 65°C according to the annealing temperature of the primer, 1 min at 72°C, and a final extension for 10 min at 72°C. All the PCR products were digested with EcoRI and XbaI and cloned into pPICZ EcoRI and XbaI cut vector. The validity of all the constructs was confirmed by automated DNA sequencing. Five micrograms of each of the SacI-digested pPICZ VCP deletion constructs was then integrated into Pichia pastoris as per the manufacturer's protocol. The integration of the deletion mutants in Pichia were confirmed by amplifying the respective genomic DNA using the 5'AOX1 and 3'AOX1 primers and with a combination of gene-specific primers. Further, the validated integrants were used for expression of the recombinant proteins.
Expression and purification of vaccinia virus complement control protein and its deletion mutants. The vaccinia virus complement control protein cloned in Pichia pastoris was expressed and purified as previously described (2). Expression of VCP mutants was performed as described below. A single colony of recombinant Pichia pastoris expressing the respective VCP mutant was inoculated in 10 ml of BMGY medium (100 mM potassium phosphate, pH 6.0, 10 g/liter yeast extract, 20 g/liter peptone, 13.4 g/liter yeast nitrogen base, 0.4 mg/liter biotin, and 1% glycerol) and incubated overnight at 30°C in a shaking incubator. This inoculum was then added to 400 ml of BMGY and incubated for 48 h at 30°C with shaking. The cells were centrifuged, resuspended in 400 ml of BMMY (BMGY containing 0.5% methanol, but without 1% glycerol), and incubated at 30°C for an additional 96 h with vigorous shaking; 0.5% methanol was added after every 24 h. After incubation, cells were pelleted and the supernatant containing the VCP mutant was collected for purification.
For purification, the supernatants containing VCP mutants were concentrated by ultrafiltration and precipitated with 80% ammonium sulfate at 0°C. The pellets were then suspended and dialyzed in PBS. For further purification, CCP 1-3 and CCP 1-2 mutants were passed through a PD-10 column (Amersham Pharmacia Biotech) in 10 mM sodium phosphate (pH 7.4), loaded onto a heparin-agarose column in the same buffer, and eluted with 500 mM NaCl. The eluted proteins were then exchanged into 20 mM Tris (pH 7.9), loaded onto a Mono Q column, and eluted with a linear gradient of 0 to 500 mM NaCl. For purification of CCP 2-4 and CCP 2-3 mutants, the samples were passed through DEAE-Sephacel (Sigma, St. Louis, Mo.) preequilibrated with 10 mM sodium phosphate, pH 7.4, containing 150 mM NaCl. The flowthrough was collected, passed through a PD-10 column in 20 mM Tris (pH 7.9), and loaded onto a Mono Q column. The bound proteins were eluted with a linear gradient of 0 to 500 mM NaCl. CCP 3-4 mutant was purified by being loaded onto DEAE-Sephacel in 10 mM sodium phosphate, pH 7.4, containing 150 mM NaCl. The flowthrough containing CCP 3-4 mutant was collected and buffer exchange was performed using a PD-10 column preequilibrated with 5 mM sodium acetate, pH 4.0. The sample was then loaded onto a Mono S column, and the bound proteins were eluted with a linear salt gradient of 0 to 500 mM NaCl. Eluted fractions in all the above purifications were analyzed by SDS-PAGE and Western blotting using anti-VCP antibodies; fractions containing pure proteins were pooled, dialyzed into PBS, and concentrated by ultrafiltration.
Mass analysis and protein sequencing. The molecular mass determination and sequencing of the VCP mutants were performed at the Biomolecular Research Facility (University of Virginia, Charlottesville, VA) as described below. The molecular weights of the VCP mutants were determined by matrix-assisted laser desorption ionization-time of flight mass spectrometry (MS) using an Applied Biosystems Voyager DE-Pro. The samples were dissolved in 70% acetonitrile-30% water, mixed with sinapinic acid matrix, and spotted on the matrix-assisted laser desorption ionization plate. Approximately 200 spectra were averaged for each sample.
The sequencing of VCP mutants was performed as described below. The samples were run on a 12% SDS-PAGE gel and stained with Coomassie blue. The protein bands were cut, transferred to a siliconized tube, washed, and destained overnight in 200 μl of 50% methanol. The gel pieces were then dehydrated in acetonitrile, rehydrated in 30 μl of 10 mM dithiolthreitol (DTT) in 0.1 M ammonium bicarbonate, and reduced at room temperature for 30 min. The DTT solution was then removed, and the samples were alkylated in 30 μl of 50 mM iodoacetamide in 0.1 M ammonium bicarbonate at room temperature for 30 min. The reagent was removed, and the gel pieces were dehydrated in 100 μl of acetonitrile. The gel pieces were subjected to two cycles of dehydration and rehydration in acetonitrile and 0.1 M ammonium bicarbonate and then dried by vacuum centrifugation. The gel pieces were rehydrated in 20 ng/μl trypsin (second band for each sample in chymotrypsin) in 50 mM ammonium bicarbonate on ice for 10 min, excess enzyme solution was removed, and 20 μl of 50 mM ammonium bicarbonate was added. The samples were digested overnight at 37°C, and the peptides formed were extracted from the polyacrylamide in two 30-μl aliquots of 50% acetonitrile-5% formic acid. These extracts were combined and evaporated to 25 μl for MS analysis, which was done using a liquid chromatography-MS system consisting of a Finnigan LCQ ion trap mass spectrometer with a Protana nanospray ion source interfaced to a self-packed 8 cm by 75 μm (inner diameter) Phenomenex Jupiter 10-μm C18 reversed-phase capillary column. The digest was analyzed using the double-play capability of the instrument acquiring one full scan mass spectrum to determine peptide molecular weights, followed by four product ion spectra to determine amino acid sequences in sequential scans. The data were analyzed by database searching using the Sequest algorithm and by manual interpretation.
SPR measurements. The kinetics of binding of VCP and VCP mutants to human C3b and C4b were determined by using the SPR-based biosensor BIACORE 2000 (Biacore AB, Uppsala, Sweden). Binding experiments were performed using physiologic ionic strength buffer (PBS-T, 10 mM sodium phosphate, 145 mM NaCl, pH 7.4, containing 0.05% Tween-20) at 25°C. Addition of 0.05% Tween-20 blocked the nonspecific adsorption of VCP to the sensor chips (50). In this assay, both C3b and C4b were oriented in their physiological orientation on a streptavidin chip (Sensor Chip SA, Biacore AB) by labeling their free -SH group with biotin (2). In brief, 914 response units (RUs) of C3b and 916 RUs of C4b were immobilized on FC-2 and FC-3, respectively, and FC-1 served as a control flow cell. Binding was measured at 50 μl/min to avoid the mass transport effect, and it was measured for 120 s. Dissociation was measured for 180 s. The sensor chip surfaces were regenerated with 30-s pulses of 0.2 M sodium carbonate, pH 9.5. Sensograms obtained for the control flow cell (Fc-1) were subtracted from the data for the flow cell immobilized with either C3b or C4b, and the SPR data were analyzed by BIAevaluation software version 4.1 using global fitting.
Measurement of factor I cofactor activity. Both qualitative and quantitative analyses of factor I cofactor activity of VCP and its mutants for C3b and C4b were determined using 10 mM phosphate buffer, pH 7.4, containing 145 mM NaCl. In these assays, 2.7 μg of C3b or 2.9 μg of C4b was mixed with 100 ng of factor I and VCP (as indicated in the figure legends) or its mutants and incubated at 37°C for 4 h in a total volume of 15 μl. The reactions were stopped by the addition of sample buffer containing DTT and electrophoresed on a 9% SDS-PAGE gel for determining C3b cleavages and a 10% SDS-PAGE for C4b cleavages. The cleavage products were visualized by staining the gel with Coomassie blue. For quantitative analysis, gels were scanned for densitometric analysis by using the ChemiDoc XRS system (Bio-Rad, Segrate, Italy) to calculate the percentage of '-chain. The data obtained were normalized by considering 100% '-chain to be equal to the '-chain intensity obtained in the absence of factor I (control). The binding data were fit using nonlinear regression analysis (GraFit; Erithacus Software, London, United Kingdom) and a four-parameter logistic analysis was performed to identify the best-fit 50% inhibitory concentration value (the concentration of VCP causing 50% '-chain cleavage).
Measurement of C3 convertase decay-accelerating activity. The classical pathway decay-accelerating activity of VCP and its mutants was determined by forming EAC142 (37). In brief, the classical pathway C3 convertase EAC142 was made by incubating 100 μl of EA (4 x 109/ml) in DGVB++ with 4 μg C1 for 20 min at 30°C in a total volume of 120 μl. The cells were then washed with ice-cold DGVB++, resuspended in 100 μl DGVB++, and incubated for 20 min at 30°C with 8 μg of C4. After the incubation, they were mixed with 4 μg of C2 and further incubated for 4 min at 30°C to form EAC142. The convertase formation was then stopped by adding EDTA to a final concentration of 10 mM. To define the effect of VCP and its mutants on the decay of EAC142, 10 μl of EAC142 (109/ml) was mixed with various concentrations of VCP or its mutants in a total volume of 25 μl for 5 min at 22°C. The reaction mixture was then mixed with 125 μl of DGVB and 100 μl of guinea pig serum diluted 1:100 in DGVB containing 40 mM EDTA, incubated 30 min at 37°C, and centrifuged. The percentage of lysis was determined by measuring the absorbance at 405 nM.
The alternative pathway C3 convertase decay was measured by forming C3b,Bb on rabbit (ER) or sheep (ES) erythrocytes. ERC3b and ESC3b were generated by incubating ER or ES with C3, factor B, and factor D in the presence of NiCl2 as previously described (43, 47). C3b,Bb on ER or ES was made by mixing 150 μl of C3b-coated rabbit or sheep erythrocytes (109/ml) with 45 μg of factor B and 0.76 μg of factor D in 210 μl of GVB containing 3.5 mM NiCl2. The reaction mixture was then incubated for 5 min at 30°C and stopped by the addition of EDTA to a final concentration of 10 mM. Immediately thereafter, 10 μl of ERC3b,Bb or ESC3b,Bb (109/ml) was mixed with various concentrations of VCP or its mutants in a total volume of 50 μl and incubated for 10 min at 37°C. The reaction mixture was then mixed with 50 μl of human serum diluted 1:5 in GVB containing 20 mM EDTA and incubated at 37°C for 20 min. After incubation, the reaction mixture was mixed with 200 μl of GVBE and centrifuged. The percentage of lysis was determined as above. The data generated were normalized by setting 100% lysis equal to the lysis that occurred in the absence of an inhibitor.
RESULTS
Design, expression, purification, and characterization of VCP deletion mutants. Three aspects were taken into consideration while designing the mutants. First, since VCP is a soluble protein, mutants were expressed as soluble proteins and not as membrane-bound fusion proteins as described in an earlier study, where attempts were made to localize the C3b-binding site (45). Second, it has previously been shown for the human complement regulators that apart from the CCP domains, the interdomain linker regions may also play a role in the ligand binding (14); therefore, each construct started with the first Cys of the starting domain and ended with the last residue of the interdomain linker. This design kept the entire linker region at the C-terminal side of each mutant (see the amino acid sequences listed in Table 1). Third, several previous studies on the human regulators have shown that two or three contiguous domains (5, 14, 19), as well as their native configuration (i.e., distance between the successive domains) (4, 24), is essential for their complement regulatory activities; therefore, mutants designed contained either two or three successive CCP domains.
Earlier, we and others have shown that the Pichia expression system produces a high yield of properly folded VCP (36, 46). Thus, we utilized the same expression system for expressing VCP mutants. The culture supernatants containing the expressed mutants were purified by using a series of chromatographic procedures as described in Materials and Methods. The purified mutants were homogeneous and >95% pure as determined by SDS-PAGE analysis (Fig. 1). To confirm the identity of the mutants, they were subjected to sequencing by mass spectrometry. The amino acid sequences of the expressed mutants were consistent with the predicted sequences confirming the identity of each mutant (Table 1). The addition of EAEAEF in the beginning of each mutant was the result of cloning into the Pichia vector. The molecular masses of the expressed mutants were analyzed by SDS-PAGE and mass spectrometry. In SDS-PAGE, all the proteins migrated higher than their molecular weights; however, this was not due to differences in glycosylation because the primary sequence of VCP does not contain any glycosylation site (23). The data obtained using mass spectrometry confirmed that the molecular masses of the mutants were similar to the calculated masses (Table 1).
Characterization of factor I cofactor activity of VCP mutants. A fluid-phase assay was utilized to characterize the factor I cofactor activity of VCP mutants for C3b and C4b. In this assay, C3b or C4b was incubated with the mutant proteins and factor I, and cleavage of the '-chain was assessed by running the samples on SDS-PAGE gels. VCP and the CCP 1-3 mutant supported cleavage of the '-chain of C3b (Fig. 2), as well as C4b (Fig. 3), but none of the other mutants showed any detectable activity. These data indicate that CCPs 1 to 3 comprise the minimum region required for the factor I cofactor activity for both C3b and C4b. Incubation of the CCP 1-3 mutant with C3b and factor I did not result in complete cleavage of the '-chain of C3b (Fig. 2). We therefore sought to analyze the relative factor I cofactor activity of VCP and the CCP 1-3 mutant for C3b and C4b. In this assay, C3b or C4b was incubated with factor I and increasing concentrations of either VCP or CCP 1-3 mutant. Figure 4 shows that higher concentrations of the CCP 1-3 mutant than that of VCP were required to cleave the '-chains of C3b and C4b. The concentrations of VCP required to cleave 50% of the '-chain of C3b and C4b were 0.12 μM and 0.038 μM, respectively, while the concentrations of the CCP 1-3 mutant required to cleave 50% of the '-chain of C3b and C4b were 4.9 μM and 0.58 μM, respectively. These data indicate that in comparison to VCP, the CCP 1-3 mutant has 41- and 15-fold-lower factor I cofactor activity for C3b and C4b, respectively.
Characterization of decay-accelerating activity of VCP mutants. A previous study had demonstrated that apart from factor I cofactor activity, VCP also possesses significant decay-accelerating activity for the classical and limited decay-accelerating activity for the alternative pathway C3 convertases (29). To identify the domains required for the decay-accelerating activity, we measured the activity of VCP mutants using hemolytic assays. For measuring the decay-accelerating activity for the classical pathway C3 convertase C4b,2a, the enzyme was formed on the sensitized sheep erythrocytes and allowed to incubate with the mutant proteins. The remaining C3 convertase activity was quantitated by measuring hemolysis after adding EDTA sera (a source of C3 to C9). VCP showed significant decay-accelerating activity even at a 100 nM concentration. On a molar basis, VCP was only 1.8-fold-less active than sCR1. Among the mutants, CCP 1-3 and CCP 1-2 mutants showed decay-accelerating activity, while the other mutants did not show any activity (Fig. 5), indicating that CCPs 1 to 2 comprise the minimum region required for the classical pathway. It should be pointed out here that the concentration of CCP 1-2 mutant required to inhibit the decay-accelerating activity was 5,600-fold higher than the concentration of VCP (Fig. 5), suggesting that in addition to CCPs 1 to 2, CCPs 3 and 4 also contribute significantly to the classical pathway decay-accelerating activity.
For measuring the alternative pathway decay-accelerating activity of VCP and its mutants, the alternative pathway C3 convertase C3b,Bb was formed on rabbit erythrocytes. The enzyme was incubated with VCP or the mutant proteins, and the remaining enzyme activity was detected by adding EDTA sera and measuring hemolysis. Surprisingly, neither VCP nor its mutants showed any alternative pathway decay-accelerating activity even at a 50 μM concentration. In a parallel assay, factor H and sCR1 showed significant inhibition at 100 nM and 2.5 nM concentrations, respectively (Fig. 6A). Since the previous study which reported limited alternative pathway decay-accelerating activity of VCP utilized sheep erythrocytes for forming C3b,Bb (29) and not the rabbit erythrocytes, we tested if the lack of inhibition in our assays was due to the differences in the cells used. Thus, we formed C3b,Bb on sheep erythrocytes and measured the decay-accelerating activity of VCP and its mutants. On sheep cells, VCP accelerated the decay of alternative pathway C3 convertase, though the concentration of VCP required to decay 50% alternative pathway C3 convertase was 64 μM, which was 2,400-fold higher than that required to decay the classical pathway C3 convertase (Fig. 6B). Among VCP mutants, only the CCP 2-4 mutant showed inhibition, although like VCP at a similarly high concentration, suggesting that the limited AP decay-accelerating activity is imparted by CCPs 2 to 4.
Effect of membrane sialic acid on the decay-accelerating activity of VCP. From the data presented in Fig. 6, it is clear that VCP shows limited decay-accelerating activity against the alternative pathway C3 convertase present on sheep erythrocytes but not against the convertase present on rabbit erythrocytes. Previously, differences in the decay-accelerating activity against the alternative pathway C3 convertase present on sheep versus rabbit erythrocytes have been reported for human complement regulator factor H. It has been shown that factor H displays enhanced decay-accelerating activity towards the alternative pathway C3 convertase present on sheep erythrocytes; this has been attributed to the interaction of its heparin-polyanion-binding site with sialic acid present on the erythrocyte membrane (rabbit erythrocytes contain approximately 13% of the sialic acid present on sheep erythrocytes) (8, 40). Since VCP contains a heparin-binding site on CCP 4 (9), we sought to analyze if the differences in its decay activity on sheep versus rabbit erythrocytes were due to interaction of its heparin-polyanion-binding site with sialic acid present on sheep erythrocytes. For this purpose, we treated sheep erythrocytes with neuraminidase and compared the decay-accelerating activity of VCP against the C3 convertase present on untreated and treated erythrocytes; factor H was included as a positive control. As expected, factor H showed reduced alternative pathway C3 convertase decay-accelerating activity on neuraminidase-treated cells compared to untreated cells (Fig. 7); the decay activity was 7.4-fold less on the treated than on the untreated sheep cells. However, VCP did not show any difference in the decay activity, thereby indicating that like factor H, the heparin-binding site on VCP does not help in enhancing the decay of the alternative pathway C3 convertase on sheep cells. Thus, the observed differences in the decay activities of VCP on sheep and rabbit erythrocytes were not due to the differences in the membrane sialic acid content.
Binding of VCP mutants to C3b and C4b. From the data presented above, it is clear that CCPs 1 to 2 and 2 to 4 are the minimum regions required for the decay-accelerating activities and that CCPs 1 to 3 are required for cofactor activities. To determine whether reduced decay-accelerating activities and cofactor activities of the VCP mutants correlate with binding to C3b and C4b, we performed a quantitative analysis of binding of VCP and its mutants to C3b and C4b using a surface plasmon resonance assay. In this assay, C3b and C4b were oriented on a streptavidin chip by labeling their free -SH groups with biotin (2). This setup mimicked the physiological orientation of these proteins and also allowed accurate determination of the binding constants (2). When VCP or its deletion mutants were allowed to flow over the sensor chip, only VCP and the CCP 1-3 mutant bound to C3b and C4b (Fig. 8). Further analysis showed that the CCP 1-3 mutant had 36- and 228-fold-lower affinity for C3b and C4b, respectively, than VCP (Fig. 9 and Table 2). This decrease was primarily due to reduction in the on-rates of the mutant (Table 2). These data suggest that poor factor I cofactor activities of the CCP 1-3 mutant could be due to its lower affinity for C3b and C4b. It is, however, interesting that the CCP 1-2 and CCP 2-4 mutants, which showed the classical and alternative pathway decay-accelerating activities, respectively, showed no detectable binding to C4b and C3b.
DISCUSSION
In the present study, we utilized a series of VCP deletion mutants to address the contribution of each CCP domain to its cofactor activities, decay-accelerating activities, and binding to C3b and C4b. Our data on the localization of domains required for factor I cofactor activity for C3b and C4b indicate that a minimum of three CCPs (CCPs 1 to 3) are essential for these activities, because further deletion of either CCP 1 or CCP 3 completely abolished the cofactor activities (Fig. 2 and 3). These data are consistent with the previous observations on mapping of the human RCA proteins (factor H, C4bp, MCP, and CR1) for cofactor activities, which indicated that a minimum of three CCPs are required for cofactor activities (4, 11, 14, 16, 19, 20, 44, 52). There is now a consensus that factor I-mediated inactivation of C3b and C4b involves two steps: (i) recognition of C3b or C4b by the cofactor and (ii) interaction of factor I with C3b or C4b and the cofactor (48, 54). Thus, it is conceivable that the recognition sites for C3b or C4b and factor I on the complement regulators are spatially conserved and require three CCPs. This, however, does not seem to be a general phenomenon as our recent data on kaposica, the complement regulator of the Kaposi's sarcoma-associated herpesvirus, indicated that only two CCPs are enough to impart cofactor activities in kaposica (35). Since CCPs 1 to 3 are required for both C3b and C4b cofactor activities, it can be envisaged that the recognition site for C3b and C4b on VCP is formed by similar residues. This possibility, however, appears to be less likely for VCP as it binds to different regions on C3b and C4b; its binding site on C3b is located on the C3dg region, while its binding site on C4b is located on the C4c region (2).
VCP has previously been shown to possess efficient decay-accelerating activity for the classical pathway and weak decay activity for the alternative pathway C3 convertases (29). Our data show that CCPs 1 to 2 of VCP form the minimum region essential for its classical pathway decay-accelerating activity, but CCPs 3 and 4 are required for its complete activity (Fig. 5). Earlier studies of DAF showed that CCPs 2 to 3 are important for its classical pathway decay-accelerating activity (5). It is pertinent to point out here that CCPs 1 to 2 of VCP show maximum sequence similarity to CCPs 2 to 3 of DAF, which possesses classical pathway decay-accelerating activity (5). Our data show that VCP possesses limited decay-accelerating activity only against the alternative pathway C3 convertase present on the sheep erythrocytes and not against the C3 convertase present on rabbit erythrocytes (Fig. 6). Since VCP contains a heparin-binding site like factor H (9), it is likely that similar to factor H (31, 49), the heparin-binding site on VCP helps it to enhance its decay-accelerating activity against the alternative pathway C3 convertase present on the sheep erythrocytes, which contains a high surface density of sialic acid. However, this premise did not hold true, as the decay rate of VCP did not reduce on neuraminidase-treated sheep erythrocytes (Fig. 7). The weak decay activity was also observed with the CCP 2-4 mutant, suggesting that CCPs 2 to 4 are important for this activity. Like VCP, the Kaposi's sarcoma-associated herpesvirus complement control protein kaposica also contains very weak decay-accelerating activity against alternative pathway C3 convertase (35, 55). Thus, it seems that viral homologs of complement control proteins primarily contain three out of four C3 convertase regulatory activities of the host complement regulators.
Our data on binding of VCP to target proteins measured using SPR identified CCPs 1 to 3 as the smallest structural unit capable of binding to C3b and C4b (Fig. 8). The presence of CCP 4, however, significantly enhanced the affinities for C3b and C4b (Fig. 9 and Table 2). These findings are consistent with the previous data on binding of the human RCA proteins to C3b and C4b, which essentially suggested that C3b or C4b binding primarily resides within three or four CCPs (4, 14, 19, 20, 52). Previously, two studies attempted to localize the C3b-binding site on VCP. Both these studies suggested that all the four CCPs are required for C3b binding. In the first study, various domains of VCP were expressed as membrane-bound CR2 fusion proteins (45). The lack of binding of the CCP 1-3 mutant to C3b in this study (as opposed to our present study) could have been due to lower sensitivity of the assay used. Alternatively, it is possible that the fusion of CCP 3 to CR2 could have sterically hindered the access to residues present on CCP 3, which are critical for binding to C3b. Unlike the first study, the second study utilized soluble proteins and SPR technology for detecting binding but failed to test binding of the CCP 1-3 mutant to C3b due to lack of this mutant (53).
Even though ligand binding is a prerequisite for cofactor and decay activities, a large body of evidence indicates that ligand binding does not always correlate well with cofactor and decay activities (26, 41). Our data support this premise. For example, the CCP 1-3 mutant showed a 36-fold decrease in affinity for C3b and a 228-fold decrease in affinity for C4b compared to VCP (Table 2). Although the decrease in C3b binding of the CCP 1-3 mutant correlated well with a 41-fold reduction in its C3b cofactor activity, the dramatic reduction in its affinity for C4b did not correlate with the 15-fold decrease in C4b cofactor activity. Similarly, CCP 1-2 and CCP 2-4 mutants which exhibited the classical and alternative pathway decay-accelerating activities, respectively, did not show detectable binding to C4b and C3b. It is likely that like DAF, these mutants might have better affinity for the C3 convertases than for their subunits C4b and C3b; DAF has been shown to bind to C4b,2a with 1,000-fold higher affinity than to C4b alone (38).
The data discussed above point out that although only two or three CCPs are enough to mediate the cofactor and decay activities of VCP, the entire length of the protein is essential for its full functional activity. It is likely that the residues scattered on two or three CCPs together form a binding site for C3 convertases or their subunits (C3b or C4b) and that the other CCPs contribute to the conformational stability of the complexes. Alternatively, a larger surface spanning all the four domains of VCP contributes to forming the recognition site for the convertases and C3b or C4b. These possibilities assume that functional domains are arranged in a linear fashion. This, however, may not be true; it is also likely that the recognition sites are discontinuous and that the middle domains or regions serve as essential spacers. These data, along with our recent observation on kaposica, which also indicated that the entire length of the protein is important for its complete function (35), explain why the four-CCP structure of viral CCP is highly conserved in viruses.
The important question in vaccinia virus biology is how VCP helps to protect the vaccinia virus from complement-mediated neutralization in vivo. The complement system is known to neutralize viruses in the presence and the absence of specific antibodies. Previously, it has been shown that extracellular enveloped virus, which is important for the virus dissemination, is protected from the alternative pathway due to the presence of host complement regulators on its envelope (57). However, this form remains susceptible to classical pathway-mediated neutralization (57). Thus, vaccinia virus must encode an effective regulator against the classical pathway to overcome the classical pathway-mediated neutralization. Earlier studies have shown VCP to be a better regulator of the classical pathway than even the host complement regulators factor H and C4-binding protein (21, 46); in the present manuscript, we have shown that VCP is only twofold-less effective than sCR1 in decaying the classical pathway C3 convertase (Fig. 5). The question is whether this inhibition of the classical pathway is also seen in vivo? In an earlier in vivo study, it was shown that the lesions formed by the VCP deletion mutant and the wild-type virus were similar for the first few days, but the lesion size was significantly reduced after 5 days in the mutant virus (15). Further, the appearance of antibody capable of mediating complement-enhanced neutralization correlated well with this timing (15), suggesting that VCP effectively inhibits classical pathway-mediated neutralization in vivo. In the present manuscript, we have shown which of the domains of VCP are required for each of its complement regulatory activities. Further delineation of the vital control points in VCP important for inhibition of the classical pathway decay-accelerating activity and C4b cofactor activity and targeting these control points would allow evaluation of the relative importance of these activities in immune evasion and may provide an alternative approach to manage vaccinia virus vaccine-related complications.
ACKNOWLEDGMENTS
We thank John D. Lambris (Department of Pathology and Laboratory Medicine, University of Pennsylvania, Philadelphia, PA) for his continuous support; Michael K. Pangburn (Department of Biochemistry, University of Texas Health Science Center, Tyler, Texas) for his support and the generous gift of complement proteins factors B, D, H, and I; Henry Marsh (AVANT Immunotherapeutics, Inc., Needham, Mass.) for providing sCR1; Nicholas E. Sherman (Biomolecular Research Facility, University of Virginia, Charlottsville, Va.) for sequencing VCP mutants; and Gabriela Canziani (Protein Interaction Facility, University of Utah, Salt Lake City, Utah) for advice on maintenance of Biacore.
This work was supported by a Wellcome Trust Senior Research Fellowship in Biomedical Science in India to A.S. The authors also acknowledge financial assistance to A.K.S. by the Council of Scientific and Industrial Research of India.
J.M. and J.B. contributed equally to this work.
REFERENCES
Alcami, A., and U. H. Koszinowski. 2000. Viral mechanisms of immune evasion. Trends Microbiol. 8:410-418.
Bernet, J., J. Mullick, Y. Panse, P. B. Parab, and A. Sahu. 2004. Kinetic analysis of the interactions between vaccinia virus complement control protein and human complement proteins C3b and C4b. J. Virol. 78:9446-9457.
Bernet, J., J. Mullick, A. K. Singh, and A. Sahu. 2003. Viral mimicry of the complement system. J. Biosci. 28:249-264.
Blom, A. M., L. Kask, and B. Dahlback. 2001. Structural requirements for the complement regulatory activities of C4BP. J. Biol. Chem. 276:27136-27144.
Brodbeck, W. G., D. Liu, J. Sperry, C. Mold, and M. E. Medof. 1996. Localization of classical and alternative pathway regulatory activity within the decay-accelerating factor. J. Immunol. 156:2528-2533.
Carroll, M. C. 2004. The complement system in regulation of adaptive immunity. Nat. Immunol. 5:981-986.
Cooper, N. R. 1998. Complement and viruses, p. 393-407. In J. E. Volanakis and M. M. Frank (ed.), The human complement system in health and disease. Marcel Dekker, Inc., New York, N.Y.
Fearon, D. T. 1978. Regulation of membrane sialic acid of ?1H-dependent decay-dissociation of amplification C3 convertase of the alternative complement pathway. Proc. Natl. Acad. Sci. USA 75:1971-1978.
Ganesh, V. K., S. A. Smith, G. J. Kotwal, and K. H. Murthy. 2004. Structure of vaccinia complement protein in complex with heparin and potential implications for complement regulation. Proc. Natl. Acad. Sci. USA 101:8924-8929.
Gewurz, B. E., R. Gaudet, D. Tortorella, E. W. Wang, and H. L. Ploegh. 2001. Virus subversion of immunity: a structural perspective. Curr. Opin. Immunol. 13:442-450.
Gordon, D. L., R. M. Kaufman, T. K. Blackmore, J. Kwong, and D. M. Lublin. 1995. Identification of complement regulatory domains in human factor H. J. Immunol. 155:348-356.
Hammer, C. H., G. H. Wirtz, L. Renfer, H. D. Gresham, and B. F. Tack. 1981. Large scale isolation of functionally active components of the human complement system. J. Biol. Chem. 256:3995-4006.
Henderson, C. E., K. Bromek, N. P. Mullin, B. O. Smith, D. Uhrin, and P. N. Barlow. 2001. Solution structure and dynamics of the central CCP module pair of a poxvirus complement control protein. J. Mol. Biol. 307:323-339.
Hourcade, D., M. K. Liszewski, M. Krych-Goldberg, and J. P. Atkinson. 2000. Functional domains, structural variations and pathogen interactions of MCP, DAF and CR1. Immunopharmacology 49:103-116.
Isaacs, S. N., G. J. Kotwal, and B. Moss. 1992. Vaccinia virus complement-control protein prevents antibody-dependent complement-enhanced neutralization of infectivity and contributes to virulence. Proc. Natl. Acad. Sci. USA 89:628-632.
Iwata, K., T. Seya, Y. Yanagi, J. M. Pesando, P. M. Johnson, M. Okabe, S. Ueda, H. Ariga, and S. Nagasawa. 1995. Diversity of sites for measles virus binding and for inactivation of complement C3b and C4b on membrane cofactor protein CD46. J. Biol. Chem. 270:15148-15152.
Jha, P., and G. J. Kotwal. 2003. Vaccinia complement control protein: multi-functional protein and a potential wonder drug. J. Biosci. 28:265-271.
Johnston, J. B., and G. McFadden. 2003. Poxvirus immunomodulatory strategies: current perspectives. J. Virol. 77:6093-6100.
Jokiranta, T. S., J. Hellwage, V. Koistinen, P. F. Zipfel, and S. Meri. 2000. Each of the three binding sites on complement factor H interacts with a distinct site on C3b. J. Biol. Chem. 275:27657-27662.
Kalli, K. R., P. H. Hsu, T. J. Bartow, J. M. Ahearn, A. K. Matsumoto, L. B. Klickstein, and D. T. Fearon. 1992. Mapping of the C3b-binding site of CR1 and construction of a (CR1)2-F(ab')2 chimeric complement inhibitor. J. Exp. Med. 174:1451-1460.
Kotwal, G. J. 1994. Purification of virokines using ultrafiltration. Am. Biotechnol. Lab. 12:76-77.
Kotwal, G. J., S. N. Isaacs, R. Mckenzie, M. M. Frank, and B. Moss. 1990. Inhibition of the complement cascade by the major secretory protein of vaccinia virus. Science 250:827-830.
Kotwal, G. J., and B. Moss. 1988. Vaccinia virus encodes a secretory polypeptide structurally related to complement control proteins. Nature 335:176-178.
Kuhn, S., C. Skerka, and P. F. Zipfel. 1995. Mapping of the complement regulatory domains in the human factor H-like protein 1 and in factor H. J. Immunol. 155:5663-5670.
Lachmann, P. J. 2002. Microbial subversion of the immune response. Proc. Natl. Acad. Sci. USA 99:8461-8462.
Lambris, J. D., A. Sahu, and R. Wetsel. 1998. The chemistry and biology of C3, C4, and C5, p. 83-118. In J. E. Volanakis and M. Frank (ed.), The human complement system in health and disease. Marcel Dekker, Inc., New York, N.Y.
Lu, J., C. Teh, U. Kishore, and K. B. Reid. 2002. Collectins and ficolins: sugar pattern recognition molecules of the mammalian innate immune system. Biochim. Biophys. Acta 1572:387-400.
Lubinski, J., T. Nagashunmugam, and H. M. Friedman. 1998. Viral interference with antibody and complement. Semin. Cell Dev. Biol. 9:329-337.
Mckenzie, R., G. J. Kotwal, B. Moss, C. H. Hammer, and M. M. Frank. 1992. Regulation of complement activity by vaccinia virus complement-control protein. J. Infect. Dis. 166:1245-1250.
Means, R. E., S. M. Lang, Y. H. Chung, and J. U. Jung. 2002. Kaposi's sarcoma associated herpesvirus immune evasion strategies. Front. Biosci. 7:e185-e203.
Meri, S., and M. K. Pangburn. 1990. Discrimination between activators and nonactivators of the alternative pathway of complement: regulation via a sialic acid/polyanion binding site on factor H. Proc. Natl. Acad. Sci. USA 87:3982-3986.
Moss, B. 2001. Poxviridae: the viruses and their replication, p. 2849-2883. In D. M. Knipe, P. M. Howley, D. E. Griffin, R. A. Lamb, M. A. Martin, B. Roizman, and S. E. Straus (ed.), Fields virology, 4th ed. Lippincott Williams and Wilkins, Philadelphia, Pa.
Mullick, J., J. Bernet, A. K. Singh, J. D. Lambris, and A. Sahu. 2003. Kaposi's sarcoma-associated herpesvirus (human herpesvirus-8) open reading frame 4 protein (kaposica) is a functional homolog of complement control proteins. J. Virol. 77:3878-3881.
Mullick, J., A. Kadam, and A. Sahu. 2003. Herpes and pox viral complement control proteins: ‘the mask of self.’ Trends Immunol. 24:500-507.
Mullick, J., A. K. Singh, Y. Panse, V. Yadav, J. Bernet, and A. Sahu. 2005. Identification of functional domains in kaposica, the complement control protein homolog of Kaposi's sarcoma-associated herpesvirus (human herpesvirus-8). J. Virol. 79:5850-5856.
Murthy, K. H., S. A. Smith, V. K. Ganesh, K. W. Judge, N. Mullin, P. N. Barlow, C. M. Ogata, and G. J. Kotwal. 2001. Crystal structure of a complement control protein that regulates both pathways of complement activation and binds heparan sulfate proteoglycans. Cell 104:301-311.
Pan, Q., R. O. Ebanks, and D. E. Isenman. 2000. Two clusters of acidic amino acids near the NH2 terminus of complement component C4 alpha'-chain are important for C2 binding. J. Immunol. 165:2518-2527.
Pangburn, M. K. 1986. Differences between the binding sites of the complement regulatory proteins DAF, CR1, and factor H on C3 convertases. J. Immunol. 136:2216-2221.
Pangburn, M. K. 1987. A fluorimetric assay for native C3. The hemolytically active form of the third component of human complement. J. Immunol. Methods 102:7-14.
Pangburn, M. K., and H. J. Müller-Eberhard. 1978. Complement C3 convertase: cell surface restriction of ?1H control and generation of restriction on neuraminidase-treated cells. Proc. Natl. Acad. Sci. USA 75:2416-2420.
Pangburn, M. K., K. L. Pangburn, V. Koistinen, S. Meri, and A. K. Sharma. 2000. Molecular mechanisms of target recognition in an innate immune system: interactions among factor H, C3b, and target in the alternative pathway of human complement. J. Immunol. 164:4742-4751.
Ploegh, H. L. 1998. Viral strategies of immune evasion. Science 280:248-253.
Pryzdial, E. L., and D. E. Isenman. 1986. A reexamination of the role of magnesium in the human alternative pathway of complement. Mol. Immunol. 23:87-96.
Reilly, B. D., S. C. Makrides, P. J. Ford, H. C. Marsh, and C. Mold. 1994. Quantitative analysis of C4b dimer binding to distinct sites on the C3b/C4b receptor (CR1). J. Biol. Chem. 269:7696-7701.
Rosengard, A. M., L. C. Alonso, L. C. Korb, W. M. Baldwin III, F. Sanfilippo, L. A. Turka, and J. M. Ahearn. 1999. Functional characterization of soluble and membrane-bound forms of vaccinia virus complement control protein (VCP). Mol. Immunol. 36:685-697.
Sahu, A., S. N. Isaacs, A. M. Soulika, and J. D. Lambris. 1998. Interaction of vaccinia virus complement control protein with human complement proteins: factor I-mediated degradation of C3b to iC3b1 inactivates the alternative complement pathway. J. Immunol. 160:5596-5604.
Sahu, A., T. R. Kozel, and M. K. Pangburn. 1994. Specificity of the thioester-containing reactive site of human C3 and its significance to complement activation. Biochem. J. 302:429-436.
Sahu, A., and J. D. Lambris. 2001. Structure and biology of complement protein C3, a connecting link between innate and acquired immunity. Immunol. Rev. 180:35-48.
Sahu, A., and M. K. Pangburn. 1993. Identification of multiple sites of interaction between heparin and the complement system. Mol. Immunol. 30:679-684.
Sahu, A., A. M. Soulika, D. Morikis, L. Spruce, W. T. Moore, and J. D. Lambris. 2000. Binding kinetics, structure-activity relationship, and biotransformation of the complement inhibitor compstatin. J. Immunol. 165:2491-2499.
Seet, B. T., J. B. Johnston, C. R. Brunetti, J. W. Barrett, H. Everett, C. Cameron, J. Sypula, S. H. Nazarian, A. Lucas, and G. McFadden. 2003. Poxviruses and immune evasion. Annu. Rev. Immunol. 21:377-423.
Sharma, A. K., and M. K. Pangburn. 1996. Identification of three physically and functionally distinct binding sites for C3b in human complement factor H by deletion mutagenesis. Proc. Natl. Acad. Sci. USA 93:10996-11001.
Smith, S. A., R. Sreenivasan, G. Krishnasamy, K. W. Judge, K. H. Murthy, S. J. Arjunwadkar, D. R. Pugh, and G. J. Kotwal. 2003. Mapping of regions within the vaccinia virus complement control protein involved in dose-dependent binding to key complement components and heparin using surface plasmon resonance. Biochim. Biophys. Acta 1650:30-39.
Soames, C. J., and R. B. Sim. 1997. Interactions between human complement components factor H, factor I and C3b. Biochem. J. 326:553-561.
Spiller, O. B., D. J. Blackbourn, L. Mark, D. G. Proctor, and A. M. Blom. 2003. Functional activity of the complement regulator encoded by Kaposi's sarcoma-associated herpesvirus. J. Biol. Chem. 278:9283-9289.
Stoiber, H., C. Speth, and M. P. Dierich. 2003. Role of complement in the control of HIV dynamics and pathogenesis. Vaccine 21(Suppl. 2):S77-S82.
Vanderplasschen, A., E. Mathew, M. Hollinshead, R. B. Sim, and G. L. Smith. 1998. Extracellular enveloped vaccinia virus is resistant to complement because of incorporation of host complement control proteins into its envelope. Proc. Natl. Acad. Sci. USA 95:7544-7549.
Warren, L. 1959. The thiobarbituric acid assay of sialic acids. J. Biol. Chem. 234:1971-1975.(Jayati Mullick, John Bern)