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Factor H (fH) is essential for complement homeostasis in fluid-phase and on surfaces. Its two C-terminal domains (CCP 19-20) anchor fH to self surfaces where it prevents C3b amplification in a process requiring its N-terminal four domains. In atypical hemolytic uremic syndrome (aHUS), mutations clustering towards the C-terminus of fH may disrupt interactions with surface-associated C3b or polyanions and thereby diminish the ability of fH to regulate complement. To test this we compared a recombinant protein encompassing CCP 19-20 with sixteen mutants. The mutations had only very limited and localized effects on protein structure. While we found four aHUS-linked fH mutations that decreased binding to C3b and/or to heparin (a model compound for cell-surface polyanionic carbohydrates), we identified five aHUS-associated mutants with increased affinity for either or both ligands. Strikingly, these variable affinities for the individual ligands did not correlate with the extent to which all the aHUS-associated mutants were found to be impaired in a more physiological assay that measured their ability to inhibit cell surface complement functions of full-length fH. Taken together, our data suggest that disruption of a complex fH-self surface recognition process, involving a balance of affinities for protein and physiological carbohydrate ligands, predisposes to aHUS.
Atypical hemolytic uremic syndrome (aHUS) is a disease characterized by hemolytic anemia, thrombocytopenia and acute renal failure (1). In 1998, Warwicker et al. described mutations in the complement regulator, factor H (fH), associated with aHUS (2). This observation was subsequently confirmed in five independent cohorts of aHUS patients (3-7). The majority of these mutations are heterozygous and cluster in the C-terminus of fH. Incomplete penetrance is described in all reports, suggesting that a combination of mutations, risk haplotypes, and precipitating factors, such as infection or pregnancy, are required for aHUS to manifest (8-12).
Factor H, an abundant serum glycoprotein (~500 μg/ml), plays a key role in the homeostasis of the complement system. It accelerates the decay of C3b,Bb complexes that, in the alternative pathway, proteolytically convert C3 to its activated form, C3b (13). It is also a cofactor for factor I-mediated cleavage and inactivation of C3b (14). Factor H is composed of 20 complement control protein modules (CCPs) that resemble beads on a flexible string in the transmission electron microscope (15, 16). The N-terminal four CCPs (i.e. CCP 1-4) encompass the region capable of cofactor and decay accelerating activity (17-20). A second principal C3b-binding site lies within the C-terminal pair of CCPs (CCP 19-20) and also interacts with iC3b and C3d (18, 21-23). At least two sites on fH (i.e. CCP 7 and CCP 20) interact with heparin and other polyanions (24-30). The C-terminal site additionally binds sialic acids (24, 28).
In the absence of fH, spontaneous activation of the alternative pathway of complement occurs in plasma, which leads to consumption of complement components C3 and factor B. In addition to its function as a regulator of alternative pathway activation in fluid phase, fH is essential for complement control on many self-surfaces. For example, it binds to human endothelial cells and basement membranes (31) protecting them from complement-mediated attack (32). Indeed, fH has a ten-fold higher affinity for C3b on host cells and other non-activators of complement versus activators, which is dependent on the presence of sialic acid clusters or other polyanions on the surface (33, 34). The specific chemical nature of human host markers has not been identified, but it is assumed to be polyanionic since sheep cells, which do not activate complement, activate the alternative pathway after surface sialic acid is removed (33, 34). Previous studies have shown that the C3b- and polyanion-binding sites in CCPs 19-20 are key for fH interactions with host surfaces (21, 22, 28, 31, 35-37). Furthermore, a recombinant form of these C-terminal CCPs (rH19-20) has been shown to compete with full-length fH for binding to C3b and host polyanions (35). This competition results in increased complement activation on host surfaces, without affecting control of complement in plasma (35).
The C-terminal module of fH is the most commonly mutated one in individuals with aHUS and functional analysis to date has demonstrated an associated impairment of binding to C3b and/or GAGs (38). Moreover, a transgenic mouse line expressing fH missing its C-terminal region regulated C3 activation in plasma but spontaneously developed aHUS (39). The truncated murine fH failed to bind to endothelial cells. Thus, in aHUS, the mutant form of fH is believed to regulate the alternative pathway of complement in the fluid phase, but fails to be recruited by glomerular endothelial cells and/or the glomerular basement membrane.
Controversy has arisen over the interpretation of the limited structure-function studies that attempted to assign disease-linked mutations in the fH C-terminus to binding sites on the fH C-terminus for specific ligands. Jokiranta et al. (40) concluded that aHUS-linked mutations lie in a C3d/b-binding site while the polyanion-binding site is not affected. On the other hand, Herbert et al. (41) presented evidence that shows that mutations cluster within an electropositive polyanion-binding site. Moreover, none of the disease-linked mutants have been analyzed with respect to their structural integrity; thus it remains possible that the mutations are destabilizing or result in unfolded protein. In the current work we set out to resolve these issues and to thereby address the question: why do aHUS-linked mutations prevent fH from functioning properly?
To avoid difficult-to-interpret complications likely to arise from the presence of additional C3b- and heparin-binding sites that reside within the N-terminal 18 CCPs of fH, we mutated and analyzed a recombinant construct containing only the CCPs 19 and 20 (rH19-20). We measured the affinity of each of sixteen rH19-20 mutants for C3b and, separately, for polyanions. Because C3b and host polyanions reside together on surfaces, we also used assays designed to measure the interactions with these ligands simultaneously in a physiological setting on the cell surface.
Blood from sheep and from two healthy human adults was collected by venipuncture and the red cells were frozen at −80°C (42). The University of Texas Health Science Center institutional review board approved protocols, and written informed consent was obtained from all human donors. The buffers used were: PBS (10 mM sodium phosphate, 140 mM NaCl, 0.02% NaN3, pH 7.4); veronal buffered saline (VBS), 5 mM veronal, 145 mM NaCl, 0.02% NaN3, pH 7.3; GVB, VBS containing 0.1% gelatin; GVBE, GVB containing 10 mM ethylenediaminetetraacetic acid (EDTA); DGVB, half-physiological ionic-strength buffer prepared by diluting GVB two-fold with 5% dextrose in water; MgEGTA, 0.1 M MgCl2, 0.1 M EGTA (ethyleneglycoltetraacetic acid), pH 7.3.
Complement proteins fH (43), C3 (44, 45), factor B (46), and factor D (47) were purified from normal human plasma as described. A fragment consisting of C-terminal residues 1107-1231 of fH (domains 19-20, i.e. rH19-20) was cloned, expressed and purified as described (41). Site directed mutagenesis was used to generate sixteen mutant versions of rH19-20, using the QuickchangeR mutagenesis system (Stratagene, Cedar Creek, TX). Ten of these are associated with aHUS (D1119G, R1182S, W1183R, T1184R, L1189F, L1189R, S1191L, R1210C, R1215G, and double mutant S1191L;V1197A) (48); the remaining six mutants (E1172R, K1188Q, R1203S, R1203Y, R1210S, and K1230A) have not so far been observed in aHUS patients, but were generated to probe C3b- and polyanion-binding sites. The residues chosen for the design of these mutants lie in putative C3b- and polyanion- binding sites (40, 41) and cover a wider surface region within domain 20, which is where most aHUS-associated mutations in fH lie. The proteins were cloned and expressed using the Pichia pastoris strain KM71H as previously described (41). Mass spectra of recombinant proteins (purified by cation-exchange chromatography and stored at −80°C in PBS as described previously (41)) were consistent with predicted molecular weights. A 15N-labelled sample of each mutant was prepared by using (15NH4)2SO4 (Isotec) as the sole source of nitrogen during cell growth. Protein concentrations were determined (280 nm) using theoretical E1%1 cm values (49). Factor H and an aliquot of wildtype rH19-20 (100 μg) were radiolabeled with 125I to specific activities of 4 to 6 μCi/μg as described (35).
For NMR, samples of 15N labeled proteins were 100 μM in 20 mM deuterated sodium acetate, 10% D2O, pH 4.5. In each case a [15N,1H] heteronuclear single quantum coherence (HSQC) spectrum was collected at 37°C on a Bruker AVANCE 800 MHz spectrometer fitted with a 5 mm CPDCI cryoprobe with Z gradients. Based on existing spectral assignments for wild type rH19-20 (41), analyses were conducted to reveal any changes in the 15N and 1H chemical shifts of each backbone amide consequent upon mutagenesis.
Protein samples (3 μM, 1 ml) in 20 mM potassium phosphate buffer, pH 7.4, at 20°C, were loaded individually onto a HiTrap heparin-affinity chromatography column (7 mm × 25 mm, GE Healthcare, Piscataway, NJ), equilibrated with the same buffer, and subsequently eluted with a 20-column volume gradient from 0–1 M NaCl in 20 mM potassium phosphate buffer, pH 7.4. Protein elution was monitored using absorbance at 280 nm. Similar experiments were carried out for the aHUS-associated mutants (0.6 μM, 5 ml) in 20 mM HEPES, pH 7.0, at 4°C, and eluted with a 20 column volume gradient from 0-1 M NaCl in 20 mM HEPES, pH 7.0.
Oligosaccharides were prepared for GMSA from low-molecular weight heparin by partial digestion with heparinase I followed by size-fractionation on a BioGel P10 gel-filtration column (BioRad, Hercules, CA) (50). In the case of the tetrasaccharide, dp4, strong anion-exchange chromatography (AS-17 Ionpac, Dionex, Sunnyvale, CA) was subsequently performed to isolate only fully sulfated product (ΔUA2S-GlcNS6S-IdoA2S-GlcNS6S). Due to the heterogeneity associated with the length of the hexasaccharide (dp6) this purification step was omitted in dp6 preparations resulting in variable sulfation patterns. Fluorophore-labeled species were produced by attachment of 2-aminoacridone to the oligosaccharide reducing end (50), and GMSAs were performed, as described previously (51).
A BIAcore 3000 instrument was used to perform SPR. Human C3b (800-2000 response units (RU)) was amine coupled to a CM5 sensor chip (GE Healthcare). In addition, a streptavidin (SA) sensor chip (GE Healthcare) was coated with biotinylated C3b, generated as recently described (52). For each type of sensor chip, three flow cells (Fc2, Fc3 and Fc4) were independently coupled with C3b. Then 20 μl injections of the rH19-20 mutants and wildtype rH19-20 were performed at 0-12 μM over the flow cells at 5 μl/min, at 22°C. The running and sample buffer consisted of PBS with 0.005% Tween. The maximum equilibrium binding of the proteins to C3b was measured by subtracting RUs obtained for the reference cell (Fc1) from the C3b-coupled flow cell RUs. The equilibrium binding values for each concentration were plotted using the Grafit 5 program (Erithacus), which determined the apparent Kd by fitting the data to a single-site saturation curve. Since the wildtype rH19-20 interacts with C3b with a 1:1 stoichiometry, the Kd values of the mutant proteins were calculated by setting the capacity (as a constant) to that of the average (n=12) capacity of wildtype rH19-20, for each C3b-coupled sensor chip.
Deposition of C3b on sheep erythrocytes (ES), and zymosan particles (Sigma-Aldrich, St. Louis, MO) was accomplished using purified C3, factor B and factor D as previously described (53). The number of bound C3b molecules was determined to be 125,000 per Es cell or 100,000 per zymosan particle by measuring binding of radiolabeled factor Bb (53).
Sheep erythrocytes (ESC3b) bearing a total of 1 μg C3b were incubated with ~20 ng of radioiodinated human fH and 0 to 8 μM non-labeled rH19-20 (wildtype or mutants) in 100 μl half-ionic strength buffer (DGVB). Alternatively, the cells were incubated with ~20 ng 125I-rH19-20 and the non-labeled proteins. After 20 min at 22°C, the bound and free radiolabeled proteins were separated as described (34). The competition experiments with C3b-coated zymosan (ZymC3b) were carried out as described above, except only the IC50 dose of the unlabeled wildtype rH19-20 (1 μM) was compared to a 1 μM dose of various rH19-20 mutants.
Lysis of human erythrocytes (35) (EH) was measured by pre-incubating 5 × 106 EH with 7 μg/ml anti-CD59 monoclonal antibody (clone MEM43; Abcam, Cambridge, MA) for 20 min at 4°C, and then mixing, on ice, with GVB, normal human serum (NHS; 40% final), and 0.1 M MgEGTA (5 mM final concentration) in the presence of 1 μM wildtype or mutant rH19-20. This concentration of wildtype rH19-20 is enough to achieve ~50% hemolysis (IC50) of anti-CD59 treated cells, due to inhibition of fH-mediated protection. The mix was immediately transferred to a 37°C water bath and incubated for 20 min. To determine the extent of hemolysis induced by the mutants versus the wildtype rH19-20, 200 μl cold GVBE was added, the samples were centrifuged, and the optical density of the supernatant was determined at 414 nm. The percent lysis was determined by subtracting the A414 of the background lysis control, and dividing by the maximum lysis in water.
Conventional Student's t tests (2-tailed) were used to assess the statistical significance of the difference between wildtype rH19-20 and the mutants in their ability to bind to C3b (BIAcore), to C3b on zymosan, and to lyse EH in the presence of NHS and p values <0.05 were considered statistically significant.
A total of sixteen mutations of rH19-20 were expressed in P. pastoris, as summarized in Figure 1A. Ten of these correspond to aHUS-linked sequence variants (38). As previously noted (41) these, to a large extent, congregate on one face of module 20 (Fig. 1A). A further six mutants, that have not been identified in aHUS patients, but that cover a wider surface region of the disease-linked domain 20, were designed to help delineate binding sites for C3b and heparin. Based on previous studies (40, 41), the residues chosen for these “designer” mutants are hypothesized to lie within C3b- or polyanion-binding sites. All the mutants were homogeneous as judged by SDS-PAGE under both oxidizing and reducing conditions (not shown). The expected molecular weight of all the mutants was confirmed by electrospray ionization mass spectrometry (not shown).
Samples of wildtype-sequence rH19-20, and of the mutant forms, were isotopically enriched with 15N. Each sample was used to collect a 2D 15N,1H-HSQC spectrum. Nearly every amino acid residue within wildtype rH19-20 yielded a distinct peak in such a spectrum, as reported previously (41) and as expected for a properly folded polypeptide. For fifteen out of sixteen mutants, the 15N,1H-HSQC spectra (not shown) were also consistent with folded proteins; the exception was the designed mutant R1203Y, which was judged to be unfolded by 1H NMR. Furthermore, a comparison of HSQC spectra showed good conservation of chemical shifts with variations confined to the vicinity of the mutated residues. Moreover, lineshapes and spectral quality were consistent with monomeric protein for all proteins tested. These observations suggest that none of the aHUS-linked mutations resulted in major structural changes.
Previous work suggests that host GAGs form at least part of the target recognized by the C-terminal anchoring region of fH (25, 28, 31, 35, 37, 41). Thus it was of interest to assess the extent to which our mutations affect GAG binding. Heparin is commonly used as an experimental model compound for GAGs that are found on cell surfaces such as heparan sulfate and dermatan sulfate. The wildtype and mutant forms of rH19-20 were loaded onto a heparin-affinity column and eluted using a salt gradient (Fig. 2A-B). Some aHUS-linked mutants bound to the resin more tightly, others bound more weakly and the remainder eluted similarly to wildtype rH19-20. For example, the strongest interaction was found with W1183R (eluting at 520 mM NaCl), while the weakest was R1182S (eluting at 325 mM NaCl). The wildtype rH19-20 eluted at 370 mM NaCl. In general, mutations that resulted in gain of positive charge increased affinity (aHUS mutants W1183R, T1184R, L1189R; designed mutant E1172R), while mutations causing a loss of positive charge decreased affinity (aHUS mutants R1182S, R1210C, R1215G; designed mutants K1188Q, R1203A, R1210S, K1230A). To investigate further, GMSAs were performed. In these assays, the migration of a fluorescently tagged defined-length GAG fragment through an agarose gel was monitored in the presence of wildtype rH19-20 or the mutants. The wildtype protein binds and hence retards the mobility of both a chemically pure fully sulfated heparin-derived tetrasaccharide (dp4, data not shown), as well as heparin-derived hexasaccharides (dp6) that contain heterogeneous sulfation patterns (Fig. 2C). The experimental conditions were optimized so that the wild type showed some free dp6 migrating towards the anode. Smaller amounts of observed free dp6 for some mutants indicated stronger binding, while larger amounts corresponded to weaker binders. The ranking of affinities of rH19-20 mutants (aHUS-associated and designed mutants) for GAGs, as inferred from the GMSA (Fig. 2C), was very similar to the ranking derived from heparin-affinity chromatography (Fig. 2A-B). In summary, there is no uniform link between the affinity for heparin of the aHUS-associated mutants and aHUS pathology. Thus, some of the aHUS-associated mutants (W1183R, T1184R, L1189R, L1189F, S1191L, and S1191L;V1197A) were either unaffected or had increased affinity for heparin.
We measured the affinity of each mutant protein for C3b using SPR. C3b was immobilized on a CM5 sensor chip by amine coupling. Figure 3A summarizes the value of Kd obtained for each mutant. Four of the aHUS mutants (D1119G, R1182S, R1210C, and R1215G) bound with lower affinity to immobilized C3b, compared to the wildtype. The other aHUS mutants were either unaffected in their ability to bind (V1197A in the context of the S1191L;V1197A double mutant) or had a slightly (L1189F) or significantly (W1183R, T1184R, L1189R, S1191L) increased affinity for C3b. It is noteworthy that most of the aHUS mutants that clearly bind more strongly to C3b (i.e. W1183R, T1184R, L1189R) correspond to those with a higher affinity for heparin, while most of those with low affinity for C3b (i.e. R1182S, R1210C, R1215G) correspond to those exhibiting weaker heparin binding, as summarized in Figure 4. Interestingly, the aHUS mutant S1191L, which bound with greater affinity to C3b (Fig. (Fig.33--4),4), maintained a normal ability to bind heparin (Figs. (Figs.22 and and44).
A second type of sensor chip, pre-immobilized with streptavidin (SA chip), was used to immobilize C3b molecules that had been biotinylated at the cysteine-SH within the thioester site. This mode of immobilization emulates more closely the natural orientation of C3b on surfaces (52). Figure 3C shows representative equilibrium binding single-site saturation curves for rH19-20 wildtype as well as examples of a high-affinity (L1189R) and a low-affinity (R1215G) mutant binding to C3b. The Kd values obtained with the SA-C3b sensor chip (Fig. 3B) were very similar to those obtained on the CM5 chip (Fig. 3A). These findings indicate that the mode of C3b coupling did not bias the measured Kd values.
In order to further investigate the unexpected increase in affinity for C3b of some mutants as observed by SPR, C3b-coated zymosan particles (ZymC3b; which are devoid of polyanions) were incubated with radiolabeled wildtype rH19-20 in combination with unlabeled wildtype or the mutants with the highest and lowest affinity for C3b (by SPR). Figure 3D shows that aHUS mutants W1183R, T1184R, and L1189R as well as designed mutant E1172R, competed with radiolabeled rH19-20 more efficiently for binding to surface-bound native C3b than the unlabeled wildtype rH19-20, while aHUS mutants R1182S and D1119G were less efficient. The differences between the ability of the wildtype versus each of the mutants to bind to ZymC3b were all significant (p<0.05). These results agree with the SPR data, confirming that some disease-linked mutants do indeed bind with higher affinity to C3b. Taken together, the C3b-binding data show that there is no correlation between affinity for C3b and the association of the mutations with aHUS.
Considering our panel of sixteen rH19-20 mutations as a whole, fourteen neither introduced a cysteine (the exception being R1210C) nor resulted in structural disruption (the exception being R1203Y). Of these fourteen, nine had a positive or negative effect, compared to wildtype rH19-20, on both heparin and C3b binding, while one (V1197A, within the context of S1191L;V1197A double mutant) had no significant effect (Fig. 4). Two other mutations (R1210S and K1230A) decreased heparin binding, but did not alter C3b binding. Mutant D1119G significantly decreased C3b binding, but had no effect on heparin binding. On the other hand, S1191L increased C3b binding, while having no effect on heparin binding. Since these mutations resulted in only localized structural changes it seems valid to summarize their various functional consequences on the 3D structure of wildtype rH19-20 (Fig. 1B) and to infer that the wildtype residue is directly involved in ligand-binding if its substitution alters affinity. On this basis, Figure 1B illustrates that there is an overlapping binding patch on one face of module 20 for heparin and C3b. Two of the mutants that had minor or negligible effect on heparin binding (L1189F and V1197A, respectively) both involve conservative substitutions of buried side chains.
A clear message of the results described above is that while ten mutants included in this study have been linked with occurrence of aHUS (48), not all of them are negatively affected in their ability to bind to C3b and polyanion ligands (summarized in Fig. 4). We therefore tested whether these mutations interfere with the ability of fH to bind, simultaneously, to C3b and host-like polyanions in a more physiological setting using a competitive binding assay, as previously described (35). In this assay, rH19-20 competes with radiolabeled full-length fH, inhibiting its binding to C3b and host-like polyanions on sheep erythrocytes (ES), which carry high levels of sialic acid-containing polysaccharides (33, 34) (as do most human cells and tissues). We showed previously that such competition impairs fH complement-regulatory functions and increases complement activation on host surfaces (35). Thus, the C-terminal module-pair is important for anchoring fH to self-surfaces following C3b deposition. Data in Figure 5A confirms that wildtype-sequence rH19-20 inhibits, in a dose dependent manner, the binding of full-length fH to C3b-coated host-like cells, with an IC50 of 0.2 μM. The aHUS-linked mutant forms of rH19-20, on the other hand, were significantly impaired in their ability to compete, displaying from < 2.5% (R1215G, D1119G and R1182S) to 40-50% (T1184R and R1210C) of the inhibitory activity of the wildtype rH19-20 (Fig. 5A). Only the designed mutant (E1172R) retained more than 70% activity in this assay. The presence of the V1197A mutation within the double mutant S1191L;V1197A did not further impair its ability to compete versus S1191L alone.
We also determined the effect of the mutations on the ability of rH19-20 to compete with radiolabeled wildtype rH19-20, i.e. without the potential interference of the other C3b/polyanion-binding sites or higher-order architecture of full-length fH, and very similar results were obtained (Fig. 5B). Thus, the four aHUS mutants that were most affected (D1119G, R1182S, W1183R, and R1215G) and the two least affected mutants (T1184R and R1210C) in the competition assay with full-length 125I-fH, ranked comparably in the competition assay with 125I-rH19-20, and can be compared in summary Figure 4.
Experiments were performed in which complement-mediated lysis of human erythrocytes was measured in the presence of normal human serum and rH19-20 mutants (Fig. 6). Wildtype rH19-20 competes with fH, inhibiting fH-mediated protective action more effectively than any of the aHUS-linked mutants, apart from R1210C and T1184R, which are comparably effective. This is largely consistent with the cell-surface binding results (Fig. 5) and provides further evidence that anchoring of fH to self-surfaces via its C-terminus is essential for its protective function.
Thus, the competition data strongly indicate that a common feature of aHUS-linked fH mutants is the reduced ability to engage, in a functionally effective way, with the fH target (C3b/C3d), when in the context of polyanion-rich self-surfaces. Intriguingly, this behavior on physiological cell surfaces does not correlate with the increased affinity that some of the aHUS mutants displayed for C3b (Fig. 3) and/or for the GAG-model compound heparin (Fig. 2), when these interactions were analyzed individually.
If a mutation were to impair the ability of fH to regulate complement at self-surfaces it would have particularly significant consequences for tissue and cell surfaces thought to rely on fH for protection from complement-mediated damage. An example is the glomerular basement membrane that lacks membrane-bound complement regulators, but is exposed to plasma (and therefore the alternative pathway of complement), by fenestrations between the endothelial cells of the glomerular microvasculature. Thus, fH mutations clustered within the surface-anchoring C-terminal region, which have been detected in a significant proportion of aHUS patients, may be expected to directly trigger or exacerbate pathogenesis. Understanding the precise properties of disease-linked mutants of fH will therefore help to clarify the molecular basis of aHUS. For example, disease-associated mutations in the C-terminal region may disrupt its three-dimensional protein structure. Alternatively, they could interfere with the recognition site in CCP 20 of fH for polyanionic self markers, or they may perturb the C-terminal binding site for surface-bound C3b, or they could affect both polyanion and C3b-binding sites simultaneously. In the present study, we sought to distinguish between these various possibilities.
Our NMR analyses show that none of the ten disease-linked mutations in the current study result in anything other than highly localized effects on the structures of CCPs 19 or 20. Consequently, any functional effects of these mutations may reasonably be ascribed to their location within, or near to, critical binding sites on the wildtype protein. C3b- and heparin-binding data collected on these and five further, structurally intact, “designer” mutations in the context of rH19-20 allow the most comprehensive delineation of binding sites for these ligands to date. This exercise (summarized in Figs. Figs.1B1B and and4)4) suggests that positively charged binding sites for heparin and C3b overlap on one face of rH19-20. The binding surface for heparin (and by extension, for other glycosaminoglycans) appears to extend towards the C-terminus of fH while the C3b-binding surface extends into module 19. Changes in the electrostatic nature of the surface had the largest influences (gain of positive charge or loss of negative charge appeared to correlate with increases of both heparin and C3b-binding), thus it is possible that some of the residues concerned are involved in electrostatic steering effects rather than direct contacts with ligands. These findings are in partial agreement with previous reports (40, 41) but do not support a suggestion of simultaneously occupied binding sites on opposite faces of CCP 20 (41).
Functional analyses of some aHUS-associated mutants reveal effects consistent with an intuitively feasible model of aHUS pathogenesis; i.e. their reduced ability to bind to a polyanion-rich C3b-coated host-like surface correlates with their diminished performance in assays that measure directly affinity for C3b or for heparin. For example, although D1119G (involving loss of a negative charge) is unaffected in its ability to bind heparin, it has significantly diminished affinity for C3b (Fig. (Fig.33--4)4) and this mutant was also the weakest of all the mutants in the cell surface assays. Moreover, analysis of some of the “designer” mutants (i.e. those engineered for this study to help delineate C3b- and heparin- binding sites, but not identified in any aHUS patients to date) also produced results that appear to link an affinity for C3b and/or heparin to a capacity for blocking fH binding to a C3b-coated erythrocyte surface. Thus, the designer mutant R1210S binds C3b with wildtype affinity and is only slightly diminished in affinity for heparin, while maintaining near-wildtype activity in the cell surface assay. The case of the aHUS-linked R1210C is complicated by the likelihood that it forms disulfide bonds with itself or other proteins (54) (recombinant R1210C was found to be glutathionylated in our study). In all, seven mutations (aHUS-linked mutants R1215G and R1182S, and designer mutants K1188Q, R1203S, R1210S, K1230A) display both weaker heparin and C3b binding (Figs. (Figs.22--3)3) and, as expected, their performance is significantly impaired in the cell-surface assays (Fig. 5). It should be noted that the reduced heparin binding obtained with R1182S contrasts with the unaffected heparin binding previously observed with a mutant, R1182A, which has not been found in aHUS patients (40).
Indeed all the aHUS-associated mutants exhibit a weaker than wildtype ability to compete with fH for erythrocyte surface binding (Fig. 5). For example, L1189R and W1183R perform very poorly in cell-surface assays (Fig. (Fig.55--6),6), while T1184R and R1210C are the only aHUS-mutants that are nearly as good at inhibiting the protective effects of fH as the wildtype rH19-20. What is striking is that several of these mutations have a higher affinity both for heparin (Figs. (Figs.22 and and4)4) and for C3b (Fig. (Fig.33--4).4). Such is the case for aHUS mutants W1183R, T1184R, L1189R, and the designer mutation E1172R, all of which involve an introduction of one or more positive charges. Note that in the case of the W1183R mutant, the observed increased affinity for C3b and heparin differed from what had been observed previously with another aHUS-linked mutant, W1183L (40) that had reduced binding to C3b and heparin; unlike W1183L (40), the W1183R mutant ran as a single band by SDS-PAGE (not shown). The L1189F mutation also shows increased C3b- and heparin-binding, albeit to a much lesser extent than L1189R (Figs. (Figs.33--4).4). Another mutant (S1191L) with weaker than wildtype activity in the cell surface assays (Figs. (Figs.55--6),6), has enhanced affinity for C3b (Fig. 3), contrary to previously reported results (54, 55). This same mutant had no effect on heparin binding (Fig. 2), in agreement with a previous study (55). Interestingly, the presence of V1197A in the S1191L;V1197A double mutant has no additional effect in any of the assays tested versus the S1191L mutant.
An overall summary of the heparin and C3b binding results (where a wide range of affinities for C3b and heparin were observed), and the cell-surface complement functional assays (where all aHUS-linked mutants were impaired), is illustrated in Figure 4. Taken together these results unexpectedly show that, measured separately, affinities for C3b and heparin do not correlate consistently with the strength of binding to a polyanion-rich erythrocyte surface (on which C3b has been deposited); nor do they correlate with the disease-risk phenotype.
In the case of heparin it is conceivable that this material is not in fact representative of a putative host tissue-specific polyanionic ligand. The presence in such a ligand of particular patterns or densities of sulfation, for example, might result in differential binding amongst the mutants that correlates better with cell-surface assays. Studies aimed at identifying physiological polyanion species, as well as elucidating their chemical nature, are warranted.
The lack of correlation between the separate ligand binding assays and the cell based assays also suggests that a mutation may affect a third functional site in CCPs 19-20, perhaps different from those involved in C3b or polyanion binding. For example, a mutation could affect the region involved in the formation of putative dimers or tetramers (40, 56-58), which may be important for fH binding/function on the cell surface. In this regard, we have recently determined that polyanions greatly promote formation of fH dimers and tetramers, and that this dimerization/tetramerization leads to an increase in fH binding and complement control activity at the cell surface (58). In addition, we have determined that this polyanion-induced self-assembly is mediated, at least in part, by C-terminus to C-terminus complex formation, specifically through CCP 18-20 (58).
A further possibility derives from consideration of the relative affinities of fH for each ligand in the setting of the cell surface, and the order of binding events. An increased affinity between fH and polyanions, due to CCP 19-20 mutations, may result in a decrease in the ability of fH to diffuse normally over the cell surface, impairing its ability to efficiently encounter and bind C3b. Thus, such mutants would be less effective than normal CCP 19-20 that binds with the correct affinity to the combination of both ligands (polyanions and C3b/C3d). Likewise, the mutants that were found to have increased affinity for C3b may get retained/delayed in complexes with the products of decay acceleration and cofactor activity (i.e. C3b/iC3b).
Thus, we propose that aHUS-linked mutations disturb a complex relationship between affinities of fH for its individual ligands (C3b and physiological polyanions) or possibly for itself, in the context of the cell surface.
The authors express their appreciation to Connie Elliot for her excellent technical assistance and to Dr. Pierre Neuenschwander for helpful advice on operation of the BIAcore 3000 instrument.
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One of the authors (M.K.P.) is an officer of and has a financial interest in Complement Technology, Inc. (www.ComplementTech.com), a supplier of complement reagents.
1This research was supported by American Heart Association National Scientist Development Grant 0735101N (V.F.), D.K. is a Kidney Research UK Clinical Training Fellow, National Institutes of Health Grant DK-35081 (M.K.P.), Wellcome Trust Grant 078780/Z/05/Z (D.U. and P.N.B.).