Factor I Autoantibodies in Patients with Atypical Hemolytic Uremic Syndrome: Disease-Associated or an Epiphenomenon?

Summary

Background and objectives

Atypical hemolytic uremic syndrome is a disease associated with mutations in the genes encoding the complement regulators factors H and I. In addition, factor H autoantibodies have been reported in ∼10% of patients with atypical hemolytic uremic syndrome. This study searched for the presence of factor I autoantibodies in atypical hemolytic uremic syndrome.

Design, setting, participants, & measurements

This study screened 175 atypical hemolytic uremic syndrome patients for factor I autoantibodies using ELISA with confirmatory Western blotting. Functional studies using purified immunoglobulin from one patient were subsequently undertaken.

Results

Factor I autoantibodies were detected in three patients. In one patient with a high titer of autoantibody, the titer was tracked over time and was found to have no association with disease activity. This study found evidence of an immune complex of antibody and factor I in this patient, but purified IgG, isolated from current serum samples, had only a minor effect on fluid phase and cell surface complement regulation. Genetic analysis of the three patients with factor I autoantibodies revealed that they had two copies of the genes encoding factor H–related proteins 1 and 3 and therefore, did not have a deletion commonly associated with factor H autoantibodies in atypical hemolytic uremic syndrome. Two patients, however, had functionally significant mutations in complement factor H.

Conclusions

These findings reinforce the concept of multiple concurrent risk factors being associated with atypical hemolytic uremic syndrome but question whether autoantibodies per se predispose to atypical hemolytic uremic syndrome.

Introduction

Atypical hemolytic uremic syndrome (aHUS) is a disease characterized by overactivation of the alternative complement pathway (1). Mutations in the genes encoding complement regulatory proteins complement factor H (CFH) (2–5), complement factor I (CFI) (6–10), and membrane cofactor protein (CD46) (6,11–14) and complement components C3 (C3) (15) and complement factor B (CFB) (16) are associated with aHUS.

As well as inherited defects in complement regulation, acquired defects in the form of autoantibodies to CFH have been described (17–20). These autoantibodies mainly bind to the C-terminal end of CFH, where aHUS-associated mutations cluster (21). This region of the molecule binds to C3b and glycosaminoglycans and is responsible for cell surface complement regulation (22). CFH autoantibodies have been shown to impair cell surface complement regulation, thus mimicking the action of the CFH mutations seen in aHUS (17–20,23).

CFI is a serine protease that cleaves C3b and C4b in the presence of its cofactor proteins, CFH (24), C4 binding protein (25), CD46 (26), and complement receptor 1 (27). By inactivating C3b and C4b through limited proteolytic cleavage and thereby preventing the formation of the C3 and C5 convertases, CFI inhibits the alternative and classic complement pathways. CFI consists of a light chain (which carries the catalytic site) and a heavy chain (of unclear function) linked by a disulphide bond.

Mutations in CFI have been reported in 2–12% of aHUS patients (6–10). Although they are distributed throughout the molecule, they do cluster in the serine protease domain (21). Most aHUS-associated CFI mutations result in decreased secretion, resulting in a quantitative defect in complement regulation. Functional analysis of CFI mutants that are secreted normally has revealed a loss of alternative and classic pathway cofactor activity, both in the fluid phase and on cell surfaces (7,28,29).

Here, we describe the presence of CFI autoantibodies in the Newcastle aHUS cohort, investigate their functional impact, and show that these autoantibodies occur in the presence of additional genetic risk factors.

Materials and Methods

Subjects

Paired serum and DNA samples were available from 175 patients with aHUS and 100 healthy blood donors (blood donor controls). The study was approved by the Northern and Yorkshire Multi-Center Research Ethics Committee, and informed consent was obtained in accordance with the Declaration of Helsinki.

Genetic Screening

In individuals with CFI autoantibodies, mutation screening of CFH, CD46, CFI, CFB, C3, and thrombomodulin (THBD) was or had previously been undertaken by direct fluorescent sequencing as described (2,8,12,15,30,31). Variants discovered in these genes were assessed in DNA samples from 300 normal control individuals within the Wellcome Trust Patient Control Consortium (32,33). Genotyping of the following single nucleotide polymorphisms was undertaken by direct sequencing: CD46 −652A>G (rs2796267), CD46 −366A>G (rs2796268), CD46 c.4070T>C (rs7144), CFH −331C>T (rs3753394), CFH c.2016A>G p.Gln672Gln (rs3753396), and CFH c.2808G>T p.Glu936Asp (rs1065489).

CFHR1 and -3 copy number was measured by multiplex ligation-dependent probe amplification with a kit from MRC Holland (SALSA MLPA kit P236-A1 ARMD). CFHR4 copy number was measured by multiplex PCR assay as described (20). Screening for CFH autoantibodies was performed as previously described (20,34).

ELISA

The anti-CFI ELISA was carried out essentially as previously described for factor H (34), except that 5 µg/ml CFI (purified from pooled serum samples) (35) was substituted for CFH herein and a standard curve was generated using a polyclonal goat anti-CFI (Comptech) followed by rabbit anti-goat horseradish peroxidase (HRP) (Stratech Scientific). The OD₄₅₀ value for the 1/5000 dilution of goat anti-human CFI was given an arbitrary value of 100,000 relative units (RU). Alternatively, protein A/G column was used to isolated patient and control Ig from sera following manufacturer’s instructions (Pierce, United Kingdom), and the presence of CFI in the samples was detected using 1 µg/ml Medical Research Council of the United Kingdom (MRC) OX21 (gift from Bob Sim, Oxford, United Kingdom) by standard sandwich ELISA of samples.

Western Blotting

Purified CFI (35 µg/ml) was diluted in solubilizing buffer, and 20 ml was loaded onto a 10% SDS-PAGE preparative gel and transferred to nitrocellulose, which was then cut into 0.5- to 1-cm-wide strips. After blocking in 5% nonfat milk/PBS, strips were then incubated with individual sera samples (1/25 to 1/100 as appropriate) overnight at 4°C. After extensive washing in PBS/Tween 0.02%, bound autoantibody was detected using goat anti-human IgG-HRP (Stratech Scientific). Alternatively, for CFI immune complex detection, pre- or postcolumn sera (equivalent to 1/20 dilution of fresh serum) or purified Ig (using protein A/G column; Pierce; Thermo Scientific) was concentrated (using 30-kD cutoff spin columns; Sartorius Stedim Biotech) and adjusted to 1 mg/ml after quantification by bicinchoninic acid assay (Pierce; Thermo Scientific) was loaded on SDS-PAGE and blotted. MRC OX21 was used to identify the presence of CFI. Blots were developed using an enhanced chemiluminescence substrate according to the manufacturer’s specifications (Pierce; Thermo Scientific).

Complement Assays

C3 and C4 levels were measured by rate nephelometry (Beckman Array 360). CFH and CFI levels were measured by radioimmunodiffusion (Binding Site). Cell surface expression of CD46 was measured by flow cytometry as previously described (12,36).

Alternative Pathway Assays

Cell-bound complement activity was carried out essentially as previously described with minor modifications (37). Briefly, rabbit red blood cells (TCS Biologic) were washed several times in PBS and subsequently transferred to alternative pathway buffer (APB; 3.12 mM Barbital, 0.9 mM Na Barbital, 145 mM NaCl₂, 7.83 mM MgCl₂, 0.25 mM CaCl₂, 10 mM EGTA, and 0.1% w/v gelatin, final pH 7.2). Cells were resuspended at 0.1% v/v, and 100 µl were plated out on round-bottomed 96-well plates containing 100 µl triplicate serial dilutions of normal human serum or patient sera in APB. Wells were supplemented with purified patient or normal control Ig (100 µg/well), CFI (0.7 µg/well), or CFH (5 µg/well) before adding rabbit red blood cells. Plates were incubated at 37°C for 30 minutes before red cells were pelleted at 500 g for 5 min. Absorbance of supernatant was measured at OD₄₁₀.

Fluid Phase C3b Breakdown

Fluid phase complement activity was established as previously described (38). Briefly, purified C3b (4 µg), CFH (0.5 µg), and CFI (titration from 1 µg; obtained from Comptech, TX) were mixed with APB. Patient and control Ig (25 µg) in APB were mixed with the C3b- and CFH-containing solution before addition of CFI. Samples were then taken to 37°C for 3 min before being heated to 95°C for 5 min; 10% SDS-PAGE gels were stained with Coomassie blue or alternatively subjected to Western blotting as appropriate. C3b breakdown was visualized using a sheep anti-C3 at 1/500 (gift from B. P. Morgan, Cardiff, United Kingdom) followed by 1/2000 donkey anti-sheep HRP (Stratech Scientific). Western blots were visualized for 30 seconds. Image analysis was carried out on scanned gels and autorad films as follows. A set grid was used to compare pixel intensity (grayscale gradient) for each band within a lane. Results were standardized for loading based on the β-chain of C3.

Results

CFI Autoantibodies

In both the blood donor controls (BDCs) and aHUS patients, CFI autoantibody titer (in RU) was not normally distributed (Figure 1A) (determined by the Kolmogorov–Smirnov test). The median (range; mean) antibody titer in BDC and aHUS patients was 64 RU (32–504; 97) and 56 RU (22–34,921; 310), respectively. Compared with the BDC group, aHUS patients had increased levels of CFI autoantibody (P<0.02). We used the 0.975 fractile of the BDC group to determine autoantibody positivity as recommended by the International Federation of Clinical Chemistry for data with non-normal distribution (39). This use equated to 423 RU, and six patients were above this threshold (Figure 1A). However, our experience with anti-CFH autoantibodies has suggested that it is prudent to confirm presence of autoantibodies using a second technique (34). Thus, Western blotting was used to confirm the presence of autoantibodies in samples with a titer ≥423. Only the three patients with the highest titer of CFI autoantibodies were confirmed to have CFI autoantibodies after Western blotting analysis. Detection of autoantibodies in patient 1’s serum required a shorter film exposure (Figure 1B) and was readily detectable when less serum was used than for patients 2 and 3 (Figure 1C), consistent with the ELISA results. Additional Western analysis indicated the CFI autoantibody detected in patient 1 bound to the heavy chain of CFI (Figure 1D). Using specific HRP-conjugated secondaries (mouse monoclonal antibodies: MH17–15, -22, -32, -42; Invitrogen, United Kingdom) in our ELISA, P1 and P3 autoantibodies were established as being predominately IgG₁ subclass, whereas P2 autoantibodies were IgG₃ subclass (data not shown). Thus, of the 175 aHUS patients that we screened, 3 patients were confirmed to possess significant levels of CFI autoantibodies.

Figure 1.

Complement factor I (CFI) autoantibodies are found in atypical hemolytic uremic syndrome (aHUS) patients. (A) Antibodies against purified CFI in aHUS patients were detected using ELISA as described in Materials and Methods; 100 normal healthy blood donor controls (BDCs) and 175 aHUS patients were screened, and nonspecific background signal was subtracted. A standard curve was generated from a polyclonal control antibody and given an arbitrary titer of 100,000 relative unit (RUs) from a 1/5000 dilution. The horizontal bar represents the threshold for positivity based on the 97.5 percentile of the control cohort (423 RU). Purified CFI was run out on 10% SDS-PAGE and transferred to nitrocellulose. Strips of nitrocellulose were then incubated with sera collected from subjects. Patients are listed one to six, and the dilution of serum used is indicated below blots. Enhanced chemiluminescence Western blotting substrate was used to visualize bound antibody. Autorad film was exposed for 20 seconds (B) and 10 minutes (C) before developing. A known CFI autoantibody negative (−ve) was used as a control. (D) shows a Western blot of a preparative gel with reduced purified CFI loaded. Sera from patient 1 (lane 1) and a goat anti-CFI as positive control (lane PC) were incubated with the nitrocellulose, and a 30-second exposure is shown. Molecular mass markers are shown, and autorad pictures are of the individual strips of a single blot reassembled/aligned before exposure. Irrelevant intervening strips have been cropped out where appropriate. Data shown is representative of several blots.

Clinical Details: Genotyping and Background Analysis of the Three Patients with CFI Autoantibodies

A summary of the clinical details of the three patients with CFI autoantibodies is shown in Table 1, with a full summary available in Supplemental Material. Briefly, all required renal replacement therapy at initial presentation and plasma therapy was instituted. However, only patient 3 recovered renal function. In the remaining two patients, a total of three renal transplants were undertaken in the absence of any specific therapy to remove autoantibodies. Patient 1 showed recurrent aHUS in the allograft. Table 1 also shows the values for serum levels of C3, C4, CFH, and CFI, the results of screening for CFH autoantibodies, and the measurement of CD46 expression on the original samples. In all individuals, systemic alternative pathway complement activation was shown with low levels of C3, but in only one individual was the C4 level low. Mutation screening showed that patient 1 had a heterozygous mutation in CFH (c.3468dupA) and a sequence variant in CFI (c.1657 C>T; p.Pro553Ser) that was present in normal controls at a frequency of 5/574 chromosomes (Table 2). Patient 2 had a heterozygous mutation in CFH (c.2018G>A; p.Cys673Tyr). No mutations were detected in genes previously associated with aHUS in patient 3, and all three patients had two copies of CFHR1, -3, and -4. Genotyping for CFH and CD46 susceptibility factors revealed that both patients 1 and 2 are heterozygous for the at-risk haplotype CFH_TGTGGT (H3) haplotype (20), whereas patient 2 is homozygous and patient 1 is heterozygous for the at-risk CD46_GGAAC haplotype (40) (Table 3).

Table 1.

Clinical details of patients with factor I autoantibodies

Patient ID	1	2	3
Age at presentation	26 yr	5 yr	13 mo
Sex	Female	Female	Female
Precipitant	Postpartum	Unknown	C. difficile and rotavirus diarrhea
Family history	Yes	No	No
Outcome	ESRF	ESRF	Recovered renal function
Renal transplant	Yes	Yes	N/A
	One lost to recurrent aHUS at 1 mo	No recurrence at 65 mo	N/A
	One lost to transplant glomerulopathy at 50 mo		N/A
C3 (0.68–1.38 g/L)	0.66	0.61	0.56
C4 (0.18–0.60 g/L)	0.32	0.22	0.08
Factor H (0.35–0.59 g/L)	0.34	0.52	0.58
Factor H auto-Ab	Negative	Negative	Negative
Factor I (38–58 mg/L)	89	48	54
MCP expression	Normal	Normal	Normal

Normal range in parentheses. aHUS, atypical hemolytic uremic syndrome; ESRF, end stage renal failure; MCP, membrane cofactor protein.

Table 2.

Mutation screening of the genes encoding complement factor H, membrane cofactor protein, complement factor I, complement factor B, complement components C3, and thrombomodulin and measurement of the copy number of the genes encoding complement factor H–related proteins 1, 3, and 4

Patient ID	CFH	CD46	CFI	CFB	C3	THBD	CFHR1 Copy Number	CFHR3 Copy Number	CFHR4 Copy Number
1	c.3468dupA	nmd	c.1657 C>T; p.Pro553Ser	nmd	nmd	nmd	2	2	2
2	c.2018G>A; p.Cys673Tyr	nmd	nmd	nmd	nmd	nmd	2	2	2
3	nmd	nmd	nmd	nmd	nmd	nmd	2	2	2

CFH, complement factor H; CD46, membrane cofactor protein; CFI, complement factor I; CFB, complement factor B; C3, complement component C3; THBD, thrombomodulin; CFHR1, 3, and 4; complement factor H–related proteins 1, 3, and 4; nmd, no mutation detected.

Table 3.

CD46 and CFH susceptibility factors

Patient ID	CD46 −652A>G (rs2796267)	CD46 −366A>G (rs2796268)	CD46 c.4047T>C (rs7144)	CFH −332C>T (rs3753394)	CFH c.2016A>G Q672Q (rs3753396)	CFH c.2808G>T E936D (rs1065489)	CFH c.1204 C>T H402Y (rs1061170)
1	AG	AG	TC	TT	GG	GT	TT
2	GG	GG	CC	CT	GG	TT	TT
3	AG	AA	TT	CC	AG	GT	CT

Time Course of Ig Class Switching and Titer

Patient 1 showed a high titer of IgM CFI antibodies at the time of initial presentation with aHUS (Figure 2), but by the time of the first renal transplant (2 years after presenting with aHUS), the titer of IgM CFI autoantibodies was at background levels, where they have remained. Notably, the first renal transplant was lost to recurrent aHUS, and at that time, there were no detectable IgM or IgG CFI autoantibodies. By 42 months after the initial presentation and 24 months after the post-transplant recurrent episode of aHUS, IgG CFI autoantibodies are present in high titers. This finding coincides with the second renal transplant, which was not associated with recurrent aHUS. The sample used in our cohort analysis was 6 months after the peak IgG response (Figure 2), and levels had dropped to 50% maximal detected response by this time. The titer of the IgG CFI autoantibodies then declined slowly and currently rests at the ELISA positive cutoff.

Figure 2.

Patient 1 autoantibody titer changes markedly over time and does not associate with disease. Antibodies against purified CFI in archived samples from patient 1 were detected using ELISA as described in Materials and Methods. Both IgG and IgM autoantibodies were assayed. Nonspecific background signal was subtracted. OD results were transformed as previously described. Results shown are the mean of three separate ELISAs. The chronology of the two transplants and two episodes of aHUS are shown with arrows underneath the x axis. The sample assayed in Figure 1A is denoted (§).

CFI and CFI Autoantibodies Form a Circulating Immune Complex

Despite the apparent lack of correlation between CFI autoantibody titer and the clinical course of the disease in patient 1, we wished to establish if the presence of CFI autoantibodies might be disease-modifying. We hypothesized that immune complexes of CFI and CFI autoantibodies could both lead to the generation of additional proinflammatory stimuli (41) and elude detection by standard ELISA. Western blot analysis of a column-purified IgG sample from freshly obtained patient 1 serum (i.e., anti-CFI at 650 RU) showed that CFI had remained associated with IgG (Figure 3A). The post-IgG affinity column sample from patient 1 had less CFI than the normal control sample (despite similar precolumn levels established by anti-CFI sandwich ELISA), suggesting that it was bound out of the serum with IgG and then eluted from the IgG during the wash. In a second approach, we determined that Ig was associated with captured CFI extracted from archived serum samples from patient 1 (Figure 3B). Therefore, we consistently detect immune complexes of IgG autoantibody and CFI in patient 1, which may contribute to disease.

Figure 3.

Complement factor I (CFI) and autoantibodies are found in immune complexes. (A) Both P1 and control serum, confirmed to have equivalent CFI levels using sandwich ELISA, were loaded onto a protein A/G affinity column to purify IgG. Pre- and postcolumn samples and purified Ig were run out on 10% SDS-PAGE and transferred to nitrocellulose. The presence of CFI in these samples was determined using anti-CFI (Medical Research Council of the United Kingdom OX21; Bob Sim, Oxford, United Kingdom) and goat anti-mouse IgG horseradish peroxidase (HRP). Enhanced chemiluminescence Western blotting substrate was used to visualize bound antibody. Column 1, pure CFI (1 µg). Column 2, P1 precolumn serum. Column 3, control precolumn sample. Column 4, P1 postcolumn sample. Column 5, control postcolumn sample. Column 6, 25 µg purified control Ig. Column 7, 1 µg purified P1 Ig. Column 8, 25 µg purified P1 Ig. Molecular mass markers are shown, and the data are representative of three independent experiments. (B) Serum from P1 or normal controls (normal human serum) were incubated on anti-CFI (MRC OX21) -coated ELISA plates and doubly diluted from an initial 1/25 load. Goat anti-human IgG HRP was then used to detect CFI-associated IgG. Data shown is mean ± SD of combined data from three independent serum collections for patient 1 or controls. Data shown is representative of two experiments.

Purified Patient Ig Interferes with CFI Function in a Fluid Phase C3b Breakdown Assay

Detection of immune complexes suggests that CFI autoantibodies readily associate with CFI in the fluid phase. Therefore, we next assessed whether purified total Ig (containing CFI autoantibodies) isolated from patient 1 could alter CFI function in a fluid phase C3b breakdown assay. Purified C3b, CFH, and CFI were incubated with control or patient total Ig. The presence of Ig from patient 1 slowed the breakdown of C3b in the fluid phase (Figure 4). A 50% reduction in C3b breakdown over the 3-minute time period was found when limiting concentrations of CFI were mixed with patient Ig before the addition of the C3b to the reaction mixture (Figure 4C).

Figure 4.

Fluid phase breakdown of complement components C3b is modified after addition of purified P1 IgG. Purified C3b (4 µg), complement factor H (CFH; 0.5 µg), and CFI (4×1/10 double dilution from 1 µg, left to right on gels, and no CFI; designated by −) were incubated with patient or control Ig (25 µg) at 37°C for 3 minutes before the addition of 2× reducing SDS-PAGE sample solution and incubation at 95°C for 5 minutes. (A) Samples (10 µl) were loaded on 10% SDS-PAGE gel for staining with Coomassie blue. (B) Alternatively, 10 µl 1/20 dilution of sample were used for SDS-PAGE destined for Western blotting. C3b breakdown was then visualized using a sheep anti-C3 at 1/500 followed by appropriate secondary. Shown is a 30-second autorad exposure. M indicates the marker lane; sizes are illustrated on the left side, and C3 breakdown fragments are illustrated by labeled arrows on the right. The data shown is representative of four experiments. (C) Image analysis was carried out on scanned autorad films and Coomassie gels using a defined area to compare the pixel intensity (on a grayscale gradient) of bands in each lane. Results shown are the values obtained for the α′-band after being standardized for loading variation using the values obtained for the β-chain of C3. Data shown are the mean plus SD of the four experiments. Mann–Whitney t test was used to establish statistical significance at each concentration. *P<0.05, **P<0.01.

Alternative Pathway Hemolytic Activity Significantly Increased in Patient 1 Largely Because of Lower CFH Levels

From the fluid phase assays, there was evidence that CFI autoantibody did interfere with CFI function. However, cell surface complement regulation is critical in aHUS (42). Using standard alternative pathway hemolysis assays (37), freshly isolated patient 1 serum had similar hemolytic activity compared with normal human serum (NHS) (Figure 5A). This finding was counterintuitive considering that the patient serum had both lower CFH levels and CFI autoantibody, albeit currently at low levels. We surmised that the low serum C3 levels in this patient could be undermining this assay. Therefore, we mixed patient and NHS serum 1:1 to replenish C3 levels. In this analysis, the 1:1 mix gave significantly greater lysis than NHS only, suggesting failure to control complement activation in serum from patient 1. Addition of purified CFI (Figure 5B) or CFH (Figure 5C) into this mixed sample indicated that the majority of the defect was caused by the loss of CFH. To test whether the CFI autoantibody had any effect on complement regulation, we supplemented NHS with 100 µg purified patient or control Ig. Addition of patient Ig had no impact on hemolysis compared with control Ig (Figure 5D). These findings were also replicated in the sheep red blood cell alternative pathway assay (43), and addition of purified CFI autoantibody to a standard classic pathway hemolytic assay (37) again showed no clear increase in lysis as a result of interference with factor I function (data not shown).

Figure 5.

Complement factor I (CFI) autoantibodies had minimal/no detectable effect on cell surface complement regulation. (A) Alternative pathway mediate rabbit red blood cell lysis was established as per standard protocols. Patient 1 sera (P1), normal human sera (NHS), and a 1:1 mix of patient 1 and NHS (P1:NHS) are shown. (B) A 1:1 mix of patient 1 and NHS (P1:NHS) or NHS alone was supplemented with purified CFI (0.7 µg/well) before the addition of cells and hemolytic assay. (C) A 1:1 mix of patient 1 and NHS (P1:NHS) or NHS alone was supplemented with purified CFH (5 µg/well) before the addition of cells and hemolytic assay. (D) NHS was supplemented with purified Ig (100 µg/well) from a normal control (N Ig), patient 1 (P1 Ig), or buffer only as indicated. Percentage lysis was calculated from the OD₄₁₀ readings of released haem compared with 100% lysis of rabbit red blood cells in dH₂O. Mean and SD of triplicate analysis are shown. Results are representative of several independent experiments. Mann–Whitney test was used to establish significance. **P<0.01.

Discussion

In this study, we report for the first time the presence of CFI autoantibodies in aHUS patients. There was, however, little evidence to correlate the genesis of IgG isotype CFI autoantibodies with the course of aHUS in the patient with the highest recorded titer of CFI autoantibodies. Furthermore, functional analysis of freshly isolated CFI autoantibodies suggests that, currently, their presence results in only a minor modification of complement regulator capacity of CFI. This finding leads us to question whether these autoantibodies are disease-modifying or an epiphenomenon.

At a frequency of ∼2% (3/175) in the Newcastle aHUS cohort, CFI autoantibodies are less frequent than CFH autoantibodies (5–10%) (17–20). In patient 1, we have for the first time seen evidence of the class switch event in a complement protein autoantibody. Intriguingly, there is a large interval between the initial IgM and IgG response (Figure 2). Review of the patient’s history has provided no clear indication of the trigger for either the initial or subsequent events, and critically, the levels of autoantibody do not seem to be associated with the course of the disease. That we show that CFI autoantibodies exist both free in solution and in an immune complex with CFI suggests the autoantibodies may have low to intermediate affinity to native CFI.

A deletion incorporating CFHR1 and -3 is strongly associated with CFH autoantibodies in aHUS (19,44). Here, we found that all three individuals with CFI autoantibodies have two copies of CFHR1 and -3, suggesting that their absence only plays a specific role in the development CFH autoantibodies. That two of three individuals with CFI autoantibodies also carried mutations in CFH supports the hypothesis that chronic increased complement activation may predispose to the generation of autoantibodies against complement components. In patient 1, there was a heterozygous duplication of a single base pair (c.3468dupA), leading to a frame shift and premature stop codon in CFH. This finding is consistent with the low levels of CFH seen in this patient. Furthermore, patients with CFI deficiency also display low CFH levels. Therefore, the low levels of CFH seen in this patient could result from both immune complex removal of CFH associated with excess C3b (45) as well as the effects of the CFH mutation. Patient 2 has a heterozygous nonsynonymous CFH mutation (c.2018G>A; p.Cys673Tyr) that would be predicted to result in failure of secretion from that allele (3), but CFH levels, presumably produced from other allele, were found to be in the normal range (Table 1). In addition to mutations and autoantibodies, several studies have now identified CFH and CD46 risk haplotypes for aHUS. Intriguingly, both patients 1 and 2 do possess at-risk haplotypes on CFH and CD46 (Table 3) (CFH_TGTGGT [H3] haplotype [20] and CD46_GGAAC haplotype [40]), whereas patient 3, who had the best clinical outcome, did not have additional genetic risk factors for aHUS, a phenomenon previously noted in aHUS patients with CFI mutations (29).

Thus, as in patients with CFH autoantibodies, individuals with CFI autoantibodies may have additional genetic risk factors predisposing to aHUS. Furthermore, despite having a functionally significant mutation in CFH, this latent predisposition to disease did not manifest in patient 1 until the age of 26 years when she developed the disease in association with pregnancy (46). Likewise, patient 3 only developed aHUS after a diarrheal illness (not shiga toxin-associated). Thus, these three patients also show that, in addition to inherited and acquired susceptibility factors, a trigger is needed for the disease to be manifest. Recent analysis of cohorts of aHUS patients with complement mutations has identified upper respiratory tract infections (6), viruses (36), pregnancy (6,47), drugs (6), and non-Escherichia coli diarrheal illnesses (48) as potential triggers.

We have found CFI autoantibodies in ∼2% of the Newcastle aHUS cohort. These autoantibodies were associated with other susceptibility factors and support the theory that multiple hits are necessary in most patients before aHUS presents clinically (49). Importantly, CFI autoantibodies do not seem to track with disease in the patient with highest antibody titer, raising the possibility that these autoantibodies could be a marker of disease rather than a direct factor in disease development.

Disclosures

T.H.J.G. is a scientific advisor for Alexion Pharmaceuticals and has received grant support from the same company.