Comparison of Three <scp>ELISA</scp> Assays for the Detection and Quantification of Autoantibodies Against Complement Factor H

Complement-mediated thrombotic microangiopathy (CM-TMA) also known as atypical hemolytic uremic syndrome (aHUS) is a rare form of thrombotic microangiopathy (TMA), marked by a triad of clinical features: thrombocytopenia due to platelet consumption, mechanical hemolytic anemia, and organ impairment of varying severity, most commonly affecting the kidneys [1]. CM-TMA is primarily driven by dysregulation of the alternative complement pathway. About 60% of cases involve genetic mutations in complement regulatory proteins [1], 10% of cases are associated with autoantibodies targeting complement factor H (FH) [2-4]. The remaining 30% of cases are idiopathic, with no identifiable genetic mutation or autoimmune cause [5]. In patients with anti-FH autoantibodies, particularly those with high titers, immunosuppressive therapy (e.g., rituximab or corticosteroids) may be added to plasma exchange (PEX) to suppress B-cell activity and reduce antibody production [3]. Consequently, detecting anti-FH autoantibodies is essential for guiding treatment as prompt diagnosis and targeted therapy are essential to minimize irreversible organ damage and improve patients' outcomes [6]. Therefore, it is essential to use diagnostic methods that are both sensitive and specific for detecting and quantifying anti-FH autoantibodies. The enzyme-linked immunosorbent assay (ELISA) remains the most widely employed method for this purpose [7, 8]. However, the increasing availability of commercial kits presents challenges for clinical laboratories seeking reliable and reproducible results. The standardization of protocols have been recommended to reduce inter-laboratory variability [8, 9] with Watson et al. endorsing the Paris protocol as the standard assay, for its high specificity and reduced cross-reactivity [8]. The Hemostasis Laboratory of CHU Sainte-Justine (CHUSJ) developed an in-house ELISA assay derived from this protocol [10]. The objective of this study was to evaluate and compare the performance of our in-house method with two commercial ELISA kits. These assays were assessed for their ability to detect and quantify anti-human Factor H immunoglobulin G (IgG) in plasma samples from patients referred to the CHUSJ Hemostasis Laboratory for suspected CM-TMA. Concordance between assays was evaluated using Pearson correlation for quantitative comparison, and Cohen's kappa for categorical agreement. Statistical significance was defined as a p-value ≤ 0.05. Statistical analyses were performed using IBM SPSS statistics (version 28.0.0.0). A total of 75 citrated plasma samples from 51 patients were included in the present comparison study. These samples were selected from approximately 1100 specimens received between 2015 and 2025 at the Hemostasis Laboratory of CHUSJ, all previously tested for anti-FH autoantibodies due to suspected CM-TMA. The selection criteria ensured a representative distribution of patient demographics (median age: 32 years; range: 4–83 years; 27 females and 24 males) and anti-FH IgG levels, including both initial diagnostic and follow-up samples. Various clinical conditions such as confirmed aHUS, membranous glomerulonephritis (GN), complement 3 GN (C3GN), Crohn's disease, bone marrow transplant, or familial history of genetic aHUS were covered to ensure the inclusion of diverse pathological contexts. External samples (56 from 40 patients) received at our laboratory facility, without clinical data were included to reflect real-world laboratory settings where medical history may be limited. Blood samples were collected in 0.109 mol/L buffered sodium citrate tubes (Vacutainer BD). Platelet-poor plasma (PPP) was obtained by double centrifugation at 2685 × g for 10 min at room temperature and stored at −80°C until analysis. One in-house direct ELISA titration assay (in-house ELISA) and two commercial quantitative ELISA kits, the Anti-Faktor H assay (#4067; Generic Assays; Dahlewitz, Germany; ELISA-1) and the CFH IgG ELISA kit (#KA1477, Abnova, Taipei City, Taiwan; ELISA-2), were used to detect antibodies targeting FH. The commercial kits were used according to the manufacturer's instructions. ELISA-1 was optimized for use in PPP. Results were expressed in arbitrary units (AU/mL). For the in-house ELISA, 96-well microtiter plates (Immulon 4HBX, ThermoFisher Scientific) were coated with 2 μg/mL of purified human FH (Sigma-Aldrich St-Louis, MO), diluted in carbonate–bicarbonate buffer (pH 9.6), and incubated overnight at 4°C. Wells were then blocked for 1 h at room temperature (RT) with phosphate-buffered saline (PBS: 137 mM NaCl, 8 mM Na2HPO4, 1.5 mM KH2PO4, 2.7 mM KCl, pH 7.2) containing 2% bovine serum albumin (BSA; Sigma-Aldrich). After blocking, 100 μL of PPP from patients, diluted 1:50 in PBS containing 0.1% Tween 20 and 2% BSA were added and incubated for 2 h at RT in the dark. Plates were then washed five times with PBS (pH = 7.4) containing 0.1% Tween 20. Subsequently, wells were incubated for 1 h with HRP-conjugated anti-human IgG (Sigma-Aldrich, ref: A2290) diluted 1:5000 in HEPES buffer (125 mM HEPES, 125 mM NaCl, 2% BSA, 0.1% Tween 20) at RT. After five washes, color development was performed using the o-phenylenediamine dihydrochloride (OPD) substrate, and the reaction was stopped with H2SO4 (1.4 M). Absorbance at 492 nm was measured using a Synergy H4 microplate reader (BioTek, Winooski, VT). To enhance analytical specificity, a BSA-coated blank control was included. This control enabled the identification and subtraction of background signals attributable to Ig reactive with non-target proteins, thereby confirming positivity. Control plasmas from 12 healthy donors (6 females and 6 males, aged 18–55 years) were included on each plate to establish the positivity threshold, defined as the mean absorbance of healthy controls plus three standard deviations (SD). For samples tested positive, antibody titration was performed using serial plasma dilutions ranging from 1:50 to 1:6400. A known anti-FH IgG-positive sample was included as a positive control. The antibody titer was defined as the reciprocal of the highest dilution of the patient sample yielding a positive signal. All samples were analyzed simultaneously using the three ELISA methods to minimize potential bias related to storage time. Both ELISA-1 and ELISA-2 demonstrated good assay performance for negative controls, with intra- and inter-assay coefficients of variation (CV) below 20%. ELISA-1 showed strong reproducibility, with low intra-assay CV (4.2% and 8.7%) and inter-assay CV (14.9% and 4.6%) for negative and positive controls, respectively. In contrast, while ELISA-2 had acceptable precision for negative controls (intra-assay CV: 10.3%; inter-assay CV: 13.8%), the inter-assay CV for the positive controls was elevated at 46.9%, indicating substantial variability for positive samples. Out of 75 samples tested, 31 were positive for anti-FH using the in-house assay, 38 were negative and 1 fell within the gray zone (titer of 1) but was considered positive according to clinical interpretation. Five false-positive samples were identified, one of which tested positive with ELISA-1 and all were negative with ELISA-2. These results highlight the importance of including a BSA-coated well as a negative control. The Venn's diagram shows the overlapping distributions of anti-FH autoantibodies' positive and negative results for each ELISA kit (Figure 1). Among the 32 positive cases (median titer: 8 AU/mL; range: 1–128), identified by the in-house ELISA, 27 (84%) were also positive with ELISA-1, and 26 (81%) were positive with ELISA-2. Similarly, of the 43 negative results by the in-house ELISA, 36 (84%) were also negative with both ELISA-1 and ELISA-2, indicating a good but not complete agreement between assays. A concordance analysis was then performed to assess the level of agreement among the three ELISA using different positivity thresholds (Table 1). We observed a risk of under-detecting positive cases when applying the manufacturer's positivity threshold. As recommended by the manufacturers, we chose the positivity threshold based on the characteristics of our study population. Although the use of a 2 SD threshold is considered less stringent than 3 SD, the sensitivity was higher (> 80%) for both the ELISA-1 and ELISA-2, when thresholds were set at 3.7 AU/mL and 9.0 AU/mL respectively (corresponding to 2 SD). The concordance between the two thresholds (2 SD and 3 SD) was comparable and the reduction in specificity was minimal when using 2 SD rather than 3 SD. The highest concordance was observed between the in-house ELISA and the ELISA-1, with 85% agreement, a Cohen's kappa (κ) of 0.69, and a Pearson correlation coefficient (r2) of 0.61, indicating good agreement and linear relationship. In comparison, ELISA-2 showed 82% agreement with the in-house ELISA and a κ of 0.65, suggesting good agreement but the correlation of quantitative values was weak (r2 = 0.11). Finally, the concordance between ELISA-1 and ELISA-2 was 74.7% with κ = 0.49 and r2 = 0.25, reflecting moderate agreement and a weak correlation. Linear regression comparing anti-FH autoantibody titers measured by the in-house ELISA and ELISA-1 yielded a slope (β) of 1.06 and an intercept (α) of 7.56, indicating a strong proportional relationship with a slight positive bias (Figure 2). In contrast, the comparison with ELISA-2 showed a slope of 0.36 and an intercept of 13.19, suggesting that ELISA-2 underestimates higher titers while introducing a positive baseline offset. Finally, the regression between ELISA-1 and ELISA-2 showed a slope = 0.41; intercept = 9.15, highlighting the limited proportionality and agreement between these two commercial assays. Bland-Altman plots confirmed these discrepancies. The in-house ELISA had the closest agreement with ELISA-1, with a moderate mean bias and tighter limits of agreement (LOA), therefore indicating greater consistency. In contrast, ELISA-2 tended to underestimate values and showed higher variability. The greatest variability was observed between ELISA-1 and ELISA-2, with ELISA-2 generally underestimating anti-FH titers. These findings suggest that, although average anti-FH levels may appear similar, individual results differ significantly, and ELISA-1 and ELISA-2 should not be considered interchangeable for quantitative analysis. Our study has several limitations. First, the ELISA methods detect only free anti-FH autoantibodies that are not bound to circulating FH [9], possibly omitting positive patients with circulating immune complexes. Additionally, the study was conducted using plasma samples, although this matrix was not evaluated by the manufacturer of ELISA-1. Despite potential matrix effects, the assay showed strong reproducibility and consistent performance, with intra- and inter-assay CVs within acceptable limits. In conclusion, our findings support the need to optimize diagnostic practices in CM-TMA associated with anti-FH autoantibodies. This comparative evaluation of two commercial ELISA kits against a validated in-house method revealed variability in performance. Given assay discrepancies, we strongly recommend the use of individual blank controls to exclude false positives. Confirming a positive result using a different kit, when feasible, may also be instrumental to support clinical decision making. S.T.-B., C.M., A.-L.L., and A.C. analyzed data and wrote the manuscript. E.P., L.B., and R.Y. collected and analyzed data. G.-E.R., S.T., and A.B. designed the study and wrote the manuscript. The study was approved by the Research Ethics Committee of CHUSJ (2019-2184). The authors declare no conflicts of interest. The data that support the findings of this study are available from the corresponding author upon reasonable request.

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