The presence in the sera of celiac disease (CD) patients of anti-actin autoantibodies (AAAs) has been suggested as a marker of severe intestinal villus atrophy (1). AAAs have been detected with an immunofluorescence (IF) technique and seem to contribute to villus cytoskeleton damage and to the pathogenesis of intestinal damage in CD (2).
The aims of the present study were to evaluate the relationship between the presence of serum IgA AAAs and severity of intestinal mucosa damage in CD patients and to compare the IF assay with a new ELISA for IgA AAA determination.
We enrolled 150 individuals in the study. IgA AAAs were assayed in 58 consecutive CD patients diagnosed between January and December 2003: 30 adults (10 male; median age, 32 years; range, 18–56 years) and 28 children (14 male; median age, 18 months; range, 1–12 years). The sera were collected at CD diagnosis, after overnight fasting, and were frozen at −80 °C for a mean of 9 months before AAA determination. In 20 patients, AAAs were reassayed after 6–12 months of gluten-free diet. CD diagnosis was based on the revised criteria of the European Society of Pediatric Gastroenterology and Nutrition (3). We enrolled 64 patients as “healthy” controls [34 adults evaluated for suspected hypercholesterolemia (15 male; median age, 35 years; range, 18–56) and 30 children with recurrent pharyngotonsilitis (14 male; median age, 3 years; range, 1–12 years)]. None of these controls had symptoms or laboratory signs suggesting CD, and all were negative for anti-endomysium (EmA) and anti-transglutaminase (anti-tTG). We enrolled an additional 28 adults with autoimmune or gastrointestinal diseases other than CD as “disease” controls: type 1 autoimmune hepatitis (AH; 6 cases), type 2 AH (4 cases), systemic lupus erythematosus (4 cases), Sjögren disease (3), primary biliary cirrhosis (2), Crohn disease (4), multiple food intolerance (3), autoimmune enteropathy (1), and refractory sprue (1).
Finally, to test whether the anti-actin ELISA used was specific for F-actin, we assayed for IgG anti-F-actin in 18 sera from patients with type 1 AH, positive by IF for IgG anti-F-actin on HEP-2 cells (titer range, 1:160–1:1280), and 30 negative sera from healthy controls.
Modified Hep-2 cells (HEP II actin; INOVA; ref. 508090) were incubated at room temperature for 30 min with serum serially diluted (1:10 to 1:1280) with phosphate-buffered saline (pH 7.2) in a covered moist chamber, washed twice in phosphate-buffered saline for 5 min, incubated at room temperature for 30 min, with fluorescein isothiocyanate-conjugated goat anti-human serum [anti-IgA; F(ab′)2 Anti-Human IgA; ref. 30240; Bio-Rad], washed twice as before, and read with a fluorescence microscope. The affinity-isolated rabbit AAAs were used as positive controls in all experiments (Sigma Chemical); rabbit monoclonal AAAs were used at a 1:50 dilution during the first incubation followed by a second incubation with the 1:100-diluted fluorescein isothiocyanate-conjugated anti-rabbit immunoglobulins (Sigma).
The IgA AAA enzyme immunoassay (ELISA) was performed with a commercially available method for anti-actin IgG determination (F-Actin Smooth Muscle; INOVA; ref. 708785) and an anti-serum anti-human IgA conjugate (INOVA; ref. 508549). We added 100 μL of diluted serum (1:101) to the wells and incubated for 30 min at room temperature. After the wells were washed three times with buffer, 100 μL of the IgA conjugate was added to each well, and the plates were incubated for 30 min and washed again. We added 100 μL of 3,3′,5,5′-tetramethylbenzidine (TMB) chromogen to each well and incubated the wells in the dark for 30 min at room temperature. Stopped reactions were read at 450 and 620 nm. The typical threshold value for AAA absorbance was arbitrarily fixed as equal to the mean value + 2 SD displayed by control sera (0.270). The control wells (no serum) introduced in each plate had an absorbance of <0.025. Intra- and interassay CV were calculated on 20 samples.
Serum IgA AAAs evaluated by IF were positive in 54 of the 58 (93%; 95% confidence interval, 88–98%) untreated CD patients; samples from 3 CD children and 1 adult were negative. The titer of IgA AAA-positive samples ranged between 1:20 and 1:640 (median, 1:80). Serum IgA AAAs evaluated by ELISA were positive in samples from 51 of the 58 CD patients (86%; 79–93%); the samples from the same 4 patients negative by IF were also negative by ELISA, and samples from 3 additional CD children were negative. The correlation between AAA results by IF and ELISA was high: r = 0.819 (Spearman correlation coefficient; see Fig. 1⇓ in the Data Supplement that accompanies the online version of this Technical Brief at http://www.clinchem.org/content/vol51/issue5/). The within-assay imprecision (CV) of the ELISA method for IgA AAAs was 4.3%, and the between-assay CV was 9.2%.
None of the healthy controls had AAA values, as assayed by ELISA, higher than the cutoff, and values were significantly higher in CD patients [mean (SD), 0.974 (0.924)] than in controls [0.157 (0.057); Mann–Whitney test, z = 6.4; P <0.0001]. Among the patients with autoimmune diseases or with various intestinal diseases, 10 had values above the cutoff: 6 patients with type 1 AH [all positive for anti-smooth muscle antibodies (ASMAs)]; 2 patients with primary biliary cirrhosis (both positive for anti-mitochondrial antibodies and 1 positive for ASMAs), and 2 patients with multiple food intolerance. The AAA values of CD patients were significantly higher than those of the disease controls (Mann–Whitney test, P <0.001). Regarding the IF AAA assay results for the 2 control groups, there were 8 positives in the healthy controls (all with a titer of 1:20) and 9 positives in the disease controls (titers, 1:40–1:160), 7 of whom also were positive by ELISA.
Intestinal histology of the CD patients revealed that 8 had mild villus atrophy, 23 had marked villus atrophy, and 27 had total villus atrophy. All patients with marked or total villus atrophy were positive for AAAs in the ELISA, whereas all 7 patients negative for AAA had mild villus atrophy. There was a significant inverse correlation between AAA values by ELISA and IF and the villus-height/crypt-depth ratio (r = −0.447 by ELISA, r = −0.404 by IF; P <0.0001; Fig. 1⇓ ). Furthermore, the CD patients negative for AAAs by ELISA had significantly higher villus/crypt ratios than the patients positive for AAAs [mean (SD), 1.41 (0.32) vs 0.52 (0.15); P <0.01].
Anti-tTG values and EmA titer showed a significant, but lower, inverse correlation with severity of intestinal mucosa damage [Spearman correlation, r = −0.324 for anti-tTGs, r = 0.315 for EmAs). We found no correlation between anti-tTG and AAA values.
In 19 of the 20 CD patients, AAA assay results were negative, by both ELISA and IF, after 6–12 months on gluten-free diet.
IgG AAA results were positive by ELISA (absorbance range, 0.300–0.935) in 17 of the 18 patients with type 1 AH showing anti-F-actin antibodies with IF, whereas none of samples from healthy controls gave positive results.
Gluten ingestion in CD is associated with alterations of the intestinal mucosa and interactions between gliadin and several extracellular matrix components, leading to the formation of neo-epitopes, which act as autoantigens (8)(9). AAAs, described in CD very recently (1), are among the autoantibodies consequent to the formation of these autoantigens in CD patients.
Our results show that AAAs correlate with severity of intestinal villus damage in CD patients: The AAA ELISA was positive in 51 of the 58 CD patients studied, and all patients with severe intestinal mucosa damage were positive for AAAs. The 7 patients negative for AAAs had mild villus atrophy. Furthermore, there was a highly significant inverse correlation between AAA values (assayed both by ELISA and IF) and the villus/crypt ratios in CD patients. A similar correlation coefficient (0.56) between AAA titers and severity of intestinal mucosa lesions has been reported in a multicenter study on the usefulness of AAA assays in CD (10).
Both AAA assays (IF and ELISA) were also positive in several disease controls, mainly in patients with type 1 AH and primary biliary cirrhosis. In general, the ELISA for IgA AAAs that we used in the present study showed good reproducibility and a higher sensitivity than the previously described ELISAs (1)(11)(12), and its use can be recommended. This probably depends on the use of F-actin and not G-actin as antigen in the ELISA: In vitro incubation of intestinal epithelial cells with gliadin caused intracellular actin polymerization with an increase in F-actin (2), which is the real neo-epitope recognized by AAAs. The major problem in using an F-actin ELISA is the difficulty of getting, in vitro, 100% of actin in its polymerized form (F-actin) and avoiding the unpolymerized form (G-actin), which could interfere with the quality of the test. However, the materials we used seemed to be specific for F-actin antibodies as 17 of the 18 patients with type 1 AH and none of the controls were IgG AAA-positive with the ELISA. Additional studies involving a greater number of anti-F-actin-positive patients are needed to confirm this result.
In conclusion, AAAs are a reliable marker of severe intestinal mucosa damage in CD patients, and a simple ELISA technique offers an accurate method for their determination. We propose that, after exclusion of coexistent CD and type 1 AH, intestinal biopsy is not necessary for villus/crypt evaluation in EmA/AAA-positive patients.
We are indebted to Professor A. Craxı̀ and his research group (Gastroenterology Division of Palermo University, Palermo, Italy), who provided the stored sera from patients suffering from type 1 AH. We thank Carole Greenall for excellent revision of the English. We are particularly grateful to M.G. Clemente (Pediatric Department, Cagliari University, Italy) for technical advice on the AAA IF assay. We also thank C. Scalici and G. Di Fede for assistance in revising the manuscript. This work was supported by a grant from the Ministry of Universities and Scientific Research (decr 86 Ric, dated January 30, 2002) and of the Ministry of Agriculture of Italy (DM 224/7303/02, dated June 10, 2002; project “ALICE”).
- © 2005 The American Association for Clinical Chemistry