Heat treatment prior to testing allows detection of antigen of Dirofilaria immitis in feline serum
© Little et al.; licensee BioMed Central Ltd. 2014
Received: 2 November 2013
Accepted: 17 December 2013
Published: 13 January 2014
Diagnosis of Dirofilaria immitis infection in cats is complicated by the difficulty associated with reliable detection of antigen in feline blood and serum samples.
To determine if antigen-antibody complex formation may interfere with detection of antigen in feline samples, we evaluated the performance of four different commercially available heartworm tests using serum samples from six cats experimentally infected with D. immitis and confirmed to harbor a low number of adult worms (mean = 2.0). Sera collected 168 (n = 6), 196 (n = 6), and 224 (n = 6) days post infection were tested both directly and following heat treatment.
Antigen was detected in serum samples from 0 or 1 of 6 infected cats using the assays according to manufacturer’s directions, but after heat treatment of serum samples, as many as 5 of 6 cats had detectable antigen 6–8 months post infection. Antibodies to D. immitis were detected in all six infected cats by commercial in-clinic assay and at a reference laboratory.
These results indicate that heat treatment of samples prior to testing can improve the sensitivity of antigen assays in feline patients, supporting more accurate diagnosis of this infection in cats. Surveys conducted by antigen testing without prior heat treatment of samples likely underestimate the true prevalence of infection in cats.
KeywordsAntigen test Cat Dirofilaria immitis Heartworm Heat treatment
Detection of antigen of Dirofilaria immitis is a sensitive and specific means of diagnosing heartworm infection in dogs, but these tests have been considered much less reliable in cats [1–3]. Lack of antigen detection in cats has been attributed to low circulating antigenemia due to the low number of worms often seen in feline infections, a higher likelihood of male only infections, and stunted development of D. immitis adults in an aberrant host [4–6]. In dogs, these assays are considered capable of detecting infections with as few as 1 – 3 adult female D. immitis[1, 7].
When antigen-based assays were first developed for detecting D. immitis in dogs, serum was pre-treated with heat and/or EDTA to destroy immune complexes prior to testing, as antigen-antibody complexes were recognized as inhibiting detection of antigen in some canine samples [8–10]. However, this step is no longer included in protocols of commercial D. immitis antigen tests, including those labeled for use in cats. We recently described that heat treatment of canine serum samples prior to testing can reveal antigen of D. immitis, a phenomenon which may interfere with detection of infection in as many as 7% of dogs in the southern United States [S. Little, unpublished data]. In canine serum samples which test false negative on commercial assays, antigen is apparently trapped in immune complexes, preventing detection. Once these complexes are disrupted, the assays are able to accurately detect antigen [8–10].
To determine the role antigen blocking may play in detection of D. immitis antigen in feline samples, we tested serum samples from cats experimentally infected with a low number of heartworms with four different commercially available assays before and after heat treatment of sera.
Large volume serum samples were available from six (3 male and 3 female) domestic short-haired cats experimentally infected at 10 months of age with Dirofilaria immitis by subcutaneous inoculation of third-stage larvae (L3) and confirmed to be infected by recovery of adult worms at necropsy or confirmation of histological lesions. Briefly, trickle infection of a total of 100 L3 of D. immitis was performed by subcutaneous inguinal inoculation of each cat a total of four times, on study days 7, 14, 21, and 28, with 25 L3 (Missouri strain) harvested shortly prior to inoculation from infected Aedes aegypti mosquitoes (Liverpool strain). Whole blood samples were collected from the jugular vein or, rarely, the cephalic vein, of each cat on days 84, 112, 140, 168, 196, and 224 directly into vacuum tubes containing either EDTA or no additive. Cats were cared for through Oklahoma State University’s Association for Assessment and Accreditation of Laboratory Animal Care-accredited animal resources program throughout the study; all research procedures followed a detailed animal care and use protocol approved by Oklahoma State University’s Institutional Animal Care and Use Committee. Anti-coagulated whole blood was assayed for microfilaria by modified Knott’s test and by real-time PCR for Wolbachia spp. as previously described [11, 12]. For tubes without additive, blood was allowed to clot, the serum separated by centrifugation, placed into aliquots, and stored at −80°C until further use.
Antibody testing was conducted using a commercial assay (Solo Step® FH, HESKA) according to manufacturer’s instructions, and by a fee-for-service reference laboratory (ANTECH Diagnostics) using a commercially available microtiter plate assay (Synbiotics Corporation, Zoetis). The reference laboratory tested each sample in triplicate and provided optical density (O.D.) results from each well as determined on spectrophotometry as well as corresponding positive and negative control sample wells.
All antigen testing was conducted using commercial assays according to manufacturers’ instructions, with the exception that samples were tested before and after heat treatment on each assay. For heat treatment, serum samples were placed in a heat block at 103°C for 10 minutes, the resultant coagulum centrifuged, and the supernatant used in each commercial assay. Test kits evaluated before and after heat treatment included enzyme linked immunosorbent assay (ELISA) in microtiter plate formats (DiroCHEK®, Synbiotics Corporation, Zoetis; PetChek® Heartworm PF Antigen Test, IDEXX Laboratories, Inc.), membrane bound ELISAs (SNAP® Feline Heartworm® Test, IDEXX Laboratories, Inc.), and lateral flow immunochromatographic tests (WITNESS® HW, Synbiotics Corporation, Zoetis). In addition, O.D. readings were obtained by spectrophotometry before and after heat treatment for one of the microtiter plate assays (PetChek® Heartworm PF Antigen Test, IDEXX Laboratories, Inc.) according to manufacturer’s directions.
Comparison of test results
Optical density from microtiter plate assay (PetChek® Heartworm PF Antigen Test, IDEXX Laboratories, Inc.) and mean O.D. of triplicate antibody tests (ANTECH Diagnostics) for each cat for all days on which antigen was detected in any one cat and for cats on each of three individual study days (study days 168, 196, and 224) were compared using one-way ANOVA in Excel 2007 (Microsoft Office) with significance assigned at alpha = 0.05 . An antigen/antibody ratio was also determined for each cat on days 168, 196, and 224 using O. D. and mean O. D., respectively.
Results from testing feline serum samples collected 196 and 224 days after experimental infection with 100 third-stage larvae of Dirofilaria immitis(positive results in bold)
Cat (number of worms recovered)
Antigen before heating (O. D.)
Antigen after heating (O. D.)
Antigen before heating (O. D.)
Antigen after heating (O. D.)
After heat treatment of samples, the microtiter well assays identified antigen in 1, 5, and 5 of 6 cats on days 168, 196, and 224, respectively. Antigen was also detected in 4 of 6 cats after heating samples collected on day 224 using both the membrane bound ELISA and the lateral flow immunochromatographic test. Results (O.D.) on one of the microtiter plate assays (PetChek® Heartworm PF Antigen Test, IDEXX Laboratories, Inc.) before and after heating, and of antibody assay from the same samples, for days 196 and 224 are shown in Table 1. Results (O.D.) of antigen test (PetChek® Heartworm PF Antigen Test, IDEXX Laboratories, Inc.) after heating and antibody tests were significantly associated when all days were considered together (R 2 = 0.40, F = 16.6, P < 0.001) and on samples from day 224 (R 2 = 0.70, F = 16.1, P =0.005) but not for days 168 (F = 2.6, P =0.15) or 196 (F = 0.36, P =0.56) alone. The antigen/antibody ratio was highest in the single cat for which antigen was detected prior to heating on day 196 (Table 1), although the limited number of samples precluded meaningful statistical analysis.
In the present study, we showed that heat treatment of feline serum prior to antigen testing resulted in a dramatic increase in detection of D. immitis antigen. Although only 1/6 (16.7%) samples from cats infected with a low number of adult heartworms was antigen positive prior to heat treatment, as many as 5/6 (83.3%) became positive after heat treatment when the most sensitive microtiter well plate assays were used, presumably due to destruction of antibody and release of antigen from antibody-antigen complexes . Accordingly, antigen blocking resulted in a false negative result from the majority (80%) of cats harboring both adult D. immitis and circulating antigen. Given the findings in the present study, it seems prudent to heat treat serum samples from cats with a high index of suspicion for heartworm infection prior to testing for antigen of D. immitis.
Although the American Heartworm Society feline guidelines specifically state that “antigen tests are more reliable [in cats] than generally credited,” many still have the impression that because cats generally have very low worm burdens or male-only worm infections, antigen tests are not useful in cats [4, 6]. The data in the present study are exciting in that they suggest that once antigen-antibody complexes are disrupted, antigen tests may be of great value in confirming feline infection with D. immitis, particularly the microtiter well-based assays. Heat treatment of samples prior to testing may have clinical utility, particularly given the low level of circulating microfilariae usually present in cats and difficulties in interpreting antibody test results [2, 5]. This finding may be of use not only in confirming an initial clinical suspicion of feline heartworm infection, but also for evaluating the phenomenon of self-cure in cats, which previously required evaluation of antibody levels 12 months after antigen of D. immitis was no longer identified . The prevalence of D. immitis antigen in feline samples in the United States is estimated to be 0.9% , but the data in the present study suggest that the actual prevalence is likely higher, particularly in areas where canine heartworm infection and thus spillover into cats, is endemic or hyperendemic.
Interestingly, heat treatment of sera allowed us to detect antigen in almost every infected cat included in the present study. The basis for interference with antigen detection is not fully understood, but in other systems this process is thought to be related to antigen-antibody complex formation [16, 17]. Cats infected with D. immitis develop a remarkable degree of inflammation which manifests as severe lung pathology characterized by villous arteritis, medial hypertrophy of the pulmonary arterioles, and pulmonary parenchymal damage, collectively referred to as heartworm-associated respiratory disease (HARD) [2, 3, 5]. This chronic inflammatory process likely results in hypergammaglobulinemia, which may prevent detection of D. immitis antigen by either specific or non-specific formation of antigen-antibody complexes in vivo[8, 18]. Elucidation of the mechanisms responsible for inhibition of antigen detection in feline samples as well as the best approach to disrupt these complexes will require additional research.
Antigen tests for D. immitis have long been considered of limited utility in cats. The data in the present study suggest this concept should be revisited. Heating feline serum samples prior to testing greatly facilitated detection of D. immitis antigen, presumably due to disruption of antigen-antibody complexes.
Funding to support the feline experimental infections which provided the samples used in this study was provided by Bayer HealthCare, Animal Health. Additional funding was provided by the Krull-Ewing Endowment at Oklahoma State University and the National Center for Veterinary Parasitology, which receives annual support from Bayer HealthCare, Animal Health, Novartis Animal Health, and Merial, Limited, a Sanofi Company. Dr. Lindsay Starkey, Anne Barrett, Kaylynn Gruntmeir, and the Laboratory Animal Resources staff at the Center for Veterinary Health Sciences provided outstanding technical assistance.
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