Serologic Test Kits

Serologic testing for FeLV is important to identify infected cats and prevent transmission of the virus ( see Feline Leukemia Virus). Testing is also recommended before vaccination against FeLV, because vaccination of infected cats will not limit development or transmission of disease. Most currently available in-house diagnostic kits are designed to detect soluble FeLV-specific gag protein p 27, which is produced in large amounts during viremia. The time between infection and the presence of detectable antigen varies but is likely to be within 28–30 days. Vaccination against FeLV will not yield a positive test result when antigen tests are used, nor will the presence of maternally derived antibodies in kittens. Testing serum, plasma, or whole blood is considered more reliable than testing saliva or tears. Most test kits have positive and negative controls incorporated into the kit, so technical problems with running the test may be detected.

A negative result on a screening test is very reliable; however, false-positive results may occur with the tests of soluble antigen, especially in a population of cats with a low incidence of disease. A true positive result may reflect either transient or persistent viremia, so clinical decisions should not be based on a single test result. Confirmation of positive results, especially in asymptomatic cats, should be pursued by additional testing such as soluble antigen testing using a kit from a different manufacturer or immunofluorescent antigen testing on blood or bone marrow. Cats with discordant results should be retested using both test methods in 60 days and should probably be considered potentially infective until their status is clarified.

Some cats previously thought to have cleared infection have been identified using real-time PCR, which is a very sensitive method to detect cell-associated FeLV proviral DNA or viral RNA. These cats are proviral DNA positive but FeLV antigen negative and are unlikely to shed virus or develop clinical disease. However, proviral FeLV could be transmitted in blood transfusions, so potential feline blood donors should be PCR negative for FeLV.

Because the concentration of FIV antigen in the blood of infected cats is often very low, in-house diagnostic tests are designed to detect anti-FIV antibodies rather than antigen. Test kits yield a positive or negative result for antibody rather than a titer. Antibodies usually develop within 60 days of infection, but this period is quite variable, and a few infected cats do not develop detectable antibody levels. Once present, antibodies appear to be present for >2 yr, except for those transiently detectable in kittens that have maternally derived antibodies. A kitten < 6 mo of age that tests positive should be retested after 6 mo of age for this reason. If the test is still positive, the kitten is most likely infected.

With the currently available ELISA in-house test kits, sensitivity is high. However, these tests cannot distinguish between antibodies produced in response to vaccination with FIV vaccine and those produced by natural infection. An ELISA test that appears to successfully discriminate between antibodies produced in response to vaccination and those produced in natural infections has been developed but is not available for commercial use. A reasonable approach is to use the in-house test as a screening test. A negative result is a fairly reliable indicator that the cat is not infected. Positive results, especially in asymptomatic cats, should be confirmed by another test such as Western blot if available. Kittens born to vaccinated queens are antibody-positive for a variable length of time.

Both ELISA and immunomigration test kits are available for detection of CPV antigen in feces. These tests are fairly specific for the virus, but dogs that have been recently vaccinated may transiently shed antigens detected by the test kits. The sensitivity is somewhat lower for several reasons. Fecal shedding occurs for only ~7–10 days, beginning on day 3–5 after exposure, so virus is not always detectable in dogs with clinical signs. Blood in the feces as well as the formation of antigen-antibody complexes, with antibodies from the blood or exudate present in the gut, may be associated with false-negative results. It appears that in-house test kits may have decreased sensitivity to the new CPV2c strain.

Although in-house test kits for diagnosis of CPV are not licensed for use in cats, several studies have shown they can be used to detect feline panleukopenia virus in cat feces. The specificity and sensitivity are not known. Some cats test positive after vaccination, even when inactivated virus vaccines are used.

In-house tests for assay of anti-CPV antibodies have become available to evaluate the need for revaccination and the presence of possibly interfering maternal antibodies. These tests are not intended to diagnose parvovirus infection and do not distinguish between natural exposure and vaccine-induced antibodies. Currently available test kits include ELISA and immunochromatographic tests. They are semiquantitative and use color changes in positive control wells compared with color changes in samples to determine relative antibody levels. The level of anti-CPV antibody protective against infection is not known, but results are interpreted based on protective titers measured by other methods. PCR is widely available for detection of virus in feces.

In-house test kits for determination of antibodies to CDV are available, usually in combination with canine parvovirus antibody testing. These tests are semiquantitative ELISA for anti-canine distemper IgG in serum or plasma. They may be used to evaluate the need for revaccination and to determine the level of maternal antibody present in puppies. They are less useful for diagnosis of infection with CDV. An immunochromatography-based assay for CDV antigen has been developed and has good sensitivity and specificity, especially when conjunctival swabs are the sample used; sensitivity and specificity are somewhat less for blood and nasal swabs. This assay is not currently available as an in-house test kit.

Lyme disease ( see Lyme Borreliosis) is caused by the tickborne spirochete B burgdorferi. In general, diagnostic tests for this infection have had several potential problems, including relatively high levels of positive results in clinically healthy animals, persistence of antibodies after apparent resolution of clinical disease, interpretation of antibody titers in vaccinated dogs, and the fact that detectable antibody may not be present until 4–6 wk after exposure. An in-clinic ELISA test kit for detection of anti–B burgdorferi antibodies in serum or plasma minimizes some of these problems by using a synthetic peptide (C₆) based on a conserved B burgdorferi–specific protein. This protein is important in inducing a humoral immune response by 3–5 wk after natural infection in dogs. Vaccination does not induce a cross-reactive immune response to this antigen, so assays using the C₆ protein are able to distinguish vaccine-induced antibodies from those produced in response to natural infections. C₆-based assays are also more specific than previous tests, because there is no cross-reactivity with other tickborne agents or with autoimmune antibodies.

A currently available in-house test assay for anti–B burgdorferi antibodies is a screening test that gives only a positive or negative result and is thus not quantitative. Although licensed for dogs, this test kit also has been validated in cats and horses. Quantitative assays, performed in reference laboratories, can be used to monitor treatment efficacy (ie, detection of decreasing antibody titers), but these results are not always conclusive.

Canine ehrlichiosis ( see Ehrlichiosis, Anaplasmosis, and Related Infections in Animals) is a tickborne disease caused by one of several species of Ehrlichia. It can be associated with thrombocytopenia, anemia, and neutropenia as well as other nonspecific clinical signs. Although identification of Ehrlichia morulae in leukocytes can be diagnostic for this infection, serologic assay for detection of antibodies is more common and has better sensitivity. In-house tests for ehrlichiosis are either qualitative (giving only a positive or negative result) or semiquantitative tests for antibodies to E canis. Those currently available are ELISA to be performed on serum, plasma, or whole blood. The test kits use either recombinant analogues of major outer membrane proteins of E canis or whole E canis proteins as the antigen. The different Ehrlichia species stimulate antibodies that appear to cross-react in these assays, for the most part. Because anti−E canis antibodies may be very long-lived and may be present in subclinical infections, detection of these antibodies does not distinguish between exposure and Ehrlichia-induced illness and cannot reliably indicate success of response to therapy.

B canis is not a problem in some countries, and tests are not always readily available. However, infection may be subclinical, or it may cause a variety of clinical signs, including abortion, infertility, and discospondylitis ( see Brucellosis in Dogs). Infection most often occurs during mating; thus, testing of breeding animals is important in disease prevention. A rapid slide agglutination test that includes a 2-mercaptoethanol (2-ME) step to reduce false-positive results by eliminating nonspecific IgM reactions is available as an in-house diagnostic kit. This assay detects serum antibody to surface antigens of the bacteria. Reports of sensitivity and specificity of this test vary depending on the confirmatory test used. ELISA and immunochromatographic assays have been developed for qualitative and semiquantitative tests for anti–B canis antibodies. Positive results obtained using any of these test kits should be confirmed with PCR or blood culture.

Serologic testing for heartworm antigen in dogs is more sensitive than screening for microfilaremia and, in addition, can detect occult infections. Heartworm antigen is first detectable at ~5 mo after infection and will usually precede microfilaremia by a few weeks; however, this time frame may be shifted later in animals on macrocyclic lactone preventives and those with low numbers of worms. Antigen test formats available include ELISA and immunochromatographic assays. Serum or plasma may be used in all kits, and some may be performed using whole blood. Some test kits can be stored at room temperature, whereas others must be stored refrigerated and brought to room temperature before use. Batch testing and single sample testing are both available. All of the test kits have very high specificity. False-positive results are most often due to technical problems such as inadequate washing or failure to read the results at the optimal time. Manufacturer’s instructions should be closely followed with any of the test formats.

Sensitivity varies from one test to another and is affected by worm load, worm gender (a female antigen is detected, thus male-only infections will not be detected), and maturity of female worms. None of these tests has 100% sensitivity. When unexpected false results are obtained on an antigen test, additional testing, using a different format, is recommended.

After adulticide treatment, antigenemia should become undetectable by 6 mo. However, it can take more than a month for some adult heartworms to die, so a positive antigen test at 5–6 mo after treatment does not necessarily indicate treatment failure. The test should be repeated 2–3 mo later.

Heartworm disease in cats is substantially different from the disease in dogs, and recommendations for serologic testing are consequently different. Cats tend to have a much lower worm burden—often only one or two worms. Cats also have single-sex infections more frequently than do dogs. Circulating microfilaremia is rare in cats, and microfilariae have a shorter life span. These differences are often attributed to a more effective feline immune response to heartworm infection. However, heartworms can cause serious disease in cats.

Heartworm antigen tests are less sensitive for detection of infection in cats than they are in dogs, with a reported sensitivity of 50%–80%. Therefore, a negative antigen test does not exclude heartworm infection. Antibody testing is also available and can be useful as a screening test, but the presence of antibody does not confirm feline heartworm disease. Transient exposure to larvae will stimulate production of antibody; many early infections are cleared spontaneously, and adult heartworms never develop. A combination of testing for both heartworm antigen and anti-heartworm antibody results in much higher sensitivity and specificity than using either test method alone. Combination testing is warranted in cats with clinical signs of heartworm disease when results of a single test are not conclusive.

Test kits for detection of heartworm antigen and anti-heartworm antibody in cats are marketed in several formats. These can use either plasma or serum and are qualitative or semiquantitative.

The only known pregnancy-specific hormone in the bitch and queen is relaxin, which is produced by the placenta when a fertilized egg is implanted. Relaxin is first detectable in the plasma around day 20–25 after fertilization. It is not present in pseudopregnant or nonpregnant bitches or queens. Relaxin levels peak by approximately day 40–50 of gestation and drop at parturition, but they may remain detectable for up to 50 days during lactation. In-clinic test kits to measure relaxin include microwell ELISA and immunomigration qualitative tests requiring serum, plasma, or whole blood. The assays appear to have very good specificity, ie, a detectable level of relaxin is not found in nonpregnant animals. False-negative results have been reported in some bitches carrying very small litters or in those with one or more nonviable puppies.

In the bitch, serum luteinizing hormone (LH) is normally present in very low levels except for a dramatic rise just before ovulation. Ovulation occurs 2 days after this LH surge, and the LH level returns to baseline within 24–40 hr of peaking. Serum progesterone levels begin to rise at the time of the LH surge. A bitch will be fertile between 4–7 days after the LH surge, with the most fertile period on days 5 and 6. In addition, the LH surge determines the gestation period, with parturition occurring between day 64 and 66 after the surge. The LH surge may occur anywhere from 3 days before to 5 days after the onset of estrous behavior, so it cannot be reliably predicted by behavior.

Ovulation timing by measurement of LH requires daily testing, which usually begins when >50% of vaginal epithelial cells are cornified, based on vaginal cytology. There is occasionally a false LH peak not followed by ovulation, but it will also not be followed by an increase in progesterone. For that reason, assays of both LH and progesterone are recommended for most accurate ovulation detection. An increased LH concentration not followed by increased progesterone levels is considered a proestrus fluctuation, and testing for ovulation should continue.

A currently available in-clinic kit to measure LH concentrations in serum is essentially qualitative, with a serum level of < 1 ng/mL being read as negative and a level of >1 ng/mL considered positive. The test is an immunochromatographic assay.

Ovariectomized bitches and queens have serum concentrations of LH >1 ng/mL. Because this LH concentration is also seen in intact bitches about to ovulate, a single high LH concentration does not confirm reproductive status, but a low LH concentration indicates a nonovariectomized animal.

Serum progesterone begins to increase after the LH surge with a modest increase on the day before ovulation and a further increase to 4–10 ng/mL on the day of ovulation. Progesterone continues to increase and stays increased throughout pregnancy or diestrus. As the rise in progesterone is more constant compared with LH, daily testing is not necessary. It is recommended that testing begin in late proestrus and continue every 2–3 days until a high range is reached, indicating ovulation.

The test kits for in-clinic progesterone testing are semiquantitative ELISA with preovulatory concentrations designated according to the kit used. Some kits are designed to give two additional progesterone ranges (intermediate and high), whereas others indicate only preovulatory and “ovulatory day or later” levels. In comparison with other test methods, the in-house test kits are less accurate, particularly in the range of roughly 1.5–10 ng/mL, which is the range of interest for earliest detection of ovulation. Accuracy is greater at higher progesterone levels. As mentioned above, measurement of both LH and progesterone is recommended for most accurate breeding management.

Assay of total serum thyroxine (T₄) may be used as a screening test for canine hypothyroidism or as a diagnostic test for feline hyperthyroidism. (Also see The Thyroid Gland.) In addition, T₄ concentrations are measured when monitoring therapy for hypo- or hyperthyroidism. An in-house ELISA test kit, which uses an additional instrument to read results, provides semiquantitative information about T₄ concentration in canine and feline sera. The assay is run at one of two “dynamic ranges,” depending on the predicted range of results. Published studies comparing this assay with other test methods for T₄ vary in their conclusions. It is recommended that T₄ concentrations be periodically measured in stored pooled serum of cats and dogs to monitor whether assay performance is consistent. It is probably still better to use a commercial laboratory for thyroxine measurement.

Almost all (99%) of total T₄ is bound to protein in the blood and therefore not biologically accurate. Free T₄ levels are a more accurate test of thyroid function in cats and dogs.

Measurement of foal serum IgG concentration within the first 24 hr after birth can be useful in preventing disease related to failure of passive transfer of IgG in colostrum from mare to foal. Use of foal-side testing procedures can facilitate prompt diagnosis and treatment. A foal serum IgG level of >800 mg/dL is generally considered optimal, with < 200 mg/dL indicating failure of passive transfer. Concentrations of 200–800 mg/dL are considered evidence of partial transfer.

Although radial immunodiffusion is considered the most accurate test for IgG concentration, it takes much longer (5–24 hr) than many of the other test methods and thus is not as useful as an indicator of the need for therapeutic intervention. More rapid screening tests include the zinc sulfate turbidity test, glutaraldehyde clot test, and ELISA test kits.

The zinc sulfate test estimates IgG in serum based on its precipitation when added to a zinc-containing solution. Turbidity generally becomes visible when IgG levels are 400–500 mg/dL. This test takes ~1 hr and may be performed with zinc sulfate solutions made in the clinic, or reagents for the test may be purchased as a test kit.

The glutaraldehyde clot test requires the addition of 1 volume of serum to 10 volumes of 10% glutaraldehyde and examination of the tubes at timed intervals up to 60 min. The presence of IgG in the serum causes a solid clot to form in the tube. Clot formation in < 10 min generally correlates to an IgG concentration of >800 mg/dL, whereas a positive reaction within 60 min is interpreted as an IgG concentration of >400 mg/dL. Both of these test methods use serum, rather than plasma, so time for the blood to clot and separation of serum must be added to the time needed to perform the test.

ELISA test kits for in-house or foal-side testing can use either serum or whole blood as the sample and take ~10 min. Most are semiquantitative, with color changes corresponding to IgG concentrations of < 400 mg/dL, 400–800 mg/dL, or >800 mg/dL. One quantitative handheld colorimetric immunoassay is currently available.

Reports of sensitivity and specificity among these methods vary. In general, sensitivity in identifying foals with IgG concentrations < 400 mg/dL appears to be acceptable for all methods. Thus, there are few foals with total failure of passive transfer that would not be identified by any of these methods. Specificity is less acceptable for some of the methods, and treatment of some foals that did not need it might be instituted based on the results.

Measurement of IgG concentration in neonatal calf serum is important for the same reasons described for foals. However, IgG levels in calves are different from those in foals, with >1,000 mg/dL considered evidence of adequate passive transfer and < 1,000 mg/dL indicating failure of passive transfer. Radial immunodiffusion is the gold standard for accurate measurement of serum IgG in calves (as in foals), but the length of time required to complete this assay makes it less useful than other methods. Other methods include zinc sulfate and sodium sulfite turbidity tests, glutaraldehyde coagulation test, measurement of total serum solids by refractometry, and a lateral flowthrough ELISA test kit.

The sodium sulfite and zinc sulfate turbidity tests are both based on the precipitation of high-molecular-weight proteins in these solutions. Serum is used as the sample, and tests are read after 15–30 min of incubation. Results of these tests vary in sensitivity and specificity depending on the endpoint chosen, and technical difficulties with reagents can decrease test performance. The glutaraldehyde clot test is performed as in foals, with no clot formation at 60 min indicating failure of passive transfer.

Measurement of serum total protein by refractometer is a fairly reliable indicator of adequate passive transfer in healthy, well-hydrated calves. A level of 5.2 g/dL is roughly equivalent to an IgG concentration of 1,000 mg/dL.

A commercially available ELISA test kit uses serum as the sample and takes ~20 min. The assay is qualitative in that it indicates an IgG concentration of >1,000 mg/dL or < 1,000 mg/dL. Sensitivity and specificity are reasonably good, and this test is less influenced by factors such as calf dehydration or reagent instability than some other methods. Manufacturer recommendations for storing and using the kits must be followed.