Table 1: Diagnostic methods for detection of infectious diseases and their use; Source: Laboklin

Method	Direct pathogen detection	Detection of antibodies
Microscopy/Electron microscopy	X
Microbiological culture	X
Immunohistochemistry	X
Polymerase-Chain-Reaction (PCR)	X
Enzyme Linked Immunosorbent Assay (ELISA)	X	X
Immunfluorescence test (IFT)	X	X
Virus Neutralisation test (VNT)		X
Western Blot (WB)		X
Lateral Flow tests	X	X
And others

 

Different test methods can lead to differing test results and there is a great variety and discrepancy in test sensitivity and specificity. It is important to keep in mind that no test will ever provide 100% sensitivity and specificity. In many cases, a combination of different diagnostic methods will be the most helpful and meaningful.

Direct pathogen detection

There is a variety of different test methods for the direct detection of pathogens. Examples are microscopy, snap-tests, Immune Fluorescence Tests (IFT) and Enzyme Immunoassays (EIA), which can detect pathogen antigens. In addition, there is also Polymerase Chain Reaction (PCR, detection of pathogen specific genome sequences) and microbiological culture. For all these diagnostic tools, there is a distinction between targeted and non-targeted methods.

The choice of sample material is of essential importance and dependent on where the pathogen is to be expected at the time of sampling. Depending on infectious agent and test method, different materials would be valid: blood, swabs, faeces, urine, aspirate, tissue, skin scrapings, hair and so on. Which of these materials can be used is subject to various factors. Knowledge of the pathogenesis of specific infectious diseases, especially target organs and route of shedding, are crucial. In addition, the current stage of infection at the time of sampling as well as age and immune (and vaccination) status of the patient are of importance.

Preanalytics

Before submitting a sample for testing, it is important to know whether or not the infectious agents have to be alive and able to replicate. Depending on this, prompt processing of the sample (without time delay) or special transport media might be necessary, which is especially important for microbiological cultures. It is important to follow the specific instructions for the respective sample by the commercial laboratory of choice, to receive the most meaningful test results possible. For example, for PCR tests, we recommend to submit dry swabs without transport media, EDTA whole blood, fluids in sterile, uncoated sample tubes as well as unfixed native tissue.

In the best-case scenario, sampling should be done as early in the infection as possible and always before any therapy attempts have started.

While any negative test result can never rule out an infection with absolute certainty, a positive test result usually confirms the presence of an infectious agent. However, a positive result does not always present a correlation of pathogen and clinical disease. Careful attention has to be given to the interpretation of results of particularly sensitive test methods. The ability to replicate and therefore the infectivity of certain infectious agents can only be validated in microbiological culture.

Antibody detection

Serum antibodies can be of different origins:

• Maternal antibodies: These can be present in foals up to 6-8 weeks (rarely up to 6 months). This has to be taken into account when interpreting positive results in this age group.
• Vaccination: In general, no direct differentiation between antibodies formed by vaccination and infection is possible (exception are so called marker vaccines).
• Infection: Antibody titres persist for a long time after infection.

It takes a certain amount of time for the immune system to develop specific antibodies after contact with a pathogen (vaccination or infection). The first sub-group of immune globulins to appear are IgM-antibodies, which can be detected after around 1-2 weeks, depending on the infectious agent and the immune status of the host. IgG-antibodies can be detected after around 3 weeks and in most cases persist over a long period of time (Figure 1). For infections with certain pathogens, these time periods can vary and some infectious agents, for example mycoplasms, will not initiate antibody formation.

Therefore, a positive antibody titre can only be expected after a certain period of time. In the case of acute and peracute stages of disease, no antibodies can be detected and especially for viral infections, only a retrospective diagnosis is possible.

For some pathogens (e.g. West Nile virus, TBE virus and Borrelia), a simultaneous detection of both IgM- and IgG-antibodies is possible. Since both are present at different stages of infection, a concurrent detection of both will assist in determining if a recent infection is present (IgM ↑) or if the infection has persisted for a longer time (IgG ↑).

However, for most pathogens, only tests determining IgG are available. A single test of an individual sample will not provide a meaningful result. For example, a positive result could either point to a previous infection or a vaccination. If the sample has been taken too early, false negative results are possible.

In contrast, a comparison of a serum pair, that was taken around 2-4 weeks apart, will assist with diagnosis and facilitate the interpretation of test results, although in some cases, this will only be possible retrospectively. A 4-fold increase of the antibody titre, or a comparative significant rise of a detection value, indicates a recent exposition (vaccination or acute infection; Figure 2).

In cases with unclear results, or if only a direct pathogen detection has to be performed, it would be advisable to take a serum sample at the onset of the disease. This serum sample can be frozen at -20°C and can be examined as an “acute” sample together with a “convalescence” sample taken at a later date, to complete the serum pair.

Different diagnostic laboratories use different test values and also vary in test procedures and technical equipment, so results will not be directly comparable between different laboratories.
Therefore, serum pair samples have to be analyzed using the same setup and methods at the same laboratory.

To detect antibodies in serum, the most commonly used methods are Enzyme-linked immunosorbent assays or immunofluorescence tests. In-house rapid tests are also available for veterinary practice, usually in the shape of lateral-flow-devices. For more specific enquiries, Western Blot is also available for some pathogens, which can be used to confirm other test results. The most specific serological diagnostic method is the virus neutralization test, which determines a cytopathic effect in cell cultures. It detects antibodies which neutralize viruses and therefore inhibit the infectivity of these viruses. Therefore, this test will also provide an indication for the presence of protective antibodies in vivo. In contrast, the detection of antibodies using ELISA is no reliable indication of the presence of protective antibodies. Depending on the pathogen, cellular immune response might be of equal or higher importance as the humoral immune response.

For serological tests, the most appropriate sample materials are serum and plasma. These should be processed as soon as possible after blood sampling, to avoid hemolysis which can interfere with the tests. In some cases, antibodies can also be detected in other liquids, like cerebrospinal fluid or aqueous humor. Since antibodies are relatively stable in the samples, processing at the laboratory is not time-critical. Therefore, serum samples can be stored for a longer period of time, either refrigerated or frozen at -20°C.

The presence of a positive antibody titre against a specific pathogen does not provide conclusive evidence that said pathogen is indeed the causative agent of a disease. For the interpretation of these results, the whole picture, including clinical symptoms and, if available, epidemiological data, has to be taken in account.

Serological tests may offer advantages compared to direct pathogen detection, particularly, if the pathogens in question are only present in the peripheral blood seasonally or periodically or if they are only present in tissue stages.

Take home message

Due to the large variety of infectious agents, it is not possible to offer a generalised recommendation of which diagnostic test to choose. Both direct pathogen and antibody detection are valid and are of clinical use. Depending on the problem, they can be used side-by-side and complement each other (Table 2). In many cases, it is useful and necessary to perform several different test methods.

Before submitting samples for testing at diagnostic laboratories, veterinarians have to collect a sufficient anamnesis, perform a clinical exam and formulate a preliminary diagnosis, while also determining the likely stage of infection. In the end, pathogenesis of a respective infectious agent as well as available test methods will decide which diagnostic test can be performed. For the most meaningful results, the following points are of importance:

• Time of sampling
• Type, quality and quantity of the sample
• Potentially required stabilisers or transport media
• Time and conditions of transport

Dr. Michaela Gentil

 

You can find our wide range of services relating to direct and indirect pathogen diagnostics in horses in the “Infectious diseases” section at: www.laboklin.com

 

Table 2: Short and to the point – a few examples of infectious agents and possible detection methods (bold= method of choice); Source: Laboklin

Pathogen	Direct pathogen detection	Antibody detection
Equine herpesvirus 1	PCR of EDTA blood (only during fever phase!), deep nasal swab, dead fetuses and placenta, cerebrospinal fluid, all depending on clinical symptoms	Disease peracute to acute, high seroprevalence in the horse population due to widespread distribution of the virus and vaccination; retrospective detection via serum pair is possible
Borrelia burgdorferi	Direct detection is difficult; PCR can be attempted from synovia, skin- or joint biopsies, if necessary	IgM- and IgG-antibodies as screening test, Western Blot for confirmation and useful to differentiate vaccination / infection antibodies
West Nile virus	PCR of EDTA-blood often low clinical value, since viremia has subsided by the time clinical symptoms appear; virus detection by PCR can be attempted in cerebrospinal fluid or tissue (post-mortem)	Concurrent detection of IgM- and IgG-antibodies by ELISA; cross-reactions with other flaviviruses (e.g. TBE, Usutu) are possible! In positive cases, differentiation is carried out using VNT
Dermatophilus congolensis	Detection in dandruff and scab material, using cytology (limited sensitivity) or PCR	No test available
Equine infectious anaemia virus (EIA)	High genetic variability of the virus, detection difficult	Once infected, animals remain virus carriers for life (persistent infection) and seropositive; diagnosis via Coggings-test or c-ELISA. However: Incubation period is up to 3 months, tests will have to be repeated during this time! Important: Foals of infected mares can be positive for up to 6 months due to maternal antibodies.

Richard J. Piercy (RVC): Myopathies – PSSM 1/2 and atypical myopathy

1. PSSM 1/2
According to Piercy, if myopathy is suspected, it can initially be confirmed by detecting elevated serum or plasma activities of the muscle enzymes creatine kinase (CK) and aspartate aminotransferase (AST). However, (apparently) subclinical elevations of CK and AST are often found in horses with poor performance. It is usually difficult to assess the clinical relevance of these elevations. In some types of myopathies, CK and AST activities are actually within the reference range, even though the horses show clinical signs. CK and AST can be important indicator of the degree and timing of muscle damage. CK activity is highest 6 – 12 h after muscle damage and then decreases with a half-life of about 12 h. In contrast, AST activity is highest after about 24 h and may also remain elevated for several days to weeks. A CK activity of 10,000 IU/L indicates 400 – 500 mg of damaged muscle mass.
If exertional rhabdomyolysis is suspected, an exercise test should be performed. Unfortunately, there are no standardized protocols. Piercy recommends 20 minutes of light to moderate exercise (trotting) on the lunge and measuring CK and AST activities in advance, after 4 h and after 24 h. However, in horses with myopathy, the percentage increase in CK activity after exercise can vary greatly – in some horses, there may be no change at all even though there is severe muscle damage.

Polysaccharide storage myopathy (PSSM) is an exercise-induced congenital myopathy. Animals heterozygous for PSSM often have normal muscle enzyme activities. Homozygous animals, however, may have increased muscle parameters. A genetic test is available for the diagnosis of PSSM1. Equine malignant hyperthermia (EMH) and immune-mediated myositis can also be excluded by genetic testing. As far as PSSM2 or myofibrillar myopathy (MFM) are concerned, there is now information that contradicts the validity of the genetic tests available.
The diagnosis of PSSM2 can still only be confirmed by muscle biopsy and is based on amylase-resistant inclusions, myopathic muscle fibres and a negative genetic test for PSSM1. Desmin aggregates, on the other hand, confirm the diagnosis of MFM. According to Piercy, muscle biopsy is the best method for confirming equine motor neuron disease, sarcocystosis or myopathy in horses with occasional or mild increases in CK or AST activity and in horses showing other clinical signs (such as paresis) without elevated muscle enzymes. In exertional rhabdomyolysis, a muscle biopsy is not helpful as it only provides information about the severity and chronicity of the disease. However, in sport horses, a muscle biopsy may be of interest to assess the prognosis of rhabdomyolysis.
According to Piercy, non-specific myopathies with mild CK and AST elevations occur in Icelandic horses, Connemara ponies or warmblood horses.

His conclusion was: Atypical myopathies should be considered in horses with non-specific elevated CK and AST activities. For pathological examination, muscle biopsies should be sent unfixed and refrigerated in a plastic container to a specialised laboratory.

2. Atypical myopathy
The prevalence of atypical myopathy (AM) has increased over the past few years. The disease is caused by the toxin hypoglycin A (HGA) contained in the seeds and seedlings of the sycamore tree (Acer pseudoplatanus), which horses may ingest while grazing in autumn, spring and winter. The mortality rate is about 60 – 70 %. Horses with AM may show various signs of muscle pain, stiffness, weakness, head and neck hanging down, muscle fasciculations or tremors, breathing difficulties, lethargy, colic-like symptoms and, typically, myoglobinuria. Susceptibility to HGA varies between horses. This also means that horses that seem to show no clinical signs can have high levels of HGA in their blood and vice versa. According to Piercy, it was possible to detect HGA in the serum of horses with slightly increased CK activity (CK < 1000 IU/L) without any apparent cause. Subclinical cases are therefore possible, the only clinical sign may be poor performance. For the diagnosis of atypical myopathy, serum or plasma HGA as well as the metabolites can be determined. HGA is known to have a half-life of 2 days in blood. If large amounts of acylcarnitine are produced as metabolites, the survival rate of affected horses is significantly reduced.

S. Möller (Laboklin): Detection of colchicine in suspected meadow saffron poisoning
Meadow saffron poisoning is caused by the ingestion of tropolone alkaloids (e. g. colchicine) from leaves (spring), seed capsules (summer) or flowers (autumn) (Fig. 1). Horses often eat dried plant parts in hay. The intake of high doses of colchicine can lead to colic, bloody diarrhoea, circulatory disorders or even death. According to the literature, a dose of 0.17 mg/kg bw already leads to severe diarrhoea, while the lethal dose for horses is 1 mg/kg bw (per os).
The aim of the study was to develop a valid test procedure for the detection of colchicine poisoning. A total of 91 left-over urine samples from horses sent to Laboklin were analysed for cholchicine. 28 urine and blood samples (EDTA, serum) were analysed from a farm where horses became ill after ingesting contaminated hay. The horses suffered from recurrent colic of unknown cause, diarrhoea or faecal water, gastric ulcers, hypoproteinaemia, oedema as well as lameness of unknown origin. Colchicine was detected in all of the urine samples of the suspected horses (13.20 ± 32.12 ng/ml, max. 152.80 ng/ml). The blood samples, however, were tested negative for colchicine. This study shows that it is possible to detect colchicine in urine and thus support the diagnosis of meadow saffron poisoning.

Part II: AAEP Annual Convention, November 2022, San Antonio/Texas

During the “Kester News Hour”, a number of important publications from the past year were presented.

K. Thane et al.: Effect of varying sampling times on the results of the TRH stimulation test in the diagnosis of PPID
Conclusion: The second blood sample should be taken exactly 10 minutes after the TRH injection. Sampling after 9 or 11 minutes leads to approx. 10 % differences in the results and to incorrect interpretations in about 20 % of the cases.

N. Pusterla: Role of Sars-CoV-2 in the equine population
Horses are susceptible to the virus but do not develop clinical signs. They can, however, seroconvert. In a racing stable with a large number of sick jockeys, 3.5 % of the horses were seropositive, but none had a positive PCR result.

C. B. Fernandes: Behaviour and some perinatal parameters of mule foals
Worldwide, there are approx. 10 – 11 million mules, which are considered to be extremely hard-working, persevering, feed-efficient, intelligent and not easily scared. For mule foals, too, the first days of life are the most critical ones because of the change in foetal circulation and nutrition to pulmonary respiration and enteral feeding. Mule foals have a higher APGAR score than horse foals; they are faster at seeking the udder, standing and nursing. Mares with mule foals eliminate the placenta faster than mares with horse foals. However, meconium discharge is significantly later in mule foals: up to 72 hours p. p. Overall, mule foals have a faster neurological and hormonal adaptation to extra-uterine life (“hybrid vigor phenomenon”, “genetic improvement”). The gestation period until mule foals are born is the same as for horse foals.

L. Huggins: Retrospective evaluation of mares with abnormal behaviour and their endocrinological test results, especially the diagnosis of granulosa cell tumours
31,981 blood samples from mares with abnormal behaviour were included in the study. In 86 % of the mares, the endocrinological results were not suspicious. The sensitivities of the tests were 90 % for AMH, 80 % for inhibin B and 40 % for testosterone. Abnormal rectal examination findings as well as “stallion-like behaviour” in the clinical history, however, correlated with abnormal hormone concentrations.
Conclusion: If there are problems with mares showing undesirable behaviour, they are rarely found in the reproductive tract.

Equine Endocrinology Group: What is new?
Testing for PPID in horses without any signs is not recommended!
The reference ranges for ACTH were raised slightly. This leads to fewer positive but more borderline results. ACTH levels that are borderline do not allow a direct diagnosis of PPID, but these horses should be monitored further. If the ACTH levels of treated horses are still above the reference range, even though the clinical signs of the horses are much better, the dose does not automatically have to be increased! Instead, the patient’s clinical presentation should be checked more often. For borderline results where the clinical picture is also inconclusive, short-term diagnostic treatment should be considered.
For horses that do not respond well to pergolide tablets or do not tolerate them at all, a cabergoline preparation is available in the USA (human) which is injected once every two weeks and leads to an improvement in clinical signs. H. C. Schott reported on long-term treated PPID patients where he observed that adjustments in ACTH levels can still occur years after starting pergolide treatment. Overall, pergolide improves the quality of life, but not the life expectancy.
EMS: For the oral Karo Light corn syrup test, blood samples should be taken after 60 and/or 90 minutes. Insulin and glucose are determined. Alternatively, an insulin resistance test can be carried out. Fasting not required! How to proceed: baseline blood sample to determine glucose, immediately followed by injection of 0.10 IU/kg insulin.
2nd blood sample after 30 minutes: The glucose concentration should have decreased by 50 %.
It is also possible to first determine the insulin response to the usual food: Give the horses their normal food or let them graze for 5 – 6 hours. Collect a blood sample after 2 hours to determine the insulin level.

Obese horses (Fig. 2) with normal insulin: insulin regulation still works. Maybe determine leptin, but this is not yet commercially available.
T. Sundra et al. from Australia presented a very promising approach to the treatment of EMS. The group used ertugliflozin to treat hyperinsulinaemia and laminitis. It is a sodium-glucose co-transporter-2 inhibitor which promotes glucose excretion via the kidney. The study was carried out on 36 ponies. Treatment was initially given for 6 weeks, or longer in some horses. The results were promising: In all ponies, the insulin concentration was considerably reduced and the horses lost a lot of weight. Lameness also mostly disappeared with the radiographs of the horses being unchanged. No further episodes of laminitis occurred. Hepatic parameters and triglycerides should be monitored closely. Triglycerides were frequently elevated, but no horse developed hyperlipaemia. Some horses developed PU/PD during treatment. Long-term studies and controls are still lacking and the medicine seems to be very expensive. However, the results and images shown suggest that there is a promising approach to treating EMS.

Dr Antje Wöckener, Dr Svenja Möller