Approximately 1 – 3% (1) or 5 – 12% (2, 3) of cats infected with feline enteric coronavirus (FECV) develop feline infectious peritonitis (FIP). Feline coronavirus (FCoV) is the term used to refer to FECV and its mutated form – feline infectious peritonitis virus (FIPV).
To date, not all stages of the pathogenesis of FIP have been clarified and finding a diagnosis is only possible by invasive methods or by including several laboratory tests, especially for the non-effusive or dry form of FIP.

Epidemiology and shedding of FECV

FCoV can be found in many households (4) or animal shelters worldwide, especially in multi-cat households (2, 5). Cats younger than 12 months are 2.5 times more likely to shed FECV in the faeces than cats aged 1 – 5 years (4). Most cats get infected at 6 – 10 weeks of age (usually via the mother). Normally, faecal excretion does not occur before the age of 9 weeks, but shedding from 4 weeks of age onwards has also been proven (4).

FECV may spread through the faeces for a period of 18 months after infection. In general, approximately 10 – 13% of cats become chronic carriers (chronic viral shedding) after infection, 70 – 80% of cats have a transient infection, i.e. they are intermittent shedders, 5 – 10% of cats develop resistance (1, 6). Cats that are chronic carriers of FECV may help to spread the virus within the cat population but appear to be less likely to develop FIP themselves (1).

FCoV mutation

RNA viruses usually have a very large genome and their polymerase is prone to reading errors during viral replication, so, in general, they are very likely to mutate (5).
According to current knowledge, the amino acid sequences of mutated (FIPV) and non-mutated (FECV) strains of FCoV only differ in very few individual sequence positions (2). Yet, these few changes in the amino acid sequence can still lead to a change in cell tropism of FCoV. It is assumed that FIPV, unlike FECV, does not invade intestinal enterocytes, but rather infects macrophages and monocytes where it replicates. As a result, after the mutation, it is no longer shed in the faeces. Thus, a cat suffering from FIP cannot transmit the mutated virus to other cats.
So far, no mutation is known which reliably leads to FIP when infected. Four regions are thought to be responsible for possible FCoV gene mutations causing the change in viral cell tropism. These include the open reading frame (ORF) “3a-c ORF” (the significance of the mutation is not yet clear; a virus with mutations in this sequence region is no longer excreted in the faeces), the “7a-b ORF” (the significance of the mutation is not yet clear; however, it is detected discontinuously in cases of FIP), the M gene (which is responsible for a membrane protein of the virus) and the S gene for the so-called “spike protein” (this protein is responsible for the ability of the virus to “enter” the cells) (5).

The main focus of research is currently on the detection of mutations in the spike protein, as this is thought to be the main reason for the change in cell tropism. Although a mutation in the spike protein was found in 91% of the tissue samples from cats with clinical FIP, 9% of the cats with clinical FIP had no mutation in the spike protein. Furthermore, a mutation in the spike protein was also detected in 89% of the tissue samples from cats without clinical FIP (3). It was therefore concluded that a mutation in the spike protein can more likely be used as a marker for systemic virus spread than for confirming the diagnosis of FIP (3, 7). A PCR result negative for mutations must be critically assessed, as there may still be a mutation of FCoV. A possible reason could be that the mutation is located at a different sequence site or it is just not present in the supplied material. PCR results which are positive for mutations must also be viewed critically, since cats without clinical disease can also carry an FCoV mutation, as described above.
There are other studies which suggest that several mutations must be present for the cat to develop the clinical picture of FIP (5). Thus, the exact mutation mechanisms and their effects resulting in FIP, and eventually the benefits and methods of mutation detection are not fully understood.

Risk factors for the development of FIP

In the literature, various risk factors are described that may be related to the development of FIP. The age of the cat is considered to be the most important one. Cats younger than 2 years have the highest risk of developing FIP (4, 5). After that, the risk for FIP, especially the dry form, only seems to increase again at an older age (6).
Stress in any form, e.g. change of owner, transfer to a shelter, surgery or changes in hierarchy, is another factor which is certain to have an impact. Moreover, many cats with FIP come from households with a high population density (5). According to studies, the excretion of FECV in a cat’s faeces increases 10-fold after changing its home (change of owner or shelter) – in some cats even up to 106-fold (1). Intact males have a higher risk, whereas neutered cats are less likely to develop FIP (6). Furthermore, a genetic factor is being discussed, more precisely the number of alleles encoding the feline leukocyte antigen (FLA), which is supposed to vary in different breeds. For example, Burmese cats are said to have fewer alleles than other breeds (1). This could result in a loss of FLA diversity and could consequently cause these cats to develop a poorer immune defence (1). However, there are many different studies on breed-specific FIP, with sometimes contradictory statements about the same breeds or in which a breed predisposition could not be replicated (1, 4, 6).
The theory suggesting that the interferon-γ-gene and its variants are associated with the risk of developing FIP has not yet been confirmed (4).

Diagnosis of FIP

So far, the gold standard of FIP detection remains staining the viral antigen in macrophages, which are surrounded by pyogranulomatous lesions, by means of histopathology or immunohistochemistry (7). Unfortunately, the high degree of certainty of this method is contrasted by the invasive nature of obtaining tissue samples.
As a further diagnostic component, PCR can be performed to detect FCoV (typically, real-time PCR on fluids from body cavity effusions has the highest sensitivity). According to current knowledge, all fluid or tissue samples which are PCR-positive for mutations also have a positive FCoV PCR (3, 7). Since in case of FIP, FECV is no longer excreted in the faeces due to the mutation of the virus, and since a cat can be reinfected with non-mutated FCoV at the same time despite suffering from FIP (7), FCoV PCR from faecal samples is of little help in making a diagnosis.
In general, the result of the FCoV PCR must always be assessed together with the results of other tests. Thus, for example, the Rivalta test, serum protein electrophoresis, cytology of CSF or of body cavity effusions and possibly an ultrasound scan continue to be important building blocks for the diagnosis of FIP (1, 6).

Diagnosis of FECV (non-)shedders

When identifying chronic and intermittent shedders, it is important to note that after an initial infection with FECV, the virus can be shed for over 18 months. FCoV PCR can therefore be positive over a long period of time without the cat necessarily being a chronic carrier.
There is no general recommendation on how long the test period should be (i.e. in which time frame repeated FCoV PCR tests on faeces should be carried out) through which the cat can be identified as a non-shedder. Different information can be found on this, ranging from more than 5 – 30 days (4) to at least 5 months (6) or even 9 months (1).
Of course, the general rule is: the longer the period chosen, the more certain the status of the animal.

Treatment

So far, there is no therapeutic option to avoid the fatal outcome of FIP. Only few data is available on treatment attempts with e.g. corticosteroids, chlorambucil and cyclophosphamide, polyprenyl immunostimulant or pentoxifylline (6). Furthermore, for many drugs, there are no suitable control studies or sufficient numbers of cases (6).
A small molecule from the group of nucleoside analogues, GS-441524, is currently being discussed as the most promising therapeutic option. Its mechanism of action is that this molecule is incorporated as an alternative substrate into the viral RNA chain during replication, thereby stopping the elongation of the RNA chain, as no further ribonucleic acids can be added. According to first studies, effective levels can also be achieved in the ocular chamber and the CSF. In vitro and after initial infection trials, a daily subcutaneous injection of GS-441524 seems to reduce the clinical signs of FIP, improve the general condition of the cats and significantly increase their lifespan by 8 to 17 months after diagnosis (8, 9).

Prevention

The best and only safe prevention of FIP is to prevent the cat from becoming infected with FCoV.
If a new FCoV-negative cat is supposed to join the household after a cat has died, it is best to wait for 3 months to ensure that any remaining FCoV in the household has lost its infectivity (6). In a dry environment, FCoV can be infectious for at least 7 weeks. The virus is, however, sensitive to almost all common detergents. Bleach has been described as particularly suitable (1).
Another recommendation to reduce the viral load is to clean the litter trays daily; if possible, the litter trays should be in different rooms than the food and water bowls (6).
Recent studies have shown that the choice of cat litter can help to reduce the viral load and viral transmission. According to these studies, cat litter variants that are based on clay minerals prevent, in vitro, the infection of cells with FECV and reduce the virus titre (10). However, these results are rather attributed to virus-binding properties (since clay mineral usually binds proteins and fats) than to any virus-neutralising capacity (10). It is questionable whether these virus-binding properties are also fully effective if a cat does not completely cover its faeces with cat litter. Those types of cat litter whose raw material is based on sawdust do not seem to have any virus-binding or virus-neutralising properties.
Further field research is needed to determine the effectiveness (10).
Possibly, adapted feeding, i.e. a reduction of unsaturated fatty acids and a reduction of the omega-6 to omega-3 ratio, can contribute to a less pro-inflammatory environment in the intestine and in the animal. If the animal’s condition is less pro-inflammatory, monocytes and macrophages show a lower tendency to adhere and migrate, so that the contact between virus and immune cells decreases and, thus, possible penetration and virus replication in monocytes or macrophages is reduced (1).

Dr. med. vet. Eva-Maria Wittauer

Proliferative lesions of the oral cavity are regularly seen in both dogs and cats. In contrast to nodules in the skin, which are easily visible, proliferative lesions in the oral cavity often go unnoticed for a long period. Once they reach a certain size, secondary changes like increased salivation or a decreased food-intake indicate presence of a lesion in the oral cavity. A regular and thorough inspection of the oral cavity is therefore important.

In addition to inflammatory lesions, various epithelial, mesenchymal, melanocytic or round cell neoplasia can occur in the oral cavity – similar to other locations and organ systems. However, odontogenic tumours are unique to the oral cavity and do not occur elsewhere in the body. They originate from the various components of the teeth, periodontal apparatus and surrounding stroma. Based on the cell of origin, currently a rather descriptive nomenclature for odontogenic tumours is used, which can be quite long and complicated. The nomenclature is based on the human WHO classification of odontogenic tumours, even though this classification is not always directly applicable to the veterinary odontogenic counterparts. Based on comparative pathology and continuing extending knowledge, the nomenclature has been subject to a constant change in the last decades. The unique features of a selection of four odontogenic proliferative lesions are discussed below.

Canine acanthomatous ameloblastoma

Canine acanthomatous ameloblastomas (CAA) (Figure 2) are unique to dogs, and are formerly known as acanthomatous epulis or peripheral ameloblastoma. It is the most common odontogenic tumour in dogs. CAA can occur anywhere in the oral cavity adjacent to teeth, but in about 50% of the cases the rostral mandible is affected. Especially medium to large breed dogs are reported to have CAA, although all breeds can be affected.
In general, CAA is considered as a benign neoplasm, since metastases have not been reported so far. However, despite being a rather benign lesion, CAA can show highly variable biological behavior, varying from slow growth and minimal local invasion, to rapid and extensive local invasive growth with high tropism for local bone invasion. Especially predominantly intra-osseous CAA are associated with highly aggressive biological behavior. These tumours can therefore cause marked destruction and osteolysis of the underlying mandibular or maxillary bone, with additional tooth loss, which can be identified on radiographs.

In recent years, our understanding of infectious diseases in reptiles has grown immensely. It is also clear that the connection between infection and disease is often dependent on multiple factors, including pathogen specific factors (e.g. strain specific properties, virulence factors) and host specific factors (e.g. species, age, sex), as well as environmental conditions (e.g. temperature, hygiene, time of year) and coinfections with other agents. Infections in reptiles also often persist, making quarantine particularly important in this group of animals. The diagnosis of infectious agents in reptiles has also made great strides in recent years. The following pages contain an overview of select infectious agents found in tortoises and turtles, snakes, and lizards. The overview is mainly focused on microorganisms (viruses, bacteria, and fungi), although some parasites are also included. If you are unsure what tests or what types of samples are best in a specific case, we are happy to consult with you on cases.

Dogs with otitis externa (O. e.) are frequently presented to the small animal practice. Clinical signs are head shaking, scratching, restlessness and an unpleasant odour. In most patients with O. e., it is a multifactorial disease. For one, there may be predisposing factors (breed, “swimmer’s ear”, incorrect ear cleaning). Apart from that, it can be distinguished between primary causes (especially foreign bodies, parasites, allergies) and secondary causes which maintain the disease (untreated/ incorrectly treated inflammation, pathological changes). A small amount of bacteria belonging to the physiological skin microbiome (coagulase-negative staphylococci, alpha- and non-haemolytic streptococci, Bacillus spp., Corynebacterium spp.) can be detected in the healthy external auditory canal. Furthermore, a small number of bacteria classified as pathogenic (S. pseudintermedius, beta-haemolytic streptococci, E. coli, Proteus spp., pseudomonads) can be found there. A low level of yeasts (M. pachydermatis) is also considered normal. As a result of the changed conditions in the affected ear, especially in chronic inflammation, both bacteria and yeasts can multiply well and it is sensible to take swabs from the affected ear(s) and let them be examined microbiologically. At the beginning, a cytological specimen should be prepared. On the one hand, cytology shows the amount of bacteria, and on the other hand, it is possible to differentiate between cocci and bacilli as well as to detect Malassezia and inflammatory cells. If suspicious structures are detected in cytology, it is always advisable to grow a culture and subsequently perform an antibiogram. If antibiotic and/or antimycotic treatment is necessary, various commercial ear preparations with different active ingredients are available.

Pathogen spectrum: The analysis includes 8896 ear swab samples from dogs which were examined by culture in 2016 as part of routine diagnostics. 19% of the samples did not show any bacterial growth (culture negative). Bacteria could be detected in 81% of the samples (culture positive). Of these, a pure culture could be grown in 32.5% of cases, a mixed culture in 31.7% and only a physiological microbiome in 16.8% of cases (Fig. 1).

The most common bacterium detected in the bacteriologically positive samples was S. pseudintermedius (38.7%). Almost half of it could be detected in pure culture. Beta-haemolytic streptococci were cultivated in 12.7%, P. aeruginosa in 12.3% and “other” pathogens in 11.9% of the samples. This was followed by E. coli with 6%, P. mirabilis with 5.1% and enterococci with 3.8% (Fig. 2).

Mycological examination: Of the total number of samples, 78% were examined bacteriologically and mycologically and 22% by bacteriology only. 62.8% of the mycological samples were culture positive and 37.2 % were culture negative. M.pachydermatis accounted for the largest proportion of mycologically positive samples at 98.6%. Figure 3 shows the detection rate of Malassezia in combination with each type of bacteria detected as well as the number of negative results. Candida spp. and moulds (“others”) were only detected in individual cases.

Level of resistance: the resistance of the most frequently detected bacteria to the active substances contained in ear preparations available at the time of evaluation (Fig. 4 – 9) were examined.

• Marbofloxacin and orbifloxacin: Both antibiotics belong to the group of fluoroquinolones (gyrase inhibitors). There was a higher percentage of P. aeruginosa isolates that were resistant to marbofloxacin. Resistance to orbifloxacin was increased in P. aeruginosa, P. mirabilis and E. coli. Overall, a favourable level of resistance could still be seen for both agents.
• Gentamicin and neomycin are aminoglycoside antibiotics. Both agents have a very similar resistance pattern. The level of resistance to gram-negative bacteria is good. Beta-haemolytic streptococci and enterococci show a natural resistance to aminoglycosides.
• Chloramphenicol and florfenicol belong to the fenicol group of antibiotics. They have a very broad spectrum of activity. In addition to P. aeruginosa isolates, S. pseudintermedius and P. mirabilis also show a higher percentage of resistant isolates here.
• Polymyxin B (polypeptide antibiotic) is effective against many gram-negative bacteria. Gram-positive bacteria are mostly resistant. In this analysis, gram-negative pathogens have a high percentage of resistant isolates (30% and more). All gram-positive bacterial isolates are resistant.

Diagnosing infectious diseases in rabbits is not always easy. For one thing, rabbits show clinical signs rather late and for another, there is often a time delay before they are presented to the veterinarian. Especially the diagnostic workup of infectious diseases with non-specific signs is a challenge for the practitioner. The first step is a detailed clinical history. Here, it is necessary to ask about how the animal is kept (individually or in a group), its age, vaccination status and any unusual occurrences. A thorough clinical examination and a general blood test are just as essential as the laboratory diagnosis of the infection itself. Sample material and detection methods depend on the suspected infection.

Below, some selected infectious diseases with non-specific, respiratory and/or mainly gastrointestinal signs are described in more detail. For the sake of clarity, the most important facts are presented in tabular form.

Infections with non-specific signs

In patients with non-specific signs such as apathy, lethargy and anorexia or in cases of sudden death, it is often difficult to associate them directly with a specific infectious agent. Common lethal infectious diseases in this category are rabbit haemorrhagic disease (RHD) and tularaemia.

Tularemia

Even though tularaemia is rare, recent cases in Germany and its zoonotic potential keep putting it in the spotlight again and again.
Tularaemia, also called “rabbit fever”, is a notifiable bacterial zoonosis (causative agent Francisella tularensis). The main reservoir in Germany are European hares. Yet, rabbits, rodents, squirrels and wild ruminants can also be infected.

Despite its low incidence in humans (17 – 72 cases per year in Germany) [1]), tularaemia is also notifiable in humans (§ 7 par. 1 German Infection Protection Act) and is considered an occupational disease of hunters and persons who trade in hares or process them into food products [1]. Because of the low infectious dose of 10 – 50 (!) pathogens, the severe course of the disease in some cases and good chances of recovery if treatment is started early in humans, detection should be performed soon if there is any suspicion. Infection of humans occurs via the oronasal route, the conjunctiva or via lesions of the skin or the mucous membrane. Sources are evisceration of carcasses, contaminated, insufficiently heated food, contaminated water, aerosols as well as bites/stings of arthropods (ticks, mosquitoes, horseflies).
Typical clinical symptoms in humans include: non-specific, flu-like symptoms, skin ulcerations, swollen and suppurative lymph nodes, fever, conjunctivitis and pneumonia. Aminoglycosides, fluoroquinolones, tetracyclines, chloramphenicol or rifampicin are recommended for treatment [2]. Prophylaxis in humans includes: avoiding unprotected contact with wild animals, maintaining occupational hygiene when handling sick or dead wild animals and game, only eating game dishes which are well cooked.

Table 1: Fact sheet tularaemia in rabbits and hares [1, 3 – 5]

Tularaemia		Data and facts
Causative agent		• Francisella tularensis, family Francisellaceae (y-proteobacteria) • gram-negative, non-motile, pleomorphic bacillus • host range: mainly hares (reservoir: rodents and others); Zoonosis! Notifiable!
Infection		incubation period: 3 – 5 (14) days [2, 3] • biting insects, ticks
Shedding		• secretions and excretions •  vector: ticks (can host for months, pathogen multiplication and transovarial transmission [4]), horseflies, mosquitoes
Clinical picture		mild – lethal •  acute course: apathy, fever, tachypnoea, ruffled fur, loss of shyness, swollen lymph nodes, diarrhoea, vomiting, dyspnoea, sepsis [1] •  chronic course: emaciation, spleen abscesses, liver abscesses [5]
Course		• mild to severe (lethal), death usually after 2 – 13 d due to sepsis
Diagnosis		• suspicion: clinical picture and history: contact to the “wild animal” • direct pathogen detection by PCR: EDTA blood, swab, lymph nodes, spleen • pathological examination
Treatment		none, killing!
Prophylaxis		• high tenacity (0 – 10 °C: weeks, < 0 °C: months) • easy to kill with bactericidal disinfectants (RKI list, VAH list) • there is no vaccine approved in Europe

 

Infections with respiratory signs

Respiratory signs, such as sneezing, coughing, nasal and ocular discharge and, above all, dyspnoea, occur when there are disorders in the upper and/or lower respiratory tract and inflammatory changes in the non-respiratory area that impair gas exchange. The causes are manifold.

Apart from cardiological, traumatic, degenerative and/or neoplastic changes, infections are the most frequent cause.

Snuffles (Rhinitis contagiosa cuniculi)

Snuffles is one of the most common reasons why rabbits are presented in practice.

“Snuffles” is a globally spread, ageindependent, predominantly bacterial mixed infection of the upper and sometimes lower respiratory tract. Many rabbits are asymptomatic carriers. In times of immunosuppression, triggered by stress or other underlying diseases, it may become clinically apparent. In addition to changes in husbandry and inadequate housing conditions, breed-related factors such as brachycephaly can also lead to an increased incidence [6].
Pasteurella (P.) multocida and Bordetella (B.) bronchiseptica, for example, are considered to be co-pathogens of snuffles but, just like mycoplasmas [7, 8], are also often found in the upper respiratory tract of healthy rabbits [16]. Infections with P. multocida typically cause rhinitis with mucopurulent nasal discharge, but can also manifest as otitis, conjunctivitis, pneumonia, abscesses, genital infections and septicaemia [9]. Because of their wide host range, both P. multocida and B. bronchiseptica are said to have a certain zoonotic potential and thus pose some risk to immunocompromised people or children [10, 11].

Table 2: Fact sheet snuffles [6 – 7, 12 – 13]

Snuffles		Data and facts
Causative agent		manifold, depending on stock and husbandry: •  Hein et al. 2021: 32% Pasteurellaceae, 28% Enterobacteriaceae, 13% Pseudomonaceae, 12% Staphylococcaceae [12] •  Villa et al. 2001: 43% Mycoplasma spp., 39% Bordetella bronchiseptica, 14% Pasteurella multocida, 14% Chlamydia spp., 10% Staphylococcus aureus, 6% Escherichia coli [7] •  Rougier et al. 2006: 55% Pasteurella multocida, 52% Bordetella bronchiseptica, 28% Pseudomonas spp., 17% Staphylococcus spp. [13]
Infection		• oronasal, airborne
Shedding		• respiratory secretions
Clinical picture		•  uni- or bilateral nasal stridor; sneezing; watery, later mucopurulent nasal discharge; sticky front paws; varying degrees of dyspnoea •  imparied drainage and ascending infections → conjunctivitis, otitis media/interna partly with vestibular syndrome [14]; encephalitis •  severe courses → pneumonia and sepsis
Course		• mild to severe depending on the pathogen
Diagnosis		•  nasal lavage sample –  from the deeper parts of the upper respiratory tract (to avoid contamination with intestinal/ environmental pathogens) –  clean the rhinarium with an alcohol-soaked swab – lavage sample collection with physiological saline solution using a 2- to 3-ml syringe and attached intravenous cannula –  lavage sample of the nasolacrimal duct in case of dacryocystitis •  bacteriological examination: lavage sample/swab in tube with medium •  PCR (Mycoplasma spp.): lavage sample in sterile screw cap tube without medium
Treatment		• depending on pathogen/antibiogram, mucolysis, immunostimulation
Prophylaxis		•  improving husbandry and housing conditions (excessively dry air due to heating in winter, draughts, lack of hygiene) [6]

 

Infections with gastrointestinal signs

Diarrhoea is a common problem in rabbits. In addition to dietary causes, it often results from infections with endoparasites. Apart from clinical history and clinical examination, faecal examination is indispensable for workup!

Coccidiosis

Coccidiosis is an infection caused by protozoa which is associated, to some extent, with high mortality rates, especially in young animals (see Table 3).

Table 3: Fact sheet coccidiosis [15, 16]

Snuffles		Data and facts
Causative agent		•  genus Eimeria (E.), protozoa •  intestinal coccidiosis: more than 25 different species of Eimeria, especially E. intestinalis, E. magna, E. media, E. perforans [15, 16] •  hepatic/bile duct coccidiosis: Eimeria stiedai •  host-specific and non-specific species
Infection		• peroral intake of oocysts (contaminated water, food)
Shedding		• enteral (intracelluar proliferation in intestinal mucosa) • adult animals often chronic carriers for months
Clinical picture		•  intestinal coccidiosis: mainly caecal tympany; watery, foul-smelling diarrhoea; inappetence; apathy [15] •  bile duct coccidiosis: hepatopathy, fatigue, reduced food intake, diarrhoea or constipation, ascites, icterus
Course		•  epidemic, high mortality (especially young animals), depending on infectious dose, pathogenicity and individual constitution
Diagnosis		•  faecal examination: microscopic (fresh, flotation)
Treatment		• sulfonamides, toltrazuril (not approved for small mammals)
Prophylaxis		•  oocysts remain infectious for months after sporulation in the outside world •  repeated thorough cleaning and disinfection of the surrounding areas [6]

 

Helminthiasis

Worm infestations sometimes occur in rabbits. In rabbits, particularly nematodes (roundworms/threadworms) are relevant (see Table 4). Cestodes are rare in wild rabbits and even less common in domestic rabbits. Detection is done by microscopic faecal analysis of a fresh sample and after flotation – if trematode larvae are suspected also after sedimentation.

Table 4: Fact sheet helminthiasis in rabbits [15]

Helminthiasis		Data and facts
Nematodes		Passalurus ambiguus (“rabbit pinworm”, oxyuriasis) •  frequent, parasitise mainly in the caecum •  larvae hatch in the rectum and can then be seen at the anus and/or on the faeces •  signs only after severe infestation (usually asymptomatic) •  diagnosis: tape impression from anus, eggs in faeces (fresh, flotation) Graphidium strigosum (stomach worm), Trichostrongylus retortaeformis Strongyloides spp., Trichuris leporis •  rare, especially in young animals when fed contaminated herbage •  signs: apathy, inappetence, enteritis, mucous-watery diarrhoea, cachexia, subacute to chronic catarrhal inflammation of the intestine in massive infestations •  diagnosis: flotation
Nematodes		Anaplocephalidae •  rare in wild rabbits, very rare in domestic rabbits •  intermediate host: moss mite, beetle mite ingested with the herbage •  signs young animals: catarrhal enteritis with diarrhoea, cachexia, developmental disorders, constipation in case of severe infestation •  diagnosis: flotation
Trematodes		Fasciola hepatica (common liver fluke), Dicrocoelium dendriticum (lancet liver fluke) •  rarity, usually not significant •  infection through herbage contaminated with metacercariae or infected ants •  signs common liver fluke: hepatitis, cholangitis, inappetence, cachexia, icterus, oedema formation •  signs lancet liver fluke: unnoticed, no clinical signs •  diagnosis: combined sedimentation-flotation method, most often necropsy findings

 

Conclusion

The importance of infectious diseases in rabbits should not be underestimated. With the knowledge about possible pathogens and the corresponding detection methods, diagnosis can be made without delay and treatment can be optimised.

Jana Liebscher, Dr. Jutta Hein

Pet reptiles are frequently presented to the practice for various types of skin lesions. There are many possible causes for these lesions and they are often multifactorial. These include, first and foremost, husbandry issues, such as inappropriate temperature or humidity, unsuitable substrate or furnishings or poor hygiene. A detailed clinical history and an exact diagnosis are essential for successful treatment.

The purpose of this Laboklin aktuell is to give you a brief overview of the most important infectious causes of skin lesions (excl. parasites). Mixed infections are common and many factors influence how infective dermatitis progresses. A difinitive diagnosis often requires tests such as cytology and histopathology combined with direct pathogen detection and interpretation of results in the context of the clinical presentation.

Viral causes

There are a few viruses which are frequently found in association with skin lesions in reptiles. In some cases, they are considered the primary cause, while in others, their role in the development and progression of the disease is less clear.

Iridoviridae

Iridoviruses of the genera Ranavirus and Iridovirus have been found in reptiles. Both have been described in association with of skin lesions, especially in lizards.

As the largest organ of the body, the skin is easily accessible for various examinations, yet the diagnostic work-up of dermatology patients can be extremely frustrating. It is therefore best to always examine dermatology patients according to a fixed diagnostic plan. A full medical history – if possible by means of a case history form – and a clinical dermatological examination are followed by a more detailed assessment of the lesions detected in the dermatological examination. A microscopic examination of material from the skin surface or from hair is the first step in further diagnosis. This way, bacteria, fungi, yeasts or parasites as well as inflammatory cells can quickly be identified.

Generally, sampling can be carried out easily, but the efficiency of the possible tests greatly depends on selecting the right sample material as well as on how the samples are taken.

A tape test is the best method for detecting Cheyletiella (Fig. 1). These mites cause dry seborrhoea, especially on the back. The sample material is obtained by repeatedly pressing the adhesive tape on the affected skin areas. Alternatively, scales can first be collected with a comb or, if there is excessive scaling, the material can also be collected directly from the examination table. Then, a drop of paraffin oil is applied to a microscope slide and the adhesive tape is stuck over it. The tape preparation is completely sampled with the 4x objective while the condenser is swung out/lowered, as there are usually only a few mites present.

To detect Sarcoptes mites and the less common Notoedres mites in cats, a superficial skin scraping is done. Cheyletiella, which also live on the skin surface, can sometimes be detected in the superficial skin scraping; less frequently, Demodex mites or fungal infection of hair can be found this way.

Take a used/blunt scalpel blade (to minimise the risk of injury) and paraffin oil to scrape off as many scales as possible from the previously shaved skin areas. Since Sarcoptes mites are usually only present in small numbers, it is recommended to collect the sample material from several sites (especially from the edges of the ears and on the elbow) and a large area. Moreover, sampling fresh papules increases the diagnostic accuracy.

The sample obtained is placed on a slide with another drop of paraffin oil, covered with a coverslip and examined under a microscope (4x or 10x objective with the condenser swung out or lowered).

For the diagnosis of sarcoptic mange, PCR on skin scrapings (without paraffin oil) or the determination of serum antibody titres are also available.

Deep skin scrapings are used to diagnose demodicosis.

With a scalpel blade or even a sharp spoon and paraffin oil for moistening, the skin is scraped in the direction of hair growth until capillary bleeding occurs. Between scrapings, the skin should be squeezed repeatedly to bring the mites from the depth of the hair follicles to the surface. It is best to sample those areas that exhibit efflorescences such as redness, alopecia, scaling, comedones or follicular casts. The scraped material is immediately spread on a slide and mixed well with the oil as the sample material could coagulate into lumps due to the bleeding and thus become more difficult to analyse. A coverslip is placed on top and the sample is then examined with a 4x or 10x objective. Demodex mites (Fig. 2) are sometimes present in large numbers and can also often be found along the hair roots on the specimen.

Apart from microscopic detection, there is also a PCR test available for the diagnosis of demodicosis which covers all species described in dogs (D. canis, D. injai, D. cornei) and cats (D. cati, D. gatoi, D. felis).

In some cases, skin scrapings are also recommended for cytological examinations if the skin surface is intact. However, these specimens are often not diagnostic as the cells are usually completely destroyed during sampling.

Impression smears for the cytological examination of the skin surface are prepared particularly with regard to the type of inflammation involved or secondary infections present (Fig. 3). A slide is pressed onto the skin lesion and then air-dried. When making an impression smear, it is necessary to ensure that the slide is not wiped over the skin lesions, as this destroys the cells and only nuclear remnants and chromatin fibres will be visible under the microscope. As in surgery, the motto “dab, don’t wipe” also applies here. In the microscope, the specimen is first scanned at low magnification (10x or 20x objective) to find diagnostically conclusive areas which are then evaluated in detail at high magnification (40x to 100x, oil immersion).

In Europe, cowpox virus is the most relevant representative of the genus Orthopoxvirus in the family Poxviridae. Cowpox viruses have zoonotic potential and occur endemically in Europe as well as in Northern and Central Asia.

Numerous species have been identified as hosts, including humans, pet rats and cats. Moreover, there have also been individual cases in dogs, primates, elephants, rhinos and exotic felids. Wild rodents, especially voles (e.g. bank voles), are considered the natural reservoir of cowpox virus. Domestic animals with close contact to humans pose the greatest zoonotic risk.

Feline cowpox virus infections occur rather rarely. However, they are often not recognised clinically and have become an important route of transmission from animals to humans in recent years.

Infections are typically observed in outdoor cats. They mainly occur in late summer and autumn, since this is the time the rodent population is at its peak.

The primary infection is usually the result of local bite wounds from infected prey, especially on the forelimbs, the chest as well as in the face. Initially, these are small local changes which can worsen due to secondary infections. Multiple/ generalised skin lesions may develop within a few weeks as a result of leukocyte-associated viraemia. Systemic changes are uncommon in immunocompetent cats, but fatal pneumonia is seen in kittens, immunocompromised animals and exotic felids (cheetahs).

It must be pointed out that there have been individual case reports which describe respiratory signs alone or only secondary, atypical skin lesions.
In these cases, diagnosis could only be made after taking a biopsy of the affected site or through further examination.

In dogs, tumours mostly occur at the age of 9 – 12 years. However, some young dogs (1 – 3 years) also develop neoplastic masses. So far, only a few studies have been conducted on neoplasms in dogs up to 12 months of age (Kessler & v. Bomhard 1997; Schmidt et al. 2010). Most of these neoplasms (almost 90%) were benign canine cutaneous histiocytomas, but some cases of malignant tumours (sarcomas, lymphomas, carcinomas) were described as well (Kessler & v. Bomhard 1997, Schmidt et al. 2010).

LABOKLIN’s sample material  2016 – 2019

As part of a large interdisciplinary joint research project (FORTiTHer), 170,000 data sets obtained from routine histopathological examinations (2016 – 2019) were analysed. For the results presented here, we evaluated data of dogs up to 3 years of age, for which breed and age were indicated in the clinical history and where the quality of the sample was sufficient for diagnosis (n=18,389). Overall, 225 breeds were included; the 15 most common ones are shown in Fig. 1.

Non-tumorous changes (inflammation, cysts, hyperplasia, degenerative lesions) were detected in more than half (57%) of the submitted samples.
Polyps, dysplasia or epulides, for example, were classified as tumour-like lesions (5%).
Tumours were found in 38% of all submitted samples (Fig. 2).
Within the group of tumours, 81% were benign (mainly histiocytomas, papillomas, benign mammary tumours, hair follicle tumours, lipomas, Fig. 3).
However, this also means that in dogs under 3 years of age, 19% of all tumours submitted were (semi)malignant (Fig. 4)!

Below, a few selected tumours are presented as an example.

Canine cutane histiocytoma

Canine cutaneous histiocytomas are, by far, the most common tumours in young dogs (Fig. 3). Histiocytomas are benign and undergo spontaneous regression in many cases. They are diagnosed by cytology or histology.

Mast cell tumours

Generally, it must always be presumed that a mast cell tumour has malignant potential. In dogs up to 3 years of age, they were the most frequent (semi)malignant tumours (39%, Fig. 4).
Often, the initial diagnosis of mast cell tumours is made cytologically. If possible, the following excision is done paying particular attention to the resection margins and with tumour bed biopsies being collected. Grading and examination of the tumour margins are done histologically.

In our sample material of dogs up to 3 years of age, 44.6% of the mast cell tumours were classified as grade I, 52.7% as grade II and 2.7% as grade III according to Patnaik et al. (1984) (or 97.3% low-grade and 2.7% high-grade, respectively, according to Kuipel et al. 2011).

Lymphomas

Lymphomas account for 8% of the (semi)malignant tumours in our sample material. The submitted samples mainly originated from the lymph nodes. However, the gastrointestinal tract, the internal organs and the skin were affected as well (Fig. 5).
Lymphomas are usually diagnosed by cytology and/or histology. Further tests (e.g. clonality analysis, immunohistology, immunotyping) provide for a more accurate characterisation.

Soft tissue sarcomas

In our sample material of dogs up to 3 years of age, soft tissue sarcomas were relatively common making up 10% of the (semi)malignant tumours.
Similar to mast cell tumours, a cytological diagnosis of soft tissue sarcomas can be helpful in planning the surgical procedure. Grading (McSporran et al. 2009) is then done histologically. Two thirds of the soft tissue sarcomas in our sample material were diagnosed as grade 1 (Fig. 6).

Bone tumours

A total of 199 samples from bones were sent in, of which 41% showed inflammation, 23% tumours and 18% callus formation. Nearly all bone tumours (91%) were malignant (osteosarcomas). It is interesting to note that the 46 osteosarcomas primarily came from Labradors (n=7), Golden Retrievers (n=5) and Rhodesian Ridgebacks (n=5), but only one Great Dane was included.

Bone tumours are therefore an important differential diagnosis in young animals, too.

Diagnosis is made by cytology or by histology with a representative sample being of utmost importance. In 11% of the submitted bone samples, the samples were not representative and no morphological correlate to the clinical or radiological changes could be found.

Mammary tumours

In our sample material, 69% of the mammary tumours in dogs up to 3 years of age (n=306) were benign.
But also simple and complex carcinomas (28%) as well as malignant mixed tumours (3%) were diagnosed. The breed distribution can be seen in Fig. 7.

Human coeliac disease is a gluten-induced enteropathy characterised by a specific genetic genotype (HLA-DQ2/HLA-DQ8 genes) and the detection of autoantibodies against gluten (Leonard et al. 2017).

Gluten is the main storage protein of grains (wheat, rye, barley, spelt; Fig. 1) and a complex mixture of hundreds of similar yet slightly different proteins, especially gliadin and glutenin (Biesiekierski 2017).

In humans, the clinical picture of coeliac disease was first described by Samuel Gee in 1887, although wheat was not identified as a possible trigger until about 60 years later, in 1941, by William Diecke (Diecke 1941).

Clinically, those affected mainly suffer from malabsorption, diarrhoea and children mostly from growth disorders (Andersen 1947). Prevalence in the total population is about 1%, with regional differences (Husby et al. 2012).

Two clinical pictures have been described that are associated with gluten intolerance in dogs. One is a symptom complex in Irish Setters with mainly digestive disorders, and the other is a clinical picture in Border Terriers mostly characterised by seizures.

Recently, however, mixed forms of the clinical pictures have also received greater scientific attention (Lowrie 2017).

In connection with gluten intake, digestive disorders with inappetence, chronic diarrhoea and weight loss as well as growth retardation in young animals have been described in Irish Setters similar to coeliac disease in humans. Usually, the onset of clinical signs is at the age of six months (Daminet 1996). Pathologically, increased intestinal permeability, partial villous atrophy and intraepithelial infiltration with lymphocytes can be detected. However, mucosal damage is typically less pronounced in Irish Setters than in people suffering from coeliac disease (Pemberton et al. 1997).

This gluten sensitivity in dogs, which is associated with digestive problems, appears to be breed-specific to the Irish Setter (Daminet 1996) and, according to Garden et al. (2000), is inherited in an autosomal recessive manner.

After changing to a gluten-free diet, all clinical signs usually improve significantly and also immediately (Pemberton et al. 1997).

Although elevated gliadin IgG antibody titres are detectable in people with coeliac disease and are also used as a screening test (Leonard et al. 2017), a first study by Hall et al. from 1992 did not measure any elevated gliadin IgG antibody titres in Irish Setters with gluten-sensitive enteropathy. Hall et al. (1992) speculated that immune complex binding may be a possible explanation for this. However, this connection, which so far has only been investigated in the study mentioned above, is questionable and should be examined more closely in further studies. This is because the test for the presence of antibodies could also be a valuable diagnostic tool for gluten-induced enteropathy in Irish Setters (manuscript in preparation).