69 Canine Infectious Diseases
A multitude of infectious diseases are encountered in small animal veterinary practice and, as such, numerous diagnostic tests and techniques are used in their diagnosis. Some of the commonly used techniques and tests available in small animal infectious disease diagnosis are discussed in this chapter. Specific tests for the diseases discussed in this chapter are summarized in Table 69-1.
Serologic testing refers to the detection of endogenous antibodies directed against a particular infectious agent or detection of actual infectious antigens. Numerous techniques are used in this process. Serologic antigen detection is considered more diagnostic of infection than antibody detection because antibodies can be present in animals that were previously infected. Additionally, serologic antibody tests are more likely to have cross-reactivity with antibodies directed at other, closely related agents. Therefore high single antibody titers, or demonstration of a fourfold increase in serum antibody titers, are generally required to support active infection.
The enzyme linked immunosorbent assay (ELISA) is a commonly used rapid method of detection of antibodies or antigens. This methodology is used in commercial test kits intended for “in-house” use. Dirofilaria (antigen), Ehrlichia (antibody), and Borrelia (recombinant product that detects antibody) infections can be detected in practice settings with these kits. Individual test characteristics are important for the practitioner to understand, because false-positive and false-negative tests results are not uncommon when there is a lack of quality control in their performance.
Fluorescent antibodies (FAs) can also be used to detect antibodies or antigen. These are not performed as an in-house test. A fluorescing molecule is first bound to antibodies specific to an organism or antibody being evaluated. These are then applied to serologic, histologic, or cytologic specimens. If the organism or antibody of interest is present in the sample, fluorescence will be seen when evaluated. Fluorescent techniques have some subjectivity to their interpretation and the practitioner should be confident of individual laboratories performing these interpretations. Diseases diagnosed with this methodology include distemper, rabies, pseudorabies, Lyme disease, Rocky Mountain spotted fever (RMSF), ehrlichiosis, plague, and tularemia.
The ability of antigen or antibody to cause agglutination of red cells or other materials is used for detection of some infectious diseases. Latex agglutination is used in a kit for cryptococcal antigen detection. Agar gel immunodiffusion (AGID) and complement fixation (CF) are used to detect antibodies against some of the pathogenic fungi. In general, these two tests lack sensitivity and specificity and therefore identification of the organism is important.
The availability of polymerase chain reaction (PCR) as a method of detection is gradually increasing in veterinary medicine. Minute amounts of DNA or RNA can be detected with nucleic acid primers specific to the organism and then multiplied via rounds of replication to produce enough material for electrophoretic analysis. As a result, PCR tests can detect infections in tissues with low concentrations of antigen and detect differences in species within a genus. Along with the incredible sensitivity of these tests comes a greater effect on results if contamination occurs. Whole blood can be collected from dogs and tested via PCR for Bartonella and Ehrlichia species. Blood or joint fluid can be tested for Borrelia. Urine or renal tissue can be tested for Leptospira. Other PCR tests for canine infectious diseases are likely to become commercially available in the near future.
Lymph node and tissue aspiration are simple, inexpensive, and frequently rewarding techniques used to diagnose cases of fungal infection. A 22-gauge needle is applied to a 12-cc syringe. The needle is then inserted into the affected tissue, and either it is redirected through the tissue numerous times (“woodpecker” technique), or a light amount of suction is applied by drawing back to the 2- or 3-cc mark on the syringe. Suction is released and the needle withdrawn. The needle is removed from the syringe, air drawn into the syringe, the needle reattached, and the fluid forced out of the syringe onto a slide, culture swab, or into a glass tube depending on the intended purpose of the sample. In the case of cytology, a simple smear is made. Cytologic samples can be used for direct visualization of Histoplasma, Blastomyces, Coccidioides, Cryptococcus, Neorickettsia helminthoeca, Ehrlichia, and Clostridium. With application of FA techniques, cytologic samples can be used to verify Yersinia and Tularemia infections.
Conjunctival scrapings can be used for cytologic diagnosis of distemper. Scrapings are collected by first removing mucus and tears from the inferior conjunctiva with a cotton tip applicator. A heat-sterilized blunt metal spatula or dull scalpel blade is then used to repeatedly scrape the superficial layer of conjunctiva until a small amount of tissue can be lifted off, applied to a glass slide, and smeared. This usually causes erythema of the conjunctiva and a slight amount of bleeding. Slides are sent for distemper fluorescent antibody staining. Using routine in house stains, the slides can be evaluated for characteristic intracytoplasmic distemper virus inclusions.
Rectal scrapings can be useful for detection of histoplasmosis (Fig. 69-1). This is performed by passing a gloved finger or cotton-tipped applicator into the rectum and scraping the mucosa repeatedly. The finger or applicator is removed and the tip lightly rolled on a slide for staining with routine in house stains.
Figure 69-1 Rectal scraping from a dog presented with clinical signs of chronic large bowel diarrhea and weight loss. Cytologic examination reveals large numbers of intracellular and free Histoplasma capsulatum organisms.
Tracheal washes can be used for diagnosis of fungal and bacterial infections. A 16- to 18-gauge, 12-inch through-the-needle catheter is needed. The needle is either inserted between tracheal rings either in the distal third of the cervical trachea large dogs or in the cricothyroid ligament (space just cranial to the firm ventral ridge of the cricoid cartilage) in medium-size dogs. The area of puncture is prepared aseptically and sterile gloves are worn. Dogs that weigh less than 20 pounds are best sampled by passing a sterile red rubber catheter through a sterile endotracheal tube. After the needle is inserted and the catheter advance through it, the needle is withdrawn a short distance out of the skin. One to 10 ml of sterile saline is infused. When the animal coughs, aspirate to collect wash material. This can be repeated several times if needed. The catheter is removed and typically no bandaging is needed. Subcutaneous emphysema can occur. The fluid is used for cytologic analysis and culture as indicated.
CSF is collected in a much more refined manner and must be handled with care. Cytologic preparations are best interpreted by an experienced cytologist and made after cytocentrifugation to concentrate the low quantity of cells into a small area. The fluid must be analyzed within a short period before cellular degradation. Therefore a laboratory capable of handling CSF should be identified before collection and be reachable within 30 minutes. The atlantooccipital cistern is the most useful site of collection for most central nervous system (CNS) infections. After anesthesia has been induced and the site aseptically prepared, the dog is placed in lateral recumbency (right lateral for a right dominant person) with the neck flexed to open the atlantooccipital cistern. The space is identified as a subtle depression in the center of a triangle made by placing the left thumb on the right wing of the atlas, the left middle finger on the left wing, and the index finger on the occipital protuberance. This correlates to the cranial aspect of the atlas. A 22-gauge, 1.5- to 2.5-inch spinal needle is then slowly inserted into this space with the needle nearly 90° to the spine and directly on midline with the bevel directed cranially. It is uncommon to completely insert a 1.5-inch needle into a dog of any size. The stylet is removed intermittently to evaluate for flow so that the needle is not advanced into the cord or brain stem. If frank blood appears, the needle has been placed off midline and the needle should be removed. After CSF flows from the needle, it should be aseptically collected by allowing it to drip into an empty glass tube. Approximately 1 to 2 ml is required for most diagnostic tests. The needle is then removed. Cell counts, cytologic evaluation, and fluorescent antibody techniques can be applied to CSF for detection of canine distemper virus antigens. Titers for distemper, Ehrlichia canis, RMSF, Cryptococcus, and other infections can be determined and compared with serum concentrations for diagnosis. Fluid can be submitted for cultures.
The virus attaches to the upper respiratory epithelium. Local replication occurs in mononuclear cells and spreads to local lymph nodes and tonsils, and then to other lymphoid containing organs and epithelial tissues. Hematogenous dissemination results in infection of other epithelial organs and the CNS. Animals that mount an adequate humoral and cell-mediated response clear the virus. If immunity of the dog is inadequate, then spread of the virus to other tissues, skin, glandular tissue, epithelium of the gastrointestinal tract, and respiratory and urinary system occurs and clinical signs become evident. Shedding of virus from infected dogs may be of short or long duration, depending on the dog’s immunologic status and response.
Ocular and nasal discharges are present in early stages of the disease, and are usually associated with concomitant constitutional signs such as fever, lethargy, and anorexia. If the dog’s immunity is inadequate, secondary bacterial infections result in purulent ocular and nasal discharges and cough. Respiratory distress may be observed in severely infected animals.
Gastrointestinal signs such as vomiting and diarrhea may be observed because of the local replication of virus in these tissues. Enamel hypoplasia can be noted in young animals that were previously infected with canine distemper and subsequently develop neurologic signs.
Dermatologic signs such as impetiginous dermatitis in young dogs, or nasal and footpad hyperkeratosis may be noted. Pustular dermatitis is reported to be a good prognostic indicator but is not a consistent observation.
Neurologic signs in dogs are due to acute or chronic encephalitis. Manifestations include seizures, cerebellar and vestibular signs, paraparesis, tetraparesis, and myoclonus. Myoclonus and the classical “chewing gum fits” are features that are almost pathognomonic for this disease, but other CNS disease can produce similar clinical manifestations. However, if these signs are observed in a susceptible, unvaccinated young dog, it would lend more credence to canine distemper infection. Ocular manifestations associated with canine distemper infection include conjunctivitis, keratitis, optic neuritis, and keratoconjunctivitis sicca.
Leukopenia may be observed early in the disease and may persist in dogs with progressive disease and early neurologic manifestations. A mild anemia and thrombocytopenia have also been reported in the early stages of infection. Leukocytosis and neutrophilia result when secondary opportunistic infections are present.
Viral inclusions can be observed in erythrocytes, lymphocytes, and other mononuclear cells. These appear as round to oval gray/blue intracytoplasmic bodies. No major biochemical abnormalities are consistently noted.
A compatible history and clinical signs are usually supportive of a diagnosis of acute canine distemper. Lack of proper vaccination in a young animal should also increase the clinical suspicion. Chronic neurologic disease from canine distemper infection is difficult to differentiate from other causes of acquired seizures and paresis, based solely on historical and physical findings.
CSF analysis in dogs with acute encephalitis usually shows increases in protein concentration and increased mononuclear cells, predominately lymphocytes. Detection of anti-CDV antibody in the CSF is the most definitive evidence of distemper encephalitis, provided that blood contamination is minimal during collection. If blood contamination is present in CSF, serum, and CSF antibody titers for CDV and canine parvovirus can be measured and their corresponding CSF to serum ratios computed. If the ratio for CDV antibodies exceeds that of parvovirus, then a diagnosis of CDV infection is plausible (see Chapter 4). Immunocytology and immunohistochemistry can be used to detect viral antigen in tissue samples. Conjunctival, tonsillar, and respiratory samples may reveal a positive fluorescence test early in the course of disease before antibody titers increase. This technique may also be applied to tissue samples obtained at the time of biopsy or necropsy.
Serum titers can be detected by ELISA methodology. Immunoglobulin (Ig)M antibodies can be detected in both acute and chronic encephalopathies from canine distemper and is more definitive than IgG antibodies. IgG antibodies only denote that the animal has been exposed to the virus, either naturally or through vaccination.
There is no known effective antiviral therapy that moderates the disease. Supportive therapies consist of control of opportunistic infections and include, antibacterial therapy, and antiprotozoal therapy. Opportunistic infections reported include Bordetella spp., Salmonella spp., Toxoplasma gondii, and Neospora caninum.
Attention to general hygiene, fluid balance, and nutrition is also important. Cleaning of ocular and nasal discharges, application of ophthalmic preparations to prevent exposure keratitis, and a good high protein and caloric diet are all beneficial.
Vaccination is typically started at approximately 6 weeks to avoid lapse in protection in puppies with maternal antibody titers that are waning. Boosters are administered every 2 to 3 weeks until the dog is 16 weeks of age. The decline of maternal antibodies is predictable based on the maternal antibody titers by use of nomogram that shows the rate of antibody decline and the expected time for optimal vaccination. At least two vaccines should be administered to all dogs in the initial preventative program.
Several different vaccines are available that provide active immunization of animals. These include modified live, inactivated whole virus, subunit products, and recombinant vector–based vaccine. Use of heterologous vaccination with measles virus can provide temporary immunity in animals younger than 6 weeks.
There are two different and distinct adenoviruses that infect dogs. CAV-1 causes infectious hepatitis and upper respiratory disease, whereas CAV-2 predominately results in an acute upper respiratory disease. CAV-2 is one viral agent incriminated in the canine “kennel cough” complex.
These viruses are shed in most body secretions from acutely infected dogs. Contact of susceptible dogs with contaminated fomites results in oronasal exposure. The virus replicates in tonsils and lymphoid tissues of the oral cavity and then disseminates hematogenously to other tissues and organs.
CAV-1 infections can be commonly asymptomatic infections, based on serologic surveys, or cause acute or chronic disease syndromes. These syndromes are related to organ injuries of the liver, kidneys, eyes, and respiratory tract. The predominant organ injury with CAV-1 is the liver, where it causes hepatic necrosis. Clinical signs related to hepatic injury include fever, vomiting, diarrhea, hepatic enlargement, and abdominal pain. If the dog mounts an effective immune response early in the disease, hepatic injury may be limited. In dogs with severe hepatic necrosis, disseminated intravascular coagulation (DIC) may result and cause a hemorrhagic diathesis. Partial immune responses in infected CAV-1–infected dogs can result in a chronic hepatopathy, characterized by persistent hepatic inflammation.
Renal injury also occurs in CAV-1 infection but is subclinical. The virus localizes in the glomerular vasculature and later leads to immune complex deposition and proteinuria. Virus persists in the renal tubular epithelium and a mild interstitial nephritis can develop. Clinical signs of renal involvement are not usually significant.
Ophthalmologic signs occur in some naturally infected dogs. Keratitis and uveitis develop from virus replication and antibody production within this tissue. Mild to severe corneal edema can develop. CAV-1 can eventually result in secondary glaucoma from obstruction of the ocular drainage angle from inflammatory debris. Ocular changes may be the only clinical sign noted in mild cases of CAV-1 infection. These ocular lesions are not always reversible in the dog on its recovery.
Respiratory involvement with CAV-1 and CAV-2 infections result in mild to moderate respiratory signs and produce a dry hacking cough and abnormal lung sounds. Interstitial pneumonia may be complicated with secondary bacterial infection in severely infected, susceptible dogs.
Laboratory abnormalities depend on the severity of injury to the liver, kidneys, eye, and vascular endothelium. Hemogram changes include leukopenia and thrombocytopenia. In severe infections, DIC and associated coagulation abnormalities can be observed with increases in activated partial thromboplastin time, one-stage prothrombin time, fibrin degradation products, and d-dimer concentrations.
Hepatic enzyme elevations depend on the severity of infection and the host immune response to infection. Increases in alanine aminotransferase (ALT) and alkaline phosphatase (ALP) are usually present, with ALT concentrations being greater than ALP concentrations. Hyperbilirubinemia is not a feature of the acute hepatic syndrome because most dogs that recover do so rapidly. In animals with viral persistence causing chronic hepatic inflammation, this finding may be present along with other biochemical abnormalities associated with a chronic hepatopathy (decreased serum urea nitrogen, albumin and glucose, increased ALT, ALP, and total bilirubin).
Dogs infected with CAV-2 infections do not have any consistent laboratory abnormalities, except in cases of secondary respiratory bacterial infection in which an inflammatory leukogram may be present.
A clinical diagnosis of CAV-1 infection is difficult, but should be suspected when young, unvaccinated dogs are presented with an acute onset of gastrointestinal signs and elevated liver enzymes. Dogs presenting with this compatible history are often suspected of having toxicosis. Evidence of a hemorrhagic diathesis in dogs with the aforementioned signs and laboratory abnormalities consistent of DIC lend additional support for an acute hepatopathy with activation of the clotting pathways. CAV-1 can be isolated by viral isolation from oropharyngeal swabs, but this is not a viable diagnostic test for most clinical cases.
Histologic examination of the liver reveals an acute centrilobular to panlobular necrosis. Evidence of hemorrhage may be widespread in dogs that exhibit signs compatible with DIC. Intranuclear, acidophilic inclusion bodies can be identified in dogs that die from CAV-1 infection.
Vaccination with CAV-1 and CAV-2 modified live vaccines are highly effective in preventing disease. CAV-1 modified live virus (MLV) may result in ocular and renal lesions; therefore, a CAV-2 MLV is used to provide a heterotypic antibody response that is protective. Administration of two doses of MLV vaccine beginning at 8 to 9 weeks of age is recommended. Because of the duration of immunity with MLV, booster vaccination is not always required.
Infection is acquired when the dog ingests infected tissues. Whereas other mammals can be transient reservoirs of the virus, pigs are the predominant reservoirs of the disease and can have subclinical infections. Dogs can also contract pseudorabies by biting infected pigs or eating raw infected meat. Dogs are not a source of infection for other dogs. After ingestion, the virus ascends nerve fibers in the area of inoculation in a retrograde fashion, ultimately infecting the brain stem and cranial nerve nuclei.
Ptyalism was the most common clinical sign of pseudorabies in dogs and was reported in 100% of cases in one retrospective study. In the same report, all dogs had exposure to pigs. Restlessness, anorexia, ataxia, wandering aimlessly, tachypnea, dyspnea, vocalizing, and pruritus were associated with the disease in more than 50% of dogs with pseudorabies. Muscle spasms, vomiting, aggressiveness, trismus, dysphagia, and abnormal papillary light responses can also be seen. Death routinely occurs within a short period after the first clinical signs become apparent.
Clinical signs and a history of exposure to pigs should raise a degree of suspicion of the disease. Antemortem diagnosis is uncommon as the disease is rapidly fatal. Postmortem diagnosis is accomplished with fluorescent antibody testing or virus isolation from affected tissues.
After the virus is injected into tissue, the virus replicates within tissues myocytes and then enters surrounding nervous tissue. The virus spreads in motor and sensory neurons by axonal spread to the central nervous system. After the central nervous system is infected, the virus disseminates to other body tissues via peripheral, sensory, and motor neurons. Salivary glands eventually become infected; from there, the viral disease can then be transmitted to other susceptible animals. Not all animals infected with rabies are efficient in the transmittal of this disease through their saliva.
Depending on the region of the country, skunks, foxes (gray, Arctic, and red), raccoons, and bats account for most of the wildlife species reported with rabies. The number of cases reported in 2001 increased among bats, cats, skunks, rodents/lagomorphs, and swine and decreased among dogs, cattle, foxes, horses/mules, raccoons, and sheep/goats. The relative contributions of the major groups of animals were as follows: raccoons (37.2%; 2767 cases), skunks (30.7%; 2282), bats (17.2%; 1281), foxes (5.9%; 437), cats (3.6%; 270), dogs (1.2%; 89), and cattle (1.1%; 82).
Clinical signs vary and are mainly neurologic in nature. Ptyalism, dysphagia, and an inability to swallow may suggest pharyngeal, esophageal, or a gastrointestinal disorder, but these are the result of neurologic dysfunction. In the early states of rabies, behavioral changes are present. These changes are subtle and are not always noticed by the owner or veterinarian.
Furious and paralytic forms of the disease are described in dogs. These clinical manifestations are arbitrary and are used to describe changes in the dog’s behavior during progression of the disease. The furious form of the disease in dogs is characterized by aggression to people, animals, or inanimate objects. Dogs may appear excitable, bark, or attack imaginary objects. Ataxia or incoordination precedes the more dramatic clinical neurologic signs. Seizures, coma, and paralysis are evident before the animal succumbs of the disease.
On the basis of current methods, the most rapid and sensitive test for detecting rabies is the direct fluorescent test on brain tissue. The head from the suspected dog should be cooled and maintained on ice and not frozen. Specimens of the medulla, cerebellum, and hippocampus are used for the direct FA test.
Conventional histopathologic examination of brain tissue does not always provide confirmation of the disease because changes are mild. Acute polioencephalitis is observed early in the course of the disease, with a necrotizing encephalitis in progressive infections. Presence of intracytoplasmic inclusions, or Negri bodies, may be observed in the hippocampus and Purkinje cells of affected dogs. Special stains may be required to demonstrate these structures more readily.
The first step is to prevent contact of dogs with wildlife reservoirs such as skunks, foxes, and raccoons. Vaccination of dogs is highly effective and should be performed based on current recommendations by the National Association of State Public Health Veterinarians.
Vaccination is recommended with a vaccination approved by the Food and Drug Administration at 3 months of age and 1 year later. Booster vaccinations after the initial two vaccinations should be performed in accordance with local public health laws with an approved product, either annually or every 3 years.
The taxonomy of the Ehrlichia species has recently changed and is based on an objective classification scheme of the sequences of 16S ribosomal genes. The genus Ehrlichia belongs in the family Anaplasmataceae, which contains three other genera: Anaplasma, Neorickettsia, and Wolbachia. The Ehrlichia genus contains Ehrlichia canis, Ehrlichia chaffeensis, and Ehrlichia ewingii. Ehrlichia phagocytophila, Ehrlichia equi, and Ehrlichia platys are now classified in the genus Anaplasma, along with the agent of human granulocytic ehrlichiosis. These latter three Ehrlichia species have similar nucleotides and are considered by some to be the same organism.