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December 9, 2015

Chicago Veterinary Medical Association Shaping the Future of Veterinary Medicine - Promoting the Human-Animal Bond

Proudly Presents:

INFECTIOUS DISEASES

With: JANE E. SYKES

BVSC, PHD, DACVIM(SAIM)

Co-Sponsored by:

Chicago Veterinary Medical Association – December 9, 2015

Infectious Diseases Jane E. Sykes BVSc, PhD, DACVIM Page 1 of 33

ANTIBIOTIC USE GUIDELINES: WHICH DRUG, WHAT DOSE AND HOW LONG FOR?

Jane E. Sykes, BVSc(Hons), PhD, DACVIM(SAIM) Professor of Small Animal Internal Medicine – University of California, Davis

Introduction In recent years there has been a dramatic rise in the prevalence of multidrug resistant bacteria in dogs and cats. Because of this, whenever possible veterinarians should make attempts to confirm a suspected bacterial infection by requesting microscopic evaluation of direct smears, culture and susceptibility by a laboratory before the choice is made to administer an antimicrobial drug. A Gram stain prepared from the specimen can permit the rapid preliminary diagnosis of infection, and provide information regarding whether the organism(s) present are gram-positive or gram-negative. This helps guide the clinician to select an appropriate empiric therapy, if necessary, while awaiting the results of culture and susceptibility testing. Important terms used to describe resistant bacterial infections are as follows:

• Beta lactam: antimicrobial drug that includes a beta-lactam ring (all penicillins, cephalosporins and carbapenems such as meropenem). These bind to penicillin binding proteins (PBPs) (bacterial enzymes that catalyze bacterial cell wall formation) and cause bacterial lysis.

• Beta lactamases: bacterial enzymes that destroy the beta lactam ring (associated with resistance to beta-lactams). These include a variety of penicillinases. Beta lactamase inhibitors are drugs that inhibit these enzymes and include clavulanic acid and sulbactam.

• ESBLs: extended spectrum beta lactamases. These are bacterial enzymes that destroy critical beta-lactam drugs needed for treatment of resistant bacterial infections in humans (by definition, third generation cephalosporins such as cefuroxime, cefotaxime, ceftazidime). They are generally expressed by gram-negative enteric bacteria such as E. coli and Klebsiella.

• MRS: methicillin resistant staphylococcus. These organisms express an altered penicillin binding protein (PBP2a) that does not bind beta-lactam drugs. Therefore they are resistant to penicillins, cephalosporins and carpapenems.

• MDR: multidrug resistance. By definition, this is resistance to 3 or more CLASSES of antimicrobial drugs (e.g., cephalosporins, fluoroquinolones, and aminoglycosides).

Methods of Susceptibility Testing Clinical microbiology laboratories will perform susceptibility testing for most aerobic bacteria, with the exception of streptococci. Streptococci from dogs and cats are almost always susceptible to penicillins. Most laboratories also do not routinely perform susceptibility testing on anaerobes, which also mostly have predictable susceptibilities, although resistance in anaerobes is increasing and some anaerobes, such as Bacteroides fragilis, have a high prevalence of β-lactamase enzyme production. Susceptibility testing can be performed using dilution methods or diffusion methods. The minimum inhibitory concentration (MIC) is the lowest concentration of antimicrobial drug that inhibits visible growth of an organism over a defined incubation period, most commonly 18 to 24 hours, and is



determined using dilution methods, which involve exposing the organism to 2-fold dilutions of an antimicrobial drug. The concentration range used varies with the drug and the organism being tested. Standard protocols are published by the Clinical and Laboratory Standards Institute (CLSI) that specify medium composition and pH, inoculum size (determined on the basis of turbidity measurements), inoculation procedures, agar depth and incubation conditions, as well as quality control requirements. Because failure to comply with these protocols can lead to erroneous results, veterinarians should always attempt to use laboratories that follow CLSI protocols. The most widely used dilution method in North America is broth microdilution, whereby 2-fold dilutions of antimicrobials are made in a broth media in a microtiter plate. Pre-prepared frozen or freeze-dried plates are available commercially for inoculation (e.g., Sensititre plates, TREK Diagnostic Systems). The results can be determined using visual examination of the plates for the inhibition of bacterial growth, or by the use of semi-automated or automated instrumentation. The MIC for each antimicrobial drug tested against the organism is reported to the clinician on the susceptibility panel. It is the lowest concentration of antibiotic (usually in µg/mL) that inhibits growth of the organism in vitro, and the lower the MIC, the more potent the antimicrobial is at inhibiting bacterial growth. Diffusion methods include gradient diffusion (also known as Etest®) and disk diffusion. The Etest involves use of a plastic strip coated with an antimicrobial gradient on one side and an MIC interpretive scale on the other side. An agar plate is inoculated with the organism of interest so that subsequent growth of the organism will form a “lawn”, rather than individual colonies. The strips are applied to the surface of the plate, with the lowest concentration towards the center. The antimicrobial drug diffuses into the medium, which results in an elliptical zone of growth inhibition around the strip. The MIC is read at the point of intersection of the ellipse with the MIC scale on the strip. Although the strips are expensive, Etests have the advantage of being adaptable to use with fastidious organisms and anaerobes if susceptibility testing of these organisms is deemed necessary. Disk diffusion involves application of commercially available drug-impregnated filter paper disks to the surface of an agar plate that has been inoculated to confluence with the organism of interest, and is also known as Kirby-Bauer antibiotic testing. Commercially available, mechanical disk-dispensing devices can be used to apply several disks simultaneously to the surface of the agar. The drug diffuses radially through the agar, the concentration of the drug decreasing logarithmically as the distance from the disk increases. This results in a circular zone of growth inhibition around the disk, the diameter of which is inversely proportional to the MIC. The zone diameters are interpreted on the basis of guidelines published by CLSI and the organisms are reported as susceptible, intermediate or resistant. Breakpoints and Definition of Susceptible vs. Resistant Organisms Once susceptibility testing has been performed, organisms are classified on the susceptibility panel report as “susceptible” (S), “resistant” (R), and, in some cases, of “intermediate” (I) susceptibility. This refers to a predicted in vivo situation, rather than in vitro susceptibility. The growth of “susceptible” isolates should be inhibited by concentrations of antimicrobial agent that are usually achievable in blood and tissues using normal dosage regimens. "Intermediate" isolates have MICs that approach usually attainable blood and tissue levels and for which response rates may be lower than those for susceptible



isolates. This category implies clinical efficacy in body sites where the drugs are normally concentrated (e.g., enrofloxacin and amoxicillin in urine) or when a higher-than-normal dose of a drug can be used, and acts as a buffer zone in order to prevent technical factors from causing major discrepancies in interpretations. “Resistant” isolates should continue to grow in the face of the usually achievable concentrations of the drug in blood and tissues. In order to determine if an in vivo response is likely, the laboratory refers to breakpoints, or clinical cut-off MICs (or, for disk diffusion testing, cut-off zone diameters), which are established, published, and revised regularly by committees associated with standards agencies such as the CLSI. If the MIC determined in the microbiology laboratory is lower than the published breakpoint, then the organism is defined as susceptible. The breakpoint is not reported to the clinician. Breakpoints are established on the basis of multiple factors, which include 1) a knowledge of MIC distributions and resistance mechanisms for each organism-drug combination, 2) clinical response rates in humans and animal models, 3) how the drug is distributed and metabolized in the body (pharmacokinetics), and 4) whether the drug is concentration-dependent or time-dependent as it relates to antibacterial effect (pharmacodynamics). Zone diameter breakpoints for disk diffusion testing are determined by correlation with MIC values. For simplicity, breakpoints are established for bloodstream infections, and are based on a specific dosage regime for the antimicrobial drug tested, which are selected by the standards agency involved. Because some antimicrobials are concentrated extensively in urine, some veterinary laboratories may report urine MIC panels, which provide breakpoints for lower urinary tract infections, which are higher than corresponding serum MIC breakpoints. These have been controversial because the possibility of concurrent pyelonephritis cannot always be ruled out. Breakpoints are often re-evaluated when new mechanisms of resistance appear in bacteria or when new data are generated that improve understanding of the pharmacokinetics and pharmacodynamics of an antimicrobial drug. The Clinician’s Role in Interpretation of Susceptibility Panels The veterinary clinician should always remember that the list of drugs reported in the susceptibility panel is simply just a list of drugs tested. They are not suggestions from the laboratory for patient care. The clinician should always ask 4 main questions, in order, when faced with a susceptibility panel:

1. Is this organism that was cultured likely to be the cause of disease? (i.e., should I treat this organism?)

Once a positive culture has been obtained, the veterinarian must consider the significance of the positive test result, even if susceptibility test results are reported. The detection of bacterial organisms within a sample does not always imply that the organism is causing the animal’s clinical signs. Contamination is the most common cause of false positive cultures. Isolation of only one or two colonies of coagulase-negative staphylococci, Bacillus spp., Corynebacterium spp., and propionibacteria commonly suggest contamination. Isolation of large numbers of a single type of bacteria from a normally sterile site is generally clinically significant, especially when supported by cytologic examination of a stained smear that demonstrates the presence of bacteria within leukocytes.

2. Are any of the drugs shown as “susceptible” the appropriate drugs for treatment of the bacterial species cultured?



Laboratories often (but not always) report results for specific antimicrobials on the basis of the organism being tested (e.g., cephalosporins may not be reported for enterococci because of intrinsic resistance). Certain antimicrobials should be generally be reserved for treatment of multiple-drug resistant organisms that cause life-threatening infections (e.g., vancomycin, linezolid, meropenem).

3. Assuming the drugs are active against the bacterial species isolated, are the drugs the right

drugs for the patient in question? a. Will they achieve adequate concentrations at the site of infection? b. What route of administration is necessary and can the antimicrobials be administered by the

route that is most appropriate for my patient? c. Could adverse drug reactions occur in this patient with these antimicrobials? d. Could drug interactions occur in this patient with these antimicrobials?

For infections in sites such as the CNS, the clinician needs to consider whether or not an antimicrobial to which the organism is reported as susceptible will penetrate that site. The clinician should also consider other factors, such as immunosuppression, pregnancy and other concurrent illness or drug therapy, when treating infections on the basis of antimicrobial susceptibility test results.

4. Is the antimicrobial drug currently being administered the most appropriate for the infection I am trying to treat?

Because antimicrobial susceptibility testing results are generally not available until 2-3 days after submission of a specimen for culture, in animals that are critically ill, antimicrobial therapy may already have been initiated by the time those results are available. The susceptibility results may show that the organism is resistant to a drug being used, in which case the drug should be changed to one that the organism is susceptible to. The susceptibility pattern can also aid in choosing an alternate drug when the patient does not tolerate the initial drug prescribed. Susceptibility testing may indicate that the organism is susceptible to a more narrow-spectrum (and generally less expensive) antimicrobial drug than the drug initially prescribed, in which case the treatment should be changed to minimize the development of antimicrobial resistance. Below is a brief summary of the current ISCAID Antimicrobial Use Guidelines for urinary tract infections and superficial pyoderma for practitioners’ reference. Doses of specific antimicrobial drugs are listed in table format in the guidelines themselves, which are available at www.iscaid.org. Guidelines for Urinary Tract Disease Simple Uncomplicated Urinary Tract Infection (UTI) Definition: Sporadic bacterial infection of the bladder in an otherwise healthy individual with normal urinary tract anatomy and function.

• A clinically significant infection implies the presence of dysuria, pollakiuria, and/or stranguria. Diagnosis of UTI cannot be made on the basis of clinical signs alone.

• Sediment analysis alone is not adequate for diagnosis because of the variable quality of interpretation. It is supporting evidence for the presence of UTI.



• Complete urinalysis and quantitative aerobic C&S testing should be performed for all cases. Free-catch samples should not be used.

• For cystocentesis specimens, counts ≥ 103 CFU/mL indicate UTI. For catheterized specimens, counts ≥ 104 in males and ≥ 105 CFU/mL in females are significant.

• Bacterial isolation should only be attempted in clinics with appropriate laboratory facilities, proper biosafety containment and waste management, and adequately trained individuals. A recent study showed that in-house “urine paddles” may be useful to rule out the presence of infection but these do not reliably identify bacteria and can generate false negative results (Ybarra et al, 2014).

• Treatment is indicated to relieve patient discomfort while awaiting C&S test results. Recommendations for initial treatment are amoxicillin (11 – 15 mg/kg PO q8h) or trimethoprim-sulfonamide (15 mg/kg PO q12h). Additional information obtained from the CLSI since publication of the guidelines suggests that amoxicillin could be used q12h.

• Veterinarians are encouraged to document and monitor resistance patterns among isolates from their hospital.

• If C&S testing reveals a resistant isolate and there is a lack of clinical response, treatment should be changed to an appropriate antimicrobial drug.

• Treatment is typically for 7 to 14 days, but shorter treatment times (<= 7 days) may be effective. Since publication of the guidelines, two studies have provided evidence to support this. In one study, a 3-day course of once daily, high dose enrofloxacin was as effective for treatment of uncomplicated UTIs in dogs as 14 days of clavulanic acid-amoxicillin (Westropp et al, 2012). In another study, a 3-day course of trimethoprim-sulfamethoxazole was as effective for treatment of uncomplicated UTIs in dogs as 10 days of cephalexin (Clare et al, 2014), although long-term microbiologic cure rates in both groups of dogs were less than 50%.

• There is no evidence that intra- or post-treatment urinalysis or urine culture is indicated in the absence of ongoing clinical signs of UTI.

Complicated UTI Definition: a UTI that occurs in the presence of an anatomic or functional abnormality or a comorbidity that predisposes the animal to persistent infection, recurrent infection, or treatment failure. Recurrent UTIs, as defined by the presence of 3 or more episodes of UTI during a 12-month period, also indicate complicated infection.

• The same general principles as for uncomplicated UTI apply • Efforts should be made to identify the underlying cause; consider referral • Treatment should be based on the results of C&S testing • Although 4 weeks is generally used for treatment, more studies are needed that evaluate the

shortest duration of treatment for different comorbidities. • Urine culture could be considered after 5-7 days of treatment to ensure treatment has been

effective, and should be performed 7 days after treatment is discontinued. If there is a lack of clinical response, additional investigation and management of the underlying cause should be made.



• There is insufficient evidence to recommend “pulse” or chronic low-dose treatment, urinary antiseptics, and nutritional supplements such as cranberry juice extract for prevention of UTIs.

Subclinical Bacteriuria Definition: presence of bacteria in the urine as determined by positive bacterial culture, in the absence of clinical signs of UTI.

• Treatment may not be necessary, but could be considered if there is a high risk of ascending or systemic infection (e.g. patients with underlying renal disease)

• Diagnosis and management of the underlying cause is critical Urinary Catheters

• Clinical signs of UTI absent: no culture or treatment indicated. • Removal of urinary catheters: urine culture is reasonable if the risk and implications of a UTI are

high. There is no indication for routine use of prophylactic antimicrobials. • Clinical signs of UTI present: perform a culture after replacement of the urinary catheter with a

new catheter. Several mL of urine should be removed to clear the catheter before a specimen is obtained for culture. Alternatively, remove the catheter and perform a cystocentesis. Culture from the collection bag, and culture of the catheter tip after removal are not recommended. Treatment should follow the guidelines for complicated and uncomplicated UTIs, and is more likely to be successful after catheter removal.

Pyelonephritis

• C&S testing should always be performed. • Treatment should be initiated while awaiting culture results, using antimicrobials effective

against Gram-negative Enterobacteriaceae. A fluoroquinolone is a reasonable first choice, after which treatment should be based on C&S results. If combination treatment was used initially and C&S results indicate that both drugs are not required, the spectrum should be narrowed.

• Treatment for 4 to 6 weeks is recommended until further information becomes available. • Culture is recommended 1 week after starting treatment and 1 week after treatment is

discontinued. Guidelines for Superficial Bacterial Folliculitis in Dogs (Superficial Pyoderma) The cause of most superficial bacterial folliculitis (SBF) in dogs is Staphylococcus pseudintermedius. In the past, S. pseudintermedius infections could be routinely treated successfully with beta-lactam antimicrobials. However, methicillin-resistant S. pseudintermedius (MRSP), which is often highly multi-resistant, has now spread rapidly throughout the world. This has focused attention on the need to ensure that diagnosis and treatment of SBF is managed in an optimal way so as to resist the spread of this organism, the proliferation of resistant clones, and the wasteful and ineffective use of antimicrobial drugs to which this organism is resistant.



• Staphylococci are not able to invade normal skin and thus SBF only occurs when there is an underlying problem such as allergic dermatoses, hyperadrenocorticism, hypothyroidism, or demodicosis. These need to be differentiated from SBF.

• Early signs of SBF are papules and pustules associated with the hair follicles. Subsequently, annular areas of alopecia, scaling, erythema and hyperpigmentation may appear, commonly surrounded by epidermal collarettes.

• Diagnosis of SBF should be supported by cytological examination and the demonstration of coccoid bacteria associated with inflammatory cells and within phagocytes. Inflammatory infiltrates and phagocytosis may be reduced or absent in patients with immunosuppression associated with underlying diseases or therapy. Cytology is also important for identification of co-infection with Malassezia pachydermatis.

• SBF must be distinguished from other causes of folliculitis and pustular disease. It should be differentiated from demodicosis by deep skin scrapings and from dermatophytosis by fungal culture. Such tests are required when history and clinical signs are suggestive but not typical of SBF, or when antibacterial treatment fails.

• Culture is essential if there is a poor response to 2 weeks of appropriate systemic antimicrobial therapy, emergence of new lesions 2 weeks or more after the initiation of such therapy, presence of residual lesions after 6 weeks of therapy combined with cytology demonstrating infection with cocci, when cytology reveals intracellular bacterial rods, and when there has been a history of a multidrug resistant infection in the dog or a dog that shares the household. C&S testing is encouraged in cases of recurrent SBF infection or when there has been a history of treatment with antimicrobial drugs.

• Owners should be taught how to recognize when progress is not being made so that they may seek veterinary advice prior to the end of a course of treatment.

• Specimens for culture should be taken from pustules. If pustules are not found specimens may be taken from pus beneath crusts, and from papules or epidermal collarettes. No surface disinfection should be carried out.

• Contact with owners should be maintained to promote effective compliance and to determine if and when failure to respond to treatment, or recurrence, occurs.

• In recurrent infection, an effective diagnostic plan should be formulated to identify and resolve underlying disease.

• Topical antimicrobials (ointments, gels and creams) should be considered, which can be applied to localised lesions 2 to 3 times daily. Such products may contain antibiotics (e.g. mupirocin), silver sulfadiazine, benzoyl peroxide and hydroxyl acids (e.g. lactic acid). More extensive areas of infection can be treated with shampoos, lotions, sprays and rinses. These contain antiseptics such as chlorhexidine, benzoyl peroxide, ethyl lactate, povidone iodine, triclosan and hydroxyl acids; they are commonly used 2 to 3 times weekly and until 7 days after lesions resolve, with contact times of 10 minutes if possible before rinsing or conditioners. They can then be used for prophylaxis on a weekly or less frequent basis depending on response.

• Selection of appropriate drugs for systemic therapy depends on availability, safety, cost, local prevalence of resistant staphylococci and patient specific factors. Empirical choices can be made



in non-recurrent cases and when there has been no history of antimicrobial drug exposure. Otherwise, drug selection should be based on C&S tests. First-tier empirical drugs include clindamycin, first generation cephalosporins, potentiated sulphonamides, and lincomycin. When first-tier drugs are not appropriate and topical therapy cannot be used, second tier drugs may be chosen; these include fluoroquinolones, doxycycline, chloramphenicol and rifampin. Most of the Working Group members felt that third generations cephalosporins should be second tier drugs, but the evidence to support their placement in this category was lacking, and so they were listed as first or second tier drugs.

• Systemic antimicrobial drugs given at subtherapeutic doses or as pulse therapy have been used to prevent or delay recurrence but such protocols are likely to promote the development of antimicrobial resistance and are strongly discouraged.

References 1. Weese JS, Blondeau JM, Boothe D, Breitschwerdt EB, Guardabassi L, Hillier A, Lloyd DH, Papich MG,

Rankin SC, Turnidge JD, Sykes JE. Antimicrobial Use Guidelines for treatment of urinary tract disease in dogs and cats: antimicrobial guidelines working group of the International Society for Companion Animal Infectious Diseases. Vet Med Int 2011; Epub Jun 27.

2. Hillier A, Lloyd DH, Weese JS, Blondeau JM, Boothe D, Breitschwerdt EB, Guardabassi L, Hillier A, Lloyd DH, Papich MG, Rankin SC, Turnidge JD, Sykes JE. Guidelines for the diagnosis and antimicrobial therapy of canine superficial bacterial folliculitis (Antimicrobial Guidelines Working Group of the International Society for Companion Animal Infectious Diseases). Vet Dermatol 2014;25(3):163-e43.

3. Ybarra WL, Sykes JE, Wang Y, Byrne BA, Westropp JL. Performance of a veterinary urine dipstick paddle system for diagnosis and identification of urinary tract infections in dogs and cats. J Am Vet Med Assoc 2014;244(7):814-819.

4. Westropp JL, Sykes JE, Irom S, Daniels JB, Smith A, Keil D, Settje T, Wang Y, Chew DL. Evaluation of the efficacy and safety of high dose short duration enrofloxacin treatment regimen for uncomplicated urinary tract infections in dogs. J Vet Intern Med 2012;26(3):506-512.

5. Clare S, Hartmann FA, Jooss M, Bachar E, Wong YY, Trepanier LA, Viviano KR. Short- and long-term cure rates of short-duration trimethoprim-sulfamethoxazole treatment in female dogs with uncomplicated bacterial cystitis. J Vet Intern Med 2014;28(3):818-826.



UPDATE ON CANINE INFECTIOUS RESPIRATORY DISEASE COMPLEX


Introduction Canine infectious respiratory tract disease (CIRD) remains a major problem in shelter and boarding kennel environments, despite widespread vaccination against the disease. As a result of improvements in diagnostic testing, there is increasing awareness of mixed infections in affected animals. In environments such as shelters, co-infections with a variety of different viruses and bacteria may be more common that infections with a single pathogen. In addition, several pathogens have emerged in recent years as important contributors to CIRD in kennel and shelter situations. Pathogens causing CIRD can help each other to infect the host. For example, canine distemper virus causes profound immunosuppression, which predisposes dogs to infection with other respiratory viruses and bacteria. Severe disease is more likely to be associated with co-infections. Single infections may be present in some animals that show no signs of illness. Similar findings have been reported in children with community-acquired pneumonia. Differential Diagnosis Understanding the differential diagnosis for CIRD is important because it aids selection of appropriate diagnostic tests, the design of rational therapy, and permits institution of proper preventative measures for CIRD. There are now at least 9 organisms known to play a role in canine infectious respiratory disease. Bacterial causes of canine infectious respiratory disease include Bordetella bronchiseptica, Streptococcus equi subspecies zooepidemicus, and Mycoplasma spp. Viral causes of canine infectious respiratory disease include influenza viruses, canine distemper virus, canine respiratory coronavirus, canine parainfluenza virus, canine adenovirus (especially canine adenovirus-2), and canine herpesvirus. Establishment of a diagnosis may not be necessary in dogs that are otherwise healthy but just have the characteristic, ‘honking’ cough of the kennel cough syndrome. The vast majority of these dogs will have self-limiting infections, with clinical signs generally resolving within 5-7 days without antimicrobial therapy. Some dogs may require a short course of antimicrobial therapy, but it is recommended that antibiotic treatment be withheld if uncomplicated infection is present and clinical signs have been present for less 10 days. A cough suppressant such as hydrocodone could be considered in this situation, but cough suppression is contraindicated in dogs with complicated disease (moist cough, pulmonary infiltrates, fever, lethargy, inappetence). Diagnostic testing is indicated if

• An outbreak has occurred. • Affected dogs are systemically unwell.



• The cough is persisting despite treatment. Establishment of a diagnosis can help with control and prevention in kennel situations, and appropriate antimicrobial therapy for dogs with bacterial infections, e.g., Bordetella bronchiseptica infections. Some B. bronchiseptica infections can be refractory to treatment with systemic antimicrobial drugs. This may result from antimicrobial resistance or inadequate drug penetration to the site of infection. Clinical signs are not useful for diagnosis of a specific infectious agent, because the signs are overlapping and non-specific, and mixed infections are commonly present. Diagnostic tests available for diagnosis of canine infectious respiratory disease include culture for bacteria and mycoplasmas, blood tests (serology) for antibody against canine influenza virus and canine distemper virus, and polymerase chain reaction testing of throat swabs or respiratory lavage specimens for the DNA and RNA of respiratory viruses and bacteria. Many laboratories now offer canine respiratory disease PCR panels. This has led to increased detection of canine infectious respiratory pathogens and an increasing awareness of co-infections. Virus isolation in culture is cumbersome and is not widely offered for routine diagnostic purposes. Sometimes a diagnosis is best obtained by combining multiple different diagnostic modalities. Culture remains a useful test for bacteria such as Bordetella bronchiseptica and mycoplasmas, although the growth of mycoplasmas can be slow and unreliable. Culture also allows susceptibility testing for B. bronchiseptica, as some strains may demonstrate antibiotic resistance. The results of serologic testing may be difficult to interpret as a result of prior vaccination. Vaccination can help to reduce the severity of disease but does not prevent infection. EMERGING AND RE-EMERGING RESPIRATORY PATHOGENS OF DOGS Bordetella bronchiseptica This is the most common bacterial agent causing CIRD, and tends to cause moderate signs of CIRD. Infection is best diagnosed via transtracheal washing or bronchoalveolar lavage, but occasionally throat swabs or nasal washings/swabs will be positive. Both culture and PCR assays are available for detection of Bordetella bronchiseptica. Parenteral, intranasal or oral vaccines are available to help to prevent bordetellosis, but the relative efficacy of these vaccines remains unclear. Dogs vaccinated with parenteral vaccines require two doses given 4 weeks apart for initial protection, and protection does not become effective until one week after the second dose. Only a single dose of an intranasal vaccine is required. Annual boosters are indicated thereafter for both parenteral and intranasal vaccines. Inadvertent administration of intranasal or oral Bordetella vaccines can lead to cutaneous abscesses or life-threatening systemic infections and death, so it is particularly important to pay attention to the vaccine type and the proper route of administration. If inadvertent administration of these vaccines occurs, immediate treatment with doxycycline is indicated; immediate subcutaneous administration of gentamicin and crystalloids at the site of inoculation has also been advocated. Bordetella bronchiseptica has the potential to cause respiratory disease in immunocompromised humans, but there is no clear evidence that the organisms in canine avirulent live vaccines are capable of contributing to human illness.



Streptococcus spp. Streptococcus equi subspecies zooepidemicus is a beta-hemolytic streptococcus that has caused outbreaks of acute suppurative or necrotizing hemorrhagic pneumonia in shelter situations. Streptococcus canis can be found in the lungs of both healthy dogs and dogs with kennel cough, whereas S. equi is rarely found in healthy dogs. Whether it acts as a primary pathogen or secondary invader is not clear, but in a recent outbreak from California, the consistent presence of co-infection was not documented. It is rarely isolated from household pets. No vaccine is available. Mycoplasmas Mycoplasmas are normal flora in the respiratory tract of dogs, but are occasionally isolated from dogs with infectious respiratory disease without evidence of coinfection. The primary mycoplasma associated with lower respiratory disease in dogs may be Mycoplasma cynos. Other mycoplasmas have been isolated from the respiratory tract of dogs, but these have not been definitively associated with lower respiratory disease. Molecular techniques have improved our ability to detect mycoplasmas, but we still have trouble knowing whether a positive result is associated with disease. No vaccine is available. Influenza viruses Influenza viruses are enveloped viruses with segmented single-stranded RNA genomes that belong to the family Orthomyxoviridae. Influenza viruses that cause disease in domestic animals belong to the genus Influenzavirus A. Influenza A viruses are classified based on the composition of their hemagglutinin (H) and neuraminidase (N) genes. To date, 18 H types and 11 N types have been identified, each of which are antigenically distinct. Genomic rearrangements that occur within influenza A viruses allow for occasional cross-species transmission. These occur when two different viruses simultaneously infect a host, with subsequent genetic reassortment. Occasionally, cross-species transmission occurs without alteration of the viral genome. The names of influenza viruses are specified as follows: influenza genus (A, B or C)/host/geographic origin/strain number/year of isolation and, in parentheses, H and N type. For example, A/canine/Florida/43/2004 (H3N8). In the USA, canine influenza virus (CIV) emerged in racing greyhounds in Florida in 2003 and 2004, where it caused hemorrhagic pneumonia and a high mortality. Serological evidence of infection in the greyhound dog population dates back to 1999. Infections spread slowly and have subsequently been reported in racing greyhounds and non-greyhounds in at least 38 US states. Outbreaks have continued to occur in shelter situations for nearly a decade after the virus was discovered. The virus that has circulated in the USA is an H3N8 virus that resembles an equine influenza virus, which suggested that an interspecies jump occurred without genetic reassortment. Instead, accumulation of point mutations with minor amino acid changes occurred, followed by sustained transmission among dogs. The most significant outbreaks of disease due to CIV have occurred in Florida, New England, Colorado, Wyoming, and Texas. In other states, sustained transmission of the virus from one dog to another has not occurred. The most significant risk factor for infection has been indoor housing. Virtually all cases to date have involved dogs in kennels, animal shelters or dog day care facilities. Dogs of all ages and breeds are susceptible, but to date severe hemorrhagic pneumonia has only occurred in greyhounds. The virus



is shed for up to 7 to 10 days, but is typically shed for just a few days. In some dogs, shedding may have ceased when clinical signs are most apparent. CIV can still infect horses, but horses develop only mild disease or no clinical signs. Although infections with influenza viruses may be more likely to produce signs of fever and lethargy than dogs infected with other respiratory pathogens (e.g., Bordetella bronchiseptica, canine respiratory coronavirus, canine distemper virus, canine herpesvirus, canine adenovirus 2, canine parainfluenza virus), it is not possible to diagnose influenza virus infections in dogs based on clinical signs alone. The high prevalence of co-infections and increased severity of disease when multiple pathogens are present further complicates diagnosis. A history of exposure to other animals with respiratory disease can raise suspicion for the diagnosis. When outbreaks occur, attempts to make a diagnosis are indicated. Collection of multiple specimen types (oropharyngeal swabs, nasal swabs, and if possible transtracheal or bronchoalveolar lavage specimens) from several dogs with and without clinical signs can facilitate diagnosis and allow interpretation of the significance of positive test results. Organism detection methods, such as PCR, are likely to be of highest yield early in the course of illness (e.g., the first 1 to 3 days), or in exposed dogs that have not yet developed clinical signs. Using a combination of serology and organism detection methods (culture or PCR) may also facilitate diagnosis. Necropsies can provide valuable information, and should be performed as soon as possible after death or euthanasia occurs by a veterinary pathologist. Tissues should be submitted for histopathology (in formalin), bacterial and virus cultures (fresh tissue), and/or PCR for respiratory viruses and bacteria. Despite the increased availability of molecular diagnostic assays, virus isolation is still offered to veterinarians for routine diagnostic purposes by some veterinary diagnostic laboratories that specialize in virology (e.g., the Animal Health Diagnostic Laboratory at Cornell University in the USA). Panels of real-time PCR assays that detect respiratory pathogens may include assays for CIV. Unfortunately, false negative PCR results are common because of transient or low-level shedding of many respiratory viruses. In addition, because influenza viruses are RNA viruses, false negatives may result from degradation of viral RNA during specimen transport. Point-of-care assays are available for detection of nucleoprotein antigen to human influenza A viruses. Unfortunately, such assays have limited sensitivity and specificity for diagnosis of influenza virus infections in dogs. Serological assays for CIV exposure are based on serum neutralization or hemagglutination-inhibition. Serology is of limited use for diagnosis, because of vaccine titer interference in regions where vaccination is performed, and the high prevalence of subclinical exposure in regions where infection is endemic. Titers may be negative in the first 10 days of illness. Despite these limitations, serological assays have been key to identification of outbreaks of disease caused by CIV, when the disease is not endemic and widespread immunization has not yet been performed. Analysis of paired serum specimens collected 2 weeks apart can be used to document recent infection. In some dogs, no other



diagnostic test may be useful for antemortem diagnosis because virus shedding is so transient and difficult to detect. Assays for CIV that use equine influenza virus antigen for antibody detection have suboptimal sensitivity. Treatment of influenza virus infections is supportive. The efficacy and optimal dosage of neuraminidase inhibitors like oseltamivir is unknown, and because oseltamivir is a first line treatment for pandemic influenza in humans, it should not be used to treat dogs with respiratory disease, even when CIV infection is known to be present. In the United States, inactivated, parenteral vaccines are available for reduction of disease caused by H3N8 CIV and viral shedding. Their use has been recommended for dogs that may contact other dogs in regions where CIV is endemic. Vaccination against CIV is also required for importation of North American dogs to Australia. The initial vaccine may be given as early as 6 weeks of age. Because CIV vaccines are inactivated, 2 initial doses are required 3 to 4 weeks apart, and maximum immunity does not occur until 1 week after the second dose. As a result, CIV vaccines may not protect dogs that enter shelters with endemic canine influenza. There is currently no evidence of zoonotic transmission of CIV. However, a recent study revealed that a variety of human influenza viruses infect the canine trachea, and that reassortment of these viruses with CIV results in viable viruses. Thus dogs have the potential to be sources of novel viruses that could lead to influenza virus pandemics in humans. Avian-lineage H3N2 CIV emerged in South Korean dogs in 2007, and a similar virus was subsequently isolated from dogs in China. Experimental evidence exists that cats may also be susceptible to infection by this virus. A novel H3N1 virus was also detected in South Korean dogs that lacked clinical signs of respiratory disease. A novel H5N2 influenza virus was detected in a dog with respiratory disease in China, and was shown to be transmissible to other dogs, cats and chickens. Dogs are susceptible to infection with human influenza virus H1N1, avian H5N1, and avian H6N1, but sustained transmission of these viruses in the dog population has not been reported. Serologic evidence of exposure of feral dogs in China to avian influenza virus H10N8 has been reported. Limited infection of dogs with equine H3N8 viruses was detected in hounds in England and during an equine influenza outbreak in Australia. In England, disease was so severe that several hounds had to be euthanized, and subacute bronchointerstitial pneumonia was detected at necropsy. The Australian dogs developed inappetence, lethargy, nasal discharge, and a cough that persisted for several weeks, but dog-to-dog transmission was not identified. Experimental transmission of H3N8 influenza virus from horses to dogs was documented in Japan, but the infected dogs did not show clinical signs of illness. Canine influenza virus H3N2 was first detected in March in Illinois and Michigan and was probably imported from Asia. The largest numbers of cases, so far, have been detected in Illinois and Georgia, but cases have been detected in more than 20 states. Canine Respiratory Coronavirus Canine respiratory coronavirus is a newly discovered virus that represents another cause of respiratory disease in dogs worldwide. It has similarity to a cow coronavirus but is distinct from canine enteric



coronavirus (for which vaccines are available). Its presence tends to correlate with mild disease, but it has been detected in outbreaks of severe respiratory tract disease. Infection with canine respiratory coronavirus may predispose to other bacterial and viral infections, but may also potentially be a primary pathogen. It can be detected using PCR on transtracheal or bronchoalveolar lavage specimens, or throat swabs. Currently, no vaccines are available to prevent this infection. Canine Distemper Virus Canine distemper virus is another important cause of kennel cough, and it can also cause neurologic or gastrointestinal signs. However, many dogs with distemper lack neurologic or gastrointestinal signs. It is probably vastly undiagnosed as a cause of kennel cough in dogs. Canine distemper virus can be detected using PCR on respiratory specimens. It can also be detected using PCR on whole blood or conjunctival scrapings. Canine distemper virus vaccines are part of the core vaccine series. Three initial doses (6-8 weeks, 10-12 weeks, and 16-18 weeks) are required, after which an annual booster is indicated followed by boosters every 3 years. Recently, increasing numbers of distemper cases have been described in adult, previously vaccinated dogs, including in outbreak situations. The reason for this remains unclear, but careful handling and storage of vaccines that contain canine distemper virus is important to preserve their efficacy. Administration of the third puppy dose earlier than 16 weeks of age may also contribute to vaccination failure due to interference by maternal antibody. Canine Parainfluenza Canine parainfluenza virus remains the most important viral cause of CIRD in dogs, and intranasal and parenteral non-core vaccines are available and in widespread use for prevention of infection. Again, the relative efficacy of these types of vaccines is not well understood. Other viral pathogens include canine adenovirus (for which vaccination is available and used as a core vaccine for prevention of infectious canine hepatitis), and canine herpesvirus. Canine herpesvirus has recently been documented as a potential cause of ocular disease in dogs. SUMMARY In conclusion, an increasing number of pathogens have been recognized as causes of CIRD in dogs, and co-infections with multiple pathogens are commonly present. It is important not to overlook the possibility of coinfections, which may contribute to severe disease or result in a failure to respond as expected to therapy. Prevention is assisted by proper attention to hygiene and quarantine, minimizing overcrowding within kennels and shelters, and use of vaccines for CIRD. Because of growing concerns about antibiotic resistance, antibiotic treatment should be withheld unless dogs are systemically unwell and show signs of mucopurulent nasal discharge, lethargy, or have evidence of secondary bacterial pneumonia. Dogs with uncomplicated ‘kennel cough’ (i.e. those that are otherwise bright and alert but just coughing) typically recover without treatment over 1-4 weeks. Cough suppression may be indicated to help the dog (and the owner) sleep at night. If antibiotics are deemed indicated, doxycycline should be considered as a first-line treatment because of its activity against Bordetella and mycoplasmas.



RETROVIRUS RIDDLES


Introduction Feline immunodeficiency virus (FIV) and feline leukemia virus (FeLV) are retroviruses that belong to the genera Lentivirus and Gammaretrovirus, respectively. They remain important causes of mortality in cats through their ability to cause immunosuppression and neoplasia. Screening for retrovirus infections continues to be underperformed by veterinarians. Cats infected by both viruses may live long periods of time with good health, and so a positive test result should not alone be a reason for euthanasia. Feline Immunodeficiency Virus Infection Epidemiology FIV establishes a chronic, persistent infection that, in some cats, can culminate in immunodeficiency and/or tumor formation. There are at least 6 different subtypes of FIV, designated A through F. The existence of many strains of the virus complicates the design of molecular diagnostic tests and vaccines for FIV. FIV is shed in high concentrations in saliva, and the major mode of transmission is through bites. Studies worldwide have consistently shown that seropositivity (which is equivalent to infection) is associated with a history of bite wounds, older age, male sex, illness, and outdoor access. Indoor housing decreases transmission of FIV but does not eliminate it. Worldwide, the seroprevalence of FIV in domestic pet cats currently ranges from around 1 to 12%. Higher prevalences are found in feral and free-ranging cats and sick cats. Pathogenesis The main cellular target for FIV is the CD4+ T cell. However, FIV also infects CD8+ T cells, B cells, macrophages, and dendritic cells, microglia and astrocytes. Three phases of disease have been delineated, acute (primary), subclinical, and terminal. Knowledge of these phases can help practitioners understand diagnostic test results for FIV. After inoculation, the virus replicates in lymphoid tissues, and virus can be detected in the blood in high concentrations 2 weeks after infection. A peak of viremia occurs 8 to 12 weeks after infection. There is a decline in CD4+ and CD8+ T cells in peripheral blood, and this may be associated with transient illness, which lasts 3 to 6 months and is often unrecognized by cat owners. Some cats may show signs of lethargy, fever, anorexia, diarrhea, stomatitis, weight loss and lymphadenopathy during this phase. Most cats survive the acute phase of infection. In the subclinical phase, the numbers of CD4+ T cells rebound, and the virus load in the plasma declines to very low levels that can be difficult to detect even with PCR assays. Cats remain subclinically infected, often for years or even for life. There is a slow and progressive decline in the number of CD4+ T cells, reduction in the CD4+:CD8+ T cell ratio, and in some cats, hyperglobulinemia, which results from B cell hyperactivation. The rate of progression of this subclinical phase depends on factors such as the virus strain, environmental factors such as co-infections with other agents that activate virus transcription, and host immunity.



Ultimately, in some cats, these changes lead to the terminal phase of disease, which is characterized by the appearance of clinical manifestations of opportunistic infections, neoplastic disease, myelosuppression, and neurologic disease. However, many infected cats never develop FIV-related clinical signs, even when CD4+ T cell counts are low, and instead die from other causes. Lymphomas are the most commonly reported FIV-associated tumor, but other tumors can also occur. Immune dysregulation and increased circulating immune complexes can also lead to immune-mediated disorders. Myelodysplasia may develop in some cats. Neurovirulent strains of FIV can cause progressive behavioral changes, tremors, sleep disturbances, anisocoria, delayed reflexes, abnormal cranial nerve function, urinary and fecal incontinence, and seizures. The extent to which chronic FIV infection contributes to disease of other organs, such as the heart muscle and the kidneys, is not well understood, because the prevalence of cardiomyopathy and interstitial nephritis in geriatric cats not infected with FIV is high. Diagnosis The initial assay of choice for diagnosis of, and screening for, FIV infection is an ELISA assay that detects antibody to FIV. Provided there has not been a history of vaccination for FIV and the tested cat is less than 6 months of age, positive antibody test results equal infection, because the virus establishes a life-long, persistent infection. Point-of-care, lateral flow ELISA assays and diagnostic laboratory-based ELISA assays have rapid turnaround times and high sensitivities and specificities in cats. Positive test results in the absence of antibody to FIV may occur as a result of operator error or non-specific reactivity against tissue culture components after vaccination. Confirmation of positive screening test results has been recommended because of the low prevalence of infection in healthy cats and the higher possibility that false positive test results may occur. It is especially important to confirm positive test results if they are likely to result in euthanasia or rehoming for disease control purposes. Confirmation can be done using a test from a different manufacturer or Western blotting. False positive test results for infection can occur in cats vaccinated for FIV or kittens less than 6 months of age that possess maternal antibody, the latter either as a result of infection or vaccination of the queen. Kittens that test positive should be retested after 4 to 6 months of age. Cats with a history of vaccination with the inactivated FIV vaccine develop false positive test results that can persist for more than 4 years. Infection can occur in the face of vaccination, so positive test results in a vaccinated cat may represent infection or historical vaccination. Currently, molecular testing with PCR assays is required to identify infection in these cats, but some infected cats may test PCR-negative because of the low levels of virus present. Cats that are seropositive should probably be excluded as blood donors regardless of their vaccination history. False negative serological test results can occur early in the course of illness, because some cats take up to 60 days to develop an antibody response to the virus. Thus when recent exposure is possible, a second serological test should be performed a minimum of 2 months after the initial test. False negative test results can also occur in cats in the terminal phase of disease, as a result of impaired antibody production, or in kittens with rapidly progressive infections. These cats often have high plasma viral loads. Thus, if FIV infection is suspected on the basis of clinical signs, negative antibody test results



should be followed by virus detection using molecular methods (see below). However, negative serology when serology is used as a screening test is considered to be highly reliable because of the high sensitivity of the test and the low prevalence of infection in most populations of healthy cats. IFA assays for antibodies to FIV have also been described, which have had sensitivities and specificities between 95 and 100% when compared with Western immunoblotting. False positive results may occur if inexperienced laboratory personnel interpret non-specific fluorescence as a positive result. Molecular Diagnosis Using the Polymerase Chain Reaction. A variety of PCR assays have been developed for diagnosis of FIV infection. Assays may detect viral RNA (RT-PCR), proviral DNA, or both RNA and proviral DNA. Because the vaccine is inactivated, PCR should not detect vaccine virus in cats with a history of vaccination. Compared with serology, PCR can be insensitive (sensitivity < 80%), because viral loads in healthy cats are often low, and some strains may not be detected because of variability in the sequence of the viral genome among FIV isolates. Sensitivity may be higher in cats in the acute and terminal phase of disease when viral loads are expected to be higher. False positive test results have the potential to occur as a result of contamination, and in some commercial laboratories, unacceptable sensitivities and specificities have been reported. Regardless of the assay used, given the limitations of its interpretation, PCR should never be performed in the absence of concurrent serological testing. PCR-positive, seronegative cats may reflect a false positive PCR result or the terminal phase of disease with failure to produce antibody. Prognosis A limited number of studies have shown no significant difference in lifespan between cats infected with FIV and uninfected cats. In 1 study, the median survival times of these 2 groups after testing for FIV infection were 3.9 and 5.9 years, respectively. The progression and severity of disease is related to virus strain and host immunity. Because the lifespan of FIV-infected cats may not be considerably different from that of uninfected cats, no cat should be euthanized on the basis of a positive FIV test alone. Once terminal FIV-related disease occurs, lifespans are typically less than 1 year. Feline Leukemia Virus Infection Feline leukemia virus (FeLV) belongs to the genus Gammaretrovirus of the family Retroviridae. Despite a reduction in the prevalence of infection in recent years, FeLV remains an important cause of mortality in cats as a result of its ability to cause immune suppression, bone marrow disorders and hematopoietic neoplasia. FeLV infection progresses more rapidly than FIV infection and is more pathogenic, so virtually all cats that have progressive, productive infections ultimately die of FeLV-related disease. However, in contrast to FIV infection, many cats infected with FeLV regress to a permanent state of viral latency. Thus, a positive test result for FeLV in an apparently healthy cat does not always imply that FeLV-related disease and mortality will occur.



There are 4 different subtypes of FeLV: FeLV-A, FeLV-B, FeLV-C, and FeLV-T. Each subtype uses a different receptor to enter cells. All cats infected with FeLV-B, FeLV-C, and FeLV-T are co-infected with FeLV-A, and only FeLV-A is transmitted between animals. The other subtypes, which are more pathogenic, arise from FeLV-A. The FeLV subtype influences the clinical expression of disease. For example, FeLV-T, a T-cell tropic variant, is associated with immunodeficiency in cats, whereas FeLV-C is associated with non-regenerative anemia. Transmission of FeLV-A primarily occurs as a result of prolonged, close contact with salivary secretions, such as through licking, mutual grooming, and shared food and water dishes. The prevalence of infection has declined over the last 2 decades with more extensive testing and immunization for the infection. Currently, the overall prevalence of infection in mixed populations of cats is approximately 1 to 6%. The median age of cats infected with FeLV is 3 years. This reflects the phenomenon of age-related resistance to FeLV. Pathogenesis The outcome of FeLV infection depends strongly on the virus strain involved and factors that influence host immune function. The virus replicates in oral lymphoid tissue and then circulates in a few monocytes and lymphocytes within peripheral blood. Some cats may develop systemic signs, such as fever, lethargy and lymphadenopathy during this period. A small number of infected lymphocytes then travel to the bone marrow, where the virus infects rapidly dividing precursor cells and subsequently lymphoid and epithelial cells throughout the body. Infection of the bone marrow is a critical step in the pathogenesis of FeLV infection. Once infection of epithelial cells within the intestinal crypts and salivary glands occurs, the virus is shed in massive quantities in the saliva and feces; it can also be shed in urine. There are several possible outcomes of infection with FeLV. The immune system of some infected cats is able to suppress productive viral infection within a few weeks after infection, before significant infection of the marrow occurs. These cats develop a regressive infection whereby proviral DNA is present in the host cell genome but production and shedding of virus no longer occurs. Regressive infection usually occurs for life, but it may be reactivated with immunosuppression. Later in life, an unknown percentage of cats with regressive infections may develop FeLV-negative malignancies as a result of integration of viral DNA within host cellular oncogenes. Transfusion of blood from cats with regressive infections to naïve cats has the potential to be followed by reactivation of FeLV in the transfused cat. Cats develop progressive infection once involvement of the marrow is established and persistent viremia and progressive FeLV-related disease results. Progressive FeLV infection leads to opportunistic infections, neoplasia, anemia, immune-mediated disease, neurological disorders, enteritis, and reproductive disease. The most common types of neoplasia in cats infected with FeLV are lymphoma and leukemia. Anemia in cats infected with FeLV may occur as a result of multiple different mechanisms, including decreased RBC production and increased RBC destruction.



Diagnosis Infection with FeLV is often diagnosed during screening efforts. Screening should be performed with ELISA assays for FeLV antigen. The retrovirus status of all cats should be known regardless of the presence of absence of illness. Even though some cats that test positive for FeLV antigen have no clinical signs or physical examination abnormalities, a CBC, chemistry panel and urinalysis should be obtained from these cats (and at a minimum, a complete CBC with blood smear evaluation) to assess for underlying abnormalities that may signal the presence of FeLV-related disorders. The initial assay of choice for diagnosis of FeLV infection is an ELISA that detects soluble p27 capsid protein antigen in blood. When virus isolation in culture was used as the gold standard, the sensitivity of 7 different assays ranged from 92.1 to 96.8%, and the specificity ranged from 95.4 to 99.2%. ELISA. When ELISA assays are used as screening tests, confirmation of positive test results is recommended because of the low prevalence of infection in healthy cats and the higher possibility that false positive test results may occur. Positive test results in the absence of FeLV antigen occur rarely as a result of operator error or non-specific reactivity. There are several options to confirm a positive test result:

• Perform another ELISA antigen test using an assay from a different manufacturer. However, it should be remembered that in contrast to FIV infection, cats that test truly positive for FeLV antigenemia early in the course of infection may ultimately still control the infection.

• Perform an IFA assay on peripheral blood smears, because cats with positive IFA results have infection of the bone marrow and are almost always progressively infected.

• Retest with ELISA 1 month later. If the antigen test remains positive, progressive infection is likely. Because in some cats, antigenemia may persist for 4 months before regression occurs, the test should be repeated 3 months later or monthly if client finances permit so long as the cat remains healthy.

• Perform a full CBC. If hematological abnormalities exist, progressive infection is likely. False negative ELISA assay results can occur in the first month after exposure, before sufficient virus can be detected in the peripheral blood. Cats that test negative within 30 days of possible exposure to the virus should be retested 1 to 2 months later. IFA. IFA assays can be performed on fresh peripheral blood smears or bone marrow. IFA is less sensitive than ELISA and, depending on the laboratory, is more prone to false negative and positive results and so is not recommended for screening purposes. False negative test results can occur in cats with progressive infection when there are inadequate blood cells in the periphery, such as in neutropenic cats. Performance of IFA on bone marrow rather than peripheral blood may help to overcome this problem.



Polymerase Chain Reaction. Several different PCR assays have been developed for detection of FeLV. Currently the major clinical indications for PCR are 1) to screen potential blood donors in association with antigen testing, 2) to test for regressive infection when FeLV is strongly suspected as the cause of disease but antigen tests are negative. PCR assays may detect viral RNA (RT-PCR), proviral DNA, or both RNA and proviral DNA. At the current time, PCR should never be used in the absence of antigen testing in order to screen for or diagnose FeLV infection. In addition, it is important that the clinician understand if the laboratory assay used detects proviral DNA, viral RNA (RT-PCR), or both, because the clinical significance of a positive viral RNA assay may differ from that of a positive proviral DNA assay. The clinical outcome for cats with a positive proviral PCR test result but negative soluble antigen test, which in 1 study represented about 10% of cats with negative antigen test results, is currently unclear. The sensitivity and specificity of commercially available PCR assays is likely to vary depending on assay design, and assays offered commercially have not been well validated or their use has not been well published, so caution is always warranted when interpreting the results of PCR assays for FeLV infection. FeLV Vaccines Several vaccines are available for prevention of FeLV infection. No vaccine provides 100% protection against FeLV infection, and even when protection against progressive infection occurs, regressive infections still occur after challenge. However, vaccination can protect cats from progressive FeLV infection, and so it is indicated for all cats that are at risk of infection. Two doses are given 3 to 4 weeks apart from 8 to 9 weeks of age, followed by a booster at 1 year and then every 1 to 3 years thereafter. Testing for FeLV should be performed before each booster if exposure to FeLV was likely before booster immunization was required (which would be true for most cats vaccinated for FeLV). Vaccination does not interfere with diagnostic test results for FeLV infection. References and Suggested Readings 1. Goldkamp CE, Levy JK, Edinboro CH, et al. Seroprevalences of feline leukemia virus and feline

immunodeficiency virus in cats with abscesses or bite wounds and rate of veterinarian compliance with current guidelines for retrovirus testing. J Am Vet Med Assoc 2008;232:1152-1158.

2. Levy J, Crawford C, Hartmann K, et al. 2008 American Association of Feline Practitioners' feline retrovirus management guidelines. J Feline Med Surg 2008;10:300-316.

3. Hosie MJ, Addie D, Belak S, et al. Feline immunodeficiency. ABCD guidelines on prevention and management. J Feline Med Surg 2009;11:575-584.

4. Hartmann K. Clinical aspects of feline immunodeficiency and feline leukemia virus infection. Vet Immunol Immunopathol 2011;143(3-4):190-201 (for a review of molecular pathogenetic mechanisms of disease).

5. Lutz H, Addie D, Belak S, et al. Feline leukaemia. ABCD guidelines on prevention and management. J Feline Med Surg 2009;11:565-574.



VECTOR-BORNE DISEASES: LYME, ANAPLASMOSIS, AND EHRLICHIOSIS


Vector-borne infectious diseases have been identified more frequently in humans and domestic animals in recent years and include some of the most important emerging and re-emerging infectious diseases. Infectious diseases that may be transmitted to dogs and cats via a vector include viral, bacterial, and protozoal diseases. Examples include Lyme borreliosis, anaplasmosis, ehrlichiosis (especially caused by Ehrlichia canis and Ehrlichia ewingii), Rocky Mountain spotted fever, babesiosis, hepatozoonosis, plague, tularemia, leishmaniasis, bartonellosis, and possibly hemoplasmosis. The vectors most commonly are arthropods, such as fleas, ticks, sandflies (leishmaniasis), other biting flies, and mosquitoes (eg, West Nile virus). The epidemiology of vector-borne infections depends on the distribution of the arthropod vector; travel history is an important consideration when examining dogs with possible vector-borne infectious diseases. The development of molecular diagnostic techniques has contributed to improved recognition of a number of zoonotic vector-borne infectious agents, such as Bartonella and Borrelia. Improved surveillance for these diseases has also improved recognition. Human and animal travel is increasing worldwide and has contributed to emergence of many infectious diseases. Habitat changes, deforestation, and urbanization have contributed to the emergence of a number of important vector-borne diseases in humans. Climate change may expand the range for some vectors, which places more hosts at risk. However, increased temperatures may result in more people and their pets staying indoors in an air-conditioned environment, which would tend to decrease the risk for vector-borne infectious disease. Humidity, rainfall, wind, and the duration of daylight are also important considerations. Co-infection with multiple zoonotic vector-borne infectious agents is a relatively common phenomenon that is being increasingly recognized. Co-infections can complicate the clinical picture, and in some situations the presence of one organism may affect the pathogenicity of another. Documentation of exposure to one vector-borne infectious agent should be a cue to search for exposure to other agents transmitted by similar vectors and should raise suspicion for the possibility of other potentially zoonotic agents. Dogs can also serve as sentinels for human exposure to vector-borne infectious diseases, but in the majority of instances (with the exception of Bartonella, plague, and tularemia), dogs and cats do not transmit these infections directly to people. However, many of these diseases have the potential to be transmitted to humans through exchange of blood between infected animals. This could occur as a result of aggressive interactions between animals, blood transfusion, common hospital accidents such as needle stick injuries or injuries that occur during necropsy of infected animals, or possibly even contact between an infected animal’s blood and abraded skin. Organisms that cause chronic, persistent infections such as Bartonella species are of special concern because their presence is less likely to be suspected. Absence of a history of vector exposure also does not rule out the presence of a vector-



borne infectious disease. Increased awareness of the zoonotic potential of the conditions listed below and precautions to prevent accidental exposure to blood and body fluids should reduce the risk that veterinarians may become exposed to these agents. Lyme Disease Borrelia burgdorferi is the cause of Lyme disease, the most common vector-borne disease in humans. Ecologic changes—primarily farmland reforestation and residential development in wooded areas, an associated explosion in deer populations, and close proximity of reservoirs, ticks, humans, and dogs—have all likely contributed to increasing recognition of Lyme disease. The tick vectors in the United States are Ixodes scapularis (upper midwest and northeast) and Ixodes pacificus (west). The primary mammalian reservoir in the upper midwest and northeast regions is Peromyscus leucopus, the white-footed mouse. Western gray squirrels may be an important reservoir in California. Following inoculation of spirochetes into the host by the tick, the organisms migrate through connective tissue from the bite site, causing fever, inappetence, thrombocytopenia, and lameness due to neutrophilic polyarthritis in some dogs approximately 2 to 5 months after infection. As a result, seronegative Lyme disease is rare in dogs. A small percentage of dogs may develop glomerulonephritis (so-called Lyme nephritis), which may be accompanied by thrombocytopenia and in some cases, polyarthritis. The vast majority of dogs (> 90%) show no signs of illness. In humans, B. burgdorferi causes erythema migrans, arthritis, and neurologic signs (neuroborreliosis). Dogs have been used as a sentinel for human exposure. A specific diagnosis is generally obtained through serology for detection of antibodies. Diagnosis is often difficult, however, because of the widespread subclinical exposure in endemic areas. In dogs, diagnosis can involve detection of antibodies to the C6 peptide, which can be performed using the in-house SNAP 4Dx Plus kit (IDEXX Laboratories, Westbrook, ME) or the IDEXX quantitative C6 ELISA. The C6 peptide represents a portion of the Borrelia VlsE surface protein, which is only expressed during natural infection. Therefore assays that detect antibodies to this protein are able to differentiate between the immune response to vaccination and the immune response to natural infection. Other assays detect antibodies against multiple Borrelia surface proteins, and to some extent, the pattern of antibody reactivity to each surface protein can provide information on whether infection is acute or chronic, and whether the immune response is to vaccine antigens or natural infection. These assays include the Antech Accuplex 4 assay (which detects antibodies to OspA, OspC, OspF, P39 and SLP), the Cornell fluorescent bead-based assay (OspA, OspC and OspF), and Abaxis VetScan Lyme (VlsE, OspC and p41). OspA is an outer surface protein that facilitates adherence of the spirochete to the tick midgut. Expression of OspA is shut down by the spirochete when it enters a dog at the time of natural infection, so antibodies to OspA usually represent a response to vaccination. When ticks ingest a blood meal from a vaccinated dog, antibodies to OspA lyse the spirochete before it enters the dog, so vaccines that contain OspA primarily act in the tick rather than in the dog. OspC is expressed by the spirochete as it enters the tick’s salivary glands and then early in the course of infection. Therefore, antibodies to OspC appear shortly after infection, whereas antibodies to OspF appear several weeks after infection (late antibody response). We still have much to learn about how clinically useful these patterns of reactivity



are. Polymerase chain reaction (PCR) assays are available as part of vector-borne disease PCR panels but have limited sensitivity when performed on blood because the organism resides primarily in connective tissues. Apparently healthy, seropositive dogs should probably be evaluated for proteinuria and co-infections with other tick-borne pathogens, and tick control should be recommended (see below). Generally, treatment of sick dogs involves administration of doxycycline (5 mg/kg PO q12h) for 4 weeks, but complete elimination of the spirochete may not occur. The author does not advocate serially monitoring quantitative C6 antibody titers because there is no evidence that this correlates with clinical recovery or elimination of infection, and no evidence that persistent positive antibody titers lead to relapse of disease or Lyme nephritis. In addition, subclinical reinfections may be possible with other strains of Borrelia, as has been shown to occur in humans. Dogs with Lyme nephritis are being treated with some success using immunosuppressive drugs. Vaccination for Lyme disease is controversial because polyarthritis is readily treated with doxycycline and evidence that vaccination prevents Lyme nephritis is lacking. In addition, some internal medicine specialists fear that vaccination theoretically has the potential to predispose dogs to Lyme nephritis should they become infected after vaccination (or be vaccinated after natural infection that persists subclinically). Recombinant OspA Lyme vaccines were efficacious in humans, but concerns that they contributed to immune-mediated arthritis led to their withdrawal from the market, despite the relative lack of evidence for this. Currently there is also no evidence that vaccination contributes to chronic immune-mediated consequences of infection in dogs. Because humans can experience recurrent erythema migrans lesions as a result of reinfection, there does seem to be some basis for vaccination of seropositive dogs to prevent infection by new strains of Borrelia (remembering that the vaccine works in the tick to prevent infection). Current vaccines are either inactivated whole spirochete vaccines that express OspA and OspC (Zoetis, Merck) or recombinant OspA vaccines (Merial). There are many variants of OspC and antibodies to one OspC variant may protect against others. Therefore, the value of inclusion of OspC in vaccines has been debated. Granulocytic Anaplasmosis Anaplasma phagocytophilum is the cause of granulocytic anaplasmosis. This disease was first recognized in humans and dogs in the upper Midwest in the early 1990s. The emergence of this disease has been postulated to be due to repopulation of habitats previously devoid of ticks by deer, small rodents, and humans, together with improved diagnostic capabilities. The vectors appear to be the same as those for B. burgdorferi. A variety of species may be infected, including horses, ruminants, dogs, cats, and wildlife. The largest number of cases has been seen in the upper midwest and northeastern United States, and the disease is increasingly reported from other areas, including northern California, southwestern Oregon, the northwest coast, and Vancouver Island in British Columbia. Clinical signs of granulocytic anaplasmosis include fever, lethargy, inappetence, peripheral lymphadenopathy, and lameness due to neutrophilic polyarthritis. Neurologic signs have been reported anecdotally but are rare, and include neck pain, placing deficits, and even seizures. Laboratory abnormalities may include leukopenia, thrombocytopenia, and elevated alkaline phosphatase (ALP) activity and sometimes alanine aminotransferase (ALT) activity.



Morulae may be seen occasionally on blood smears within granulocytes, most commonly neutrophils. Diagnosis is best made by visualization of morulae (in endemic areas where they can be distinguished from morulae of Ehrlichia ewingii) or using PCR. Diagnosis can also be based on acute- and convalescent-phase serology using an indirect immunofluorescent antibody (IFA) assay. Serology is not the preferred method for diagnosis of this disease though, because acute titers are often low or negative, so diagnosis is retrospective using this method. Serologic panels such as the SNAP 4Dx Plus test kit and the Accuplex assay detect antibodies to Anaplasma species. A positive result should prompt the clinician to perform IFA serology so that acute and convalescent titers may be determined. Positive ‘blue dots’ for Anaplasma usually suggest exposure to an Anaplasma species in the past, and should prompt the clinician to discuss the need for tick prevention, rather than being considered a sign of active Anaplasma infection (because many dogs are subclinically exposed). Treatment is with doxycycline for 2 weeks, which usually results in resolution of clinical signs within 24-48 hours (and clinical improvement within hours). Whether persistence of this organism occurs in dogs is controversial; there is evidence for persistence of European strains in horses and ruminants. Ehrlichiosis Ehrlichioses of dogs are a group of tick-transmitted diseases caused by intracellular, gram-negative, bacteria that include Ehrlichia canis, Ehrlichia ewingii, Ehrlichia chaffeensis, and the recently discovered organism Ehrlichia muris. An organism related to Ehrlichia ruminantium, the cause of heartwater disease in cattle, has also been detected in ill dogs from South Africa. Ehrlichia canis infects monocytes and causes canine monocytic ehrlichiosis (CME), one of the most important tick-borne diseases of domestic dogs worldwide. Ehrlichia ewingii is an unculturable bacterium that infects granulocytes and causes canine granulocytic ehrlichiosis in the mid-western and southeastern United States. Ehrlichia chaffeensis causes human monocytic ehrlichiosis; dogs are a proposed reservoir for this organism. Ehrlichia muris was recently discovered as a cause of febrile illness in dogs from the upper Midwest of the United States. The geographic distribution of each pathogen is generally restricted to that of their vectors and mammalian reservoir hosts. E. canis is transmitted primarily by Rhipicephalus sanguineus. Different strains of E. canis exist worldwide that may vary in virulence. Because of chronic, subclinical infection, dogs can be transported to non-endemic regions and subsequently develop disease years later. Ticks acquire infection by feeding as larvae or nymphs on infected dogs. Wild canids may serve as reservoir hosts. No age or sex predilection for CME has been documented, but there is some evidence that purebred dogs may be more susceptible to disease. The course of CME is divided into acute, subclinical and chronic phases, although these phases may not be clearly distinguishable. Clinical signs of acute CME occur 8 to 20 days after infection. Immune-mediated mechanisms appear to be important in the pathogenesis of disease. Lethargy, inappetence, fever and weight loss are the most common signs. Replication of the organism in reticuloendothelial tissues is associated with generalized lymphadenopathy and splenomegaly. Ocular and nasal discharges, peripheral edema and less commonly, mucosal and cutaneous hemorrhages may also occur. Neurologic



signs may occur. Dogs can recover from the acute phase within 2-4 weeks without treatment, after which they may eliminate the infection or remain subclinically infected. Subclinical infection may persist for months to years. Chronic CME is typified by the finding of pancytopenia, which results from hypoplasia of all bone marrow cells. Clinical signs include lethargy, inappetence, bleeding tendencies, mucosal pallor, fever, weight loss, lymphadenopathy, splenomegaly, dyspnea, anterior uveitis, retinal hemorrhage and detachment, polyuria/polydipsia, and edema. Polymyositis occurs in some dogs, which can be manifested by diffuse muscle wasting and tetraparesis. Secondary opportunistic infections such as viral papillomatosis, protozoal infections, and bacterial urinary tract infections can also develop, although the precise underlying mechanism of immunosuppression has not yet been elucidated. Marked granular lymphocytosis and bone marrow plasmacytosis may occur, sometimes accompanied by a monoclonal gammopathy, which may lead to misdiagnosis of lymphocytic leukemia or multiple myeloma, respectively. Protein-losing nephropathy may develop as a result of immune-complex glomerulonephritis. Diagnosis of CME can be made based on visualization of morulae within circulating monocytes, but this is insensitive, especially in dogs with chronic disease. In one study, after careful searching, morulae were found in only 2 of 19 dogs with CME. More often, the diagnosis is made using serology, which may be performed using IFA testing, ELISA technology, or Western blotting. Using IFA, antibodies can be detected between 7 and 28 days after initial infection. Dogs with acute disease may initially have negative test results if sufficient time has not elapsed for antibody production to occur. PCR may be helpful for diagnosis in this situation. If the initial serum antibody titer is positive, this may reflect previous exposure, and may not correlate with the presence of disease. Re-testing should be performed 2 to 3 weeks later to demonstrate seroconversion. Dogs with chronic E. canis infection frequently have extremely high IFA titers, sometimes > 1:600,000. Seroconversion does not generally occur in dogs with chronic disease, although antibody titers may decline in some dogs with treatment. Serologic cross-reactivity to other Ehrlichia species occurs. Cross-reactivity to Anaplasma antigens can occur to a lesser extent. The sensitivity of PCR for diagnosis of CME when performed on bone marrow in dogs with chronic ehrlichiosis appears to be less than 70%. A variety of ELISA assays have been developed for detection of antibodies to E. canis. If affordable to the client, the incidental finding of Ehrlichia seroreactivity in dogs screened using ELISA should prompt performance of a thorough physical examination and basic laboratory testing including a CBC, chemistry panel and urinalysis to evaluate for thrombocytopenia, hyperglobulinemia, and proteinuria. When sick dogs test positive, serologic testing with IFA should be performed so that a titer can be obtained; acute and convalescent testing may be needed. The treatment of choice for CME is doxycycline (10 mg/kg PO q24h) for a minimum of 28 days. Most dogs with acute disease show clinical improvement within 24 to 48 hours. Dogs with severe chronic disease may not respond to therapy, or cytopenias may gradually resolve over a period of several months. Treatment of seroreactive, but otherwise apparently healthy dogs is controversial, because treatment has not been shown to change the outcome for these dogs and has the potential to lead to antimicrobial resistance or adverse effects of drug therapy.



E. canis DNA has been detected in some human patients with clinical signs of human monocytic ehrlichiosis, suggesting that E. canis may be a cause of monocytic ehrlichiosis in people. Thus, precautions should be taken when handling engorged ticks as well as blood and tissue specimens from infected dogs.

Bartonellosis Bartonella species include Bartonella henselae and Bartonella vinsonii subspecies berkhoffii. These are curved, gram-negative bacteria that reside within erythrocytes and endothelial cells and are transmitted by a variety of arthropod vectors, including fleas and probably to some extent, ticks, lice, and biting flies. Infection of cats by B. henselae in endemic areas may be as high as 30% and is most often subclinical. Endocarditis has been reported rarely in cats in association with Bartonella infection, and the organism is suspected to play a role in uveitis, lymphadenomegaly, gingivostomatitis, and possibly lower urinary tract disease in some cats, although the role of Bartonella in these syndromes has been controversial because of the high prevalence of infection in apparently healthy cats. In dogs, infection with Bartonella is of lower seroprevalence (generally <5%) and has been associated most commonly with endocarditis. The prognosis for Bartonella endocarditis is poor when compared with that due to other bacterial pathogens. Bartonella causes cat-scratch disease in immunocompetent humans and a variety of serious disease manifestations in the immunocompromised, such as endocarditis, peliosis hepatis (multiple vascular proliferations throughout the liver), and bacillary angiomatosis (vascular proliferations in the skin). Diagnosis is generally made using serology, but veterinarians must be aware that some animals with active infection are seronegative, and positive serology does not imply active infection. Blood culture (using special media for Bartonella, usually in a specialized diagnostic laboratory) and PCR of blood is useful in cats for detection of bacteremia but has low sensitivity in dogs, which tend to have lower organism loads. More sensitive culture-PCR methods have been developed at North Carolina State University and are available commercially for diagnosis of Bartonella infections in a variety of animal species (readers are referred to the Galaxy Diagnostics website for more information). Because both dogs and especially cats may have subclinical bacteremia, a positive PCR or culture result does not imply that Bartonella is the cause of clinical signs. Bartonella infection may be difficult to eliminate using antimicrobial therapy. Doxycycline and azithromycin have been the most common antibiotics used. Resistance to azithromycin has been documented. Concerns about treating cats that lack clinical signs suggestive of bartonellosis have been raised, even if they live with immunocompromised people. This is because humans may become scratched during administration of medication, and no medication effectively eliminates infection. PREVENTION OF VECTOR-BORNE DISEASE TRANSMISSION Prevention of vector-borne infectious diseases in dogs and cats in the home environment involves a combination of the following:

• Reduction of exposure to vectors through avoidance of infested environments and housing cats indoors



• Removal of habitats that harbor arthropod vectors in the environment around homes (such as brush, woodpiles, and standing water)

• A search for and removal of ticks within hours after exposure of dogs to potentially infested environments

• Application of topical ectoparasiticides with activity against ticks when possible • Use of appropriate vaccines where available

Prompt removal of ticks prevents transmission of vector-borne pathogens because there is a delay of at least 12 hours before pathogens are transmitted by the tick to the mammalian host. In dogs, use of ectoparasiticides has been shown to effectively prevent transmission of important tick-borne infectious diseases. The use of products with activity against fleas effectively prevents transmission of B. henselae. Traditionally, use of ectoparasiticides with activity against ticks in cats has been difficult because of cats' susceptibility to the adverse effects of permethrin-based products. However, a flumethrin-containing ectoparasiticide (Seresto, Bayer) is approved for tick and flea prevention in dogs and cats in Europe and the United States, which provides a solution to this problem. Unlike permethrin and deltamethrin, metabolism of flumethrin is not dependent on hepatic glucuronidation, so cats are less susceptible to toxic adverse effects of flumethrin. Other new flea and tick preventatives on the market for dogs only include the isoxazalines, afoxolaner (Nexgard, Merial) and fluralaner (Bravecto, Merck). These are oral chewables that have activity for one month or three months, respectively. Clients should be educated about the reasons for application of ectoparasiticides, including prevention of zoonotic vector-borne diseases. They should also be told that while ectoparasiticides reduce the risk for vector-borne infections, they do not completely prevent these infections, especially in heavily infested environments. In the hospital environment, care should be taken to avoid direct contact with vectors found on pets during physical examination. Ticks should be removed with curved forceps or other tick-removal devices and disposed of in alcohol. Infested animals should be treated immediately with ectoparasiticides. Blood donors should be screened for vector-borne diseases using appropriate combinations of serologic and PCR assays. In most cases, use of both serology and PCR assays is recommended. When handling potentially infected animals (which is effectively all veterinary patients because of the potential for subclinical infections), care should be taken to avoid sharps injuries and exposure to blood and even saliva. Handwashing should be practiced after handling every patient, and gloves should be worn for all procedures that involve non-intact skin. In order to avoid needle-stick injuries, sedation should be considered for procedures that involve venipuncture or fine-needle aspiration, especially if the patient is skittish or uncooperative or a zoonotic infection is suspected. Because many infections can be transmitted through bites and scratches, staff should be educated on bite and scratch avoidance, such as the use of suitable restraint devices and protective gloves and warning signage on cages and medical records. Should bites or scratches occur, they should be vigorously flushed and washed immediately with soap and water or 0.05% chlorhexidine solution, and the bite reported to appropriate officials. If a bite occurs, medical attention should be sought as soon as



possible. All bites or scratches must be documented and consideration given to why the injury occurred, so that procedures or training to prevent future injuries can be implemented if necessary.



LEPTOSPIROSIS: DIAGNOSIS, TREATMENT AND PREVENTION


Introduction Leptospirosis is caused by infection with various serovars of Leptospira interrogans sensu lato. Organisms are transmitted by direct contact with infected urine, bite wounds or ingestion of infected tissues, or indirectly, through contact with infected water, soil, food or bedding. Survival of leptospires is promoted by stagnant warm water, a neutral or slightly alkaline pH, and temperatures between 0 and 25°C. The seasonality of the disease is variable depending on local climactic conditions, especially rainfall. In areas with year-round rainfall, the disease may occur throughout the year. There are over 200 pathogenic serovars, which are grouped into antigenically-related serogroups. Serovars known to infect and cause disease in dogs include Canicola, Icterohaemorrhagiae, Grippotyphosa, Pomona, Ballum, Bratislava, Autumnalis, Bataviae, Australis, and Hardjo. In addition, classification of leptospires is gradually moving from predominantly serovar-based classification to that based on genetic typing (genotype-based classification). Each serovar (and more accurately, each genotype) is adapted to a one or more mammalian host species (maintenance hosts). Other hosts act as incidental hosts. Disease in incidental hosts tends to be more severe and the duration of shedding is generally shorter. Maintenance hosts include dogs (Canicola); rats (Icterohaemorrhagiae); small wildlife mammalian species such as voles and raccoons (Grippotyphosa); cattle and pigs (Pomona); pigs (Bratislava); cattle (Hardjo); and mice (Ballum). The prevalence of infection with a serovar/genotype in dogs depends on the degree of contact between the dog population and the maintenance host for that serovar/genotype. The most common serovars thought to infect dogs before the introduction of the Leptospira vaccines were Icterohaemorrhagiae and Canicola. Vaccines containing only serovars Icterohaemorrhagiae and Canicola do not protect against infection by other serovars. Since introduction of the bivalent bacterins containing these two serovars, in North America and Europe, there have been decreasing reports of disease associated with seroconversion to Canicola and Icterohaemorrhagiae, and increasing reports of disease associated with seroconversion to serovars Pomona, Grippotyphosa, Autumnalis and Bratislava (in North America) and Sejroe, Australis and Grippotyphosa (in Europe). Vaccine pressure, increasing contact between dogs and certain wildlife reservoir hosts and increased testing have been suggested as reasons for this change. In truth, the actual serovars causing disease in dogs worldwide remain uncharacterized because the disease is diagnosed by serology, and serologic test results are not predictive of the infecting serovar. Pathogenic leptospires penetrate abraded skin or mucus membranes and multiply rapidly in the bloodstream and tissues, causing renal failure, hepatic injury (usually not hepatic failure) and vasculitis. The disease is multisystemic and may also involve the pancreas (pancreatitis), gastrointestinal tract (gastroenteritis), eye (uveitis) and lungs (leptospiral pulmonary hemorrhage syndrome, or LPHS). In



humans, Leptospira can also cause meningitis, which is most commonly manifest as a severe headache. Clinical manifestations may also depend on the age of the host, the infectious dose, and the strain of Leptospira involved. Clinical Manifestations Most infections are subclinical. Younger, large breed, outdoor adult dogs are commonly affected, but the disease can occur in any dog breed and at any age; dogs that live in cities may become infected as a result of exposure to rodent reservoir hosts. One recent study showed an increase in the percentage of small breed dogs diagnosed with leptospirosis between 1970 and 2009 (Lee et al, 2014). Younger animals tend to be more severely affected. Male dogs may be predisposed. Lethargy, anorexia, vomiting, pyrexia, dehydration, abdominal pain and increased thirst and urination are common signs of acute leptospirosis. Reluctance to move due to myositis, icterus, and uveitis may be noted. Respiratory difficulty may result from pulmonary hemorrhage, which is often associated with the development of moderate anemia. Laboratory Findings Leukocytosis, thrombocytopenia, azotemia, hypoalbuminemia and mild to moderately elevated liver enzyme activities are common. Urinalysis may reveal isosthenuria, proteinuria, glucosuria and casts. Although it occurs with other causes of renal tubular damage, glucosuria in addition to azotemia can be a “red flag” for a diagnosis of leptospirosis. Proteinuria is typically low-level (urine protein:creatinine ratio < 5), which helps differentiate leptospirosis (interstitial nephritis) from Lyme nephritis (which involves the glomerulus). Thoracic radiography may reveal a focal or diffuse interstitial to bronchointerstitial pattern; alveolar patterns may represent pulmonary hemorrhage. Occasionally mild pleural effusion is evident. Hepatomegaly, splenomegaly, renomegaly and/or peritoneal effusion may be evident from abdominal radiography. Hyperechoic renal cortices and mild renal pelvis dilation are occasionally seen with abdominal ultrasound. Diagnosis Canine leptospirosis is an underdiagnosed disease in some regions/practices and an overdiagnosed disease in other regions/practices. Identification of leptospirosis requires a high clinical suspicion for the disease based on knowledge of the range of clinical presentations that suggest leptospirosis. This is because currently accurate diagnosis is retrospective and generally based on serology using the microscopic agglutination test (MAT). Respective titers are provided for each of several different serovars in order to increase the chance of antibody detection. Studies in humans and dogs have shown that the serovar with the highest titer can vary over time and that paradoxical cross-reactivity to multiple serovars occurs after exposure to a single serovar. Thus, the MAT does not accurately predict the infecting serovar, and therefore should not be used for this purpose. Titers may be negative in the first week of illness because of the short incubation period and delay in antibody production. Low positive or negative titers after at least one week of illness suggest leptospirosis is not present.



Over-diagnosis results from misinterpretation of positive test results. Positive titers early in the course of an illness may reflect residual post-vaccinal titers or prior subclinical infection, and are not diagnostic for the disease. Demonstration of a fourfold rise in titer is required over a 1-2 week interval. In acutely ill dogs (< 1 week of illness), it is the author’s opinion that leptospirosis serology should only be performed in a paired fashion or not at all, because of the limited utility of a single positive titer, regardless of its magnitude. Postvaccinal titers against Icterohaemorrhagiae, Canicola, Grippotyphosa and Pomona occasionally rise as high as 1:6400 for a few months after vaccination, and these can interfere with interpretation. The results can also vary dramatically between laboratories (Miller et al, 2011). Use of a laboratory with a high level of quality control is recommended, or a laboratory that participates in the International Leptospirosis Society’s proficiency testing scheme. More information on leptospirosis diagnosis at the author’s institution can be found on the UC Davis leptospirosis laboratory testing website (http://www.vetmed.ucdavis.edu/foley_lab/leptospira/index.cfm). Recently, an LipL32-based ELISA assay has become available in North America for detection of antibodies to pathogenic leptospires in dogs. In the future, rapid in-clinic kits for leptospirosis serology may also become available. These assays yield qualitative (positive or negative) results, and may be most useful for screening dogs for the presence or absence of antibodies. Should these kits yield negative results, then the clinician should consider whether it may be too early for the animal to have developed antibodies (as can occur with the MAT). Another test should be performed one week later to see if the animal seroconverts. Should these kits yield positive results, then the clinician should consider whether previous vaccination has occurred (assuming they cross-react with vaccine-induced antibody, as is the case for the LipL32-based ELISA). Previous exposure without clinical disease should also be considered as a reason for positive results. Thus, in the author’s opinion, a positive result using this kit should stimulate reflex testing with acute and convalescent quantitative serology using the MAT. Darkfield microscopy of the urine is not recommended as sole test for diagnosis because of the large number of false positives and false negatives. Silver staining and fluorescent antibody or immunoperoxidase staining of tissue specimens can also yield false negatives, and do not help identify the infecting serovar. Culture is difficult because of the fastidious growth requirements of leptospires and the need for specialized media, but is the only way to truly identify an infecting serovar. Cultures must be incubated for several weeks. Repeated specimen collection may be required due to intermittent shedding. The sensitivity of PCR assays is still not well established, and they do not provide information about the infecting serovar, although they have been used to provide information on genotype. The author’s experience is that PCR may be insensitive for diagnosis of canine leptospirosis, but the sensitivity and specificity may vary geographically depending on the serovars present and shedding patterns that occur for those serovars. PCR assays are best performed on blood AND urine concurrently because urinary shedding begins 10 days after the onset of infection. UC Davis now offers a multimodality approach to diagnostic testing for leptospirosis that includes serology with or without culture and PCR.

http://www.vetmed.ucdavis.edu/foley_lab/leptospira/index.cfm



Treatment Specific treatment involves initial use of parenteral penicillin derivatives for leptospiremia. In the author’s hospital, ampicillin is generally used (20 mg/kg IV q6-8h, adjusting dose down if severe azotemia is present) for up to 14 days or as long as the patient is vomiting or appears nauseated. It is recommended that treatment then be changed to doxycycline (5 mg/kg PO q12h) for 2 weeks, in order to eliminate the carrier phase. Doxycycline can be used instead of penicillins if vomiting does not occur after administration. Supportive therapy is also indicated for acute renal failure (eg. IV fluids, H2 blockers, antihypertensives, gastric protectants, antiemetics, phosphate binders, packed red cells and nutritional support). The use of hemodialysis can improve survival in dogs with severe renal failure. Approximately 50% of the patients with leptospirosis at the author’s institution are dialyzed, and the average number of treatments required before polyuria and recovery occurs is 3. Euthanasia or death due to leptospirosis is recorded in 18% of our dogs. Prevention In North America, vaccines are available for serovars Canicola, Icterohaemorrhagiae, Pomona and Grippotyphosa and in widespread use. The vaccines are generally safe and efficacious and studies suggest they provide a 1-year duration of immunity (Minke et al, 2009; Klaasen et al, 2003). A recent study at our institution that examined over 130,000 dogs seen at a mobile vaccine clinic showed that although administration of a Leptospira vaccine increased the risk of adverse reactions, the risk of a reaction was still extremely low (0.45%, compared with 0.28% for all vaccines). Moreover, when broken down by type of adverse event, the rate of hypersensitivity-type events (most severe) increased from 7.2/10,000 dogs to 9.1/10,000 dogs with administration of Leptospira vaccine and this increase was not significant (Yao et al, 2015). Although it was prevalent when the two-way (Canicola and Icterohaemorrhagiae) vaccines were in widespread use, vaccine failure appears to be extremely rare with the current 4-serovar vaccines (Hennebelle et al, 2013). In Europe, only 2-way vaccines have been available until very recently, and disease has been occurring in vaccinated and unvaccinated dogs. New vaccines are being introduced that contain three (Icterohaemorrhagiae, Canicola, and Grippotyphosa), or four (Icterohaemorrhagiae, Canicola, Grippotyphosa, and Bratislava) serovars. Leptospira bacterins have been associated with occasional acute, severe allergic reactions, but the incidence of these reactions has decreased dramatically in recent years, and reaction rates appear to be approaching those of distemper-hepatitis-parvovirus vaccines, even in small breed dogs. Vaccination against pathogenic leptospires is strongly recommended for dogs living in areas where leptospirosis occurs (ie. throughout the US), and are recommended even for small breed dogs that are confined to urban backyards, because of the possibility of infection as a result of rodent exposure. Minimizing access to rodents, farm animals and other wild animals also should help to prevent infection. Public Health Risk Leptospirosis remains an important zoonosis, although most documented human leptospirosis in North America results from recreational activities that involve water, rather than contact with dogs. Because dogs are generally incidental hosts they may not shed for significant periods of time, although more



studies are required to confirm this, and there are anecdotal reports of leptospirosis in staff that work in veterinary hospitals. Human leptospirosis is typically a ‘flu-like illness’, but in some cases may be associated with vomiting, diarrhea, shock, jaundice, renal failure, pneumonia, meningitis, or abortion. Any animal with acute renal failure should be treated as a suspect. Warnings should be placed on cages, gloves should be worn while handling these dogs and bleach or iodine-based disinfectants should be used to clean areas soiled with urine. Owners should be warned that without specific treatment, leptospires may be shed in the urine for months despite clinical recovery. The ACVIM has published consensus guidelines for the diagnosis, treatment, and prevention of leptospirosis in dogs (Sykes et al, 2011). References 1. Miller MD, Annis KM, Lappin MR, et al. Variability in results of the microscopic agglutination test in

dogs with clinical leptospirosis and dogs vaccinated against leptospirosis. J Vet Intern Med 2011;25(3):426-432,

2. Minke JM, Bey R, Tronel JP, et al. Onset and duration of protective immunity against clinical disease and renal carriage in dogs provided by a bi-valent inactivated leptospirosis vaccine. Vet Microbiol 2009;137:137-145.

3. Klaasen HL, Molkenboer MJ, Vrijenhoek MP, et al. Duration of immunity in dogs vaccinated against leptospirosis with a bivalent inactivated vaccine. Vet Microbiol 2003;95:121-132.

4. Yao PJ, Stephenson N, Foley JE, et al. Incidence rates and risk factors for owner-reported adverse events following vaccination of dogs that did or did not receive a Leptospira vaccine. J Am Vet Med Assoc 2015;247:1139-1145.

5. Hennebelle JH, Sykes JE, Carpenter TE, et al. Spatial and temporal patterns of Leptospira infection in dogs from northern California: 67 cases (2001-2010). J Am Vet Med Assoc 2013;242:941-947.

6. Sykes JE, Hartmann K, Lunn KF, et al. 2010 ACVIM small animal consensus statement on leptospirosis: diagnosis, epidemiology, treatment and prevention. J Vet Intern Med 2011;25(1):1-13.

7. Lee HS, Guptill L, Johnson AJ, Moore GE. Signalment changes in canine leptospirosis between 1970 and 2009. J Vet Intern Med 2014;28(2):294-299.

8. Ball C, Dawson S, Williams N. Leptospira cases and vaccination habits within UK vet-visiting dogs. Vet Rec 2014;174(11):278.

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