Bacterial Diseases of Nonhuman Primates

GI diseases are a major problem in captive nonhuman primates. Clinical signs include the following:

• watery or mucoid, blood-tinged feces

• rapid dehydration

• weight loss or emaciation

• lethargy or prostration

• rectal prolapse

The bacteria most commonly associated with GI disease in nonhuman primates are Shigella spp, Campylobacter jejuni, Yersinia spp, and Escherichia coli. Occasionally, Klebsiella pneumoniae, Salmonella spp, Pseudomonas aeruginosa, Lawsonia intracellularis, Aerobacter aerogenes, and Aerobacter hydrophila are implicated. Nonhuman primates can be intermittent, subclinical carriers of any of these organisms.

Helicobacter spp have been implicated as a cause of gastritis, anorexia, and vomiting.

Viral etiologies, such as noroviruses and rotaviruses, are often underrecognized and underreported as causes of acute GI upset with or without infectious bacteria. Helminths or pathogenic protozoa can also be complicating factors, especially in juvenile animals.

Smaller primates dehydrate and become hypoglycemic very rapidly. Mortality rates can be extremely high in acute outbreaks of GI disease, unless treatment is instituted promptly to restore and maintain normal fluid and electrolyte balance. Severe clinical signs and death are generally due to dehydration, hypokalemia, and metabolic acidosis. The most common lesions at necropsy are hemorrhagic enteritis, enterocolitis, colonic ulcers, or simply colitis.

With many privately owned nonhuman primates, dietary indiscretion is common, and the owner should be asked about irregular feeding habits. Fecal flotation and trichrome stains should be performed, and appropriate anthelmintics should be administered. Fecal culture or PCR assay should be performed to determine the presence of infectious agents, and culture and susceptibility testing should be performed if antimicrobial susceptibility data are not available.

Abdominal ultrasonography could inform treatment by identifying the type of inflammatory lesions by location (ie, gastric, small intestinal, cecal, or large intestinal) and the presence of contributory conditions such as neoplasia.

Endoscopy with biopsy or gastric lavage with culture can be used to differentiate gastric trichobezoars from Helicobacter or Shigella infection in cases of vomiting.

Hydration should be restored and maintained with parenteral electrolyte solutions. Although medications might be more easily administered parenterally, potassium, B vitamins, electrolytes, bismuth subsalicylate (40 mg/kg, PO, every 8–12 hours as needed for up to 14 days), and antimicrobial agents can be administered orally or by nasogastric tube in most nonhuman primates.

Bland diets, bismuth subsalicylate, fiber supplementation, and oral vitamins should be instituted until culture results are obtained.

For patients with anorexia, antiemetic agents (maropitant: 1 mg/kg, SC, once; or ondansetron: 0.1–0.2 mg/kg, IM or IV, once) or analgesic treatment (buprenorphine: 0.01–0.03 mg/kg, IM or IV, every 8–12 hours for great apes and Old World monkeys, or 0.005–0.01 mg/kg, IM or SC, every 6–8 hours for New World monkeys; or buprenorphine extended-release formulation: 0.2–0.25 mg/kg, SC, every 2–5 days) should be strongly considered.

Many nonhuman primates are carriers of potentially infectious, opportunistic microorganisms that can be transmitted to other animals and humans and that can develop resistance to antimicrobial treatment. Therefore, mild to moderately affected nonhuman primates and those without clinical dehydration should be treated with supportive care only, without antimicrobials, to avoid increasing antimicrobial resistance in these organisms.

Empirical antimicrobial treatment is generally indicated for the following conditions:

• for acute shigellosis: azithromycin (40 mg/kg, PO, once; then 20 mg/kg, PO, every 24 hours for 4 days) or enrofloxacin (10 mg/kg, PO or IV, every 24 hours for 7 days)

• for Campylobacter-associated diarrhea: azithromycin or tylosin (10–20 mg/kg, PO, every 24 hours for 14 days)

Upper respiratory diseases of bacterial origin can cause widespread illness, and bacterial pneumonia is associated with increased mortality rates, particularly in newly imported or immature nonhuman primates. Causative agents include Streptococcus pneumoniae, K pneumoniae, Bordetella bronchiseptica, Haemophilus influenzae, various other Streptococcus species, and various Staphylococcus and Pasteurella species.

Both New World and Old World primates are highly susceptible to cross-species transfer of respiratory pathogens from humans. Often, caretakers have had a history of an upper respiratory infection, so obtaining an adequate history on presentation is important.

Hand-reared infants frequently present with aspiration pneumonia from bottle feeding, and radiography and culture (via transtracheal lavage or nasal swab) are indicated.

Pneumonia can accompany or follow other primary diseases (eg, dysentery or respiratory viral infection) in nonhuman primates. Clinical signs include the following:

• coughing

• sneezing

• dyspnea or tachypnea

• mucoid or mucopurulent nasal discharge or epistaxis

• fever

• lethargy

• anorexia

• weight loss

Empirical antimicrobial treatment with one of the following drugs is generally indicated:

• azithromycin: 40 mg/kg, PO, once; then 20 mg/kg, PO, every 24 hours for 4 days

• penicillin: 20,000–60,000 IU/kg, SC, every 24 hours for 7–14 days (higher dosage for lemurs)

• a cephalosporin: cephalexin (25 mg/kg, PO, every 12 hours for 5–7 days) or cefazolin (25 mg/kg, IM or IV, every 12 hours for 5–7 days)

Cultures from pharyngeal swabs or transtracheal lavage are useful to isolate the causative agent of pneumonia and determine the specific antimicrobial susceptibility. Commercially available human respiratory PCR panels can assist in differentiating viral from bacterial etiologies. Intensive nursing and other supportive therapy, such as fluids and oxygen administration, can also aid recovery in select cases.

Most nonhuman primate species have cervical air sacs that affect respiratory disease susceptibility and course, especially in some captive species, such as orangutans and howler monkeys. Air sacculitis can be subclinical to life-threatening and can become chronic. Clinical signs include nasal discharge, halitosis, and distention of the air sacs with exudate. Animals with respiratory infections should be monitored for these signs and treated with antimicrobials indicated by culture and susceptibility test results and with surgical drainage of the air sac, if necessary.

Infection with Clostridium tetani, particularly as a consequence of fighting, parturition, frostbite, and other forms of skin trauma, is a risk with free-ranging and outdoor-housed monkeys.

For populations at risk, immunization with tetanus toxoid, using an adsorbed human product or an equine product at 0.25 mL per animal, IM, every 5 years starting at 1 year old, should be considered. (Also see Tetanus in Animals.)

Prior immunization is critical, because tetanus is life-threatening. As in other species, in nonhuman primates treatment involves administration of antitoxin, appropriate antimicrobials (eg, penicillin), and supportive care.

All nonhuman primates are susceptible to tuberculosis caused by Mycobacterium bovis or Mycobacterium tuberculosis complex; however, major differences exist in the prevalence of cases in different species. Most cases of tuberculosis in nonhuman primates have been reported in Old World primates such as macaques. Reports in New World species such as squirrel monkeys are much less frequent. The incidence of tuberculosis is < 1% among quarantined Old World primates, but 45% of cases (especially in cynomolgus monkeys) might not be diagnosed until after the first 30 days of quarantine.

Clinical signs are not a reliable indication of the severity of tuberculosis in monkeys. A monkey that appears healthy could have extensive miliary disease involving thoracic and abdominal organs. Signs of debilitation might appear only shortly before death. However, advanced tuberculosis should be suspected in animals that cough, lose appetite or weight, or have enlarged or draining lymph nodes, skin wounds that fail to heal, or abdominal masses.

A testing program is mandatory. Tuberculin tests should be performed on all nonhuman primates on arrival at the facility and at 2-week intervals thereafter until negative results have been recorded on at least three consecutive tests for the entire group. The last test should be administered within 14 days of release from quarantine and introduction into an established colony. After their release from quarantine, all nonhuman primates should be skin-tested at least semiannually.

A tuberculin skin test (intradermal tuberculin skin test), using mammalian old tuberculin (MOT), is currently the primary diagnostic method for routine surveillance. Intradermal injection of 0.1 mL of MOT using a 26-gauge needle in the upper palpebrae is the standard method in most species. Injection into the eyelid (see tuberculosis test site image) enables monitoring for swelling at the injection site without recapture and restraint. In smaller species such as marmosets, tamarins, and prosimians, the skin of the abdomen is used rather than the eyelid.

Tuberculosis test site, capuchin

Image

Courtesy of Dr. Terri Parrott.

The injection sites are observed at 24, 48, and 72 hours after injection. Grading systems can be found in the National Institutes of Health Guideline for the Prevention and Control of Tuberculosis in Nonhuman Primates.

The presence of palpebral swelling with a lid droop can be interpreted as a positive tuberculosis test result. In some species (eg, orangutans), false-positive skin test results are common.

A combination of testing, including thoracic and spinal radiography, ELISA, and interferon-gamma assays, might be required to differentiate tuberculosis from nontuberculous mycobacterial infections or general reactivity to the skin test. Comparative tests using both mammalian and avian purified protein derivative tuberculin, cultures, and PCR assay of gastric or bronchial lavage, feces, or tissues can further eliminate confusion.

The time from initial infection to skin test conversion depends on the route of exposure, the infecting inoculum, and the bacterial strain; in rhesus monkeys it is usually 4–8 weeks. Late in the disease, anergy (lack of immune response) can lead to a negative skin test result. Anergy can also be induced by concurrent viral infections, such as measles or immunosuppressive diseases.

Infected nonhuman primates can also develop latent tuberculosis. These animals appear healthy, are not infectious, and often have negative skin test results. Latent tuberculosis can be reactivated in response to environmental stressors or other immune modulation, leading to active disease and potential transmission.

Chest radiographs can aid diagnosis of tuberculosis in well-established cases; however, radiographs are unreliable in diagnosis because tuberculosis lesions rarely calcify or cavitate in nonhuman primates as they do in humans. Tracheobronchial lymph node enlargement and lung consolidation might be the only radiographic signs, and these might be clear only on thoracic CT images.

In questionable cases of tuberculosis in nonhuman primates, additional tests, such as culture, PCR assays, staining of gastric lavage or transtracheal wash samples, ELISAs, and comparative abdominal skin testing with avian and atypical tuberculins, could also aid diagnosis.

Biopsies of positive or suspicious abdominal skin tests could be useful to identify true delayed-type hypersensitivity reactions in nonhuman primates. Because of the public health risk, euthanasia is recommended for all animals with positive skin test results. Tuberculosis should then be confirmed at necropsy. When performing necropsies on suspect cases, all personnel should use appropriate safety equipment and personal protective equipment, such as biosafety cabinets rated for Biosafety Level 3 work, N95 respirators, and/or powered air-purifying respirators.

When a positive case of tuberculosis is identified, the entire group of exposed nonhuman primates should be quarantined, with skin testing performed at 2-week intervals for a minimum of 4–6 weeks. For animals in quarantine, the quarantine period should be restarted, and testing should be continued.

Privately owned animals should be quarantined away from small children and untrained personnel. All personnel working with nonhuman primates or in primate facilities should have regular skin tests or blood tests (interferon-gamma release assays) at least annually to ensure that subclinical human carriers do not transmit tuberculosis to the nonhuman primates.