Infectious Disease

Infectious Disease

Bradley T. Simon1 and Jusmeen Sarkar2

1 Department of Small Animal Clinical Sciences, College of Veterinary Medicine and Biomedical Science, Texas A&M University, College Station, TX, 77843, USA

2 Anesthesia and Pain Management Service, Veterinary Specialty Center, Buffalo Grove, IL, 60089, USA


The One Health initiative alleges “to promote, improve, and defend the health and well‐being of all species” [1]. It utilizes collaborative endeavors integrating fields of science to educate and improve human and animal quality of life. Anesthetists play a vital role in this initiative. Zoonosis, iatrogenic administration of contaminated anesthetic drugs, unsafe injection practices, multidrug‐resistant bacterial infections, postoperative surgical site infections (SSIs), work‐area‐ and equipment‐related infectious transmission, and active infectious diseases compromising a multitude of body systems are among some of the considerations for anesthetists. Understanding how these considerations integrate into the perioperative period allows the anesthetist to reduce risk factors predisposing animals to morbidity and mortality.

Contamination and Cleaning of Anesthetic Equipment

Breathing Circuits

Bacterial colonization of anesthetic breathing circuits is reported in human [27] and veterinary medicine [8], most prevalent in the breathing circuit components closest to the patient (i.e., mask or elbow adapter) [2, 8, 9]. Opinions have varied on whether breathing circuits are considered safe for extended use [3, 4, 8, 10, 11]. One veterinary teaching hospital evaluated bacterial colonization in anesthetic breathing circuits subjected to routine use over a 2‐month period [8]. The presence of microorganisms of low pathogenicity was reported (Bacillus spp., coagulase‐negative Staphylococcus spp., or Micrococcus spp.), but none likely to cause postoperative pneumonia even in immunocompromised patients [8]. Cooler temperatures of anesthetic and compressed gases, evaporation following breathing circuit disconnection from the patient, bactericidal effects of high oxygen concentrations, bacteriostatic properties of the circuit, and alkaline condensate in the carbon dioxide absorbent all reduce the risk for bacterial and viral colonization associated with reusable breathing circuits [2, 8, 12, 13]. A filter placed between the patient and breathing circuit [8] or between the breathing expiratory limb and anesthesia machine may reduce the risk for bacterial and viral colonization (figure 16.1). Buildup of moisture within anesthetic circuit breathing filters may increase airway resistance, impeding adequate ventilation [14], and filters should be replaced with buildup of moisture, prolonged use, or excessive coughing.

Schematic illustration of a particulate-air filter placed at the end of the expiratory limb between rebreathing circuit and the anesthesia machine.

Figure 16.1 A particulate‐air filter placed at the end of the expiratory limb between rebreathing circuit and the anesthesia machine (red arrow). These filters are used to protect the internal surfaces of the breathing circuit and do not allow more than 5% of particles 0.3 μm in diameter or larger to penetrate through the filter.

Endotracheal Tubes

Endotracheal tubes are recommended for single use, as they come in direct contact with the patient’s tracheal mucosal surfaces. Single‐use endotracheal tubes may not always be practical in the veterinary setting; however, it is the anesthetist’s obligation to ensure that adequate decontamination protocols are followed (Table 16.1) [15]. Disinfection of experimentally contaminated endotracheal tubes with Streptococcus zooepidemicus and Bordetella bronchiseptica was achieved with a 5‐min 0.5% chlorhexidine gluconate solution soak [16]. Spraying all surfaces of the endotracheal tube with an accelerated hydrogen peroxide solution is effective to a lesser extent [16]. A cold‐water soak is not effective in disinfecting endotracheal tubes and is not recommended. Intubation equipment, laryngoscope blades, and laryngoscope handles also require disinfection, as they are a potential source of contamination [17]. Decontamination with a 2% chlorhexidine/70% alcohol wipe was effective against several species of microorganisms including methicillin‐resistant Staphylococcus aureus (MRSA) [18].

Anesthetic Equipment and Monitors

Blood contamination on anesthetic equipment and monitors is a source for infectious disease transmission. A total of 418 surfaces on anesthesia machines, anesthesia carts, and monitors frequently handled by anesthesia personnel were evaluated in one human hospital for blood contamination [19]. Prior to examination, each surface was deemed ready for use by a member of the anesthesia staff. Of the surfaces examined, 137 (33%) were positive for blood contaminants and only three had blood visible to the unaided eye [19]. The highest prevalence of blood contamination was found on anesthesia monitoring cables (82%), drawer handles (64%), and pulse oximeter probes (59%) [19]. Commercially available disinfecting wipes containing sodium hypochlorite as the active ingredient are effective in cleaning most types of microbes from anesthesia equipment [20, 21]. Wiping with 70% isopropanol does not adequately clean and disinfect reusable pulse oximetry sensors [21]. Utilizing standard operating procedures (SOPs) for routine cleaning of anesthesia equipment and monitors may improve the efficiency and quality of care, and reduce incidences of nosocomial transmission during the perianesthetic period.

Multidrug‐Resistant Organisms and Surgical Site Infection

Multidrug‐Resistant Organisms

The emergence of multidrug‐resistant organisms (MDRO) in hospital settings is of growing concern and is an important cause of morbidity and mortality [22]. Currently,, extended‐spectrum beta‐lactamase Enterobacteriaceae and MDR Enterococcus, Acinetobacter, and Pseudomonas spp. are of most concern [22]. Factors predisposing patients to the development of MDROs include duration and type of antibiotic use, disruption of intrinsic protective barriers of the body, age extremes, presence of diabetes mellitus, and immune suppression [15]. To decrease MDRO transmission and SSIs, anesthetists should receive training in identifying patient risk factors, aseptic techniques and antibiotic regimens, and appropriate decontamination procedures. Surgical patients receiving antibiotics for their MDRO should receive an additional dose [15] 30–60 min before surgical incision [23, 24]. Locations and transport of anesthetized patients diagnosed with MDROs should be kept to a minimum. The anesthesia team should wear disposable gowns, gloves, surgical caps and masks, and avoid contact to nearby surfaces, staff, and patients. An in‐line breathing circuit filter can be added to reduce the risk of cross‐contamination [25]. Staff members without MDRO patient contact should assist in opening doors, cages, kennels, and elevators as required. Anesthetic equipment should be cleaned and disinfected as previously described, but, in MDRO patients, breathing circuits and endotracheal tubes should be kept to single use.

Table 16.1 Cleaning processes, the microbiological organisms they render nonviable, and methods for implementation.

Source: Adapted with permission from Einav and Wiener‐Well [15].

Process Microorganisms rendered nonviable Methods Required for items Examples
Bacteria Viruses Fungi Bacterial spores
Sterilization Yes Yes Yes Yes Pressurized steam, dry heat, ethylene oxide and hydrogen peroxide Intended to contact a normally sterile body area/cavity
That may inadvertently be inserted into a normally sterile body area/cavity
Surgical equipment and single‐use disposables (e.g., needles, scalpels, ventilation tubing and filters, endotracheal tube, urinary catheter and drains)
Esophageal stethoscope
High‐level disinfection Yes Yes Yes No Aldehydes, peracetic acid and chlorine dioxide That contact mucous membranes, but do not penetrate the membrane or come in touch with blood Ventilation masks, components of the laryngoscope, endoscopes, oral and nasal airways, and connectors
Intermediate‐level disinfection Yes Most
Not small viruses without envelopes
Yes No Alcohol, sodium, hypochlorite, phenols, and iodophors That contact the external parts of the body (i.e., skin and hair) Blood pressure cuffs, stethoscopes, pulse oximeter
probes, head rests and straps, and ECG
Low‐level disinfection Most Some Some No Alcohol and quaternary ammonium compounds Noncritical patient‐care surfaces and equipment that contacts intact skin Bedrails, over‐the‐bed table or stand, and medical devices and equipment

All contaminated equipment/devices must first be cleaned (thus decreasing their bacterial load) in order to achieve appropriate disinfection or sterilization. When items have been within the vicinity of a patient with a multidrug‐resistant organism (MDRO; i.e., in the same room), disposable items should generally be discarded even if they have not been used and multiple‐use items should be assumed to have been in direct contact with the patient.

Surgical Site Infections

SSIs have tremendous clinical and financial impact in veterinary medicine [26, 27]. In humans, it is estimated that SSIs reach more than $3.5–$10 billion annually in health care expenditures [28]. The Centers for Disease Control and Prevention (CDC) Healthcare‐Associated Infections guidelines have defined SSI to be an infection that occurs in the part of the body (incision or organ) where the surgery took place [28]. The rate of SSI in small‐animal veterinary medicine is approximately 5.5–5.8% [29, 30]. Staphylococcus spp., a bacteria commonly found within the skin, is the most common cause for SSI in dogs and cats [31]. The incidences of SSI in dogs and cats for clean, clean‐contaminated, contaminated, and dirty procedures are 2.5–4.9%, 4.5%, 5.8–9.1%, and 17.8–18.1%, respectively [29, 32]; with the potential of being as high as 13.3% following clean orthopedic procedures [33]. Minimally invasive surgery may be associated with a lower incidence of SSI when compared to open surgery in dogs and cats [34]. Dogs with SSIs, when compared to those without infections, have higher mean postoperative costs, longer duration from surgery to final closure of the case, and more postoperative visits [26]. Currently there is no association between SSI and long‐term effects on functional outcome in dogs following orthopedic surgery [35]; however, additional studies are required. Anesthetists are an important factor in the prevalence of SSI during the perioperative period (Table 16.2) [36]. Anesthesia‐specific factors associated with the development of SSI include hypoxemia, increasing American Society of Anesthesiologists (ASA) physical status, hypothermia, hyperglycemia, administration of contaminated propofol, inadequate antibiotic prophylaxis, prolonged anesthesia duration, contaminated anesthesia gurneys, and postoperative anemia [27, 31,3741]. Additional factors include concurrent endocrinopathy (diabetes mellitus, hyperadrenocorticism, hypothyroidism), prolonged surgery time, increasing number of people in the operating room, additional days spent in the intensive care unit (ICU), obesity, extremes in age, intact males, dirty surgical site, surgical site clipped before anesthetic induction, surgery performed during the warmer summer months, and prolonged use of antibiotics in clean wounds [29, 30,4245]. Inhalant‐based anesthesia techniques also may increase the risk for SSI when compared to neuraxial techniques [46, 47]; however, there are conflicting reports [48]. The influence of forced‐air warming blankets on the spread of nosocomial infections has been of much debate [49, 50]. Currently, there is no strong evidence to suggest that these warming devices increase the risk for SSI [49, 51]. On the contrary, forced‐air warming devices may provide benefit in reducing SSI by minimizing perioperative hypothermia [50, 52]. Pharmacological and nonpharmacological factors that reduce the incidence of SSI include appropriate volume resuscitation and fluid therapy, normothermia, adequate analgesia, intraoperative and postoperative oxygen supplementation, normoglycemia, and prophylactic antibiotic therapy [53, 54]. Furthermore, lidocaine, thiopental, and diphenhydramine all possess bacteriostatic properties, and, when combined with propofol, may decrease the potential risk for bacterial contamination [5558]. Recommendations from the CDC guidelines of SSI are represented in Table 16.3 [27, 28, 59].

Table 16.2 Risk factors for the development of surgical site infections.

Source: Adapted with permission from Barie and Eachempati [36]. Data from: National Nosocomial Infections Surveillance System (NNIS) System Report: Data Summary from January 1992–June 2001, issued August 2001. Am J Infect Control 2001; 29:  404–21.

Patient factors
Chronic inflammation
Corticosteroid therapy
Diabetes or hyperglycemia
Extremes of age
Peripheral vascular disease
Postoperative anemia
Prior site irradiation
Remote infection
Skin carriage of Staphylococci spp.
Skin disease in the area of infection
Surgical site clipped before anesthetic induction
ASA physical status > II
Environmental factors
Contaminated medications (i.e., propofol)
Contaminated anesthesia gurneys
Inadequate disinfection/sterilization
Inadequate skin antisepsis
Inadequate ventilation
Dirty surgical site
Surgery performed during
warmer summer months
Increased number of
people in the surgical
Treatment factors
Emergency procedure
Inadequate antibiotic prophylaxis
Oxygenation (controversial)
Prolonged preoperative hospitalization
Prolonged operative time
Prolonged anesthesia time
Prolonged use of antibiotics in clean wounds

Suboptimal timing, duration, and choice of perioperative antibiotics are still being practiced in veterinary medicine [33, 60]. Inappropriate use of prophylactic antibiotics may be due to failure of the veterinarian to keep up to date on current practices or failure to transition to evidence‐based practices [60]. Examples of inappropriate use include the use of antibiotics for clean surgical procedures (except surgeries involving surgical implants), administration of prophylactic antimicrobials postoperatively, and the continued administration of antibiotics for longer than 24 h following surgery [61]. In contrast to the latter examples, recent evidence suggests that postoperative antibiotics may reduce the incidence of SSI following clean orthopedic implant surgery in dogs [62, 63]; however, duration of administration may be an important factor in their effectiveness [64].

Table 16.3 Recommendations from Centers for Disease Control and Prevention guidelines for the prevention of surgical site infection, 2017 [27, 28, 59].

  • Parenteral antimicrobial prophylaxis

    • Administer antimicrobial agents only when indicated based on published guidelines

    • Time administration such that bactericidal concentration is established in serum and tissues at initial incision

  • Glycemic control

    • Implement perioperative glycemic control using blood glucose target levels <200 mg dl−1 in patients with and without diabetes

  • Normothermia

    • Maintain perioperative normothermia

  • Oxygenation

    • Administer increased fractions of inspired oxygen intraoperatively and in the immediate postoperative period following extubation for all patients with normal pulmonary function undergoing general anesthesia with endotracheal intubation

  • Antiseptic prophylaxis

    • Instruct owners to perform full body bathing the night before surgery with either soap or an antiseptic agent. Allow complete drying of densely haired patients to avoid skin irritation

    • Clipping of hair adjacent to the surgical site with clean, appropriately serviced clippers after induction of anesthesia and immediately before the surgical procedure

    • Intraoperative skin preparation should be performed with an antiseptic agent containing alcohol unless contraindicated

    • Nail polish, jewelry, and artificial nails should be avoided or removed in all members of the anesthesia and surgery team prior to handling of the patient during the perioperative period

In most veterinary settings, the anesthetist administers the prophylactic antibiotics to the surgical patients. Withholding prophylactic antibiotics can increase the rate of SSI. For most antibiotics, complete distribution into tissues can occur within 30–60 min following intravenous administration [61]. In a patient not previously exposed to antibiotics, an initial IV dose of cefazolin administered 10–30 min before skin incision and at the completion of the procedure is recommended. For procedures lasting longer than 2 h, additional intraoperative doses should be administered every 90 min following the initial dose [61]. For most surgical procedures, a first‐generation cephalosporin such as cefazolin is effective. For surgeries in which the bowel is entered, coverage of Gram‐negative organisms is important, and anaerobic coverage is appropriate if the large bowel or female genital tract is entered. Additional studies are needed to determine the recommended duration of postoperative prophylactic antibiotic administration in veterinary medicine.

Respiratory Infections

Aspiration, Bacterial, and Viral Pneumonia

Aspiration Pneumonia

Aspiration pneumonia is one of the leading causes of infectious pneumonia in human patients hospitalized in the ICU [65], with an overall mortality rate of approximately 45% [66]. Mortality rate is lower in dogs than in humans but can be as high as 25% [67, 68]. A retrospective study evaluating over 140 000 dogs anesthetized or sedated over a 10‐year period reported a 0.17% incidence of postanesthetic aspiration pneumonia [69]. Several factors may predispose a patient to the development of aspiration pneumonia (Table 16.4) [6972]. Two or more anesthesia‐related events, increased anesthetic duration, regurgitation and hydromorphone administration at induction, postanesthetic vomiting and regurgitation, and procedures such as laparotomy, upper airway surgery, neurosurgery, thoracotomy, magnetic resonance imaging (MRI), and endoscopy were associated with the development of aspiration pneumonia [69, 70, 73, 74]. Megaesophagus, history of preexisting respiratory (i.e., laryngeal paralysis or brachycephalic syndrome) or neurologic disease, preanesthetic tetraparesis, and cervical lesions were also associated with aspiration pneumonia [69, 70, 73, 75]. There are no reported incidences of postanesthesia aspiration pneumonia in cats; however, one retrospective analysis reported an 11% mortality rate following aspiration pneumonia [72]. Risk factors in cats are likely similar to those observed in dogs.

Table 16.4 Factors associated with aspiration pneumonia in dogs and cats [6972].

Source: Adapted with permission from O’Hara et al. [59].

Gastrointestinal disease
Recent history of vomiting
Foreign body obstruction
Chronic gastrointestinal disease
Enteral nutrition
Neurologic disease
Myasthenia gravis
Conditions leading to prolonged recumbency
Preanesthetic tetraparesis
Esophageal disease
Esophageal motility disorder
Hiatal hernia
Esophageal stricture
Upper airway surgery
Magnetic resonance imaging
Pharyngeal and laryngeal disease
Muscular dystrophy
Oropharyngeal dysphagia
Laryngeal disease or trauma
Increased duration of anesthesia
Recent history of anesthesia
Two or more anesthesia‐related events
Regurgitation during anesthesia
Administration of hydromorphone at induction
Respiratory disease
Upper respiratory disease
Postprocedural upper airway obstruction
Brachycephalic (Bulldog, Pug)
Golden Retriever
Cocker Spaniel
English Springer Spaniel
Domestic Shorthair

Inhalation of sterile (aspiration pneumonitis) or septic (aspiration pneumonia) acidic orogastric contents irritates the pulmonary mucosa and fosters an environment in which bacterial colonization can occur. Damage associated with aspiration pneumonia can be categorized into two distinct phases. The first phase results from the direct chemical burn of acidic gastric contents resulting in pulmonary parenchymal inflammation. Pulmonary gas exchange is compromised, and the lungs become susceptible to bacterial colonization leading to bacterial pneumonia [76, 77]. Acidic contents stimulate alveolar macrophages and type II pneumocytes to release inflammatory cytokines, which occurs during the second phase. Inflammatory cytokines recruit neutrophils in high concentrations which release proteases and oxidants, compromising the integrity of the vascular endothelial walls in the pulmonary parenchyma [78]. This increases permeability to proteins and fluid, with associated accumulation of edema, fibrin, hemorrhage, and neutrophils. These infiltrates result in the development of atelectasis, airway collapse, and disruptions to the mucociliary action causing bacterial entrapment and infection [76, 78].

Bacterial and Viral Pneumonia

Bacterial pneumonia is one of the most common clinical diagnoses in dogs with respiratory disease and is often associated with viral infections or aspiration of gastrointestinal (GI) contents. In dogs, it is often due to canine infectious respiratory disease (CIRD) complex, a syndrome that involves a multitude of viral (canine respiratory coronavirus, influenza virus, herpesvirus, pneumovirus, parainfluenza, bocavirus, hepacivirus, picornavirus) and bacterial (B. bronchiseptica, Streptococcus canis, Streptococcus equi subsp. zooepidemicus, and Mycoplasma cynos) pathogens [71, 79]. Pathogens associated with infectious pneumonia in cats include Pasteurella spp., Escherichia coli, Staphylococcus spp., Streptococcus spp., Pseudomonas spp., B. bronchiseptica, and Mycoplasma spp. [71]. If the innate immune system is unable to clear the pathogens from the lower respiratory system, bronchopneumonia or bronchointerstitial pneumonia develops, damaging the type I pneumocytes and the respiratory epithelium. Progression of the disease is similar to that seen during aspiration pneumonia [71, 80].

Clinical Signs

Clinical signs are dependent on the patient’s physical status and severity and chronicity of disease. In cases of aspiration pneumonia, clinical signs also depend on the volume and pH of aspirated orogastric contents. Anesthetized or neurologic patients have a less pronounced cough reflex, which makes them prone to aspiration and the development of acute respiratory distress syndrome (ARDS) [69, 81]. Fatalities in dogs and cats with ARDS following aspiration pneumonia are approximately 84% and 100%, respectively [81]. Early clinical signs include soft coughing, but as the disease worsens, productive coughing, dyspnea, exercise intolerance, lethargy, decreased appetite, syncope, cyanosis, and orthopnea may develop. Cats exhibit less pronounced clinical signs when compared to dogs. Increased respiratory effort, anorexia, vomiting, and lethargy were among the most common presenting complaints for cats diagnosed with pneumonia [72]. Other clinical signs in cats include coughing, drooling, oral bleeding, regurgitation, weight loss, and tachypnea with short, shallow breaths and nasal flaring [71, 72]. Increased work of breathing from decreased pulmonary compliance and increased pulmonary shunting contributes to the development of hypoxemia in patients with pneumonia [72, 80].


A thorough history and physical examination are extremely important in the diagnosis of pneumonia. Recent history of an anesthetic event may be indicative of aspiration pneumonia. Anesthetists should pay close attention to the patient’s respiratory rate and effort. Thoracic auscultation may reveal harsh lung sounds, crackles, and/or wheezes. Thoracic radiographs are an important tool in diagnosing and determining the severity of pneumonia in dogs and cats (Figure 16.2). Patients with pneumonia can develop concurrent upper airway disease (nasal congestion or discharge) and therefore upper auscultation should also be performed prior to sedation or anesthesia [71]. Fever does not always accompany pneumonia in dogs and cats and is not a reliable indicator of disease [68, 72]. In cats with pneumonia, 71%, 18%, and 11% had an alveolar pattern, unstructured interstitial pattern, and mixed alveolar–interstitial pattern, respectively, with the right middle lung and left cranial lung lobes most affected [72]. Cats commonly had an inflammatory leukogram on complete blood count with occasional band neutrophils [72].

Photo depicts radiographic evidence of a dog diagnosed with aspiration pneumonia.

Figure 16.2 Radiographic evidence of a dog diagnosed with aspiration pneumonia.

Source: Photo courtesy of Dr. Jessica Vallone.

Anesthetic Management

In cases of acute pneumonia, anesthesia and surgery should be postponed until the patient is deemed stable following antimicrobial therapy, intravenous fluids to correct hydration deficits, nebulization and coupage, oxygen supplementation, and oral mucolytic agents such as N‐acetylcysteine. Patients diagnosed with pneumonia that require emergent anesthesia should be ventilated using protective ventilation strategies. Goals for protective ventilation include optimization of patient–ventilator synchrony, reduction in iatrogenic injury, and supporting gas exchange (Figure 16.3) [82]. Tidal volume should not exceed 6–8 ml kg−1 of lean body weight with a rate of 8–15 breaths min−1 to achieve adequate oxygenation (SpO2 >95%) and ventilation (end‐tidal carbon dioxide [EtCO2] between 35 and 50 mmHg) [82]. Hypercapnia is generally well tolerated when pH is maintained between 7.20 and 7.45 in nonpregnant hemodynamically stable patients with normal intracranial pressures. A fractional inspired oxygen concentration of 100% is recommended in moderate to severe cases of pneumonia. Positive end‐expiratory pressure (PEEP) of 5–10 cm H2O can be provided in hypoxemic patients (PaO2 <60 mmHg; SpO2 <90%) (Figure 16.4). To decrease the likelihood of desaturation during anesthesia induction, preoxygenation via face mask at 100 ml kg−1 min−1 for a minimum of 3 min is recommended [83]. Elevating the head approximately 30° and placement of an orogastric tube can prevent the aspiration of gastric contents [84]. Endotracheal intubation with a cuffed endotracheal tube inflated to 20 cm H2O is recommended for the prevention of aspiration pneumonia. Active suctioning should be readily available to the anesthetist to remove any buildup of secretions within the oropharynx and endotracheal tube. Prevention of aspiration is key in patients with predisposing risk factors. Reductions in pure μ‐opioid receptor agonist administration, minimizing changes in position during intubation and recovery, prophylactic treatment of nausea with maropitant, metoclopramide, ondansetron, proton‐pump inhibitors, or H2‐receptor antagonists, providing optimal timing for endotracheal tube cuff deflation, and aseptic handling of intubation equipment are preventative measures in reducing the development of pneumonia [74].


Lactate and the pneumonia severity index (PSI) have been used to determine morbidity and mortality in human patients diagnosed with pneumonia (Table 16.5) [80, 85]. For example, more intensified monitoring and diagnostic evaluations are recommended in human patients with lactate levels >1.8 mmol l−1 or PSI scores >71 (Class II) [80, 85]. Mortality rate is approximately 25% in dogs [67, 68] and 11% in cats [72].

Canine Influenza A Virus

Canine influenza viruses (CIVs) are among the family Orthomyxoviridae that includes the Influenza A viruses (IAVs). The two major IAVs found in dogs include H3N2 and H3N8. Not until recently did evidence exist that dogs could be responsible for an epidemic spread and considered a natural host. H3N8 appears to be isolated among animal shelter dogs [86]. In 2015, another IAV, H3N2, was first introduced into the United States and is currently an ongoing widespread epidemic [87]. Transmission of H3N2 from dogs to cats has been reported in Korea and the United States but appears to be mainly found in shelter populations [88]. Humans are not susceptible to either form of IAV. Typically, these viruses are inactivated within minutes to hours in most environments; however, there is potential for them to persist longer (days) in cool, dark, damp conditions [86]. Transmission via the respiratory route usually requires direct contact and often is associated with animal shelters and boarding kennels [89]. Most commercially available disinfecting wipes and hot water will inactivate the IAV.

Schematic illustration of goals for providing protective ventilation in dogs and cats with aspiration or bacterial pneumonia.

Figure 16.3 Goals for providing protective ventilation in dogs and cats with aspiration or bacterial pneumonia [82]. Vt: tidal volume; RR: respiratory rate; PEEP: positive end‐expiratory pressure; FiO2: fractional inspired oxygen; SPO2: blood oxygen saturation; PaCO2: arterial carbon dioxide pressure.

The H3N8 and H3N2 CIVs predominately bind to the surface of epithelial cells in the bronchus, nasal mucosa, trachea, and type II pneumocytes [90]. This results in necrosis and exfoliation of the tracheal and bronchial mucosa (tracheitis and bronchitis, respectively), promoting the infiltration of lymphocytes and serous effusion in the alveolar spaces [89, 90].

Clinical Signs

CIV infections result in rapid onset of clinical signs associated with upper respiratory tract disease (coughing, sneezing, lethargy, fever, ocular and nasal discharge, and inappetence) [91]. Initially, CIV‐positive patients develop a persistent, dry, and nonproductive cough which can last for several weeks. As the disease continues and lung defenses become compromised via damage to the ciliary epithelium [92], secondary pneumonia may develop [89, 92]. In some instances, vomiting, diarrhea, and respiratory distress have been reported [91]. Despite the latter signs, both forms are associated with a low fatality rate (<5%) [91].

Photo depicts a positive end-expiratory pressure valve of 5 cm H2O mounted vertically on the expiratory side of a circle system (red arrow).

Figure 16.4 A positive end‐expiratory pressure valve of 5 cm H2O mounted vertically on the expiratory side of a circle system (red arrow).


A confirmed diagnosis of CIV is usually difficult due to the short incubation (2 days) and shedding (2–4 days) periods of the virus. Using reverse transcription polymerase chain reaction (RT‐PCR) on nasal swabs is the test of choice to detect CIV. Alternatively, hemagglutination inhibition titers are rapid and can detect antibody responses in dogs at 7 days post infection [89].

Anesthetic Management

CIV has the potential to spread rapidly among immunologically naïve or compromised animals. Patients suspected or diagnosed with CIV should be kept in isolation with limited contact to other patients. Changes from routine anesthetic management are dependent on the patient’s history, physical examination, and severity of clinical signs. Dehydration should be quantified and managed with a balanced crystalloid solution prior to general anesthesia. Oxygenation and ventilation should be assessed prior to anesthesia via use of capnometry, pulse oximetry, and/or arterial blood gas analysis. Side‐stream capnography can be used to acquire an estimate of the end‐tidal carbon dioxide concentration prior to induction (Figure 16.5). High (>50 mmHg) EtCO2 values may indicate inadequate ventilation due to upper airway disease, nasal congestion, or severe lower airway disease. Clinical signs of hyperventilation (EtCO2 <25 mmHg) or increased respiratory rate may be an indicator for hypoxia. Oxygen supplementation may be required following anesthesia. Clinical signs associated with hypoxia include tachypnea, tachycardia, restlessness, and cyanosis. The anesthetist should closely monitor ventilation and oxygenation during the recovery period. Preoxygenation and protective ventilation strategies are recommended especially in patients diagnosed with a secondary pneumonia. A balanced anesthetic regimen with the use of local regional techniques would minimize the respiratory depressant and immunosuppressant effects associated with inhalant anesthetics. Opioids may be beneficial in patients feeling pain with a dry nonproductive cough due to their high analgesic efficacy and antitussive properties, respectively [93, 94]. All airway management equipment should be sterilized prior to and following use to avoid secondary bacterial pneumonia or cross‐contamination, respectively.

Table 16.5 A pneumonia severity index used to determine the prognosis in patients diagnosed with pneumonia.

Factors Points Score and stratification
Neoplastic disease
Congestive heart failure
Cerebrovascular disease
Renal disease
Liver disease
Altered mental status
Systolic blood pressure <90 mmHg
Temperature <35 °C or ≥40 °C
Arterial pH <7.35
BUN ≥30 mg dl−1
Sodium <130 mmol l−1
Glucose ≥250 mg dl−1
Hematocrit <30%
PaO2 60 mmHg or SpO2 90%
Age +10
Point total
≤70 low risk
71–90 low
91–130 moderate
>130 high
Risk class

Adapted with permission from: Rider and Frazee [80].

Photo depicts the use of side-stream capnography (blue arrow) prior to induction of anesthesia to estimate the patient's end-tidal carbon dioxide concentration (red arrow) and ability to ventilate.

Figure 16.5 The use of side‐stream capnography (blue arrow) prior to induction of anesthesia to estimate the patient’s end‐tidal carbon dioxide concentration (red arrow) and ability to ventilate.

Feline Respiratory Disease Complex

Feline respiratory disease complex (FRDC) or feline upper respiratory tract infection (URI) refers to an acutely contagious respiratory or ocular disease caused by one or multiple pathogens (viral, bacterial, and/or fungal) [95]. Cats housed in crowded or densely populated housing are most often affected, and it is rarely diagnosed in singly housed indoor cats [95]. Stress and poor air quality may further exacerbate the severity of the disease and increase the transmission of FRDC [96]. For example, approximately 54% of cats were reported having mild to severe clinical signs associated with FRDC in eight United States animal shelters [97]. The most common pathogens include feline calicivirus, feline herpesvirus‐1 (feline rhinotracheitis), Chlamydophila felis, B. bronchiseptica, and Mycoplasma spp. [97]. Cryptococcus neoformans var. neoformans and Cryptococcus gattii have also been reported [98]. These organisms are often spread via bodily secretions (respiratory, ocular, and oral). Pending on the etiology, aerosol transmission can occur, but most often is due to contact with contaminated surfaces or direct exposure to infected cats [95].

Clinical Signs

Clinical signs are similar among etiologies and can range from mild to severe. Patients with FRDC often present with one or more of the following clinical signs: serious, mucoid, or mucopurulent nasal discharge, sneezing, conjunctivitis and ocular discharge, ulcerations of the lips, tongue, gums, or nasal planum, hypersalivation, coughing, fever, lethargy, and inappetence [95]. In severe cases, secondary bacterial pneumonia may also be present.


Most often, if FRDC is suspected, the causative agent need not be determined because supportive treatment is similar among pathogens [95]. Determining the etiology in patients with lower respiratory tract disease has been previously discussed in this chapter. If a definitive diagnosis is desired, PCR testing, bacterial cultures, serologic assays, and viral isolation have been used, but false‐negatives and false‐positives commonly occur [95].

Anesthetic Management

Patients with FRDC may be at an increased risk of respiratory complications, particularly those with excessive ocular–nasal discharge and nasal congestion. Those with systemic signs of infection undergoing elective procedures are also at considerable risk for perioperative morbidity. Elective procedures should be delayed until clinical signs of disease have resolved. In humans, it has been suggested that anesthesia should not commence for at least 4–6 weeks following URIs to allow for adequate healing of the airways [99]. Anesthetic management should be focused around maintaining adequate hydration and reducing oronasal secretions and handling of the infected airway. Adverse events such as bronchospasm, laryngospasm, airway obstruction, and hypoxia may occur during the perianesthetic period [99]. Airway management is a priority; additional endotracheal tubes, airway equipment, and induction drugs should always be readily available before, during, and after the anesthetic event. Following extubation, hypoxia is alleviated with supplemental oxygen. Equipment should be disinfected as with all infectious diseases and was previously discussed.

Gastrointestinal Infections

Infectious Gastroenteritis

Infectious gastroenteritis (GE) may be of viral, bacterial, or parasitic origins. Common viral etiologies include canine parvovirus, canine coronaviral enteritis, feline parvoviral enteritis, feline immunodeficiency virus (FIV)‐associated diarrhea, and feline leukemia virus (FeLV)‐associated panleukopenia. Campylobacter jejuni, Salmonella spp., Helicobacter spp., Clostridium perfringens, and Clostridium difficile are among some causes of bacterial GE. Except for parvovirus, FIV, FeLV, and parasites, the primary pathogen is rarely confirmed; however, clinical features and management are similar among pathogens.

Clinical Signs

Vomiting and diarrhea are the two most common clinical signs associated with infectious GE. Vomiting is a result of the activation of the emetic center in the reticular formation of the medulla oblongata, the chemoreceptor trigger zone located in the fourth ventricle of the brain, and the stimulation of receptors and autonomic nervous system within the abdominal viscera [100]. Reduced intestinal absorption and/or increased intestinal secretion results in an accumulation of fecal water leading to diarrhea [100]. Infectious agents compromise the GI mucosa by blunting the absorptive properties of the villi or disrupting the ion pumps in the GI epithelium [100]. A thorough history and physical examination should be performed in patients with GE. The anesthetist should look for historical indications (chronic vomiting and diarrhea) and clinical signs associated with dehydration (Table 16.6) [100]. Skin tenting should be performed over the dorsum of the neck or lateral thorax [101]. Fever, depression, anorexia, and abdominal pain are also associated with infectious GE.


Patients exhibiting mild clinical signs of GE typically do not require extensive diagnostic tests. In patients with moderate to severe illness, parasites can be identified using centrifugal flotation with zinc sulfate flotation solution and direct fecal examination, fecal enzyme‐linked immunosorbent assay (ELISA) for canine parvovirus, and serologic analysis for FeLV and FIV. Gastric biopsy from the body, fundus, and antrum of the stomach may be helpful in diagnosing Helicobacter in cats; however, this requires general anesthesia.

Table 16.6 Subjective parameters used to assess the degree of dehydration.

Source: Adapted with permission from: Tello and Perez‐Freytes [100].

Estimated degree of dehydration Clinical signs
<5% (subclinical) Nonapparent on physical examination
5% (mild) Tacky or dry mucous membranes (MM)
6–8% (moderate) Dry MM
Decreased skin elasticity
Normal pulse quality
Normal arterial blood pressure
8–10% (severe) Dry MM
Further decrease in skin elasticity
Small increase in capillary refill time (CRT)
Weak pulses
Approximately 12% (hypovolemia) Dry MM
Increased CRT
Skin tent does not return to normal
Tachycardia or bradycardia
Weak to absent pulses
Altered mentation
Cold extremities

Anesthetic Management

Dehydration from GI losses must be assessed and treated prior to general anesthesia. Appropriate fluid therapy is dependent on the patient’s history, physical examination, and diagnostic tests (complete blood count, serum biochemistry, and acid–base status). General anesthetics negatively impact the cardiovascular system, and it is best to volume‐resuscitate and stabilize the patient prior to anesthesia. This may require delaying anesthesia and surgery if possible. A balanced crystalloid solution (lactated Ringer’s solution [LRS], Plasma‐Lyte, Normosol‐R, etc.) should be administered intravenously, with the dose and rate dependent upon the extent of dehydration and colloidal oncotic pressure or total protein. In severe cases, colloids can be administered at doses of 3 ml kg−1 and 5 ml kg−1 over 15 min to cats and dogs, respectively. Care should be taken to not exceed approximately 20 ml kg−1 day−1 of these solutions, as colloids have been reported to interfere with the function of platelets, von Willebrand’s factor, and factor VIII in critically ill patients [102]. Vomiting is routinely associated with hyponatremia, hypochloremia, and hypokalemia. Diarrhea also contributes to hypokalemia via the loss of potassium ions in the feces. Potassium chloride (KCl) may be added to a replacement isotonic crystalloid solution to correct hypokalemia (Table 16.7) [100]. The rate of potassium administration should not exceed 0.5 mEq kg−1 h−1. The anesthetist should be cautious when bolusing fluids containing potassium chloride. If bradycardia or electrocardiogram changes associated with hyperkalemia are observed, the bolus should be discontinued, and patient’s serum potassium concentrations evaluated. Elevations in lactate, packed cell volume (PCV), and total protein may also be good laboratory indicators for dehydration. Blood pressure should be assessed prior to and during general anesthesia. Low blood pressure (mean arterial pressure <70 mmHg) may indicate hypovolemia and warrants immediate fluid therapy prior to induction of anesthesia. Alternatively, pulse pressure variation (PPV) and plethysmographic variability index (PVI) can be useful for identifying a patient’s response to volume replacement with increases in stroke index and cardiac index [103]. Briefly, cyclic oscillations in the amplitude of the plethysmographic waveform recorded by pulse oximetry during positive pressure ventilation are a result of decreased venous return (Figure 16.6). This is often associated with hypovolemia and additional fluid therapy is recommended. A metabolic acidosis routinely accompanies patients with GI disease and dehydration. This can be corrected with a balanced crystalloid solution such as LRS or Plasma‐Lyte. The crystalloid solution 0.9% sodium chloride is considered more acidic (pH is approximately 5.0) than other balanced solutions (LRS, pH is approximately 6.5) and is not recommended for resuscitation in patients with a metabolic acidosis.

Table 16.7 Recommended amount of potassium chloride and rate of infusion so as not to exceed 0.5 mEq kg−1 h−1.

Source: Adapted with permission from: Tello and Perez‐Freytes [100].

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Oct 18, 2022 | Posted by in SUGERY, ORTHOPEDICS & ANESTHESIA | Comments Off on Infectious Disease

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Serum potassium concentration (mEq l−1)