Raphaë1 Vézina Audette1 and Stephen D. Cole2 1 Portland Veterinary Emergency and Specialty Care, Portland, Maine, USA 2 Clinical Infectious Disease Laboratory, Department of Pathobiology, School of Veterinary Medicine, University of Pennsylvania, Philadelphia, Pennsylvania, USA Veterinary patients that undergo anesthesia face a genuine risk of infection, even when well‐established preventative measures are precisely executed. Postoperative infections in domestic animals (dogs, cats, and horses) may manifest in various ways ranging from skin and soft tissue infections involving the surgical site, to osteomyelitis, pneumonia, urinary tract infection, and catheter‐associated complications. Postoperative infections often result from a combination of several factors, including patient immunosuppression, disease, concomitant drug administration, breach of aseptic technique, prolonged procedures, prolonged recumbency, insufficient infection prevention measures, and postoperative surgical site management. The use of antimicrobial drugs perioperatively can impact host susceptibility to nosocomial infections but must be weighed against the risks of antibiotic resistance. This chapter explores the relationship between general anesthesia and perioperative infections and discusses the vital role anesthetists play in the prevention of nosocomial infections. Skin and soft tissue infections at incision sites are the most common postsurgical infections among all veterinary species. Mild infections are easily managed but severe infections can progress to abscessation, cellulitis, and systemic disease or compromise surgical wounds leading to dehiscence. Bacteria of the genus Staphylococcus are the most common bacteria associated with incisional infections [1,2]. Staphylococcus pseudintermedius, which is part of the normal skin and oral cavity flora of dogs and cats, is the most common species associated with small animal incisional infections. Infection caused by this organism may originate from the patient itself or contaminated fomites. In contrast, Staphylococcus aureus is not considered normal flora of most dogs and cats (although it is more common in cats than dogs) and infections caused by S. aureus are most likely from an anthropogenic source (e.g., surgeon, anesthetist, and care staff) as it commonly constitutes a part of the cutaneous and respiratory flora in humans [3]. In horses, methicillin‐resistant S. aureus (MRSA) postsurgical infection is much more common as a result of horse‐adapted strains [4]. Enteric organisms (i.e., Escherichia coli and Klebsiella pneumoniae) have been reported to be more common in incisional infections following abdominal surgery [5]. Rates of postsurgical infection vary between procedures, facilities, and surgeons. Table 7.1 illustrates the variability of reported rates among common procedures for dogs, cats, and horses. Surgical wounds can be characterized as clean, clean‐contaminated, contaminated, or dirty (infected) (see Box 7.1 for definitions). Infection rates for procedures characterized as clean may be lower than those characterized as contaminated or dirty. Table 7.1 Wound classifications and associated infection rates in dogs and horses. All veterinarians must practice infection prevention to uphold their professional oath to protect animal health and welfare and to promote public health. The American Animal Hospital Association (AAHA) has described a hierarchy of controls for veterinary facilities, and this is shown in an adapted illustration (Fig. 7.1) [18]. The hierarchy ranks the following types of control (from most to least effective): elimination controls, engineering controls, administrative controls, and personal protective equipment (PPE). Elimination controls are the most effective, but often the most difficult to implement. Elimination controls remove or prevent the entry of a threat of infection (i.e., a pathogen) completely. Engineering controls are aspects of hospital design and setup that remove hazards or improve compliance with infection prevention procedures. Administrative controls are the policies and procedures focused on infection prevention. PPE includes the use of specialized attire or equipment to limit the spread of pathogens and is considered the least effective of these controls because it is the most dependent on human factors such as compliance and education. However, PPE is still a critical piece of an infection prevention program, but it cannot be the sole tool. Specific examples of controls relevant to the veterinary anesthesia team are highlighted in Fig. 7.1. Figure 7.1 Hierarchy of infection controls. PPE, personal protective equipment; SOP, standard operating procedures. The success of an infection control program is dependent on the people participating in it. It is critical that veterinary anesthetists are champions of a facility’s efforts to implement and use infection prevention strategies. Beyond the delivery of anesthesia and analgesia and ensuring a safe recovery from anesthesia, the anesthesia team is concerned with preoperative patient evaluation, including the identification of factors that may impact patient outcomes. It is fundamental to have a thorough understanding of factors that may influence patient safety and anesthetic risk, including nosocomial infections, so that risks can be mitigated, and patient safety improved [19]. Training is key to success but should not stand alone without some form of audit and feedback. For example, personnel responsible for reprocessing equipment should be trained at least annually with updates on procedures conducted when necessary. A review process should also be in place. Audit and feedback should never be punitive but viewed as an opportunity to demonstrate proficiency and identify areas for improvement. The preoperative patient preparation area and operating rooms (ORs) are high‐task‐density environments where omission of any critical steps risks severely compromising patient outcomes. Successfully getting a patient through surgery requires the meticulous orchestration of completion of each task in the correct fashion and order. Carrying out a wide variety of procedures, each with specific steps that must be remembered and executed properly, is paramount to ensure patient safety. Observance of evidence‐based perioperative guidelines, particularly antimicrobial use, has been described in veterinary medicine albeit in a limited number of publications [20,21]. The use of surgical checklists in human hospitals has been adopted globally and has contributed to a significant improvement in perioperative outcomes [22]. In veterinary medicine, the implementation of surgical checklists has also been studied, specifically relating to critical outcomes, including infection and mortality. In one study carried out in a private practice referral center, the implementation of a surgical checklist reduced the risk of incisional infections in dogs undergoing exploratory laparotomy for gastrointestinal foreign body obstructions [11]. Another study from a different academic institution determined that surgical checklist implementation decreased surgical complications such as missed perioperative antibiotic doses and operations at the wrong surgical site, decreased the rate of unexpected return to the OR, and improved implementation of safety measures; however, it did not affect morbidity or mortality rates or the incidence of incisional infections during the study [23]. Frequent hand hygiene is the single most important action that an individual can take to protect their patients’ and their own health. Wearing gloves is never a substitute for hand hygiene. For hands without visible soiling, the use of alcohol‐based hand sanitizers is considered an effective approach. Important exceptions include when working with patients known to be infected with non‐enveloped viruses (e.g., canine parvovirus and equine rotavirus), fungal pathogens (e.g., dermatophytes), or spore‐forming bacteria (e.g., Clostridium spp.). Handwashing with soap and water is the best strategy in cases of these pathogens or for people with visibly soiled hands. Box 7.2 describes best practices for the execution of hand hygiene. Auditing of hand hygiene practices allows for the identification of barriers to implementing hand hygiene (e.g., too few hand sanitizer stations) and it has been shown to be effective at improving compliance in veterinary facilities. For anesthetists that do not work within the sterile field, the use of routine hand hygiene strategies in combination with clean, non‐sterile gloves for most procedures suffices. Handwashing or disinfection should always be performed before and after putting on gloves. Specialized and extensive hand hygiene, colloquially described as “scrubbing in,” should be performed by all surgeons and other professionals whose hands will enter a sterile field during a surgical procedure. Outside of the surgical suite the anesthetist should wear clean scrubs in patient care areas. Long hair should be tied back, and stethoscopes and lanyards should not dangle as they can act as fomites. Personnel should change into and out of scrubs when entering and leaving the facility, respectively, and always change when visibly soiled. Clean laboratory coats or water‐resistant aprons are appropriate for protecting scrubs when performing procedures where extensive direct contact with animals (e.g., during physical restraint or when clipping hair) or biological materials will occur. Laboratory coats should never be worn outside of the hospital or inside a surgical suite. It is best practice to have work‐dedicated footwear. In cases of animals with zoonotic or highly transmissible diseases, additional PPE including coveralls, gowns, face shields/masks, and shoe covers (“booties”) should be considered on a case‐by‐case basis. Inside the surgical suite, additional PPE is needed in addition to clean scrubs. For individuals not entering the sterile field, bouffant/surgical caps, face masks, non‐sterile gowns, non‐sterile gloves, and shoe covers have all been recommended. Surgeons and others entering the sterile field should use sterile gowns and gloves; in some cases, two pairs of gloves are worn (“double gloved”). Protective eyewear should be used particularly for contaminated and dirty (infected) wounds and for procedures involving irrigation. Sterilization and disinfection are both processes used to allow for the safe reuse of medical devices and equipment (also known as “reprocessing”). Cleaning refers to the act of removing visible soilage from the surface of an object and is a critical first step that must be completed prior to disinfection or sterilization. Disinfection reduces or eliminates most (if not all) pathogenic microbes on the surfaces of inanimate objects with the important exception of bacterial or fungal spores. Sterilization, however, kills all microbial life, including spores. Whether a device needs to be cleaned and either disinfected or sterilized prior to reuse is based on the Spaulding classification system, which classifies devices as critical, semicritical, or noncritical [24]. Table 7.2 defines these classifications and provides examples of devices used by anesthetists or surgeons. Reprocessing instructions for medical devices can be found within the manufacturer’s instructions for use (MIFU). Table 7.2 Spaulding’s classification of medical devices. Disinfection is primarily achieved with the use of liquid chemicals called “disinfectants.” Disinfectants can be further categorized as low‐level (kills vegetative bacteria, some viruses, and some fungi), intermediate‐level (kills vegetative bacteria, most viruses, and most fungi and mycobacteria), or high‐level (kills all microbial life, except for large numbers of spores). Some high‐level disinfectants (e.g., 2% glutaraldehyde) can be considered chemical sterilants if used with prolonged exposure times (3–12 h). The efficacy of disinfectants is dependent on the removal of all organic materials prior to use along with appropriate wet contact time and proper dilution (if required). The MIFU should always be followed exactly when using a disinfectant and any deviation may lead to incomplete efficacy. Certain devices such as thermometers and ultrasound probes can be used with specially designed plastic covers, which may decrease their need for high‐level disinfection following use. Some facilities designate certain devices (i.e., thermometers) that would need frequent disinfection for use with a single patient throughout their stay before discarding or sending home with the patient. Sterilization is achieved by the use of an autoclave (steam with high heat under pressure), dry heat sterilizer, or ethylene oxide gas. Hydrogen peroxide liquid plasma and chemical sterilants may also be used. Sterility assurance monitoring (SAM) should be performed to ensure adequate processes are in place. A good SAM program always includes adequate documentation of results. SAM should include a multimodal approach to testing that includes biological indicator testing, chemical indicator testing, and mechanical monitoring. Biological indicator tests are the only tests that assure sterilization by assessing the killing of spore‐forming organisms (e.g., Bacillus spp. or Geobacillus spp.) and are typically performed weekly. Chemical indicator testing involves chemical changes when the correct physical conditions are met that lead to a color change in an indicator (e.g., lead carbonate to lead (II) oxide). Chemical indicator tests should be included for each package of instruments to be sterilized (external indicators such as autoclave tape can also be used). Mechanical monitoring should follow guidelines from the sterilizer’s manufacturer but often includes checking a variety of parameters on instrument gauges, screens, or printouts. Chemical indicator testing and mechanical monitoring do not ensure sterilization was achieved, but they can be used to detect procedural errors such as incorrect packaging or overloading. Common reasons for failure of sterilization include incorrect operation, incorrect settings, improper loading or overloading, and excessive packing. Certain medical devices are labeled as “single‐use,” which implies they cannot be reused [24]. Laryngoscopes are considered an essential instrument to perform safe and atraumatic orotracheal intubation in cats and dogs [25]. Laryngoscopes, because they are expected to encounter a patient’s mucosal surface, are considered semicritical devices according to Spaulding’s classification and thus require, at a minimum, high‐level disinfection between uses in different patients. Though no reports of hospital‐acquired infections involving laryngoscopes exist in companion animals, disinfection protocols should be followed. Anecdotal reports of an epidemic of respiratory infectious disease in horses associated with the use of a mouth rinse plunger to flush the mouth prior to orotracheal intubation illustrate the necessity to ensure high‐level decontamination in semicritical items. In one North American study, respiratory pathogens such as equine herpes virus 1 and 4 (EHV‐1, EHV‐4), equine influenza virus (EIV), equine rhinitis B virus (ERBV), and/or MRSA were detected in 22% of healthy horses presented for routine dental care [26]. These pathogens can be reasonably expected to contaminate the tip‐of‐mouth rinse plungers or bite blocks and orotracheal tubes. Endotracheal tubes (ETTs) and laryngeal masks offer ways to maintain airway patency in veterinary patients during general anesthesia or deep sedation. Additionally, they are indispensable for the protection of a patient’s airways against the potentially lethal risk of aspiration pneumonia. In 2018, an outbreak of carbapenem‐resistant E. coli was linked to reuse of ETTs at a veterinary teaching hospital [27]. Most commercially available ETTs are labeled as single‐use only. In the context of veterinary anesthesia, it may not be economically feasible to use a new ETT for each patient at all facilities. However, since ETTs encounter a patient’s mucosal surface, they are deemed semicritical devices according to Spaulding’s classification, and thus require high‐level disinfection. While these devices are widely reused in the veterinary field [28], best practices for sterilization or disinfection have not been established. Crawford and Weese studied the efficacy of various disinfection strategies for ETTs against in vitro inoculated Streptococcus zooepidemicus and Bordetella bronchiseptica [29]. Although soaking in chlorhexidine gluconate or accelerated hydrogen peroxide solutions significantly reduced growth on direct and enrichment cultures compared to water or triclosan‐containing soap solutions, high‐level disinfection was not achieved with either method. A recent study evaluated the efficacy of four different protocols for ETT disinfection and found no difference in bacterial growth following cleaning of the ETT with either a water scrub, detergent scrub, detergent scrub and chlorhexidine gluconate soak, or detergent scrub and bleach soak all of which yielded significant bacterial growth [28]. Reprocessing single‐use items may affect the integrity and efficacy of the device. For example, a study of ETT sterilization demonstrated that the physical integrity of the ETT cuff may become compromised following glutaraldehyde or ethylene oxide sterilization. If a facility chooses to reuse ETTs or other single‐use devices against the MIFU, then they must include these factors in their risk assessment for the patient. Minimum standard monitoring equipment for patients under general anesthesia in veterinary medicine includes electrocardiography (ECG), pulse oximetry, capnography, and temperature probes [25]. Many different configurations exist for each instrument and the anatomical site on which they are used on patients will dictate the need for levels of disinfection or sterilization. For example, though rarely used in everyday practice, intravascular thermistors for measurement of core body temperature would be considered a critical device and thus require sterilization before use, whereas esophageal thermometers are considered semicritical and may only require high‐level decontamination between patients. If rectal thermometers are in use, appropriate hygiene measures such as the use of plastic sleeves and high‐level disinfection between patients are recommended. For this reason, instruments that do not represent an increased level of invasiveness may be preferred; for example, surface ECG monitors require low‐level decontamination of leads applied to intact skin, compared to esophageal ECG leads, which require a higher level of decontamination. Catheters used for vascular cannulation are considered critical items. As such, they must only be used when sterile. Prior to the placement of vascular catheters, sterile preparation of the placement site by removing hair with clippers followed by the use of an antiseptic solution can prevent infectious complications [30,31]. For placement of peripheral intravenous (IV) catheters, a clean technique may suffice as long as appropriate hand hygiene is observed [32]. The current trend in human medicine is to remove IV catheters only when clinical signs of inflammation are observed. Commonly reported risk factors for infectious catheter complications in humans include the type of catheter used, blood sampling through the catheter, the type of IV infusate administered, the duration the catheter was in place, and the catheter location. A recent prospective study evaluated these proposed risk factors in cats and dogs in a private referral practice intensive care unit and did not identify any of them as being associated with an increased risk of a positive bacterial culture from the catheter tip [33]. In cats and dogs, one study suggested that a dwell time greater than 72 h was not associated with an increased risk of IV catheter contamination [34], whereas others reported increased rates of infections after a period of as little as 36 h [35]. An association between lack of experience and risk of catheter infection has been reported in dogs [36]. Concurrent corticosteroid therapy and the presence of phlebitis were also positively associated with a positive catheter tip culture [36]. A prospective evaluation of microbial contamination of anesthesia breathing circuits used in the context of canine anesthesia demonstrated that in the absence of a biofilm, the environmental contaminants that colonize the breathing circuits are likely of low pathogenicity and short viability [37]. Evidence from simulation experiments supports the notion that anesthesia machines can be contaminated by bacterial pathogens, and that heat and moisture exchange filters can effectively prevent bacterial contamination [38]. Further, they demonstrated that volatile anesthetics (halothane) and carbon dioxide absorbents (soda lime) did not provide any demonstrable bactericidal action. Uncertainty remains about whether viral pathogens can contaminate an anesthesia breathing circuit after use in a sick animal shedding respiratory viruses, which would then be transferred to subsequent patients. Factors involved in the potential transfer of pathogens from corrugated tubing to a patient’s airways are ill‐defined. Bacterial filters may not be efficient against viral pathogens and, as such, it is recommended to discard breathing circuits when a risk for viral cross‐contamination exists. As part of the presurgical evaluation, close attention should be paid to the skin. If there is evidence of cutaneous infections (e.g., bacterial pyoderma and dermatophytosis) therapy should be initiated and the surgery rescheduled after clinical resolution. For emergent surgeries, antimicrobial therapy should encompass the suspected pathogens. A swab for bacterial culture and susceptibility can be collected for submission prior to antibiotic administration to guide postoperative care. Skin cannot be sterilized or disinfected but antiseptic chemicals can be used to decrease the overall microbial burden of the skin. While protocols may vary, the effective and efficient preparation of the surgical site is a critical part of the aseptic technique. It has been demonstrated that prolonged time from clipping to incision is associated with an increased incidence of postsurgical infection [39,40]. After induction of anesthesia, hair on the surgery site should be clipped with electric clippers leaving a sufficient border around the intended incision site. Use of a size 10 blade over a size 40 blade has been advocated as it results in less irritation and lower numbers of colony‐forming units (CFUs) [31]. Shaving should not be done as it is traumatic to the skin and is associated with higher rates of infection. Care should be used to avoid cross‐contamination of patients with clipper blades. The next step after clipping is to remove gross organic material from the skin (cleaning) with a surgical scrub containing a detergent. Gauze swabs are used to scrub the skin in a concentric pattern away from the intended surgical site. Following an initial scrub, surgical sites are then rinsed with 70% alcohol, which doubles as an additional bacterial kill and removes the detergent. The final step is to use a surgical antiseptic (e.g., chlorhexidine gluconate or povidone‐iodine). Box 7.3 addresses the preparation of the epidural injection site.
7
Infection Prevention and Control in Anesthesia
Introduction
Postoperative infections
Procedure
Wound classification
Dogs
Horses
References
Ovariohysterectomy
Clean
1.3–5.7%
NA
[6]
Castration
Clean
2.0–3.4%
2.2–3.6%
[6–8]
Enterotomy or gastrotomy
Clean‐contaminated
7.0–19.9%
19–40.4%
[5,9–11]
Orthopedic surgery
Clean
5.4–6.6%
3.7–14.2%
[12–15]
Cystotomy
Clean‐contaminated
0.0–3.0%
NA
[16,17]
Infection control and prevention
Important tools and steps for perianesthetic infection control
Perioperative checklists
Hand hygiene
Attire
Reprocessing of anesthetic and surgical equipment
Class
Definition
Example(s)
Sterilization versus disinfection
Critical
Devices that contact or enter sterile tissues or bloodstream
Surgical instruments (i.e., forceps and scalpels), implants, biopsy tools
Sterilization
Semicritical
Devices that come into contact with mucous membranes but do not enter sterile tissues or the blood stream
Laryngoscope blades, endoscopes
Sterilization or high‐level disinfection
Noncritical
Devices that come into contact with patient skin but not mucous membranes. Some devices that may not come in direct contact with patients may also require disinfection between patients
Laryngoscope handles, stethoscopes, blood pressure cuffs
Low‐ to intermediate‐level disinfection
Laryngoscopes and mouth rinse plungers
Endotracheal tubes
Monitoring equipment
Intravenous catheters
Breathing circuits and ventilators
Preparation of the surgical site and maintenance of the sterile field

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