Surgical Facility Location
Vendors Commonly Used by Comparative Medicine
All survival surgery will be performed using aseptic techniques, including surgical gloves, masks, sterile instruments, and aseptic techniques. Animal Welfare Act regulations.
Appropriate attention to pre-surgical planning, personnel training, aseptic and surgical technique, animal well-being, and physiological status during all phases of protocol will enhance the outcome of surgery. Guide for the Care and Use of Laboratory Animals.
Before beginning a surgical procedure there should be a meeting of the surgical team to develop a surgical plan. Depending on the complexity of the procedure, the surgical team should consist at least of the following: a surgeon, anesthetist, veterinarian, surgical technicians, animal care staff, and investigator. For very complex procedures there may be more than one surgeon and a sterile and non-sterile assistants.
Regardless of the nature of the surgery, the surgical plan should identify: personnel involved, their roles and training, type of operative procedure, equipment and supplies needed, identify location of operating room, preoperative health assessment, intra-operative monitoring, operative technique, post-operative care, need for antibiotics, and a mechanism for keeping all relevant records.
Depending on their roles, personnel should have adequate training in appropriate surgical technique including but not limited to asepsis, gentle tissue handling, minimal dissection of tissue, appropriate use of instruments, effective hemostasis, correct use of suture materials and patterns.
- Sterilization kills all microorganism (life), achieved by physical or chemical means
- Disinfection destroys vegetative bacteria
- Antiseptic prevents or arrests growth of microorganisms by destruction or inhibition on skin or mucus membranes
- Asepsis is freedom from infection or prevention of contact with microorganisms
A dedicated surgical facility is required for large animals e.g. primates, dogs, cats, rabbits. Regardless of the location, when an area is being used for surgery no other activity should take place. In general the surgical facility should have the following components: a) animal preparation room, b) instrument preparation room, c) surgeon preparation room, d) holding and recovery room and e) an operating room.
Lighting in the operating room should be bright, focused and non-glare. Lighting should be sufficient to perform the procedures but should not be make the operating area hot.
Animal preparation e.g. clipping of hair, scrubbing and anesthesia induction should not be performed in the operating room but rather in the animal preparation room. The surfaces in the room including floors, walls and ceiling must be non porous, sealed, durable and sanitizable. The area should be maintained clean and free from clutter and access limited to people involved in the procedures being performed. A schedule for cleaning and sanitizing the rooms should be maintained and adhered to. Rooms should be cleaned “daily” i.e. at the end of the procedures on that day by damp dusting flat surfaces, lights and operating room furniture. Clean all organic debris with soap and disinfect after each surgery. Wet vacuum or mop the floors at the end of the day. Mops should be laundered after use or soaked in germicidal solution. Clean the scrub sinks and soap dispensers, and empty waste buckets. A more though cleaning should be performed “weekly” including wiping down walls, ceilings, cabinets and equipment with germicidal cleaning solution after removing organic debris and soiling.
Examples of common hard surface disinfectants for cabinets, surgery tables, lights, etc:
- Alcohols (70% ethyl alcohol, 85% isopropyl alcohol) for 15 min in absence of organic mater or gross contamination.
- Quaternary ammonium compounds (e.g. Roccal®, Quatricide®) are rapidly inactivated by organic mater and may support growth of gram negative bacteria.
- Aldehydes e.g. glutaraldehyde (e.g. Cidex®, Cide Wipes®, Cetylcide-G®) rapidly disinfects surfaces. Toxic. Follow OSHA exposure limits.
- Phenolics (e.g. Lysol®, TBQ®) less affected by organic materials than other disinfectants.
- Sodium hypochlorite (e.g. Clorox 10% solution) corrosive, activity reduced by organic matter.
- Chlorine dioxide (e.g. Clidox®, Alcide®) kills vegetative organisms within 3 min, corrosive, activity reduced by organic matter, must be made fresh.
- Chlorhexidine (e.g. Novalsan®, Hibiclens®) rapidly bactericidal and persistent also effective against many viruses, active in the presence of blood.
- Always follow manufacturers’ recommendations.
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Clean all instruments and materials prior to sterilization. Dirt, blood and gross contaminants should be completely removed as they compromise sterilization. Always soak instruments in water with detergent immediately after surgery to remove blood and debris, and to facilitate cleaning. Instruments can be washed manually with a brush or in a washer sterilizer. In certain cases it may be necessary to use an ultrasonic cleaner “for difficult to clean” instruments.
Rinse instruments thoroughly after washing to remove any residues from the cleaning agents. Dry and safely store the instruments after cleaning.
Dry instruments and supplies should be loosely packed in standard or special packs in order of use. Take special care to avoid damaging delicate instruments. Packs should be dated. Sterilization indicators e.g. autoclave tapes or test cultures should always be included. Note that autoclave tapes only indicate that the surface reached the required temperature.
Packs should be stored in an appropriate manner after sterilization. Wrapped and sterilized packs are good for 6 months if properly stored. Storing packs in sealed in plastic bags prolongs their shelf life.
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It is critical that you start your surgery with sterile instruments. Instruments may be sterilized by physical or chemical means. Steam sterilization in an autoclave (121°C for 15 min or 131°C for 3 min) is extremely effective. Autoclaves should not be used for temperature sensitive instruments. Some corrosion may occur and sharp instruments may be dulled. Packs should not be removed from the autoclave until they are completely dry. Sterilization failure may occur if residual air is not evacuated during the operation of the autoclave. Dry heat in a chamber is a good alternative to a steam autoclave. It is non-corrosive and penetrates most materials and closed containers. Dry heat does not dull instruments, but the high temperatures attained may damage some materials. Instruments must be cooled before contacting tissues. Other physical means of sterilization include gamma irradiation primarily for heat or moisture sensitive items such as syringes, suture materials, scalpel blades and medical devices.
- Too tight packs
- Improperly loaded autoclave
- Defective autoclave
- Insufficient temperature and pressure
- Too short exposure time
- Sterilization is the complete reduction of microbial life, which may be accomplished by physical (e.g. heat, radiation) or chemicals means.
- Sterilants are essentially the same as sporocides. They kill all microorganisms including bacterial endospores. A sporocidal product kills all microorganisms including bacterial endospores.
- Disinfectants, on the other hand, kill 100% of vegetative (actively growing) bacteria (of certain species) under conditions specified by the Environmental Protective Agency, but are not efficacious against fungi, viruses, Mycobacterium tuberculosis or bacterial spores. These agents are only effective if used according the manufacturers instruction and may be inactivated by organic matter such as blood, body fluids or tissues.
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A variety of chemicals can be used to sterilize heat sensitive materials. All chemicals must be rinsed from the instruments using sterile saline or sterile water to avoid tissue damage. Often long contact times are required for sterilization. Organic materials may decrease effectiveness of these chemicals. All surfaces must be exposed and tubing must be filled with the solution. Effectiveness is dependent on adequate contact time with the instruments, proper mixing, age of the solution, and absence of organic material from the instruments. Most chemical sterilants have to be “activated” in order to be effective. Follow manufacturers instructions and avoid mixing incompatible compounds, and remember that most of these chemicals are hazardous agents. Disinfectants should not be used as sterilants. Residual chemicals must be rinsed off before using the instruments.
Examples of common chemical sterilants include:
- 2% Glutaraldehyde for 10 hours (Cidex®, Abcocide®). Shelf life 14-28 days after activation depending on type.
- 8% Formaldehyde plus 70% alcohol 18 hours.
- 7% stabilized hydrogen peroxides 8 hours (Accelerated Hydrogen Peroxide®, Virox STF®, Sporox®). Shelf life 21 days.
- 7.35% hydrogen peroxide and 0.23% peracetic acid 3 hours (EndoSpor® plus). Shelf life 14 days.
- Chlorine dioxide 1:5 solution 6 hours. Must be mixed daily (Clidox®).
- 1.37% Sodium hypochlorite 6 hours. Shelf life 14 days after activation (Alcide®).
- Always follow manufacturers’ recommendations.
Ethylene oxide gas sterilizes instruments in 3-7 hours when applied at 21-60°C at 40-60% relative humidity using specialized equipment. Ethylene oxide is flammable, explosive, toxic, and very irritating to tissues. All instruments must be aerated for 8-12 hours at 50-60°C or 1-7 days at room temperature.
- Physical methods – thermocouples placed with load
- Chemical methods – packed within load or autoclave tape
- Biological methods – bacterial spores, used once a week
- Bacillus stearothermophilus for steam autoclaves
- Bacillus subtilis for ethylene oxide
- Micrococcus radiodurans for gamma radiation
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Prior to scrubbing hands, the surgeon should don a surgical cap, facemasks, surgical scrubs and appropriate shoes. Remove jewelry items and ensure that fingernails are trimmed short.
Scrubbing should be thorough beginning at the tip of the fingers all the way to the elbows using a surgical scrub containing a germicide e.g. chlorhexidine. Vigor and exposure times are critical, 3-15 minutes or 5-20 brush strokes per surface, with at least two thorough scrubs and rinses.
At the end of the scrub dry your hands with a sterile towel beginning at the tip of the fingers to the elbow. Rotate the towel and repeat the procedure on the other hand.
After drying the hands, proceed to put on a sterile gown. Lift the gown, unfold away from the table, and insert your arms into the sleeves. The assistant closes the back of the gown, and the surgeon closes the waist tie.
Open the paper covering (outer covering should have been previously opened) on the gloves as illustrated. Insert the gloves as shown making sure not to touch any non-sterile surfaces. If you accidentally touch a non-sterile surface with your gown or gloves discard them and re-gown and/or re-glove as appropriate.
Always maintain a zone of sterility in front of you. Clasp your hands in front of you making sure the hands are above the table, above your waist and no higher than your shoulders.
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Prior to surgery it is important that the subjects are properly identified. Obtain the weight, age, sex, colony history, and health status. Determine whether the animals have been acclimatized to the facility, generally 3-5 days rest after arriving from the vendor should be sufficient. In some instances this period may need to be up to two weeks.
Perform a physical examination to determine if the animal is healthy. If possible perform a complete blood count, blood chemistry and urine analysis.
If indicated, bathe the animal the day before surgery. Withhold food for 6-12 hours before surgery. Apply ophthalmic ointment to the eyes following induction of anesthesia to prevent corneal drying. Provide airway support, vascular access and provide pre-emptive analgesia as indicated.
Always prepare an area approximately twice the surgical area you will need. Preparation should take place in the animal preparation room. Hair should be removed from the surgical site using clippers with #40 blade or a depilatory cream (depilatory creams can irritate the skin, so rinse the area thoroughly after using the cream) followed by a surgical scrub alternating between disinfectant (e.g. iodophors or chlorhexidine) and alcohol. Iodophors (e.g. Betadine®, Prepodyne®, Wescodyne®) inactivate a wide range of microbes but their activity is reduced in the presence of organic mater. Chlorhexidine (e.g. Novalsan®, Hibiclens®) are rapidly bactericidal, persistent and active against many viruses. They are active even in the presence of blood.
A gauze sponge can be used for scrubbing. During the scrub, the process should begin along the incision line and extend outward and never from outward (dirty) towards the center (clean). Do not go over the incision site with the same scrub.
At the end of the scrub apply a coat of germicide. Reapply the germicide in the operating room.
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The decision to drape depends on the nature of the procedure being done. For extensive procedures it is necessary to drape, using towels, stockinettes or plastic wraps. Drapes help to maintain a sterile field and preserve body heat. While drapes play an important role in reducing contamination of the surgical site, faulty technique may increase contamination. Drapes must cover the animal and table.
Anesthesia alters thermoregulation and reduces metabolism. Loss of heat also occurs from open body cavity and evaporation of body fluids. Loss of heat can significantly prolong the duration of anesthetic, which in turn increases the risk of complications. Animals should always be kept warm with a hot water blanket.
Animals can experience extensive fluid loss during surgery. Fluid loss occurs primarily as a result of evaporation from body cavities and due to blood loss. Reduce intra-operative fluid loss by irrigating the operative field with warmed sterile saline, and by administration of warm, sterile isotonic fluids parenteraly during the surgery. Control blood loss during surgery by cauterizing or ligating potential bleeders. Monitor water and food intake, body condition and animal weight post-surgically.
Operating Room Conduct
- Reduce contamination in the environment
- Gentle tissue handling
- Minimal tissue handling
- Effective hemostasis
- Correct suture material and techniques
- Appropriate use of instruments
- Adequate monitoring of the animal
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Animals should be monitored every 15 minutes to ensure that the animal is alive and doing well. It is critical that assessment not completely rely on instrumentation to monitor the animals. Check the color of the mucus membranes, response to reflexes, heart and pulmonary functions. Instruments can be used to monitor level of oxygenation, expired carbon dioxide, electrocardiogram, electroencephalogram and temperature.
Select the correct surgical instruments for the procedure to be performed. Surgical instruments should be handled to minimize contamination for example placing on sterile drape, segregation according to function helps insure sterility for example instruments used on the skin should not be used within the abdominal cavity.
Tissues should be handled gently avoiding unnecessary trauma or drying out. Only minimal dissection with appropriate instruments should be done. Blood vessels that are likely to bleed should be ligated. Avoid contamination of incisions sites.
Wounds should be closed with appropriate suture material and techniques using the right kind of needles. Non-cutting (atraumatic) taper point or round needles have no cutting edges and should be used for soft tissue like peritoneum, intestines, kidney etc. Cutting or reverse cutting needles provide a cutting edge through dense, difficult to penetrate tissues like skin.
In general absorbable sutures (e.g. cat gut, vicryl®, dexon®) should be used for soft tissues. Blood vessels should be ligated with slowly absorbable (e.g. vicryl®, dexon®, PDS®, maxon®) or non-absorbable sutures (e.g. nylon, silk). Non-absorbable sutures (e.g. ethilon®, prolene®, dermalon®), surgical glue or stainless steel wound clips and staples should be used for the skin. Good surgical techniques will prevent post-surgical complications like infection, hemorrhage or even death. Proper surgical and post-surgical records should be maintained.
Non-absorbable suture materials used to close skin wounds should be removed as soon as the wound is healed (7-10 days) or within two weeks, whichever occurs first.
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Administer warmed sterile isotonic fluids and keep the animals warm using hot water blankets, hot water bottle or heat lamp (avoid burns). Animals should be checked frequently preferably every 10-15 minutes, and turned from side to side until recovered. Monitor recovery from anesthesia closely and be prepared to provide respiratory support. In the pre-surgical planning phase you should have discussed with the veterinarian anticipated outcomes, for example how long it will take the animal to recover from anesthesia post-operatively, what to expect and what not to expect. Is there bleeding from the incision site? Is the color of the animal’s mucus membranes pink (good) or is it bluish (bad)? Do you anticipate lameness e.g. after orthopedic surgery or not?
Monitor food and water intake after recovery from anesthesia and provide nutritional support. Is the incision site swollen? Is there discharge from the site? Swelling, discharge and discoloration of the incision site signals the need for veterinary attention. Does the animal have a fever? If the animal has a fever consult a veterinarian so treatment plan can be initiated. Administer analgesic and check for signs of discomfort or pain. The principal investigator is responsible for ensuring that post-procedural care is provided as described in the approved animal use protocol. The post-procedural monitoring and care plan should be developed in consultation with the veterinary staff. The surgical plan should include when animal is expected to return to normal behavior. If there is any question as to whether or not the animal is doing well consult a veterinarian immediately.
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Samuel Perkins Co. Inc., 497 Beale Street, Quincy, MA. firstname.lastname@example.org
Barber Vet Supply, 1-800-552-5698
AJ Buck & Sons, 1-800-638-8672
Penn Vet Supply, 1-800-548-4490
Hospital Pharmacy, 410-955-6591
Clinical Engineering Services, 114 Brady Building 410-955-5639
- Anesthesia and analgesia in laboratory animals. Kohn DF, Wixson SK, White WJ, Benson GJ (eds.) Academic Press. San Diego. 1997
- Laboratory animal anesthesia. Flecknell PF. Academic Press. London. 1996.
- Formulary for laboratory animals. 2nd Edition. Hawk CT, Leary SL. Iowa State University Press, Ames. 1999.
- Lumb & Jones’ Veterinary Anesthesia. Thurmon JC, Tranquilli WJ, Benson GJ (eds.). Williams & Wilkins, Baltimore. 1996.
- Pain Management in Animals. Flecknell P and Waterman-Pearson A (eds.). W.B. Saunders, London. 2000.
- Principles of Surgical Technique. The Art of Surgery. 2nd Edition. Wind G.G. and Rich N.M. Urban & Schwarzenberg, Baltimore-Munich. 1987.
- Animal Phsiologial Surgery. Lang C.M. Springer-Verlag, New York. 1976.
- Fundamentals Techniques in Veterinary Surgery. Knetch C.D. Saunders, Philadelphia. 1987.
- Textbook of Small Animal Surgery. 2nd. Edition. Slatter D.H. W.B. Saunders. 1993.
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