by Tory Hampshire, DVM, NINDS and Judy Davis, DVM, NINDS Future articles will discuss new techniques for ventilation, anesthesia, and drug delivery systems for rodents. For more info, e-mail davis or call her at 301-496-9354.

Hot Mouse Tips: A Three-Part Series
PART 1. IMPROVING THE MOUSE PATIENT:
PERIOPERATIVE TIPS FOR BETTER OUTCOMES

Now that the dawn of mouse phenotyping has arrived, so, too, have more complicated procedures. More often than not, scientists struggle with the issue of anesthesia because of outdated equipment or technique. In this first of a three-part series, we offer you hot methods and tips that may shake up your assumptions and expedite your results inside the mouse facility!

Before You Start

It is important to overcome the tendency to focus on the procedure while being less attendant to the patient’s physiological status. Aggressive preparation is critical in small animals–particularly when one is doing simultaneous procedures or back-to-back procedures.

First, the effects of anesthesia on rodent hormones and metabolism have not been fully evaluated. What is known is that any procedure that will produce pain, distress, hypothermia, hypo-volemia, dehydration, hypoglycemia, acid-base disturbance, infection, or adrenocortical stress (surgery and anesthesia are big ones) will affect your results in a bigger way than will staving off these adverse effects.

Therefore, consider:

1. Preanesthetic Techniques. Rodents, and mice in particular, have high metabolic rates and small surface areas. Compared with larger animals, they generally require higher anesthetic dosages to achieve an effective level of anesthesia, and the duration is typically shorter. They are also less likely to survive respiratory arrest from overdosage.

Metabolic rate also influences the onset of hypothermia and dehydration from exposed membranes. And even a small amount of surgical blood loss in a mouse may represent a substantial percentage of total blood volume.

Solutions:

Plan to make provisions for warmth under and over your rodent patient. Heating pads, heat lamps, or even pocket warmers
can provide these sources. Accidental burning can be averted by covering the warming pads with polar fleece.

Two Ways to Warm a Mouse: (above) an isolated, warmed holding area (Thermocare, Inc., Incline Village, NV, $945–$1060) accommodates several rodents at a time and can receive oxygen through any side port, or (below) a hand-sewn triple-layer of polar fleece can hold a standard single-use pocket warmer in one pocket and a mouse in the other.

Assume that your patient will not return to normal drinking patterns immediately and will experience some dehydration. Warmed Lactated Ringers Solution at 60–70 mL/kg/day bolused under the skin before surgery (just after induction of anesthesia) will provide maintenance hydration and help to insulate a mouse during the procedure.

Drug effects are dose-dependent; an accurate weight is critical! We see this as the most common cause of anesthetic over- and underdosing. Daily weighing postoperatively will also aid in evaluating hydration status and general well-being. Purchase a gram scale and use it daily from the day before surgery to three to five days after. You will be amazed at how much weight rodents lose 12 hours after surgery. Up to 20 percent of body weight can be lost due to decreased fluid intake and increased loss of fluids during surgery.

2. Respiration. Most of the time inhalant anesthetic is administered by mask to the mouse because intubation is a challenge given the mouse’s small mouth opening, large incisors, large immobile tongue, and large cheek folds. Because rodents are burrowing animals, they have very compliant rib cages. In contrast to larger animals, their functional residual air capacity approximates their vital capacity. When anesthetized, the frequency of respiration decreases while tidal volume remains low. Therefore, minute ventilation is normally maintained with high frequency, low tidal volume, which renders the animal susceptible to respiratory acidosis and hypoxemia. Other considerations include a rodent’s propensity to hold its breath and stress-induced catecholamine release.

Solutions:

If you are stuck on injectable anesthesia cocktails, try Dopram (doxapram hydrochloride), a CNS respiratory stimulant, as part of your preanesthetic regimen (one drop, full strength on the tongue).

Enlist the help of your veterinary section and try to gain access to isoflurane anesthesia and scavenging (downdraft table or hood). Masks made with syringe cases work well. Rodent masks can also be purchased from most veterinary suppliers.

For procedures likely to last more than 45 minutes, invest in a ventilator and learn to intubate your mouse or rat (see "part 2" in our series).

3. Perioperative Stress and Pain. Diazepam is not effective as a sedative, muscle relaxant, or tranquilizer in most rodents. Alpha-agonists (xylazine, medetomidine) have the potential of undesirable side effects: Sedation often outlasts analgesia, and bradycardia, hypothermia, hyperglycemia, and respiratory depression are not uncommon. Alpha-agonists can be countered, however (consult your veterinarian for reversal agents), and therein lies a strong advantage for their use. Whatever analgesic you choose, you should consider providing it preemptively before noxious stimuli allow pain perception.

Preoperative medications include anticholinergics, anxiolytics, analgesics, respiratory stimulants, and anti-inflammatory drugs. The use of anticholinergics in rodents is controversial. Although they protect against vagal-mediated bradycardia, they increase the viscosity of airway secretions and the potential for obstruction of small airways or the trachea. If you use anticholinergics, you should ventilate the animal vigorously; if you don’t use them, you need to monitor heart rate closely.

Solutions:

To prevent bradycardia, atropine 0.04 mg/kg subcutaneously (SQ) or intramuscularly (IM) 10 minutes before anesthetic induction may be used in concert with Dopram as the respiratory stimulant.

For narcotic analgesia: buprenorphine hydrochloride 10–20 µg/kg may be given IM or SQ at onset of immobility and every 6–8 hours thereafter.

To reduce pain from local irritants such as stereotactic apparatus, instill lidocaine 2 percent gel in the ear canal and on the ear bars at onset of immobility.

For procedures expected to cause severe swelling or visceral pain, Banamine (flunixin) 1 mg/kg SQ may be given twice daily starting at immobility onset [Lactated Ringers fluid and an H2 blocking agent such as Zantac (ranitidine) 0.5 mg/kg IM or Pepcid (famotidine) 2.5 mg/kg SQ are also recommended to protect against renal and gastrointestinal damage, respectively].

To prevent drying of the cornea (keratitis sicca), use a petroleum-based artificial tear ointment. Keratitis sicca is a frequent side effect of anesthesia and is exacerbated by the irritating effect on the eye of anesthesia delivered via face mask. Medications can be obtained from the ORS/VRP pharmacy at 435-2780.

4. Anesthesia Induction. The most consistent and reliable anesthetic protocols for rodent neonates and adults couple inhalants with vigilant monitoring. Those still using Metofane (methoxy flurane) in a bell jar, think again! Metofane provides little benefit and high risks because of its poor liquid-to-gas coefficient. Rodents are usually either too deep or too light to facilitate safe surgery.

Anesthetic responses to predetermined amounts of injectable anesthetic range from inadequate to death. If you oversee technicians who perform your anesthetic procedures using agents such as barbiturates [Nembutal (pentobarbital sodium) or Pentothal (thiopental sodium)], get them into the habit of using a standard worksheet. We ask our technicians to weigh animals the day before surgery and organize their individual doses for each mouse or rat in small, identified Ziploc bags. Injectable anesthetics have a low margin of safety, often fail to mute peripheral reflexes, and are associated with protracted recovery. If injectables are used, you should monitor heart rate, respiration, and body temperature until the animal is actively moving around the cage. It is common to mistake sternal positioning for "recovery" and to place the animal back in the animal room, which is probably (hopefully) colder than the recovery area; the animal then becomes hypothermic–the metabolic rate falls, and residual anesthesia effectively anesthetizes the animal. Unfortunately, rodents lack the physiological capability to overcome these events, and the outcome can be catastrophic for investigators.

This small Plexiglas box and inhalant anesthesia system work for rapid induction and recovery under isoflurane gas anesthesia. Here it is used under a hood, but the system also works nicely over a down-draft surface, provided the operator guards against a chilled mouse bottom.

Solutions:

If you use the alpha-agonist xylazine, you can use the reversal agent yohimbine (Yobine) 0.25–0.5 mg/kg intravenously to shorten recovery periods. If you are not expert at tail vein injections, you can give this drug IM using a 25-gauge or smaller needle.

Use of a small Plexiglas box is preferred for inhalant anesthesia induction. Once the animal is anesthetized, it is removed from the chamber and switched to a face mask or a multiple-mouse port manifold for maintenance (if you need up to six animals anesthetized at once). This achieves the benefit of using oxygen flow over liquid anesthetic to produce the plane of anesthesia desired. Recovery from inhalation anesthesia is also very rapid. As a tip, consider oxygenation (100 percent) for 5 minutes prior to induction and on recovery to ensure that hypercapnia does not develop. Isoflurane with a calibrated vaporizer is the method of choice in terms of patient safety and lack of environmental contamination (details on vaporizers and components next article).

5. Anesthesia Maintenance. Solutions:

Facilitate good depth of anesthesia with injectable combinations like ketamine-xylazine cocktails by using a topical local anesthetic whenever possible along the incision line—0.1–0.2 mL of 2 percent mepivacaine or lidocaine under the skin at the surgical site will enhance analgesia and better facilitate immobility. It will also lower the needed anesthetic dose.

Whenever possible, use nonrebreathing systems with oxygen flow at least three times the minute ventilation (10 mL/kg or about 0.2 mL/mouse) to lower CO2.

Take precautions against patient cooling, which is increased with high oxygen flow rates. Because the mean alveolar concentration of anesthetic (MAC) needed falls with falling body temperature, you must lower your anesthetic concentration to prevent anesthetic overdosage if your mouse chills. Monitor body temperature, and adjust MAC accordingly.

Surgical procedures invite pH disturbance. Remember: Rodents normally maintain minute ventilation by high respiratory rates and low tidal volumes. During anesthesia, both tidal volume and minute ventilation fall; therefore, if you don’t ventilate, you risk respiratory acidosis with or without hypoxemia.

If you anticipate long procedures (greater than 45 minutes), learn to ventilate your rat or mouse. Special machines can be purchased from several suppliers to achieve high-frequency ventilation in small tidal volumes for rodents (details next article).

6. Recovery Support. High metabolic rate, a high ratio of surface area to weight, and high oxygen flow rate lead to a faster rate of cooling in rodents than in larger animals. It is critical that they are kept warm until they are fully ambulatory. If you have a means of monitoring temperature, all the better! pH disturbances are also common and can be kept minimal by shortening anesthesia time; reliable pH monitoring is now possible in rats but not yet in mice.

Dedicated rodent thermometers can be purchased (Physiotemp, Clifton, NJ) with rectal probes for more accurate temperature assessment.

Solutions:

Consider purchasing a pulse oximeter to measure saturated oxygen concentration. This equipment has reasonable efficacy in the rat when body temperature is conserved to the tail. In mice, however, blood flow in the tail, foot, and tongue is not great enough to yield reliable readings.

Environmental space room temperature should be kept with the mouse in mind, not the surgeon. Warm water-circulating blankets, warm lavage intraoperatively, plastic wrap for insulation, pocket warmers, or used pediatric isolation units make good provisions for warmth. Keep a small rodent recovery room or area that can be separated from the standard mouse or rat room. The isolette shown above is also useful for this purpose if you do not have a separate room. Some companies are now engineering mouse racks with thermal elements so that individual rows can be warmed.

Remember that your mouse or rat patient may not feel well enough to eat or drink for the next few days. Add fresh fruit, Jell-O cubes, or extra doses of subcutaneous fluid boluses to your postoperative regime.

Work with your institute veterinary staff to develop pain scoring for your mouse or rat based on subjective and objective indicators. This will help in making analgesic redosing decisions.

You should worry about infections if your procedure is long and complications occur. You can run a complete blood count (Clinical Center lab 496-3386) by obtaining an orbital blood sample in a heparinized hematocrit tube. Your institute vet can also prescribe a prophylactic antibiotic for your patient.

Institution of as many of these tips for periprocedural care of the mouse and rat will result in less wasted time, greater survival rates, and, best of all, comfortable animals. Common use of these materials and procedures can’t be anything other than a win-win situation in the world of animal research and research support.

High-Calorie Mouse Jell-O -2 cups boiling water -1 pkg. raspberry-flavor Jell-O -60 mLStat-VME (VRP Pharmacy) -20 mLPediasure (VRP Pharmacy) -2 scoops Designer Protein (GNC Health Food Store) Blend well and refrigerate in ice-cube trays. Serve one-quarter cube per mouse per day.

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