Assessing Changes in Volatile General Anesthetic Sensitivity of Mice after Local or Systemic Pharmacological Intervention

Summary

Loss of the righting reflex has long served as a standard behavioral surrogate for unconsciousness, also called hypnosis, in laboratory animals. Alterations in volatile anesthetic sensitivity caused by pharmacological interventions can be detected with a carefully controlled high-throughput assessment system, which may be adapted for delivery of any inhaled therapeutic.

Abstract

One desirable endpoint of general anesthesia is the state of unconsciousness, also known as hypnosis. Defining the hypnotic state in animals is less straightforward than it is in human patients. A widely used behavioral surrogate for hypnosis in rodents is the loss of righting reflex (LORR), or the point at which the animal no longer responds to their innate instinct to avoid the vulnerability of dorsal recumbency. We have developed a system to assess LORR in 24 mice simultaneously while carefully controlling for potential confounds, including temperature fluctuations and varying gas flows. These chambers permit reliable assessment of anesthetic sensitivity as measured by latency to return of the righting reflex (RORR) following a fixed anesthetic exposure. Alternatively, using stepwise increases (or decreases) in anesthetic concentration, the chambers also enable determination of a population's sensitivity to induction (or emergence) as measured by EC₅₀ and Hill slope. Finally, the controlled environmental chambers described here can be adapted for a variety of alternative uses, including inhaled delivery of other drugs, toxicology studies, and simultaneous real-time monitoring of vital signs.

Introduction

General anesthetics are defined by their ability to cause a reversible state of hypnosis in a wide variety of species, yet an explanation as to how such a diverse class of drugs can all elicit a singular endpoint remains elusive. A number of theories have been posited over the years, starting with the Meyer-Overton correlation between anesthetic potency and lipid solubility, which suggested general membrane disruptions as the basis for hypnosis^1,2. More recent evidence suggests that protein targets affecting neuronal signaling contribute to anesthetic effects. Mice have proven to be an indispensable model for exploring these theories because of the homology between murine and human anesthetic responsiveness. Though a mouse cannot be asked about its subjective awareness under general anesthesia, certain primitive reflexes serve as useful surrogate measures of rodent hypnosis. In the first few days following birth, mice develop a reflexive righting response that prevents them from being passively placed in a supine position³. The dose of anesthesia at which a mouse loses its righting reflex correlates well with human hypnotic doses⁴.

Assessment of loss of righting reflex (LORR) has become a widely used laboratory standard for testing anesthetic sensitivity in mice as well as a variety of other species including rat, guinea pig, rabbit, ferret, sheep, and dog^5-8. The dose of a given anesthetic at which LORR will occur for members of a species is extremely consistent, but it can be shifted significantly by environmental factors. For example, sleep-deprived rats are more sensitive to both volatile and intravenous anesthetics⁹ and rats with high aerobic capacity are less sensitive to isoflurane¹⁰. Hypothermia has also been shown to decrease the dose of numerous anesthetics required for hypnosis in a large spectrum of species^11-14. In order to reliably identify the anesthetic dose at which LORR occurs in a group of experimental animals, it is critical that the assessment environment be carefully controlled to minimize stress, maintain euthermia, and deliver equal amounts of drug to all subjects. Not surprisingly, genetic factors are also known to alter anesthetic sensitivity^15-18. Consequently, careful consideration should also be given to controlling for genetic background¹⁹.

We have developed an apparatus that ensures identical gaseous anesthetic delivery to each of 24 mice while maintaining a constant 37 ^oC environment. The transparent cylindrical design of our exposure chambers allows for fast LORR assessment and easy integration of telemetric physiological measurements. This system has been shown to accurately measure isoflurane, halothane, and sevoflurane induction EC₅₀ and time to emergence in wild-type mice²⁰. We have also used this system to observe changes in anesthetic sensitivity in mice with genetic mutations and targeted hypothalamic lesions^21-23. Here we describe two ways in which anesthetic sensitivity may be assessed after a pharmacological intervention using our controlled environment apparatus. Steady-state phenotyping of volatile anesthetic induction and emergence sensitivity requires 8-10 hours and is consequently best tailored for studies in which experimental conditions do not change, such as in chronic or long-acting pharmacological interventions. However, for short-acting treatments whose effects dissipate significantly over time we also present a simple procedure to evaluate changes in righting reflex following stereotactically-targeted microinjections or intravenous drug treatments that significantly impact anesthetic emergence. These tests represent a small subset of the potential applications for this controlled environment system, which could be adapted for any number of subjects of a variety of species to receive any type of inhaled therapeutic.

Protocol

All procedures involving animals outlined herein have been approved by the University of Pennsylvania's Institutional Animal Care and Use Committee.

1. Overview of the Testing Apparatus

1. The testing apparatus consists of 24 clear acrylic cylindrical chambers 10 cm in length and 5 cm in diameter (total volume of 200 ml). This size is appropriate for a typical 25 g adult mouse. Chambers have ports at each end for gas inlet and outlet. The outlet end is removable so that animals may be easily loaded into the chamber. Gas port openings are carefully sealed with Teflon tape, while rubber o-ring gaskets are used to seal the removable end of the cylindrical chambers.
2. Each chamber is mounted on a rack that sits inside a water bath. The rack is fitted so that only the lower portion of the chambers (below the gas inlets) is submerged. For stability, the back end of the chamber rests on a support so that the entire chamber sits horizontally. This ensures even contact of the entire chamber with the bath.
3. Polyethylene tubing connects an oxygen tank to an anesthetic vaporizer, and then passes through a 10 L/min flow meter. The tubing splits into 25 small-diameter resistors of equal length to ensure equal flow is delivered to each of the 24 chambers and to an agent analyzer.
4. Vacuum lines exit each chamber at the opposite end of the gas inlet. This promotes unidirectional flow that eliminates rebreathing of exhaled carbon dioxide. The vacuum lines combine at a manifold to connect to an in-house suction line. A pop-off valve along the main vacuum line ensures atmospheric pressure conditions within each chamber.
5. The bath is filled with enough water to fully contact the bottom of each chamber. The water is circulated through the bath and maintained at a constant 37 ^oC by a pump.

2. Check the System Prior to Exposure

1. Check that the temperature of the water bath is 37 ^oC throughout the bath.
2. Flow oxygen at a rate of 5 L/min (200 ml/min per chamber + agent analyzer). Submerge each chamber under water and look for bubbles or entry of water into the chamber, both of which are indicative of leaks. Seal any leaks before beginning the experiment.
3. For each chamber, connect a 500 ml/min flow meter in line after the chamber to make sure that flows are balanced across each of the 25 gas lines. This ensures that the input 5 L/min flows will be distributed evenly so that each chamber receives 200 ml/min flow. Any chamber not receiving the expected flow should have its inflow and outflow tubing checked for obstructions.
4. Calibrate the agent analyzer to ensure a reading of 0.00% isoflurane when 100% oxygen is flowing.

3. Implant Temperature Transponder

1. One week prior to habituation, anesthetize each mouse with 2% isoflurane.
2. Sterilize the dorsal neck area with betadine.
3. Inject a temperature transponder subcutaneously between the shoulder blades using the sterile, prepackaged injector needle.
4. Monitor the injection site daily for infection and migration of the transponder.

4. Habituate Animals to Testing Chambers

1. Four days before the first assessment, place all mice into individual chambers for 2 hr with 100% oxygen flowing.
2. Repeat step 4.1 daily for the four days prior to assessment to avoid the confounding effects of stress due to a new environment.

5. Perform the Pharmacological Intervention that You Wish to Test for Effects on Anesthetic Sensitivity

1. This intervention may be a stereotaxic injection into a specific part of the brain²⁴, an intravenous or intraperitoneal injection²⁵, or delivery of a drug to a specific brain area via cannula²⁶.
2. Because these procedures themselves may change anesthetic sensitivity compared to a naïve animal, a proper control group should undergo the same procedure with vehicle injections.
3. Ensure that the pharmacological intervention has an appropriately long duration of action if you are planning to do a stepwise increasing and/or decreasing determination of anesthetic sensitivity as shown in step 6 below; otherwise, skip to step 7.

6. Assess Anesthetic Sensitivity using the Stepwise EC₅₀ Determination for Induction and Emergence

1. Place each animal into individual chambers with 100% oxygen flowing.
2. Set the isoflurane concentration to 0.4%* for 15 min. During the last 2 min of this period, assess each animal's righting reflex by gently rolling the chamber until the mouse is placed on its back. The righting reflex is considered to be intact if and only if the mouse is able to restore all of its paws to the floor of the chamber within 2 min.
   1. *Note that 0.4% isoflurane is a subhypnotic dose in C57BL/6J mice. If any mice lose their righting reflex at the first step, the initial dose was too large and should be reduced on subsequent days.
3. Record the state of righting reflex for each mouse and scan each mouse for temperature data. A template record is shown in Table 1.
4. Increase the isoflurane concentration by ~0.05% for 15 min and repeat step 7.2. Continue to do this until all animals have lost their righting reflex.
5. Optional: repeat the same procedure for decreasing step-wise isoflurane doses until all animals have regained their righting reflex (see step 6.3).
6. To end the experiment, turn off the isoflurane and flush the entire system with 100% oxygen for 15 min. This will help to prevent hypoxia as the mice recover before being returned to their home cages and will protect the experimenter from any anesthetic exposure.
7. Optional: if the number of animals or the number of anesthetic concentrations are limited due to resource or time constraints, the curve-fit parameter estimates-particularly the Hill slope-may have underappreciated, falsely low error estimate. In such cases, it may be necessary to repeat the anesthetic sensitivity measurement described in steps 6.1-6.6 on up to two additional experimental days to fully obtain the true Hill slope's parameter and its corresponding error estimation.

7. Assess Short-term Changes in Anesthetic Sensitivity with Time to Emergence

1. Place each animal into individual chambers with 100% oxygen flowing.
2. Set the isoflurane concentration to 1.2%, which corresponds to the induction ED₉₉ for wild-type C57BL/6J mice²⁰. Maintain for 30-60 min depending upon the expected duration of action of the acute intervention.
3. Confirm LORR in all animals by gently rolling each chamber until mice are placed on their backs.
4. Turn off isoflurane and flow 100% oxygen. Measure the time until each animal regains its righting reflex. This is defined by placement of all four paws on the floor of the chamber and confirmed by the presence of three consecutive tests with an intact righting reflex.

Results

Figure 1 demonstrates the utility of the step-wise LORR assay for determining long-term effects of a pharmacological intervention. Ibotenic acid (IBA) is an agonist of the glutamatergic N-methyl-D-asparate (NMDA) receptor that is often used as an excitotoxin to cause permanent neuronal lesions. Here we injected 10 nl of 1% IBA bilaterally into the ventrolateral preoptic area (VLPO) of C57BL/6J mice one week prior to testing. The majority of neurons in this nucleus exhibit low rates of firing during wakef...

Discussion

Though assessment of LORR in a single mouse is a seemingly straightforward task, it is nevertheless essential to maintain identical physiological conditions between subjects in order to collect reliable data from a group of animals. The tightly regulated, high-capacity LORR apparatus presented here offers a way to standardize experiments and maximize efficiency. By following the basic tenets of thermoregulation and equal flow distribution, this system can be easily recreated and customized to fit individual experimenters...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
Name of the Reagent	Company	Catalogue Number	Comments
Oxygen	Airgas	OX300
Isoflurane	Butler Schein		Any volatile anesthetic of interest may be substituted
Name of Material	Company	Catalogue Number	Comments
Mass flow meter- 10 SLPM	Omega Engineering	FMA-A2309
Mass flow meter- 500 SCCM	Omega Engineering	FMA-A2305
Anesthetic agent analyzer/gas indicator	AM Bickford	FI-21 Riken
Heating water pump	Fisher Scientific	13-874-175
Temperature transponders	BMDS	IPTT-300
RF temperature reader	BMDS	DAS-6007