Systematic Assessment of Well-Being in Mice for Procedures Using General Anesthesia

Summary

We developed a protocol to assess well-being in mice during procedures using general anesthesia. A series of behavioral parameters indicating levels of well-being as well as glucocorticoid metabolites were analyzed. The protocol can serve as a general aid to estimate the degree of severity in a scientific, animal-centered manner.

Abstract

In keeping with the 3R Principle (Replacement, Reduction, Refinement) developed by Russel and Burch, scientific research should use alternatives to animal experimentation whenever possible. When there is no alternative to animal experimentation, the total number of laboratory animals used should be the minimum needed to obtain valuable data. Moreover, appropriate refinement measures should be applied to minimize pain, suffering, and distress accompanying the experimental procedure. The categories used to classify the degree of pain, suffering, and distress are non-recovery, mild, moderate, or severe (EU Directive 2010/63). To determine which categories apply in individual cases, it is crucial to use scientifically sound tools.

The well-being-assessment protocol presented here is designed for procedures during which general anesthesia is used. The protocol focuses on home cage activity, the Mouse Grimace Scale, and luxury behaviors such as burrowing and nest building behavior as indicators of well-being. It also uses the free exploratory paradigm for trait anxiety-related behavior. Fecal corticosterone metabolites as indicators of acute stress are measured over the 24-h post-anesthetic period.

The protocol provides scientifically solid information on the well-being of mice following general anesthesia. Due to its simplicity, the protocol can easily be adapted and integrated in a planned study. Although it does not provide a scale to classify distress in categories according to the EU Directive 2010/63, it can help researchers estimate the degree of severity of a procedure using scientifically sound data. It provides a way to improve the assessment of well-being in a scientific, animal-centered manner.

Introduction

EU Directive 2010/63¹ stipulates that the 3R Principle (Replacement, Reduction, Refinement) developed by Russel and Burch² is to be applied whenever animal experimentation is necessary. The ultimate goal of the EU Directive is to phase out all animal testing, but the Directive acknowledges that, for the time being, some animal experiments are still needed to conduct research that will protect human and animal health. Thus, if an animal experiment cannot be replaced by any alternative method, only the minimum number of laboratory animals is to be used to obtain reliable results. In addition, the amount of pain, suffering, and distress accompanying experimental procedures should be minimized using appropriate refinement measures. EU Directive 2010/63 stipulates that the severity of a procedure must be prospectively classified as non-recovery, mild, moderate, or severe¹. As severity classification is decided on a case-by-case basis, it is important to have scientifically sound tools to estimate the severity of a given procedure.

Score sheets as proposed by Morton and Griffith³ are an essential tool in detecting any deviations from normal status, including negative effects on well-being⁴. Score sheets are used to retrospectively determine pain, suffering, and distress caused by an experiment and focus on visible changes in the physical state of the individual animal (e.g., body weight, fur, gait). Although, Annex VIII of EU Directive 2010/63 provides examples of each severity category, researchers still lack tools to estimate the degree of severity of a given procedure using scientifically based data.

The absence of indicators showing negative well-being is not the only way to determine the state of the animal; the presence of indicators pointing to positive well-being is also important^5,6,7,8. For example, animals display luxury behaviors like burrowing and nest building behavior only when all their essential needs are met. If well-being is reduced, luxury behaviors are the first to decline^5,7. Protocols to be used in assessing well-being should include indicators pointing to the physical, physiological/biochemical, and psychological states of animals in order to evaluate their well-being in a detailed and comprehensive manner⁹.

Within the context of refinement, a protocol was developed to meet these requirements and to assess the effects of a procedure involving general anesthesia on well-being of mice¹⁰. At the same time, the goal was to minimize any additional stress to enable the easy integration of the protocol into a given experiment. The protocol considers burrowing behavior, home cage behavior such as activity, food intake, and nesting, and trait anxiety-related behavior. In addition, it includes the Mouse Grimace Scale (MGS), and the non-invasive analysis of corticosterone metabolites in feces. The protocol is designed to facilitate the assessment of well-being in a scientific and animal-centered manner and to provide information on well-being that supports the classification of the degree of severity. In addition to score sheets, it can provide useful information for the severity classification of a procedure. As the protocol is easy to carry out and does not require extensive equipment, it can be integrated into an ongoing experiment without influencing the results of a study. It should be noted that the Animal Research: Reporting of In Vivo Experiments (ARRIVE) guideline¹¹ is to be observed in all studies involving animal experiments, with the goal of improving design, analysis, and reporting.

Protocol

The study was performed according to the guidelines laid down by the German Animal Welfare Act and was approved by the Berlin State Authority ("Landesamt für Gesundheit und Soziales", permit number: G0053/15).

NOTE: The main objective of this protocol was to investigate the effect of repeated anesthesia on glucocorticoid metabolites. A sample size calculation was performed to determine the number of animals to be used: n ≥ 2 × (s/µ₁- µ₂)² × (z_α + z_β)². µ₁- µ₂ is the difference between population means at which power and sample size calculations are made (α = 5%, β = 80%); z_α = 1.96 and z_β = 0.84 are the quantiles of the standard normal distribution. Figure 1 illustrates the time line of this protocol. If a parameter of the protocol shows a difference with the control level, the animal should be closely monitored, and the parameter should be measured again after a suitable period. For example, if trait anxiety-related behavior is increased, this behavior should be tested again a week later, in order to help determine the period until full recovery. Time points and periods defined in this protocol can be adapted for use with other procedures. When changing time points, habituation periods should be kept as described in the protocol. In order to reduce factors that might influence the mice's behavior, tests requiring more manipulation should be conducted after tests that do not disturb the normal behavior of mice. Figure 2 summarizes all tests of the protocol using a summary scoring sheet. Figure 3 provides simplified scales of the grade of well-being, which give an overview of how to interpret the test results.

1. Habituating mice to handling by experimenter

1. Allow mice to habituate to the animal facility for at least 2 weeks after they have been obtained from another facility or vendor.
2. House mice in groups and maintain them under standard conditions (room temperature 22 ± 2 °C; relative humidity 55 ± 10%) on a light:dark cycle of 12:12 h.
3. Provide all groups with a tunnel and cotton nestlets as standard enrichment, and provide food and water ad libitum.
4. Habituate all mice to the tunnel and/or cup handling at least a week prior to testing¹².
   NOTE: Picking up mice by the tail can induce stress or anxiety, which in turn affects well-being and also has an impact on the results of this protocol¹².

2. Preparing the behavioral testing room and apparatuses

NOTE: Provide a separate room for testing, ideally near the room where the animals are kept. Transport the mice in their home cages to the testing room at least 60 min before the procedure is conducted. If possible, conduct all tests of this protocol in the same testing room where the procedure is carried out.

1. Prepare an observation cage to test burrowing behavior⁸ and to take the photographs for use in the MGS¹³ (Figure 4).
   1. Use a glass box with a floor area of approximately 220 mm × 290 mm and a height of 390 mm.
   2. Cover the floor of this box with approximately 0.5 cm of bedding material.
   3. Scatter a handful of used bedding material from the home cage on top of the new bedding material to reduce distress caused by the new environment.
   4. Provide food, the same kind that is normally supplied as diet, and water.
      NOTE:If possible, use water bottles, because mice may fill water bowls with bedding material .
2. Prepare a cage (type III: 420 mm × 260 mm × 150 mm) for the 24-h observation period, for which mice are housed individually (Figure 5).
   NOTE: In order to minimize the duration of individual housing, collect data for nest building behavior, home cage activity, food intake, and fecal corticosterone metabolites (FCM) during this period.
   1. Place new bedding material in the cage (approximately 0.5 cm deep) and scatter a handful of used bedding material without feces from the home cage on top of the new material, in order to reduce distress.
   2. Provide a standardized square cotton nestlet of a defined weight, as environmental enrichment only (see table of materials)¹⁴.
      NOTE: Commercial nestlets might differ in weight. Therefore, we modified the weight of the nestlet described by Deacon and used 2.0 g instead of 2.7 g¹⁴.
   3. Mount the infrared sensor on the top of the cage, when using an infrared sensor to measure home cage activity (see table of materials).
   4. Provide food, the same kind that is normally supplied as diet, and water ad libitum.

3. Mouse Grimace Scale

NOTE: Photographs for the MGS are taken in the observation cage at three time points: (i) 2 days prior to the procedure to record baseline MGS levels, (ii) 30 min after the procedure, and (iii) 150 min after the procedure. When well-being is impaired, scores on the MGS increase. If increased MGS scores are still observed after 150 min, take additional photographs at a later stage.

1. Use a high-definition camera for photography.
2. Gently transfer the mouse into the observation cage and allow the mouse to habituate to the new environment for at least 30 min.
3. Continuously take about 30 - 40 photographs for each time point within 1 - 2 min.
4. Sort all photographs by selecting the sharp frontal or lateral photographs and discarding blurry photographs or photographs that show mouse faces from other perspectives than frontal or lateral view.
5. Randomly select one photograph from each time point, (i.e. 2 days prior to the procedure, 30 min after the procedure, and 150 min after the procedure) for each mouse.
6. Crop the photographs to display only the head of the mouse so that the body position is not visible¹³.
7. Create a spreadsheet file with one sheet for each photograph and add a table including the five facial action units of the MGS to each sheet.
   NOTE: The file contains baseline photographs as well as photographs post procedure.
8. Randomize the order of the sheets.
9. Present the file on a computer screen to three independent persons, who were previously trained to use the MGS developed by Langford et al. and have them score the facial action units using a 3-point-scale (0 = not present, 1 = moderately present, 2 = obviously present).
   NOTE: Scoring is based on following parameters¹³: Orbital tightening ("narrowing of the orbital area, with a tightly closed eyelid or an eye squeeze"); nose bulge ("rounded extension of skin visible on the bridge of the nose"); cheek bulge ("convex appearance of the cheek muscle"); ear position ("ears pulled apart and back from their baseline position or featuring vertical ridges that form owing to tips of ears being drawn back"); whisker change ("movement of whiskers from their baseline position either backward, against the face or forward, as if standing on end; whiskers may also clump together").
10. Analyze scores, as follows (adapted from Langford et al.¹³).
    1. Average all facial action units for each photograph to generate the MGS score.
       NOTE: If one of the facial action units could not be scored, average the remaining facial action units.
    2. Subtract the mean for the baseline photographs from the mean for the photographs post procedure to obtain a MGS difference score for each mouse.
    3. Test for differences in the MGS difference scores between the persons (nonparametric test for related samples).
       NOTE: If there is a significant difference (p < 0.05), determine whether scores of all photographs or only scores of a few photographs differ between the persons. If the latter is true, repeat scoring of these photographs. Otherwise, the persons should repeat the MGS training and then score the photographs again.
    4. Average the MGS difference scores obtained from the different scorers for each mouse, if results of all persons do not significantly differ.
    5. Use a nonparametric statistical test to compare the MGS difference scores averaged between the study groups.

4. Burrowing behavior^8,15,16

1. Prepare burrows by placing 140 ± 2 g food pellets normally supplied as diet in a standard opaque plastic water bottle (250 mL, 150 mm length, 55 mm diameter, 45 mm diameter of bottle neck)⁸.
   NOTE: As mice prefer wide tubes, burrows with a diameter of 68 mm can be used as described by Deacon¹⁶.
2. Place the burrow filled with food pellets in the home cage 5 days prior to the procedure for acclimatization.
   NOTE: The regular food-dispensing unit in the cage should not be emptied but should also remain filled with food pellets, as mice are used to this.
3. Carry out the test twice, 2 days prior to the procedure (baseline); carry out the last 30 min post procedure as well.
   1. Let the mouse habituate for at least 30 min to the observation cage where photographs for the MGS were taken.
   2. Place the plastic water bottle filled with food pellets parallel to the back wall of the observation cage.
   3. Weigh food pellets (g) remaining in the burrow after 2 h.
4. Calculate the weight of food pellets removed from the burrow by mice relative to initial weight (%).

5. 24-h observation period

NOTE: Mice are housed individually, as described in 2.2. (Figure 5), for a period of 24 h, in order to measure food intake, home cage activity, nest building behavior, and FCM levels. The 24-h observation takes place twice: (i) 2 days prior to the procedure for baseline levels, (ii) on the day of the procedure.

1. Food intake
   1. Weigh the mice at regular intervals (e.g. 2 days before anesthesia, immediately before anesthesia, 2 days after anesthesia and weekly after anesthesia), in order to evaluate any changes in body weight (part of the score sheet).
      NOTE: Body weight is required to calculate food intake per gram of body weight. Water intake can also be measured during the 24-h observation period. If food intake is reduced, well-being may be impaired.
   2. Determine the initial weight of standard food diet (grams) provided in the food unit of the cage (approximately 100 g).
   3. Determine the weight of standard food diet at the end of the 24-h observation period.
   4. Scan the cage side beneath the food unit carefully for food spillage and add any extra food pellets found to the weight of food pellets remaining in the food unit.
   5. Calculate food intake per unit body weight.
2. Home cage activity
   NOTE: The following instructions refer to the use of an infrared sensor (see table of materials), but home cage activity can also be assessed with alternative programs. Deviation of home cage activity from control levels (e.g. hypoactivity, hyperactivity) may be a sign of impaired well-being.
   1. Start the program.
   2. Choose a sample interval of 1 min and an acquisition time of 24 h, meaning that impulses are recorded every minute for 24 h.
      NOTE: If the experimenter enters the room several times after recording started, only use data from periods, when mice were not disturbed (i.e. during the dark period).
   3. Sum up 10-min intervals of impulses.
   4. Calculate the area under the time curve (impulses × min).
3. Nest building behavior
   NOTE: Complex and high nests can serve as an indicator of well-being.
   1. Place a square cotton nestlet (see Table of Materials) with a defined weight (e.g. 2.0 g) in the middle of the cage.
   2. Score the nest on a 5-point scale (see below) according to Deacon¹⁴ the following morning, approximately 2 h after the light turns on. Weigh any untorn nestlet pieces that are at least 5% of the initial nestlet weight. Score the nests as follows¹⁴
      1. Assign score of "1" if 90% of the nestlet intact.
      2. Assign score of "2" if it is 50 - 90% intact.
      3. Assign score "3" if 50 - 90% of the nestlet is shredded.
      4. Assign score "4" if more than 90% is shredded but nest is flat, and less than 50% of its circumference is higher than mouse body height when curled up.
      5. Assign score "5" if more than 90% nestlet is shredded and nest is high, and more than 50% of its circumference is higher than body height of a curled up mouse.
4. Fecal Corticosterone Metabolites
   NOTE: Increases of FCM above the control level reflect acute stress levels over the 24-h postanesthetic period.
   1. Collect all dry fecal pellets from the cage by using forceps at the end of the 24-h observation period and eliminate wet pellets contaminated with urine.
   2. Extract FCM according to Palme et al.¹⁷, as follows.
      1. Dry fecal samples at a temperature of 60 - 70 °C.
      2. Homogenize fecal samples using a mortar.
      3. Shake an aliquot of 0.05 g with 1 mL of 80% methanol in a centrifuge tube for 30 min on a multi-vortex.
      4. Centrifuge samples at 2500 x g for 15 min.
      5. Pipette 0.5 mL of supernatant into another centrifuge tube.
      6. Store fecal samples (and extracts) at a minimum of -18 °C.
      7. Analyze FCM using a 5α-pregnane-3b,11b,21-triol-20-one enzyme immunoassay (EIA)^18,19 or another fully validated EIA.
   3. Calculate the percentage change of FCM concentrations relative to the baseline FCM concentrations.

6. Free exploratory paradigm

1. Take the home cage out of the rack and place it on a table surface at the end of the 24-h observation period.
2. Place a gridded cage top (without food or water bottles) in the cage at an angle of 45° to the longer side of the cage.
   NOTE: Do not destroy the nest, which serves as a hiding place for the mouse, but place the cage top diagonally above the nest.
3. Monitor or video-record the mice for 10 min from a distance of approximately 1.5 m.
   1. Start the timer.
   2. Note all times when the mouse climbs onto the cage top (with all four paws on the cage top) or leaves the cage top (with one or more paws on the cage floor).
      NOTE: Some mice may climb up the cage top and leave it in order to walk along the edge of the cage. Some mice also rear on the cage top. Treat these cases as if the mice were still on the cage top.
4. Evaluate parameters following Bert et al.²⁰.
   1. Analyze latency to first exploration (in seconds).
   2. Analyze number of explorations.
   3. Analyze total duration (seconds) of exploration.
      NOTE: A high latency to first exploration, a low number of explorations, and a low total duration of exploration can indicate higher trait anxiety levels.

Results

This protocol was originally developed to assess well-being of C57BL/6JRj mice following a single experience of isoflurane anesthesia (one 45-min anesthesia session, n = 13 females) or repeated isoflurane anesthesia (six 45-min anesthesia sessions with 3 - 4 days between the anesthesia sessions, n = 13 females) compared with the well-being of control mice (n = 6 females)¹⁰, which received no anesthesia but were tested according to the same measures. We assessed the...

Discussion

The protocol was originally developed to assess well-being of C57BL/6JRj mice that received either a single anesthesia or repeated isoflurane anesthesia. The results confirm that tests of luxury behaviors, as well as other measures (e.g. the free exploratory paradigm, the MGS, burrowing food intake) were sensitive methods for assessing well-being. Repeated isoflurane anesthesia caused short-term effects on trait anxiety-related behavior, the MGS, and burrowing behavior. Moreover, repeated isoflurane anesthesia a...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
Isofluran	CP-Pharma Handelsgesellschaft mbH	1214
InfraMot - Sensore Units	TSE Systems	302015-SENS
InfraMot - Control Units	TSE Systems	302015-C/16
InfraMot - Software	TSE Systems	302015-S
Nestlet N	Ancare - Plexx	NES3600
Camera EOS 350D	Canon