Thanks to Sabine Jacobs for assisting with the sample collection, Edith Klobetz-Rassam for analysis of FCM, PD Dr. med. vet. habil. Roswitha Merle for assisting with statistical analysis, and Wiebke Gentner for proofreading the manuscript. The study is part of the Berlin-Brandenburg research platform BB3R (www.bb3r.de) and was funded by the German Federal Ministry of Education and Research (grant number: 031A262A) (www.bmbf.de/en/index.html).&#160;

The study was performed according to the guidelines laid down by the German Animal Welfare Act and was approved by the Berlin State Authority (&#34;Landesamt f&#252;r Gesundheit und Soziales&#34;, 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 &#8805; 2 &#215; (s/&#181;1- &#181;2)2 &#215; (z&#945; + z&#946;)2. &#181;1- &#181;2 is the difference between population means at which power and sample size calculations are made (&#945; = 5%, &#946; = 80%); z&#945; = 1.96 and z&#946; = 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&#39;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
<ol>
	<li>Allow mice to habituate to the animal facility for at least 2 weeks after they have been obtained from another facility or vendor.</li>
	<li>House mice in groups and maintain them under standard conditions (room temperature 22 &#177; 2 &#176;C; relative humidity 55 &#177; 10%) on a light:dark cycle of 12:12 h.</li>
	<li>Provide all groups with a tunnel and cotton nestlets as standard enrichment, and provide food and water ad libitum.</li>
	<li>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.</li>
</ol>
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.
<ol>
	<li>Prepare an observation cage to test burrowing behavior8 and to take the photographs for use in the MGS13 (Figure 4).
	<ol>
		<li>Use a glass box with a floor area of approximately 220 mm &#215; 290 mm and a height of 390 mm.</li>
		<li>Cover the floor of this box with approximately 0.5 cm of bedding material.</li>
		<li>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.</li>
		<li>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 .</li>
	</ol>
	</li>
	<li>Prepare a cage (type III: 420 mm &#215; 260 mm &#215; 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.
	<ol>
		<li>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.</li>
		<li>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.</li>
		<li>Mount the infrared sensor on the top of the cage, when using an infrared sensor to measure home cage activity (see table of materials).</li>
		<li>Provide food, the same kind that is normally supplied as diet, and water ad libitum.</li>
	</ol>
	</li>
</ol>
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.
<ol>
	<li>Use a high-definition camera for photography.</li>
	<li>Gently transfer the mouse into the observation cage and allow the mouse to habituate to the new environment for at least 30 min.</li>
	<li>Continuously take about 30 - 40 photographs for each time point within 1 - 2 min.</li>
	<li>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.</li>
	<li>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.</li>
	<li>Crop the photographs to display only the head of the mouse so that the body position is not visible13.</li>
	<li>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.</li>
	<li>Randomize the order of the sheets.</li>
	<li>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 (&#34;narrowing of the orbital area, with a tightly closed eyelid or an eye squeeze&#34;); nose bulge (&#34;rounded extension of skin visible on the bridge of the nose&#34;); cheek bulge (&#34;convex appearance of the cheek muscle&#34;); ear position (&#34;ears pulled apart and back from their baseline position or featuring vertical ridges that form owing to tips of ears being drawn back&#34;); whisker change (&#34;movement of whiskers from their baseline position either backward, against the face or forward, as if standing on end; whiskers may also clump together&#34;).</li>
	<li>Analyze scores, as follows (adapted from Langford et al.13).
	<ol>
		<li>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.</li>
		<li>Subtract the mean for the baseline photographs from the mean for the photographs post procedure to obtain a MGS difference score for each mouse.</li>
		<li>Test for differences in the MGS difference scores between the persons (nonparametric test for related samples). 
		NOTE: If there is a significant difference (p &#60; 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.</li>
		<li>Average the MGS difference scores obtained from the different scorers for each mouse, if results of all persons do not significantly differ.</li>
		<li>Use a nonparametric statistical test to compare the MGS difference scores averaged between the study groups.</li>
	</ol>
	</li>
</ol>
4. Burrowing behavior8,15,16
<ol>
	<li>Prepare burrows by placing 140 &#177; 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.</li>
	<li>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.</li>
	<li>Carry out the test twice, 2 days prior to the procedure (baseline); carry out the last 30 min post procedure as well.
	<ol>
		<li>Let the mouse habituate for at least 30 min to the observation cage where photographs for the MGS were taken.</li>
		<li>Place the plastic water bottle filled with food pellets parallel to the back wall of the observation cage.</li>
		<li>Weigh food pellets (g) remaining in the burrow after 2 h.</li>
	</ol>
	</li>
	<li>Calculate the weight of food pellets removed from the burrow by mice relative to initial weight (%).</li>
</ol>
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.
<ol>
	<li>Food intake
	<ol>
		<li>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.</li>
		<li>Determine the initial weight of standard food diet (grams) provided in the food unit of the cage (approximately 100 g).</li>
		<li>Determine the weight of standard food diet at the end of the 24-h observation period.</li>
		<li>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.</li>
		<li>Calculate food intake per unit body weight.</li>
	</ol>
	</li>
	<li>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.
	<ol>
		<li>Start the program.</li>
		<li>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).</li>
		<li>Sum up 10-min intervals of impulses.</li>
		<li>Calculate the area under the time curve (impulses &#215; min).</li>
	</ol>
	</li>
	<li>Nest building behavior 
	NOTE: Complex and high nests can serve as an indicator of well-being.
	<ol>
		<li>Place a square cotton nestlet (see Table of Materials) with a defined weight (e.g. 2.0 g) in the middle of the cage.</li>
		<li>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
		<ol>
			<li>Assign score of &#34;1&#34; if 90% of the nestlet intact.</li>
			<li>Assign score of &#34;2&#34; if it is 50 - 90% intact.</li>
			<li>Assign score &#34;3&#34; if 50 - 90% of the nestlet is shredded.</li>
			<li>Assign score &#34;4&#34; 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.</li>
			<li>Assign score &#34;5&#34; 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.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Fecal Corticosterone Metabolites 
	NOTE: Increases of FCM above the control level reflect acute stress levels over the 24-h postanesthetic period.
	<ol>
		<li>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.</li>
		<li>Extract FCM according to Palme et al.17, as follows.
		<ol>
			<li>Dry fecal samples at a temperature of 60 - 70 &#176;C.</li>
			<li>Homogenize fecal samples using a mortar.</li>
			<li>Shake an aliquot of 0.05 g with 1 mL of 80% methanol in a centrifuge tube for 30 min on a multi-vortex.</li>
			<li>Centrifuge samples at 2500 x g for 15 min.</li>
			<li>Pipette 0.5 mL of supernatant into another centrifuge tube.</li>
			<li>Store fecal samples (and extracts) at a minimum of -18 &#176;C.</li>
			<li>Analyze FCM using a 5&#945;-pregnane-3b,11b,21-triol-20-one enzyme immunoassay (EIA)18,19 or another fully validated EIA.</li>
		</ol>
		</li>
		<li>Calculate the percentage change of FCM concentrations relative to the baseline FCM concentrations.</li>
	</ol>
	</li>
</ol>
6. Free exploratory paradigm
<ol>
	<li>Take the home cage out of the rack and place it on a table surface at the end of the 24-h observation period.</li>
	<li>Place a gridded cage top (without food or water bottles) in the cage at an angle of 45&#176; 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.</li>
	<li>Monitor or video-record the mice for 10 min from a distance of approximately 1.5 m.
	<ol>
		<li>Start the timer.</li>
		<li>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.</li>
	</ol>
	</li>
	<li>Evaluate parameters following Bert et al.20.
	<ol>
		<li>Analyze latency to first exploration (in seconds).</li>
		<li>Analyze number of explorations.</li>
		<li>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.</li>
	</ol>
	</li>
</ol>

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N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Hippocampal+cytotoxic+lesion+effects+on+species-typical+behaviours+in+mice.">Hippocampal cytotoxic lesion effects on species-typical behaviours in mice.</a> Behav Brain Res. 132 (2), 203-213 (2002).</li><li>Filali, M., Lalonde, R., Rivest, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Subchronic+memantine+administration+on+spatial+learning,+exploratory+activity,+and+nest-building+in+an+APP/PS1+mouse+model+of+Alzheimer's+disease.">Subchronic memantine administration on spatial learning, exploratory activity, and nest-building in an APP/PS1 mouse model of Alzheimer's disease.</a> Neuropharmacology. 60 (6), 930-936 (2011).</li><li>Guenther, K., Deacon, R. M., Perry, V. H., Rawlins, J. N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Early+behavioural+changes+in+scrapie-affected+mice+and+the+influence+of+dapsone.">Early behavioural changes in scrapie-affected mice and the influence of dapsone.</a> Eur J Neurosci. 14 (2), 401-409 (2001).</li><li>Deacon, R. M., Reisel, D., Perry, V. H., Nicholas, J., Rawlins, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Hippocampal+scrapie+infection+impairs+operant+DRL+performance+in+mice.">Hippocampal scrapie infection impairs operant DRL performance in mice.</a> Behav Brain Res. 157 (1), 99-105 (2005).</li><li>Jirkof, P., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Assessment+of+postsurgical+distress+and+pain+in+laboratory+mice+by+nest+complexity+scoring.">Assessment of postsurgical distress and pain in laboratory mice by nest complexity scoring.</a> Lab Anim. 47 (3), 153-161 (2013).</li><li>Atanasov, N. A., Sargent, J. L., Parmigiani, J. P., Palme, R., Diggs, H. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Characterization+of+train-induced+vibration+and+its+effect+on+fecal+corticosterone+metabolites+in+mice.">Characterization of train-induced vibration and its effect on fecal corticosterone metabolites in mice.</a> J Am Assoc Lab Anim Sci. 54 (6), 737-744 (2015).</li><li>Voigt, C. C., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Hormonal+stress+response+of+laboratory+mice+to+conventional+and+minimally+invasive+bleeding+techniques.">Hormonal stress response of laboratory mice to conventional and minimally invasive bleeding techniques.</a> Anim Welf. 22 (4), 449-455 (2013).</li><li>Walker, M. K., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+less+stressful+alternative+to+oral+gavage+for+pharmacological+and+toxicological+studies+in+mice.">A less stressful alternative to oral gavage for pharmacological and toxicological studies in mice.</a> Toxicol Appl Pharmacol. 260 (1), 65-69 (2012).</li><li>Miyashita, T., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Social+stress+increases+biopyrrins,+oxidative+metabolites+of+bilirubin,+in+mouse+urine.">Social stress increases biopyrrins, oxidative metabolites of bilirubin, in mouse urine.</a> Biochem Biophys Res Commun. 349 (2), 775-780 (2006).</li><li>Bains, R. S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Analysis+of+individual+mouse+activity+in+group+housed+animals+of+different+inbred+strains+using+a+novel+automated+home+cage+analysis+system.">Analysis of individual mouse activity in group housed animals of different inbred strains using a novel automated home cage analysis system.</a> Front Behav Neurosci. 10 (106), (2016).</li><li>Saibaba, P., Sales, G. D., Stodulski, G., Hau, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Behaviour+of+rats+in+their+home+cages:+daytime+variations+and+effects+of+routine+husbandry+procedures+analysed+by+time+sampling+techniques.">Behaviour of rats in their home cages: daytime variations and effects of routine husbandry procedures analysed by time sampling techniques.</a> Lab Anim. 30 (1), 13-21 (1996).</li></ol>

The authors have nothing to disclose.

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...

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 important5,6,7,8. 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 decline5,7. 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.

<table><tbody><tr><td>Isofluran</td><td>CP-Pharma Handelsgesellschaft mbH</td><td>1214</td><td></td></tr><tr><td>InfraMot - Sensore Units</td><td>TSE Systems</td><td>302015-SENS</td><td></td></tr><tr><td>InfraMot - Control Units</td><td>TSE Systems</td><td>302015-C/16</td><td></td></tr><tr><td>InfraMot - Software</td><td>TSE Systems</td><td>302015-S</td><td></td></tr><tr><td>Nestlet N</td><td>Ancare - Plexx</td><td>NES3600</td><td></td></tr><tr><td>Camera EOS 350D</td><td>Canon</td><td></td><td></td></tr></tbody></table>

systematic assessment of well-being in mice for procedures using general anesthesia

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.

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...

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

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.

In keeping with the 3R Principle (Replacement, Reduction, Refinement) developed by Russel and Burch, scientific research should use alternatives to animal ...

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Research

JoVE Journal

Behavior

12.1K Views.  Freie Universität Berlin. 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.

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

Nest Building as an Indicator of Health and Welfare in Laboratory Mice

The minimization and alleviation of suffering has moral and scientific implications. In order to mitigate this negative experience one must be able to identify when an animal is actually in distress. Pain, illness, or distress cannot be managed if unrecognized. Evaluation of pain or illness typically involves the measurement of physiologic and behavioral indicators which are either invasive or not suitable for large scale assessment. The observation of nesting behavior shows promise as the basis of a species appropriate cage-side assessment tool for recognizing distress in mice. Here we demonstrate the utility of nest building behavior in laboratory mice as an ethologically relevant indicator of welfare. The methods presented can be successfully used to identify thermal stressors, aggressive cages, sickness, and pain. Observation of nest building behavior in mouse colonies provides a refinement to health and well-being assessment on a day to day basis.

We demonstrate the utility of nest building behavior in laboratory mice as an indicator of welfare. Nest scoring is a sensitive technique that is altered by temperature, illness, and aggression. The time to integrate into nest test (TINT) is a simple cage-side assessment that can detect postoperative pain.

The minimization and alleviation of  suffering has moral and scientific implications. In order to mitigate this  negative experience one must be able to ...

Automated, Quantitative Cognitive/Behavioral Screening of Mice: For Genetics, Pharmacology, Animal Cognition and Undergraduate Instruction

We describe a high-throughput, high-volume, fully automated, live-in 24/7 behavioral testing system for assessing the effects of genetic and pharmacological manipulations on basic mechanisms of cognition and learning in mice. A standard polypropylene mouse housing tub is connected through an acrylic tube to a standard commercial mouse test box. The test box has 3 hoppers, 2 of which are connected to pellet feeders. All are internally illuminable with an LED and monitored for head entries by infrared (IR) beams. Mice live in the environment, which eliminates handling during screening. They obtain their food during two or more daily feeding periods by performing in operant (instrumental) and Pavlovian (classical) protocols, for which we have written protocol-control software and quasi-real-time data analysis and graphing software. The data analysis and graphing routines are written in a MATLAB-based language created to simplify greatly the analysis of large time-stamped behavioral and physiological event records and to preserve a full data trail from raw data through all intermediate analyses to the published graphs and statistics within a single data structure. The data-analysis code harvests the data several times a day and subjects it to statistical and graphical analyses, which are automatically stored in the &#34;cloud&#34; and on in-lab computers. Thus, the progress of individual mice is visualized and quantified daily. The data-analysis code talks to the protocol-control code, permitting the automated advance from protocol to protocol of individual subjects. The behavioral protocols implemented are matching, autoshaping, timed hopper-switching, risk assessment in timed hopper-switching, impulsivity measurement, and the circadian anticipation of food availability. Open-source protocol-control and data-analysis code makes the addition of new protocols simple. Eight test environments fit in a 48 in x 24 in x 78 in&#160;cabinet; two such cabinets (16 environments) may be controlled by one computer.

Fully automated system for measuring physiologically meaningful properties of the mechanisms mediating spatial localization, temporal localization, duration, rate&#160;and probability estimation, risk assessment, impulsivity, and the accuracy and precision of memory, in order to assess the effects of genetic and pharmacological manipulations on foundational mechanisms of cognition in mice.

We describe a high-throughput, high-volume, fully automated, live-in 24/7 behavioral testing system for assessing the effects of genetic and ...

Use of the Open Field Maze to Measure Locomotor and Anxiety-like Behavior in Mice

Animal models have proven to be invaluable to researchers trying to answer questions regarding the mechanisms of behavior. The Open Field Maze is one of the most commonly used platforms to measure behaviors in animal models. It is a fast and relatively easy test that provides a variety of behavioral information ranging from general ambulatory ability to data regarding the emotionality of the subject animal. As it relates to rodent models, the procedure allows the study of different strains of mice or rats both laboratory bred and wild-captured. The technique also readily lends itself to the investigation of different pharmacological compounds for anxiolytic or anxiogenic effects. Here, a protocol for use of the open field maze to describe mouse behaviors is detailed and a simple analysis of general locomotor ability and anxiety-related emotional behaviors between two strains of C57BL/6 mice is performed. Briefly, using the described protocol we show Wild Type mice exhibited significantly less anxiety related behaviors than did age-matched Knock Out mice while both strains exhibited similar ambulatory ability.

A protocol is provided to use an Open Field Maze to access general locomotor activity, anxiety and emotionality in a laboratory mouse model.

Animal models have proven to be invaluable to researchers trying to answer questions regarding the mechanisms of behavior. The Open Field Maze is one of ...

Assessment of Social Interaction Behaviors

Social interactions are a fundamental and adaptive component of the biology of numerous species. Social recognition is critical for the structure and stability of the networks and relationships that define societies. For animals, such as mice, recognition of conspecifics may be important for maintaining social hierarchy and for mate choice 1.
A variety of neuropsychiatric disorders are characterized by disruptions in social behavior and social recognition, including depression, autism spectrum disorders, bipolar disorders, obsessive-compulsive disorders, and schizophrenia. Studies of humans as well as animal models (e.g., Drosophila melanogaster, Caenorhabditis elegans, Mus musculus, Rattus norvegicus) have identified genes involved in the regulation of social behavior 2. To assess sociability in animal models, several behavioral tests have been developed (reviewed in 3). Integrative research using animal models and appropriate tests for social behavior may lead to the development of improved treatments for social psychopathologies.
The three-chamber paradigm test known as Crawley's sociability and preference for social novelty protocol has been successfully employed to study social affiliation and social memory in several inbred and mutant mouse lines (e.g. 4-7). The main principle of this test is based on the free choice by a subject mouse to spend time in any of three box's compartments during two experimental sessions, including indirect contact with one or two mice with which it is unfamiliar. To quantitate social tendencies of the experimental mouse, the main tasks are to measure a) the time spent with a novel conspecific and b) preference for a novel vs. a familiar conspecific. Thus, the experimental design of this test allows evaluation of two critical but distinguishable aspects of social behavior, such as social affiliation/motivation, as well as social memory and novelty. "Sociability" in this case is defined as propensity to spend time with another mouse, as compared to time spent alone in an identical but empty chamber 7. "Preference for social novelty" is defined as propensity to spend time with a previously unencountered mouse rather than with a familiar mouse 7. This test provides robust results, which then must be carefully analyzed, interpreted and supported/confirmed by alternative sociability tests. In addition to specific applications, Crawley's sociability test can be included as an important component of general behavioral screen of mutant mice.

Here we describe a detailed protocol for examination of sociability in mice by using Crawley's sociability and preference for social novelty test. We describe the advantages and possible applications for this procedure, including critical details important for correct interpretation of the results.

Social interactions are a fundamental and adaptive component of the biology of numerous species. Social recognition is critical for the structure and ...

Neuroscience

Implantation of Radiotelemetry Transmitters Yielding Data on ECG, Heart Rate, Core Body Temperature and Activity in Free-moving Laboratory Mice

The laboratory mouse is the animal species of choice for most biomedical research, in both the academic sphere and the pharmaceutical industry. Mice are a manageable size and relatively easy to house. These factors, together with the availability of a wealth of spontaneous and experimentally induced mutants, make laboratory mice ideally suited to a wide variety of research areas.
In cardiovascular, pharmacological and toxicological research, accurate measurement of parameters relating to the circulatory system of laboratory animals is often required. Determination of heart rate, heart rate variability, and duration of PQ and QT intervals are based on electrocardiogram (ECG) recordings. However, obtaining reliable ECG curves as well as physiological data such as core body temperature in mice can be difficult using conventional measurement techniques, which require connecting sensors and lead wires to a restrained, tethered, or even anaesthetized animal. Data obtained in this fashion must be interpreted with caution, as it is well known that restraining and anesthesia can have a major artifactual influence on physiological parameters1, 2.
Radiotelemetry enables data to be collected from conscious and untethered animals. Measurements can be conducted even in freely moving animals, and without requiring the investigator to be in the proximity of the animal. Thus, known sources of artifacts are avoided, and accurate and reliable measurements are assured. This methodology also reduces interanimal variability, thus reducing the number of animals used, rendering this technology the most humane method of monitoring physiological parameters in laboratory animals3, 4. Constant advancements in data acquisition technology and implant miniaturization mean that it is now possible to record physiological parameters and locomotor activity continuously and in realtime over longer periods such as hours, days or even weeks3, 5.
Here, we describe a surgical technique for implantation of a commercially available telemetry transmitter used for continuous measurements of core body temperature, locomotor activity and biopotential (i.e. onelead ECG), from which heart rate, heart rate variability, and PQ and QT intervals can be established in freeroaming, untethered mice. We also present pre-operative procedures and protocols for post-operative intensive care and pain treatment that improve recovery, well-being and survival rates in implanted mice5, 6.

A surgical technique for implantation of commercially available telemetry transmitters used for continuous measurement of biopotential (one-lead ECG), heart rate, core body temperature and locomotor activity in freely moving mice is shown. Recommendations and protocols for post-operative care and pain relief, improving recovery, well being and survival rate are also presented.

The laboratory mouse is the animal species of choice for most biomedical research, in both the academic sphere and the pharmaceutical industry. Mice are a ...

Medicine

A Behavioral Test Battery for the Repeated Assessment of Motor Skills, Mood, and Cognition in Mice

Pharmacological and toxicological studies in neurodegeneration require comprehensive behavioral analysis in mice because motor dysfunctions and dysfunctions in mood and cognition are common and often shared symptoms in neurodegenerative diseases. Shown here is a behavioral test battery for motor, mood, and cognition, which can be repeatedly tested in a longitudinal study. This battery assesses the overall behavioral phenotype in mice by examining each domain of behavior with at least two independent well-accepted tests (i.e., open-field test and rotarod test for motor function, social interaction test, elevated plus maze test, and forced swim test for emotional function, and Morris water maze test and novel object recognition test for cognitive function). Therefore, this sensitive and comprehensive test battery is a powerful tool for the study of behavioral alternation in neurodegeneration.

A comprehensive behavioral test battery of motor skills, mood&#8212;including social interaction, depression, and anxiety&#8212;and cognition is designed for the repeated assessment of neurodegeneration-related behavioral changes in mice.

Pharmacological and toxicological studies in neurodegeneration require comprehensive behavioral analysis in mice because motor dysfunctions and ...

A Chronic Immobilization Stress Protocol for Inducing Depression-Like Behavior in Mice

Depression is not yet fully understood, but various causative factors have been reported. Recently, the prevalence of depression has increased. However, therapeutic treatments for depression or research on depression is scarce. Thus, in the present paper, we propose a mouse model of depression induced by movement restriction. Chronic mild stress (CMS) is a well-known technique to induce depressive-like behavior. However, it necessitates a complex procedure consisting of a combination of various mild stresses. In contrast, chronic immobilization stress (CIS) is a readily accessible chronic stress model, modified from a restraint model that induces depressive behavior by restricting movement using a restrainer for a certain period. To evaluate the depressive-like behaviors, the sucrose preference test (SPT), the tail suspension test (TST), and the ELISA assay to measure stress marker corticosterone levels are combined in the present experiment. The described protocols illustrate the induction of CIS and evaluation of the changes in behavior and physiological factors for the validation of depression.

This article provides a simplified and standardized protocol for induction of depressive-like behavior in chronically immobilized mice by using a restrainer. In addition, behavior and physiological techniques to verify induction of depression are explained.

Depression is not yet fully understood, but various causative factors have been reported. Recently, the prevalence of depression has increased. However, ...

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

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.

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.

One desirable endpoint of general anesthesia is  the state of unconsciousness, also known as hypnosis. Defining the hypnotic state in animals is  less ...

Assessment of Mouse Judgment Bias through an Olfactory Digging Task

Judgment biases (JB) are differences in the way that individuals in positive and negative affective/emotional states interpret ambiguous information. This phenomenon has long been observed in humans, with individuals in positive states responding to ambiguity 'optimistically' and those in negative states instead showing 'pessimism'. Researchers aiming to assess animal affect have taken advantage of these differential responses, developing tasks to assess judgment bias as an indicator of affective state. These tasks are becoming increasingly popular across diverse species and fields of research. However, for laboratory mice, the most widely used vertebrates in research and a species heavily relied upon to model affective disorders, only one JB task has been successfully validated as sensitive to changes in affective state. Here, we provide a detailed description of this novel murine JB task, and evidence of its sensitivity to mouse affect. Though refinements are still necessary, assessment of mouse JB opens the door for answering both practical questions regarding mouse welfare, and fundamental questions about the impact of affective state in translational research.

This article provides a detailed description of a novel mouse judgment bias protocol. Evidence of this olfactory digging task's sensitivity to affective state is also demonstrated and its utility across diverse research fields is discussed.

Judgment biases (JB) are differences in the way that individuals in positive and negative affective/emotional states interpret ambiguous information. This ...

Behavioral Assays for Autism Spectrum Disorder in Mouse Models

Strategies for Assessing Autistic-Like Behaviors in Mice

Autism spectrum disorder (ASD) is a neurobiologically complex condition with a heterogeneous genetic etiology. Clinically, ASD is diagnosed by social communication impairments and restrictive or repetitive behaviors, such as hand flapping or lining up objects. These behavioral patterns can be reliably observed in mouse models with ASD-linked genetic mutations, making them highly useful tools for studying the underlying cellular and molecular mechanisms in ASD. Understanding how genetic changes affect the neurobiology and behaviors observed in ASD will facilitate the development of novel targeted therapeutic compounds to ameliorate core behavioral impairments. Our lab has employed several protocols encompassing well-described training and testing procedures that reflect a wide range of behavioral deficits related to ASD. Here, we detail two assays to study the core features of ASD in mouse models: self-grooming (a measure of repetitive behavior) and the three-chamber social interaction test (a measure of social interaction approach and preference for social novelty).

Rodent models are valuable tools for studying core behaviors related to autism spectrum disorder (ASD). In this article, we expound on two behavioral tests for modeling the core features of ASD in mice: self-grooming, which assesses repetitive behavior, and the three-chamber social interaction test, which documents social impairments.

Autism spectrum disorder (ASD) is a neurobiologically complex condition with a heterogeneous genetic etiology. Clinically, ASD is diagnosed by social ...