Materials and Reagents

Group-housed mice
50 mL conical tubes with holes cut into the sides (for restraint stress–induced anorexia; Figure 1)

Figure 1. Setup of restraint stress–induced anorexia apparatus. Separate cages are used for male and female mice, respectively
Clean mouse cages
Standard chow mouse diet (Formulab Diet 5008 or similar approved mouse diet)
Standard home cage water bottles
Standard food hoper
Cage enrichment (such as nestlets)
Scale for measuring food intake and body weight (Satorius Entris II Essential Precision Balance or similar)
Lab notebook for recording food intake and body weight

Instructions to make restraint stress apparatus

A simple restraint stress–anorexia apparatus can be constructed from 50 mL conical tubes (Figure 1). Make 6–12 holes, depending on how many you can fit, on the sides of the conical tubes to enable mice to breathe. Care should be taken to make the holes big enough for air flow, but not big enough for the mice to fit their nose into, so they do not hurt themselves in the borders. Holes can be made by using a tool such as a heat pen for melting plastic. The mouse identifying information (i.e., ear tag number and sex) can be written on the side of each tube. During restraint sessions, the mice can be placed into the restrainers within their resident cage (Figure 1). Following each restraint session, the restraint tubes should be cleaned with soap and water and allowed to dry before re-using on each mouse.

Procedure

Restraint stress–induced anorexia
1. Group house mice, 3–4 per cage and separated by sex; acclimate the mice to housing conditions for one week prior to beginning food intake measurements. During this period, mice should be handled daily by the investigator to acclimate the mice to handling-associated stress.
2. To acclimate the mice to handling, pick up each mouse and scruff gently. Remove the water bottle and food for one hour prior to baseline feeding measurements (to control for the one-hour restraint period in which mice will not have access to food and water). Place mice in a clean cage immediately following the removal of the food and water so that mice do not have access to any food crumbs that may be on the bottom of the cage.
  Note: All feeding measurements should be performed at the same time of day in the same experimental location.
3. After one hour of food and water removal, add a water bottle to the clean cage with fresh bedding and mouse enrichment (i.e., nestlet or wooden block) and between 20 and 30 g of food to the food hopper (mice eat approximately 3–5 g/day/mouse). Measure the body weight of the mice and amount of food immediately prior to adding the food and water to the cages.
4. Measure the amount of food left in the food hopper 1, 2, 4, and 24 h after adding the food. Food intake per mouse is calculated at each time point by dividing the amount of food consumed by the number of mice in each cage.
5. Measure the body weight of the mice 24 h after adding the food.
6. Repeat steps A2–A4 for 3 to 5 days to establish baseline levels of food intake for each mouse prior to restraint stress.
7. On restraint stress day (following baseline food intake measurements), grab the mouse by the tail and place each mouse in a 50 mL conical tube (see Figure 1) for sixty min, inserting the head of the animal first followed by the rest of its body, including the tail. For the control animals, only remove the food and water to give them a similar environment of food and water restriction as the restraint mice for sixty minutes.
  Notes:
  1. Institutional animal care and use approval (IACUC) must be obtained to perform experimental work on mice. It is essential to monitor each mouse during the restraint period to ensure that mice are breathing adequately.
  2. To obtain the food intake of individual mice with and without restraint stress, the above procedure (steps 1–7) may also be performed on singly housed mice. However, mice should be single-caged for a minimum of one week prior to starting restraint stress–anorexia experiments in order to acclimate mice to single housing.
8. Following sixty minutes of restraint, return the mice to the home cages with their littermates and add food and water as described in step A3.
9. Measure food intake as described in step A4.
10. Repeat steps A7–A9 for an additional 7–10 days to examine the effects of chronic restraint–stress on food intake and body weight.
Social isolation–induced anorexia
1. Group house mice with 3–4 per cage and acclimate the mice to housing conditions for one week prior to beginning food intake measurements. During this period, mice should be handled daily by the investigator to acclimate the mice to handling-associated stress.
2. Change mice to a fresh cage, with fresh bedding and mouse enrichment. Measure the body weight of each mouse. Add 20–30 g of food to the cages and measure the amount of food added to each cage. Return to measure the amount of remaining food 1, 2, 4, and 24 h later. Calculate the amount of food consumed by each mouse by dividing the total amount of food consumed by the number of mice in the cage.
  Note: All feeding experiments should be performed at the same time of day in the same experimental location. Since mice are nocturnal and consume most of their food during the dark cycle, optimal results will be obtained by performing feeding experiments at the onset of the rodent’s dark cycle.
3. Repeat step B2 for 3–5 consecutive days to establish a baseline measurement of food intake and body weight prior to social isolation.
4. At the same time of day as steps B2-B3, single-cage each mouse, measure each mouse’s body weight, and add 10–15 g of food to each cage. Return to measure the amount of food in each cage 1, 2, 4, and 24 h later. For the control group, continue to calculate the amount of food consumed by each mouse by dividing the total amount of food consumed by the number of mice in the cage.
  Note: Mice consume food based on a circadian structure. Care must be made to perform all feeding measurements at the same time each day.
5. Repeat step B4 for an additional 3–14 days.

Data analysis

Exclusion criteria: During stress-induced anorexia experiments, researchers must abide by IACUC approved exclusion criteria for removing animals from experimental conditions. These exclusion criteria will vary at each research institute and individual investigators should discuss exclusion criteria with veterinary staff prior to initiating experiments. Generally, mice are removed from experimental sessions when signs of pain or distress, such as reduced activity or piloerection, are observed or if any of the animal’s body weight drops by more than 20 percent.

Following the completion of restraint stress induced–anorexia or social isolation–anorexia experiments, food intake and body weight is compared for each mouse following no-restraint vs. restraint-stress (paired, within samples comparisons). It is expected that restraint stress will reduce food intake and body weight in mice. For restraint stress–induced and social isolation–induced anorexia, we typically observe 10%–20% changes in food intake in the hours immediately following restraint stress, although results will vary depending on mouse strain, sex, and age of mice. The effects of pharmacological or other manipulations to reduce or enhance stress-induced anorexia can also be evaluated by determining the effect of the manipulation on food intake and body weight. The specific experimental design will vary slightly depending on the specific research question.

Conclusion

In this article, we describe two simple mouse models for inducing stress-induced anorexia. These models provide a valuable experimental tool for researchers interested in determining the neural mechanisms connecting stress with feeding behavior.

Acknowledgments

This work was funded in part by awards R00 DK127065 (P.S.), Brain and Behavior Research Foundation Young Investigator Award (P.S.), and the Foundation for Prader Willi Research (P.S.).

The original methods in this paper are based on our prior publication (Sweeney et al., 2021; DOI: 10.1126/scitranslmed.abd6434).

Competing interests

There are no conflicts of interest or competing interests.

Ethics

All experiments were approved by the University of Illinois Institutional animal care and use committee (IACUC).

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