Assessment of the Metabolic Effects of Isocaloric 2:1 Intermittent Fasting in Mice

Summary

The current article describes a detailed protocol for isocaloric 2:1 intermittent fasting to protect and treat against obesity and impaired glucose metabolism in wild-type and ob/ob mice.

Abstract

Intermittent fasting (IF), a dietary intervention involving periodic energy restriction, has been considered to provide numerous benefits and counteract metabolic abnormalities. So far, different types of IF models with varying durations of fasting and feeding periods have been documented. However, interpreting the outcomes is challenging, as many of these models involve multifactorial contributions from both time- and calorie-restriction strategies. For example, the alternate day fasting model, often used as a rodent IF regimen, can result in underfeeding, suggesting that health benefits from this intervention are likely mediated via both caloric restriction and fasting-refeeding cycles. Recently, it has been successfully demonstrated that 2:1 IF, comprising 1 day of fasting followed by 2 days of feeding, can provide protection against diet-induced obesity and metabolic improvements without a reduction in overall caloric intake. Presented here is a protocol of this isocaloric 2:1 IF intervention in mice. Also described is a pair-feeding (PF) protocol required to examine a mouse model with altered eating behaviors, such as hyperphagia. Using the 2:1 IF regimen, it is demonstrated that isocaloric IF leads to reduced body weight gain, improved glucose homeostasis, and elevated energy expenditure. Thus, this regimen may be useful to investigate the health impacts of IF on various disease conditions.

Introduction

Modern lifestyle is associated with longer daily food intake time and shorter fasting periods¹. This contributes to the current global obesity epidemic, with metabolic disadvantages seen in humans. Fasting has been practiced throughout human history, and its diverse health benefits include prolonged lifespan, reduced oxidative damage, and optimized energy homeostasis^2,3. Among several ways to practice fasting, periodic energy deprivation, termed intermittent fasting (IF), is a popular dietary method that is widely practiced by the general population due to its easy and simple regimen. Recent studies in preclinical and clinical models have demonstrated that IF can provide health benefits comparable to prolonged fasting and caloric restriction, suggesting that IF can be a potential therapeutic strategy for obesity and metabolic diseases^2,3,4,5.

IF regimens vary in terms of fasting duration and frequency. Alternate day fasting (i.e., 1 day feeding/1 day fasting; 1:1 IF) has been the most commonly used IF regimen in rodents to study its beneficial health impacts on obesity, cardiovascular diseases, neurodegenerative diseases, etc.^2,3. However, as shown in previous studies^6,7, and further mechanistically confirmed in our energy intake analysis⁸, 1:1 IF results in underfeeding (~80%) due to the lack of sufficient feeding time to compensate for energy loss. This makes it unclear whether the health benefits conferred by 1:1 IF are mediated by calorie restriction or modification of eating patterns. Therefore, a new IF regimen has been developed and is shown here, comprising of a 2 day feeding/1 day fasting (2:1 IF) pattern, which provides mice with sufficient time to compensate for food intake (~99%) and body weight. These mice are then compared to an ad libitum (AL) group. This regimen enables examination of the effects of isocaloric IF in the absence of caloric reduction in wild-type mice.

In contrast, in a mouse model that exhibits altered feeding behavior, AL feeding may not be a proper control condition to compare and examine the effects of 2:1 IF. For example, since ob/ob mice (a commonly used genetic model for obesity) exhibit hyperphagia due to the lack of leptin regulating appetite and satiety, those with 2:1 IF exhibit ~20% reduced caloric intake compared to ob/ob mice with AL feeding. Thus, to properly examine and compare the effects of IF in ob/ob mice, a pair-feeding group as a suitable control needs to be employed.

Overall, a comprehensive protocol is provided to perform isocaloric 2:1 IF, including use of a pair-feeding control. It is further demonstrated that isocaloric 2:1 IF protects mice from high fat diet-induced obesity and/or metabolic dysfunction in both wild-type and ob/ob mice. This protocol can be used to examine the beneficial health impacts of 2:1 IF on various pathological conditions including neurological disorders, cardiovascular diseases, and cancer.

Protocol

All methods and protocols here have been approved by Animal Care Committees in The Animal Care and Veterinary Service (ACVS) of the University of Ottawa and The Centre for Phenogenomics (TCP) and conform to the standards of the Canadian Council on Animal Care. It should be noted that all procedures described here should be performed under institutional and governmental approval as well as by staff who are technically proficient. All mice were housed in standard vented cages in temperature- and humidity-controlled rooms with 12 h/12 h light/dark cycles (21–22 °C, 30%–60% humidity for normal housing) and free access to water. Male C57BL/6J and ob/ob mice were obtained from the Jackson Laboratory.

1. 2:1 Isocaloric IF Regimen

1. For lean and diet-induced obesity mouse models, prepare either a normal diet (17% fat, ND) or high fat diet (45% fat, HFD).
   NOTE: 60% HFD can be used to induce severe diet-induced obesity; yet, due to the softness of the food pellet, it is relatively difficult to accurately measure daily food intake. An automated continuous measurement system can improve versatility for multiple types of diets.
2. Measure baseline body weight and body composition of each mouse at 7 weeks of age using a scale and EchoMRI, respectively.
   NOTE: Refer to section 3 for body composition measurement.
3. Based on body weight and body composition results, randomly and equally divide 7 week-old male C57BL/6J mice into two groups: ad libitum (AL) and intermittent fasting (IF) groups.
4. Place two to three mice per cage and ensure free access to drinking water.
   NOTE: The number of mice per cage can affect food intake behavior. It is recommended to maintain an equal number of mice per cage in all groups during the study.
5. Provide 1 week of acclimation to the new cage environment and diet before starting the IF regimen.
6. Fasting period: move mice to a clean cage with fresh bedding at 12:00 PM. Do not add food for the IF group, while providing a weighed amount of food to the AL group.
   NOTE: For each fasting cycle, it is important to change cages for both AL and IF groups to ensure that both groups are exposed to the same amount of handling time.
7. After 24 h, measure the weights of mice in both groups and leftover food in AL cages.
   NOTE: Make sure to include the weight of food crumbs on the food hopper and bottom of the cage, especially when using HFD, as mice often remove small pellets or fragments of food from the hopper and keep them near nest sites. The average energy intake per mouse at the end of each 2:1 cycle (3 days) is around 35 kcal, equivalent to ~10 g for a normal diet (3.3 kcal/g) and ~7 g for HFD (4.73 kcal/g).
8. Feeding period: provide a weighed amount of food at 12:00 PM for both AL and IF groups.
9. After 48 h of providing the food, measure the weight of leftover food and mice.
10. Repeat steps 1.6–1.10 for the duration of the study (e.g., 16 weeks).

2. Pair-feeding (PF) Control Group

NOTE: For an IF experiment in which altered feeding behavior is observed in a mouse model (e.g., hyperphagia in ob/ob mice), it is necessary to have a pair-feeding group as a control for proper calorie-independent comparison to IF.

1. For the PF control group, stagger the experiment schedule such that the same amount of food consumed by the IF group is offered to the PF group (Figure 2).
2. Measure the amount of food consumed by the IF group over 2 days of refeeding period.
3. Divide this amount of consumed food in the IF group evenly into three proportions and provide it daily to the PF group at 12:00 PM.
   NOTE: Providing an equal amount of food daily is critical. In the case of mice with hyperphagia, if the pair-fed mice are provided with an amount of food less than their voluntary consumption at once, they will likely consume all provided food and become effectively fasted. This may then prevent proper comparison to IF-treated mice and confound the result.
4. Repeat steps 2.1–2.3 for the duration of the study.

3. Body Composition Analysis

NOTE: Since long-term IF affects body weight in mice, body composition can be measured at appropriate cycles (e.g., every 3 or 4 cycles) using a body composition analyzer to quantify fat and lean mass in live, non-anesthetized mice.

1. Turn on the body composition analyzer.
   NOTE: Before starting the program, leave the machine on for at least 2–3 h to warm up.
2. Run a system test on the body composition analyzer to test its measurement accuracy. If necessary, calibrate the system using canola oil and water samples.
3. Measure the body weight of each mouse.
4. Place the mouse in a small animal cylindric holder.
5. Insert a delimiter to constrain physical movement of the mouse during the measurement and place the holder into the body composition analyzer.
6. Run the scanning program.
   NOTE: It takes approximately 90–120 s to analyze.
7. After measurement, remove the holder from the equipment and bring the mouse back to the cage.
   NOTE: A more detailed protocol can be found in a previous publication⁹.

4. Glucose and Insulin Tolerance Tests

1. For glucose tolerance test (GTT), measure body weight and body composition of each mouse before subjecting to fasting and mark the tail with a permanent marker for easy and rapid indexing.
2. Place mice in new cages without food at 7:00 PM for overnight fasting.
   NOTE: Overnight fasting is the standard protocol, yet due to mouse physiology (e.g., increased glucose utilization after prolonged fasting^10,11), shorter fasting (~6 h) can be used as described for ITT.
3. After fasting 14–16 h (9:00 AM in the following morning), measure body weight and body composition of each mouse and calculate the amount of glucose dosage based on body weight.
   NOTE: To avoid overestimation of glucose intolerance in obese mice, lean mass obtained from the body composition analysis can be used to calculate glucose dosage^12,13.
4. For each mouse, cut the tip of the tail (0.5–1.0 mm) using clean surgical scissors. After wiping off the first drop of blood, draw a fresh drop of blood from the tail and measure baseline fasting blood glucose level with the glucometer.
   NOTE: Additional tail cutting is not required for every blood glucose measurement during GTT or ITT. The wound can be freshened by abrading it with gauze to draw a drop of blood.
5. Subject mice to an intraperitoneal (i.p.) injection of glucose (1 mg/g of body weight).
   NOTE: Based on the objective of an experiment (e.g., examining incretin effects), oral administration of glucose can be performed by oral gavage. The protocol for oral GTT (OGTT) can be found in another study¹⁴.
6. Measure blood glucose from the tail at 0, 5, 15, 30, 60, and 120 min post-glucose injection.
7. After finishing the GTT, provide a sufficient amount of food.
8. For the insulin tolerance test (ITT), remove food at 9:00 AM.
   NOTE: Since both GTT and ITT are stress-inducing experiences for mice that can elevate blood glucose levels and change physiology, it is recommended to perform ITT after providing at least 2–3 days of recovery after GTT experiment.
9. After fasting for 6 h (3:00 PM), measure baseline blood glucose from the tail as described in step 4.4.
10. Subject mice to i.p. injection of insulin (0.65 mU/g of body weight).
11. Measure blood glucose from the tail at 0, 15, 30, 60, 90, and 120 min post-insulin injection.
12. After finishing ITT, provide a sufficient amount of food.

5. Indirect Calorimetry

NOTE: Energy metabolism of IF-treated mice can be further evaluated through indirect calorimetry over a single cycle of IF. This will measure oxygen consumption (VO₂), carbon dioxide production (VCO₂), respiratory exchange ratio (RER), and heat (kcal/h).

1. Turn on the power of the indirect calorimeter system at least 2 h before running the experiment.
   NOTE: This system warm-up is important for accurate measurement.
2. Prepare cages with clean bedding, fill water bottles, and add the pre-weighed amount of chow to the food hoppers.
3. Check the condition of the Drierite and lime soda. If a color indicator of the Drierite appears pink, which indicates that the Drierite has absorbed a high amount of moisture, it is necessary to replace or top with fresh Drierite.
4. Calibrate the system using a gas with the specific composition (0.5% CO₂, 20.5% O₂).
5. Measure body weight and body composition of each mouse, which will be used to normalize VO₂ and VCO₂ data.
6. Gently place one mouse per cage.
7. Assemble metabolic cages, place them in the temperature-controlled environment chamber, and connect to gas lines and activity sensor cable.
8. After setting up the experiment profile by adding appropriate experimental parameters using the software, run the program for measurement. The purpose of the first day's measurement is to provide a period of acclimatization and measure baseline energy metabolism.
9. At 12:00 PM the following day, subject mice to 24 h of fasting by removing food and crumbs from the hopper and bottom of the cage. If necessary, replace with clean bedding.
10. After 24 h, add the pre-weighed amount of chow to the food hopper for the refeeding period.
11. Continue to measure for the next 48 h. Check regularly whether the system is running without hardware or software interruption.
12. After completing measurement, terminate the program and bring mice back to their original cages. Measure the amount of leftover food to examine food intake.
13. The detailed protocol for indirect calorimetry can be found in a previous study⁹.

Results

Figure 1 shows the feeding analyses after 24 h fasting and the comparison between 1:1 and 2:1 intermittent fasting. A 24 h fasting period resulted in a ~10% reduction in body weight, which was fully recovered after 2 days of refeeding (Figure 1A). A 24 h fasting period induced hyperphagia during the subsequent 2 days of refeeding (Figure 1B). Nevertheless, the comparison of energy intake between 1:1...

Discussion

It has been well-documented that IF provides beneficial health effects on various diseases in both humans and animals^{8,15,16,17,18,19}. Its underlying mechanisms, such as autophagy and gut microbiome, have recently been elucidated. The presented protocol describes an isocaloric 2:1 IF regimen in mice for investigating calorie-...

Materials

Name	Company	Catalog Number	Comments
Comprehensive Lab Animal Monitoring System (CLAMS)	Columbus Instruments		Indirect calorimeter
D-(+)-Glucose solution	Sigma-Aldrich	G8769	For GTT
EchoMRI 3-in-1	EchoMRI	EchoMRI 3-in-1	Body composition analysis
Glucometer and strips	Bayer	Contour NEXT	These are for GTT and ITT experiments
High Fat Diet (45% Kcal% fat)	Research Diets Inc.	#D12451	3.3 Kcal/g
High Fat Diet (60% Kcal% fat)	Research Diets Inc.	#D12452	4.73 Kcal/g
Insulin	El Lilly	Humulin R	For ITT
Mouse Strain: B6.Cg-Lepob/J	The Jackson Laboratory	#000632	Ob/Ob mouse
Mouse Strain: C57BL/6J	The Jackson Laboratory	#000664
Normal chow (17% Kcal% fat)	Harlan	#2918
Scale	Mettler Toledo		Body weight and food intake measurement