Body Composition and Metabolic Caging Analysis in High Fat Fed Mice

Summary

This protocol describes the use of a body composition analyzer and metabolic animal monitoring system to characterize body composition and metabolic parameters in mice. An obesity model induced by high-fat feeding is used as an example for the application of these techniques.

Abstract

Alterations to body composition (fat or lean mass), metabolic parameters such as whole-body oxygen consumption, energy expenditure, and substrate utilization, and behaviors such as food intake and physical activity can provide important information regarding the underlying mechanisms of disease. Given the importance of body composition and metabolism to the development of obesity and its subsequent sequelae, it is necessary to make accurate measures of these parameters in the pre-clinical research setting. Advances in technology over the past few decades have made it possible to derive these measures in rodent models in a non-invasive and longitudinal fashion. Consequently, these metabolic measures have proven useful when assessing the response of genetic manipulations (for example knockout or transgenic mice, viral knock-down or overexpression of genes), experimental drug/compound screening and dietary, behavioral or physical activity interventions. Herein, we describe the protocols used to measure body composition and metabolic parameters using an animal monitoring system in chow-fed and high fat diet-fed mice.

Introduction

Metabolism underpins many aspects of normal cellular, organ, and whole-body physiology. Consequently, in the setting of various pathologies, alterations to metabolism may directly contribute to the underlying condition or may be adversely impacted as a side-effect of the pathology. Traditionally, metabolic research and studies into energy balance have been concentrated on the field of obesity and related conditions such as insulin resistance, pre-diabetes, glucose intolerance, cardiovascular disease, and diabetes. This research is warranted given the escalating prevalence of such conditions worldwide and the individual, societal, and economic costs these conditions inflict. As such, the development of prevention strategies and new therapeutics to target obesity is a continuing goal in research laboratories around the world and preclinical mouse models are heavily relied upon for these studies.

While weighing mice provides a reliable assessment of weight gain or loss, it does not provide a breakdown of the different components that make up whole-body composition (fat mass, lean mass, free water as well as other components such as fur and claws). The weighing of fat pads at the completion of studies once the mouse is deceased provides an accurate measure of different fat depots but can only provide data for a single time point. As a consequence, it is often necessary to enroll multiple cohorts to investigate the development of obesity over time, significantly increasing animal numbers, time, and costs. The use of dual-energy X-ray absorptiometry (DEXA) provides an approach to assess body fat and lean tissue contents and allows the researcher to obtain data in a longitudinal fashion. However, the procedure requires mice to be anesthetized¹, and repeated bouts of anesthesia may impact the accumulation of adipose tissue or impact other aspects of metabolic regulation. EchoMRI utilizes nuclear magnetic resonance relaxometry to measure fat and lean mass, free water, and total water content. This is achievable due to the creation of contrast between the different tissue components, with differences in the duration, amplitude and spatial distribution of generated radio frequencies allowing the delineation and quantification of each tissue type. This technique is advantageous as it is non-invasive, quick, simple, requires no anesthesia or radiation, and, importantly, has been positively validated against chemical analysis².

A key consideration of obesity and related research is the energy balance equation. While fat accumulation is more complicated than purely energy in (food intake) versus energy out (energy expenditure), they are vital factors to be able to measure. Daily energy expenditure is the total of four different components: (1) basal energy expenditure (resting metabolic rate); (2) the energy expenditure due to the thermic effect of food consumption; (3) the energy required for thermoregulation; and (4) the energy spent on physical activity. As energy expenditure generates heat, measuring heat production by an animal (known as direct calorimetry) can be used to assess energy expenditure. Alternatively, measurement of inspired and expired concentrations of O₂ and CO₂, allowing for determination of whole-body O₂ consumption and CO₂ production, can be utilized as a way to indirectly measure (indirect calorimetry) heat production and consequently calculate energy expenditure. An increase in food intake or a decrease in energy expenditure will predispose mice to weight gain and observations of changes in these parameters can provide useful information of likely mechanisms of action in particular models of obesity. A related metabolic parameter of interest is the respiratory exchange ratio (RER), an indicator of the proportion of substrate/fuel (i.e., carbohydrate or fat) that is undergoing metabolism and being utilized to produce energy. Consequently, measurement of food intake (energy consumed) combined with physical activity levels, O₂ consumption, RER, and energy expenditure can provide a broad understanding of an organism's metabolic profile. One method to gather such data is to use a comprehensive laboratory animal monitoring system (CLAMS), which is based on the indirect calorimetry method to measure energy expenditure and has the added capabilities of determining physical activity levels (beam breaks) and food intake via scales incorporated into the measurement chamber.

In this protocol we provide a straight-forward description of the use of a body composition analyzer to assess body composition in mice and a metabolic animal monitoring system to measure aspects of metabolism. Considerations and limitations for these techniques will be discussed as well as suggested methods of analysis, interpretation, and data representation.

Protocol

All experiments described were approved by the Alfred Medical Research Education Precinct Animal Ethics Committee (AMREP AEC) and mice were provided humane care in line with the National Health and Medical Research Council (NHMRC) of Australia Guidelines on Animal experimentation. Animals were administered their prescribed diet and water ad libitum and housed in a temperature-controlled environment (~21 - 22 °C) with a 12 h light and 12 h-dark cycle. Seven week old male mice (on a C57Bl/6J background) were fed either regular normal chow diet (energy content 14.3 MJ/kg, consisting of 76% of kJ from carbohydrate, 5% fat, 19% protein; see Table of Materials) or for the high fat-feeding group, a high fat diet (HFD) (energy content 19 MJ/kg, consisting of 36% of kJ from carbohydrate, 43% fat, 21% protein, Specialty Feeds) for 3 weeks. Body weight and body composition measurements using an EchoMRI machine were made weekly while the metabolic monitoring analysis took place in a CLAMS after 3 weeks of the diet.

1. Body Composition Analyzer Procedure

Note: To function optimally, the EchoMRI 4-in-1 used in this protocol should be contained within a room where the air temperature is stable and does not fluctuate. Ideally this should be constantly monitored. Moving of the machine and interruptions to power should also be avoided if possible. If the power supply has been interrupted and the system has to be restarted, allow at least 2 - 3 h for the machine to warm up before using it again. Before starting, ensure that you are wearing correct personal protective equipment.

1. Prior to scanning mice, perform a system test on the body composition analyzer machine. This involves using a calibration standard (referred to as a canola oil system test sample (COSTS)) to test the precision of the instrument and to ensure there has been no drift in its accuracy.
   1. Open the system software, then click the System Test toolbar button or pressing "Alt + Y" simultaneously.
   2. Before the system test is carried out by the computer, wait for a reminder to verify that the correct COSTS (in this case the mouse-specific COSTS) has been placed inside the gantry of the system ( Figure 1). Once confirmed that this is indeed the case, accept to proceed with the test, which will take few minutes to complete.
2. Once the system test has been passed, continue forward with scanning.
   1. If the system test fails, repeat the system test.
   2. If the machine continues to be out of range (indicating a deviation has occurred), calibration may be necessary to rectify the situation. Complete this by following the prompts or as described in the user manual provided at time of purchase. If the problem persists, check the manual³ or report the issue to the manufacturer's support team and seek further instruction.
3. Place the mice in a small animal specimen holder (long cylinder) to keep them contained while in the machine. To do this, place the holder horizontally, pick up the mouse and insert it into the opening of the cylinder head first. Slowly and carefully bring the holder to the vertical position so that the mouse is at the bottom of the cylinder and ready for analysis.
4. Once within the holder, insert a delimiter to limit the movement of the mouse during the measurement period. In some circumstances, with extremely active mice, it may be necessary to hold the delimiter in place with your fingertip.
   NOTE: Familiarize the mice with placement in the specimen holders prior to their initial analysis to reduce stress. The use of a red colored animal specimen holder can also reduce the potential stress response, as the mice feel that they are in the dark.
5. Within the software, select a folder (folder toolbar) to save the data to and create a file name.
6. If necessary, reduce the amount of random noise in the fat and lean measurements by increasing the number of primary accumulations of the scan. Once the software is initiated, the primary accumulations is set to a recommended default value for general everyday use; unless there is a specific reason to change these parameters, the default settings will give the necessary level of precision to users.
7. If not interested in obtaining data for free water and total water, turn off the water stage by selecting the tab to say no. Doing so will reduce the scan duration significantly and improve throughput.
8. Initiate the scan by selecting "start scan" or by pressing F5 on the keyboard. Enter all relevant data about the animal (e.g., animal ID, body mass, etc.) and press "ok" or F5 to commence the scan, which will take approximately 1 min.
9. After data have been obtained, remove the animal holder containing the mouse from the machine and place the animal back in its cage. Once all animals have been scanned, export the data for further analysis and collation.
10. Before and after use, thoroughly clean the animal holders as per the instructions of the manufacturer. As these holders are constructed from acrylic plastic, isopropyl alcohol and ethyl alcohol should be avoided as they can cause cracking of the holders and/or rapid deterioration of the holder, thereby increasing the likelihood of breakage. Instead, either use warm dishwashing water solution, or, if further disinfectant is required, use F10 (at a 1:125 dilution) or other disinfectant or cleaning sprays (see Table of Materials) and then wipe off.

2. Metabolic Animal Monitoring System Procedure

NOTE: The system requires ~2 h to warm up and stabilize. If the machine has been turned off, it must be switched on to allow the Zirconia cell to be heated to 725 °C. Also we generally place mice in the body composition analyzer a day prior to entering the animal monitoring system to avoid any issues with restraint stress.

1. Ensure the computer attached to the animal monitoring system is turned on and open the control program. Select the "Oxymax Utility" option from the tool menu to initiate the pumps.
2. Fill water bottles with appropriate water, weigh and inspect the health of the mice, and organize food. If measuring food intake in the system, consider powdering the food. Fill the food hoppers by depressing the spring-loaded platform and tip food into the hopper. Ensure that the food hopper and water bottle are completely full to ensure that there is enough food and water to last the allotted experimental time.
3. Check the status of the drierite/desiccant; if using a color indicator, it should be blue and therefore dry, but if it is pink/purple, it has had significant moisture absorbance and should be replaced or topped up.
4. Check the condition of the ammonia trap and soda lime and replace if required. If the ammonia trap is connected two at a time, when the second trap displays signs of a color change, replace the first one. An increase in the CO₂ offset can also signify the need to replace the soda lime.
   NOTE: Desiccant can be dried in an oven and reused, however we follow the recommendations from the manufacturer of the system to use fresh each time.
5. Assemble the chambers. To do this, place the food hopper on the balance, then place the chamber on top with the perforated platform that becomes the floor of the chamber inserted. Carefully place the mouse in the chamber and attach the lid of the system with the front and back clips and secure before positioning the water bottle and fastening. As a precaution, re-check all chamber lids, mice, and water (Figure 2A-D).
   NOTE: Depending on the size of the mice being examined, it may be necessary to adjust the height of the spaces above the food hopper so that the mice have access to the food but not enough space that they can sleep directly on top of the feeder.
6. As it is recommended that the gas sensors be calibrated before each experiment, calibrate the system.
   1. Use a gas of known composition (0.5% CO₂, 20.5% O₂, balance nitrogen). Connect the calibration gas tank to the system via a regulator and hose. Turn on and ensure the tank output pressure is reading 5 - 10 psi.
      Note: Some systems will have a second tank, hose and regulator for use of pure Nitrogen as an "offset" gas. The system we operate instead utilizes soda lime to generate CO₂ free air.
   2. Follow procedures to calibrate both the O₂ and CO₂ sensors. Select "calibration" from the tools menu and sequentially calibrate both the O₂ and CO₂. Before calibrating ensure that 1) sample and reference flows are 0.400LPM, 2) the Zirconia O₂ sensor temperature is 725 °C (± 1 °C), 3) the sample and reference drier and air pumps are on, and 4) the calibration gas is attached and turned on.
   3. If necessary, when calibrating the O₂ sensor, slightly adjust the offset control on the front of the zirconia oxygen sensor to achieve an O₂ ratio value of 1.0000 (± 0.0002). This is to ensure it is within acceptable limits (highlighted in green font in the software display on the computer screen).
   4. After successful O₂ and CO₂ sensor calibration, turn off the calibration gas cylinder and disconnect the hose from the regulator. After calibration, O₂ for reference air (atmospheric) should read 20.92 (± 00.02). If calibration is out of tolerance, repeat, and refer to trouble shooting guides from the manufacturer. Failing this, contact the manufacturer for further instructions.
7. Proceed with experimental set-up. Select "experimental file open" from the experiment menu. Select the appropriate template (e.g., mouse). Under "setup" in the experiment menu define the parameters of the experiment that should be recorded (e.g., mouse ID, weight, group, etc.) De-select any chambers not in use and select the location for the experiment to be saved.
8. Ensure the scales have been tared if measuring food intake and start the capturing of data by selecting "run" in the experiment menu. Data is captured for different lengths of time depending on the phenotype, institutional guidelines on animal isolation, and system usage.
   NOTE: In our hands, the experiment is routinely run for 48 h, with the first 24 h used as acclimatisation to the new environment and the second 24 h used for data analysis. The data collection period is based on how long the investigator wishes to keep their mice singly housed and dependent on animal ethics approval. Alternatively, if provisions exist, mice may be acclimatised in the chambers prior to being placed into the system and connected. Each chamber is measured approximately once every 13 min when a 12 chamber system is in use.
9. Regularly check and monitor the results that are obtained while the mice are in the system to ensure animal welfare and that appropriate data is being collected. Any issue may be able to be identified at this stage and rectified. Check on each mouse every morning and evening when they are in the system.
10. Check the metabolic tab at the top of the data file page for the data collected in real time for each mouse in regards to the oxygen consumption, RER, and energy expenditure. Meanwhile, beam breaks and food consumption data can be located in the activity and feeding tabs, respectively. Check that the "O₂ In" is reading around 20.90 - 20.94, the "CO₂ In" is around 0.040 - 0.050, the RER is between 0.7 and 1, and the flow rate is constant at 0.5 - 0.6 L/min.
11. At regular intervals, check that the mice have access to food and water and that they are consuming each. Ensure they are not demonstrating any signs of distress such as digging at the perforated flooring. Also, monitor the results which are displayed.
12. At the completion of the allocated experimental time, select "stop" from the experiment menu and export the results (as CSV files, File>Export>Generate Subject CSV's) for analysis.
13. Inspect the health of the mice, weigh them and then return to their home cages.
    1. Mice may be hostile towards each other after separation, so monitor once they are housed together again.
    2. Disassemble the cages, remove excess food from hoppers, and tip out any feces, urine, and food from the cages. Submerge bottles and sippers in diluted T-bac solution, soak, and clean the other components in diluted bleach solution. Rinse with clean water and leave to air dry.
14. Calculate metabolic parameters with the software. The software utilizes a number of equations to provide the final data output⁴.
    For calculation of oxygen consumption and carbon dioxide production: Oxygen consumption: VO₂ (LPM)_{= Vi}O_2i - V_oO_2o; Carbon dioxide production: VCO₂ (LPM)_=VoCO_2o-V_iCO_2i
    Where: V_i = the input ventilation rate (LPM), V_o = the output ventilation rate (LPM), O_2i = the O₂ concentration at input, O_2o = the O₂ concentration at output, CO_2i = the CO₂ concentration at input, CO_2o = the CO₂ concentration at output.
    For calculation of RER: RER = VCO₂ / VO₂. Note that protein oxidation was not measured and therefore the RER was not adjusted for this.
    For calculation of energy expenditure: Energy Expenditure: CV = 3.815 + 1.232* RER
    Heat (Kcal/h)) = CV * VO_2. Where: CV is the calorific value (the relationship between heat and the volume of oxygen consumption). This is derived from "The Elements of the Science of Nutrition" referred to as the Lusk Table, comprised by Graham Lusk.

Results

The results seen in Figure 3 display a typical change in body composition parameters upon high fat feeding, as measured via EchoMRI. At baseline there was no difference in any parameter measured (Figure 3A-F). However, after just 1 week of high fat feeding, there was a significant increase in body weight, fat mass, and fat mass percentage in the HFD group (Figure 3A,B

Discussion

Critical steps

The protocols described herein provide an example of ways in which to measure body composition and various metabolic parameters in mice using a body composition analyzer and a metabolic animal monitoring system. For both techniques, it is critically important to ensure that the machines are working optimally, and to do this, it is imperative that the researcher performs a system test for the body composition analyzer and calibrates to a known gas composition fo...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
4 in 1 system	EchoMRI	4 in 1 system	Whole body composition analyser
Canola oil test sample (COSTS)	EchoMRI	Mouse-specific (contact company for cat number)
Animal specimen holder	EchoMRI	103-E56100R
Delimiter	EchoMRI	600-E56100D
12 chamber system	Columbus Instruments	Custom built	Metabolic Caging System; includes control program
Drierite	Fisher Scientific	238988	CLAMS consumable
Calibration gas tank	Air Liquide	Mixed to order	Gas calibration (0.5% CO2, 20.5% O2, balance nitrogen).
Normal chow diet	Specialty Feeds	Irradiated mouse and rat diet
High fat diet	Specialty Feeds	SF04-001
Balance	Mettler Toledo	PL202-S	Balance for weighing mice
TexQ Disinfectant spray	TexWipe
Hydrogen Peroxide cleaning solution	TexWipe	TX684