Development of a Mobile Mitochondrial Physiology Laboratory for Measuring Mitochondrial Energetics in the Field

Summary

We designed and constructed a mobile laboratory to measure respiration rates in isolated mitochondria of wild animals captured at field locations. Here, we describe the design and outfitting of a mobile mitochondrial laboratory and the associated laboratory protocols.

Abstract

Mitochondrial energetics is a central theme in animal biochemistry and physiology, with researchers using mitochondrial respiration as a metric to investigate metabolic capability. To obtain the measures of mitochondrial respiration, fresh biological samples must be used, and the entire laboratory procedure must be completed within approximately 2 h. Furthermore, multiple pieces of specialized equipment are required to perform these laboratory assays. This creates a challenge for measuring mitochondrial respiration in the tissues of wild animals living far from physiology laboratories as live tissue cannot be preserved for very long after collection in the field. Moreover, transporting live animals over long distances induces stress, which can alter mitochondrial energetics.

This manuscript introduces the Auburn University (AU) MitoMobile, a mobile mitochondrial physiology laboratory that can be taken into the field and used on-site to measure mitochondrial metabolism in tissues collected from wild animals. The basic features of the mobile laboratory and the step-by-step methods for measuring isolated mitochondrial respiration rates are presented. Additionally, the data presented validate the success of outfitting the mobile mitochondrial physiology laboratory and making mitochondrial respiration measurements. The novelty of the mobile laboratory lies in the ability to drive to the field and perform mitochondrial measurements on the tissues of animals captured on site.

Introduction

To date, studies designed to measure mitochondrial energetics have been limited to laboratory animals or animals captured near established physiology laboratories, which precluded scientists from performing mitochondrial bioenergetic studies in tissues collected from animals during such activities as migration, diving, and hibernation^1,2,3,4,5,6. While many investigators have successfully measured the basal and peak metabolic rates and daily energy expenditures of wild animals^7,8, the capacity of researchers to measure the performance of mitochondria has remained limited (but see^1,4,9). This is partly due to the need for fresh tissue for isolating mitochondria and a laboratory facility to perform the isolations within about 2 h of obtaining the fresh tissue. Once the mitochondria have been isolated, the mitochondrial respiration measurements should also be completed within ~1 h.

Isolated mitochondrial respiration rates are usually performed by measuring oxygen concentration in a sealed container connected to a Clark electrode. The theory behind this method is founded on the basic observation that oxygen is the last electron acceptor of mitochondrial respiration during oxidative phosphorylation. Therefore, as oxygen concentration falls during an experiment, it is assumed that adenosine triphosphate (ATP) production occurs¹⁰. Consumed oxygen is a proxy for produced ATP. Researchers can create specific experimental conditions using different substrates and initiate adenosine diphosphate (ADP)-stimulated respiration (state 3) by adding predetermined amounts of ADP to the chamber. Following the phosphorylation of the exogenous ADP to ATP, the oxygen consumption rate decreases, and state 4 is reached and can be measured. Furthermore, the addition of specific inhibitors allows information regarding leak respiration and uncoupled respiration to be obtained¹⁰. The ratio of state 3 to state 4 determines the respiratory control ratio (RCR), which is the indicator of overall mitochondrial coupling^10,11. Lower values of RCR indicate overall mitochondrial dysfunction, whereas higher RCR values suggest a greater extent of mitochondrial coupling¹⁰.

As previously stated, the collection of biological material, mitochondrial isolation, and measurement of respiration rates must be completed within 2 h of obtaining tissue. To accomplish this task without transporting animals over large distances to established laboratories, a mobile mitochondrial physiology laboratory was constructed to be taken to field locations where these data can be collected. A 2018 Jayco Redhawk recreational vehicle was converted into a mobile molecular physiology laboratory and named the Auburn University (AU) MitoMobile (Figure 1A). A recreational vehicle was selected because of the built-in refrigerator, freezer, water storage tank and plumbing, electricity powered by 12-volt batteries, gas generator, propane tank, and self-leveling system. Further, the recreational vehicle provides the capability of staying at remote sites overnight for data collection. The front of the vehicle was not altered and provides the driving and sleeping quarters (Figure 1B). Previously installed bedroom amenities (bed, TV, and cabinet) in the rear of the vehicle and the stovetop were removed.

Custom-made stainless-steel shelving and a custom quartz countertop supported by 80/20 aluminum framing were installed in place of the bedroom amenities and stovetop (Figure 1C). The laboratory benches provide adequate space for data collection (Figure 1D). Power consumption of each piece of equipment (i.e., refrigerated centrifuge, mitochondrial respiration chambers, plate readers, computers, homogenizers, scales, portable ultra-freezer, and other general laboratory supplies) was taken into consideration. To support the large voltage and current demands of the centrifuge, the electrical system was upgraded to that of aircraft-grade equipment. An external compartment in the rear of the vehicle was converted into a liquid nitrogen storage bay, which meets the United States Department of Transportation's guidelines for liquid nitrogen storage and transport. This storage unit was constructed with stainless steel and has proper venting to keep any expanding nitrogen gas from leaking into the passenger compartment of the vehicle.

To confirm that the mobile laboratory can be used in mitochondrial bioenergetic studies, mitochondria were isolated, and mitochondrial respiration rates from wild-derived house mice (Mus musculus) hindlimb skeletal muscle were measured. Because Mus musculus is a model organism, the mitochondrial respiration rates of this species are well-established^12,13,14. Although previous studies have documented mitochondrial isolation via differential centrifugation^15,16,17, a brief overview of the methods used in the mobile mitochondrial physiology laboratory methods is described below.

Protocol

The following sections describe the mitochondrial laboratory methods. All animal handling and tissue collection procedures were approved by the Auburn University Institutional Animal Care and Use Committee (#2019-3582).

1. Description of buffers used for data collection

NOTE: These buffers can be prepared in a stationary laboratory and moved to the mobile laboratory prior to the field trip (unless otherwise noted below).

1. Prepare the skeletal muscle mitochondrial isolation buffer with bovine serum albumin (BSA), as seen in Table 1.
   1. Dissolve chemicals in deionized water (~ 90% volume) except for the fatty acid-free BSA. Place the buffer in the refrigerator until the temperature is 4 °C.
   2. Adjust the solution to a pH of 7.5 while maintaining the temperature at 4 °C.
   3. Add the fatty acid-free BSA and bring up the volume to 100%. Aliquot the solution into 50 mL conical tubes. Store this solution at -20 °C until use.
2. Prepare the skeletal muscle mitochondrial isolation buffer without BSA as seen in Table 1.
   1. Dissolve the chemicals in deionized water (~ 90% volume). Place the buffer in the refrigerator until the temperature is 4 °C.
   2. Adjust the solution to a pH of 7.5 while maintaining the temperature at 4 °C.
   3. Bring up the volume to 100%. Aliquot the solution into 50 mL conical tubes. Store this solution at -20 °C until use.
3. Prepare the skeletal muscle resuspension buffer as seen in Table 1.
   1. Dissolve the chemicals in deionized water (~ 90% volume). Place the buffer in the refrigerator until the temperature is 4 °C.
   2. Adjust the solution to a pH of 7.4 while maintaining the temperature at 4 °C.
   3. Bring up the volume to 100%. Aliquot the solution into 50 mL conical tubes. Store this solution at -20 °C until use.
4. Prepare the skeletal muscle respiration buffer as seen in Table 2.
   1. Dissolve the chemicals in deionized water (~ 90% volume) except for the fatty acid-free BSA. Heat the buffer until the temperature is 37 °C.
   2. Adjust the solution to a pH of 7.0 while maintaining the temperature at 37 °C.
   3. Add the fatty acid-free BSA and bring up the volume to 100%. Aliquot the solution into 50 mL conical tubes. Store this solution at -20 °C until use.
5. Prepare the respiration substrates as seen in Table 2.
   1. Ensure that these substrates are made fresh on the day of data collection in 100 mM Tris-HCl, pH 7.4. Store on ice until use.
      ​NOTE: The provided values are to make a sufficiently concentrated solution for enough substrate to be taken up by the mitochondria. The final concentrations of the substrates are 2 mM pyruvate, 2 mM malate, 10 mM glutamate, and 5 mM succinate.

2. Performing mitochondrial isolation (Figure 2)

NOTE: Mitochondrial isolation and mitochondrial respiration measurements are performed in the laboratory bench area of the mobile laboratory, and all solutions should be kept at 4 °C unless otherwise noted.

1. Park the mobile laboratory on flat ground. Turn on the generator and level the vehicle. Extend the slide and set up the equipment.
2. Thaw the desired amounts of buffers.
   NOTE: Generally, 30 mL of skeletal muscle isolation buffer and 10 mL of skeletal muscle isolation buffer without BSA are needed per muscle.
3. Set up and calibrate the mitochondrial respiration chambers to the desired temperature of experiments and current barometric pressure per manufacturer's instructions. See the Table of Materials for specific chambers used in experiments.
4. Euthanize the animal via decapitation.
   NOTE: The current study used decapitation for euthanasia. Some gases, such as carbon dioxide and isoflurane, affect mitochondrial function^18,19,20; these effects should be considered when selecting the best method of euthanasia for each study. Which method should be performed for each study will be determined by the scientific question being asked.
5. Excise skeletal muscle, quickly trim away fat and connective tissue, weigh, and place the muscle in skeletal muscle isolation buffer with BSA (at least 1/10 w/v) (e.g., 1 g of skeletal muscle to 10 mL of buffer).
6. Mince the skeletal muscle with scissors on ice.
7. Transfer the minced tissue to a 50 mL centrifuge tube using a cut 5 mL pipet tip. Homogenize it with a blade (see the Table of Materials) at 50% power for 5 s. Add protease (5 mg/g wet muscle) and digest for 7 min, mixing the solution every 30 s. Terminate the reaction by adding an equal volume of isolation buffer with BSA.
8. Centrifuge the homogenate at 500 × g for 10 min. Transfer the supernatant through double-layered cheesecloth using a cut 5 mL pipet tip into a clean 50 mL centrifuge tube. Centrifuge the supernatant at 3,500 × g for 10 min to precipitate a brown mitochondrial pellet.
9. Pour out the remaining supernatant. Add the same volume of isolation buffer with BSA to the centrifuge tube. Resuspend the mitochondrial pellet with a flexible scraper (policeman) by gently working the mitochondrial pellet off the walls of the centrifuge tube. Centrifuge at 3,500 × g for 10 min.
10. Pour out the remaining supernatant. Add the same volume of isolation buffer without BSA to the centrifuge tube. Resuspend the mitochondrial pellet by gently working the mitochondrial pellet off the walls of the centrifuge tube with a clean policeman. Centrifuge at 3,500 × g for 10 min.
11. Decant the supernatant and resuspend the mitochondrial pellet in resuspension buffer by gently working the mitochondrial pellet off the walls of the centrifuge tube with a clean policeman.
    NOTE: The volume of the resuspension buffer will depend on the size of the mitochondria pellet.
12. Transfer the resuspended mitochondria to a Dounce homogenizer with a cut 1 mL pipet tip. Using the Dounce homogenizer, carefully homogenize the suspension with 4-5 passes.
13. Place the mitochondrial suspension in a labeled 2 mL microcentrifuge tube using another cut 1 mL pipet tip.

3. Mitochondrial respiration measurements (Figure 3)

1. Complex I substrates
   1. Add 945 µL of respiration buffer to the chamber. Ensure that the stirrer is spinning, and the buffer temperature is maintained at 37 °C. Start the recording of the data collection.
   2. After the oxygen concentration has stabilized, add 20 µL of the mitochondria and place the lid on the chamber. In the software, denote that mitochondria were added to the chamber.
   3. Add 10 µL of 1 M glutamate, 10 µL of 200 mM malate, and 10 µL of 200 mM pyruvate to the chamber with individual syringes and wait until the signal stabilizes. In the software, denote that substrates have been added.
      NOTE: These substrates are typically used to measure carbohydrate-driven respiration. For other combinations of substrates to be used to measure fat-driven respiration, see²¹.
   4. Add 5 µL of ADP with a separate syringe and observe the rapid oxygen consumption (state 3). In the software, denote that ADP was added.
      NOTE: Following the phosphorylation of the added ADP, the oxygen consumption rate will plateau to state 4.
   5. After 4 min of state 4 data collection, terminate the recording. Save the data file.
2. Complex II substrates
   1. Add 963 µL of the respiration buffer to the chamber. Ensure that the stirrer is spinning, and the buffer temperature is maintained at 37 °C. Start the recording of the data collection.
   2. After the oxygen concentration has stabilized, add 20 µL of mitochondria and place the lid on the chamber. In the software, denote that mitochondria were added to the solution.
   3. Add 2 µL of 4 µg/µL rotenone followed by 10 µL of 500 mM succinate to the chamber using separate syringes and wait until the signal stabilizes. In the software, denote that substrates have been added.
   4. Add 5 µL of ADP using a separate syringe and observe the rapid oxygen consumption (state 3). In the software, denote that ADP was added.
      NOTE: Following the phosphorylation of the added ADP, the oxygen consumption rate will plateau to state 4.
   5. After 4 min of state 4 data collection, terminate the recording. Save the data file.

Results

The current manuscript investigated the mitochondrial respiration of wild-derived Mus musculus (n = 7, male = 5, female = 2; age = 1.30 ± 0.2 years) in a mobile mitochondrial physiology laboratory (Figure 1). To measure skeletal muscle mitochondrial respiration, the entire hindlimb, thus aerobic and anaerobic muscle, was used for mitochondrial isolation (Figure 2). Examples of raw mitochondrial respiration data are shown in Figure 3...

Discussion

The mobile mitochondrial physiology laboratory enables researchers to isolate mitochondria and measure mitochondrial respiration rates within 2 h of tissue collection at remote field sites. The results presented herein suggest that measurements of mitochondrial respiration made in the AU MitoMobile are comparable to measurements made in a university research laboratory. Specifically, the values for state 3, state 4, and RCR for wild-derived Mus musculus presented here are comparable with previously published res...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
1.7 mL centrifuge tubes	VWR	87003-294
2.0 mL centrifuge tubes	VWR	87003-298
50 mL centrifuge tubes	VWR	21009-681	Nalgene Oak Ridge Centrifuge Tube
ADP	VWR	97061-104
ATP	VWR	700009-070
Bradford	VWR	7065-020
Clear 96 well plate	VWR	82050-760	Greiner Bio-One
Dounce homogenizer	VWR	22877-284	Corning
EGTA	VWR	EM-4100
Filter paper			Included with Hansatech OxyGraph
Free-fatty acid BSA	VWR	89423-672
Glucose	VWR	BDH8005-500G
Glutamate	VWR	A12919
Hamilton Syringes	VWR	60373-985	Gaslight 1700 Series Syringes
Hansatech OxyGraph	Hansatech Instruments Ltd		No Catalog Number, but can be found under Products --> Electrode Control Units
KH2PO4	VWR	97062-350
Malate	VWR	97062-140
Mannitol	VWR	97061-052
Membrane			Included with Hansatech OxyGraph
MgCl2	VWR	97063-152
MOPS	VWR	80503-004
Policeman	VWR	470104-462
Polytron	Thomas Scientific	11090044
Potassium chloride (KCl)	VWR	97061-566
Protease	VWR	97062-366	Trypsin is commonly used; however, other proteases can be used.
Pyruvic acid	VWR	97061-448
Sodium Dithionite	VWR	AA33381-22
Succinate	VWR	89230-086
Sucrose	VWR	BDH0308-500G
Tris-Base	VWR	97061-794
Tris-HCl	VWR	97061-258