Robust Mitochondrial Isolation from Rodent Cardiac Tissue

Summary

Bioenergetic and metabolomic studies on mitochondria have revealed their multifaceted role in many diseases, but the isolation methods for these organelles vary. The method detailed here is capable of purifying high-quality mitochondria from multiple tissue sources. Quality is determined by respiratory control ratios and other metrics assessed with high-resolution respirometry.

Abstract

Mitochondrial isolation has been practiced for decades, following procedures established by pioneers in the fields of molecular biology and biochemistry to study metabolic impairments and disease. Consistent mitochondrial quality is necessary to properly investigate mitochondrial physiology and bioenergetics; however, many different published isolation methods are available for researchers. Although different experimental strategies require different isolation methods, the basic principles and procedures are similar. This protocol details a method capable of extracting well-coupled mitochondria from a variety of tissue sources, including small animals and cells. The steps outlined include organ dissection, mitochondrial purification, protein quantification, and various quality control checks. The primary quality control metric used to identify high-quality mitochondria is the respiratory control ratio (RCR). The RCR is the ratio of the respiratory rate during oxidative phosphorylation to the rate in the absence of ADP. Alternative metrics are discussed. While high RCR values relative to their tissue source are obtained using this protocol, several steps can be optimized to suit the individual needs of researchers. This procedure is robust and has consistently resulted in isolated mitochondria with above-average RCR values across animal models and tissue sources.

Introduction

Mitochondria are subcellular organelles that establish cytoplasmic energetic conditions optimized for specific cell functions. While cellular, tissue, and organism-level studies can provide insights into mitochondrial function, isolating the organelles offers a level of experimental control not possible otherwise. Mitochondrial isolations have been performed since the 1940s, allowing mechanistic studies of metabolism and respiration across a variety of cells and tissues^1,2. The historical relevance of mitochondria is also well-documented³. As the main producers of ATP, mitochondria play many key roles that are vital for optimal cellular and organ function⁴. Within the mitochondrial matrix, substrates are oxidized by the TCA cycle, producing reducing equivalents and mobile electron carriers such as NADH and UQH₂^5,6. Cytochrome C is the third main mobile electron carrier in the mitochondrial biochemical reaction network⁷. These molecules are then oxidized by the transmembrane complexes of the electron transport system (ETS) embedded in the inner mitochondrial membrane⁸. Redox reactions of the ETS are coupled with proton translocation from the matrix to the intermembrane space. These processes establish an electrochemical proton gradient that is used to phosphorylate ADP with Pi by F₁F₀ ATP synthase to produce ATP^9,10. The individual processes that occur at each complex can be explored with high-resolution respirometry using Clark-type electrodes or microplate oxygen consumption assays^11,12. Additionally, disease models and treatments using isolated mitochondria can determine the impact or importance of mitochondrial function in the progression of certain pathologies. This has proven fruitful in the field of cardiology, where alterations in fuel and substrate delivery have been used to elucidate how mitochondrial dysfunction influences heart failure^13,14,15,16. Mitochondria are also known to impact the development of other disease states such as diabetes, cancer, obesity, neurological disorders, and myopathies^17,18. Therefore, the use of isolated or purified mitochondria enables mechanistic investigations of oxidative metabolism and ATP production in the source tissue.

There is no shortage of mitochondrial isolation protocols due to their importance in bioenergetic research. Additionally, highly specific methods tailored to subpopulations of mitochondria within tissues and cells can be found^19,20. The basic procedural steps are similar between isolation methods, but variations can be made to buffer composition, homogenization steps, and centrifugation spins to improve the amount and quality of mitochondria. Changes to these aspects are based on the metabolic demand of the tissue, overall organ function, mitochondrial density, and other factors. In tissues such as the liver and skeletal muscle, handheld homogenizers are used to preserve mitochondrial integrity and limit damage to the mitochondrial membranes²¹. However, when isolating from kidneys, some protocols suggest using manually driven homogenization or commercial kits to yield better results²². Although both methods yield functional mitochondria, the quality of the organelles can become compromised by the additional time it takes to complete isolations using these protocols. Centrifugation is also vital to the extraction of mitochondrial protein, as it separates cellular components such as nuclei and other organelles from mitochondria²³. During the isolation process, it is debated whether differential or density-based centrifugation should be implemented to obtain purer isolates²⁴. While density centrifugation can separate mitochondria from organelles of similar specific gravity, such as peroxisomes, it is not well-established if mitochondria from these methods better represent in situ organelle function compared to mitochondria isolated using differential centrifugation. In the field of mitochondrial physiology, density-based centrifugation is preferred and can be easily altered to increase isolate purity. Whether changes to g-forces, centrifugation duration, and the number of centrifugation spins are incorporated should be considered before experimentation due to their influence on mitochondrial purification. Furthermore, mitochondrial resuspension, arguably the most crucial step during isolation, differs greatly between studies, with the use of scraping, vortex-based mixing, or homogenization^23,25,26. Mechanical resuspension of these types can be too abrasive and impair the membrane integrity of mitochondria. For this reason, gentle washing should be performed to correct this. Despite the plethora of modulations and suggested methodologies, there are fewer comprehensive protocols with high reproducibility and adaptability for rodent models.

The methods described herein outline a detailed, robust, and highly reproducible protocol that yields purified, well-coupled mitochondria from small animal cardiac tissue. As demonstrated, this method can be easily adapted to accommodate the specific needs of each experiment and/or laboratory environment for isolating mitochondria from kidneys, liver, and cultured cells. Further modifications can be made to isolate mitochondria from tissues and other animals not listed here. Buffer recipes used for all isolations are provided and can be modified if needed. Similar to other published protocols, motorized homogenization, and differential centrifugation are implemented; however, adjustments are made to both the shearing time and the force at which the samples are centrifuged to consistently deliver high-quality mitochondrial isolates, depending on the isolation source. Notably, this protocol differs from others by using gentle washing via pipetting to resuspend pelleted mitochondria, which helps preserve mitochondrial membrane integrity and maintain the overall functionality of the organelles. Mitochondrial protein is quantified either by total protein determination or by measuring citrate synthase activity. The utility and broad applicability of this isolation method are further supported by the values of respiratory control ratios (RCR) achieved across various organisms and tissue sources.

Protocol

The use and treatment of all vertebrate animals were performed in accordance with approved protocols reviewed and accepted by the Institutional Animal Care and Use Committee (IACUC) at Michigan State University. This protocol was designed using both male and female Hartley albino guinea pigs and Sprague Dawley (SD) rats. For the isolation of cardiac mitochondria from guinea pigs, animals were sacrificed at ages between 4-6 weeks (300-450 g). Cardiac mitochondria from SD rats of both sexes were obtained between the ages of 10-13 weeks (250-400 g). Recipes for buffers are described in Table 1 and should be prepared in advance. Details of the reagents and equipment used in this study are listed in the Table of Materials.

1. Experimental preparation

1. Before starting the isolation, label two 50 mL centrifugation tubes as "Spins 1 and 2" and "Spin 3".
2. Place tubes, freshly thawed isolation buffer (IB), sharp mincing scissors, assembled homogenization probe, and a 5 mL beaker or equivalent-sized container on ice.
3. Place freshly thawed respiration buffer (RB) in an incubator to warm for subsequent respiratory assays.
4. Pre-chill a refrigerated centrifuge to 4 °C.
5. Ensure that all equipment is arranged and 20 µL of 7-15 U/mg protease from Bacillus licheniformis has been added to the tube labeled "Spins 1 and 2".
6. Set up a gravity-dependent pressure system for perfusion via cannulation using Cardioplegia buffer (CB, Table 1), glass cannula, and plastic tubing with a stopcock valve attached.
7. Arrange surgical tools and include proper scissors and forceps for dissection and cannulation of the heart.
   NOTE: All water should be of pure quality (18.2 MΩ·cm)

2. Tissue dissection

1. Inject animals with sterile heparin sulfate intraperitoneally at a dose of 500 U/kg.
2. Allow the animal to sit in the induction chamber for 15 min after heparin administration. During this time, supply 2 L/min of pure O₂ gas to fully oxygenate the animals, calm them, and minimize any stress that may have adverse consequences on tissue of interest.
3. Start anesthesia induction by a continuous flow of isoflurane at 0.5%. After 1 min, increase to 1%. Continue toincrease by 1% every minute until 5% is reached. Once at 5%, wait for 1 min and monitor the animal's breathing pattern.
4. Once breathing has slowed and becomes labored (approximately at the 6.5 min mark) turn off isoflurane and oxygen flow.
5. Quickly remove the animal from the induction chamber and check for appropriate anesthetic depth by squeezing a paw and checking for the corneal reflex. If the animal responds to either stimulus, then place it back in the induction chamber, readminister anesthesia, and repeat the anesthetic depth check.
6. Once the proper depth of anesthesia is reached, quickly decapitate with a guillotine to severe the cervical spine and place the prone body on ice.
7. Make two parallel vertical incisions from the clavicles proceeding along the lateral rib cage down the length of the thorax. Ensure that the incisions are deep enough to cut through the ribs on the lateral thorax, but avoid damaging intrathoracic structures such as the heart or great vessels.
   NOTE: Vertical incision sizes depend on the animal being used. If using guinea pigs and rats, cut to the diaphragm (approximately 6.35 cm for rats and 11 cm for guinea pigs). If using mice, perform a standard thoracotomy²⁷.
8. Expose the heart using a hemostat to displace the anterior thorax and pack the exposed thoracic cavity with ice. This step minimizes warm ischemia time and enhances the viability of the isolated organelles.
9. Use tweezers to bluntly dissect the thymus and pericardium and fully expose the heart.
10. Using forceps, apply gentle inferior traction on the heart to expose and identify the aorta. The aorta is the thickest great vessel branching out from the base of the heart. Other large vessels, such as the pulmonary vein, are noticeably more translucent than the aorta.
11. Cut the aorta approximately 4-6 mm above the aortic root but below the carotid branching points.
12. Cannulate the aorta and perfuse the heart^28,29 with ice-cold cardioplegia (CB) solution using a gravity-dependent pressure head until the coronary arteries are cleared of blood and the organ appears blanched.
    NOTE: For large rodents, a cannula diameter of 1.8-2.2 mm works well, while for smaller rodents, a diameter range of 1.4-1.8 mm is recommended. Retro-perfusion should take no more than 15-30 s for the coronary vessels to clear of blood.
13. Isolate the ventricular myocardium by dissecting away the atria, cartilaginous valvular tissue, and fatty tissues.
14. Place ventricles in a pre-chilled 10 mL beaker containing 0.1-0.2 mL ice-cold IB.
15. Mince tissue with sharp surgical scissors until pieces are approximately 1 mm³.

3. Mitochondrial purification and protein quantification

1. Transfer the minced tissue into the pre-chilled centrifuge tube labeled "Spins 1 and 2" containing the protease solution.
2. Add ice-cold IB to a final volume of 25 mL.
3. Using a motorized handheld rotor-stator homogenizer, disperse the tissue at 18,000 rpm on ice for 20-25 s.
4. Centrifuge homogenized tissue in a tube labeled "Spins 1 and 2" at 8,000 x g for 10 min at 4 °C.
5. Discard the supernatant (which contains protease) by pouring it into the waste carboy and gently rinse the pellet with 5 mL of IB to remove residual protease.
6. After discarding the rinse, resuspend the pellet with fresh ice-cold IB to a final volume of 25 mL by gentle vortex.
7. Centrifuge at 800 x g for 10 min at 4 °C.
8. Remove the supernatant (contains mitochondria) by gently pouring it into a pre-chilled 50 mL tube labeled "Spin 3". While pouring, take care to avoid dislodging the loose upper portion of the pellet. As an alternative, a transfer pipette or stripette can be used to collect the supernatant.
9. Centrifuge the supernatant at 8000 x g for 10 min at 4 °C.
10. Discard the resulting supernatant and retain the mitochondria-containing pellet.
11. Use a lint-free wipe to absorb excess supernatant from the inside wall of the tube, taking care to avoid disturbing the pellet. Keep the pellet at 4 °C on ice as much as possible.
12. To resuspend the mitochondria, add 80 µL of ice-cold IB to the bottom of the tube. Gently resuspend the pellet by repeatedly washing IB over the pellet.
13. To avoid creating bubbles, set a micropipette to aspirate and deliver between 40-60 µL of volume.
14. As the mitochondrial pellet disperses, increase the micropipette volume to efficiently resuspend the pelleted mitochondria. Avoid touching the pellet with the pipette tip, and avoid making bubbles.
15. Once resuspended, transfer the mitochondria to a pre-chilled microcentrifuge tube and label it as stock mitochondria. Make a note of the total resuspension volume.
16. To determine the mitochondrial protein concentration in the sample, conduct a total protein assay using the well-known BCA or Bradford protein assays as defined by the manufacturer's instructions.
    NOTE: An alternative or complementary strategy to assess yield is to determine the citrate synthase activity. For reference, mitochondrial content can be quantified by following the protocol described in Vinnakota et al.³⁰.

4. Quality control checks

1. In a pre-chilled microcentrifuge tube, dilute the mitochondrial stock to the desired working concentration with IB.
   NOTE: Mitochondrial stocks are diluted to 40 mg/mL to work at 0.1 mg/mL final concentration for respirometry assays when using isolates from cardiac tissue.
2. Rinse oxygraph chambers, stoppers, and microliter syringes ten times with distilled water to clean them before use in respirometry assays.
3. To test the viability and quality of mitochondrial isolates, load 2.3 mL of respiration buffer (RB) into oxygraph chambers and allow the oxygen consumption signal to equilibrate at 37 °C for about 10 min or when the rate is near 0.
4. Once the signal is equilibrated, push down the stoppers and aspirate the excess buffer that emerges from the capillary of the stopper.
5. Add 1 mM EGTA using a microliter syringe to chelate any residual calcium in the buffers or mitochondrial sample.
6. To fuel respiration, add 5 mM sodium pyruvate and 1 mM L-malate.
7. Following the addition of substrates, add a bolus of diluted mitochondria to achieve working concentration and allow respiration to occur for 5 min. This period is termed LEAK or State 2 respiration.
8. At the 5 min mark, add a bolus of 500 µM ADP to initiate State 3 respiration, also termed OXHPOS, and allow the mitochondria to respire until anoxia.
9. Calculate the respiratory control ratio (RCR) by dividing the maximal rate of oxygen consumption during State 3 by the rate of oxygen consumption just before the addition of ADP in State 2 (see Figure 1).
   NOTE: An alternative RCR expression of 1 - 1/RCR can also be used as a metric of quality, which is bounded between 0 and 1; however, it makes it difficult to differentiate quality using this metric when the RCR > 10 (see Figure 2).
10. Rinse chambers and stoppers 10 times with pure water to clean the oxygraph for subsequent assays. If respirometry is complete, fill chambers with 70% ethanol and place stoppers in chambers until the next use.

Results

Upon completion of mitochondrial isolation, the quality and functionality of the isolates should be tested via quantifying rates of oxygen consumption (J_O2) using high-resolution respirometry. To do so, mitochondrial stocks were diluted to 40 mg/mL to allow for working concentrations of 0.1 mg/mL in 2 mL of RB for all respirometry assays using isolated cardiac mitochondria. Respiration was fueled by 5 mM sodium pyruvate and 1 mM L-malate in the presence of 1 mM EGTA, a calcium chelator, and was allowe...

Discussion

Adhering to the methods concisely described in this protocol will ensure the isolation of well-coupled mitochondria from the cardiac tissue of small rodents, in addition to other tissue types and sources. Overall, the process should take a total of 3-3.5 h, during which all animal tissue, samples, and isolates should remain on ice at 4 °C as much as possible to limit degradation. This procedure is robust and can be altered in several ways to better fit experimental goals and models utilized. One modulation that can ...

Materials

Name	Company	Catalog Number	Comments
1.7 mL microcentrifuge tubes
10 mL glass beaker			For organ disection and mincing
50 mL centrifuge tubes			Centrifugation
Adenosine 5'-diphosphate monopotassium salt dihydrate	Sigma	A5285	Respirometry assays
BSA Protein Assay Kit	Thermo Scentific	PI23225	Mitochondrial protein quantification
Dextrose	Sigma	DX0145	For buffer (CB)
Ethylene glycol-bis(2-amino-ethylether)-N,N,N',N'-tetraacetic acid	Sigma	E4378	For buffers (CB and IB) and respirometry assays
Glass cannula	Radnoti		Guinea pig and rat heart perfusion
Heparin sodium porcine mucosa	Sigma	SRE0027-250KU	Animal IP injection
High-resolution respirometer			Clark-type electrode; oxygraph with 2 mL chambers
Induction chamber
Isoflurane	Sigma	792632	Anesthetic
L-malic acid	Sigma	02288-50G	Respirometry assays
Magnesium chloride hexahydrate	Sigma	M9272	For buffer (RB)
Mannitol	Sigma	MX0214	For buffer (IB)
Microliter syringes			Sizes ranging from 5–50 µL
Microplate reader			Must be able to incubate at 37 °C
MOPS	Sigma	475898	For all buffers
O2 tank
OMNI THQ Homogenizer	OMNI International	12-500	Similar rotor stator homogenizers will work
pipettes			Volumes of  2–20 µL; 20–200 µL; 200–1000 µL
Potassium chloride	Sigma	P3911	For buffers (RB and CB)
Potassium phosphate dibasic	Sigma	795496	For buffers (IB and RB)
Protease from Bacillus licheniformis	Sigma	P5459
Sodium chloride	Sigma	S9888	For buffer (CB)
Sodium pyruvate	Fisher bioreagents	BP356-100	Respirometry assays
Sucrose	Sigma	8510-OP	For buffer (IB)
Surgical dissection kit			Depends on animal and tissue source
Tabletop centrifuge			Must cool to 4 °C