Investigating the Mitochondrial Permeability Transition Pore in Disease Phenotypes and Drug Screening

Abstract Mitochondria act as ‘sinks’ for Ca2+ signaling, with mitochondrial Ca2+ uptake linking physiological stimuli to increased ATP production. However, mitochondrial Ca2+ overload can induce a cellular catastrophe by opening of the mitochondrial permeability transition pore (mPTP). This pore is a large conductance pathway in the inner mitochondrial membrane that causes bioenergetic collapse and appears to represent a final common path to cell death in many diseases. The role of the mPTP as a determinant of disease outcome is best established in ischemia/reperfusion injury in the heart, brain, and kidney, and it is also implicated in neurodegenerative disorders and muscular dystrophies. As the probability of pore opening can be modulated by drugs, it represents a useful pharmacological target for translational research in drug discovery. Described in this unit is a protocol utilizing isolated mitochondria to quantify this phenomenon and to develop a high‐throughput platform for phenotypic screens for Ca2+ dyshomeostasis. © 2019 The Authors. This is an open access article under the terms of the Creative Commons Attribution License, which permits use, distribution and reproduction in any medium, provided the original work is properly cited.


INTRODUCTION
As a universal second messenger, Ca 2+ plays a critical role in a wide range of cellular processes. At rest, intracellular [Ca 2+ ] gradients are tightly regulated; whereas extracellular [Ca 2+ ] is at ß1 mM, cytosolic [Ca 2+ ] ([Ca 2+ ] c ) and mitochondrial [Ca 2+ ] ([Ca 2+ ] m ) are maintained at close to 100 nM. This unique electrochemical gradient allows the low [Ca 2+ ] c to undergo a substantial proportional increase, with a small and therefore energetically inexpensive absolute change, in response to a stimulus. Such changes in [Ca 2+ ] c are a fundamental aspect of cell physiology in all tissues, underlying excitation contraction coupling, secretion, and motility (Denton & McCormack, 1985;Duchen, 1992;Duchen, Leyssens, & Crompton, 1998;Prudent et al., 2016).

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Ca 2+ signaling is subverted in multiple pathologies to drive cell death through opening of the mitochondrial permeability transition pore (mPTP). Although opening of the pore is triggered primarily by an increase in mitochondrial Ca 2+ load, it can also be induced by oxidative stress. Pore opening leads to enhanced permeability of the inner mitochondrial membrane (IMM) to solutes of ࣘ1.5 kDa, resulting in membrane rupture, bioenergetic failure, and ultimately cell death. Seminal studies by Crompton and colleagues in the 1980s revealed that pore opening could be inhibited by cyclosporin A (CsA), suggesting that the mPTP represents a viable drug target (Crompton, Ellinger, & Costi, 1988). The association of this phenomenon with disease has resulted in attempts to develop targeted therapies using screening assays to identify compounds that affect mPTP function (Briston, Selwood, Szabadkai, & Duchen, 2019). Although cell-permeant fluorescent dyes can assist in capturing mPTP opening in an intact cell, the results obtained can be variable, as protocols both to induce and to inhibit mPTP opening have proven to be unreliable. mPTP opening has long been studied in isolated mitochondria through very robust protocols that rely on measurements of light scattering or fluorescence measurements following bolus additions of calcium.
In this unit, mitochondrial Ca 2+ buffering is measured in isolated mitochondria (Basic Protocol 1) using a Ca 2+ -sensing fluorescent extra-mitochondrial dye (Basic Protocol 2). Often, healthy mitochondrial function is pivotal to Ca 2+ uptake, and therefore, it is crucial to ensure that the mitochondria are not damaged during isolation.
This unit details several protocols to allow the user to assay the mPTP in isolated mitochondria. Basic Protocol 1 involves isolation and purification of mitochondria from a biological sample using differential centrifugation to obtain functional mitochondria that can buffer Ca 2+ . Mitochondrial quality and functional integrity can be assessed using support protocols for the following: measurement of mitochondrial membrane potential ( m ), which is essential for Ca 2+ uptake, using the fluorescence indicator rhodamine-123 (Support Protocol 1); measurement of the oxygen consumption rate (OCR) using a Clark-type electrode (Support Protocol 2); and immunoblotting for various mitochondrial proteins as an additional surrogate for mitochondrial content and function (Support Protocol 3). These quality-control procedures are crucial during initial optimization of the isolation technique. An additional support protocol (Support Protocol 4) details cryopreservation of mitochondria using an osmolyte, trehalose, often necessary for highthroughput drug screens of compound libraries where large quantities of mitochondria might be needed regularly for multiple assays. Finally, Basic Protocol 2 describes a fundamental experiment to assay the capacity of mitochondria to accumulate Ca 2+ using Ca 2+ -sensitive fluorescent dyes and how this might vary between genotypes or treatments. The Alternate Protocol allows multiplexing of this approach with assessment of mitochondrial swelling. A workflow for the protocols is shown in Figure 1.

Figure 1
Experimental workflow depicting the order in which each protocol, whether basic or support, needs to be executed. The solid lines indicate pathways that are necessary, whereas the dashed lines indicate pathways that are optional. The information needed to determine the appropriate sequence of experiments is specified in further detail in the text. NOTE: The entire procedure should be conducted on ice, using refrigerated equipment maintained at 4°C, or in a cold room.

Dissecting scissors (World Precision
NOTE: Please ensure that you follow the appropriate guidelines set by your institution, in line with national regulations, when handling animals.
In our procedure, we sacrifice the animal using manual cervical dislocation in compliance with the guidelines set in the Animals (Scientific Procedures) Act 1986, followed by decapitation using scissors, as this minimizes damage to the mitochondria. For most experiments, if testing ࣘ25 different conditions, the liver from one adult mouse (3 to 6 months of age) is sufficient.
Alternatively, cells could be used. The number of cells needed for mitochondrial isolation will vary based on cell type. For example, the yield of mitochondria from 10 million HeLa cells or a fully confluent 10-cm culture dish is ß500 µg, which is enough for 10 technical replicates. We suggest isolating a sufficient amount of mitochondria for 12 replicates for a basic experiment, as shown in Figure 2A, as well as additional mitochondria for functional validation of the isolation technique.

Figure 2
Schematic and graphical representation of Support Protocols 1 and 2. (A) Plate map depicting the minimum conditions required to assess m using rhodamine-123 in dequench mode in a fluorescence multiwell plate reader. (B) Oxygen consumption rate (OCR) measurements using isolated mitochondria and an Oroboros instrument. The blue line and left y-axis depict the oxygen concentration in nmol/ml. The red line and the right y-axis report the slope of the blue line, that is, the OCR per milligram of protein, in pmol/s/mg. Drug additions are specified in the order that they should be performed.
2. Place animal on its dorsal side. Cut through abdomen using dissecting scissors and tweezers. Locate liver, between the ribcage and the gut. Dissect out entire liver, ensuring that all lobes are removed (Lampl, Crum, Davis, Milligan, & Del Gaizo Moore, 2015).
3. Place liver in 20 ml ice-cold isolation buffer in a 50-ml Falcon tube on ice as quickly as possible (<10 min).
4. Transfer liver from the tube to a 100-ml glass beaker. Rinse liver in ice-cold DPBS by pouring the DPBS on the liver and then carefully decanting it. Submerge washed tissue in DPBS and mince it with scissors into rice grain-sized pieces.
5. Repeat rinses with DPBS five times, until the blood is removed from the liver.
6. Weigh liver tissue and add isolation buffer with 1 mM PMSF, with the volume in milliliters equaling twice the tissue's weight in grams.
For example, use 10 ml isolation buffer for 5 g tissue.
7. Transfer tissue with the isolation buffer into the glass tube of a tissue homogenizer. Macerate tissue using an electric hammer drill for 10 to 20 plunges to obtain a uniform homogenate that contains no visible pieces of tissue.
8. Transfer homogenate to a 50-ml Oak Ridge ultracentrifuge tube and centrifuge 10 min at 800 × g, 4°C, to generate a nuclear pellet.

MEASUREMENT OF MITOCHONDRIAL MEMBRANE POTENTIAL
Healthy mitochondria maintain a negative m across the IMM by pumping protons into the intermembrane space. This requires energy from sequential reduction of components of the electron transport chain, with oxygen as the final acceptor. This electrochemical proton gradient is essential for ATP production by ATP synthase. It is also crucial for electrogenic uptake of Ca 2+ . Therefore, this mitochondrial property can be assessed using cationic fluorescent dyes such as TMRM or rhodamine-123. Rhodamine-123 is used in the protocol detailed below. This protocol can be carried out immediately after Basic Protocol 1, before proceeding to Basic Protocol 2 or the Alternate Protocol.  (Duchen, Surin, & Jacobson, 2003).

Materials
4. Resuspend mitochondria in the MSK+ containing rhodamine-123 to a final concentration of 500 µg/ml. Add 45 µl mitochondrial suspension to each well of a 96-well black opaque plate in triplicate for each condition (see Fig. 2A for an example plate design) and incubate for 30 min in a 37°C incubator.
5. Prepare two 15-ml Falcon tubes with MSK+ buffer, with one containing 10 µM FCCP (from 1 mM stock in ethanol) and the other without FCCP (as a vehicle control).
Five microliters of 10 µM FCCP (final concentration: 1 µM) or the vehicle control is injected at cycle 5 to depolarize the mitochondria.
6. Clean plate-reader injector with 190-proof ethanol and deionized water before priming the two pumps with 1 ml of the respective solutions from step 5.
7. Obtain results from samples using the plate reader once every minute until cycle 5. Continue to measure for five more cycles after addition of FCCP/vehicle control.
The light source is a high-energy xenon lamp. The emission from the excited sample is collected using a combination optic containing two liquid-filled light guides for measuring fluorescence intensity. An excitation filter of 485 nm (bandpass: 12) is used, and the emitted light is detected with a side-window photomultiplier tube after passing through a 520-nm emission filter. Integrated injectors are used for the compound additions.
8. Collect data after completion of the five cycles and plot fluorescence intensities against time using Excel or equivalent software.

MEASUREMENT OF OXYGEN CONSUMPTION RATE
Healthy coupled mitochondria utilize oxygen at complex IV of the electron transport chain to enable ATP production. Therefore, in the presence of appropriate substrates, an oxygen electrode can be employed to measure the OCR to gauge the health of mitochondria. An Oroboros Power O2k-Respirometer is used in the protocol below; this instrument contains two chambers where dissolved oxygen is measured by a Clark-type polarographic oxygen electrode as an amperometric signal that is in turn converted into a voltage. The data obtained from this protocol provide an estimate of the efficiency of oxidative phosphorylation within the isolated mitochondria (Basic Protocol 1), a parameter that should remain consistent across different isolations. This protocol can be used in conjunction with Support Protocol 1 to ascertain the viability of the mitochondria obtained from Basic Protocol 1.
The temperature and stirring can be set using the DatLab software controlling the respirometer.
2. Add 100 to 300 µg mitochondria to each chamber and close stoppers to create a closed O 2 system.  Figure 2B for an example trace to complement the following steps.

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The integrity of the outer mitochondrial membrane (OMM) is tested by observing the effect of cytochrome c addition on the OCR and comparing this OCR to State 4 respiration. An increase of <10% indicates an intact OMM. This is an important step in the functional validation of the mitochondrial isolation technique.

Add 5 mM oligomycin complex in DMSO (final concentration, 5 µM).
This step inhibits ATP synthase, thereby identifying the OCR required for ATP production. A substantial decrease in the OCR indicates an intact IMM, with minimal leakage of protons across this barrier, and identifies the rate of oxygen consumption required to sustain ATP turnover.
8. Titrate FCCP (sequential additions of 0.5 µM from a stock of 1 mM FCCP in ethanol) until maximal uncoupled respiration is attained. Add 5 mM antimycin A, a complex III inhibitor, in ethanol to a final concentration of 2.5 µM to determine the non-mitochondrial OCR.
9. Analyze data using the DatLab software after acquisition.
In brief, the average value of the OCR (see red line in Fig. 2B) after stabilization following drug addition can be extracted, and thus, the effect of the drugs can be compared across different isolations.
These assay procedures should be conducted routinely to ensure consistent and accurate mitochondrial isolation, such as every 3 to 5 isolations. A representative trace with expected values is shown in Figure 2B. In particular, ΔΨ m and the State 4 OCR should be reproducible across the preparations from the same mitochondrial source on different days, that is, within a 10% error margin.

IMMUNOBLOTTING FOR MITOCHONDRIAL PROTEINS
This protocol summarizes a procedure to quantify mitochondrial proteins using immunoblotting, thus validating the mitochondrial isolation (Basic Protocol 1) by probing for mitochondrially enriched proteins. As loss of cytochrome c from the mitochondria indicates a ruptured OMM, it is possible to diagnose problems with isolation by quantifying the levels of cytochrome c retained in the mitochondrial pellet. Other proteins that can be investigated by immunoblotting include TOM20, VDAC (OMM), subunits of the components of the electron transport chain, and subunits of ATP synthase. Shown below are a few key steps for western blotting; this procedure can be adapted for different equipment and reagents.
The sample buffer is supplied as a 4× stock, and the volume needed will be determined by the volume of mitochondrial suspension used.
Critically, a lower temperature is used to preserve the structure of target epitopes of the mitochondrial proteins.

Load samples and protein standards into the wells of a NuPAGE 4-12% Bis-Tris
Protein Gel immersed in NuPAGE MOPS SDS Running Buffer and separate proteins using a protein gel electrophoresis chamber system and a power supply according to the manufacturers' instructions.
Ideally, the gel is run at 150 V for 45 min (the dye front should have run 80% down the gel) before moving on to step 3. The same mitochondrial sample can be run in multiple wells so that multiple primary antibodies can be used to probe for different proteins. The user should probe for cytochrome c at a minimum as a measure of an intact OMM in the mitochondria in the suspension.
3. Activate an Immobilon-P PVDF membrane with methanol for 1 min. Transfer proteins from the gel to the PVDF membrane in NuPAGE Transfer Buffer with 20% methanol using a semi-dry blot transfer system at 20 V for 1 hr.
4. Incubate membrane in 5% skim milk in TBS-T for 1 hr to block the membrane.
5. Incubate membrane with the appropriate primary antibody overnight at 4°C.
6. Wash blots three times with TBS-T for 5 min each before incubating them with the corresponding HRP-conjugated IgG secondary antibody for 1 hr at room temperature.
7. Wash blots with TBS-T as in step 6 and visualize protein bands using a chemiluminescent reagent and a blot imaging system.
8. Use densitometry analysis via ImageJ2 software to determine the relative expression levels of the protein across samples.

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Current Protocols in Pharmacology 2 were satisfactory as a positive control in comparison of protein levels. Immunoblotting can also provide a parallel investigation to identify underlying causes of differences in ΔΨ m, the OCR, or Ca 2+ retention capacity. Together, Support Protocols 1 to 3 provide both quality-control checks and additional assessment of mitochondrial physiology.

CRYOPRESERVATION OF ISOLATED MITOCHONDRIA
Because of the time needed and batch-to-batch variability, the mitochondria isolation procedure (Basic Protocol 1) can be the rate-limiting step in high-throughput assessment of mitochondrial permeability transition. Long-term storage of isolated mitochondria is advisable to improve assay efficiency and consistency. Because freeze-thaw cycles can disrupt both the OMM integrity of mitochondria and bioenergetic function, cryopreservation of mitochondria with intact OMM integrity and biological functions is accomplished using trehalose, a naturally occurring osmolyte (Yamaguchi et al., 2007). Using mitochondria isolated as described in Basic Protocol 1, execute the protocol described below to prepare tissue for long-term storage and future use.

Additional Materials (also see Basic Protocol 1)
Wash buffer (see recipe), 4°C Homogenization buffer (see recipe) with cOmplete Protease Inhibitor, 4°C (Roche, 04693159001) Liquid nitrogen 1. Place tissue (or cells; see Basic Protocol 1, step 1) in ice-cold wash buffer in a 100-ml glass beaker and mince with dissecting scissors.
2. Wash tissue with wash buffer three times to remove blood.
3. Drain wash buffer and replace it with ice-cold homogenization buffer with cOmplete Protease Inhibitor. Homogenize using a tissue homogenizer as described in step 7 of Basic Protocol 1, once again using a volume in milliliters equaling twice the tissue's weight in grams.
4. Transfer homogenate to a 50-ml Oak Ridge ultracentrifuge tube and centrifuge 10 min at 800 × g, 4°C, to remove the nuclear pellet.
6. Wash mitochondrial pellet by adding the same volume of homogenization buffer as in step 3 and repeating step 5.
7. Resuspend pellet in 1 ml homogenization buffer and determine protein concentration using a Pierce BCA Protein Assay Kit (see Basic Protocol 1, step 11).
8. Using liquid nitrogen, snap freeze mitochondrial suspension at a concentration of 50 mg protein/ml in homogenization buffer and store frozen samples at −80°C for ࣘ7 months (Briston et al., 2016).
For subsequent use, thaw the mitochondria at 37°C in a water bath and maintain them on ice until functional analysis.

Investigation of Calcium Retention Capacity of Mitochondria as a Function of Amount of Ca 2+ Buffered Until Permeability Transition
This protocol is designed to determine the capacity of the isolated mitochondria (Basic Protocol 1) to accumulate Ca 2+ until the mPTP opens. Temporal fluorescence

Figure 3
Assessment of the minimum conditions needed to determine the Ca 2+ retention capacity of the isolated mitochondria, performed in triplicate. The mitochondria-free condition provides a measure of the total Ca 2+ added, and the Ca 2+ -free addition helps define background fluctuations. Cyclosporin A is a cyclophilin D-dependent inhibitor of mitochondrial permeability transition pore opening, and dimethyl sulfoxide (DMSO) is the vehicle control for the cyclosporin A. A positive control for pore inhibition, such as cyclosporin A, is needed when using the assay for identifying drug candidates.
measurements of Ca 2+ -sensing dyes in the extra-mitochondrial buffer provide a simple and robust readout. An increase in fluorescence intensity in response to Ca 2+ addition is closely followed by a decrease in fluorescence intensity as a result of an increase in mitochondrial Ca 2+ uptake. Sequential additions of Ca 2+ ultimately reach a threshold that results in opening of the mPTP, failure of the mitochondria to accumulate additional Ca 2+ , and Ca 2+ release from the mitochondria. This results in a rapid increase in fluorescence intensity. This protocol uses a fluorescence plate reader with an integrated injector system for adding sequential Ca 2+ pulses. For the purposes of drug or phenotypic screening, we recommend using CsA as a positive control for pore inhibition. We recommend using 20 ml MSK+ for a basic experiment, as described in Figure 3, and scaling up as needed for additional conditions.

Figure 4
Protocol when using a BMG Labtech Fluostar with an injection every 10 cycles, beginning from cycle 5. This sequence is repeated three times, using the inbuilt script mode, to administer a total of 12 Ca 2+ injections.

Succinate acts as a substrate for complex II, and rotenone inhibits complex I, thereby preventing reverse electron transfer through complex I and blocking accumulation of oxaloacetate, a known inhibitor of complex II activity.
3. Add 1 mM Fluo-5N (fluorescent dye) in DMSO to the MSK+ to a final concentration of 1 µM.
A single 10-µl sample of 100 µM CaCl 2 is added beginning at cycle 5, and this is repeated every 10 cycles thereafter for a total of 12 additions (Fig. 4). This results in incremental accumulation of ß10 µM Ca 2+ . Please note that the capacity of mitochondria to accumulate Ca 2+ can vary based on the source of the mitochondria, that is, different tissues or cell lines, and the concentrations of the Ca 2+ additions should be titrated accordingly to allow resolution of pore opening between controls and treatments that delay pore opening. This step can be performed in a pilot experiment, where ideally a concentration that causes pore opening between the third and ninth Ca 2+ additions should be used.

Clean integrated injector of the plate reader with 190-proof ethanol and deionized
water before priming the pump with 1 ml of the CaCl 2 solution.
6. Resuspend mitochondria in the MSK+ containing Fluo-5N from step 3 to a final protein concentration of 500 µg/ml. Add 100 µl mitochondrial suspension to each well of a 96-well clear plastic plate in triplicate for each condition (Fig. 3).

Figure 5
Typical absorbance curves for mitochondria isolated from mouse liver. In this example, a single bolus of Ca 2+ is added (arrow) to trigger Ca 2+ -induced pore opening. A loss of absorbance is observed compared to the condition with no Ca 2+ addition. Cyclosporin A (CsA) can rescue this phenotype. Because the swelling assay lacks sensitivity compared to the Ca 2+ retention capacity assay, a subtle difference in pore desensitization might be missed if using this assay alone.
7. Analyze samples with the plate reader, adjusting the gain settings using a well containing only the CaCl 2 solution as a maximum reading.
The light source is a high-energy xenon lamp. The emission from the excited sample is collected using a combination optic containing two liquid-filled light guides for measuring fluorescence intensity. An excitation filter of 485 nm (bandpass: 12) is used, and the emitted light is detected with a side-window photomultiplier tube after passing through a 520-nm emission filter. Integrated injectors are used to perform the Ca 2+ additions.
8. Analyze data using a simple spreadsheet in Excel or equivalent software, tracking the raw data for changes in fluorescence intensity over time. Given that the area under the curve for each condition represents the total Ca 2+ buffered, calculate mitochondrial Ca 2+ uptake as a fraction of the total Ca 2+ ('buffering capacity') added: Following this, perform comparisons to control untreated conditions by quantifying shift in the Ca 2+ threshold for pore opening as a percentage change: buffering capacity − buffering capacity (untreated) buffering capacity (untreated) × 100

Multiplexing of Detection of Ca 2+ Dynamics and Mitochondrial Swelling
Mitochondrial permeability transition is accompanied by mitochondrial swelling due to loss of IMM integrity. This can be measured as a decrease in light scattering. Historically, this technique was used to identify inhibitors of permeability transition. However, inhibition of mitochondrial Ca 2+ uptake can also prevent mitochondrial swelling, resulting in false positives. Nevertheless, simultaneously measuring absorbance and Ca 2+ retention capacity can help distinguish between inhibitors of mitochondrial Ca 2+ uptake and inhibitors of pore opening.
The experimental setup is the same as that described in Basic Protocol 2 and does not require additional material. The additional measurement of absorbance is conducted simultaneously at 540 nm, and the samples would need to be set up in the plate as depicted in Basic Protocol 2. A decreased sensitivity to permeability transition is indicated by a reduced loss of absorbance compared to the control (Fig. 5). Pore opening in the Ca2+ retention capacity assay is mirrored by a steep negative gradient in the absorbance curve.

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This assay makes possible the measurement of two aspects of permeability transition: that is, mitochondrial swelling due to influx of water and unregulated mitochondrial Ca 2+ release. These data provide the user with additional confidence in the results with little extra effort in setting up the experiment.

COMMENTARY Background Information
Mitochondria play a central role in [Ca 2+ ] c signaling. They accumulate Ca 2+ in an electrogenic manner, mediated by the mitochondrial calcium uniporter complex (MCU) (Baughman et al., 2011). Increased [Ca 2+ ] m increases the activity of the three rate-limiting enzymes of the tricarboxylic acid cycle, which in turn upregulates the production of NADH. This subsequently increases the rate of oxidative phosphorylation and ATP generation by providing reducing power for the electron transport chain (Griffiths & Rutter, 2009).
The mPTP was initially described in the 1970s by Haworth and Hunter as a highconductance pathway in the IMM that collapses m . Numerous studies have demonstrated protection of tissues by CsA, most notably in the context of cardiac reperfusion injury (Andreeva, Tanveer, & Crompton, 1995;Duchen et al., 1998;Hausenloy, Duchen, & Yellon, 2003).
Although it has been suggested that transient opening of the pore occurs under physiological conditions, most studies have focused on its involvement in cell death (Korge et al., 2011). As a result of this work, the mPTP has been implicated in many disease pathologies associated with necrotic cell death, ischemia/reperfusion injury, neurodegenerative disorders such as Alzheimer's and Parkinson's diseases, and several forms of muscular dystrophy and myopathy (Devalaraja-Narashimha, Diener, & Padanilam, 2009;Du et al., 2008;Dube et al., 2012;Hausenloy et al., 2003;Javadov et al., 2003;Millay et al., 2008;Palma et al., 2009;Thomas et al., 2012). The role of the mPTP has been most thoroughly documented in ischemia/reperfusion injury, particularly in tissues with a high metabolic demand, such as the brain, kidney, liver, and heart.
Although CypD is a known regulator of the mPTP and is therefore a druggable target, the uncertainty surrounding the molecular identity of the core components has been a significant hurdle in drug design and downstream validation. This has also impeded research into the involvement of the mPTP in disease and possible therapeutic interventions. Nevertheless, pharmacological targeting of elements thought to form the pore has provided positive results (for a review, see Briston et al., 2019).
As isolated mitochondria lack the physiological context of intact cells and tissues, functional validation of in vitro data in higher-order systems is needed before selecting a clinical candidate. Cell-based assays of the mPTP do not readily lend themselves to high-throughput drug discovery programs. Among these assays is the calcein/cobalt technique, which can be used to probe pore opening at the level of the single cell. For this assay, cells are pre-loaded with the fluorescent marker calcein-AM. The

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dye distributes into all compartments, including the mitochondria, and is trapped after de-esterification by nonspecific esterases. The cytosolic dye is then quenched by a short incubation with Co 2+ , which enters the cytosol but cannot enter intact mitochondria, resulting in calcein-labeled mitochondria. Induction of mPTP opening by ionophores or oxidative stress results in quenching of the mitochondrial calcein signal as Co 2+ enters the matrix through the now-permeable IMM. The loss of m following opening of the pore can also be measured using potentiometric cationic dyes such as TMRM, JC-1, or rhodamine-123. The methods used to induce pore opening can provide a false positive in these assays. For example, ionophores that transport Co 2+ as well as Ca 2+ across the IMM quench the mitochondrial calcein signal without opening the mPTP (Panel, Ghaleh, & Morin, 2017). An established inducer of the mPTP, such as ischemia/reperfusion, which provides more physiologically relevant induction of the pore by combining Ca 2+ overload and oxidative stress, might be a better approach for intact cells. Therefore, the experimental design and interpretation of results are far more complex when assaying the mPTP in intact cells.

Source of mitochondria
Mitochondria can be isolated (Basic Protocol 1) from virtually any tissue, including cultured cells. When designing the assay, if developing and testing a hypothesis that probes mitochondrial function within a tissue of interest, it is important to consider the physiological context. Other challenges, such as the ability to obtain an adequate yield of organelles and to access the tissue/cells of interest, influence the choice of isolation technique. The buffers employed as well as the instrument settings for differential centrifugation may be kept constant across different experimental setups. The ideal number of cells and/or amount of tissue needed to provide a sufficient yield must be optimized prior to initiating experiments. Furthermore, some tissue, such as muscle and heart, might require an additional protease lysis step prior to homogenization, whereas other tissues, such brain or cultured cells, require gentler homogenization. The functional integrity of the isolated mitochondria can be assessed by the quality-control assays described in Support Protocols 1 to 3.

Ca 2+ -sensing fluorescent dyes
Low-affinity cell-impermeant Ca 2+ dyes are essential for correctly measuring the kinetics of mitochondrial Ca 2+ uptake and mPTP opening (Basic Protocol 2 and Alternate Protocol). Because the Ca 2+ retention capacity of mitochondria can vary across tissues, cell types, and isolation techniques, the most appropriate dye must be identified for the assay. Ideally, the K d of the dye should be in the micromolar range. Shown in Table 1 are candidate Ca 2+ -sensitive fluorescence indicators.

Appropriate positive controls
It is crucial to include appropriate controls for assessing mitochondrial permeability transition in the assay design. CsA is the gold-standard control (see Basic Protocol 2) for desensitizing the pore to inducers of pore opening, thereby delaying CypD-dependent permeability transition. The contribution of CypD to pore opening can also be probed by genetic knock-out or knock-down. However, CypD inhibition only delays, and does not abolish, pore opening. The undefined nature of the pore complex currently makes it difficult to identify drug targets independent of CypD, although some exist, such as ER-000444793 (Briston et al., 2016).

Troubleshooting
The isolation technique (Basic Protocol 1) is often the source of technical errors encountered in executing these protocols. Overly harsh isolation conditions can damage the mitochondria and inhibit mitochondrial Ca 2+ uptake. The quality-control assays described in Support Protocols 1 to 3, and particularly retention of cytochrome c in the mitochondrial membrane (Support Protocol 3), are useful for identifying the optimum conditions for isolation of the mitochondria. Often, organelle isolation from different tissues results in a crude mitochondrial pellet that can be used in the assay of interest. However, contamination of Figure 6 Concentration-dependent response to cyclosporin A (CsA) in a Ca 2+ retention capacity assay. (A) Samples of 10 µM CaCl 2 (arrows) were added every tenth cycle after the initial addition at cycle 5 (arrows). An increase in fluorescence intensity after each addition is followed by a steep decline, indicating mitochondrial Ca 2+ uptake. (B) Graphical representation of percentage inhibition as compared to untreated mitochondria. the mitochondria with other tissue components (e.g., myelin in brain preparations) can negatively affect mitochondrial function. In such cases, a Percoll density gradient can be used to obtain enriched mitochondrial fractions with improved respiratory properties (Sims & Anderson, 2008).

Understanding Results
A graphical representation of the raw data and quantification of pore opening in the presence of CsA are shown in Figures 6A and 6B, respectively. The data were analyzed using the method described in Basic Protocol 2, step 8.

Time Considerations
The experimental workflow mentioned in Figure 1 requires ࣙ6 hr of time, which includes the time required for Basic Protocols 1 and 2. Basic Protocol 2 alone can take between 45 min and 3 hr, depending on the number of conditions per plate. Support Protocols 1 and 2 do not need to be performed for every single isolation; however, these take 1 to 2 hr each when attempted.
It is critical that freshly isolated mitochondria are used for Support Protocols 1 and 2 and Basic Protocol 2, whereas mitochondria can be stored at −20°C for use in Support Protocol 3. The time required to complete Support Protocol 3 varies based on the system used for immunoblotting, ranging from 4 hr to 3 days with current technology.
The requirement to use freshly isolated mitochondria can be circumvented by isolating mitochondria according to Alternate Protocol, which allows the user to stop after protein quantification and proceed on a later date.