I thank Dr. Yuriy Kirichok for the great science I was part of in his lab and the members of the Kirichok lab for the helpful discussions. I also thank Dr. Douglas C. Wallace for providing AAC1 knockout mice. Funding: A.M.B was supported by an American Heart Association Career Development Award 19CDA34630062.

All animal experimental procedures that were performed conform to the National Institutes of Health guidelines and were approved by the University of California Los Angeles Institutional Animal Care and Use Committee (IACUC).
NOTE: The mitochondrial isolation procedure is based on differential centrifugation and varies slightly from tissue to tissue. For example, since brown adipose tissue is extremely rich in lipids, it requires an additional step to separate cell debris and organelles from the lipid phase before harvesting the mitochondria. To avoid confusion, the two mitochondrial isolation procedures (one from the brown fat and the other from the heart) are detailed below.
1. Mitochondrial isolation from mouse interscapular brown fat (modified from Bertholet et al. 2020)15
<ol>
	<li>Euthanize C57BL/6 male mouse using CO2 asphyxiation and subsequent cervical dislocation, as recommended by the American Veterinary Medical Association Panel and the IACUC Committee.</li>
	<li>After positioning the mouse with its belly facing the table, spray alcohol to clean and wet the hair (modified from Mann et al., 2014)16.</li>
	<li>Make an incision of 2 cm in the upper back after grasping the skin with tweezers.</li>
	<li>Extract the brown interscapular fat of the mouse that corresponds to a two-lobed organ with a butterfly shape16.</li>
	<li>Transfer the brown fat to a 35 mm Petri dish filled with 5 mL of cold isolation buffer (Table 1) previously placed on ice.</li>
	<li>Clean the brown fat from the white fat under a binocular.</li>
	<li>Transfer the brown fat to a 10 mL beaker with 5 mL of cold isolation buffer (Table 1) to chop it up into thin pieces. Transfer to the ice-chilled 10 mL glass homogenizer (plastic material polytetrafluoroethylene (PTFE) pestle).</li>
	<li>Use an overhead stirrer to homogenize the pre-cut tissue on ice with six gentle strokes at a controlled speed of 275 rotations/min.</li>
	<li>Centrifuge the homogenate at 8,500 x g for 10 min at 4 &#176;C in a 15 mL ice-cold conical tube. Discard the supernatant containing the lipid phase.</li>
	<li>Resuspend the pellet in 5 mL of ice-cold isolation buffer (Table 1) and homogenize, a second time, the suspension on ice with six slow strokes at a speed of 275 rotation/min.</li>
	<li>Transfer the homogenate in a 15 mL ice-cold conical tube and centrifuge it at 700 x g for 10 min at 4 &#176;C to pellet all nuclei and unbroken cells.</li>
	<li>Collect the supernatant in a fresh 15 mL tube and place it on ice.</li>
	<li>Centrifuge the supernatant at 8,500 x g for 10 min at 4 &#176;C to obtain a pellet containing mitochondria.</li>
	<li>Resuspend pellet containing mitochondria in 3.8 mL of ice-cold hypertonic-mannitol buffer (Table 2) and incubate the mitochondrial suspension on ice for 10-15 min.</li>
</ol>
2. Mitochondrial isolation from the mouse heart (modified from Garg et al. 2019)17
<ol>
	<li>Euthanize C57BL/6 male mouse using CO2 asphyxiation and subsequent cervical dislocation, as recommended by the American Veterinary Medical Association Panel and the IACUC Committee.</li>
	<li>After positioning the mouse on its back, spray alcohol to clean and wet the hair. Then, make a 2 cm incision on the thorax after grasping the skin with tweezers.</li>
	<li>Dissect the heart from the animal&#39;s chest and rinse it to remove all the blood in a 10 mL beaker with 5 mL of the cold isolation solution (Table 1).</li>
	<li>Once the heart has been cleared of traces of blood, transfer it to another 10 mL beaker containing 5 mL of cold isolation buffer (Table 1) to chop it up into thin pieces. Then, transfer to an ice-chilled 10 mL glass homogenizer (PTFE pestle).</li>
	<li>Use an overhead stirrer to homogenize the pre-cut tissue on ice with six gentle strokes at a controlled speed of 275 rotations/min.</li>
	<li>Transfer the homogenate to a 15 mL ice-cold conical tube and centrifuge it at 700 x g for 10 min at 4 &#176;C to pellet nuclei and unbroken cells.</li>
	<li>Collect the supernatant in a fresh 15 mL tube and place it on ice.</li>
	<li>Centrifuge the supernatant at 8,500 x g for 10 min at 4 &#176;C to obtain a pellet containing mitochondria.</li>
	<li>Resuspend the mitochondrial pellet in 3.8 mL of ice-cold hypertonic-mannitol buffer (Table 2) and incubate the mitochondrial suspension on ice for 10-15 min.</li>
</ol>
3. Preparation of mitoplasts with a French Press for mechanical rupture of the OMM.
NOTE: The French press procedure allows the IMM to be released from the OMM with its integrity preserved, including the matrix and crista (Figure 2)18. Mitochondria are pre-incubated in a hypertonic-mannitol buffer (Table 2) and subjected to a lower pressure during the French press procedure to avoid any drastic stretching of the IMM when the OMM is ruptured.
<ol>
	<li>Fill the mitochondrial-hypertonic-mannitol suspension into a refrigerated mini pressure cell (piston diameter 3/8&#34;) of the French press (Figure 3A).</li>
	<li>Select the Medium mode of the French Press and compress the suspension through the mini pressure cell at 110 on the dial of the French Press for the brown fat mitochondria and at 140 for the heart mitochondria (~2,000 psi).&#160; Ensure that the suspension comes out of the mini pressure cell at a rate of about 1 drop/s.</li>
	<li>Collect the drops in a 15 mL ice-chilled conical tube.</li>
	<li>Centrifuge the suspension at 10,500 x g for 10 min at 4 &#730;C.</li>
	<li>Resuspend the mitoplasts pellet in 0.5-2 mL of ice-cold Hypertonic-KCl buffer (Table 3) and store the suspension on ice. 
	NOTE: Brown fat and heart mitoplasts are ready for patch-clamp recordings and should remain usable for about 3-6 h.</li>
</ol>
4. Electrophysiological recordings of H+ leak through UCP1 and AAC5,7,15
NOTE: Use the following electrophysiological setup (Figure 3B): inverted microscope with differential interference contrast (DIC), 60x water immersion objective, vibration isolation table and a Faraday cage, a standard amplifier supporting low-noise recordings, a standard digitizer used for electrophysiological setup, pClamp 10, a micromanipulator, bath reference electrode (3 M KCl-agar salt bridge inserted within a microelectrode holder containing a silver/silver chloride pellet molded into the holder body (described in Liu et al. 2021)19, perfusion chamber with a 0.13 mm glass coverslip bottom, connected to a gravity-fed perfusion system.
<ol>
	<li>Pull the borosilicate glass filaments on the day of recording using a micropipette puller. Set a program on the puller used for generating pipettes with a high degree of reproducibility20. 
	NOTE: This program design requires several tries to obtain pipettes optimized for the IMM patch-clamp. A standard pipette has fine tips with a progressive conic shape.</li>
	<li>Insert one glass filament within the puller and pull to obtain almost two identical patch pipettes from one borosilicate filament.</li>
	<li>Adjust the program when pipettes become inconsistent between pulling cycles due to aging of the heating box filament of the puller.</li>
	<li>Position the pipette inside the pipette polisher and place the tip near the filament under 100x magnification to fire-polish it.</li>
	<li>Press the foot pedal several times to heat the filament without clogging or damaging the tip curve.</li>
	<li>Polish until pipettes with a resistance between 25 and 35 M&#937; are obtained when filled with TMA-based pipette solution (TMA for tetramethylammonium hydroxide, Table 4).</li>
	<li>Pre-incubate coverslips (5 mm diameter, 0.1 mm thickness) with 0.1% gelatin to reduce the mitoplast adhesion and rinse them with the KCl bath solution (Table 5) before depositing the mitoplast suspension.</li>
	<li>Prepare a raw dilution by mixing ~35 &#181;L of the concentrated mitoplast suspension with 500 &#181;L of the KCl bath solution (Table 5) and place it on coverslips previously placed in a well of a 4-well plate.</li>
	<li>Incubate on ice for 15 to 20 min for mitoplasts to sediment on the coverslip.</li>
	<li>Fill the bath chamber completely with ~50 &#181;L of the KCl bath solution (Table 5).</li>
	<li>Transfer a coverslip with mitoplasts within the chamber using thin microdissection tweezers with a bent tip.</li>
	<li>Arrange the coverslip at the bottom of the chamber. Do not perfuse the chamber to keep the mitoplasts stable on the coverslip.</li>
	<li>Choose an individual non-adhesive mitoplast in the shape of 8 by scanning the coverslip under the microscope with a 60x objective.</li>
	<li>Load the pipette with the pipette solution (~50 &#181;L) and place it in the pipette holder.</li>
	<li>Bring the pipette into the bath solution with a micromanipulator and approach it just above the selected mitoplast to get close to the IMM. The amplifier program gives the resistance of the pipette once it is in the bath solution. Hold the membrane potential at 0 mV and apply 10 mV pulses using the membrane test command in the amplifier program.</li>
	<li>Apply slight negative pressure to quickly create a gigaseal with the IMM (Figure 2B).</li>
	<li>Rise the pipette with the mitoplast attached to keep them away from the coverslip to avoid the seal breakage due to the pipette drift during the experiment.</li>
	<li>Compensate the stray capacitance transients with the &#34;membrane test&#34; command in the amplifier program before testing the whole-mitoplast configuration to obtain a correct capacitance (Cm) measurement for the mitoplast membrane after the break-in.</li>
	<li>Apply short-duration (5-15 ms) voltage pulses (250-600 mV) with the amplifier program to rupture the membrane patch under the glass pipette and achieve the whole-mitoplast configuration (Figure 2C). Successful break-in is reflected by the reappearance of capacitance transients.</li>
	<li>After the break-in, fit the capacitance transients with the amplifier program&#39;s membrane test option to assess the membrane capacitance (reflecting the size of the mitoplast) and its access resistance Ra (reflecting the quality of the whole-mitoplast configuration). After the break-in, Ra should be between 40 and 80 M&#937;. Mitoplasts (2-6 &#181;m in size) used for patch-clamp experiments typically have membrane capacitances of 0.5-1.1 pF.</li>
	<li>Immediately after the break-in, replace the KCl bath solution (Table 5) with the HEPES bath solution (Table 6) by starting the perfusion.</li>
	<li>Apply an 850 ms ramp protocol designed with the amplifier program, from -160 mV to +100 mV with a 5 s interval, while holding the mitoplast at 0 mV. This protocol works for UCP16,7,15 and AAC5 studies (Figure 4 and Figure 5). 
	NOTE: It is recommended for UCP1 and AAC-dependent H+ current measurements to acquire all electrophysiological data at 10 kHz and to filter at 1 kHz using adequate software driving the amplifier and digitizer.</li>
</ol>

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The author declares no competing interests.

This method article aims to present the patch-clamp technique recently applied to mitochondria, a new approach to directly study H+ leak through the IMM responsible for mitochondrial thermogenesis5,6,7,15. This technique is not limited to tissues and can also be used to analyze H+ leak and other conductances of the IMM in different standard human and cell models such as HA...

Mitochondria are famous for being the powerhouse of the cell. Indeed, they are the major source of chemical energy, ATP. What is less known is that mitochondria also generate heat. In fact, every mitochondrion constantly generates the two types of energies (ATP and heat) and a fine balance between the two energy forms defines metabolic cell homeostasis (Figure 1). How mitochondria distribute energy between ATP and heat is certainly the most fundamental question in the field of bioenergetics, although it is still largely unknown. We do know that increasing mitochondrial heat production (called mitochondrial thermogenesis), and consequently reducing ATP production increases energy expenditure and this is one of the best ways to combat metabolic syndrome1.
Mitochondrial thermogenesis originates from H+ leak across the inner mitochondrial membrane (IMM), leading to uncoupling of substrate oxidation and ATP synthesis with consequent production of heat, hence the name &#34;mitochondrial uncoupling&#34;1 (Figure 1). This H+ leak depends on mitochondrial transporters called uncoupling proteins (UCPs). UCP1 was the first UCP identified. It is only expressed in thermogenic tissues, brown fat, and beige fat in which mitochondria are specialized for heat production2,3,4. The identity of UCP in non-adipose tissues such as skeletal muscle, heart, and liver, has remained controversial. Mitochondria in these tissues can have about 25% of the total mitochondrial energy converted into heat, which can significantly impact the physiology of the whole body1. Besides maintaining core body temperature, mitochondrial thermogenesis also prevents diet-induced obesity by reducing calories. In addition, it reduces the production of reactive oxygen species (ROS) by mitochondria to protect cells from oxidative damage1. Thus, mitochondrial thermogenesis is involved in normal aging, age-related degenerative disorders, and other conditions involving oxidative stress, such as ischemia-reperfusion. Therefore, mitochondrial thermogenesis is a powerful regulator of cellular metabolism, and a mechanistic understanding of this fundamental process will promote the development of therapeutic strategies to combat many pathologies associated with mitochondrial dysfunction.
Mitochondrial respiration was the first technique to reveal the crucial role of mitochondrial thermogenesis in cellular metabolism and is still the most popular in the community1. This technique is based on the measurement of oxygen consumption by the mitochondrial electron transport chain (ETC) that increases when mitochondrial H+ leak is activated. This technique, although instrumental, cannot directly study mitochondrial H+ leak across the IMM1, thereby making the precise identification and characterization of the proteins responsible for it difficult, particularly in non-adipose tissues in which heat production is secondary as compared to ATP production. Recently, the development of the patch-clamp technique applied to mitochondria, provided the first direct study of H+ leak across the whole IMM in various tissues5,6,7.
The mitochondrial patch-clamp of the whole IMM was first established in a reproducible way by Kirichok et al.8. They described the first direct measurement of mitochondrial calcium uniporter (MCU) currents in 2004 using mitoplasts from COS-7 cell lines8. Later, the Kirichok lab showed calcium currents from IMMs of mouse9 and Drosophila tissues9. Other labs now routinely use this technique to study the biophysical properties of MCU10,11,12,13,14. Whole IMM patch-clamp analysis of potassium and chloride conductance is also possible and has been mentioned in several papers but has not yet been the main subject of a publication6,7,9. The first measurement of H+ currents across the IMM was reported in 2012 from mouse brown fat mitochondria6, and from mouse beige fat mitochondria in 20177. This current is due to the specific uncoupling protein of thermogenic tissues, UCP16,7. Recent work published in 2019 characterized AAC as the main protein responsible for mitochondrial H+ leak in non-adipose tissues such as the heart and skeletal muscle5.
This unique approach now allows for the direct high-resolution functional analysis of the mitochondrial ion channels and transporters responsible for mitochondrial thermogenesis. To facilitate the expansion of the method and to complement other studies such as mitochondrial respiration, a detailed protocol is described below for measuring the H+ currents carried by UCP1 and AAC. Three important steps are described: 1) mitochondrial isolation from mouse brown fat to analyze UCP1-dependent H+ current and mitochondrial isolation from the heart to analyze AAC-dependent H+ current, 2) preparation of mitoplasts with a French Press for mechanical rupture of the outer mitochondrial membrane (OMM), 3) patch-clamp recordings of UCP1 and AAC-dependent H+ currents across the whole IMM.

<table><tbody><tr><td>0.1% gelatin</td><td>Millipore</td><td>ES-006-B</td><td></td></tr><tr><td>60X water immersion objective, numerical aperture 1.20</td><td>Olympus</td><td>UPLSAPO60XW</td><td></td></tr><tr><td>Axopatch 200B amplifier</td><td>Molecular Devices</td><td></td><td></td></tr><tr><td>Borosilicate glass capillaries</td><td>Sutter Instruments</td><td>BF150-86-10</td><td></td></tr><tr><td>Digidata 1550B Digitizer</td><td>Molecular Devices</td><td></td><td></td></tr><tr><td>Faraday cage</td><td>Homemade</td><td></td><td></td></tr><tr><td>French Press</td><td>Glen Mills</td><td>5500-000011</td><td></td></tr><tr><td>IKA Eurostar PWR CV S1 laboratory overhead stirrer</td><td></td><td></td><td></td></tr><tr><td>Inversed Microscope</td><td>Olympus</td><td>IX71 or IX73</td><td></td></tr><tr><td>Micro Forge</td><td>(Narishige)</td><td>MF-830</td><td></td></tr><tr><td>Micromanupulator MPC-385</td><td>Sutter Instruments</td><td>FG-MPC325</td><td></td></tr><tr><td>Microelectrode holder for agar bridge</td><td>World Precision Instruments</td><td>MEH3F4515</td><td></td></tr><tr><td>Micropipette Puller</td><td>(Sutter Instruments)</td><td>P97</td><td></td></tr><tr><td>Mini Cell for French Press</td><td>Glen Mills</td><td>5500-FA-004</td><td></td></tr><tr><td>MIXER IKA 6-2000RPM</td><td>Cole Parmer</td><td>EW-50705-50</td><td></td></tr><tr><td>Objective 100X magnification</td><td>Nikon&amp;nbsp; lens</td><td>MPlan 100/0.80 ELWD 210/0</td><td></td></tr><tr><td>pClamp 10</td><td>Molecular Devices</td><td></td><td></td></tr><tr><td>Perfusion chamber</td><td>Warner Instruments</td><td>RC-24E</td><td></td></tr><tr><td>Potter-Elvehjem homogenizer 10 ml</td><td>Wheaton</td><td>358039</td><td></td></tr><tr><td>Refrigerated centrifuge SORVALL X4R PRO-MD</td><td>Thermo Scientific</td><td>75 009 521</td><td></td></tr><tr><td>Small round glass coverslips: 5 mm diameter, 0.1 mm thickness</td><td>Warner Instruments</td><td>640700</td><td></td></tr><tr><td>Vibration isolation table</td><td>Newport</td><td>VIS3036-SG2-325A</td><td></td></tr><tr><td>&lt;strong&gt;Chemicals&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>D-gluconic acid</td><td>Sigma Aldrich</td><td>G1951</td><td></td></tr><tr><td>D-mannitol</td><td>&amp;nbsp;Sigma Aldrich</td><td>M4125</td><td></td></tr><tr><td>EGTA</td><td>&amp;nbsp;Sigma Aldrich</td><td>3777</td><td></td></tr><tr><td>HEPES</td><td>&amp;nbsp;Sigma Aldrich</td><td>H7523</td><td></td></tr><tr><td>KCl</td><td>&amp;nbsp;Sigma Aldrich</td><td>60128</td><td></td></tr><tr><td>MgCl2</td><td>&amp;nbsp;Sigma Aldrich</td><td>63068</td><td></td></tr><tr><td>sucrose</td><td>&amp;nbsp;Sigma Aldrich</td><td>S7903</td><td></td></tr><tr><td>TMA</td><td>&amp;nbsp;Sigma Aldrich</td><td>331635</td><td></td></tr><tr><td>TrisBase</td><td>&amp;nbsp;Sigma Aldrich</td><td>T1503</td><td></td></tr><tr><td>TrisCl</td><td>&amp;nbsp;Sigma Aldrich</td><td>T3253</td><td></td></tr></tbody></table>

the use of the patch-clamp technique to study the thermogenic capacity of mitochondria

Mitochondrial thermogenesis (also known as mitochondrial uncoupling) is one of the most promising targets for increasing energy expenditure to combat metabolic syndrome. Thermogenic tissues such as brown and beige fats develop highly specialized mitochondria for heat production. Mitochondria of other tissues, which primarily produce ATP, also convert up to 25% of the total mitochondrial energy production into heat and can, therefore, have a considerable impact on the physiology of the whole body. Mitochondrial thermogenesis is not only essential for maintaining the body temperature, but also prevents diet-induced obesity and reduces the production of reactive oxygen species (ROS) to protect cells from oxidative damage. Since mitochondrial thermogenesis is a key regulator of cellular metabolism, a mechanistic understanding of this fundamental process will help in the development of therapeutic strategies to combat many pathologies associated with mitochondrial dysfunction. Importantly, the precise molecular mechanisms that control acute activation of thermogenesis in mitochondria are poorly defined. This lack of information is largely due to a dearth of methods for the direct measurement of uncoupling proteins. The recent development of patch-clamp methodology applied to mitochondria enabled, for the first time, the direct study of the phenomenon at the origin of mitochondrial thermogenesis, H+ leak through the IMM, and the first biophysical characterization of mitochondrial transporters responsible for it, the uncoupling protein 1 (UCP1), specific of brown and beige fats, and the ADP/ATP transporter (AAC) for all other tissues. This unique approach will provide new insights into the mechanisms that control H+ leak and mitochondrial thermogenesis and how they can be targeted to combat metabolic syndrome. This paper describes the patch-clamp methodology applied to mitochondria to study their thermogenic capacity by directly measuring H+ currents through the IMM.

The development of the patch-clamp methodology applied to mitochondria provided the first direct study of H+ leak through the IMM and the mitochondrial transporters, UCP1 and AAC, which are responsible for it. The electrophysiological analysis of UCP1- and AAC-dependent H+ leaks can provide a first glance of the thermogenic capacity of mitochondria. The results section describes the standard procedures to measure H+ leak via UCP1 and AAC.
UCP1-dependent...

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The Use of the Patch-Clamp Technique to Study the Thermogenic Capacity of Mitochondria

This method article details the main steps in measuring H+ leak across the inner mitochondrial membrane with the patch-clamp technique, a new approach to study the thermogenic capacity of mitochondria.

Mitochondrial thermogenesis (also known as mitochondrial uncoupling) is one of the most promising targets for increasing energy expenditure to combat ...

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3.7K Views.  University of California Los Angeles. This method article details the main steps in measuring H+ leak across the inner mitochondrial membrane with the patch-clamp technique, a new approach to study the thermogenic capacity of mitochondria.

Video: The Use of the Patch-Clamp Technique to Study the Thermogenic Capacity of Mitochondria

Analyses of Mitochondrial Calcium Influx in Isolated Mitochondria and Cultured Cells

Ca2+ handling by mitochondria is a critical function regulating both physiological and pathophysiological processes in a broad spectrum of cells. The ability to accurately measure the influx and efflux of Ca2+ from mitochondria is important for determining the role of mitochondrial Ca2+ handling in these processes. In this report, we present two methods for the measurement of mitochondrial Ca2+ handling in both isolated mitochondria and cultured cells. We first detail a plate reader-based platform for measuring mitochondrial Ca2+ uptake using the Ca2+ sensitive dye calcium green-5N. The plate reader-based format circumvents the need for specialized equipment, and the calcium green-5N dye is ideally suited for measuring Ca2+ from isolated tissue mitochondria. For our application, we describe the measurement of mitochondrial Ca2+ uptake in mitochondria isolated from mouse heart tissue; however, this procedure can be applied to measure mitochondrial Ca2+ uptake in mitochondria isolated from other tissues such as liver, skeletal muscle, and brain. Secondly, we describe a confocal microscopy-based assay for measurement of mitochondrial Ca2+ in permeabilized cells using the Ca2+ sensitive dye Rhod-2/AM and imaging using 2-dimensional laser-scanning microscopy. This permeabilization protocol eliminates cytosolic dye contamination, allowing for specific recording of changes in mitochondrial Ca2+. Moreover, laser-scanning microscopy allows for high frame rates to capture rapid changes in mitochondrial Ca2+ in response to various drugs or reagents applied in the external solution. This protocol can be applied to measure mitochondrial Ca2+ uptake in many cell types including primary cells such as cardiac myocytes and neurons, and immortalized cell lines.

Here, we present two protocols for the measurement of mitochondrial Ca2+ influx in isolated mitochondria and cultured cells. For isolated mitochondria, we detail a plate reader-based Ca2+ import assay using the Ca2+ sensitive dye calcium green-5N. For cultured cells, we describe a confocal microscopy method using the Ca2+ dye Rhod-2/AM.

Ca2+ handling by mitochondria is a critical function regulating both physiological and pathophysiological processes in a broad spectrum of cells. The ...

Biochemistry

F1FO ATPase Vesicle Preparation and Technique for Performing Patch Clamp Recordings of Submitochondrial Vesicle Membranes

Mitochondria are involved in many important cellular functions including metabolism, survival1, development and, calcium signaling2. Two of the most important mitochondrial functions are related to the efficient production of ATP, the energy currency of the cell, by oxidative phosphorylation, and the mediation of signals for programmed cell death3.
The enzyme primarily responsible for the production of ATP is the F1FO-ATP synthase, also called ATP synthase4-5. In recent years, the role of mitochondria in apoptotic and necrotic cell death has received considerable attention. In apoptotic cell death, BCL-2 family proteins such as Bax enter the mitochondrial outer membrane, oligomerize and permeabilize the outer membrane, releasing pro-apoptotic factors into the cytosol6. In classic necrotic cell death, such as that produced by ischemia or excitotoxicity in neurons, a large, poorly regulated increase in matrix calcium contributes to the opening of an inner membrane pore, the mitochondrial permeability transition pore or mPTP. This depolarizes the inner membrane and causes osmotic shifts, contributing to outer membrane rupture, release of pro-apoptotic factors, and metabolic dysfunction. Many proteins including Bcl-xL7 interact with F1FO ATP synthase, modulating its function. Bcl-xL interacts directly with the beta subunit of F1FO ATP synthase, and this interaction decreases a leak conductance within the F1FOATPasecomplex, increasing the net transport of H+ by F1FO during F1FO ATPase activity8 and thereby increasing mitochondrial efficiency. To study the activity and modulation of the ATP synthase, we isolated from rodent brain submitochondrial vesicles (SMVs) containing F1FO ATPase. The SMVs retain the structural and functional integrity of the F1FO ATPase as shown in Alavian et al. Here, we describe a method that we have used successfully for the isolation of SMVs from rat brain and we delineate the patch clamp technique to analyze channel activity (ion leak conductance) of the SMVs.

F1FO ATPase Vesicle Preparation and Technique for Performing Patch Clamp Recordings of Submitochondrial Vesicle Membranes

A method to isolate submitochondrial vesicles enriched in F1FO ATP synthase complexes from rat brain is described. These vesicles allow the study of the activity of F1FO ATPase complex and its modulation using the technique of patch clamp recording.

Mitochondria are involved in many important cellular  functions including metabolism, survival1, development and, calcium signaling2. Two of the most ...

Neuroscience

Mitochondrial Functional Assessment in Brush Lizard Liver Mitochondria

Assessment of Mitochondrial Oxygen Consumption Using a Plate Reader-based Fluorescent Assay

Mitochondria serve many important functions, including cellular respiration, ATP production, controlling apoptosis, and acting as a central hub of metabolic pathways. Therefore, experimentally assessing mitochondrial functionality can provide insight into variations among different populations or disease states. Additionally, it is valuable to assess whether isolated mitochondria are healthy enough to proceed with experiments. One characteristic often used to compare mitochondrial function in different samples is the rate of oxygen consumption. Oxygen consumption and subsequent calculation of the respiratory control ratio in either intact cells or mitochondria isolated from tissue can serve all three purposes. Using mitochondria isolated from the livers of brush lizards in conjunction with a phosphorescent probe that is sensitive to the fluctuations in oxygen concentration of a solution, we measured oxygen consumption using a fluorescent plate reader. This method is not only quick and efficient but also can be conducted with a small amount of mitochondria and without the need for specialized equipment. The step-by-step protocol described here increases the accessibility of mitochondrial functional assessment to researchers.

Assessment of oxygen consumption provides integral information about mitochondrial function. Using a phosphorescent probe with a fluorescent plate reader, accurate and reproducible data can be obtained easily without specialized equipment. This assay enables any lab to measure the oxygen consumption of isolated mitochondria and calculate respiratory control ratios.

Mitochondria serve many important functions, including cellular respiration, ATP production, controlling apoptosis, and acting as a central hub of ...

Multi-parameter Measurement of the Permeability Transition Pore Opening in Isolated Mouse Heart Mitochondria

The mitochondrial permeability transition pore (mtPTP) is a non specific channel that forms in the inner mitochondrial membrane to transport solutes with a molecular mass smaller than 1.5 kDa. Although the definitive molecular identity of the pore is still under debate, proteins such as cyclophilin D, VDAC and ANT contribute to mtPTP formation. While the involvement of mtPTP opening in cell death is well established1, accumulating evidence indicates that the mtPTP serves a physiologic role during mitochondrial Ca2+ homeostasis2, bioenergetics and redox signaling 3.
mtPTP opening is triggered by matrix Ca2+ but its activity can be modulated by several other factors such as oxidative stress, adenine nucleotide depletion, high concentrations of Pi, mitochondrial membrane depolarization or uncoupling, and long chain fatty acids4. In vitro, mtPTP opening can be achieved by increasing Ca2+ concentration inside the mitochondrial matrix through exogenous additions of Ca2+ (calcium retention capacity). When Ca2+ levels inside mitochondria reach a certain threshold, the mtPTP opens and facilitates Ca2+ release, dissipation of the proton motive force, membrane potential collapse and an increase in mitochondrial matrix volume (swelling) that ultimately leads to the rupture of the outer mitochondrial membrane and irreversible loss of organelle function.
Here we describe a fluorometric assay that allows for a comprehensive characterization of mtPTP opening in isolated mouse heart mitochondria. The assay involves the simultaneous measurement of 3 mitochondrial parameters that are altered when mtPTP opening occurs: mitochondrial Ca2+ handling (uptake and release, as measured by Ca2+ concentration in the assay medium), mitochondrial membrane potential, and mitochondrial volume. The dyes employed for Ca2+ measurement in the assay medium and mitochondrial membrane potential are Fura FF, a membrane impermeant, ratiometric indicator which undergoes a shift in the excitation wavelength in the presence of Ca2+, and JC-1, a cationic, ratiometric indicator which forms green monomers or red aggregates at low and high membrane potential, respectively. Changes in mitochondrial volume are measured by recording light scattering by the mitochondrial suspension. Since high-quality, functional mitochondria are required for the mtPTP opening assay, we also describe the steps necessary to obtain intact, highly coupled and functional isolated heart mitochondria.

A spectrofluorometric protocol for the measurement of the mitochondrial permeability transition pore opening in isolated mouse heart mitochondria is presented here. The assay involves the simultaneous measurement of mitochondria Ca2+ handling, mitochondrial membrane potential and mitochondrial volume. The procedure for obtaining high-quality and functional heart mitochondria is also described.

The mitochondrial permeability transition pore (mtPTP) is a non specific channel that forms in the inner mitochondrial membrane to transport solutes with ...

One-channel Cell-attached Patch-clamp Recording

Ion channel proteins are universal devices for fast communication across biological membranes. The temporal signature of the ionic flux they generate depends on properties intrinsic to each channel protein as well as the mechanism by which it is generated and controlled and represents an important area of current research. Information about the operational dynamics of ion channel proteins can be obtained by observing long stretches of current produced by a single molecule. Described here is a protocol for obtaining one-channel cell-attached patch-clamp current recordings for a ligand gated ion channel, the NMDA receptor, expressed heterologously in HEK293 cells or natively in cortical neurons. Also provided are instructions on how to adapt the method to other ion channels of interest by presenting the example of the mechano-sensitive channel PIEZO1. This method can provide data regarding the channel&#8217;s conductance properties and the temporal sequence of open-closed conformations that make up the channel&#8217;s activation mechanism, thus helping to understand their functions in health and disease.

Described here is a procedure for obtaining long stretches of current recording from one ion channel with the cell-attached patch-clamp technique. This method allows for observing, in real time, the pattern of open-close channel conformations that underlie the biological signal. These data inform about channel properties in undisturbed biological membranes.

Ion channel proteins are universal devices for fast communication across biological membranes. The temporal signature of the ionic flux they generate ...

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

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.

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.

Mitochondrial energetics is a central theme in animal biochemistry and physiology, with researchers using mitochondrial respiration as a metric to ...

Assessment of Open Probability of the Mitochondrial Permeability Transition Pore in the Setting of Coenzyme Q Excess

The mitochondrial permeability transition pore (mPTP) is a voltage-gated, nonselective, inner mitochondrial membrane (IMM) mega-channel important in health and disease. The mPTP mediates leakage of protons across the IMM during low-conductance opening and is specifically inhibited by cyclosporine A (CsA). Coenzyme Q (CoQ) is a regulator of the mPTP, and tissue-specific differences have been found in CoQ content and open probability of the mPTP in forebrain and heart mitochondria in a newborn mouse model of fragile X syndrome (FXS, Fmr1 knockout). We developed a technique to determine the voltage threshold for mPTP opening in this mutant strain, exploiting the role of the mPTP as a proton leak channel.
To do so, oxygen consumption and membrane potential (&#916;&#936;) were simultaneously measured in isolated mitochondria using polarography and a tetraphenylphosphonium (TPP+) ion-selective electrode during leak respiration. The threshold for mPTP opening was determined by the onset of CsA-mediated inhibition of proton leak at specific membrane potentials. Using this approach, differences in voltage gating of the mPTP were precisely defined in the context of CoQ excess. This novel technique will permit future investigation for enhancing the understanding of physiological and pathological regulation of low-conductance opening of the mPTP.

This method exploits the contribution of the mitochondrial permeability transition pore to low-conductance proton leak to determine the voltage threshold for pore opening in neonatal fragile X syndrome mice with increased cardiomyocyte mitochondrial coenzyme Q content compared to wildtype control.

The mitochondrial permeability transition pore (mPTP) is a voltage-gated, nonselective, inner mitochondrial membrane (IMM) mega-channel important in ...

Patch-Clamp Techniques for Single Endolysosomal Vesicle Analysis

Endolysosomal ion channels are critical for endolysosomal ion and pH homeostasis, membrane potential regulation, and vesicle trafficking. However, electrophysiologically accessing these channels within small intracellular vesicles has been a challenge. The development of endolysosomal patch-clamp techniques has been instrumental in overcoming this barrier, allowing for the direct measurement of ion channel activity in endolysosomal membranes.
Compared to existing planar patch-clamp techniques, endolysosomal patch-clamp can simultaneously record multiple cells and easily combine with other measurement methods. Manual operation offers the advantage of visualizing targeted vesicles. It also addresses the limitation of the indispensable presence of Ca2+ on one side of the endolysosomal membrane, increasing the flexibility of experimental design. Utilizing endolysosomal patch-clamp techniques enables the direct measurement and analysis of ion channel activity within endolysosomes.
Given the close link between aberrant endolysosomal ion channel function and diseases such as neurodegenerative diseases and metabolic disorders, investigating and modulating these channels may unveil new drug targets. By restoring intracellular ion balance, we may alleviate or cure related diseases. Therefore, this technique is pivotal for discovering new drug targets and developing relevant medications.

This protocol details a method for directly measuring ion channel activity on intracellular vesicles using a manual endolysosomal patch-clamp system. We illustrate the method that involves enlarging endolysosomes and manually isolating these vesicles. This approach ensures that researchers can accurately replicate and apply the procedure.

Endolysosomal ion channels are critical for endolysosomal ion and pH homeostasis, membrane potential regulation, and vesicle trafficking. However, ...

Patch Clamp Electrophysiology

Neuron cell membranes are populated with ion channels that control the movement of charge into and out of the cell, thereby regulating neuron firing. One extremely useful technique for investigating the biophysical properties of these channels is called patch clamp recording. In this method, neuroscientists place a polished glass micropipette against a cell and apply suction to form a high resistance seal. This process isolates a small &#8220;patch&#8221; of membrane that contains one or more ion channels. Using an electrode within the micropipette, researchers can &#8220;clamp&#8221; or control the electrical properties of the membrane, which is important for analysis of channel activity. The electrode also allows for changes in the voltage across the membrane, or the flow of ions through the membrane, to be recorded. This video begins with an overview of the principles behind patch clamp electrophysiology, an introduction to the necessary equipment, and descriptions of the various patch configurations, including whole cell, cell-attached, perforated, inside-out, and outside-out patches. Next, the key steps of a typical whole-cell patch clamp experiment are outlined, in which a current-voltage (IV) curve is generated. Finally, applications of patch clamp recording are provided to demonstrate how the biophysical properties of ion channels, cell excitability, and neuroactive compounds are evaluated in neurophysiology labs today.

Patch Clamp Electrophysiology: Principle &amp; Applications

Neuron cell membranes are populated with ion channels that control the movement of charge into and out of the cell, thereby regulating neuron firing. One ...

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Mitochondrial Calcium Uptake Assay: A Plate Reader-Based Method for Measuring Calcium Uptake in Isolated Mitochondria

Source: Maxwell, J.T., et al., <a href="https://app.jove.com/v/57225">Analyses of Mitochondrial Calcium Influx in Isolated Mitochondria and Cultured Cells.</a> J. Vis. Exp. (2018).
This video describes a plate reader-based assay to measure mitochondrial calcium uptake in isolated mitochondria. The assay analyzes the fluorescence signal of a calcium-sensitive fluorescent dye to study the kinetics of mitochondrial calcium uptake and calcium overload.

Source: Maxwell, J.T., et al., Analyses of Mitochondrial Calcium Influx in Isolated Mitochondria and Cultured Cells. J. Vis. Exp. (2018).
This video ...