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

Podsumowanie

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.

Streszczenie

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.

Wprowadzenie

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 "mitochondrial uncoupling"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 production^2,3,4. 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 tissues^5,6,7.

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 MCU^{10,11,12,13,14}. 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 publication^6,7,9. 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, UCP1^6,7. 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.

Protokół

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

1. 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.
2. 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.
3. Make an incision of 2 cm in the upper back after grasping the skin with tweezers.
4. Extract the brown interscapular fat of the mouse that corresponds to a two-lobed organ with a butterfly shape¹⁶.
5. Transfer the brown fat to a 35 mm Petri dish filled with 5 mL of cold isolation buffer (Table 1) previously placed on ice.
6. Clean the brown fat from the white fat under a binocular.
7. 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).
8. Use an overhead stirrer to homogenize the pre-cut tissue on ice with six gentle strokes at a controlled speed of 275 rotations/min.
9. Centrifuge the homogenate at 8,500 x g for 10 min at 4 °C in a 15 mL ice-cold conical tube. Discard the supernatant containing the lipid phase.
10. 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.
11. Transfer the homogenate in a 15 mL ice-cold conical tube and centrifuge it at 700 x g for 10 min at 4 °C to pellet all nuclei and unbroken cells.
12. Collect the supernatant in a fresh 15 mL tube and place it on ice.
13. Centrifuge the supernatant at 8,500 x g for 10 min at 4 °C to obtain a pellet containing mitochondria.
14. 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.

2. Mitochondrial isolation from the mouse heart (modified from Garg et al. 2019)¹⁷

1. 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.
2. 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.
3. Dissect the heart from the animal'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).
4. 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).
5. Use an overhead stirrer to homogenize the pre-cut tissue on ice with six gentle strokes at a controlled speed of 275 rotations/min.
6. Transfer the homogenate to a 15 mL ice-cold conical tube and centrifuge it at 700 x g for 10 min at 4 °C to pellet nuclei and unbroken cells.
7. Collect the supernatant in a fresh 15 mL tube and place it on ice.
8. Centrifuge the supernatant at 8,500 x g for 10 min at 4 °C to obtain a pellet containing mitochondria.
9. 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.

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.

1. Fill the mitochondrial-hypertonic-mannitol suspension into a refrigerated mini pressure cell (piston diameter 3/8") of the French press (Figure 3A).
2. 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).  Ensure that the suspension comes out of the mini pressure cell at a rate of about 1 drop/s.
3. Collect the drops in a 15 mL ice-chilled conical tube.
4. Centrifuge the suspension at 10,500 x g for 10 min at 4 ˚C.
5. 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.

4. Electrophysiological recordings of H⁺ leak through UCP1 and AAC^5,7,15

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.

1. 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.
2. Insert one glass filament within the puller and pull to obtain almost two identical patch pipettes from one borosilicate filament.
3. Adjust the program when pipettes become inconsistent between pulling cycles due to aging of the heating box filament of the puller.
4. Position the pipette inside the pipette polisher and place the tip near the filament under 100x magnification to fire-polish it.
5. Press the foot pedal several times to heat the filament without clogging or damaging the tip curve.
6. Polish until pipettes with a resistance between 25 and 35 MΩ are obtained when filled with TMA-based pipette solution (TMA for tetramethylammonium hydroxide, Table 4).
7. 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.
8. Prepare a raw dilution by mixing ~35 µL of the concentrated mitoplast suspension with 500 µL of the KCl bath solution (Table 5) and place it on coverslips previously placed in a well of a 4-well plate.
9. Incubate on ice for 15 to 20 min for mitoplasts to sediment on the coverslip.
10. Fill the bath chamber completely with ~50 µL of the KCl bath solution (Table 5).
11. Transfer a coverslip with mitoplasts within the chamber using thin microdissection tweezers with a bent tip.
12. Arrange the coverslip at the bottom of the chamber. Do not perfuse the chamber to keep the mitoplasts stable on the coverslip.
13. Choose an individual non-adhesive mitoplast in the shape of 8 by scanning the coverslip under the microscope with a 60x objective.
14. Load the pipette with the pipette solution (~50 µL) and place it in the pipette holder.
15. 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.
16. Apply slight negative pressure to quickly create a gigaseal with the IMM (Figure 2B).
17. 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.
18. Compensate the stray capacitance transients with the "membrane test" 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.
19. 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.
20. After the break-in, fit the capacitance transients with the amplifier program'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 MΩ. Mitoplasts (2-6 µm in size) used for patch-clamp experiments typically have membrane capacitances of 0.5-1.1 pF.
21. Immediately after the break-in, replace the KCl bath solution (Table 5) with the HEPES bath solution (Table 6) by starting the perfusion.
22. 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 UCP1^6,7,15 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.

Wyniki

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...

Dyskusje

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 thermogenesis^5,6,7,15. 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...

Materiały

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