Assessing Mitochondrial Function in Sciatic Nerve by High-Resolution Respirometry

Marcos A. Formiga-Jr; Juliana Camacho-Pereira

doi:10.3791/63690

JoVE Journal > Biochemistry

생화학

Assessing Mitochondrial Function in Sciatic Nerve by High-Resolution Respirometry

Published: May 05, 2022

doi:

10.3791/63690

Marcos A. Formiga-Jr¹, Juliana Camacho-Pereira¹

¹Instituto de Bioquímica Médica Leopoldo de Meis, Centro de Ciências da Saúde,Universidade Federal do Rio de Janeiro

Summary

High-resolution respirometry coupled to fluorescence sensors determines mitochondrial oxygen consumption and reactive oxygen species (ROS) generation. The present protocol describes a technique to assess mitochondrial respiratory rates and ROS production in the permeabilized sciatic nerve.

Abstract

Mitochondrial dysfunction in peripheral nerves accompanies several diseases associated with peripheral neuropathy, which can be triggered by multiple causes, including autoimmune diseases, diabetes, infections, inherited disorders, and tumors. Assessing mitochondrial function in mouse peripheral nerves can be challenging due to the small sample size, a limited number of mitochondria present in the tissue, and the presence of a myelin sheath. The technique described in this work minimizes these challenges by using a unique permeabilization protocol adapted from one used for muscle fibers, to assess sciatic nerve mitochondrial function instead of isolating the mitochondria from the tissue. By measuring fluorimetric reactive species production with Amplex Red/Peroxidase and comparing different mitochondrial substrates and inhibitors in saponin-permeabilized nerves, it was possible to detect mitochondrial respiratory states, reactive oxygen species (ROS), and the activity of mitochondrial complexes simultaneously. Therefore, the method presented here offers advantages compared to the assessment of mitochondrial function by other techniques.

Introduction

Mitochondria are essential for maintaining cell viability and perform numerous cell functions such as energy metabolism (glucose, amino acid, lipid, and nucleotide metabolism pathways). As the primary site of reactive oxygen species (ROS) production, mitochondria are central in several cell signaling processes such as apoptosis and participate in the synthesis of iron-sulfur (Fe-S) clusters, mitochondrial protein import and maturation, and maintenance of their genome and ribosomes¹^,²^,³. The mitochondrial membrane dynamics network is controlled by fusion and fission processes, and they also have machinery for quality control and mitophagy⁴^,⁵^,⁶.

Mitochondrial dysfunction is associated with the appearance of several pathological conditions such as cancer, diabetes, and obesity⁷. Disturbances in mitochondrial function are detected in neurodegenerative disorders that affect the central nervous system, as in Alzheimer's disease⁸^,⁹, Parkinson's disease¹⁰^,¹¹, amyotrophic lateral sclerosis¹²^,¹³, and Huntington's disease¹⁴^,¹⁵. In the peripheral nervous system, loss of mitochondrial function in axons is observed in immune neuropathies, such as Guillain-Barré syndrome¹⁶^,¹⁷, and in association with high mitochondrial ROS production in axons, these events lead to MAP Kinase activation in Schwann cells¹⁸. This demonstrates that mitochondrial physiology may be essential not only for a site-specific cell, but for an entire tissue. In HIV-associated distal sensory polyneuropathy (HIV-DSP), mitochondria have a role in the mechanism by which the trans-activator of transcription (HIV-TAT) protein allows HIV to replicate efficiently, as well as several other roles in HIV infection pathogenesis¹⁹^,²⁰.

Evaluation of sciatic nerve mitochondrial physiology has emerged as an essential target for investigating neuropathy⁷^,²¹^,²². In diabetic neuropathy, proteomic and metabolomic analyses suggest that most molecular alterations in diabetes affect sciatic nerve mitochondrial oxidative phosphorylation and lipid metabolism⁷. These alterations also seem to be early signs of obesity-induced diabetes²¹. In a mouse model of chemotherapy-induced painful neuropathy, mitochondrial impairment in the sciatic nerve is detected as a decrease in oxidative phosphorylation²², and a reduction of mitochondrial complexes activities, membrane potential, and ATP content²³. However, although several groups have cited mitochondrial dysfunction in neuropathies, these studies are limited to the measurements of activity in mitochondrial complexes with no preservation of the mitochondrial membranes, lacking evaluation of mitochondrial integrity or measurements of ATP content as a parameter for mitochondrial ATP production. In general, a proper assessment of mitochondrial oxygen consumption and ROS production requires the isolation of mitochondria by differential centrifugation in a percoll/sucrose gradient. Isolation of mitochondria can also be a limiting factor for sciatic nerve tissue because of the large amount of tissue needed and mitochondria loss and disruption.

The present study aims to provide a protocol to measure mitochondrial physiology as mitochondrial oxygen consumption and ROS production in the sciatic nerve, preserving mitochondrial membranes and without the need for isolating mitochondria. This protocol is adapted from oxygen consumption measurements in permeabilized muscle fibers²⁴ by high-resolution respirometry (HRR). The advantages of this procedure are the possibility of evaluating mitochondria in small amounts of tissue such as the sciatic nerve and evaluating mitochondrial parameters in situ, thereby preserving the mitochondrial environment, structure, and bioenergetic profile, to obtain a physiologically trustworthy result. The mitochondrial respiratory states were determined with substrates and inhibitors after sciatic nerve permeabilization to properly assess mitochondrial bioenergetics and cytochrome c coefficient for mitochondrial membrane integrity, providing a guide for steps of the mitochondrial electron transport system (ETS) evaluation and calculation of essential parameters. This study can provide tools for answering questions in pathophysiological mechanisms in which sciatic nerve metabolism is implicated, such as peripheral neuropathies.

Protocol

The present protocol is approved by the Ethics Committee on the Use of Animals in Research, CCS/UFRJ (CEUA-101/19), and National Institutes of Health guidelines for the care and use of experimental animals. The sciatic nerve is isolated from four-month-old male C57BL/6 mice, euthanized by cervical dislocation as per the institutional guidelines. The protocol steps are optimized to avoid mitochondrial deterioration. Therefore, in this protocol, calibration of polarographic oxygen sensors was performed prior to mouse sciatic nerve tissue dissection and permeabilization.

1. Preparation of reagents

Prepare the Tissue Preservation Buffer (TP Buffer).
1. Prepare the following reagents in ultrapure water solution: 10 mM of Ca-EGTA buffer with 0.1 μM of free calcium, 20 mM of imidazole, 20 mM of taurine, 50 mM of K-MES, 0.5 mM of DTT, 6.56 mM of MgCl₂, 5.77 mM of ATP, 15 mM of phosphocreatine (see Table of Materials), pH 7.1. Store at -20°C.
Prepare the Mitochondria Respiration Buffer (MR Buffer).
1. Prepare the following reagents in ultrapure water solution: 0.5 mM of K₂EGTA, 3 mM of MgCl₂, 60 mM of MES, 20 mM of taurine, 10 mM of KH₂PO₄, 20 mM of HEPES, 110 mM of D-sucrose, 1 mg/mL of BSA (fatty acid-free) (see Table of Materials), pH 7.4. Store at -20 °C.
Prepare saponin stock solution by dissolving 5 mg of saponin (see Table of Materials) in 1 mL of TP Buffer (step 1.1) and keep it on ice. Saponin is prepared freshly.
Prepare Amplex Red by resuspending the powder (see Table of Materials) with DMSO to obtain a stock concentration of 2 mM and store at -20 °C in a vial protected from light.
NOTE: To avoid wearing out the probe by freezing and thawing, make small aliquots for storage no longer than 6 months²⁵.

2. Calibration of polarographic oxygen sensors for high-resolution respirometry (HRR)

Clean the HRR chambers of the instrument and syringes (see Table of Materials).
1. Open HRR chambers, fill with distilled water up to the top and stir for 5 min 3x. Repeat with ethanol and then again with water. Wash stoppers and syringes with water/ethanol/water 3x each.
Apply the following calibration settings in the HRR software (see Table of Materials).
1. In HRR software control, add the experimental temperature (37 °C), the parameters for oxygen sensor (gain, 2; polarization voltage, 800 mV), and for amperometric sensor (gain, 1000; polarization voltage, 100 mV).
Calibrate the oxygen sensors.
1. Pipette 2.1 mL of MR Buffer (step 1.2) into each chamber. Close with the stoppers and draw air into the chamber until a bubble is formed. Stir at 37 °C for 1 h in calibration mode until the oxygen flux per mass is stable.
2. Perform an air calibration of the polarographic oxygen sensors in the software according to the manufacturer's protocol²⁴.
  NOTE: The calibration step is only performed once before an experiment. Additional experiments in the same respiration medium and temperature can be performed after merely washing chambers (step 2.1).

3. Dissection and permeabilization of the sciatic nerve

Remove the sciatic nerve following the steps below.
1. Euthanize the animal by cervical dislocation after removing from its cage and gently restrained to resting on the bench.
2. In the euthanized animal, make an incision with scissors in the lower back, starting near the spine and proceeding down the thigh toward the foot. Remove skin and muscle attached to the nerve, and then cut and remove the entire sciatic nerve.
3. Weigh the tissue immediately and place it in a vial filled with cold TB Buffer (4 °C). Perform steps 3.2-3.3 on ice.
  NOTE: The wet tissue weight is used to normalize oxygen consumption and ROS production flow in the following steps. If the tissue cannot be weighed immediately, conserve it in cold TB Buffer. The procedure is performed in fresh tissue and must not be frozen to avoid mitochondrial damage.
For the tissue preparation, place the sciatic nerve in a Petri dish with enough TP Buffer to cover it. Hold one end of the nerve with forceps, and with another pair of forceps, pull out the nerve bundles horizontally.
NOTE: This procedure needs to be done in less than 10 min to avoid tissue deterioration. The tissue will be ready when it can be visualized as transparent foggy layers, opposite the previous white opaque tissue (Figure 1).
First, transfer the splayed tissue into a small dish containing 1 mL of TP Buffer for tissue permeabilization. To start permeabilization, transfer the tissue with forceps to a dish containing 1 mL of TP Buffer and 10 µL of saponin (from stock solution, step 1.3).
1. Shake in a microplate shaker gently for 30 min, then transfer the tissue with forceps to a fresh dish containing MR Buffer (1 mL) and shake gently for 10 min. Transfer the tissue with forceps to a calibrated HRR chamber.

4. Oxygen consumption and ROS production determination

Fill the HRR chambers with 2.1 mL of MR Buffer, add Amplex Red (step 1.4) to a final concentration of 5 µM and peroxidase to 2 U/mL, and add the permeabilized sciatic nerve (step 3).
1. Attach the instrument's fluorescence sensors, turn off the lights in the control section of the software, and press Connect to oxygraph. In "edit protocols" in the software, insert the tissue weight measured in step 3.1.3.
Go to "layout", choose the "specific flux per unit sample" option, and select Plots to simultaneously access the oxygen consumption readout and, if desired, the H₂O₂ production. Wait ~10 min.
NOTE: This time is required to stabilize the basal flow of oxygen consumption with no added substrate (basal). Before further injections, ensure that the oxygen flow is stabilized.
Inject two pulses of H₂O₂, each one to a final concentration of 260 µM, for later calibration in the chamber.
Inject 20 μL of succinate (see Table of Materials), a mitochondrial complex II substrate, to activate the mitochondrial electron transport system.
NOTE: Different mitochondrial substrates can be added at this point to evaluate mitochondrial function by different complexes. Representative results with varying substrates for mitochondrial complexes I and II are shown in Figure 2 and Figure 3. At this point, an increase in O₂ consumption and H₂O₂ production, simultaneously, is observed in Figure 3.
Add 20 µL of adenosine diphosphate (ADP) to activate adenosine triphosphate (ATP) synthesis.
NOTE: ADP stimulates ATP synthesis and reduces the membrane potential. An increase in O₂consumption and a decrease in the production of H₂O₂ are expected to be observed²⁶^,²⁷.
In sequence, add 5 µL of cytochrome c (see Table of Materials) as an indicator of membrane integrity.
NOTE: If the tissue is well prepared and permeabilized, cytochrome c should not increase oxygen consumption by more than 15%. If it occurs, check the troubleshooting section for recommendations.
Titrate with aliquots of 0.2 µg/mL of oligomycin (see Table of Materials) until no further decrease in O₂consumption is observed.
NOTE: Oligomycin acts by inhibiting ATP synthesis leading to a decrease in O₂ flow and an enhancement in H₂O₂formation favored by the high membrane potential²⁶^,²⁷.
Titrate with aliquots of 0.5 µmol/L of carbonyl cyanide 4-(trifluoromethoxy)phenylhydrazone (FCCP) (see Table of Materials), the mitochondrial uncoupler, until it is possible to observe no further increase in O₂ consumption. To finish the experiment, inject 2 µl of antimycin A to a final concentration of 5 μM and wait for flow stabilization.
NOTE: A decrease in H₂O₂ production is observed due to the membrane potential dissipation after injecting FCCP. Antimycin A inhibits complex III, thereby preventing the flow of electrons. Therefore, mitochondria-dependent O₂ consumption is impaired, decreasing oxygen flux per mass and stimulating electron leakage, increasing H₂O₂ production²⁶^,²⁷.
Go to the command bar, search for "multisensory" in the software, click on Control > Save file and Disconnect.
NOTE: The addition of other substrates and inhibitors can be performed according to the question under study. An example is described in the representative results. H₂O₂ calibration is performed after finishing the experiment, according to the manufacturer's protocol²⁸.
Open the saved file and select the "Oxygen Flux per Mass" trace to obtain the experimental oxygen consumption results. Manually select the window between injections by pressing Shift + Left mouse button.
1. Go to Marks > Statistics to visualize the results for each injection of substrate/inhibitors/uncoupled protocol. For H₂O₂ production, perform the same procedure with the "Amp-Slope" trace.
  NOTE: When selecting the window, avoid artifacts of volume injections by choosing a window where oxygen (or H₂O₂) is more stable and constant. Examples of selected windows are represented by black braces in the representative results (Figure 2 and Figure 3).

Representative Results

The mitochondrial oxygen consumption by the permeabilized sciatic nerve is represented in Figure 2. The red trace represents the O₂ flux per unit mass in pmol/s.mg. After recording a basal oxygen consumption with endogenous substrates (routine respiration), succinate (SUCC) is injected to record complex II (succinate dehydrogenase)-driven respiration, resulting in an increase in the oxygen consumption rate. In sequence, a saturating concentration of ADP is added, activating ATP synthase and driving oxidative phosphorylation. This results in a phosphorylative state of mitochondrial respiration. Phosphorylative state respiration means that ATP synthase can produce ATP from ADP + Pi (inorganic phosphate) using the membrane potential formed by the proton gradient as a proton-motive force to generate ATP²⁶. With the decrease in membrane potential promoted by ATP synthase activity, the oxygen consumption is accelerated, and an increase in the oxygen flow is observed. The addition of exogenous cytochrome c (CYTC) promoted only minimal stimulation of respiration, certifying mitochondrial outer membrane integrity for this preparation. Permeabilization of tissues can damage membrane integrity and loss of cytochrome c. Increased respiration rates of more than 10%-15% indicate damaged mitochondria and inadequate tissue preparation²⁸^,²⁹. In this representative result, there is an increase of 8.7% in respiration, indicating good quality of the tissue preparation (Table 1). The titration with oligomycin (OLIGO) needs to be done with minimal doses to avoid reaching a concentration of OLIGO that can interfere in maximal capacity evaluation³⁰. Inhibition of ATP synthase by OLIGO – or state 4 respiration by oligomycin²⁶ – resulted in decreased oxygen consumption flow. Titration with FCCP, a mitochondrial uncoupler reagent, dissipates mitochondrial membrane potential, stimulating the respiration flux and displaying the maximal capacity for electron transfer (ETS maximal capacity). At the end of the experiment, antimycin A (AA), an inhibitor of complex III, is injected to inhibit mitochondrial respiration and to record non-mitochondrial oxygen consumption (residual respiration) (Figure 2). Absolute oxygen flows recorded in this representative experiment are calculated in Table 1.

The possibility of analyzing mitochondrial oxygen consumption simultaneously with reactive oxygen production (ROS) – determined by detecting hydrogen peroxide (H₂O₂) by Amplex Red with fluorescence sensors – is a great advantage for determining a bioenergetic profile and a standard tool for the detection of mitochondrial dysfunction in diverse pathologies. The addition of different substrates for different complexes to fuel mitochondrial ETS can also yield more information about the specific sites for mitochondrial dysfunction. Figure 3 is shown with simultaneous measurement of oxygen consumption (Figure 3A) and ROS production (Figure 3B) in the presence of different substrates providing fuel for the ETS. After recording basal respiration, pyruvate + malate (PM) is added to the chamber as a substrate for mitochondrial complex I, increasing oxygen consumption flux. The addition of SUCC, the substrate of complex II, also increases respiration, but further addition of palmitoyl-carnitine (PC) does not increase oxygen flow compared to the earlier substrate additions, suggesting that PM and SUCC may already have saturated the mitochondrial ubiquinone site or a lack of fatty acids oxidation. As expected, ROS production also increases after the addition of substrates, representing the leak of oxygen that escapes from ETS and forms ROS (Figure 3B, green trace). The addition of ADP in saturating concentration increases oxygen consumption, driving ATP formation (red trace in Figure 3A) and decreasing ROS production (Figure 3B, green trace). The addition of CYTC only increases oxygen flow by 6.8%, confirming the mitochondrial membrane integrity. Titration with OLIGO decreases oxygen consumption flow (red trace in Figure 3A) and increases ROS production (green trace in Figure 3B). ADP and OLIGO effects suggest that permeabilized sciatic nerve in this protocol can replicate the standard relation in mitochondrial physiology, in which an increase in oxygen consumption leads to a decrease in membrane potential and prevents ROS production³¹^,³².

The addition of FCCP, as expected for an uncoupler, increases oxygen consumption to its maximal rate, and as a result of membrane potential dissipation, ROS production is decreased. The addition of rotenone, an inhibitor of complex I and AA, reduces oxygen consumption and increases ROS formation (Figure 3A,B), confirming that mitochondrial physiology and bioenergetic profile in the permeabilized sciatic nerve are preserved. Absolute values for oxygen flows recorded in this experiment are calculated in Table 2.

Figure 1: Sciatic nerve preparation for permeabilization of mitochondria. All the procedures need to be performed on ice. (A) The sciatic nerve is extracted from a euthanized mouse and placed in a Petri dish containing tissue preservation buffer at 4 °C. (B) Separation of sciatic nerve bundles with forceps. (C,D) The tissue is ready when it is dissected into translucent tufts, rather than the original white opaque tubular structure (A). Scale bar = 1 cm. Please click here to view a larger version of this figure.

Figure 2: Representative result of mitochondrial oxygen consumption in the permeabilized sciatic nerve of the mouse. The experiment is a representative trace of one sciatic nerve extract from a euthanized mouse. The sciatic nerve was permeabilized before, as described in the protocol section. Injections are indicated as arrows at specific times. SUCC: succinate; ADP: adenosine diphosphate; Cyt c: cytochrome c; OLIGO: oligomycin; FCCP: carbonyl cyanide 4-(trifluoromethoxy)phenylhydrazone; AA: antimycin A. Real-time oxygen concentration as [nmol/mL] (blue trace) and oxygen consumption rate as flux per unit mass [pmol/(s.mg)] (red trace) are shown. Black braces represent windows of oxygen consumption rates selected for results after each injection. Basal refers to respiration with no substrates. Please click here to view a larger version of this figure.

Figure 3: The result of mitochondrial oxygen consumption coupled to ROS production in the permeabilized mouse sciatic nerve. A representative trace of one sciatic nerve extract from a euthanized mouse is shown. The sciatic nerve was permeabilized before, as described in the protocol section. Injections are indicated as arrows at specific times. H₂O₂: hydrogen peroxide; PM: pyruvate/malate; SUCC: succinate; PC: palmitoyl-carnitine; ADP: adenosine diphosphate; Cyt c: cytochrome c; OLIGO: oligomycin; FCCP: carbonyl cyanide 4-(trifluoromethoxy)phenylhydrazone; ROT: rotenone; AA: antimycin A. (A) Real-time oxygen concentration as [nmol/mL] (blue trace) and oxygen consumption rate as flux per unit mass [O₂ pmol/(s.mg)] (red trace) are shown. (B) ROS production is demonstrated as H₂O₂ fluorescence (purple trace) and H₂O₂production slope [pmol/(s.mg)] (green trace), after calibration. Black braces represent windows of oxygen consumption rates selected for results after each injection. Basal refers to respiration with no substrates. Please click here to view a larger version of this figure.

Substrates	Additions	Oxygen Consumption Rate
Substrates	Additions	Flux per mass (pmol/[s.mg])
Basal (no addition)	none	5.9
Succinate (SUCC)	One addition of 10 mM	16.9
ADP	One addition of 1 µM	28.5
Cytochrome c (CYTC)	One addition of 10 μM	31
Oligomycin A (OLIGO)	Titration with additions of 0.2 μg/mL	21.9
FCCP	Titration with additions of 0.5 μM	31.1
Antimycin A (AA)	One addition of 5 μM	11.3

Table 1: Substrates/Uncoupler/Inhibitors/Titration (SUIT) protocol and results of oxygen consumption rates in mouse permeabilized sciatic nerve. Oxygen consumption is represented in [O₂pmol/(s.mg)], after each addition of SUIT protocol. Results are obtained by the average of selected windows marked in Figure 2.

Substrates	Additions	Oxygen Consumption (O₂pmol/[s.mg])	ROS Production (H₂O₂pmol/[s.mg])
Basal (no addition)	none	5.77	0.53
Pyruvate + Malate (PM)	One pulse of 5 mM + 2.5 mM	7.17	0.58
Succinate (SUCC)	One addition of 10 mM	14.06	0.75
Palmitoyl carnitine (PC)	Two pulses of 25 µM	14.68	0.79
ADP	One addition of 1 µM	22.9	0.25
Cytochrome c (CYTC)	One addition of 10 μM	24.36	0.09
Oligomycin A (OLIGO)	Titration with additions of 0.2 μg/mL	17.03	0.5
FCCP	Titration with additions of 0.5 μM	23.98	0.16
Rotenone (ROT)	One addition of 1μM	19.75	0.48
Antimycin A (AA)	One addition of 5 μM	4.08	0.53

Table 2: Substrates/Uncoupler/Inhibitors/Titration (SUIT) protocol and results of ROS production coupled to oxygen consumption rates mouse permeabilized sciatic nerve. Oxygen consumption is represented in [O₂pmol/(s.mg)] and ROS production by [H₂O₂ pmol/(s.mg)] after each addition of SUIT protocol. Results are obtained by the average of selected windows marked in Figure 3.

Parameters for Mitochondrial Function	Calculations from Oxygen Consumption Rates
Basal Respiration	Basal Respiration (flux with no addition of substrates)²⁸
Residual Oxygen Consumption (ROX)	[Flux after addition of AA]²⁸
Complex I leak respiration	[Flux after addition of PM] – [ROX]^28,37
Complex II leak respiration	[Flux after addition of SUCC] – [ROX]^28,37
Phosphorylative state ( Oxphos capacity)	[Flux after addition of ADP] – [ROX]^28,43
Non – phosphorylative state (Leak Respiration)	[Flux after addition of oligomycin] – [ROX]²⁸
Respiratory Control Ratio	[Phosphorylative state / Non -phosphorylative]^28,43
ETS Capacity	[Flux after addition of FCCP] – [ROX]^28,37

Table 3: Parameters calculation for evaluation of mitochondrial function in mouse permeabilized sciatic nerve. Parameters for mitochondrial physiology evaluation formula by oxygen consumption rates.

Discussion

Several diseases or conditions accompanying neuropathies have mitochondrial dysfunction as a risk factor. The evaluation of mitochondrial function in peripheral nerves is essential to elucidate how the mitochondria act in these neurodegenerative conditions. The assessment of mitochondrial function is laborious due to the difficulty of the isolation method and the scarcity of material. Thus, the development of tissue permeabilization techniques that do not require the isolation of mitochondria is essential.

To evaluate mitochondrial function in permeabilized tissues, choosing a chemical that can permeabilize the cell plasma membrane without interfering with cell respiration is critical. In peripheral nerves such as the sciatic nerve, myelin composition is about 70% lipid, most of which are cholesterol³³^,³⁴. The choice of permeabilizing agents such as saponin and digitonin that interact with cholesterol molecules results in pores that allow substrate access to mitochondrial machinery without interfering with other cellular compartments³⁵^,³⁶. In this protocol, the well-established method for muscle-fiber permeabilization by saponin described by Pesta and Gnaiger²⁴ is adapted for the sciatic nerves. The permeabilization procedure includes another critical step regarding tissue separation to allow for access of substrates as required for recording mitochondrial parameters of interest. The present work describes the steps, and the illustrations are provided for a satisfactory tissue separation in Figure 1.

Mitochondrial respiratory rates records are essential to establish differences regarding mitochondria complexes malfunction, oxidative phosphorylation disturbance, or disrupted mitochondria. Respiratory conditions and mitochondrial parameters are defined by Chance and Williams²⁶. In this article, the routine respiration is defined as basal respiration when mitochondrial oxygen consumption is recorded with no addition of exogenous substrates; complexes capacity – when substrates for complex I or II are added to fuel mitochondrial oxygen consumption, also named as leak respiration; phosphorylative state – when ADP is added in the sequence of complexes substrates and drives ATP synthesis; non-phosphorylative state – when all ADP is formed into ATP or with the addition of ATP synthase inhibitor, oligomycin (or state 4); ETS maximal capacity – when the mitochondrial uncoupler FCCP is added and; residual oxygen consumption (ROX) after addition of rotenone and antimycin A – when the oxygen consumption is not related to mitochondria. The technique described in this work for mitochondrial function evaluation in sciatic nerve tissue by HRR allows determining numerous respiratory states and calculation of intrinsic mitochondrial function within the same sample. A substrate/uncoupled/inhibitors protocol test results can be used to derivate parameters of mitochondrial function from which respiratory control ratios/factors can be calculated³⁷, as demonstrated in Table 3. Evaluation of these parameters combines absolute leak rates, coupled with maximal mitochondrial respiration, and raises data for a diagnostic value²⁴^,³⁵^,³⁶^,³⁷^,³⁸^,³⁹.

Although mitochondrial dysfunction in peripheral nerves is cited in the literature, there are limitations in demonstrating these parameters. In a chemotherapy-induced neuropathy model, a decrease in the mitochondrial membrane potential in treatment with the chemotherapy agent, oxaliplatin, and a reduction of the in vitro activity of complexes I, II, and III is observed in sensory axons of the peripheral nerve (saphenous nerve)⁴⁰^,⁴¹. However, there is a lack of information on the control exerted by membrane potential on the complexes' activities and formation of ATP by oxidative phosphorylation, an essential parameter in mitochondrial function evaluation. Also, in dissociated sciatic nerves from Sprague-Dawley rats permeabilized with digitonin, the recording of oxygen consumption is only demonstrated for the phosphorylated state, but is not compared with oxygen consumption in the presence of an ATPase inhibitor or an uncoupler²². In the present method, it is possible to calculate respiratory control ratio in the permeabilized nerve (Table 3) compared with the respiratory control ratio of isolated mitochondria, usually calculated from state3/state4 (or phosphorylative/non-phosphorylative states). With the method provided in this work, it is possible to assess complex I, II capacities – maintaining mitochondrial membrane integrity – and flux control ratios, and achieve maximal capacity respiration and residual oxygen consumption.

Different methods for sciatic nerve permeabilization for mitochondrial oxygen consumption evaluation are described in the literature. A technique for sciatic nerve permeabilization is demonstrated in a protocol with saponin treatment and permeabilization by vortex pulse; however, only phosphorylative state and maximal capacity were demonstrated⁴¹. In addition, comparing the values presented here for the same parameters, the oxygen flux observed by Cooper et al.⁴¹ is 70% lower, demonstrating that there is damage to the mitochondrial membrane related to the agitation method. In the present protocol, the tissue separation preserves mitochondrial membrane integrity, confirmed by the small increase in oxygen flux after the addition of cytochrome c. This is a feature that allows all mitochondrial parameters to be recorded, providing a complete record of mitochondrial oxidative phosphorylation and function. Even though in a sciatic nerve permeabilization method with collagenase, the oxygen flux values are comparable to those presented here, an increase of nearly 20% is observed after adding cytochrome c, suggesting some level of mitochondrial membrane damage²¹. With the protocol described in this article, an increase of only 6.3%-8.7% of oxygen flux after cytochrome c addition was observed, confirming the advantage of the present method in preserving the mitochondrial membrane integrity and optimizing oxygen consumption recording.

Mitochondria are the primary site of reactive oxygen species (ROS) generation associated with diverse signaling processes in sciatic nerve tissue and mechanisms related to pathophysiology in neuropathies⁴²^,⁴³. This protocol made it possible to access hydrogen peroxide generation with a fluorescence sensor simultaneously with oxygen consumption. As observed in Figure 3, ROS production is sensitive to alterations of mitochondrial respiratory states, confirming the mitochondrial integrity status and optimizing mitochondrial function recording.

Other methods such as extracellular flux analyzer (EFA), based on fluorescence sensors to record oxygen consumption, can evaluate peripheral-nerve mitochondrial function in cultured cells – such as Schwann cells or axons – in an automated high-throughput screening device. However, there are some advantages in using polarographic sensors such as HRR, including in particular the ability to follow the experiment in real-time, to make multiple injections of substrates/inhibitors; thereby, this permits evaluation of a greater number of complex functions in mitochondria in one experiment, and the lower cost of consumables²⁴^,³⁵^,³⁸^,³⁹. A disadvantage of using HRR compared to EFA is the absence of a sensor for media acidification (for analysis of glycolytic flux) and the limited workflow capability, which allows the use of only two samples at a time.

Limitations of tissue permeabilization techniques required for HRR are well known. They include the inability to evaluate mitochondria when ADP is limiting and a reduced uncoupled respiratory rate compared to isolated mitochondria. However, the saponin-sciatic nerve permeabilization protocol described here – as an adaptation of the muscle-fiber permeabilization protocol by Pesta and Gnaiger²⁴ – allows the assessment of mitochondrial respiratory states and mitochondrial parameters without damaging mitochondrial integrity. It also allows for simultaneously recording oxygen consumption and ROS production in the sciatic nerve, a tissue with notorious limitations for isolating mitochondria. Thus, this protocol allows for a better assessment of mitochondrial function in peripheral nerves and can provide advantages to researchers in the investigation of mitochondrial roles in pathophysiological conditions afflicting peripheral nerves.

Troubleshooting
If there is an increase in O₂consumption after adding cytochrome c to more than 15% of oxygen consumption flux, it indicates that the permeabilization procedure was very strong and caused damage to the mitochondrial structure. If that is the case, discard and reinitiate the dissection with a gentler nerve separation and repeat the steps. If some reagents injected do not demonstrate the expected effect in mitochondria oxygen consumption or H₂O₂ production, it is probably due to chamber contamination with mitochondrial inhibitors. In this case, two actions can be taken: (1) restart the experiment after washing the chambers and syringes (step 2.1) or (2) add a sample of any tissue-isolated mitochondria or homogenate before performing step 2.1. The additions of these samples will absorb mitochondrial inhibitors and improve the cleaning step.

Disclosures

The authors have nothing to disclose.

Acknowledgements

This study was financed by Instituto Serrapilheira, Fundação de Amparo à Pesquisa do Estado do Rio de Janeiro (FAPERJ), Conselho Nacional de Desenvolvimento Científico e Tecnológico (CNPq) and Coordenação de Aperfeiçoamento de Pessoal de Nível Superior-Brasil (CAPES). We are grateful to Dr. Antonio Galina Filho, Dr. Monica Montero Lomeli and Dr. Claudio Masuda for the support with laboratory facilities, and Dr. Martha Sorenson for kind and valuable comments in improving the article.

Materials

Adenosine 5' triphosphate dissodium salt hydrate	Sigma-Aldrich	A26209
Adenosine 5′-diphosphate sodium salt	Sigma-Aldrich	A2754
Amplex Red Reagent	Thermo Fisher scientific	A12222	Amplex Red is prepared in DMSO accordindly with product datasheet
Antimycin A (from Streptomyces sp.)	Sigma-Aldrich	A8674
Bovine Serum Albumin	Sigma-Aldrich	A7030	heat shock fraction, protease free, fatty acid free, essentially globulin free, pH 7, ≥98%
Calcium carbonate	Sigma-Aldrich	C6763
Carbonyl cyanide 4-(trifluoromethoxy)phenylhydrazone (FCCP)	Sigma-Aldrich	C2920
Cytochrome c	Sigma-Aldrich	C7752	(from equine heart; small hemeprotein)
DataLab version 5.1.1.91	OROBOROS INSTRUMENTS, Austria		Copyright (c) 2002 – 13 by Dr. Erich Gnaiger
Digital orbital microplate shaker 120V	Thermo Fisher scientific	88882005
DL-Dithiothreitol	Sigma-Aldrich	43819
EGTA sodium salt	Sigma-Aldrich	E8145
Hamilton syringe	Sigma-Aldrich	HAM80075	10 uL, 25 uL and 50 uL
HEPES	Sigma-Aldrich	H3375
Hydrogen peroxide solution 30% W/W	Merck	H1009
Imidazole	Sigma-Aldrich	I2399
L-(−)-Malic acid	Sigma-Aldrich	M7397
Magnesium chloride hexahydrate	Sigma-Aldrich	M2393
MES sodium salt	Sigma-Aldrich	M3885
Micro-dissecting forceps, curved	Sigma-Aldrich	F4142
Micro-dissecting forceps, straight	Sigma-Aldrich	F4017
O2K – Filter set Amplex Red	OROBOROS INSTRUMENTS, Austria	44321-01	Fasching M, Sumbalova Z, Gnaiger E (2013) O2k-Fluorometry: HRR and H2O2 production in mouse brain mitochondria. Mitochondr Physiol Network 17.17.
O2K – Fluorescence LED2 – module component Fluorscence-Sensor Green	OROBOROS INSTRUMENTS, Austria	44210-01
Oligomycin	Sigma-Aldrich	O4876	(from Streptomyces diastatochromogenes; mixture of oligomycins A, B, and C
OROBOROS Oxygraph-2k	OROBOROS INSTRUMENTS, Austria	http://www.oroboros.at
Palmitoylcarnitine (Palmitoyl-DL-carnitine-HCl)	Sigma-Aldrich	P4509
Peroxidase from horseradish	Sigma-Aldrich	P8375
Petri dishes, polystyrene	MERCK	P5606
Phosphocreatine disodium salt hydrate	Sigma-Aldrich	P7936
Potassium dihydrogen phosphate monobasic	Sigma-Aldrich	PHR1330
Potassium hydroxide	Sigma-Aldrich	221473
Rotenone	Sigma-Aldrich	R8875
Saponin	Sigma-Aldrich	SAE0073
Sodium pyruvate	Sigma-Aldrich	P5280
Sodium succinate dibasic hexahydrate	Sigma-Aldrich	S2378
Sucrose	Sigma-Aldrich	S9378
Taurine	Sigma-Aldrich	T0625

References

Pfanner, N., Warscheid, B., Wiedemann, N. Mitochondrial protein organization: from biogenesis to networks and function. Nature Reviews Molecular Cell Biology. 20 (5), 267-284 (2019).
Sena, L. A., Chandel, N. S. Physiological roles of mitochondrial reactive oxygen species. Molecular Cell. 48 (2), 158-167 (2012).
Van Der Bliek, A. M., Sedensky, M. M., Morgan, P. G. Cell biology of the mitochondrion. 유전학. 207 (3), 843-871 (2017).
Rugarli, E. I., Langer, T. Mitochondrial quality control: A matter of life and death for neurons. EMBO Journal. 31 (6), 1336-1349 (2012).
Westermann, B. Mitochondrial fusion and fission in cell life and death. Nature Reviews Molecular Cell Biology. 11, 872-884 (2010).
Pickles, S., Vigié, P., Youle, R. J. Mitophagy and quality control mechanisms in mitochondrial maintenance. Current Biology. 28 (4), 170-185 (2018).
Freeman, O. J., et al. Metabolic dysfunction is restricted to the sciatic nerve in experimental diabetic neuropathy. Diabetes. 65 (1), 228-238 (2016).
Sheng, B., et al. Impaired mitochondrial biogenesis contributes to mitochondrial dysfunction in Alzheimer’s disease. Journal of Neurochemistry. 120 (3), 419-429 (2012).
Wang, X., et al. Oxidative stress and mitochondrial dysfunction in Alzheimer’s disease. Biochimica et Biophysica Acta – Molecular Basis of Disease. 1842 (8), 1240-1247 (2014).
Li, W., Fu, Y. H., Halliday, G. M., Sue, C. M. PARK genes link mitochondrial dysfunction and alpha-synuclein pathology in sporadic Parkinson’s disease. Frontiers in Cell and Developmental Biology. 9, 1-11 (2021).
Winklhofer, K. F., Haass, C. Mitochondrial dysfunction in Parkinson’s disease. Biochimica et Biophysica Acta – Molecular Basis of Disease. 1802 (1), 29-44 (2010).
Harley, J., Clarke, B. E., Patani, R. The interplay of rna binding proteins, oxidative stress and mitochondrial dysfunction in ALS. Antioxidants. 10 (4), 552 (2021).
Nakagawa, Y., Yamada, S. A novel hypothesis on metal dyshomeostasis and mitochondrial dysfunction in amyotrophic lateral sclerosis: Potential pathogenetic mechanism and therapeutic implications. European Journal of Pharmacology. 892, 173737 (2021).
Franco-Iborra, S., et al. Mutant HTT (huntingtin) impairs mitophagy in a cellular model of Huntington disease. Autophagy. 17 (3), 672-689 (2021).
Wang, Y., Guo, X., Ye, K., Orth, M., Gu, Z. Accelerated expansion of pathogenic mitochondrial DNA heteroplasmies in Huntington’s disease. Proceedings of the National Academy of Sciences of the United States of America. 118 (30), 2014610118 (2021).
Sajic, M., et al. Mitochondrial damage and ‘plugging’ of transport selectively in myelinated, small-diameter axons are major early events in peripheral neuroinflammation. Journal of Neuroinflammation. 15 (1), 61 (2018).
Muke, I., et al. Ultrastructural characterization of mitochondrial damage in experimental autoimmune neuritis. Journal of Neuroinflammation. 343, 577218 (2020).
Rodella, U., et al. An animal model of Miller Fisher Syndrome: mitochondrial hydrogen peroxide is produced by the autoimmune attack of nerve terminals and activates Schwann cells. Neurobiology of Disease. 96, 95-104 (2016).
Han, M. M., Frizzi, K. E., Ellis, R. J., Calcutt, N. A., Fields, J. A. Prevention of HIV-1 TAT protein-induced Ppripheral neuropathy and mitochondrial disruption by the antimuscarinic pirenzepine. Frontiers in Neurology. 12, 663373 (2021).
Roda, R. H., Hoke, A. Mitochondrial dysfunction in HIV-induced peripheral neuropathy. International Review of Neurobiology. 145, (2019).
Palavicini, J. P., et al. Early disruption of nerve mitochondrial and myelin lipid homeostasis in obesity-induced diabetes. JCI Insight. 5 (21), 137286 (2020).
Zheng, H., Xiao, W. H., Bennett, G. J. Functional deficits in peripheral nerve mitochondria in rats with paclitaxel- and oxaliplatin-evoked painful peripheral neuropathy. Experimental Neurology. 232 (2), 154-161 (2011).
Lim, T. K. Y., Rone, M. B., Lee, S., Antel, J. P., Zhang, J. Mitochondrial and bioenergetic dysfunction in trauma-induced painful peripheral neuropathy. Molecular Pain. 11, 58 (2015).
Pesta, D., Gnaiger, E. High-resolution respirometry: OXPHOS protocols for human cells and permeabilized fibers from small biopsies of human muscle. Mitochondrial Bioenergetics: Methods and Protocols (Methods in Molecular Biology. 810, 25-58 (2012).
Komlódi, T., et al. Comparison of mitochondrial incubation media for measurement of respiration and hydrogen peroxide production. Methods in Molecular Biology. 1782, 137-155 (2018).
Chance, B., Williams, G. R. Respiratory enzymes in oxidative phosphorylation. III. The steady state. Journal of Biological Chemistry. 217 (1), 409-427 (1955).
Korshunov, S. S., Skulachev, V. P., Starkov, A. A. High protonic potential actuates a mechanism of production of reactive oxygen species in mitochondria. FEBS Letters. 416 (1), 15-18 (1997).
Gnaiger, E. Mitochondr Physiol Network. Mitochondrial Pathways and Respiratory Control. An Introduction to OXPHOS Analysis. 4th ed. , 80 (2014).
Kuznetsov, A. V., et al. Mitochondrial defects and heterogeneous cytochrome c release after cardiac cold ischemia and reperfusion. American Journal of Physiology-Heart and Circulatory Physiology. 286 (5), 1633-1641 (2004).
Ruas, J. S., et al. Underestimation of the maximal capacity of the mitochondrial electron transport system in oligomycin-treated cells. PLoS One. 11 (3), 0150967 (2016).
Boveris, A., Chance, B. The mitochondrial generation of hydrogen peroxide. General properties and effect of hyperbaric oxygen. Biochemical Journal. 134 (3), 707-716 (1973).
Skulachev, V. P. Membrane-linked systems preventing superoxide formation. Bioscience Reports. 17 (3), 347-366 (1997).
Majava, V., et al. Structural and functional characterization of human peripheral nervous system myelin protein P2. PLoS One. 5, 10300 (2010).
Greenfield, S., Brostoff, S., Eylar, E. H., Morell, P. Protein composition of myelin of the peripheral nervous system. Journal of Neurochemistry. 20 (4), 1207-1216 (1973).
Kuznetsov, A. V., et al. Analysis of mitochondrial function in situ in permeabilized muscle fibers, tissues and cells. Nature Protocols. 3, 965-976 (2008).
Saks, V. A., et al. Permeabilized cell and skinned fiber techniques in studies of mitochondrial function in vivo. Molecular and Cellular Biochemistry. 184 (1-2), 81-100 (1998).
Gnaiger, E. Capacity of oxidative phosphorylation in human skeletal muscle. New perspectives of mitochondrial physiology. The International Journal of Biochemistry & Cell Biology. 41 (10), 1837-1845 (2009).
Porter, C., et al. Mitochondrial respiratory capacity and coupling control decline with age in human skeletal muscle. American Journal of Physiology-Endocrinology and Metabolism. 309 (3), 224-232 (2015).
Martins, E. L., et al. Rapid regulation of substrate use for oxidative phosphorylation during a single session of high intensity interval or aerobic exercises in different rat skeletal muscles. Comparative Biochemistry and Physiology B. 217, 40-50 (2018).
Areti, A., Komirishetty, P., Kumar, A. Carvedilol prevents functional deficits in peripheral nerve mitochondria of rats with oxaliplatin-evoked painful peripheral neuropathy. Toxicology and Applied Pharmacology. 322, 97-103 (2017).
Cooper, M. A., et al. Reduced mitochondrial reactive oxygen species production in peripheral nerves of mice fed a ketogenic diet. Experimental Physiology. 103 (9), 1206-1212 (2018).
Jia, M., et al. Activation of NLRP3 inflammasome in peripheral nerve contributes to paclitaxel-induced neuropathic pain. Molecular Pain. 13, 1744806917719804 (2017).
Muller, F. L., et al. Denervation-induced skeletal muscle atrophy is associated with increased mitochondrial ROS production. American Journal of Physiology-Regulatory, Integrative and Comparative Physiology. 293 (3), 1159-1168 (2007).

Play Video

PDF

DOI

DOWNLOAD MATERIALS LIST

Cite This Article

Formiga-Jr, M. A., Camacho-Pereira, J. Assessing Mitochondrial Function in Sciatic Nerve by High-Resolution Respirometry. J. Vis. Exp. (183), e63690, doi:10.3791/63690 (2022).