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In This Article

  • Summary
  • Abstract
  • Introduction
  • Protocol
  • Results
  • Discussion
  • Disclosures
  • Acknowledgements
  • Materials
  • References
  • Reprints and Permissions

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

  1. 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.
  2. 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.
  3. 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.
  4. 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)

  1. 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.
  2. 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).
  3. 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

  1. 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.
  2. 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).
  3. 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

  1. 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.
  2. 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.
  3. Inject two pulses of H2O2, each one to a final concentration of 260 µM, for later calibration in the chamber.
  4. 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.
  5. 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 O2 consumption and a decrease in the production of H2O2 are expected to be observed26,27.
  6. 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.
  7. Titrate with aliquots of 0.2 µg/mL of oligomycin (see Table of Materials) until no further decrease in O2 consumption is observed.
    NOTE: Oligomycin acts by inhibiting ATP synthesis leading to a decrease in O2 flow and an enhancement in H2O2 formation favored by the high membrane potential26,27.
  8. 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.
  9. 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.
  10. 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).

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 sy...

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.

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

NameCompanyCatalog NumberComments
Adenosine 5' triphosphate dissodium salt hydrateSigma-AldrichA26209
Adenosine 5′-diphosphate sodium saltSigma-AldrichA2754
Amplex Red ReagentThermo Fisher scientificA12222Amplex Red is prepared in DMSO accordindly with product datasheet
Antimycin A (from Streptomyces sp.)Sigma-AldrichA8674
Bovine Serum AlbuminSigma-AldrichA7030heat shock fraction, protease free, fatty acid free, essentially globulin free, pH 7, ≥98%
Calcium carbonateSigma-AldrichC6763
Carbonyl cyanide 4-(trifluoromethoxy)phenylhydrazone (FCCP)Sigma-AldrichC2920
Cytochrome cSigma-AldrichC7752(from equine heart; small hemeprotein)
DataLab version 5.1.1.91OROBOROS INSTRUMENTS, AustriaCopyright (c) 2002 - 13 by Dr. Erich Gnaiger
Digital orbital microplate shaker 120VThermo Fisher scientific88882005
DL-DithiothreitolSigma-Aldrich43819
EGTA sodium saltSigma-AldrichE8145
Hamilton syringeSigma-AldrichHAM8007510 uL, 25 uL and 50 uL
HEPESSigma-AldrichH3375
Hydrogen peroxide solution 30% W/WMerckH1009
ImidazoleSigma-AldrichI2399
L-(−)-Malic acidSigma-AldrichM7397
Magnesium chloride hexahydrateSigma-AldrichM2393
MES sodium saltSigma-AldrichM3885
Micro-dissecting forceps, curvedSigma-AldrichF4142
Micro-dissecting forceps, straightSigma-AldrichF4017
O2K - Filter set Amplex RedOROBOROS INSTRUMENTS, Austria44321-01Fasching 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 GreenOROBOROS INSTRUMENTS, Austria44210-01
OligomycinSigma-AldrichO4876(from Streptomyces diastatochromogenes; mixture of oligomycins A, B, and C
OROBOROS Oxygraph-2kOROBOROS INSTRUMENTS, Austriahttp://www.oroboros.at
Palmitoylcarnitine (Palmitoyl-DL-carnitine-HCl)Sigma-AldrichP4509
Peroxidase from horseradishSigma-AldrichP8375
Petri dishes, polystyreneMERCKP5606
Phosphocreatine disodium salt hydrateSigma-AldrichP7936
Potassium dihydrogen phosphate monobasicSigma-AldrichPHR1330
Potassium hydroxideSigma-Aldrich221473
RotenoneSigma-AldrichR8875
SaponinSigma-AldrichSAE0073
Sodium pyruvateSigma-AldrichP5280
Sodium succinate dibasic hexahydrateSigma-AldrichS2378
SucroseSigma-AldrichS9378
TaurineSigma-AldrichT0625

References

  1. 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).
  2. Sena, L. A., Chandel, N. S. Physiological roles of mitochondrial reactive oxygen species. Molecular Cell. 48 (2), 158-167 (2012).
  3. Van Der Bliek, A. M., Sedensky, M. M., Morgan, P. G. Cell biology of the mitochondrion. Genetics. 207 (3), 843-871 (2017).
  4. Rugarli, E. I., Langer, T. Mitochondrial quality control: A matter of life and death for neurons. EMBO Journal. 31 (6), 1336-1349 (2012).
  5. Westermann, B. Mitochondrial fusion and fission in cell life and death. Nature Reviews Molecular Cell Biology. 11, 872-884 (2010).
  6. Pickles, S., Vigié, P., Youle, R. J. Mitophagy and quality control mechanisms in mitochondrial maintenance. Current Biology. 28 (4), 170-185 (2018).
  7. Freeman, O. J., et al. Metabolic dysfunction is restricted to the sciatic nerve in experimental diabetic neuropathy. Diabetes. 65 (1), 228-238 (2016).
  8. Sheng, B., et al. Impaired mitochondrial biogenesis contributes to mitochondrial dysfunction in Alzheimer's disease. Journal of Neurochemistry. 120 (3), 419-429 (2012).
  9. 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).
  10. 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).
  11. Winklhofer, K. F., Haass, C. Mitochondrial dysfunction in Parkinson's disease. Biochimica et Biophysica Acta - Molecular Basis of Disease. 1802 (1), 29-44 (2010).
  12. 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).
  13. 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).
  14. Franco-Iborra, S., et al. Mutant HTT (huntingtin) impairs mitophagy in a cellular model of Huntington disease. Autophagy. 17 (3), 672-689 (2021).
  15. 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).
  16. 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).
  17. Muke, I., et al. Ultrastructural characterization of mitochondrial damage in experimental autoimmune neuritis. Journal of Neuroinflammation. 343, 577218 (2020).
  18. 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).
  19. 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).
  20. Roda, R. H., Hoke, A. Mitochondrial dysfunction in HIV-induced peripheral neuropathy. International Review of Neurobiology. 145, (2019).
  21. Palavicini, J. P., et al. Early disruption of nerve mitochondrial and myelin lipid homeostasis in obesity-induced diabetes. JCI Insight. 5 (21), 137286 (2020).
  22. 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).
  23. 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).
  24. 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).
  25. 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).
  26. Chance, B., Williams, G. R. Respiratory enzymes in oxidative phosphorylation. III. The steady state. Journal of Biological Chemistry. 217 (1), 409-427 (1955).
  27. 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).
  28. Gnaiger, E. Mitochondr Physiol Network. Mitochondrial Pathways and Respiratory Control. An Introduction to OXPHOS Analysis. 4th ed. , 80 (2014).
  29. 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).
  30. 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).
  31. Boveris, A., Chance, B. The mitochondrial generation of hydrogen peroxide. General properties and effect of hyperbaric oxygen. Biochemical Journal. 134 (3), 707-716 (1973).
  32. Skulachev, V. P. Membrane-linked systems preventing superoxide formation. Bioscience Reports. 17 (3), 347-366 (1997).
  33. Majava, V., et al. Structural and functional characterization of human peripheral nervous system myelin protein P2. PLoS One. 5, 10300 (2010).
  34. 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).
  35. Kuznetsov, A. V., et al. Analysis of mitochondrial function in situ in permeabilized muscle fibers, tissues and cells. Nature Protocols. 3, 965-976 (2008).
  36. 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).
  37. 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).
  38. 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).
  39. 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).
  40. 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).
  41. 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).
  42. Jia, M., et al. Activation of NLRP3 inflammasome in peripheral nerve contributes to paclitaxel-induced neuropathic pain. Molecular Pain. 13, 1744806917719804 (2017).
  43. 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).

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