# MITOCHONDRIA IN SKELETAL MUSCLE HEALTH, AGING AND DISEASES

EDITED BY: Gilles Gouspillou and Russell T. Hepple PUBLISHED IN: Frontiers in Physiology

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ISSN 1664-8714 ISBN 978-2-88945-073-2 DOI 10.3389/978-2-88945-073-2

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# **MITOCHONDRIA IN SKELETAL MUSCLE HEALTH, AGING AND DISEASES**

Topic Editors:

**Gilles Gouspillou,** Université du Québec à Montréal, Canada **Russell T. Hepple,** McGill University, Canada

Immunolabeling for Endonuclease G (red; a mitochondrial pro-apoptotic protein translocating to nuclei when mitochondria trigger apoptosis), dystrophin (green), and nuclei (blue) obtained on a muscle cross section of an old men.

Image taken and adapted, with permission, from: Gouspillou G, Sgarioto N, Kapchinsky S, Purves-Smith F, Norris B, Pion CH, Barbat-Artigas S, Lemieux F, Taivassalo T, Morais JA, Aubertin-Leheudre M, Hepple RT. Increased sensitivity to mitochondrial permeability transition and myonuclear translocation of endonuclease G in atrophied muscle of physically active older humans. FASEB J. 2014 Apr;28(4):1621-33. doi: 10.1096/fj.13-242750. Epub 2013 Dec 26.

Skeletal muscle is the most abundant tissue of the human body, making up to 40 to 50% of the human body mass. While the importance of optimal muscle function is well recognized in the athletic field, its significance for general health is often underappreciated. In fact, the evidence that muscle mass, strength and metabolism are essential for our overall health is overwhelming. As the largest protein reservoir in the human body, muscles are essential in the acute response to critical illness such as sepsis, advanced cancer, and traumatic injury. Loss of skeletal muscle mass has also been associated with weakness, fatigue, insulin resistance, falls, fractures, frailty, disability, several chronic diseases and death. As a consequence, maintaining skeletal muscle mass, strength and metabolism throughout the lifespan is critical to the maintenance of whole body health.

Mitochondria are fascinating organelles regulating many critical cellular processes for skeletal muscle physiology, including for instance energy supply, reactive oxygen species production, calcium homeostasis and the regulation of apoptosis. It is therefore not surprising that mitochondrial dysfunction has been implicated in a large number of adverse events/conditions and pathologies affecting skeletal muscle health.

While the importance of normal mitochondrial function is well recognized for muscle physiology, there are important aspects of mitochondrial biology that are still poorly understood. These include mitochondrial dynamics (fusion and fission processes), morphology and processes involved in mitochondrial quality control (mitophagy). Defining the mechanisms regulating these different aspects of mitochondrial biology, their importance for muscle physiology, as well as the interrelations will be critical for expanding understanding of the role played by mitochondria in skeletal muscle physiology and health.

The present research topic provides readers with novel experimental approaches, knowledge, hypotheses and findings related to all aspects of mitochondrial biology in healthy and diseased muscle cells.

**Citation:** Gouspillou, G., Hepple, R. T., eds. (2017). Mitochondria in Skeletal Muscle Health, Aging and Diseases. Lausanne: Frontiers Media. doi: 10.3389/978-2-88945-073-2

# Table of Contents

Philip A. Kramer, Jicheng Duan, Wei-Jun Qian and David J. Marcinek

*44 Hypothesis on Skeletal Muscle Aging: Mitochondrial Adenine Nucleotide Translocator Decreases Reactive Oxygen Species Production While Preserving Coupling Efficiency*

Philippe Diolez, Isabelle Bourdel-Marchasson, Guillaume Calmettes, Philippe Pasdois, Dominique Detaille, Richard Rouland and Gilles Gouspillou

*53 Protein Kinase A Governs Oxidative Phosphorylation Kinetics and Oxidant Emitting Potential at Complex I*

Daniel S. Lark, Lauren R. Reese, Terence E. Ryan, Maria J. Torres, Cody D. Smith, Chien-Te Lin and P. Darrell Neufer

Daniel M. Craig, Stephen P. Ashcroft, Micah Y. Belew, Ben Stocks, Kevin Currell, Keith Baar and Andrew Philp

*83 On the Mechanism by Which Dietary Nitrate Improves Human Skeletal Muscle Function*

Charles Affourtit, Stephen J. Bailey, Andrew M. Jones, Miranda J. Smallwood and Paul G. Winyard

*91* **In utero** *Undernutrition Programs Skeletal and Cardiac Muscle Metabolism* Brittany Beauchamp and Mary-Ellen Harper

Terence E. Ryan, Cameron A. Schmidt, Tom D. Green, David A. Brown, P. Darrell Neufer and Joseph M. McClung

*128 Complex Coordination of Cell Plasticity by a PGC-1a-controlled Transcriptional Network in Skeletal Muscle*

Barbara Kupr and Christoph Handschin

*135 Regulation of Skeletal Muscle Mitochondrial Activity by Thyroid Hormones: Focus on the "Old" Triiodothyronine and the "Emerging" 3,5-Diiodothyronine* Assunta Lombardi, Maria Moreno, Pieter de Lange, Susanna Iossa, Rosa A. Busiello and Fernando Goglia

# Editorial: Mitochondria in Skeletal Muscle Health, Aging and Diseases

Gilles Gouspillou1, 2, 3 \* and Russell T. Hepple4, 5

1 Département des Sciences de l'Activité Physique, Faculté des Sciences, Université du Québec à Montréal, Montreal, QC, Canada, 2 Groupe de Recherche en Activité Physique Adaptée, Montreal, QC, Canada, 3 Centre de Recherche de l'Institut, Universitaire de Gériatrie de Montréal, Montreal, QC, Canada, 4 Department of Kinesiology and Physical Education, McGill University, Montreal, QC, Canada, 5 Meakins-Christie Laboratory, Research Institute of the McGill University Health Centre, Montreal, QC, Canada

Keywords: muscle, aging, mitochondria, metabolism, mitophagy, mitochondrial dynamics, bioenergetics, muscle fibers

**The Editorial on the Research Topic**

#### **Mitochondria in Skeletal Muscle Health, Aging and Diseases**

Mitochondria are fascinating organelles regulating many critical cellular processes for skeletal muscle physiology. Indeed, they play central roles in muscle cell metabolism, energy supply, the regulation of energy-sensitive signaling pathways, reactive oxygen species (ROS) production/signaling, calcium homeostasis and the regulation of apoptosis (Brookes et al., 2004). Given these multifaceted roles of mitochondria in fundamental aspects of skeletal muscle cell physiology, it is not surprising that mitochondrial dysfunction has been implicated in a large number of adverse conditions affecting skeletal muscle health. This includes for instance the agingrelated loss of muscle mass and function (Dirks and Leeuwenburgh, 2004; Short et al., 2005; Chabi et al., 2008; Gouspillou et al., 2010, 2014a,b; Picard et al., 2010; Hepple, 2014), disuse-induced muscle atrophy (Min et al., 2011), ventilator-induced diaphragmatic dysfunction (Picard et al., 2015), Duchenne and collagen muscular dystrophies (Godin et al., 2012; Bernardi and Bonaldo, 2013), long-term muscle dysfunction induced by chemotherapy treatment (Gouspillou et al., 2015; Power et al., 2016), and the development of insulin resistance (Goodpaster, 2013).

While the importance of normal mitochondrial function is well recognized for muscle physiology, there are important aspects of mitochondrial biology that are still poorly understood/investigated in the highly specialized muscle tissue. These include mitochondrial dynamics (fusion and fission processes), morphology and processes involved in mitochondrial quality control (mitophagy). Defining the mechanisms regulating these different aspects of mitochondrial biology, their importance for muscle physiology, as well as the interrelations existing between mitochondrial function, morphology, dynamics and mitophagy will be critical to further increase our understanding of the role played by mitochondria in skeletal muscle physiology and pathophysiology. The aim of the present research topic was therefore to bring together key experiments, advances, knowledge and new findings related to all aspects of mitochondrial biology in healthy and/or diseased muscle cells.

The accurate assessment of mitochondrial function is of crucial importance to dissect the role played by mitochondrial dysfunction in pathologies affecting skeletal muscles. It is also essential for the clear identification of mitochondrial adaptations to various interventions, such as nutritional or physical activity interventions. In the present research topic, Conley et al. provide a promising approach to assess mitochondrial oxidative phosphorylation in vivo by monitoring changes in mitochondrial NAD(P)H using 31P NMR spectroscopy. In their study, Conley et al. show that a unique resonance (−11.05 ppm) in the in vivo 31P spectrum provides a natural indicator of

Edited and reviewed by:

Peter J. Reiser, Ohio State University, USA

\*Correspondence: Gilles Gouspillou<EMAIL_ADDRESS>
#### Specialty section:

This article was submitted to Striated Muscle Physiology, a section of the journal Frontiers in Physiology

Received: 31 August 2016 Accepted: 20 September 2016 Published: 06 October 2016

#### Citation:

Gouspillou G and Hepple RT (2016) Editorial: Mitochondria in Skeletal Muscle Health, Aging and Diseases. Front. Physiol. 7:446. doi: 10.3389/fphys.2016.00446 mitochondrial oxidation that is sensitive to the increase in mitochondrial phosphorylation rate induced by exercise training in elderly individuals.

One of the commonly used methods to assess mitochondrial function in vitro in skeletal muscle is the preparation of permeabilized myofibers (Kuznetsov et al., 2008). To be applied in humans, this technique requires biopsies to collect muscle samples. While the Bergstrom biopsy technique (Bergstrom, 1975) is the gold standard to collect skeletal muscle samples, a growing interest now surrounds microbiospsies due to their lower level of invasiveness (Hayot et al., 2005). However, no previous in-depth validation of this method for the assessment of mitochondrial function in permeabilized myofibers and for the assessment of muscle phenotype had been performed to date. Hughes et al. therefore compared mitochondrial bioenergetics in muscle sample obtained through Bergstrom biopsies vs. microbiopsies. They reveal that microbiopsies can provide a reliable assessment of mitochondrial bioenergetics only when assay conditions are supplemented with the myosin ATPase inhibitor Blebistatin. However, the authors highlight that caution should be taken when assessing muscle fiber type composition using the microbiopsy approach, since significant differences in fiber type proportion were observed between the two approaches (Hughes et al.).

Oxidative stress is thought to play an important role in skeletal muscle dysfunction and atrophy seen in aging, disuse, and many skeletal muscle pathologies (Powers et al., 2012; Johnson et al., 2013). Because they are considered as one of the main sources of ROS production, mitochondria are a key focus in the field of oxidative stress. While reactive oxygen and nitrogen species were initially only seen as detrimental for muscle cells, it is now recognized that these reactive species are essential for normal skeletal muscle physiology (Sohal and Orr, 2012), mainly through the reversible redox post-translational modifications they can induce. The ability to accurately quantify reversible redox post-translational modifications is therefore critical to investigate the mechanisms by which mitochondrial oxidative stress contributes to skeletal muscle dysfunction in diseases. In their article, Kramer et al. provide a detailed review of the available literature on reversible redox post-translational modifications and mitochondrial and skeletal muscle function. They then provide critical review on current approaches to assess reversible redox post-translational modifications (Kramer et al.).

Several studies have implicated altered kinetic properties of the adenine nucleotide translocator (ANT) in the agingrelated impairment in mitochondrial energetics in skeletal muscle cells (Yan and Sohal, 1998; Gouspillou et al., 2014b). In the present research topic, Diolez et al. formulate the interesting hypothesis that these alterations in ANT could represent a protective mechanism to limit ROS production in aged muscle mitochondria while moderately disrupting mitochondrial energetics. Considering the importance of ROS as therapeutic targets, this hypothetical mechanism deserves further study.

The present research topic also provides readers with fundamental advancement in our understanding of the regulation of mitochondrial function in skeletal muscle cells. Indeed, in an elegant study, Lark et al. provide evidence that Protein Kinase A (PKA) can regulate mitochondrial energetics and H2O2 emission. Using PKA inhibitors and various mitochondrial substrates, they show that this regulation occurs at the level of Complex I. Finally, they provide new insights on how mitochondrial cyclic adenosine monophosphate (cAMP) production, cAMP being a positive regulator of PKA, is regulated (Lark et al.).

Understanding how nutrition modulates mitochondrial biology in muscle cells is of tremendous importance in the field of medicine. For instance, mitochondrial dysfunction has been suggested to be causally involved in obesity-induced insulin resistance and in the pathophysiology of type II diabetes (Goodpaster, 2013). Precisely defining how skeletal muscle mitochondria respond to obesogenic diet feeding is therefore of critical importance. In the present research topic, Putti et al. provide readers with a mini-review focused on the impacts of different dietary fat sources on mitochondrial bioenergetics, morphology and dynamics in skeletal muscle cells in the context of insulin-resistance. They also highlight the pressing need for mechanistic studies to confirm in vivo the relationship between mitochondrial morphology and dynamics and the development of insulin-resistance (Putti et al.).

Besides being of particular interest for the field of medicine, defining the impact of nutrition on mitochondrial biology is also an important research topic in the field of exercise physiology. The present research topic features two important review articles in this field. The first one, written by Craig et al. critically reviews the available literature on the utilization of small nutrients, such as caffeine, green tea extracts, polyphenols and amino-acids to enhance the impact of exercise training on mitochondrial biogenesis. They also provide recommendations and guidance for future studies that are required to explore the efficacy of these nutrients in humans, as well as the exercise setting in which they may prove beneficial (Craig et al.). The second review, written by Affourtit et al., first provides a state-of-the-art review of the available literature on the beneficial effects of dietary nitrate on human performance. Affourtit et al. then critically review the available experimental data in relation to the underlying mechanisms, with a particular emphasis on the impact of nitrate on mitochondrial bioenergetics.

It is now well established that low birth weight is associated with an increase in the risk of developing disease in later life such as coronary heart disease, diabetes, hypertension and stroke (see de Boo and Harding, 2006; Gluckman et al., 2008 for detailed reviews). Maternal undernutrition ranks as one of the most important causes that can lead to low birth weight (de Boo and Harding, 2006). In the present research topic, Beauchamp and Harper review the available evidence indicating that in utero undernutrition results in a metabolic reprogramming in both cardiac and skeletal muscles, characterized for instance by a reduction in mitochondrial content and respiration in the offspring. They also provide insights into the underlying mechanisms and provide a rationale linking the metabolic alterations resulting from in utero undernutrition to the increased risk of developing metabolic diseases (Beauchamp and Harper).

As mentioned earlier, mitochondrial dysfunction has been implicated in many conditions associated with muscle atrophy, including aging, cancer cachexia, and disuse-induced muscle atrophy (Dirks and Leeuwenburgh, 2004; Short et al., 2005; Chabi et al., 2008; Gouspillou et al., 2010, 2014a,b; Picard et al., 2010, 2015; Min et al., 2011; Hepple, 2014; Argilés et al., 2015). While the field to date has mainly focused on mitochondrial function (i.e., energetics, ROS production and mitochondrial mediated apoptosis), a growing interest now surrounds mechanisms involved in mitochondrial quality control (i.e., mechanisms responsible for the degradation of damaged / dysfunctional mitochondria). In the present research topic, Romanello and Sandri provide readers with a thorough and critical review on the current knowledge linking mitochondrial function, dynamics and quality control in the regulation of muscle mass. They also highlight several research avenues and challenges that will undoubtedly stimulate the field (Romanello and Sandri).

In the present research topic, Ryan et al. contributed a very interesting review paper focused on skeletal muscle and endothelial cell mitochondria in the setting of Critical Limb Ischemia. The latter is the most severe clinical presentation of peripheral arterial disease and manifests as chronic ischaemic "rest pain" and/or ischaemic skin lesions (Minar, 2009). Unfortunately, there is no effective way to treat the muscle myopathies caused by this disease. In their review, Ryan et al. first highlight the importance of skeletal muscle in the manifestation of Critical Limb Ischemia, and then provide a strong and exciting rationale for a role for endothelial cell and skeletal muscle mitochondria in the patient outcomes. As such, they propose that limb muscle and endothelial cell mitochondria should be considered as targets for novel therapeutic intervention (Ryan et al.).

Skeletal muscles have an impressive capacity to adapt to mechanical and physiological challenges by changing their phenotype in terms of size, fiber type, capillarization levels and aerobic capacity. Although skeletal muscle plasticity has been the focus of intense research effort, the molecular mechanisms underlying muscle plasticity are still incompletely understood.

#### REFERENCES

Since the seminal paper published by Spiegleman's group in 2002 (Lin et al., 2002), in which PGC-1α over-expression was shown to result in an increase in the proportion of slow-oxidative fibers, PGC-1α has attracted sustained attention. In the present research topic, Kupr and Handschin discuss recent advances in our understanding of how PGC-1α regulates skeletal muscle cell plasticity in health and disease. They also highlight further avenues of research to fully decrypt how PGC-1α influences muscle plasticity in health and disease (Kupr and Handschin).

Although often overlooked, our endocrine system exerts an important control on the regulation of mitochondrial mass and function in skeletal muscle cells, especially through thyroid hormones (Salvatore et al., 2014). Amongst thyroid hormones exerting control on mitochondrial biology in skeletal muscle cells, 3,5,3′ -Triiodo-L-thyronine (T3) has been extensively studied (Salvatore et al., 2014). In the present research topic, Lumbardi et al. review the experimental evidence indicating that 3,5-diiodo-L-thyronine (T2), and emerging iodothyronines also impact mitochondrial metabolism in skeletal muscle cells (Lombardi et al.). These novel aspects of thyroid physiology reviewed in Lombardi et al. open new perspectives for understanding the involvement of skeletal muscle mitochondria in systemic consequences of hypo- and hyper-thyroidism.

Because of the diversity and the quality of the articles compiled herein, we feel the present research topic was a success. We hope that it will not only provide readers with new insights and viewpoints on the role played by mitochondria in skeletal muscle health, aging and diseases, but will also serve as a platform to stimulate new ideas, experiments and research projects for further advances in the field.

#### AUTHOR CONTRIBUTIONS

GG and RH co-wrote and approved this Editorial.

#### ACKNOWLEDGMENTS

We thank all authors, reviewers and Frontiers Editorial Staffs for their precious contributions to this Research Topic.

**Conflict of Interest Statement:** The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

Copyright © 2016 Gouspillou and Hepple. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) or licensor are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.

# Mitochondrial NAD(P)H In vivo: Identifying Natural Indicators of Oxidative Phosphorylation in the 31P Magnetic Resonance Spectrum

Kevin E. Conley 1, 2, 3 \*, Amir S. Ali 1 , Brandon Flores 1 , Sharon A. Jubrias 1 and Eric G. Shankland1

*1 Department of Radiology, University of Washington Medical Center, Seattle, WA, USA, 2 Department of Physiology and Biophysics, University of Washington Medical Center, Seattle, WA, USA, 3 Department of Bioengineering, University of Washington Medical Center, Seattle, WA, USA*

Natural indicators provide intrinsic probes of metabolism, biogenesis and oxidative protection. Nicotinamide adenine dinucleotide metabolites (NAD(P)) are one class of indicators that have roles as co-factors in oxidative phosphorylation, glycolysis, and anti-oxidant protection, as well as signaling in the mitochondrial biogenesis pathway. These many roles are made possible by the distinct redox states (NAD(P)+ and NAD(P)H), which are compartmentalized between cytosol and mitochondria. Here we provide evidence for detection of NAD(P)+ and NAD(P)H in separate mitochondrial and cytosol pools *in vivo* in human tissue by phosphorus magnetic resonance spectroscopy (31P MRS). These NAD(P) pools are identified by chemical standards (NAD+, NADP+, and NADH) and by physiological tests. A unique resonance reflecting mitochondrial NAD(P)H is revealed by the changes elicited by elevation of mitochondrial oxidation. The decline of NAD(P)H with oxidation is matched by a stoichiometric rise in the NAD(P)+ peak. This unique resonance also provides a measure of the improvement in mitochondrial oxidation that parallels the greater phosphorylation found after exercise training in these elderly subjects. The implication is that the dynamics of the mitochondrial NAD(P)H peak provides an intrinsic probe of the reversal of mitochondrial dysfunction in elderly muscle. Thus, non-invasive detection of NAD(P)+ and NAD(P)H in cytosol vs. mitochondria yields natural indicators of redox compartmentalization and sensitive intrinsic probes of the improvement of mitochondrial function with an intervention in human tissues *in vivo*. These natural indicators hold the promise of providing mechanistic insight into metabolism and mitochondrial function *in vivo* in a range of tissues in health, disease and with treatment.

Keywords: magnetic resonance spectroscopy, 31P MRS, nicotinamide adenine dinucleotide, NAD+, NADP+, muscle, exercise

# INTRODUCTION

Nicotinamide adenine dinucleotide is a co-enzyme that is integral to cell and mitochondrial metabolism (Williamson et al., 1967; Stubbs et al., 1972; Nicholls and Ferguson, 2002; Ying, 2008; White and Schenk, 2012) and has a signaling role in mitochondrial biogenesis (Imai, 2011; Canto and Auwerx, 2012). NADH is a key player in oxidative phosphorylation in mitochondria

Edited by:

*Russell T. Hepple, McGill University, Canada*

#### Reviewed by:

*Douglas Root, University of North Texas, USA Giorgos K. Sakkas, University of St. Mark & St. John, UK*

> \*Correspondence: *Kevin E. Conley<EMAIL_ADDRESS>
#### Specialty section:

*This article was submitted to Striated Muscle Physiology, a section of the journal Frontiers in Physiology*

Received: *29 October 2015* Accepted: *01 February 2016* Published: *30 March 2016*

#### Citation:

*Conley KE, Ali AS, Flores B, Jubrias SA and Shankland EG (2016) Mitochondrial NAD(P)H In vivo: Identifying Natural Indicators of Oxidative Phosphorylation in the 31P Magnetic Resonance Spectrum Front. Physiol. 7:45. doi: 10.3389/fphys.2016.00045* (Nicholls and Ferguson, 2002) and in glycolysis (Williamson et al., 1967; Stubbs et al., 1972), while the related metabolite, NADP, is important in anti-oxidant protection (Ying, 2008). Optical methods detect the sum of these compounds (NAD+ and NADP+ = NAD(P)+) and can resolve their cellular location to provide natural indicators of mitochondrial function and cytosolic redox state (NAD(P)H/NAD(P)+) (Jobsis and Duffield, 1967; Barlow and Chance, 1976; Wendt and Chapman, 1976; Scholz et al., 1995; Mayevsky and Rogatsky, 2007; Gandra et al., 2012; Claflin et al., 2015). These measures have lead to insight into the role of NAD(P) redox in oxidative stress (Ying, 2008; Murphy, 2009; Aon et al., 2010; Massudi et al., 2012a), as an index of oxygenation states (Barlow and Chance, 1976; Mayevsky and Rogatsky, 2007), and as a probe of mitochondrial (dys)function in age and disease (Heikal, 2010; Massudi et al., 2012b; Claflin et al., 2015). However, optical measures are typically limited to isolated tissues and are limited in depth penetration in vivo. A non-invasive measure of NAD(P) in vivo would open a window on cell and mitochondrial metabolism in age and disease as well as the impact of treatments to reverse metabolic dysfunction.

Non-invasive magnetic resonance methods now make detecting NAD(P)+ and NAD(P)H in vivo possible both in animals and humans (de Graaf and Behar, 2014; Zhu et al., 2015). A distinct spectral position (chemical shift, CS) in the phosphorus spectrum (31P MRS) is found for each metabolite representing the NAD(P)+ and NAD(P)H redox states (Lu et al., 2014). It may also be possible to distinguish the cellular location of these NAD(P) redox states since several metabolites are reported to show distinct CS in the 31P MR spectrum between these two compartments (Garlick et al., 1983; Hutson et al., 1989, 1992). For example, both ATP and NADP+ show striking differences in CS with Mg2+ binding at the low metal concentration in the cytosol vs. the 10-fold higher level found in mitochondria (Mas and Colman, 1984; Gout et al., 2014). Inorganic phosphate also shows a significant chemical shift in vivo as a result of the pH difference in cytosol vs. mitochondria (Garlick et al., 1983; Hutson et al., 1992; Kan et al., 2010). There are also promising studies suggesting that NAD(P) redox states can be distinguished between the mitochondria vs. the cytosol. These studies found that phosphoryl compounds related to NAD(P) showed CS differences between erythrocytes vs. the bathing solution that are large enough to be detected by 31P MRS (Kirk and Kuchel, 1988a,c). Thus, it may be possible to detect a NAD(P)H CS difference inside mitochondria relative to the cytosol in the 31P MR spectrum thereby providing a natural indicator of key player in oxidative phosphorylation.

Here we describe a unique resonance in the 31P MRS spectrum that conforms with the properties of NAD(P)H in mitochondria in human muscle in vivo. The aim of this study was to test the nature of this new resonance and determine if it provides a natural indicator of mitochondrial oxidative function. The first test compared this unique resonance to chemical standards of metabolites known to be present in the same region of the 31P MR spectrum, including NAD+, NADP+, NADH, and NADPH (Kushmerick et al., 1986). No overlap in the spectral position (chemical shift) of the unique resonance vs. that of known metabolites was found. Two additional tests were performed to determine whether dynamics elicited in this peak by exercise conformed with changes expected for mitochondrial NAD(P)H. This functional test was designed to elevate oxidative phosphorylation and involved 40 elderly subjects before and after exercise training. The pre-training test revealed a reciprocal decline in the unique resonance vs. elevation in NAD(P)+, which are changes consistent with oxidation of mitochondrial NAD(P)H to NAD(P)+. The posttraining test found a greater change in the unique resonance after exercise training, which is consistent with elevated oxidation and the greater phosphorylation rate reported in these subjects. Taken together, these results provide evidence that this MRS detectable resonance conforms to the properties of mitochondrial NAD(P)H. Thus, this unique resonance holds promise as the first natural indicator of the inner workings of oxidative phosphorylation in the 31P MRS spectrum. It also holds promise as a sensitive indicator of the mitochondrial response to treatments designed to improve mitochondrial function.

# METHODS

### Subjects

An elderly group consisted of 40 subjects (18 male, 22 female) ranging in age from 65 to 80 years (68.8 ± 5.9 years, means ± S.E.M.). Subjects were not involved in formal exercise training, were in good health and had no significant cardiac, neurological, or musculoskeletal disease, as we have described (Conley et al., 2000; Jubrias et al., 2001). All subjects voluntarily gave informed, written consent as approved by the University of Washington Human Subjects Review Committee and in accordance with the Declaration of Helsinki.

#### MR Methods

A General Electric 1.5 T Signa imager/spectrometer was used for the in vivo spectra as described (Conley et al., 2000). A 9 cm diameter surface coil tuned to the phosphorus frequency (25·9 MHz) was placed over the vastus lateralis muscle of the thigh. The B1 field homogeneity was optimized by off resonance proton shimming on the muscle water peak. The unfiltered PCr linewidth (full width at half-maximal height) was typically 4–8. Each subject had a high resolution 31P MR spectrum of the resting vastus lateralis muscle taken under conditions of fully relaxed nuclear spins (16 free-induction decays (FID) with a 16 s interpulse delay) using a spectral width of ±1250 Hz and 2048 data points. Since volume selective methods were not used in collection, the spectra represent signal from a hemispherical volume defined by the coil radius. No volume selective methods were used to collect the spectra. Dynamic changes during stimulation and recovery were made using a standard 1-pulse experiment with partially saturated nuclear spins (1.5 s interpulse delay). Artifacts due to movement were reduced by stabilizing the limb during the muscle contractions. All fully relaxed spectra were zero filled from 2048 to 4096 points, Fourier transformed with 15 Hz apodization, baseline corrected, and manually phased using the Mnova software package (Mestrelab Research, Santiago de Compostela (Spain)). No other spectral treatment was employed.

# Chemical Phantoms

Metabolites that resonate in the NAD(P) region of the 31P MR spectrum were identified from muscle extracts (Kushmerick et al., 1986). Solutions containing these chemicals were prepared using binding constants taken from the USA National Institute of Standards and Technology (NIST) Critically Selected Stability Constants of Metal Complexes Database (see Kushmerick, 1997). All solutions contained (in mmol/L): EGTA 15, MOPS 80, free Mg2+ 1, Na+ 83, and K+ 52. The following were varied in individual solutions: ATP, ADP, Pi , Creatine Phosphate (PCr), ADP, NAD+, NADH, NADP+, or UDP-Glucose. The ionic strength was maintained at 0.175 M, pH = 7.0 (36◦C) in all solutions. High-resolution MR spectra of individual solutions in an NMR tube were taken at 4.7T (sweep width = 10,000 Hz, 16 K complex points, 128 FIDs, 5 s delay between pulses).

# Peak Simulations

Resonances were simulated at 1.5T using the spin parameters and chemical shifts for the simulations determined from NAD+, NADP+, and NADH in physiological solution at 4.7T. The Spin Simulation feature of the Mnova software (Mestrelab Research, Santiago de Compostela (Spain)) permitted determining the spectral properties of these resonances. The line-width of the simulated resonances were fixed to that of α-ATP due to the similar chemical environment of the two phosphate groups in NAD(P). The details of this process have been described (Lu et al., 2014).

### NAD(P) Region In vivo

The −10.4 to −11.3 region containing NAD+ and NADP+ was extracted from the fully relaxed spectra by fitting a Lorenzian line shape to the α–ATP peak (**Figure 1**). The spectral intensity of this region was integrated as well as three sub-regions: −10.5 to −10.68 ppm (NADH), −10.69 to −10.92 (NAD(P)+) and −10.93 to −11.30 (hereafter, −11.05 ppm). The dynamic changes in these regions were determined by subtraction spectra normalized to the α–ATP level between stimulated vs. resting muscle and during recovery vs. resting muscle.

#### Stimulation and Recovery Protocol

The quadriceps muscles were activated by transcutaneous electrical stimulation of the femoral nerve, as previously described (Blei et al., 1993; Conley et al., 2000). Spectra were collected during rest, stimulation and recovery to measure the PCr, Pi , ATP, and the NAD(P) region peaks. Spectra were averaged during each of the three periods to analyze the NAD(P) region peaks:

Control period (60 s, 10 spectra): Baseline data were obtained during resting muscle conditions to establish initial metabolite peak levels.

Stimulation period (120 s, 20 spectra): A 3 Hz electrical stimulation period was used to decrease [PCr] and activate NAD(P)H synthesis.

Aerobic recovery (300 s, 50 spectra): upon cessation of stimulation, PCr recovery was followed until restoration of baseline levels during which NAD(P)H oxidation was determined.

# Statistics

Pre- vs. post-training differences relative to zero were tested with a 2-tailed Student's t-test. No adjustment for multiple comparisons was made as per reference (Perneger, 1998). Statistical differences are reported at P < 0.05. Means are reported ±SEM.

# RESULTS

Here we show evidence for a unique resonance in the 31P spectrum corresponding to NAD(P)H in mitochondria. To identify this resonance, we first compare the unique resonance against known metabolites in muscle. Second, we test the link between the unique resonance vs. the change in mitochondrial content that came with exercise training in these subjects. Third, we use an acute exercise bout to activate NAD(P)H oxidation to test for changes in the resonance and determine whether these changes (oxidation) parallel the improvement in phosphorylation found after exercise training in these subjects.

# Unique Resonance in 31P Spectrum In vivo

The presence of an unidentified peak in the 31P MR spectrum of human muscle is illustrated in **Figure 1**. An expansion of the spectrum in the α–ATP region shows the NAD(P) peak region from −10.4 to −11.3 ppm. This spectrum was taken from an elderly vastus lateralis muscle under MR conditions in which peak areas reflect their chemical content (i.e., fully relaxed conditions). The resonances for chemical standards of NADH (−10.6 ppm), NAD+ (−10.75 ppm), and NADP+ (−10.84 ppm) are superimposed on the NAD region from the in vivo spectrum in Panel B [UDP-glucose is also present (Kushmerick et al., 1986) and resonates in this region at −10.84 in in vitro solutions that emulate in vivo conditions]. This comparison highlights the presence of an additional resonance centered at −11.05 ppm in the NAD(P) region. Thus, the unique resonance at −11.05 ppm does not correspond with resonances for metabolites in this spectral region that have been identified in tissue extracts (Kushmerick et al., 1986).

# Unique Resonance is 40% of Total NAD(P) Integral

The NAD(P) integral is 11% of the area of α–ATP (11.5 ± 1.0%), which agrees with the total NAD per ATP content (7– 12%; Sahlin, 1983; Henriksson et al., 1986) reported from biochemical analyses in human vastus lateralis muscle (Sahlin, 1983; Henriksson et al., 1986; Ren et al., 1988). Based on the ATP level in these muscles (5.8 mM; Conley et al., 2000), the full integral (0.7 mM) contains 0.4 mM NAD(P) (60% of the integral) vs. 0.5 mM measured for total NAD in young muscle (Henriksson et al., 1986). The unique resonance at −11.05 ppm is the second largest peak in the NAD(P) integral (**Figure 1C**), which translates to 0.25 mM (38% of the total). This is close to the value (0.26 mM) estimated from the NADH content of muscle mitochondria (heart, 10 nmole/mg mitochondrial protein, Alano et al., 2007) and the mitochondrial volume density of the elderly muscle in this study [3% (Conley et al., 2000), i.e., 10 nmole/mg × mg

protein/µl (Vinnakota and Bassingthwaighte, 2004) × 0.03µl mito/µl muscle = 0.26 nmole/µl or 0.26 mM]. Thus, two key segments of the NAD(P) region in vivo are consistent in size with the contents of total NAD+ assayed in muscle and NADH

Content of the three resonances *in vivo* [NAD(P)] per ATP content, mean % ± SEM, *n* = 39 (Conley et al., 2000).

estimated from mitochondrial volume density of human muscle. In contrast, the small resonance at −10.6 ppm associated with NADH in solution was 5% of the total integral in the NAD(P) region or ∼0.01 mM. A low NADH content in the cell is expected from the oxidized NAD redox state maintained by enzyme equilibria linked to the phosphorylation potential (Williamson et al., 1967; Stubbs et al., 1972). This low value agrees with the NADH levels in older bioluminescent assays on human vastus lateralis tissue (0.02 mM at rest, Henriksson et al., 1986). These findings suggest that the chemical standard for NADH at −10.6 ppm identifies the resonance for the small cellular NADH pool, while the larger −11.05 ppm resonance is more consistent with the mitochondrial NAD(P)H pool. Thus, the metabolite content represented by the −10.4 ppm to −11.3 ppm integral, which includes the unique resonance at −11.05 ppm, is in reasonable agreement with the total NAD(P) content reported from biochemical analysis of vastus lateralis muscle tissue.

# Increase in −11.05 ppm Peak and Mitochondrial Volume with Exercise Training

Our second test of the nature of the unique resonance took advantage of the increased mitochondrial content that accompanied a 6-mo exercise training program in these subjects (Jubrias et al., 2001). We tested whether the −11.05 ppm resonance increased in proportion to the elevation in mitochondrial volume density found with resistance training (RT) in these subjects (131%). **Figure 2** shows the significant changes in the −11.05 ppm resonance with RT in elderly subjects (139%), while no change was found on average in either property in control (CN) or endurance trained (ET) groups. Also, no change was found in the NADH peak area at −10.60 ppm in the RT group (0.1 ± 0.5% ATP). Thus, the adaptation in the −11.05 ppm peak was exclusively associated with that of mitochondrial volume density with exercise training.

# Reciprocal Change in −11.05 ppm Peak vs. NAD(P)+ with Elevated Oxidation

A third test involved an exercise and recovery bout designed to alter the NAD(P) resonances by elevating oxidative phosphorylation, as shown in **Figure 3**.

**Figure 3A** shows the changes in creatine phosphate (PCr), a natural indicator of cell ATP use and re-synthesis, during an exercise and recovery bout. Exercise consisted of a 2 min muscle stimulation period that increased ATP use resulting in depletion of PCr level from the resting state (red line). The curvilinear shape of the PCr breakdown curve indicates that activation of ATP synthesis by mitochondria reduces the net breakdown toward the end of the stimulation period. Once stimulation ends and contractile ATP demand ceases, oxidative phosphorylation predominates (blue line) and PCr is resynthesized to quickly return to resting levels.

**Figure 3B** shows the 31P spectra of the NAD(P) region taken from 10 subjects during these two periods. The spectrum collected from resting muscle (black line) represents the low oxidative phosphorylation condition. The spectrum collected from muscle during stimulation is shown in by red lines (left hand panels) and that collected during the recovery period of elevated oxidative phosphorylation is shown by the blue lines

(right hand panels). The solid arrows show the reciprocal changes in the −10.75 ppm and in the −11.05 ppm region.

**Figure 3C** shows the net differences in these regions with subtraction of these spectra. The change in the −10.75 ppm region is outlined by several dashed peaks. These peaks represent the resonances of the following chemical metabolites in solution: NADH (−10.6 ppm), NAD+ (−10.75 ppm), and NADP+ (10.83 ppm). Note that the decline in the −10.75 ppm region with stimulation (red arrows) is matched by an equal rise in the −11.05 ppm region. The opposite changes are seen during recovery from stimulation (blue line): the −11.75 ppm region rises and the −11.05 ppm region declines.

**Figure 3D** shows the integration of the peak areas in the difference spectra. Equal and opposite change in the −10.75 vs. −11.05 ppm region is confirmed by no significant difference in a paired t-test of the absolute integrated areas of the two regions in stimulation or recovery (P > 0.15 each). Such reciprocal changes are expected for a trade-off of oxidized and reduced forms of NAD(P)+. The correspondence of the dashed peaks representing the resonances for NAD+ and NADP+ and the change in the −10.75 ppm region is strong support that this region represents net changes in NAD(P)+. Similarly, the reciprocal change in the −11.05 ppm region suggests that change in NAD(P)H occurs in the region of the unique resonance. These changes are consistent with the dynamics expected for reduction and oxidation of mitochondrial NAD(P)H. Taken together, these results point to mitochondrial NAD(P)H dynamics occurring at the downfield spectral position (−11.05 ppm) from the resonance defined by the chemical NADH (−10.6 ppm) that likely defines the cell NADH.

# Exercise Training Accelerates NAD(P) Dynamics and Phosphorylation

Our final test was whether the −11.06 ppm peak changes with an independent measure of oxidative phosphorylation. These subjects showed increased phosphorylation capacity after endurance (ET) and resistance (RT) training (Jubrias et al., 2001). This capacity was measured during the stimulation-recovery experiment shown in **Figure 3**, which was undertaken in each subject and repeated after exercise training. **Table 1** shows the changes in the −11.05 ppm peak in the three groups in this training experiment and reveals that both ET and RT groups show a net decline in the −11.05 ppm peak after training. No change is apparent in the control group. A corresponding net rise that balances the decline in the −11.05 ppm peak area is apparent in the −10.6 ppm peak (ET) or the −11.75 ppm peak (RT) with training. This rise in the integrated peaks in the −10.5 to −10.9 ppm region vs. the −11.05 ppm region demonstrates a net stoichiometry in the changes of the NAD(P) peaks with training. This net decline in the −11.05 ppm region during elevated oxidation following training further supports this spectral region as representing the mitochondrial metabolites, NAD(P)H.

**Figure 4** shows that the significant decline in the −11.05 ppm region (−10.9 to −11.3 ppm) mirrored the net increase in phosphorylation with training in these subjects. These reciprocal changes suggest that the 1-11.05 ppm region reflects the greater oxidation of NAD(P)H that is expected with the elevated phosphorylation (1PCr) found after training. Together, these results support that mitochondrial NADH oxidation in vivo is represented by the decline in the unique resonance at −11.05 ppm in the NADP region. Further, these results indicate that the −11.05 ppm peak is a sensitive indicator of the impact of an intervention on improving mitochondrial oxidation in elderly muscle.

## DISCUSSION

The metabolic role and redox state of the NAD(P) metabolites depend on their cellular location as illustrated in **Figure 5**. NADH is a critical substrate for oxidative phosphorylation and predominates over NAD+ in mitochondria. The opposite is true for the oxidized cytosol in which enzyme equilibria linked to the phosphorylation potential holds NAD(P)H content low and NAD(P)+ high (Williamson et al., 1967; Stubbs et al., 1972). Here we describe a unique resonance at −11.05 ppm in the in vivo 31P MR spectrum that conforms with the properties of mitochondrial NAD(P)H. This unique resonance has the high chemical content and the dynamic changes with oxidative phosphorylation that are expected for the NAD(P) redox state in mitochondria. In contrast, a second peak identified by an NADH chemical standard (at −10.6 ppm) is consistent with the low metabolite content expected for cytosolic NADH. Three results summarize the evidence that the −11.05 ppm peak is a natural indicator of the content and dynamics of mitochondrial NAD(P)H and thereby provides a intrinsic probe of oxidative phosphorylation in vivo:

TABLE 1 | Change in peak areas during the exercise recovery period in post- vs. pre-trained muscle.

resistance (RT) and endurance (ET) training vs. controls (CN). \**P* < 0.05.

*Units are % NAD per* α*–ATP peak for three resonances at distinct chemical shifts (ppm) in the NADP region. Values are means* ± *SE.* 1 *defines post- vs. pre-training changes in the integral of area of each resonance per integral of the* α*–ATP resonance.*

*The groups are: CN, control; RT, resistance training; ET, endurance training groups.* 1−*10.6,* 1−*10.75, and* 1−*11.05 are the chemical shifts (ppm) for the resonances defined by external standards for the reduced (NADH) and oxidized (NAD(P)*+*) forms and a third resonance not identified by external standards, respectively.* \**Bold font, P* < *0.05 post- vs. pre-training.*

#### NAD(P) In vivo vs. In vitro

Several lines of evidence indicate good agreement of the NAD(P) metabolite levels determined in the 31P MR spectrum in vivo and those reported in isolated human muscle by in vitro methods. The metabolite content represented by the peaks in the NAD(P) region (−10.5 to −11.3 ppm; 11% NAD/ATP) is in agreement with biochemical assays of NAD in human vastus lateralis (Sahlin, 1983; Henriksson et al., 1986). Chemical identification of the peaks in this region reveals that NAD(P)+ accounts for 60% of the total peak area while only 5% of the total is accounted

for by the chemically defined NADH (−10.6 ppm). The chemical content of the NAD(P)+ region (0.4 mM) is consistent with estimates for NAD+ derived from biochemical assays in young muscle (0.5 mM). The remaining 38% of the peak area of the NAD(P) region is represented by a peak centered at −11.05 ppm. Both the MR determination and that derived from independent measures estimate a NAD(P)H content of ∼0.26 mM.

## Two Resonances Reflect NAD(P)H In vivo

A large difference in peak areas is expected from in vitro results if cytosolic vs. mitochondrial NAD(P)H can be distinguished in the 31P MR spectrum (Williamson et al., 1967; Stubbs et al., 1972). The chemical content represented by the −10.60 ppm peak was low (5% of NAD(P) integral) relative to the unique resonance at −11.05 ppm (38%) as expected for cytosolic NADH, which is maintained at a low level by enzyme equilibria linked to the phosphorylation potential (Williamson et al., 1967; Stubbs et al., 1972). The larger chemical content of the −11.05 ppm corresponds with the predominance of mitochondrial NAD(P)H pool. This agreement of the size of two 31P MR resonances with the expected NAD(P)H contents provides evidence that the cytosolic vs. mitochondrial NADH pools can be distinguished by MRS in vivo (Krebs and Veech, 1969; Stubbs et al., 1972).

### Compartmentalization of NAD(P)H

The basis of the distinct spectral positions of NAD(P) is likely due to the changes in physical properties when the metabolite is located inside a compartment, such as in mitochondria. For example, enhanced fluorescence efficiency occurs with NAD(P)H inside mitochondria due to binding to complex I of the electron transport chain (Blinova et al., 2008), which permits highlighting mitochondrial NAD(P)H by optical spectroscopy and imaging. A second example is the different chemical shift in the 31P MR spectrum seen with phosphoryl compounds located inside erythrocytes (Kirk and Kuchel, 1988a,c; Larkin et al., 2007). Compounds related to NAD(P) show a 1−0.5 ppm CS in the red blood cell, which is similar to the shift of the −11.05 ppm vs. −10.6 ppm NAD(P)H peak in vivo in muscle (Kirk and Kuchel, 1988a,c; Larkin et al., 2007). The mechanism for this downfield CS in red cells is reported to be elevated hydrogen bonding resulting from the high protein concentration of the erythrocyte (predominantly hemoglobin content) (Kirk and Kuchel, 1988b; Larkin et al., 2007). A high protein content is also characteristic of mitochondria (Vinnakota and Bassingthwaighte, 2004). Thus, the segregation of phosphoryl compounds inside a high protein environment provides a physical mechanism for the downfield CS of the −11.05 ppm peak (reflecting NAD(P)H in mitochondria) relative to NAD(P)H in the cell (−10.6 ppm) in vivo.

# Increase in −11.05 ppm Peak and Mitochondrial Content with Exercise Training

A test of the identity of the −11.05 ppm peak is provided by the changes in the peaks of the NAD(P) region and in mitochondrial properties with exercise training (**Figure 2**). A shift in NAD(P) metabolite levels in correspondence with changes in mitochondria have been reported in several tissues with calorie restriction in mice in in vitro assays (Chen et al., 2008). One mechanism for an NAD+ increase is a rise in the enzyme responsible for NAD+ biosynthesis—NAMPT—that accompanies calorie restriction and exercise training (Costford et al., 2009). These adaptations in NADH and mitochondria were not uniform across tissues in calorie restricted mice (Chen et al., 2008). They also differed between training modes in the human muscle (**Figure 2**). A higher −11.05 ppm peak (139%) was found in proportion to the greater mitochondrial content with RT (131%), but neither property changed significantly with ET (**Figure 2**). In contrast to the −11.05 ppm peak, the chemically defined NADH peak at −10.6 ppm did not change despite the increased mitochondrial content in RT. This stability of the peak at −10.6 ppm in the face of a large increase in mitochondrial content is consistent with this resonance representing NAD(P)H in the cytosol. In contrast, a mitochondrial origin for the −11.05 ppm peak is supported by the unique rise of this resonance with mitochondrial volume density after exercise training.

# Activating NAD(P)H Generation and Oxidation

A functional test of the identity of the −11.05 ppm peak is provided by the dynamics of the peaks in the NADP region with exercise and recovery. These dynamics are elicited by a protocol that activates increased energy use (as shown by PCr depletion from the resting state) followed by a period of elevated oxidative phosphorylation after exercise (as shown by restoration of PCr, **Figure 3A**). The peaks in the NAD region change in concert with these PCr dynamics during exercise and recovery (**Figure 3B**) to reveal the trade-offs in the reduced vs. oxidized redox states of NAD(P) (**Figure 3C**) (Jobsis and Duffield, 1967; Wendt and Chapman, 1976). Such trade-offs are evident from the changes in the −11.05 ppm peak, which are matched by equal and opposite changes in oxidized NAD(P) (**Figure 3D**). Similar changes are apparent in optical fluorescence studies that indicate reduced mitochondrial NAD(P)H with muscle stimulation and oxidized NAD(P)H during elevated oxidative phosphorylation in recovery (Jobsis and Duffield, 1967; Wendt and Chapman, 1976; Gandra et al., 2012; Claflin et al., 2015). A rise in NAD(P)H generation under intense exercise has also been found in glycolytic type II fibers of human muscle in biochemical studies (Ren et al., 1988), while reduced NADH is found during sustained exercise in the oxidative, type I fibers of humans (Ren et al., 1988). Thus, the functional dynamics of the −11.05 ppm peak parallel the changes in mitochondrial NAD(P)H during exercise cycles reported in independent studies. These reciprocal changes in spectral regions are consistent with the −11.05 ppm peak representing mitochondrial NAD(P)H.

#### Sensitive Index of Mitochondrial Oxidative Improvements

An in vivo measure of mitochondrial NAD(P)H holds the promise of providing a natural indicator of oxidation adaptations with an intervention in human muscle. A test of the −11.05 ppm peak as such an indicator is provided by the results of exercise training that raised oxidative phosphorylation in these subjects (Jubrias et al., 2001). **Figure 4** shows a net drop in the −11.05 ppm resonance in an exercise bout after exercise training. This net decline parallels the net increase in phosphorylation rate and suggests that the −11.05 ppm peak reflects increased oxidation of mitochondrial NADH in the trained subjects. Independent findings of faster oxygen uptake after exercise training support these results. Human muscle O2 uptake during recovery from exercise is increased and the kinetics of whole-body oxygen uptake is more rapid at the onset of exercise after training in adult and elderly subjects (Zoladz et al., 2006; Murias et al., 2010). The alternative is a higher ATP flux without increased O2 uptake as a result of improvement in the coupling of generating ATP per O2, which also occurs with exercise training (Conley et al., 2013). A recent analysis concluded that both mechanisms are activated: greater electron transport chain flux and more efficient generation of ATP appeared to contribute nearly equally to the phosphorylation improvements in these subjects (Conley, 2016). The results in **Figure 4** support this independent analysis to demonstrate that elevated oxidation is a part of the improvement in mitochondrial capacity to generate ATP after training. These results indicate that the dynamics of the −11.05 ppm peak captures the mitochondrial oxidation improvements that underlie greater phosphorylation. Importantly, both the oxidation and phosphorylation measures come from the same 31P MR spectra collected in a single exercise test in human muscle in vivo.

#### Limitations and Future Directions

The low chemical content of NAD(P) related metabolites (<1 mM) in vivo is a limitation to detecting this natural indicator by 31P MRS. Measurements are possible at the low field strength of 1.5T because of the trade-off with high B1 field homogeneity at 1.5T, which provides good resolution of the individual NAD(P) peaks despite the low signal-to-noise. The large diameter coil (9 cm) used in this study and large muscle mass of the quadriceps group (∼1 kg) also helped to improve the signal for detecting these low concentration metabolites. Nonetheless, the high standard errors on some of the determinations may be due to this combination of low chemical content and low MRI field strength. However, two advances allow improved signal-to-noise over that found in the study. The first advance is the higher field strength (3T) MRIs that are now available in most medical centers that focus on human metabolic research. A second advance is the ability to decouple protons from phosphorus to increase the signal-to-noise of the NAD(P) peaks. Decoupling is a method derived from high-resolution MR spectroscopy that is now a standard feature of 3T MRIs with multi-nuclear detection (e.g., 31P MRS) capability. Thus, the availability of the higher 3T field strength for human studies and the ability to further enhance signal-to-noise with proton decoupling will make detecting of the NAD(P) peaks and their dynamic changes with an intervention possible for clinical research studies.

## SUMMARY

Here we show that a unique resonance in the in vivo 31P spectrum provides a natural indicator of mitochondrial oxidation and its improvement with treatment. This assignment comes from the known compartmentalization of NADH between the highly oxidized cell (−10.6 ppm) vs. more reduced mitochondria (−11.05 ppm). A functional test that followed NAD(P) dynamics during exercise and recovery cycles confirmed that the −11.05 ppm peak reflected oxidation and reduction of mitochondrial NAD(P)H. Finally, the greater −11.05 ppm peak changes in the exercise test paralleled the faster phosphorylation rate in these subjects after exercise training. This correspondence provides further support that this unique resonance is a natural indicator of mitochondrial oxidation. Thus, intrinsic probes are present in the 31P MR spectrum that reveal the NAD(P) redox disparity between mitochondria and cell and provide a natural indicator of oxidative phosphorylation that is sensitive to improvements in mitochondrial function with treatment.

### AUTHOR CONTRIBUTIONS

KC, SJ, and ES conceived and designed the study; all authors contributed to the collection, analysis, and interpretation of data; KC and ES were involved in drafting or revising the paper.

# FUNDING

This work was supported by NIH grants RC2AG036606, R01AR41928, R01AG10853, and a Nathan Shock Center Pilot Project award, as well as Seattle Children's Mitochondrial Guild and the Royalty Research Fund of the University of Washington.

#### ACKNOWLEDGMENTS

We thank Martin Brand and Martin Kushmerick for their insights. Michael Regnier provided physiological solutions that

#### REFERENCES

mimic the muscle cell in vivo. We also acknowledge the expert contributions of M. Elaine Cress, Elizabeth Egan, Peter Esselman, Barbara Inglin, Chris Mogadam, and Ib Odderson. Hans Hoppeler provided the micrograph in **Figure 5**.

**Conflict of Interest Statement:** The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

Copyright © 2016 Conley, Ali, Flores, Jubrias and Shankland. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) or licensor are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.

# Mitochondrial Bioenergetics and Fiber Type Assessments in Microbiopsy vs. Bergstrom Percutaneous Sampling of Human Skeletal Muscle

Meghan C. Hughes 1 , Sofhia V. Ramos 1 , Patrick C. Turnbull 1 , Ali Nejatbakhsh1 , Brittany L. Baechler 2 , Houman Tahmasebi 1 , Robert Laham1 , Brendon J. Gurd3 , Joe Quadrilatero2 , Daniel A. Kane4 and Christopher G. R. Perry 1 \*

1 Muscle Health Research Centre, School of Kinesiology and Health Science, York University, Toronto, ON, Canada, 2 Department of Kinesiology, University of Waterloo, Waterloo, ON, Canada, 3 School of Kinesiology and Health Studies, Queen's University, Kingston, ON, Canada, 4 Department of Human Kinetics, St. Francis Xavier University, Antigonish, NS, Canada

#### Edited by:

Gilles Gouspillou, Université du Québec à Montréal, Canada

#### Reviewed by:

Gordon Fisher, University of Alabama at Birmingham, USA Stuart D. R. Galloway, University of Stirling, UK

> \*Correspondence: Christopher G. R. Perry<EMAIL_ADDRESS>
#### Specialty section:

This article was submitted to Striated Muscle Physiology, a section of the journal Frontiers in Physiology

Received: 11 September 2015 Accepted: 16 November 2015 Published: 18 December 2015

#### Citation:

Hughes MC, Ramos SV, Turnbull PC, Nejatbakhsh A, Baechler BL, Tahmasebi H, Laham R, Gurd BJ, Quadrilatero J, Kane DA and Perry CGR (2015) Mitochondrial Bioenergetics and Fiber Type Assessments in Microbiopsy vs. Bergstrom Percutaneous Sampling of Human Skeletal Muscle. Front. Physiol. 6:360. doi: 10.3389/fphys.2015.00360 Microbiopsies of human skeletal muscle are increasingly adopted by physiologists for a variety of experimental assays given the reduced invasiveness of this procedure compared to the classic Bergstrom percutaneous biopsy technique. However, a recent report demonstrated lower mitochondrial respiration in saponin-permeabilized muscle fiber bundles (PmFB) prepared from microbiopsies vs. Bergstrom biopsies. We hypothesized that ADP-induced contraction (rigor) of smaller length microbiopsy PmFB causes a greater reduction in maximal respiration vs. Bergstrom, such that respiration could be increased by a myosin II ATPase-inhibitor (Blebbistatin; BLEB). Eleven males and females each received a 2 mm diameter percutaneous microbiopsy and a 5 mm diameter Bergstrom percutaneous biopsy in opposite legs. Glutamate/malate (5/0.5 mM)—supported respiration in microbiopsy PmFB was lower than Bergstrom at submaximal concentrations of ADP. 5µM BLEB reduced this impairment such that there were no differences relative to Bergstrom ± BLEB. Surprisingly, pyruvate (5 mM)-supported respiration was not different between either biopsy technique ±BLEB, whereas BLEB increased succinate-supported respiration in Bergstrom only. H2O2 emission was lower in microbiopsy PmFB compared to Bergstrom PmFB in the presence of BLEB. Microbiopsies contained fewer type I fibers (37 vs. 47%) and more type IIX fibers (20 vs. 8%) compared to Bergstrom possibly due to sampling site depth and/or longitudinal location. These findings suggest that smaller diameter percutaneous biopsies yield lower glutamate-supported mitochondrial respiratory kinetics which is increased by preventing ADP-induced rigor with myosin inhibition. Microbiopsies of human skeletal muscle can be utilized for assessing mitochondrial respiratory kinetics in PmFB when assay conditions are supplemented with BLEB, but fiber type differences with this method should be considered.

Keywords: mitochondria, respiration, permeabilized fiber, muscle biopsy, blebbistatin

# INTRODUCTION

The Bergstrom muscle biopsy technique has been an essential tool for the direct assessment of human skeletal muscle responses to a variety of physiological perturbations (Bergstrom, 1975). More recently, percutaneous microbiopsy needles have been adopted as an alternative approach to sample skeletal muscle in humans. Anecdotally, microbiopsies are often perceived as less invasive than Bergstrom given they obtain smaller samples and do not require an incision on the skin and fascia for the common procedure of sampling human vastus lateralis skeletal muscle. While these arguments have not been fully validated, interest in microbiopsies has increased nonetheless, particularly for the assessment of skeletal muscle metabolism (Hayot et al., 2005; Jorge et al., 2011; Krause et al., 2012; Votion et al., 2012). As such, validation of this methodology for metabolic assays in skeletal muscle is required given the potential influence of the sampling method on experimental results.

Permeabilized muscle fiber bundles (PmFB) are commonly used to assess mitochondrial bioenergetics in skeletal muscle. This technique requires careful separation of muscle fibers with fine forceps followed by chemical permeabilization with cholesterol-binding detergents (Kuznetsov et al., 2008; Pesta and Gnaiger, 2012; Perry et al., 2013). Surprisingly, it was recently reported that microbiopsies of pig skeletal muscle yielded lower mitochondrial respiration in PmFB which appeared to be positively correlated to needle diameter (Isner-Horobeti et al., 2014). While the exact cause of this lower respiration was not identified, it is possible that the smaller fiber length of PmFB from microbiopsies pose a challenge to preparing intact PmFB that can withstand the magnetic stirring during respirometric assessments. Loss of PmFB integrity could impair respiratory assessments given it has previously been established that preservation of mitochondrial structure and morphology are critical for optimizing respiratory assessments in PmFB (Veksler et al., 1987; Kuznetsov et al., 2008; Picard et al., 2011).

A key factor influencing PmFB intactness is a phenomenon of ADP-induced rigor, such that the very nature of assessing ADP-stimulated respiratory kinetics also induces contraction and disintegration in vitro (Ventura-Clapier and Vassort, 1985; Perry et al., 2011, 2013). Such PmFB contraction influences respiratory kinetics but can be prevented by the addition of Blebbistatin (BLEB), a myosin II ATPase inhibitor, to the assay media (Perry et al., 2011, 2012, 2013). Blebbistatin binds to the active site of subfragment 1 ATPase when ADP and phosphate are bound which stabilizes the intermediate state (Kovacs et al., 2004) lowers force production (Fedorov et al., 2007; Farman et al., 2008; Minozzo et al., 2012) and prevents shortening of muscle cell length (Fedorov et al., 2007; Farman et al., 2008; Ebrahim et al., 2013) during contraction. In fact, unpublished observations within our laboratory revealed drastic differences in mitochondrial respiration rates of PmFB in the presence (+BLEB) and absence (−BLEB) of BLEB from samples obtained using the microbiopsy technique. Moreover, microbiopsy PmFB appeared to be more prone to fiber disintegration and impaired respiratory kinetics following ADP-induced contraction than Bergstrom PmFB. If true, these observations would not only further highlight the importance of PmFB conformation when measuring mitochondrial function, but also suggest BLEB may be an effective tool for rescuing impaired respiration in microbiopsies.

The purpose of this investigation was to compare mitochondrial bioenergetics in samples obtained with the Bergstrom and microbiopsy techniques using BLEB as a tool to control the contractile state of PmFB in vitro, and furthermore, to characterize the fiber type composition of the samples obtained using each biopsy technique. We hypothesized that BLEB would increase respiration in microbiopsy PmFB in vitro by normalizing respiratory kinetics to Bergstrom PmFB. Likewise, we hypothesized that BLEB would result in similar mitochondrial H2O2 emission rates between microbiopsy and Bergstrom PmFBs. Finally, we expected fiber type composition analyses would be similar despite differences in biopsy size between both techniques.

# MATERIALS AND METHODS

#### Human Participants and Muscle Biopsies

Eleven healthy, recreationally active males (n = 5) and females (n = 6) were recruited to participate in this investigation. Their mean ± standard error of the mean (SEM) age, height, weight and BMI were 25.3 ± 0.6 years, 171.7 ± 2.4 cm, 70.6 ± 4.8 kg, and 23.8 ± 1.3 kg·m−2 , respectively. All participants were non-smokers, free of disease and not taking prescription medications or supplements. Participants were given both oral and written information about experimental procedures before giving informed consent. All experimental procedures with human participants were approved by the Research Ethics Board at York University and conformed to the Declaration of Helsinki.

With the participant lying supine on a bed, a skeletal muscle sample was obtained from the lateral aspect of the right vastus lateralis by percutaneous needle biopsy technique using a spring-loaded 14 gauge (∼1.5 mm) Medax Biofeather microbiopsy disposable needle (San Possidonio, MO, Italy) under local subcutaneous anesthesia (∼2 ml of 2% xylocaine without norepinephrine). A 12 gauge (∼2 mm) cannula was used to puncture the skin at ∼30◦ from the surface to a depth of 2 cm and guide the needle to an additional depth of 2 cm longitudinally along the vastus lateralis. Four to five cuts (10–20 mg each) were sampled with the needle rotating ∼30–40◦ between cuts over a period of ∼1 min. Each piece was removed from the needle with sterile forceps or surgical blades before the subsequent cut was made. The first three samples were used for preparation of fiber bundles, and the remaining were used for fiber type analysis (described below). After applying a bandage to the ∼2 mm diameter biopsy site, a second skeletal muscle sample was obtained from the left vastus lateralis using the Bergstrom needle biopsy technique (Bergstrom, 1975) with manual suction (Shanely et al., 2014). Three to four cuts (100–150 mg total) were sampled over a period of ∼10 s. Approximately 40 mg of sample was used for preparation of PmFB, ∼20 mg was used for fiber type analysis and the remaining sample was frozen for future work. The sequence of microbiopsy and Bergstrom sampling was randomized across participants.

# Preparation of Permeabilized Muscle Fibers (PmFB)

This technique is partially adapted from previous methods (Kuznetsov et al., 1996; Tonkonogi et al., 1998) and has previously been described (Anderson et al., 2007; Perry et al., 2011, 2012). Briefly, small portions (∼25 mg) of muscle were dissected from each biopsy and placed in ice-cold BIOPS, containing (in mM): 50 MES, 7.23 K2EGTA, 2.77 CaK2EGTA, 20 imidazole, 0.5 dithiothreitol (DTT), 20 taurine, 5.77 ATP, 15 PCr, and 6.56 MgCl2·6 H2O (pH 7.1).  Each bundle was gently separated along the longitudinal axis with a pair of antimagnetic needle-tipped forceps under magnification (Zeiss 2000, Germany). Bundles were then treated with 30µg/ml saponin in BIOPS and incubated on a rotor for 30 min at 4◦C. Saponin at 30µg/ml has previously been shown to optimize respiration in human skeletal muscle (Kane et al., 2011). Saponin is a mild, cholesterol-specific detergent that selectively permeabilizes the sarcolemmal membranes while keeping mitochondrial membranes, which contain little cholesterol, intact (Veksler et al., 1987; Kuznetsov et al., 2008). PmFB designated for pyruvate supported H2O2 emission (described below) were also treated with 1µM CDNB during the permeabilization process to remove endogenous glutathione and better isolate emission kinetics, as previously described (Treberg et al., 2010; Fisher-Wellman et al., 2013). Following permeabilization, the PmFB were washed in either MiR05 containing (in mM): 0.5 EGTA, 10 KH2PO4, 3 MgCl2·6 H2O, 60 K-lactobionate, 20 Hepes, 20 Taurine, 110 sucrose and 1 mg/ml fatty acid free BSA (pH 7.1) for respiration experiments, or Buffer Z containing (in mM): 105 K-MES, 30 KCl, 10 KH2PO4, 5 MgCl2·6 H2O, 1 mM EGTA, 5 mg/ml BSA, (pH 7.1) for H2O2 emission experiments at 4◦C until measurements were initiated (<30 min).

# Mitochondrial Respiration in Permeabilized Muscle Fiber Bundles

High-resolution O2 consumption measurements were conducted in 2 ml of respiration medium (MiR05) using the Oroboros Oxygraph-2k (Oroboros Instruments, Corp., Innsbruck, Austria) with stirring at 750 rpm. Respiration medium contained 20 mM Cr to saturate mitochondrial creatine kinase (Saks et al., 1991, 1994, 1995; Walsh et al., 2001; Anmann et al., 2006). For ADP-stimulated respiratory kinetics, either 5 mM glutamate or 5 mM pyruvate, accompanied by 0.5 mM malate, were added as complex I substrates (via generation of NADH to saturate electron entry into complex I) followed by a titration of submaximal ADP (225 and 750µM) and maximal ADP (5 mM). Succinate (20 mM) was then added under state three conditions to saturate electron entry into complex II. Cytochrome c was added to test for mitochondrial membrane integrity, with all experiments demonstrating <10% increase in respiration. All experiments were conducted in either the presence (+) or absence (−) of 5µM BLEB in the assay media which allowed for the comparison of respiratory kinetics in contracted or relaxed states for each biopsy technique. Each protocol was completed before the oxygraph chamber [O2] reached 150µM. Polarographic oxygen measurements were acquired in 2 s intervals, with the rate of respiration derived from 40 data points, and expressed as pmol/s/mg wet weight. PmFB were weighed in ∼1.5 ml of tared BIOPS (ATP-containing relaxing media) to prevent rigor that occurs when weighing PmFB in open air (unpublished observations).

# Mitochondrial H2O2 Emission in Permeabilized Muscle Fiber Bundles

Mitochondrial hydrogen peroxide (H2O2) emission was determined fluorometrically (QuantaMaster 40, HORIBA Scientific, Edison, New Jersey) in a quartz cuvette with continuous stirring at 37◦C, in 1 mL of Buffer Z supplemented with 10µM Amplex Ultra Red, 0.5 U/ml horseradish peroxidase, 1 mM EGTA, 20 mM creatine, and 40 U/ml Cu/Zn-SOD1. Either 10 mM succinate or 10 mM pyruvate were added to stimulate H2O2 emission followed by a titration of ADP in step wise increments to progressively attenuate emission. All measurements were made in the presence of 5µM BLEB to allow for the comparison of H2O2 emission in the relaxed state between biopsy techniques. Due to tissue limitations with the microbiopsy, no H2O2 emission measurements were made in -BLEB. The rate of H2O2 emission was calculated from the slope (F/min), after subtracting the background, from a standard curve established with the same reaction conditions and normalized to fiber bundle wet weight as described above.

# Microscopic Imaging of PmFB Conformation

Images of Bergstrom and microbiopsy PmFB conformation were captured prior to, and immediately following respiration experiments. Photographs were captured with a Samsung Galaxy S5 camera (South Korea) placed face-down on a transparent acrylic surface at 2.4-4X zoom focused on PmFB placed in a culture dish containing MiRO. Video recordings with the same device were made to document the change in conformation of both Bergstrom PmFB and microbiopsy PmFB +BLEB and - BLEB following the addition of 2 mM ADP (with 5 mM pyruvate and 2 mM malate) in MiR05 buffer. The media temperature was maintained at 33–37◦C by a pre-heated metal block.

# Immunofluorescence Analysis of Fiber Type

Immediately after removal from the needles, muscles were embedded in O.C.T. compound (Tissue Tek), frozen in liquid nitrogen-cooled isopentane and stored at −80◦C, until analysis. 10µm thick cryosections were cut with a cryostat (Thermo Electronic), maintained at −20◦C and transferred onto static-free microscope slides. Immunofluorescent detection of myosin heavy chain isoforms was performed as previously described (Bloemberg and Quadrilatero, 2012). In addition, fiber membranes were visualized by staining for dystrophin (DSHB, University of Iowa, Iowa City, USA). Fiber type composition analysis was performed on image composites by counting all fibers across the entire cross section. Slides were visualized with an Axio Observer Z1 fluorescent microscope equipped with an AxioCam HRm camera and associated AxioVision software (Carl Zeiss, Germany).

#### Statistics

Results are expressed as mean ± SEM. The level of significance was established at P < 0.05 for all statistics. The D'Agostino— Pearson omnibus normality test was first performed to determine whether data resembled a Gaussian distribution. Due to failed normality, Friedman's non-parametric One-way ANOVA test was performed for all respiration data. When a significant F-ratio was obtained, Dunn's multiple comparisons post-hoc analysis was performed. Paired t-tests were performed to assess differences amongst biopsy techniques with respect to H2O2 emission rates in presence of BLEB as well as fiber type proportions.

#### RESULTS

#### Comparison of ADP-Induced Contraction in Skeletal Muscle PmFB

We first compared qualitative appearances of PmFB conformation from Bergstrom vs. microbiopsy sample in the relaxed state prior to ADP-stimulated respiration (**Figure 1A**). Bergstrom PmFB appeared to have longer and more connected individual fibers forming a more cohesive bundle while microbiopsy PmFB had much shorter individual fibers. Following titrations of ADP over a period of 45–60 min at 37◦C in +BLEB or −BLEB, PmFB were retrieved from the respirometer chambers and imaged. The Bergstrom PmFB in −BLEB appeared less separated and more compact than those from +BLEB, suggesting greater ADP-induced rigor in the absence of BLEB (**Figure 1B**). However, 41% of microbiopsy PmFB from −BLEB disintegrated into multiple pieces, many of which were so small that they could not be removed from the oxygraph chamber for imaging (**Figure 1B**) whereas nearly all PmFB in +BLEB for either biopsy could be removed after the experiments.

Using video capture in separate PmFB, we observed a progressive contraction in response to ADP in −BLEB from Bergstrom (Supplementary Video Figure 1) as previously reported (Perry et al., 2011, 2012) that was also apparent in PmFB from microbiopsy (Supplementary Video Figure 3). ADPstimulated contraction was not observed +BLEB from either biopsy type (Supplementary Video Figures 2, 4). All videos were 4 min in duration and may therefore underestimate the effect of ADP on contraction that occurs during typical respiratory protocols lasting >30 min.

While force production was not measured, these results suggest the change in PmFB conformation in response to ADP was contraction, given the response was prevented by BLEB.

# BLEB Increases ADP-Stimulated Respiration in a Substrate Specific Manner

In the absence of BLEB, microbiopsy respiration was 51% lower than Bergstrom when supported by glutamate and malate at 225µM ADP (p = 0.06) and 51% lower at 750µM ADP (p < 0.05) with no differences at 5 mM ADP (**Figure 2A**). When 5µM BLEB was added to the assay media, the impaired glutamate/malate-supported respiration in microbiopsy PmFB was increased at all ADP concentrations compared to microbiopsy −BLEB (**Figure 2A**). Furthermore, there were no significant differences in respiration between microbiopsy +BLEB, Bergstrom +BLEB or −BLEB (**Figure 2A**) indicating BLEB normalized microbiopsy respiration to both Bergstrom conditions. Contrary to the findings with glutamate/malatesupported respiration, there were, surprisingly, no differences in any condition during pyruvate-supported respiration between microbiopsy and Bergstrom (**Figure 2B**). BLEB increased succinate-supported respiration in Bergstrom by 84–166% vs. both microbiopsy −BLEB and Bergstrom −BLEB, but did not significantly affect microbiopsy respiration (**Figure 2C**).

## Lower Mitochondrial H2O2 Emission Rates in Microbiopsy PmFB

Due to tissue limitations, mitochondrial H2O2 emission was measured in both biopsies only in the presence of BLEB. Succinate-induced H2O2 emission was 37% greater in Bergstrom +BLEB vs. microbiopsy +BLEB at 500µM ADP (p < 0.05) and trended greater at 0µM ADP (p = 0.08), with no difference at 50µM ADP (**Figure 3A**). Similarly, pyruvate induced H2O2 emission rates were 51–66% greater in Bergstrom vs. microbiopsy at all ADP concentrations (p < 0.05, **Figure 3B**).

## Fiber Type Analysis Reveals Differences in Myosin Expression Between Microbiopsy and Bergstrom Biopsies

Immunoflourescent analyses of fiber type composition revealed microbiopsy samples have a significantly lower proportion of type I fibers vs. Bergstrom (37.3 vs. 46.7%, p < 0.05, **Figure 4**) and a significantly higher proportion of type IIX fibers (20.1 vs. 7.6%, p < 0.05, **Figure 4**). However, no differences in relative proportion of type IIA fibers were observed (42.7 vs. 45.7%, **Figure 4**).

### DISCUSSION

This investigation demonstrates that myosin inhibition in PmFB is a novel solution to resolving the reduction in respiratory kinetics observed in microbiopsy samples of human skeletal muscle. Specifically, in −BLEB, PmFB from Bergstrom biopsies appeared more compacted and less separated following ADPstimulated respiration whereas PmFB from microbiopsies had a greater occurrence of disintegration into fragments. This in vitro change in morphology lowered glutamate-supported respiratory kinetics in PmFB from microbiopsies moreso than in Bergstrom. As hypothesized, the lower respiration in microbiopsy PmFB were increased with the myosin II ATPase inhibitor Blebbistatin,

suggesting ADP stimulated contraction of PmFB in -BLEB was responsible for the change in morphology in response to ADP and reduction in respiration vs. Bergstrom. Microbiopsies also contained fewer type I and greater type IIX fibers. However, this does not explain the lower glutamate-supported respiration given kinetics were similar with other substrates in all conditions. Mitochondrial H2O2 emission rates were detectable in samples obtained using the microbiopsy technique but absolute values were lower relative to Bergstrom, even in the presence of BLEB. Collectively, these results highlight the importance of considering needle sampling approaches when assessing mitochondrial bioenergetics in human skeletal muscle and clearly demonstrate that microbiopsies can be used to assess mitochondrial respiration in PmFB when apparent ADP-induced contraction is prevented by BLEB.

# Preventing Microbiopsy ADP-Induced Contraction in vitro Improves Glutamate-Supported Respiration

Microbiopsy sampling is seen as an attractive alternative to Bergstrom biopsies in part, because of the smaller diameter of the needle. However, concerns about the length of fibers obtained from a smaller diameter needle have been raised (Isner-Horobeti et al., 2014). Previous work demonstrating a positive correlation between needle diameter and maximal rates of respiration in PmFB suggested that this impairment in respiration is related to fiber length (Isner-Horobeti et al., 2014). This notion is supported by differences in bundle morphology observed in the present study (**Figure 1A**), which demonstrates shorter myofiber length and a disorganized conformation in microbiopsy relative to Bergstrom. Previous work with Bergstrom PmFB revealed that spontaneous ADP-induced contraction results in drastic changes in conformation including rigor and fragmentation (Perry et al., 2011, 2012). This followed previous work showing ADP causes rigor in PmFB from rodent skinned cardiac fibers (Ventura-Clapier and Vassort, 1985). Given the small fibers in microbiopsies pose a challenge to preparing intact PmFB, it is not surprising that this same apparent contraction results in more consistent and complete disintegration of PmFB

(**Figure 1B**). However, similar to previous findings in Bergstrom PmFB (Perry et al., 2011, 2012), BLEB prevents this apparent ADP-induced contraction and partially retains microbiopsy PmFB conformation, myofiber length and structural integrity (**Figure 1B**). This result demonstrates that BLEB corrects the observed reductions in glutamate-supported respiration reported previously (Isner-Horobeti et al., 2014) and therefore permits the use of microbiopsies for respiratory assessments in PmFB.

Surprisingly, these disintegration-related impairments in respiration were substrate-specific. While we generally observed similar degrees of contraction regardless of the substrate utilized (unpublished findings), this disintegration did not appear to have an impact on pyruvate-supported respiration rates given respiration was similar between both biopsy techniques with or without BLEB present (**Figure 2B**). However, succinatesupported respiration was increased by BLEB in Bergstrom PmFB which was greater than the rate obtained in microbiopsies PmFB. Speculating on the substrate-specific differences in respiration between biopsies is difficult but might suggest contraction exerts a regulatory influence on specific metabolic pathways through mechanisms that might be retained in PmFBs but are yet to be realized. While the reason for these substratespecific differences is unclear, this finding highlights the necessity of considering all experimental parameters when choosing a biopsy technique and the subsequent protocols that will be used. Given BLEB improved glutamate-supported respiration and did not impair kinetics with pyruvate, we suggest that BLEB should be supplemented in assay media during respiratory assessments supported by any of these substrates. In our hands, this approach allays concerns regarding the use of microbiopsies of PmFB as reported previously (Isner-Horobeti et al., 2014).

It should be noted that previous work demonstrating impaired respiration in microbiopsy PmFB was performed at 22◦C (Isner-Horobeti et al., 2014) whereas the present study was conducted at 37◦C. While we cannot conclude whether BLEB would normalize microbiopsy respiration to Bergstrom at 22◦C, we have previously shown BLEB is more effective at influencing respiratory kinetics in human PmFBs at 37◦C vs. lower temperatures (Perry et al., 2011). Nevertheless, it may be preferable to conduct PmFB respiration experiments at or near body temperature given that temperature is a critical regulator of metabolic control in vitro and in vivo.

# Mitochondrial H2O2 Emission is Lower in Microbiopsy vs. Bergstrom PmFB

It is tempting to speculate that the lower H2O2 emission in microbiopsies were a result of fewer mitochondria related to the greater proportion of type IIX fibers and reduced type I fibers. However, succinate- and pyruvate/malate-supported H2O2 emission has been shown to be higher in rat red gastrocnemius muscle [predominantly type IIA and IIX fibers (Bloemberg and Quadrilatero, 2012)] than soleus [predominantly type I fibers (Bloemberg and Quadrilatero, 2012)] when normalized to muscle mass (Anderson and Neufer, 2006). Surprisingly, this apparent inverse relationship between H2O2 emission and assumed mitochondrial content is opposite to our observations of lower H2O2 emission in microbiopsies with greater type IIX content. Thus, based on previous findings (Anderson and Neufer, 2006), it would appear that factors independent of mitochondrial content and fiber type might explain the lower H2O2 emission in microbiopsies.

Anecdotally, it is known that PmFB must be separated as much as possible without losing bundle integrity in order to optimize the detection of H2O2 emission. Specifically, we found Bergstrom fibers can be separated to a greater degree than microbiopsy PmFB without losing integrity (**Figure 1A**). This potentially allows for greater diffusion of substrates and ADP within the fiber, and may indicate that H2O2 emission measurements are more sensitive to the degree of separation than perhaps previously recognized. Hence, it is possible that the relatively lower degree of separation required to maintain integrity in microbiopsy PmFB may result in under-estimations of H2O2 emission.

These measurements were made in the presence of BLEB and suggest that preventing PmFB contraction does not normalize microbiopsy PmFB H2O2 emission to Bergstrom. Whether BLEB still improved this measurement in comparison to untreated PmFB is uncertain as we were not able to compare ±BLEB due to tissue limitations in microbiopsies. Ultimately, H2O2 emission is detected quite clearly in microbiopsy +BLEB which does not rule out the use of microbiopsies for this measurement. Nevertheless, potential underestimations relative to Bergstrom should be considered.

### Fiber Type Differences in Microbiopsies vs. Bergstrom: Does This Influence Respiratory Assessments?

An intriguing difference in fiber type composition existed between samples obtained from the two needles. Specifically, 7/8 subjects demonstrated an increase in percentage of type I fibers and a decrease in percentage of type IIX fibers in the Bergstrom sample relative to microbiopsy. It is well-established that the human vastus lateralis has a heterogeneous composition (Johnson et al., 1973; Komi and Karlsson, 1978; Lexell et al., 1984; Taylor and Bachman, 1999; Staron et al., 2000). In fact, the distribution varies mainly as a function of depth in that the superficial vastus lateralis contains more type II fibers whereas a greater proportion of type I fibers are found in deeper portions of the muscle (Lexell et al., 1983; Lexell and Taylor, 1989). Given the microbiopsy technique was performed at a shallower depth in the present study (∼30◦ insertion angle relative to the skin), these differences in fiber type composition might be attributed to the difference in needle depth. However, we cannot rule out potential contributions of the longitudinal location between the two needles given the shallow insertion angle of the microbiopsy generally sampled at a location more proximal to the knee.

It is tempting to speculate that the lower glutamate-supported respiration with microbiopsies was due to the lower type I and higher type IIX content, as discussed above for H2O2 emission. However, there were no differences in pyruvate or succinatesupported respiration between either needle in ±BLEB. This suggests the fiber type differences in human muscle were too small to influence the kinetics even though large differences in fiber type in rodents have been shown to influence respiration (Kuznetsov et al., 1996). Nonetheless, it would appear that the small differences in fiber type do not explain the lower glutamatesupported respiratory kinetics or H2O2 emission in microbiopsy PmFB -BLEB. We cannot rule out that such composition differences between needles could influence other metabolic assessments.

Ultimately, the differences in fiber type between sampling methods does not necessarily pose a problem for skeletal muscle metabolic assessments per se. Rather, the interpretation of results should be made in the context of sample fiber type composition inherent in each sampling method. Of course, fiber type composition could differ within a sampling approach due simply to variations in sampling depth or longitudinal consistency within trials, highlighting the need for consistency in sampling procedure regardless of needle type.

# Selecting a Muscle Biopsy Approach: Important Considerations

Categorizing one procedure as superior to the other must be considered in the context of specific parameters. The obvious advantage of the Bergstrom method is a larger sample yield with typical ranges of ∼50–250+ mg depending, in part, on the diameter of the needle, the use of suction and the number of cuts. However, we obtained a typical yield of ∼70–90 mg (and up to 120 mg) with microbiopsy by rotating the needle ∼30 degrees to uncut areas of muscle between five separate cuts while applying light pressure on the skin prior to each cut. A fast procedure is required to minimize blood contamination of muscle (<60 s including removal of muscle after each cut). Additionally, participant comfort level must also be considered. While microbiopsies are smaller in diameter, these five separate cuts also require separate insertions into the same cannula guide and therefore take longer to obtain. Conversely, the Bergstrom procedure permits multiple cuts over several seconds with a single insertion. However, 8 out of 11 participants reported preferring the microbiopsy technique over the Bergstrom when questioned following both procedures. Furthermore, reduced risks of infection are sometimes marketed by commercial suppliers of microbiopsy needles due specifically to the fact that they are disposable, thereby eliminating the need for subsequent sterilization. However, proper sterilization of Bergstrom needles obviously ameliorates this concern. These claims must also be balanced with the fact that the single-stage cut with microbiopsies requires removal of skeletal muscle from the needle with sterile forceps prior to re-insertion of the needle into the muscle for additional cuts. This requires considerable care in preventing contamination of the needle in between samples from the same entry site.

It should also be noted that our conclusions are specific to a 14 gauge needle from a specific supplier. Other models of disposable microbiopsy needles could influence the results based on needle sharpness or length of time required to obtain a sample given some needles do not have a cannula "guide" that accelerates the procedure as in the present study. Likewise, we have not compared the influence of multiple gauges, although previous work suggests a positive correlation exists between PmFB respiration and needle diameter (Isner-Horobeti et al., 2014). Finally, experimental procedures requiring intact muscle fibers should carefully consider the requirement for specific fiber lengths independent of sample size. Overall, the advantages and disadvantages to both procedures must be considered within the context of specific study designs rather than focusing on needle diameter and sample yield as the sole criteria for selecting a needle.

#### CONCLUSIONS

In conclusion, this investigation reveals that PmFB prepared from 14 gauge microbiopsies provide similar glutamate-supported respiratory kinetics to Bergstrom

#### REFERENCES

samples when assay media are supplemented with 5µM BLEB to prevent apparent ADP-induced contraction and maintain PmFB integrity. The similar pyruvate-supported respiration between biopsy approaches further suggests that microbiopsies can be used to measure respiratory kinetics in PmFB, although Bergstrom +BLEB provided the greatest rate of succinate-supported respiration. However, observed reductions in H2O2 emission and altered type I:type IIX fiber types should be considered when adopting microbiopsies. These findings highlight critical experimental parameters to consider when selecting biopsy needles and underscores the importance of preventing ADP-induced PmFB contraction by myosin inhibition with Blebbistatin in vitro.

### AUTHOR CONTRIBUTIONS

MH, BG, JQ, DK, and CP. contributed to study design. MH, SR, PT, AN, HT, RL, and CP conducted clinical trials and/or experiments while MH, BB, JQ, DK, and CP analyzed and interpreted the data. MH and CP were the primary writers of the manuscript while all authors contributed to critical interpretation and manuscript preparation. All authors approved the final manuscript.

#### FUNDING

Funding was provided to CP by National Science and Engineering Research Council (#436138-2013) with infrastructure supported by Canada Foundation for Innovation, the Ontario Research Fund and the James H. Cummings Foundation. DK was supported by NSHRF REDI (#2012-8799). MH was supported by a Canadian Institutes of Health Research CGS-M scholarship.

### ACKNOWLEDGMENTS

We thank the study participants for their efforts and dedication. We also thank Dr. Angelo Belcastro, York University for access to his human clinical laboratory and Dr. Giuseppe DeVito, University College Dublin for expert advice in microbiopsy procedures.

#### SUPPLEMENTARY MATERIAL

The Supplementary Material for this article can be found online at: http://journal.frontiersin.org/article/10.3389/fphys. 2015.00360

fatty acid-supported respiration. J. Biol. Chem. 282, 31257–31266. doi: 10.1074/jbc.M706129200

Anmann, T., Guzun, R., Beraud, N., Pelloux, S., Kuznetsov, A. V., Kogerman, L., et al. (2006). Different kinetics of the regulation of respiration in permeabilized cardiomyocytes and in HL-1 cardiac cells: importance of cell structure/organization for respiration regulation. Biochim. Biophys. Acta 1757, 1597–1606. doi: 10.1016/j.bbabio.2006.09.008

study of the fibre type arrangement in m. vastus lateralis of young, healthy males. J. Neurol. Sci. 65, 353–365. doi: 10.1016/0022-510x(84) 90098-4

**Conflict of Interest Statement:** The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

Copyright © 2015 Hughes, Ramos, Turnbull, Nejatbakhsh, Baechler, Tahmasebi, Laham, Gurd, Quadrilatero, Kane and Perry. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) or licensor are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.

# The Measurement of Reversible Redox Dependent Post-translational Modifications and Their Regulation of Mitochondrial and Skeletal Muscle Function

Philip A. Kramer 1 , Jicheng Duan2 , Wei-Jun Qian2 and David J. Marcinek 1, 3 \*

1 Department of Radiology, University of Washington, Seattle, WA, USA, 2 Biological Sciences Division, Pacific Northwest National Laboratory, Richland, WA, USA, 3 Department of Bioengineering, University of Washington, Seattle, WA, USA

#### Edited by:

Gilles Gouspillou, Université du Québec à Montréal, Canada

#### Reviewed by:

Martina Krüger, Heinrich Heine University Düsseldorf, Germany Ranganath Mamidi, Case Western Reserve University, USA

> \*Correspondence: David J. Marcinek<EMAIL_ADDRESS>
#### Specialty section:

This article was submitted to Striated Muscle Physiology, a section of the journal Frontiers in Physiology

Received: 30 September 2015 Accepted: 09 November 2015 Published: 25 November 2015

#### Citation:

Kramer PA, Duan J, Qian W-J and Marcinek DJ (2015) The Measurement of Reversible Redox Dependent Post-translational Modifications and Their Regulation of Mitochondrial and Skeletal Muscle Function. Front. Physiol. 6:347. doi: 10.3389/fphys.2015.00347 Mitochondrial oxidative stress is a common feature of skeletal myopathies across multiple conditions; however, the mechanism by which it contributes to skeletal muscle dysfunction remains controversial. Oxidative damage to proteins, lipids, and DNA has received the most attention, yet an important role for reversible redox post-translational modifications (PTMs) in pathophysiology is emerging. The possibility that these PTMs can exert dynamic control of muscle function implicates them as a mechanism contributing to skeletal muscle dysfunction in chronic disease. Herein, we discuss the significance of thiol-based redox dependent modifications to mitochondrial, myofibrillar, and excitation-contraction (EC) coupling proteins with an emphasis on how these changes could alter skeletal muscle performance under chronically stressed conditions. A major barrier to a better mechanistic understanding of the role of reversible redox PTMs in muscle function is the technical challenges associated with accurately measuring the changes of site-specific redox PTMs. Here we will critically review current approaches with an emphasis on sample preparation artifacts, quantitation, and specificity. Despite these challenges, the ability to accurately quantify reversible redox PTMs is critical to understanding the mechanisms by which mitochondrial oxidative stress contributes to skeletal muscle dysfunction in chronic diseases.

Keywords: redox signaling, post-translational modification, skeletal muscle, mitochondria, myofibrils, glutathionylation

# INTRODUCTION

Oxidative stress plays an important role in skeletal muscle health and disease. In the past, most attention has been focused on skeletal muscle dysfunction caused by oxidative damage to proteins, lipids, and DNA. This damage alters the structure of macromolecules in an irreversible manner and requires turnover and production of new macromolecules to reverse the effects. However, there is growing appreciation of the role of reactive oxygen and nitrogen species (ROS/RNS) in reversible redox post-translational modifications (PTMs) in both muscle adaptation and the effects of chronic disease and aging on skeletal muscle (Sohal and Orr, 2012). Normal skeletal muscle physiology is characterized by transient periods of elevated ROS production due to contractile activity (Powers et al., 2010). In response to repeated bouts of contraction these transient periods of elevated ROS production play an important role in mediating the adaptive benefits of exercise through activation of redox sensitive transcription factors (Powers and Jackson, 2008). On the opposite extreme, chronic elevation of oxidative stress underlies some of the pathological changes associated with disuse atrophy (Min et al., 2011; Powers et al., 2011).

In addition to these longer term adaptive changes in muscle, it is now clear that the intracellular redox environment of skeletal muscle also exerts dynamic control over skeletal muscle physiology on the time scale of seconds to hours. Mild elevation of oxidants increases skeletal muscle force production, while further increases in oxidative stress reduce force and play a role in skeletal muscle fatigue (Reid et al., 1993; Andrade et al., 1998; Matuszczak et al., 2005). Manipulation of muscle oxidative stress also induces rapid effects on mitochondrial energetics. Within 24 h of treatment to induce a mild oxidative stress in mice, paraquat impairs in vivo mitochondrial energetics and induces an energy stress similar to that observed in aged skeletal muscle (Siegel et al., 2012). Reducing mitochondrial oxidative stress by treating with the mitochondrial targeted peptide SS-31 has the opposite effect. Within 1 h of treatment with SS-31 improved mitochondrial energetics and fatigue resistance were associated with a more reduced GSH/GSSG redox state in aged mouse skeletal muscle (Siegel et al., 2013). The rapid and reversible nature of the control of this activity indicates that these effects are not dependent on turnover of oxidatively damaged molecules or protein translation, but rather are the result of a more dynamic interaction between the redox environment and skeletal muscle physiology (**Figure 1**), most likely through reversible redox-dependent PTMs. In this review we will highlight key mitochondrial, myofibrillar, and excitationcontraction (EC) coupling proteins whose activities are modified in a redox dependent manner and discuss the challenges and recent developments in strategies to quantify redox PTMs.

#### SOURCES OF ROS IN SKELETAL MUSCLE

Skeletal muscle produces ROS from both mitochondrial and nonmitochondrial sources. During muscle contractile activity there is an increase in muscle oxidant production. Mitochondria are frequently cited as the primary source for oxidant production in skeletal muscle. However, growing evidence supports a primary role for non-mitochondrial oxidant by membrane bound NADPH oxidases and, to a lesser extent, extracellular xanthine oxidase production during muscle contraction (Gomez-Cabrera et al., 2005; Frasier et al., 2013; Wadley et al., 2013; Alleman et al., 2014; Sakellariou et al., 2014). In fact, recent work demonstrates that under conditions of increased mitochondrial respiration, as in working muscle, mitochondrial superoxide generation is decreased compared to resting conditions (Anderson and Neufer, 2006; Goncalves et al., 2015). NAD(P)H oxidase enzyme complexes are found in the sarcoplasmic reticulum (SR), the transverse tubules, and plasma membrane. Superoxide produced by SR NAD(P)H oxidases has been implicated in regulation of SR Ca2+ dynamics, while plasma membrane NAD(P)H

FIGURE 1 | Theoretical graph of the transient and additive nature of reversible and irreversible redox PTMs in the context of skeletal muscle physiology and disease. (A) Acute oxidative insults (shaded area) can cause a swift and transient increase in reversible oxidative PTMs. Irreversible PTMs rely on protein degradation and turnover, while most reversible modifications can be cleared by the GSH/GSSG and Trx(red)/Trx(ox) redox couples. Subsequent acute oxidative insults can be deleterious if the modifications become irreversible, but this is not the case for reversible modifications, which, can cause transcriptional activation of the antioxidant response (adaptation, preconditioning, or hormesis) to suppress PTM formation. (B) Chronic oxidative insults can result in an oxidized redox status which persists throughout the duration of the stress. As with aging, many metabolic and functional skeletal muscle defects that result from this oxidized redox status can be rapidly reversed by restoring a more reduced redox environment. In contrast, irreversible oxidative modifications, however, can persist after the end of the oxidative insult.

oxidases, along with xanthine oxidase are the main contributors of contraction induced extracellular superoxide production as reviewed by Powers and Jackson (2008). This transient and cyclical contraction induced ROS plays an important role in regulating muscle contraction and inducing adaptive responses to exercise in skeletal (McArdle et al., 2004; Ristow et al., 2009; Powers et al., 2010) and cardiac muscle (Frasier et al., 2013).

In contrast, elevation of superoxide production associated with chronic degenerative disease appears to be primarily mitochondrial in origin.  Reducing mitochondrial H2O2 production with either genetic or pharmacological interventions targeted to the mitochondria reverses or protects against mitochondrial deficits in these conditions (Dai et al., 2011b; Siegel et al., 2013). The mitochondria contain several sites of superoxide production associated with the electron transport system (ETS) and tricarboxylic acid cycle. Complex I of the ETS appears to be the dominant source of mitochondrial superoxide under most conditions with contributions from complex II and III increasing under conditions simulating more active muscle (Goncalves et al., 2015). Complex I and II produce superoxide in the matrix of the mitochondria, while complex III generates superoxide on both sides of the inner mitochondrial membrane (Goncalves et al., 2015). Multiple dehydrogenase enzymes, including pyruvate dehydrogenase and oxoglutarate dehydrogenase, also contribute to superoxide production in the matrix (Mailloux et al., 2014a). Superoxide produced by the mitochondria is rapidly converted to H2O2 by either MnSOD (matrix) or CuZnSOD (inner membrane space). H2O2 then interacts with the GSH-thioredoxin redox buffering system or is rapidly decomposed to water and oxygen by the enzyme catalase in the cytosol.

### REDOX BUFFERING SYSTEM

Cellular redox status, as discussed in this review, is defined as the relationship of ROS producers and scavengers. A common method of determining the redox status of a cell is the reduced to oxidized glutathione ratio (GSH:GSSG). Increasing evidence suggests that redox regulation of cellular processes is the result of signaling rather than non-specific oxidation (Matés et al., 2008). H2O2, for example, interacts with the redox buffering system through the GSH/GSSG and thioredoxin (Trxred/Trxox) redox couples that are present in both the cytosolic and mitochondrial matrix compartments. This redox system, for which NADPH/NADP+ provides reducing power, transduces information on the redox status to the cell by interacting with the proteome through the thiol group on cysteine (Cys) residues. Of the greater than 200,000 Cys residues present in the proteome over 20,000 are thought to be redox sensitive, meaning that they can undergo reversible redox modification as described above (Jones and Go, 2011). The susceptibility of the thiol groups on Cys residues, referred to as redox switches, to oxidative modification varies according to their local biochemical environment. The deprotonated thiol, the thiolate anion, is much more susceptible to oxidation than the thiol group. The pKa of thiols is high (>8); however, its proximity to positively charged amino acids in proteins, arginine, and lysine, can function to decrease their pKa values and make them more reactive at physiological pH (Winterbourn and Hampton, 2008). Redox PTMs of thiols are classified as reversible or irreversible, largely depending on the degree of oxidation. The oxidation states of a low pKa thiol (−2), for example, can be readily oxidized to a reversible sulfenic acid (0) by a 2 electron oxidation with H2O2, and subsequently to sulfinic (+2) and sulfonic acid (+4) which are considered irreversible (**Figure 2**). These redox PTMs can modify protein function by altering protein structure or modifying interactions with other proteins, ligands, or DNA (Forman et al., 2014).

### THIOL-BASED REDOX PTMS

In addition to S-sulfenylation, S-nitrosylation (SNO) and Sglutathionylation (SSG) are two of the most studied reversible redox PTMs (**Figure 2**). These modifications have been wellrecognized as a fundamental mechanism of redox signaling by modulating enzyme activities and protein functions in a variety of cellular activities, such as signaling, metabolism, gene expression, and apoptosis (Janssen-Heininger et al., 2008; Winterbourn and Hampton, 2008; Brandes et al., 2009; Antelmann and Helmann, 2011). The types or levels of PTMs depend on the presence of ROS and RNS, and the redox buffering system, and the enzymatic reduction systems. Collectively, these modifications represent a continuum that relates the form of modification to the extent of ROS/RNS damage and functional consequence. However, our

current knowledge of redox PTMs and their significance is still relatively limited due to the technical challenges associated with the measurement of these labile modifications.

# APPROACHES FOR MEASURING REVERSIBLE REDOX PTMS

To measure endogenous redox PTMs, one of the major challenges is how to preserve the modifications during sample storage and prevent artificial oxidation during sample preparation. There are two general processes of introducing artifacts: (1) thiol oxidation by air through sample processing; (2) reduction of PTMs by antioxidant enzymes during sample storage or processing (Held and Gibson, 2012). Therefore, special care and sample processing protocols are required in order to accurately measure in vivo redox PTMs. To minimize oxidation artifacts during cell or tissue processing, two techniques are commonly applied. These are trichloroacetic acid (TCA) quenching and cell lysis directly with a relative high concentration of alkylation reagents, such as N-ethylmaleimide (NEM) and iodoacetamide (IAA), to block free thiols (Leichert and Jakob, 2006). TCA quenching is a fast process, which can cause precipitation and denaturation of proteins, thus suppressing reactivity of thiolate anions by lowering the pH below their pKa. In contrast, alkylation with NEM and IAM is applied to covalently block free thiols quickly, thus preserve the redox PTMs for downstream processing and measurements. NEM reacts with thiols via a faster and more specific Michael addition reaction than the nucleophilic substitution reaction with IAM. NEM is also less pH dependent compared to IAM, which makes NEM often the preferred blocking reagents for measuring redox PTMs. For measuring redox PTMs in tissues such as muscle, a detailed optimization of tissue handling and processing is particularly important to make sure that the endogenous PTMs are well-preserved during this process.

Approaches for measuring redox PTMs include both indirect and direct detection methods. Given the lack of direct detection approaches of redox PTMs, the indirect approaches are still a popular choice. **Figure 3** illustrates the general chemistry principles of most indirect and direct measurement approaches. Basically, the free thiols are initially blocked by alkylation with NEM in a denaturing buffer during tissue homogenization or cell lysis. For indirect methods (**Figure 3A**), the different types of redox PTMs are selectively reduced to free thiols by using individual sets of reagents where ascorbate is commonly used to reduce SNO (Derakhshan et al., 2007; Su et al., 2013); a glutaredoxin (Grx) reduction cocktail for SSG (Shelton et al., 2005; Zhang et al., 2011), and DTT for total reversible oxidation (Leichert et al., 2008; Paulech et al., 2013). Then, the pre-processed protein samples containing free thiols can be subjected to different enrichment strategies and to either gelbased or mass spectrometry (MS)-based approaches for profiling redox PTM. It should be noted that the selective reduction strategies used in most indirect detection methods are not perfect in terms of specificity. For example, besides reducing SNO, ascorbate is also reported to reduce disulfides in a certain degree (Kuncewicz et al., 2003; Dahm et al., 2006). Similarly, even with the use of mutated form of Grx to reduce its chance for disulfide reduction, non-specific reduction of the Grx enzyme cocktail was still observed (Su et al., 2014). In this regard, the inclusion of proper negative or positive controls when applying these indirect methods is important for more confident identifications of endogenous redox modifications (Guo et al., 2014a).

The concept of indirect detection of redox PTMs was first reported as the biotin-switch technique (BST) for Snitrosylated proteins (Jaffrey and Snyder, 2001; Jaffrey et al., 2001). This approach was also modified to detect other redox PTMs including SSG (Lind et al., 2002; Reynaert et al., 2006) and total reversible thiol oxidation (Leichert and Jakob, 2004). In this approach, the reduced thiols were labeled with reagents containing a biotin tag and labeled proteins were enriched by avidin affinity purification for biochemical (e.g., Western blotting) analyses. For example, redox differential in gel electrophoresis (redox DIGE) allows the comparison of the levels of oxidation between two samples on the same gel by using iodoaceamido or maleimide-based cyanine fluorophores (Leichert and Jakob, 2004; Hurd et al., 2007).

With the advances of MS-based proteomics a number of liquid chromatography-tandem mass spectrometry (LC-MS/MS)-based proteomics approaches have been developed for identifying and quantifying redox PTMs at the proteome level. For example, in BST the biotinylated peptides (formerly SNO-modified) were enriched by avidin chromatography and the biotin tags were removed to allow the detection of SNO modified Cys sites (Hao et al., 2006). Similarly, the OxICAT approach was developed to measure the levels of total reversible thiol oxidation by labeling the reduced and oxidized thiols with two versions of alkylating ICAT (isotope-coded affinity tags) reagents (Leichert et al., 2008). The differentially labeled thiols with ICAT are purified by biotin-avidin purification and followed by LC-MS/MS to quantify the levels of thiol oxidation on Cys residues based on the light and heavy-labeled peptide intensity ratios. Although the BST or biotin-based approaches are still popular, these approaches generally suffer from the issue of nonspecific binding during enrichment. Alternatively, our laboratory developed a resin-assisted capture (RAC) approach (originally called quantitative cysteinyl-peptide enrichment technology, Liu et al., 2004, 2005) for high-efficiency enrichment of cysteinecontaining peptides using Thiopropyl Sepharose 6B thiol-affinity resin. More recently, we and others have demonstrated the effectiveness of enriching several types of redox PTMs including SNO, SSG, and total oxidation using this approach (Forrester et al., 2009, 2011; Liu et al., 2010; Paulech et al., 2013; Su et al., 2013, 2014; Guo et al., 2014a,b). Compared to the biotin-based approaches, the resin-assisted procedure provides a simpler workflow, a very high enrichment specificity (>95% of peptides as Cys-peptides), better sensitivity than biotin avidinbased enrichment as demonstrated by a side-by-side comparison (Forrester et al., 2009). The RAC also provides the flexibility for enabling multiplex quantification by allowing on-resin digestion and isobaric labeling. For example, 4-plex (iTRAQ, isobaric tags for relative and absolute quantification), 6-plex and 10-plex TMT

(tandem mass tags) can be applied to enable MS-based sitespecific identification and quantification of redox PTMs across 4–10 biological conditions (Forrester et al., 2009; Su et al., 2013, 2014; Guo et al., 2014a,b). More recently, a new isobaric reagent iodoTMT that contains an iodoacetyl reactive group was developed to directly label free thiols to enable multiplex quantification of multiple redox PTMs (Pan et al., 2014; Qu et al., 2014). In this approach, the labeled peptides are enriched by an anti-TMT resin. Both the resin-assisted and iodoTMT approach offer the flexibility of multiplexed quantification; however, the resin-assisted approach should provide higher enrichment specificity due to the covalent capture process compared to the non-covalent immunoaffinity enrichment.

Besides the popular indirect methods, several direct measurement strategies (**Figure 3B**) have been developed for redox PTMs. For example, dimedone-based probes have been shown to specifically label SOH (Charles et al., 2007; Leonard et al., 2009). By incorporating a clickable dimedone probe with a UV-cleavable biotin, MS-based global profiling of SOH modified Cys sites was recently demonstrated (Yang et al., 2014, 2015). For SNO, organomercury resin has recently been applied for direct and site-specific identification of SNO-containing peptides by first blocking the free thiols with MMTS and capturing SNO directly by organomercury resin (Doulias et al., 2010, 2013; Raju et al., 2015). Compared to indirect methods, the direct measurement strategies are less prone to potential artifacts for redox PTMs; however, to date the indirect approaches such as RAC or iodoTMT still provide the most flexibility in multiplexed quantification over many biological samples.

Besides these modifications shown on **Figure 3**, protein Ssulfhydration (SSH) has recently been reported as a novel thiolbased PTM similarly to SNO (Paul and Snyder, 2012). A modified version of the BST method was applied to detect proteins modified by S-sulfhydration (Mustafa et al., 2009); however, the blocking reagent MMTS (methyl methanethiosulfonate) was shown to react with both free thiols and persulfides (Pan and Carroll, 2013). More recently, a tag-switch assay was reported for more specific SSH detection (Zhang et al., 2014). Further, studies are necessary to validate the specificity and effectiveness of these approaches for SSH modification.

In the study of redox PTMs, it is also important to be able to be quantify the stoichiometry of redox PTMs (i.e., ratios oxidized vs. reduced thiols) in order to better identify functionally important Cys sites. To date, such measurements were only demonstrated by a few approaches such as OxICAT (Leichert et al., 2008; Knoefler et al., 2012) and the RAC approach coupled with TMT (Guo et al., 2014b) in different biological systems. This would be an important area of further development. Furthermore, targeted quantification of redox PTMs by MS is also an interesting development in its ability to perform precise multiplexed quantification of many site-specific PTMs. The proof-of-concept was demonstrated by coupling differential alkylation of free/oxidized thiols using NEM and d5-NEM and multiple reaction monitoring (MRM) to quantify site-specific Cys oxidation status of an endogenous protein p53 (Held et al., 2010). The targeted quantification strategy should have a good potential to complement Western blotting to verify redox PTMs in specific proteins.

# CONTROL OF PROTEIN STRUCTURE AND FUNCTION BY REDOX PTMS

Changes in protein structure due to an oxidative environment, can affect the localization, proteasome clearance, and the function or activity of a protein depending on the site, type, and extent of the oxidative modification. The formation of a disulfide bond, for example, can affect the proteins tertiary structure, making it a target for proteases. Many proteases have evolved to target oxidatively modified proteins, thus turning over many of the dysfunctional or structurally abnormal proteins (Jung et al., 2013). Lon protease, for example, has been shown to preferentially target oxidized mitochondrial aconitase in skeletal muscle (Bota et al., 2002). However, heavily oxidized proteins, like aconitase, can form aggregates and become inaccessible to the proteasome (Bota and Davies, 2002). In muscle, creatine kinase has been shown to be oxidative modified in aged mice, resulting in protein aggregates and the loss of enzyme activity (Nuss et al., 2009). Oxidative and other PTMs can have a direct effect on protein function independent of structural changes. Actin, for example, has many reported PTMs that affect its polymerization, organization, and stability (Chung et al., 2013; Terman and Kashina, 2013). These PTMs can form in an enzyme active site with similar functional consequences as will be discussed in this review.