This study was supported by NIH grant PO1 HL036573 and University of Toledo, College of Medicine and Life Sciences bridge fund (LL). American Heart Association Scientist Development Grant (12SDG12050473, DK), the David and Helen Boone Foundation Research Fund (DK), and an Early Career Development Award from the Central Society for Clinical and Translational Research (DK).

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**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 Liu, Wu and Kennedy. 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.

# "Oxygen Sensing" by Na,K-ATPase: These Miraculous Thiols

Anna Bogdanova1 \*, Irina Y. Petrushanko2 , Pablo Hernansanz-Agustín3, 4 and Antonio Martínez-Ruiz 3

1 Institute of Veterinary Physiology, Vetsuisse Faculty and the Zurich Center for Integrative Human Physiology (ZIHP), University of Zurich, Zurich, Switzerland, 2 Engelhardt Institute of Molecular Biology, Russian Academy of Sciences, Moscow, Russia, 3 Servicio de Inmunología, Instituto de Investigación Sanitaria Princesa (IIS-IP), Hospital Universitario de La Princesa, Madrid, Spain, 4 Departamento de Bioquímica, Universidad Autónoma de Madrid, Madrid, Spain

#### Edited by:

Sigrid A. Langhans, Nemours Alfred I. duPont Hospital for Children, USA

#### Reviewed by:

John Cuppoletti, University of Cincinnati, USA Pablo Martín-Vasallo, University of La Laguna, Spain

> \*Correspondence: Anna Bogdanova<EMAIL_ADDRESS>
#### Specialty section:

This article was submitted to Membrane Physiology and Membrane Biophysics, a section of the journal Frontiers in Physiology

> Received: 30 May 2016 Accepted: 12 July 2016 Published: 02 August 2016

#### Citation:

Bogdanova A, Petrushanko IY, Hernansanz-Agustín P and Martínez-Ruiz A (2016) "Oxygen Sensing" by Na,K-ATPase: These Miraculous Thiols. Front. Physiol. 7:314. doi: 10.3389/fphys.2016.00314 Control over the Na,K-ATPase function plays a central role in adaptation of the organisms to hypoxic and anoxic conditions. As the enzyme itself does not possess O2 binding sites its "oxygen-sensitivity" is mediated by a variety of redox-sensitive modifications including S-glutathionylation, S-nitrosylation, and redox-sensitive phosphorylation. This is an overview of the current knowledge on the plethora of molecular mechanisms tuning the activity of the ATP-consuming Na,K-ATPase to the cellular metabolic activity. Recent findings suggest that oxygen-derived free radicals and H2O2, NO, and oxidized glutathione are the signaling messengers that make the Na,K-ATPase "oxygen-sensitive." This very ancient signaling pathway targeting thiols of all three subunits of the Na,K-ATPase as well as redox-sensitive kinases sustains the enzyme activity at the "optimal" level avoiding terminal ATP depletion and maintaining the transmembrane ion gradients in cells of anoxia-tolerant species. We acknowledge the complexity of the underlying processes as we characterize the sources of reactive oxygen and nitrogen species production in hypoxic cells, and identify their targets, the reactive thiol groups which, upon modification, impact the enzyme activity. Structured accordingly, this review presents a summary on (i) the sources of free radical production in hypoxic cells, (ii) localization of regulatory thiols within the Na,K-ATPase and the role reversible thiol modifications play in responses of the enzyme to a variety of stimuli (hypoxia, receptors' activation) (iii) redox-sensitive regulatory phosphorylation, and (iv) the role of fine modulation of the Na,K-ATPase function in survival success under hypoxic conditions. The co-authors attempted to cover all the contradictions and standing hypotheses in the field and propose the possible future developments in this dynamic area of research, the importance of which is hard to overestimate. Better understanding of the processes underlying successful adaptation strategies will make it possible to harness them and use for treatment of patients with stroke and myocardial infarction, sleep apnoea and high altitude pulmonary oedema, and those undergoing surgical interventions associated with the interruption of blood perfusion.

Keywords: Sodium-Potassium-Exchanging ATPase, redox regulation, thiols, hypoxia, S-glutathionylation, S-nitrosylation

# INTRODUCTION. OXYGEN, AND OXYGEN SENSING FROM EVOLUTIONARY AND MODERN PERSPECTIVES

# Oxygen, Redox State, Ions, Energy, and Na,K-ATPase

Sustaining of life is a process requiring high energy costs. Energy production in living systems utilizes transmembrane electrochemical gradients, those for ions and redox equivalents. No gradients can be sustained without a membrane, so membranes are the key elements of any living cell since the first proto-cell, the last universal common ancestor (LUCA), that existed more than 3 billion years ago on our planet (Sousa et al., 2013). As life evolved, more specialized compartments were formed within cells, each surrounded by its own membrane. Modern plasma membrane uses the energy of ATP produced by aerobic or anaerobic pathways to generate transmembrane Na+/K+ and Ca2+ gradients. One member of the P-type ATPases family, Na,K-ATPase, transforms the energy of phosphate bonds within the ATP to the energy of transmembrane Na/K gradient that is used to support the excitability of neurons and myocytes, control of intracellular Ca2+ levels and pH, intake of amino acids and fuel, and for sensing and signaling (Blanco and Mercer, 1998; Therien and Blostein, 2000; Blanco, 2005; Geering, 2006; Toyoshima et al., 2011; Reinhard et al., 2013; Shattock et al., 2015). It uses 1 ATP molecule to exchange 3 intracellular Na+ ions for 2 extracellular K+ ions per cycle. The number of cycles varies between 1500 and 10,000 per min (Liang et al., 2007) and the corresponding energy expenditure ranges between 20% of total energy consumption in non-excitable cells to 75% in excitable tissues under hypoxic conditions (Buck and Hochachka, 1993; Erecinska and Silver, 2001). Being a major sink for ATP under hypoxic conditions, Na,K-ATPase is capable of "sensing" the changes in O2 availability and adjusting its activity to the rates of ATP production (Bogdanova et al., 2006). This review summarizes the progress in our understanding of the mechanisms utilized by the Na,K-ATPase for "O2 sensing." Recent developments in of the field of free radical biology and medicine have provided decisive clues for dissection of these mechanisms and the role that protein thiols play in it.

# Reactive Oxygen (ROS) and Nitrogen (RNS) Species As Signal Messengers

Oxygen is a key component of the cell energy production machinery driving Cambrian explosion ∼500 Mio years ago. It plays a role of the final acceptor of the mitochondrial electron transport chain (ETC), which is coupled to ATP production by the H+-dependent reversible mitochondrial ATPase in the mitochondrial oxidative phosphorylation (OXPHOS) system. Many other metabolic reactions in the cell also use oxygen as a necessary component. Thus, cells have developed systems that can sense fluctuations in oxygen concentration and perform different adaptations, and there are several links between hypoxia sensing and redox reactions that we briefly explore here.

Oxygen produces reactive oxygen species (ROS) by successive one-electron reductions (**Figure 1**). Superoxide anion (O•− 2 ) is the first ROS formed from O2; its half-life is as short as 2 × 105 M−1 s −1 (Kalyanaraman, 2013), what makes O•− 2 particularly difficult to detect. It is not a powerful oxidant, and its effect on protein thiols is mainly limited to disruption of iron-sulfur clusters; in this regard, mitochondrial aconitase inactivation has been a classical hallmark of mitochondrial superoxide production (Fridovich, 1995; Hausladen and Fridovich, 1996). It dismutates spontaneously or by the action of cytosolic or mitochondrial superoxide dismutases (Cu,Zn-SOD or Mn-SOD respectively), producing a reduced form, hydrogen peroxide (H2O2), and an oxidized form, water (H2O). Hydrogen peroxide half-life is quite greater than that of its predecessor, 103–104 M−1 s −1 (Kalyanaraman, 2013), it can cross biological membranes and oxidize thiols within Cys residues in proteins. The latter capacity has been a useful tool for nature to design molecular sensors of H2O2; likewise several laboratories have developed different fluorescent proteins capable of detecting H2O2 through reversible oxidation of critical, sensitive thiols in their structure (Hanson et al., 2004; Belousov et al., 2006; Ermakova et al., 2014). All together, these features make H2O2 the most easily detectable ROS and the best known. A one-electron reduction of hydrogen peroxide forms hydroxyl radical (•OH), a reaction that can be catalyzed by the oxidation of Fe2+ into Fe3+ in the so-called Fenton reaction (Kalyanaraman, 2013). •OH is the most reactive and probably the most toxic free radical due to the initiation of radical chain reactions. It is worth mentioning the very fast reaction of superoxide with nitric oxide (NO• ) to form peroxynitrite (ONOO−), with a rate comparable to that of diffusion, 4–6 × 109 M−1 s −1 (Kalyanaraman, 2013). Although peroxynitrite (one of the reactive nitrogen species, RNS) is well known as an inducer of tyrosine nitration, it is a peroxide and as such it is a very effective two-electron oxidant of thiols.

#### ROS/RNS and Hypoxia

Since ROS are chemical derivatives of O2, hypoxia would lower the production of ROS due to the law of mass action, i.e., as there is a lower oxygen concentration, one would expect to observe a decrease in ROS production. However, cell systems are often more complicated than the basic chemical systems in a test tube with multiple players and reactions involved in production and scavenging of ROS and RNS. Apart of thermodynamic and kinetic restrains, compartmentalization within the cell and species/cell type-specific settings affect the capacity of electron donors to react with the oxygen molecules and time course of these reactions. Indeed, there has been a long-standing controversy in the field, as there are a number of observations reporting lower ROS production in hypoxia (Hagen et al., 2003; Acker et al., 2006; Chua et al., 2010; Fernandez-Aguera et al., 2015) but also the opposite (Chandel et al., 1998, 2000; Guzy and Schumacker, 2006), what has been called the paradoxical ROS increase in hypoxia (Turrens, 2003; Guzy and Schumacker, 2006). This controversy emerged from the differences in experimental design including timing of hypoxic exposure prior to ROS detection, cell type, tissue or organism, as well as from the methodological diversity in ROS detection. For example, two groups detecting O•− 2 in cells using superoxidesensing probe (dihydroethidium) and microscopy reported the

opposing findings (Quintero et al., 2006; Chua et al., 2010). The only decisive difference was the time of hypoxic exposure. Detection of O•− 2 in cells following 3 h of hypoxia showed no changes in superoxide levels compared to the normoxic control (Chua et al., 2010), while facilitation of O•− 2 production was observed during the first hour of hypoxia (Quintero et al., 2006). A recentreport reconciled this apparent controversy, as it showed that production of mitochondrial superoxide is upregulated exclusively during the first minutes of hypoxia and its production decreases afterwards, in what has been called a superoxide burst in hypoxia (Hernansanz-Agustin et al., 2014).

Several sources have been proposed to be the origin of ROS in hypoxia. One of the first enzymes implicated in ROS generation in hypoxia was NADPH oxidase, which plays a role in hypoxic pulmonary vasoconstriction (HPV) (Marshall et al., 1996). Together with this, xanthine oxidase was also shown to produce ROS in hypoxia in neurons, contributing to cell death in ischemic conditions (Abramov et al., 2007).

Last but not least, mitochondria-derived ROS have been also shown to increase in hypoxia (Chandel et al., 1998, 2000; Hernansanz-Agustin et al., 2014), are necessary for ROS production by NADPH oxidase in hypoxic pulmonary vasoconstriction (Moreno et al., 2014) and are prior to the xanthine oxidase-derived ROS (Abramov et al., 2007). The source of electron leakage giving rise to an excessive production of O•− 2 remains a matter of intensive investigation. Listed below are some considerations regarding the causes and possible mechanisms of hypoxia-inducible O•− 2 burst in the mitochondria.

# ROS and RNS Production in Hypoxic Mitochondria

Mitochondrial electron transport chain (mtETC) consists of four protein complexes within the mitochondrial inner membrane. Complexes I and II oxidize electron carriers NADH and FADH2, and transfer the electrons to complex III by means of ubiquinone. Complex III reduces cytochrome c, which transports the electrons to complex IV and reduces O2 to H2O. These activities of complexes I, III, and IV are coupled to the pumping of H+ across the mitochondrial inner membrane, thus creating an electrochemical gradient. The oxidative phosphorylation system (OXPHOS) includes a fifth complex, complex V or ATP synthase that transforms the energy of proton gradient to phosphorylation of ADP to ATP (Mitchell, 1961; Mitchell and Moyle, 1967).

It is suggested that ROS generation occurs mainly at the level of complex I, but also at complex III (Turrens, 2003; Murphy, 2009; Drose and Brandt, 2012). FMN group within complex I oxidizes NADH into NAD+ and transfers electrons to a series of Fe-S clusters which, in turn, reduce ubiquinone into ubiquinol (Berrisford and Sazanov, 2009; Hunte et al., 2010; Zickermann et al., 2015) in the so-called forward electron transport. On the other hand, at high mitochondrial transmembrane potential 19mt or in the presence of the complex II substrate succinate, complex I can accept electrons from ubiquinol and reduce NAD+ into NADH at the level of flavin mononucleotide FMN, together with the intrusion of H+ in a process called reverse electron transport (RET) (Vinogradov and Grivennikova, 2005, 2016; Drose and Brandt, 2012). In both forward and reverse transport there is a leakage of electrons giving rise to O•− 2 production. Inhibitors targeting the ubiquinone-binding site (e.g., rotenone; Pryde and Hirst, 2011) increase the leakage in the forward transport, but inhibit it in RET. It is also reported in the presence of high concentrations of succinate, condition associated with progression of ischemia-reperfusion injury (Drose and Brandt, 2012; Chouchani et al., 2014). RET also takes place at elevated 19mt (Korshunov et al., 1997; Drose and Brandt, 2012). O•− 2 produced by complex I will be released toward the mitochondrial matrix where it is dismutated by Mn-SOD (SOD2) to H2O2. The latter is further detoxified by GSH-Glutaredoxin2 system

within the matrix (Drose et al., 2014). Complex III can also give rise to O•− 2 production along with oxidation of ubiquinol to ubiquinone known as Q cycle (Boveris et al., 1976; Cadenas et al., 1977). This process includes formation of semiquinone as an intermediate step (Trumpower, 1990), and this step becomes rate-limiting at high 19mt or in the presence of the complex III-Qi site inhibitor Antimycin A. In case of electron leakage from complex III O•− 2 is accumulating within the intermembrane space. After dismutation by Cu,Zn-SOD (SOD1) the resulting H2O2 is extruded from the mitochondria into the cytosol through porin (Drose et al., 2014) and then detoxified by catalase and GSH/glutaredoxin 2 systems. Free radicals and H2O2 originating from these two complexes most likely serve as local signaling messengers.

Evidence obtained using transgenic animal models suggest different mitochondrial complexes of the mtETC are involved in generation of superoxide anion participating in redox signaling. Knocking down Rieske iron-sulfur protein (RISP) within the complex III indicates the key role of electron transport by this complex for HIF-1α stabilization under hypoxic conditions (Brunelle et al., 2005; Guzy et al., 2005; Mansfield et al., 2005). Silencing of the NDUFS2 protein expression within complex I compromised hypoxia-induced ROS production and arterial chemoreception in vivo (Fernandez-Aguera et al., 2015). However, none of these studies explained the paradox of ROS production in hypoxia or addressed the mechanism of their production. Our recent data point to the key roles of complex I deactivation and Na+/Ca2+ exchange through the mitochondrial Na+/Ca2+ exchanger in the hypoxic superoxide production (Hernansanz-Agustín et al., manuscript submitted).

CO, H2S, and NO production are most likely supported within the mitochondria by heme oxygenase 1 (Ryter et al., 2006), mitochondrial NO synthase (Ghafourifar and Sen, 2007), and 3-mercaptopyruvate sulfurtransferase (Li et al., 2011). All gasses were shown to protect tissues from irreversible suppression of respiratory capacity of mitochondria during ischemia-reperfusion injury (Elrod et al., 2007). In addition to blocking complex III H2S is also capable of direct binding to the heme group of complex IV (Cooper and Brown, 2008). Supplementation of H2S has been shown to inhibit HIF-1α stabilization in hypoxic, but not in anoxic conditions, which is probably related to its capacity of inhibiting mitochondrial respiration (Kai et al., 2012). Systemic administration of sulfide was shown to sustain hibernation state (Blackstone et al., 2005). Nitrite causes inhibition of complex I by S-nitrosylation (reviewed in Martinez-Ruiz et al., 2011) and decreases free radical generation in tissues exposed to ischemia-reperfusion (Shiva et al., 2007).

Genetic adaptation to hypoxia via stabilization of the alpha subunit of hypoxia-inducible factors (HIF1α and HIF2α), has been suggested to require mitochondrial ETC O•− 2 production (Chandel et al., 1998, 2000). However, how ROS act over HIFα subunits or its degrading enzymes, the prolyl-hydroxylases (EGLNs), is still unknown (Kaelin and Ratcliffe, 2008). More recently, mitochondrial complex I and ROS production have been shown to play a key role in acute oxygen sensing by carotid body (Fernandez-Aguera et al., 2015). Such increase in ROS production depolarizes glomus cells through inhibition of K+ channels and increase of cytosolic Ca2+ by extracellular Ca2+ influx. Both stabilization of HIF-α subunits and inhibition of K + channels could be influenced by ROS through oxidation of thiols. Indeed, redox balance is also modified in hypoxia since ROS reversibly oxidize thiols in the cytosolic compartment of cells which, in turn, could have a role in cell signaling and survival during hypoxia (Izquierdo-Alvarez et al., 2012). Superoxide anion generation by mitochondria upon decrease in oxygenation below "normoxic values" as well as decline in NO production by neuronal and inducible NO synthases that have low affinity for O2 (Kd 2–5 kPa; Dweik, 2005) result in an increase in oxidized glutathione in hypoxic heart tissues of animals that are hypoxia-sensitive, but not in hypoxiaresistant ones (Petrushanko et al., 2012; Yakushev et al., 2012). Oxidized glutathione joins the reactions of non-enzymatic dithiol exchange in which S-glutathionylated adducts of thiol residues are formed in multiple proteins including the Na,K-ATPase. For mitochondrial ROS/RNS to be regulators of the Na,K-ATPase function under hypoxic conditions, the ROS generators should be co-localized with the ATPase. The existence of membranebound pool of mitochondria has been confirmed (Westermann, 2015). However, the role of ROS and RNS produced by the mitochondria in control of the Na,K-ATPase activity, amplifying and complementing the signals generated as NADPH oxidases, xanthine oxidase and other free radical generators, awaits further investigation.

Therefore, mitochondria-derived ROS produced in the first minutes of hypoxia (Hernansanz-Agustin et al., 2014), may be a primary and necessary event in redox signaling in hypoxia, leading to the activation of multiple redox processes. All these events converge to targeted thiol oxidation and development of acute adaptive response (Izquierdo-Alvarez et al., 2012).

Both mitochondria and Na,K-ATPase are corner-stones in control of metabolic state of hypoxic tissue. The intimate relation between them remains to be unraveled as it does not seem to be limited to the production and consumption of ATP alone.

#### VERSATILITY OF OXYGEN SENSING. MULTIPLE SIGNALS-MULTIPLE TARGETS—MULTIPLE RESPONSES—MULTIPLE OUTCOMES

Changes in O2 availability occurring in the course if evolution trigger multiple responses in every cell of any living organism on this planet (Holland, 2006). These responses are speciesand cell type-specific. Diversity in responses matches the oxygen levels in the tissue under normoxic conditions (Bogdanova et al., 2006) and is associated with adaptations some species developed to survive hypoxic periods (Hochachka et al., 1999). A single protein, such as the Na,K-ATPase, may demonstrate essential hypoxia-insensitivity (Yakushev et al., 2012), maintain maximal activity within a narrow window of O2 concentrations (Petrushanko et al., 2007), or show linear dose-dependence of O2 concentration in the environment (Bogdanova et al., 2005; Yakushev et al., 2012) depending on the species and cell type. These responses also vary depending on the duration and severity of hypoxia (Dada et al., 2003; Fuller et al., 2003; Bogdanova et al., 2006; Petrushanko et al., 2007, 2015; Yakushev et al., 2012).

Patterns of response correlate with the changes in ROS/RNS and NO production in the cells and the corresponding shifts in redox state. Apart of the regulatory thiols within the Na,K-ATPase changes in the enzyme activity are in part mediated by its phosphorylation by multiple redox-sensitive kinases and phosphatases (Devarie-Baez et al., 2016). These versatile signaling messengers as well as the variability in free radical scavenging systems make responses to hypoxia dependent on location of free radical generators and speciation of the messengers (free radical- and gaseous-based). The mechanisms of oxygen-inducible regulation are largely restricted to the oxidative modifications of thiols (formation of mixed di-thiols with glutathione, sulfide or other protein thiols, S-nitrosylation, or proline/threonine carbonylation (Yan et al., 2013). These changes occur in multiple proteins at the same time. Finetuning of protein complexes is accomplished translating into the protection against hypoxia or reperfusion injury at the tissue and organism levels (Yan, 2014).

# OXYGEN AND REDOX-SENSITIVITY OF THE NA,K-ATPase

#### Na,K-ATPase and Its Thiols

Sodium potassium pump is formed by the 100–113 kDa catalytic α subunit, the regulatory obligatory 55 kDa β subunit and the tissue-specific regulatory proteins of 7–11 kDa belonging to the FXYD family (**Figure 2A**; Blanco and Mercer, 1998; Blanco, 2005; Geering, 2006; Toyoshima et al., 2011). Furthermore, Na,K-ATPase serves as a "docking station" for multiple other proteins of those other ion transporters, receptors, cytoskeletal proteins and members of signaling proteins are known (Reinhard et al., 2013). Each Na,K-ATPase subunit type contains cysteine residues. The only reduced thiol within the beta subunit is localized at the edge to the membrane surface, diving into and out of the membrane during the pumping cycle (White et al., 2009; Bibert et al., 2011). Muscle-specific FXYD subunit pospholemman (PLM) has two thiols (Bibert et al., 2011). The catalytic α subunit is the largest of the three subunits, forms the ATP binding site and binding sites for ions and transport pore, and has 23–24 thiols depending on the isoform (Bogdanova et al., 2006). Several of these thiols are considered to be the targets of regulatory reversible thiol modifications. These modifications make the Na,K-ATPase function extremely sensitive to the changes in redox state and oxygen availability (chronic and acute hypoxia).

#### Irreversible Oxidation of Thiols

Oxidative stress represents an imbalance between an augmented production of pro-oxidative equivalents (mostly ROS and RNS) and/or their detoxification rate by the so-called antioxidant systems (Sies, 2015). Under these circumstances oxidants attack reduced protein thiols turning them into sulfinic or sulfonic acids. Irreversible oxidation induced in the Na,K-ATPase isolated from kidneys by its exposure to 20 mM H2O2 results in a decrease in Vmax to a half and inability to form oligomeric complexes (Kurella et al., 1999). Sensitivity to oxidation varied for the Na,K-ATPase isozymes, those from the heart and the brain (astrocytes) formed by the α2 isoform of the catalytic subunit being more susceptible for oxidation than the ubiquitously expressed α1 isozyme (Xie et al., 1995). ATP was protecting the enzyme from irreversible oxidation (Xu et al., 1997). Depletion of cytoplasmic and mitochondrial GSH or exposure of cerebellar granule cells to hyperoxia also caused suppression of the enzyme activity associated with massive free radical burst (Petrushanko et al., 2006, 2007).

As reviewed below terminal irreversible oxidation of the enzyme has two more consequences. It makes the regulatory thiols inaccessible for reversible thiol modifications and therefore renders the enzyme insensitive to redox changes and alterations in oxygen levels (Petrushanko et al., 2012). Irreversible oxidation primes the proteins to degradation (Thevenod and Friedmann, 1999). In the lungs severe hypoxia (1.5 kPa) promotes ubiquitination and lysosomal degradation of the alpha1-isozyme of the Na,K-ATPase by phosphorylation of the catalytic subunit at Ser18 by PKCzeta (redox-sensitive kinase containing cysteine clusters; Dada et al., 2003, 2007; Dada and Sznajder, 2003).

#### S-nitrosylation of Thiols

S-nitrosylation (and S-nitrosation, for nomenclature see Heinrich et al., 2013) is a common thiol modification in biological systems (Martinez-Ruiz and Lamas, 2004; Martinez-Ruiz et al., 2013). Nitric oxide is unable to directly interact with reduced thiols. Several mechanisms have been described for nitrosothiol formation, which do not require the presence of specific enzymes that catalyze these reactions (Martinez-Ruiz et al., 2013).

Direct reaction of NO to nitrosylate a cysteine thiyl radical (P-S• ) whenever the latter has been formed.

$$\text{P-S}^{\bullet} + \text{ }^{\bullet}\text{NO} \rightarrow \text{ P-S-NO} \tag{1}$$

This is a fast reaction may compete with NO binding to the heme of soluble guanylate cyclase sGC heme (Madej et al., 2008; reviewed in Smith and Marletta, 2012). However, its biological significance is limited to those proteins in which the thiyl radical could be formed when NO is being produced.

NO autooxidation can also lead to S-nitrosylation, via the formation of N2O3:

$$2\text{ NO} + \text{O}\_2 \rightarrow 2\text{ NO}\_2\tag{2}$$

$$\text{NO}\_2 + \text{NO} \rightarrow \text{N}\_2\text{O}\_3\tag{3}$$

$$\text{N}\_2\text{O}\_3 + \text{RSH} \rightarrow \text{RNO} + \text{H}^+ + \text{NO}\_2^- \tag{4}$$

The rate-limiting reaction is the formation of NO2, so it requires a very high NO concentration for producing a significant amount of S-nitrosylation. This high NO concentration can be achieved in the proximity of NO sources, mainly NOS. Indeed, the reaction is 30-fold faster in hydrophobic environments (Moller et al., 2007), suggestion that it can be favored in regions close to cell membranes, where NOS enzymes localize.

schematically the cavities with trapped S-glutathionylated Cys residues inaccessible for de-glutathionylation without detergents and representing "redox memory."

Transnitrosylation in reactions involving modified cysteine residues, including S-nitrosoglutathione, has emerged as an effective and regulated mechanism forming this modifications (Nakamura and Lipton, 2013; Kohr et al., 2014):

$$\text{RSNO} + \text{R'SH} \rightarrow \text{RSH} + \text{R'SNO (trans}-\text{nitroylation,}$$

$$\text{nitroylated glutatiione may be involved}$$

Interaction of peroxynitrite with cysteine thiol residues does not give S-nitrosylated products directly but causes oxidation of thiols to sulfenic acid that in turn can form mixed disulfides, in particular with glutathione (see S-glutathionylation below; Alvarez and Radi, 2003):

$$\text{RS}^- + \text{ONOOH} \rightarrow \text{RSOH} + \text{NO}\_2^-$$

$$\text{RSO}^- + \text{GSH} \rightarrow \text{RSSG} + \text{OH}^- \tag{6}$$

Denitrosylation mainly involves two mechanisms (Smith and Marletta, 2012). GSNO concentration is controlled by enzymatically-catalyzed NADH (for GSNO reductase (GSNOR) or NADPH-dependent (carbonyl reductase 1, CBR1) reduction to GSH. Since trans-nitrosylation of thiols involves GSNO (reaction 5), this process may be considered as rate-limiting in S-nitrosylation of proteins. This is true for the Na,K-ATPase, as increase in the levels of GSNO induces the second transnitrosylation step and formation of S-glutathionylated adducts of the α (Li et al., 2001; Petrushanko et al., 2015) and β (Liu et al., 2013) subunits of the enzyme. Thioredoxin catalyzes denitrosylation of multiple targets by trans-nitrosylation of its active cysteines Cys32 and 35.

Hypoxia alters the number of S-nitrosylated cysteines in numerous proteins. In endothelial cells with high levels of eNOS (NOSIII) multiple targets get S-nitrosylated under hypoxic conditions. This NO synthase has high affinity to O2 (Kd 0.29 kPa, Dweik, 2005) and can therefore support NO production at low O2 levels. At the same time in heart tissue, where NOSI and NOSII with low O2 affinity are dominating (Kd 23–35 kPa, Dweik, 2005) hypoxia is associated with rapid cessation of NO production and decrease in S-nitrosylation of cysteine residues of the α subunit of the Na,K-ATPase (Yakushev et al., 2012). NO and S-nitrosylation were shown to protect thiols from irreversible oxidation and S-glutathionylation, and allow the enzyme to maintain high activity in the brain and heart (Petrushanko et al., 2007; Yakushev et al., 2012). Irreversible oxidation makes regulatory thiols inaccessible for S-nitrosylation. This may explain the fact that the sensitivity of the Na,K-ATPase to NO was lost in a mouse model of amyotrophic lateral sclerosis (Ellis et al., 2003).

#### S-glutathionylation of Thiols: Chemistry

S-glutathionylation (also referred to as S-glutathiolation), the formation of a mixed disulfide between a protein cysteine and that of glutathione. This thiol modification is produced in a number of chemical reactions (Klatt and Lamas, 2000; Dalle-Donne et al., 2007, 2008; Martinez-Ruiz and Lamas, 2007; Mieyal et al., 2008; Bachi et al., 2013).

Reactions resulting in formation of S-glutathionylated protein adducts include classical thiol disulfide exchange (reaction 7) that does not require any catalysis, and several mechanisms requiring prior oxidative modification of the thiol residues (either the protein thiol or cysteine residue of GSH, reactions 8–13).

$$\text{P-S}^- + \text{GSSG} \rightarrow \text{P-SSG} + \text{GS}^- \tag{7}$$

Reactions priming thiols to S-glutathionylation include prior formation of thiyl radicals with O2 as an electron acceptor (reactions 8–9), oxidation of thiols to sulphenic acid (reaction 10–11) or to nitrosothiol (reactions 12–13)

$$\text{P}-\text{S}^{\bullet} + \text{GS}^{-} + \text{O}\_{2} \rightarrow \text{P-SSG} + \text{O}\_{2}^{\bullet -} \tag{8}$$

$$\text{P}-\text{S}^- + \text{GS}^\bullet + \text{O}\_2 \rightarrow \text{P-SSG} + \text{O}\_2^{\bullet-}$$

(catalyzed by glutaredoxin) (9)

P-SOH + GSH → P-SSG + H2O (10)

P-SH + GSOH → P-SSG + H2O (11)

P-SNO + GSH → P-SSG + HNO (12)

$$\text{P-SH} + \text{GNO} \rightarrow \text{P-SSG} + \text{HNO} \tag{13}$$

S-glutathionylation of thiols in hypoxic cells is triggered by a local shift in GSSG/GSH ratio toward more GSSG (Petrushanko et al., 2012; Yakushev et al., 2012). On the other hand, oxidative stress following superoxide anion burst in hypoxic mitochondria, will promote oxidation of thiols to sulfenic acid priming them to S-glutathionylation via reaction 10.

Reversibility of S-glutathionylation is supported. Glutaredoxin1 and 2 may catalyze de-glutathionylation or glutathionylation reactions depending on the redox environment (NAD(P)H:NAD(P), GSH:GSSG, NO availability; Mieyal et al., 2008). Not all thiols in the proteins are accessible for Sglutathionylation. Whereas glutaredoxin1 is active in the cytosol and inter-membrane space, whereas glutaredoxin 2 is localized in the mitochondrial matrix, controlling the state of protein thiols in these compartments (Allen and Mieyal, 2012). Further enzymes capable to catalyze de-glutathionylation reaction include sulfiredoxin (Lei et al., 2008), glutathione transferase P (Townsend et al., 2009), and glutathione transferase omega 1 (Menon and Board, 2013).

Not all thiols are equally accessible for S-glutathionylation. Specific conditions that have to be fulfilled for the thiol to become glutathionylated are discussed below.

Localization of the Groups Favoring Glutathionylation The likelihood for a given thiol to undergo S-glutathionylation is defined by several factors: (i) dissociation constant pKa of a thiol (ii) microenvironment (amino acid composition in vicinity to the cysteine residue), (iii) accessibility of the group and steric restrictions, and (iv) redox potential of a thiol (Nagy, 2013). Nucleophilic substitution reaction in which S-glutathionylated adduct is formed involves interaction of a thiolate anion with a GSSG molecule (Allen and Mieyal, 2012). Thiol therefore has to be deprotonated to join this reaction. The average pKa of majority of cysteine residues is 8.5. Thus, probability of deprotonation of cysteine residues under physiological conditions is relatively

low making S-glutathionylation selective (Dalle-Donne et al., 2009). Reduction in pKa of distinct thiols is supported by the positive charge of three flanking amino acids such as arginine, lysine, or histidine (Allen and Mieyal, 2012; Sun et al., 2013; Zhao et al., 2015). Proximity of negatively charged amino acids to a thiol on the contrary compromises interaction with GSSG. Protein sequence analysis for the α1 subunit of the Na,K-ATPase revealed that cysteines 204, 452, 599, and 698 (corresponding to the Cys 206, 454, 601, and 700 in **Figure 2**) are prone to S-glutathionylation (Mitkevich et al., 2016).

Steric restriction and localization of cysteine residue within the protein sequence are yet other parameters that should be taken into consideration when predicting S-glutathionylation sites (Pineda-Molina et al., 2001). Hydrophilicity of GSSG suggests that the residues should be exposed to aqueous phase as the reaction with GSSG occurs. Furthermore, the exposed surface of thiol should be sufficient for docking of GSSG to form mixed dithiols (Sun et al., 2013). "Basal S-glutathionylation" of cysteines that are buried within the protein, but are S-glutathionylated, represents a special case and is discussed below.

Further restrictions limiting the number of S-glutathionylated residues are related to the redox state of a thiol. For most proteins GSH:GSSG ratio should decrease from 100:1 to 1:1 to achieve S-glutathionylation of 50% of thiols (Allen and Mieyal, 2012). Very few proteins, such as c-Jun may be glutathionylated within physiological range of half-cell redox potential for GSH:GSSG couple (Klatt et al., 1999; Allen and Mieyal, 2012). Na,K-ATPase is one of these few proteins as interaction of the cysteine residues within its catalytic α subunit with GSSG can be observed as GSSG reaches 150 µM levels in the presence of 1.5 mM GSH (Petrushanko et al., 2012; Yakushev et al., 2012).

# S-glutathionylation and Targets in the Na,K-ATPase

#### Alpha Subunit of the Enzyme

Cysteines of all three types of subunits forming Na,K-ATPase undergo S-glutathionylation. Catalytic α subunit contains 23 or more (depending on the isoform) cysteine residues (**Figure 2A**). Fifteen out of 23 Cys residues of the α1 isoform are localized within the cytosolic loops and are easily accessible for interaction with cytosolic GSSG. Out of seven cysteine residues of the regulatory β subunit 6 are forming S-S bonds with each other and only one remaining cysteine possesses a thiol accessible for S-glutathionylation (White et al., 2009). Tissue-specific FXYD subunits also contain 1–2 cysteines of which at least one undergoes S-glutathionylation (Bibert et al., 2011).

Regulatory S-glutathionylation Cysteine residues that were found S-glutathionylated within the α1 subunit isolated from duck salt gland were classified as targets for regulatory or basal S-glutathionylation (Petrushanko et al., 2012). Whereas basal S-glutathionylation is not associated with the changes in the enzyme hydrolytic activity, binding of glutathione to the regulatory cysteine residues causes the enzyme's complete inactivation that can be reversed upon de-glutathionylation (Petrushanko et al., 2012; Yakushev et al., 2012). Regulatory Sglutathionylation could be accomplished by incubation of the purified enzyme or membrane fraction with GSSG indicating that regulatory thiols were accessible for classical thiol disulfide exchange (reaction 5). Similar response was obtained for the α1 subunit within the broad range of tissues and cell type (cell line, salt gland, kidneys, heart tissue) and species (duck, rabbit, pig, mouse, rat, Spalax mole rat, trout, and human; Petrushanko et al., 2012, 2015; Yakushev et al., 2012; Juel, 2014; Xianyu et al., 2014; Juel et al., 2015; Mitkevich et al., 2016). Apart of α1, Sglutathionylation of the α2 isozyme was shown in heart and skeletal muscle. Moreover, α2 isozyme appeared to be more sensitive to GSSG-inducible inhibition in rat heart (Petrushanko et al., 2012; Xianyu et al., 2014; Juel, 2016). This observation is in agreement with the report of Xie on exceptionally high susceptibility of the α2 subunit to oxidation (Xie et al., 1995) and suggests that Cys residue(s) within this isoform are more accessible for oxidation (e.g., Cys 579 in **Table 1**). As can be seen from the **Table 1**, microenvironment for two cysteine residues (Cys 206 and 579) within the cytosolic loops of the α2 subunit favors S-glutathionylation as non-charged amino acids present in the α1 isoform are substituted by positively charged ones facilitating thereby deprotonation of the thiols. Furthermore, α2 subunit harbors the extra cysteine (Cys 236) in the actuator domain and lacks the Cys458 (present in the α1). Thus, although the number of cysteines does not differ between the α1 and α2 subunits, their location, pK and the ability to become Sglutathionylated show clear isoform-specificity.

Localization of the sites of regulatory S-glutathionylation was identified by means of mass spectrometry (Petrushanko et al., 2012). Three of them, Cys 454, 456, and 459 are proximal to the ATP binding site whereas Cys 244 is localized in the small cytosolic loop that may approach the ATP binding pocket in E2 conformation (Bogdanova et al., 2006). Binding of glutathione to the regulatory sites displaces adenine nucleotides from interaction with the enzyme (Petrushanko et al., 2012). In turn, GSSG cannot block the enzyme's hydrolytic activity in the presence of ATP or ADP (Petrushanko et al., 2012; Xianyu et al., 2014). ATP was showing maximal "protective effect" compared to the other nucleotides (Xianyu et al., 2014). Interaction with ATP (but not with the other nucleotides) brings the nucleotidebinding domain of the α subunit closer to the phosphorylation domain (E1-closed state) shielding thiols of the small and large cytosolic loops from attack by GSSG or oxidants (Petrushanko et al., 2014) and making them less prone to irreversible oxidation (Xie et al., 1995; Xu et al., 1997).

Increase in S-glutathionylation of the α subunit was triggered by hypoxia associated with mild oxidative stress and modest ATP depletion in rat heart and SC1 cell line derived from mouse fibroblasts (Petrushanko et al., 2012, 2015; Yakushev et al., 2012). Inhibition of the Na,K-ATPase and induction of S-glutathionylation could be mimicked by the modulation of intracellular redox state by exposure of cells to glutathione derivatives et-GSH, GSNO, GSSG, or depletion of the intracellular GSH (Petrushanko et al., 2006, 2015). Sglutathionylation of the α subunit cannot be sustained under anoxic conditions. This thiol modification is induced by 0.2% O2

#### TABLE 1 | Analysis of flanking amino acids' composition of selected thiols in the α1 and α2 subunits (rat sequences).

Data shown in the table are obtained as a result of sequence alignment (Clustal O(1.2.1)) for the Na,K-ATPase α1(P06685) and α2 (P06686) subunits' sequences for rat (Rattus norvegicus), published in UniProt-Knowledgebase (Swiss-Prot collection of sequences, http://www.uniprot.org/uniprot). Included into the table are the amino acids flanking the cysteine residues for which the isoform-specific differences (substitution, omission) have been observed. Asterisks stand for the conserved amino acids whereas dots and semicolon indicate the substitutions. Cysteines are color-coded in blue and the substituted amino acids in yellow.

but less pronounced at 0.05% O2 in mouse fibroblast-derived cell line (Petrushanko et al., 2015).

#### **Basal S-glutationylation of the** α **subunit**

Basal S-glutathionylation of the α subunit of the Na,K-ATPase is not associated with the changes in the enzyme function. 15 cysteine residues facing the cytosol are accessible for Sglutathionylation. Treatment of the cell lysates, membrane fractions or purified active Na,K-ATPase preparations with reducing agents (e.g., by Dithiotreitol or Tris(2-carboethyl) phosphine, TCEP) could not completely remove glutathione bound to the α subunit's thiols (Mitkevich et al., 2016). Complete de-glutathionylation could only be achieved under conditions supporting partial denaturation of the protein (8 M urea and 8% SDS). De-glutathionylation of the cysteines inaccessible for reducing agents in the protein retaining its native structure was associated with substantial loss of the enzyme activity due to unfolding. The only chance these Cys residues shielded from GSSG and reducing agents have to acquire S-glutathionylation is before the folding was completed (**Figure 2B**). This implies that S-glutathionylation of certain Cys residues is a co-translational rather than post-translational modification and it may be required for correct protein folding. Detailed analysis of the X-ray structures of the porcine α1 subunit-containing enzyme (PBD codes: 3B8E, 3KDP, 3WGU, 3WGV, 4HYT) revealed a number of isolated cavities with unresolved electron density next to the Cys residues (numbering as in **Figure 2B**) Cys 206–Cys 244; Cys 454–Cys 458–Cys 459; Cys 700–Cys 369, Cys 601 (Mitkevich et al., 2016).

These regions of relatively high residual electron density that cannot be explained by the presence of water are sufficient in size to home glutathione. However, so far no X-ray structures of the actual catalytic subunit with S-glutathionylated cysteine adduct was reported. Detection of glutathionylated residues in crystal structures of proteins is not impossible (Srinivasan et al., 2014), but much less common that detection of these modifications by means of mass spectrometry. One of the technical approaches to use for identification of S-glutathionylated cysteine residues is tracking for glutathione localisation using analysis of unresolved density next to the cysteine residues. This approach was used for identification of glutathione bound to the ABC transporter Atm1 (Srinivasan et al., 2014). In each cavity of the α subunit only one Cys residue was reported to be S-glutathionylated (basal type of S-glutathionylation, **Figure 2B**, Mitkevich et al., 2016). These findings imply that Cys454 is located in a cavity and, hence, cannot be a regulatory thiol as suggested earlier (Petrushanko et al., 2012). Its basal S-glutathionylation will contribute to the protein folding instead. Out of the Cys206-Cys244 couple Cys 206 is capable of basal S-glutathionylation whereas Cys 244 carries a regulatory thiol group. In contrast to regulatory Sglutathionylation observed within minutes after the drop in O2 availability, increase in basal S-glutathionylation is only observed after 72 h of hypoxic exposure (Mitkevich et al., 2016). This kinetics correlate with the onset of de novo protein synthesis rather than acute alterations in thiol state.

Moreover, as this modification is retained during the life-span of a protein it represents "redox memory" that reflects the cellular redox state at the moment of synthesis of this molecule. Thus, this "redox memory" may represent the process of adaptation to the alterations in redox state occurring in particular in response to deoxygenation. The fact that basal S-glutathionylation levels differ in muscle fibers being higher in oxidative fibers compared to the glycolytic fiber type suggests that it is likely to be associated with the metabolic state of the tissue (Juel, 2014).

#### S-glutathionylation of Beta Subunit

A single reduced thiol group in β subunit of the Na,K-ATPase, Cys 46 (**Figure 2A**), is the one that may undergo reversible posttranslational modifications. S-glutathionylation of this cysteine residue was reported for all three muscle types (skeletal, smooth, and heart muscles; Figtree et al., 2009; Liu et al., 2013; Juel et al., 2015). Since interaction of glutathione with Cys 46 results in down-regulation of the enzyme function this residue is a site of regulatory S-glutathionylation (Figtree et al., 2009). Inhibitory effect is achieved due to the weakening of the interaction between α and β subunits upon S-glutathionylation (Figtree et al., 2009). S-glutathionylation of β subunit does not inactivate the enzyme completely (Figtree et al., 2009). Unfortunately, the studies in which S-glutathionylation of the β subunit was detected were not presenting the information on the degree of S-glutathionylation of the α subunit, making it impossible to discriminate between the impacts of these two processes into the enzyme activity regulation (Liu et al., 2013).

In contrast to the regulatory cysteines of the catalytic α subunit that are readily interacting with GSSG Cys 46 does not join reaction of classical thiol disulfide exchange with GSSG (Petrushanko et al., 2012). This may be attributed to the localization of this Cys residue. According to the X-ray structure it is buried within the membrane in E2 2K+Pi conformation (Ogawa et al., 2009) and is only accessible for S-glutathionylation in E1ATP and E1Na(3) conformation (Liu et al., 2012). Furthermore, Cys 46 may become accessible or Sglutathionylation upon the loss of association between the α and β subunits (Garcia et al., 2015). Ouabain stabilizes the enzyme in E2 conformation and compromises S-glutathionylation of β subunit (Liu et al., 2012). De-glutathionylation can be achieved by treatment of the S-glutathionylated enzyme with Glutaredoxin.

Localization of the regulatory cysteine defines conditions required for its S-glutathionylation. It only occurs in the presence of peroxynitrite and hence involves reactions 12 and 13. Physiological and pathophysiological role of this form of regulatory thiol modification in the heart has been intensively investigated. Stimulation of O•− 2 production by NADPH oxidases, that co-localize and co-immunoprecipitate with the Na,K-ATPase, induces S-glutathionylation of the β subunit at Cys46 (Liu et al., 2013). The stimulation of NADPH oxidases could be triggered by their phosphorylation by PKC produced upon activation of β1/β2 adrenoceptors or treatment of the myocardium with angiotensin II. Furthermore, increase in S-glutathionylation of the β subunit was reported in the infarcted area in sheep heart (Figtree et al., 2009). Similar effect was achieved by exposure of cardiomyocytes to the activator of adenylate cyclase forskolin and the following activation of PKCǫ (White et al., 2010) or direct administration of ONOO- (Figtree et al., 2009). Scavenging of O•− 2 using superoxide dismutase on the contrary abolishes S-glutathionylation of the β subunit. Similar effect may be achieved by the stimulation of sGC that interfered with phosphorylation and activation of NADPH oxidases (Chia et al., 2015). Activation of β3 adrenoreceptor is a physiological stimulus decreasing S-glutathionylation of the Cys 46 (Bundgaard et al., 2010). These differential responses are easily explained based on the chemistry of S-glutathionylation of thiols. Deoxygenation is associated with suppression in production of NO by NOS1 and NOS2 as well as in reduction of activity of NADPH oxidases Kd of which for O2 is within 2 kPa range. Thus, decrease in O2 levels below this threshold reduces production of peroxynitrite and decreases S-glutathionylation of Cys 46 in β subunit. Uncoupling of oxidative phosphorylation and O•− 2 burst in the mitochondria on the contrary will facilitate GSSG production in the cytosol and induce S-glutathionylation of regulatory cysteines within the cytosolic loops of the catalytic α subunit.

#### S-glutathionylation of FXYD Subunits

Tissue-specific FXYD proteins associate with Na,K-ATPase αβ complex stabilizing it and modulating the enzyme activity (Geering, 2006). These modulatory subunits also contain 1 or 2 cysteine residues that undergo reversible thiol modifications. Most of the information on the role of these modifications in control of Na,K-ATPase function was obtained for the cardiacspecific FXYD1 subunit also known as phospholemman (PLM). It contains 2 Cys residues (C1 corresponding to Cys40 and C2 corresponding to Cys42 in human FXYD1 protein, **Figure 2A**), and S-glutathionylation of the C2 was shown to correlate reciprocally with the availability of glutathionylated form of the β subunit (Bibert et al., 2011). Localization of C2 and mechanisms of induction of S-glutathionylation of PLM are most likely shared with the β subunit of the Na,K-ATPase and involve peroxynitrite. Interaction of PLM with glutathione may be triggered by forskolin and prevented by exposure to superoxide dismutase. De-glutathionylation is also catalyzed by glutaredoxin1 or treatment with DTT. Thus, both PLM and β subunits lack cysteines that are not accessible for de-glutathionylation (redox memory). Physiological stimuli increasing the level of PLM Sglutathionylation include angiotensin II and infarction (Bibert et al., 2011).

The importance of the amino acids flanking the cysteine residue in control of accessibility of it for S-glutathionylation was emphasized as the amount of glutathionylated adducts was compared for several members of FXYD family. C2 cysteines were found to be prone to S-glutathionylation in FXYD2 (renal) and FXYD7 (brain-derived) (Geering, 2006; Bibert et al., 2011). Phospholemman (PLM, FXYD1) interacts with both catalytic and regulatory subunits of the Na,K-ATPase. Thus, interaction between α and β subunits and the enzyme function is affected by S-glutathionylation of the FXYD1. Furthermore, the reciprocal regulation of S-glutathionylation of the β and FXYD subunits suggests that FXYD may reverse the inhibitory action mediated by binding of glutathione to the regulatory β subunit on the Na,K-ATPase. It remains unclear if glutathione is transferred from the Cys46 on to the β subunit to the C2 residue in reaction of thiol disulfide exchange of the FXYD protein or if Sglutathionylation of the FXYD protein makes Cys46 inaccessible for S-glutathionylation.

# H2S and the Na,K-ATPase

Hypoxic exposure was shown to cause a rapid increase in H2S levels in a number of cells including chemoreceptory glomus cells (Peng et al., 2010; Olson, 2015; Yuan et al., 2015). Direct interaction of H2S with the Na,K-ATPase has not been reported. However, exposure of renal tubular epithelial cells to H2S causes internalization of the Na,K-ATPase in complex with other proteins. This inhibitory response of the pump is triggered by sulfhydrolation of cysteines 797 and 798 at the epidermal growth factor receptos (EGFRs) (Ge et al., 2014).

# COORDINATED REGULATION OF REDOX-SENSITIVE PROTEIN NETWORKS BY REVERSIBLE THIOL MODIFICATIONS

S-glutathionylation and S-nitrosylation are altering the function of numerous proteins that form redox sensitive networks (Sun et al., 2006; Dalle-Donne et al., 2009). Protein clusters forming these networks are the ones controlling metabolism, ion transport, cell fate and cycle, adaptation, and others. This type of free radical-based signaling is of key importance in responses to hypoxia and reoxygenation mediating acute and long-term tissue-specific coordination of function of multiple proteins. S-glutathionylation of the α subunit of the Na,K-ATPase occurs among with alterations in thiol state of other proteins. The most obvious protein partners that undergo Sglutathionylation are the adjacent β and FXYD subunits. Thiol disulfide exchange in which PLM most likely acquires glutathione from the Cys 46 subunit of the β subunit is suggested to cause dissociation of the FXYD subunit of from the Na,K-ATPase as it does not co-precipitate with the α subunit (Bibert et al., 2011). As PLM is a regulatory subunit that is shared between the Na,K-ATPase and the Na/Ca exchanger, it was suggested that Sglutathionylation as well as its redox-sensitive phosphorylation of it impacts activity of both transporters (Silverman et al., 2005; Cheung et al., 2007; Zhang et al., 2011). Calcium handling in the heart is intimately linked to the Na+ levels (Shattock et al., 2015) and, hence to the activity of the α2 isozyme of the Na,K-ATPase that is particularly prone to inhibition by GSSG (James et al., 1999; Petrushanko et al., 2012). Apart of the Na/Ca exchanger reversible thiol modifications are known to control the activity of L-type Ca2+ channel, RyR2 ryanodine receptors, SERCA2a Ca2+ pump and multiple other ion transporters (Sun et al., 2006; Bull et al., 2008; Lancel et al., 2009; Donoso et al., 2011).

These coordinated redox-driven changes in activity of multiple ion transporters may support cytoprotection and survival of the cell/tissue/organism or promote death under hypoxic conditions (see Section Regulation of the Na,K-ATPase Activity by Cardiotonic Steroids).

# OXYGEN-SENSITIVE PHOSPHORYLATION

Regulatory phosphorylation is known to control the activity, sensitivity to the cardiotonic steroids, and availability of the Na,K-ATPase on the plasma membrane. Changes in phosphorylation state of the enzyme were reported in response to hypoxic challenge and for a long time believed to be the only mechanism involved in "channel arrest" response of the Na,K-ATPase in hibernating animals (MacDonald and Storey, 1999; Ramnanan and Storey, 2006; McMullen and Storey, 2008). At present "channel arrest" responses seem to be more versatile, and yet triggered by a single mechanism of regulation, in this case not only the Na,K-ATPase itself, but also the redox-sensitive kinases, namely, reversible thiol modifications of regulatory cysteine residues or action of gasotransmitters. Phosphorylation sites at the α, β and FXYD subunits are schematically presented in **Figure 3**.

#### Protein Kinase C (PKC)

Hypoxic exposure of alveolar epithelial cells was shown to trigger actively controlled internalization of the Na,K-ATPase in the form of clathrin-coated vesicles (Dada et al., 2003). This process was induced by phosphorylation of the α subunit of the Na,K-ATPase by at atypical PKC isozyme.

Novel and classical PKC isoforms are also capable to alter activity of the Na,K-ATPase in various cell types. Classical (conventional α, β1 and β2, and γ isoforms) and novel isoforms PKC δ, ε, η, θ, and µ may be activated diacylglycerol (DAG), phorbol esthers and phosphatidylserine and are sensitive to the changes in Ca2+ in the cell. Atypical PKCι and ξ retain the sensitivity to phosphatidylserine but are Ca2+-insensitive (**Figure 3**; Newton, 1995; Parker and Murray-Rust, 2004).

Catalytic core of PKC contains critical cysteine-rich motifs that are conserved in all PKC isozymes making all proteins of this class redox-sensitive (Newton, 1995). This motif is duplicated in classical and novel PKC isoforms, whereas in atypical PKCs only one repeat is found. Oxidation of these cysteines causes transient activation of the enzyme but makes it unable to interact with DAG and phorbol esters making it Ca2+- and DAG-insensitive (Gopalakrishna and Anderson, 1989). S-glutathionylation of these regulatory thiols is inactivating α, βI, βII, γ, δ, ε, and ζ isoforms of PKC. Formation of glutathionylated adducts occurs within physiological range of GSH concentrations (0.5–10 mM) and results in the enzyme inactivation (Ward et al., 1998, 2000; Chu et al., 2001). In addition, PKC may undergo S-nitrosylation that was shown to suppress activity of PKCα (Choi et al., 2011). Depletion of intracellular GSH by exposure to conjugating agent

diethylmaleimide or inhibition of de novo GSH production by L-buthionine-S,R-sulfoximine results in reduction in activity of conventional PKC isoforms (α, βI, and βII) by 35–50%, along activation of the novel isoforms (δ and ε) (Domenicotti et al., 2000). This response of the PKCs to the changes in redox state will induce endocytosis of the Na,K-ATPase (**Figure 3**).

PKC phosphorylates the catalytic α1 subunit at Ser11 and 18 (rat sequence) at the N-terminus of the protein that forms the actuator domain. Ser18 that is considered as a major phosphorylation site is available for phosphorylation in the E2 conformation (Feschenko and Sweadner, 1997). In the E1 conformation the N-terminus is translocated approaching the small cytosolic loop that shields the phosphorylation site from interaction with the kinase (Feschenko et al., 1997; Segall et al., 2003). The analogs of Ser18 and Ser11 are missing in the α2 isoform of the catalytic subunit (compare sequences A24639 for α1 and B24639 for α2 subunit in PubMed protein sequence library http://www.ncbi.nlm.nih.gov) making this isozyme PKCinsensitive. Phosphorylation efficiency may also vary depending on the PKC isoform. Conventional isoforms are more efficient in phosphorylation of the α1 isoform of the pump isolated from rat retinal cells than the novel isoforms δ and ε (Kazanietz et al., 2001). Phosphorylation of the α subunit by PKC may have stimulatory of inhibitory effects on the Na,K-ATPase depending on the cell type (Therien and Blostein, 2000). It is suggested that phosphorylation modulates interaction of the enzyme with other proteins or trigger its internalization (Feschenko and Sweadner, 1997). For example, activation of the atypical isoforms of PKC ξ triggers internalization of the Na,K-ATPase in alveolar epithelial cells under hypoxic conditions (Dada and Sznajder, 2003). This effect is initiated by the free radical burst in the mitochondria that is followed by the activation by the 5′ - AMP-activated protein kinase (AMPK) that is phosphorylated at Thr172. The activated AMPK phosphorylates PKCξ at Thr410 and its translocation to the plasma membrane (Gusarova et al., 2009). Upon translocation PKC phosphorylates Na,K-ATPase at the Ser18 residue in the N-terminus of the α1 subunit. Endocytosis of the α1-containing isozyme of the ATPase is precluded by ubiquitination of the lysine residues next to the Ser18 (Dada et al., 2007; Lecuona et al., 2007). Ubiquitination makes the enzyme recognizable for the mu2 subunit of the adaptor protein that binds to the Tyr527 of the α1 subunit initiating its endocytosis in clathrin-coated vesicle is initiated with the following degradation of the Na,K-ATPase (Chen et al., 2006; Lecuona et al., 2007).

PLM is one more target of phosphorylation by PKCǫ at Ser36 and Ser 68. Phosphorylation it stimulated by Ca2+ and is NOdependent (Pavlovic et al., 2013). Ischemia was associated with facilitation of phosphorylation of PLM, its de-attachment from the αβ complex and increase in the Na,K-ATPase hydrolytic activity in sarcoplasmic membrane fraction (Fuller et al., 2003, 2009).

# Protein Kinase G (PKG)

Hypoxia and reoxygenation result in the alterations in NO, CO, and H2S production (see above). All gasotransmitters are capable of activation of sGC that in turn triggers activation of cGMPdependent protein kinase PKC (PKG) (Therien and Blostein, 2000; Chen et al., 2015). Similar to that for PKC, phosphorylation by PKG was reported to have diverse effects on the Na,K-ATPase activity (Therien and Blostein, 2000). It remains unclear if PKG can directly access the phosphorylation site within the α subunit, as it could only be phosphorylated in the presence of detergents in the purified protein preparation (Fotis et al., 1999; Beltowski et al., 2003). However, treatment of yolk-free homogenates of Xenopus oocytes with cGMP results in phosphorylation and activation of the Na,K-ATPase in the absence of detergents (Fotis et al., 1999). It may reflect the indirect action of PKG on the ATPase via the suppression of dephosphorylation. Yet one more report on the activation of the α2/3 isozymes of the Na,K-ATPase in the central nervous system involves activation of cGMP-PKG pathway following the stimulation with glutamate (Munhoz et al., 2005; Scavone et al., 2005). On the other hand, supplementation of NO donors was shown to inhibit the Na,K-ATPase in nonpigmented epithelial cells of porcine eyes that is associated with activation of PKG (Shahidullah and Delamere, 2006). This effect involves PKG-driven activation of src-family kinases (Shahidullah et al., 2014).

Apart of the alterations in the enzyme activity, stimulation of the cGMP-PKG pathway may affect binding of cardiotonic steroids to the Na,K-ATPase. Increases the sensitivity of the renotubular Na,K-ATPase to marinobufagenin triggered by atrial natriuretic peptide is mediated by the cGMP-PKG signaling cascade (Fedorova et al., 2012).

#### Protein Kinase A (PKA)

Cyclic AMP-sensitive protein kinase (PKA) is a redoxsensitive enzyme containing regulatory thiols (Brennan et al., 2006). In the absence of cAMP the enzyme exists in the inactive tetrameric state and the regulatory Cys199 within the C subunits is inaccessible for regulatory S-glutathionylation (Humphries et al., 2002). Binding of glutathione to this cysteine residue inactivates the kinase and enhances dephosphorylation (Humphries et al., 2005). Vector of the changes in activity of PKA in response to deoxygenation is very much dependent on the cell type and on the activity of G-protein coupled receptors signaling via cAMP-PKA transduction pathway (Jiang et al., 2011).

PKA phosphorylates the catalytic α subunit at Ser938 (rat α1 sequence) within the cytosolic M8-M9 loop. This loop may interact with the M10, C-terminus and the third Na+ binding site. In line with that phosphorylation at Ser938 decreases affinity of this Na+ binding site to Na+ and thereby suppresses the enzyme function (Einholm et al., 2016).

In the purified Na,K-ATPase protein preparation this target is accessible for phosphorylation in the presence of detergent (Feschenko and Sweadner, 1994; Lutz et al., 1996). In intact COS cells phosphorylation of the α subunit at Ser943 is triggered by β-adrenergic stimulation with the following inhibition of hydrolytic and transport activity of the Na,K-ATPase (Cheng et al., 1997).

Activation of cAMP-PKA-dependent pathways in NRK-52E and L6 cell lines may suppress activity of PKC and reduce phosphorylation of the Na,K-ATPase at the PKC binding sites (Feschenko et al., 2000). This cross-talk may result from the close proximity of the PKG binding site to that of PKC (Kruger et al., 2003).

Along with PKC, PKA may catalyze phosphorylation of PLM at Ser68 (Fuller et al., 2004, 2009) in response to ischemia or β adrenergic stimulation (Despa et al., 2005) causing release of the inhibitory action of association of the FXYD1 subunit with the Na,K-ATPase.

# REGULATION OF THE NA,K-ATPASE ACTIVITY BY CARDIOTONIC STEROIDS

Cardenolides and bufadienolides are the two classes of endogenous cardiotonic steroids that serve as hormones selectively interacting with Na,K-ATPase. These compounds are produced by midbrain and adrenocortical cells, and released into the circulation in sub-micromolar concentrations in response to various stimuli such as angiotensin II, acetylcholine, vasopressin, catecholamines, and hypoxic exposure (Bagrov et al., 2009). Interaction of these very low doses of endogenous inhibitors with the Na,K-ATPase does not compromise the transmembrane Na+ gradients, but induces activation of Src kinase and formation of protein complex in which Src kinase and Na,K-ATPase associate with epidermal growth factor receptor (EGRF) and initiate several signaling cascades (Li and Xie, 2009). Signaling modalities depend on the type of cardiotonic steroid and its dose (Dvela et al., 2007). At the molecular level these differences in physiological action of cardenolide ouabain and bufadienolide marinobufagenin are reflected by the specific pattern of conformational changes unique for each steroid upon binding to the purified Na,K-ATPase enzyme (Klimanova et al., 2015). Affinity of the Na,K-ATPase to ouabain is maximal in E2P conformation (in this conformation binding constant for ouabain exceeds that for marinobufagenin by 17-fold), whereas marinobufagenin does not discriminate between the E1 and E2 conformations when binding to the enzyme (Klimanova et al., 2015).

Notably, substitution of Cys 244 by Ala makes cells ouabainintolerant (Shi et al., 2000). This cysteine is a target of regulatory S-glutathionylation (Petrushanko et al., 2012). Furthermore, binding of cardiotonic steroids fixes the enzyme in distinct conformation depending on the type of cardiotonic steroids (Klimanova et al., 2015). Thus, binding of cardiotonic steroids may altersusceptibility of cysteine residues to S-glutathionylation and, vice versa, reversible thiol modifications may possibly alter sensitivity of the enzyme to cardiotonic steroids. This hypothesis awaits further investigations.

Local (renal) or systemic reduction in oxygen availability was shown to trigger release of cardiotonic steroids (bufadienolides or endogenous ouabain) into the circulation in rodents and humans (Zhao et al., 1995; Bagrov et al., 1998; De Angelis and Haupert, 1998; Tian et al., 2010). Apart of the induction of cytoprotective signaling pathway, endocytosis of Na,K-ATPase will support reduction of ATP consumption by hypoxic tissue (De Angelis and Haupert, 1998). Protective effect of distinct cardiotonic steroids (low doses oleandrin for the brain and ouabain or the heart) was reported for the brain and heart exposed to ischemia-reperfusion (Pasdois et al., 2007; Van Kanegan et al., 2014). Binding of ouabain in low doses to the ATPase protects from toxicity of other cardiotonic steroids (Nesher et al., 2010). Whereas micormolar doses of cardiotonic steroids trigger Ca2+ overload and promote apoptosis (Winnicka et al., 2007), nanomolar doses of some of these compounds stimulate proliferation (Winnicka et al., 2010). This anti-apoptotic action may be reflect the changes in affinity of the Na,K-ATPase-ouabain complex to other proteins interacting with the Na,K-ATPase such as PKC, BAX, and Bcl-2 (Lauf et al., 2015). Release of them from the enzyme into the cytosol protects the cells from apoptosis.

On the other hand endogenous ouabain and marinobufagenin act as powerful vasoconstrictors actively participating in development of hypertension (Bagrov and Fedorova, 1998), and have a potential to cause Ca2+ overload secondary to the inhibition of the Na+ extrusion from hypoxic tissue (Schwinger et al., 1999). At the moment, we clearly know too little about the mechanisms of cytoprotection by cardiotonic steroids to use them effectively as therapeutic agents avoiding deadly side-effects (Washam et al., 2015).

#### THE ROLE OF ACUTE REGULATION OF THE NA,K-ATPase IN ADAPTATION OR IRREPARABLE DAMAGE AT LOW O2 LEVELS

As shown above, the capacity of the Na,K-ATPase to sense and respond to the changes in oxygen availability is immensely diverse with multiple signaling cascades implicated in fine-tuning of the enzyme's activity apart and before terminal ATP depletion is reached. The choice of signaling mechanism as well as the resulting vector and amplitude of the change in the Na,K-ATPase activity depends on the cell type, severity and duration of hypoxia, and is rather species-specific. Strictly speaking, we cannot refer to Na,K-ATPase as the "oxygen-sensitive" protein as all the processes involved in its responding to hypoxia are mediated by secondary products of O2 transformation to either gaseous messengers or products of O2 reduction.

Several types of response to hypoxia include (i) inhibition by thiol modifications (ii) modulation of the enzyme function/availability at the membrane due to the changes in phosphorylation, and (iii) interaction with nanomolar doses of endogenous cardiotonic steroids.

Diversity of regulatory pathways allows for fine dosedependent regulation of the enzyme within minutes after the alteration in O2 availability. Based on the current knowledge the following scheme of responses may be suggested (**Figure 4**). Following gradual deoxygenation NO availability in cells expressing NOS1 and NOS2 becomes limited whereas O•− 2 production by NADPH oxidases is maintained while pO2 is above 2–3 kPa (Kd for NOXes ∼ 2 kPa) (Pacher et al., 2007). This is not the case for tissues in which NOS3 isoform of NO synthase with high affinity for O2 prevails (such as vascular endothelial cells). These conditions most likely favor S-glutathionylation of β subunit that is associated with activation of superoxide production by NOXes (White et al., 2009) and α subunits of the Na,K-ATPase due to the gradual accumulation of GSSG (Petrushanko et al., 2012). Further decrease in oxygenation to 1–2 kPa supports uncoupling of electron transfer and triggers mitochondrial O•− 2 production and accumulation of H2O2 in the cytosol. Oxidation of GSSG becomes more pronounced and complete inactivation of the enzyme may be achieved as soon as ATP levels decrease below 50 µM (Petrushanko et al., 2012). Further decrease in O2 to 0.2 kPa reduces S-glutathionylation and renders regulatory thiols within the α subunit oxidized to sulfinic and sulfonic acid making the enzyme insensitive to the changes in GSSG and ONOO− (Petrushanko et al., 2015). Phosphorylation may support or reduce the probability of complete reversible inactivation by S-glutathionylation of the regulatory cysteines by shifting the equilibrium between the E1 and E2 conformation of the enzyme (see Section Oxygen-Sensitive Phosphorylation).

Survival under hypoxic conditions implies that Na+ uptake particularly high in excitable tissues is balanced with equally efficient extrusion of Na+ by the pump (Hylland et al., 1997; Nilsson, 2001). Inhibition of the Na,K-ATPase is deadly when not synchronized with the closing of the cation channels and cessation of activity of the animal (channel arrest) (Buck and Hochachka, 1993; Hochachka et al., 1996; Hylland et al., 1997; Silver et al., 1997; Nilsson, 2001; Ross et al., 2006; Wilkie et al., 2008; Dave et al., 2009). If the Na,K-ATPase is suppressed and the channels are not "arrested," survival is limited to the time of Na+/K+ gradients dissipation in the brain and in the heart (Hylland et al., 1996, 1997; Silver

et al., 1997; Nilsson, 2001). It is tempting to assume that the alterations in free radical production and reversible thiol modifications are involved in the simultaneous regulation of multiple ion transporting systems coupling their activity to the mitochondrial function and ATP availability. However, data providing direct support for this hypothesis are currently missing.

The pilot studies suggest that preservation of redox state in hypoxic heart is sufficient to maintain the Na,K-ATPase activity in Spalax mole rate and trout supporting activity of both species under conditions of critical O2 shortage for at least 20 min (Yakushev et al., 2012). S-nitrosylation of the regulatory cysteine residues in the α subunit may also prevent S-glutathionylation and the enzyme inactivation by GSSG in myocardial membranes (Petrushanko et al., 2012). Whether this is the case for some or all anoxia-tolerant species (e.g., Hylland et al., 1997; Nilsson, 2001; Ross et al., 2006; Dave et al., 2009), remains to be clarified.

Maintenance of the transmembrane Na+ gradients does not only support neuronal function and heart contractility. It also regulates intracellular Ca2+ by controlling the activity of Na/Ca exchanger and that of voltage-gated Ca2+ channels. Ca2+ transport pathways are by themselves targets regulated by reversible thiol modifications in hypoxic cells (Lehotsky et al., 2002; Wang and Zheng, 2010).

Recent data revealed the existence of intimate link between the activity of the Na,K-ATPase and the gene expression in hypoxic cells and tissues. Two factors that were recently suggested to impact the gene expression under hypoxic conditions include the shift in transmembrane Na/K gradients (Koltsova et al., 2014) and the alteration in HIF1α levels upon binding of the nanomolar concentrations of cardiotonic steroids to their binding site within the Na,K-ATPase (Zhang et al., 2008; Cao et al., 2014). Decrease in HIF1α in hypoxic cells (1% O2) in the presence of <50 nM ouabain is not caused by its degradation but rather by a drop in protein synthesis (Zhang et al., 2008), most likely at the level of translation (Cao et al., 2014). Alterations of HIF1α availability by cardiac glycosides contributes to the modulation of long-term hypoxic responses of the organism. For example digoxin treatment prevents remodeling of pulmonary vasculature underlying development of hypoxic pulmonary hypertension (Abud et al., 2012). It is known that signaling cascades initiated by binding of low doses of cardiotonic steroids to the Na,K-ATPase include modulation of intracellular Ca2+ levels and free radical production as well as activation of several kinases. Molecular mechanisms linking these processes to the regulation of HIF1α translation remain unclear.

#### AUTHOR CONTRIBUTIONS

AB is the author the general schematics of review, she contributed to all the sections and assembling of the review. AM, PH were contributing to the sections Introduction. Oxygen, and Oxygen Sensing from Evolutionary and Modern Perspectives, Versatility of oxygen sensing. Multiple Signals-Multiple Targets—Multiple Responses—Multiple Outcomes, and Oxygen and Redox-Sensitivity of the Na,K-ATPase. IP contribution were the sections Oxygen and Redox-Sensitivity of the Na,K-ATPase, Coordinated Regulation of Redox-Sensitive Protein Networks by Reversible Thiol, Oxygen-Sensitive Phosphorylation, and Regulation of the Na,K-ATPase Activity by Cardiotonic Steroids. All co-authors discussed the topics

#### REFERENCES

and writing the review. All of them agree with the final text.

#### ACKNOWLEDGMENTS

The review was funded by the grants of Swiss National Science Foundation IZK0Z3\_157269/1 and 310030\_124970/1 to AB and Russian Science Foundation (Grant #14-14-01152) to IP Spanish Government grants (partially funded by the European Union FEDER/EDRF) PI12/00875 and PI15/00107 and a grant from the Fundación Domingo Martínez are supporting AM, PH received a travel grant from the Instituto de Investigación Sanitaria Princesa (to PH), and by COST actions TD0901 (HypoxiaNet) and CM1001. PH is recipient of a pre-doctoral FPU fellowship from the Spanish Government and AM is supported by the I3SNS programme (ISCIII, Spanish Government, partially funded by FEDER/ERDF).

Arch. Biochem. Biophys. 180, 248–257. doi: 10.1016/0003-9861(77)9 0035-2

significance of S-glutathionylation. Antioxid. Redox Signal. 10, 445–473. doi: 10.1089/ars.2007.1716

**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 Bogdanova, Petrushanko, Hernansanz-Agustín and Martínez-Ruiz. 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.

# Effects of Age on Na+,K+-ATPase Expression in Human and Rodent Skeletal Muscle

#### Victoria L. Wyckelsma\* and Michael J. McKenna

Clinical Exercise Science Program, Institute of Sport Exercise and Active Living, Victoria University, Melbourne, VIC, Australia

The maintenance of transmembrane Na+ and K+ concentration gradients and membrane potential is vital for the production of force in skeletal muscle. In aging an inability to maintain ion regulation and membrane potential would have adverse consequences on the capacity for performing repeated muscle contractions, which are critical for everyday activities and functional independence. This short review focusses on the effects of aging on one major and vital component affecting muscle Na+ and K+ concentrations, membrane potential and excitability in skeletal muscle, the Na+,K+-ATPase (Na+,K+-pump, NKA) protein. The review examines the effects of age on NKA in both human and rodent models and highlights a distant lack of research in NKA with aging. In rodents, the muscle NKA measured by [3H]ouabain binding site content, declines with advanced age from peak values in early life. In human skeletal muscle, however, there appears to be no age effect on [3H]ouabain binding site content in physically active older adults between 55 and 76 years compared to those aged between 18 and 30 years of age. Analysis of the NKA isoforms reveal differential changes with age in fiber-types in both rat and humans. The data show considerable disparities, suggesting different regulation of NKA isoforms between rodents and humans. Finally we review the importance of physical activity on NKA content in older humans. Findings suggest that physical activity levels of an individual may have a greater effect on regulating the NKA content in skeletal muscle rather than aging per se, at least up until 80 years of age.

Keywords: age, Na+K +-pump, single fiber, [3H]ouabain

# IMPLICATIONS OF IMPAIRED SKELETAL MUSCLE ION REGULATION IN AGING

A fundamental factor underpinning skeletal muscle contractile function is the maintenance of membrane excitability, which is heavily dependent on transmembrane sodium (Na+), potassium (K+), and chloride (Cl−) gradients and conductance's via their effects on muscle membrane potential (Em) (Hodgkin and Horowicz, 1959). The intramuscular regulation of these ions is important in both the development of and preservation against muscle fatigue (Sejersted and Sjøgaard, 2000; McKenna et al., 2008). Therefore, any disturbances in muscle ion regulation with aging are likely to impact adversely on cellular excitability and the capacity to undertake repeated muscle contractions, thereby affecting the capability to successfully complete simple daily tasks, and thus on quality of life. Preservation of muscle mass and function is critical in older individuals, due their greater risk of falls, and the consequential effects, that include ongoing physical disability,

#### Edited by:

Laura Andrea Dada, Northwestern University, USA

#### Reviewed by:

Lillian DeBruin, Wilfrid Laurier University, Canada Pablo Martín-Vasallo, University of La Laguna, Spain

\*Correspondence: Victoria L. Wyckelsma<EMAIL_ADDRESS>
#### Specialty section:

This article was submitted to Membrane Physiology and Membrane Biophysics, a section of the journal Frontiers in Physiology

> Received: 25 April 2016 Accepted: 13 July 2016 Published: 02 August 2016

#### Citation:

Wyckelsma VL and McKenna MJ (2016) Effects of Age on Na+,K+-ATPase Expression in Human and Rodent Skeletal Muscle. Front. Physiol. 7:316. doi: 10.3389/fphys.2016.00316 declines in physical and mental health and in social isolation for the individual (Stel et al., 2004). This short review focusses on the effects of aging on one major component of ion regulation in skeletal muscle, the Na+,K+-ATPase (Na+,K+-pump, NKA) protein in skeletal muscle, which is vital for the regulation of transmembrane Na+ and K+ concentration gradients, Em and excitability in skeletal muscle cells (Clausen, 2003b). This review does not cover the acute activation and regulation of the NKA, for this we direct readers to excellent reviews (Clausen, 2003b; Pirkmajer and Chibalin, 2016).

# Na+,K+-ATPase (NKA) IN SKELETAL MUSCLE

In healthy young adults, the skeletal muscle NKA content is typically around 250–350 pmol.g wet weight−1 (Clausen, 2013). Whilst neural tissue contains a higher NKA content per unit mass, skeletal muscle represents the largest pool of NKA in the body due to the very large muscle mass; thus a human weighing 70 kg is estimated to have approximately 8.4 µmol of NKA (Clausen, 2013). The NKA comprises alpha (α) and beta (β) subunits, which together constitute a functional αβ heterodimer (Clausen, 2003b; Green, 2004). The NKA α subunit (∼100–112 kDa) contains binding sites for Na+, K+, and Mg2+ ions as well as phosphate and ATP and typically undergoes both phosphorylation and oxidation (Clausen, 2003b; Lingrel et al., 2003; McKenna et al., 2006). The β subunit (∼35–55 kDa) is glycosylated and is necessary for the structural maturation of the α subunit, localisation of the NKA heterodimer to the sarcolemma and regulation of NKA activity (Cougnon et al., 2002). Each isoform is encoded by separate genes including four α isoforms (α1, α2, α3, α4), and three β isoforms (β1, β2, β3) (Blanco and Mercer, 1998). The expression of these isoforms differs across tissues, which suggests a varying function and regulation of each isoform (McDonough et al., 2002). An additional regulatory (γ) subunit associated with NKA activity, known as the FXYD family of proteins comprising seven isoforms (Geering, 2005); the predominant FXYD isoform in skeletal muscle is FXYD1, or phospholemman (Bibert et al., 2008).

To understand aging effects on NKA in muscle, it is first important to understand distributions of NKA isoforms in specific fiber types. In rat skeletal muscle, based on experiments using isoform-specific antibodies, most studies reveal that the NKA α1 isoform has a similar abundance in oxidative and glycolytic muscles (Hundal et al., 1993; Thompson and McDonough, 1996; Ng et al., 2003; Fowles et al., 2004; Zhang et al., 2006; Kristensen and Juel, 2010; Ingwersen et al., 2011). Similarly, the α2 is abundant in both oxidative and glycolytic muscles in the rat (Thompson and McDonough, 1996; Fowles et al., 2004; Kristensen and Juel, 2010; Ingwersen et al., 2011). In contrast, rat skeletal muscle exhibits distinct differences in β1 and β2 expression between fiber types, with an almost exclusive expression of β1 in muscles rich in slow twitch fibers, whereas the β2 is more abundant in fast twitch fibers (Hundal et al., 1993; Thompson and McDonough, 1996; Fowles et al., 2004; Zhang et al., 2006). The β3 isoform in rat muscle was found to be similarly abundant in red and white gastrocnemius muscles (Ng et al., 2003).

In human skeletal muscle, all isoforms except the α4 isoform are expressed (Murphy et al., 2004) and recent studies indicate that muscle fiber type NKA expression differs considerably from that in the rat. In human muscle, an initial study found no fibertype specific differences abundance for the α1, β1, or FXYD1, but revealed a greater abundance of α2 in Type II fibers, with no measures of the other NKA isoforms (Thomassen et al., 2013). Two subsequent studies from our group reported no fiber-type specific abundance in the α1, α2, and β1 isoforms, but a greater abundance of the α3 and β2 in Type II fibers (Wyckelsma et al., 2015, 2016). The FXYD1 protein was not measured in these latter studies.

The differing abundances of the NKA isoforms between fibertypes in rat muscle and to a lesser extent in human muscle, raises the important issue of understanding what the functional roles of different isoforms are in skeletal muscle. However, our understanding of the specific functions of the NKA isoforms within skeletal muscle is incomplete. Several genetically modified mouse models have been developed to address this, including mice with either one copy of either the α1 (α +/− 1 ) or α2 (α +/− 2 ) gene knocked out, leaving the mice with one-half of either the α1 or α2 isoform compared to wild-type (WT) mice (He et al., 2001), or a skeletal muscle-specific α2 gene knock out (skα2 −/−; Radzyukevich et al., 2012). The extensor digitorum longus (EDL) muscles from α +/− 1 mice showed reduced muscle force compared with WT mice, while the α +/− 2 mice had improved force (He et al., 2001). When a fatigue protocol was implemented the α +/− 2 mouse EDL muscle fatigued at a faster rate than the α +/− 1 and WT (He et al., 2001). It was suggested the improved force in basal (i.e., non-fatigued) conditions in α +/− 2 mice could have been explained by altered Ca2+ handling in these mice (He et al., 2001). In skα2 −/− mice, the resting membrane potential of the EDL was not different to WT mice (Radzyukevich et al., 2012); however, the skα2 −/− muscle showed a rapid decline in the maximum twitch and tetanic forces and fatigued earlier during a treadmill running test compared to WT mice (Radzyukevich et al., 2012). These studies now establish, at least in murine skeletal muscle, that the NKA α1 plays an important role in Na+/K+ exchange and Em regulation during basal conditions, presumably in a "house-keeping" role, whilst the α2 isoform plays a key role during muscle contractions. The α3 is the least abundant of the NKA α isoforms in rat skeletal muscle (Blanco and Mercer, 1998), but its role in skeletal muscle has not yet been established. The α3 isoform is the major NKA α isoform expressed in neurones, with NKA α3 mutations linked with dystonia-parkinsonism and alternating hemiplegia of childhood (Heinzen et al., 2014). An investigation with α3 haplo-insufficiency in mice found that while they could compete physically with WT mice, they were slower in cognitive challenges, such as navigating a water maze (Moseley et al., 2007). Thus, further research is required to determine the functional significance of NKA α3 in skeletal muscle.

The overall amount of β isoforms present in a muscle is also important for NKA activity. Muscles expressing NKA α:β isoforms in a ratio of 1:2 had a higher NKA activity than those with a 1:1 ratio, with activity measured by the K+-dependent 3-O- methylfluorescein phosphate activity assay (Lavoie et al., 1997). The alpha-beta heterodimer with a β1 isoform has a higher affinity to Na+ and lower affinity for K+ compared to β2, independent of the α isoform paired with (Crambert et al., 2000). Whilst the expression of β1 and β2 isoforms differs between muscle fiber types in the rat, the specific roles of the different β isoforms in skeletal muscle NKA regulation contractile have not yet been defined.

The NKA in muscle is adaptable with disease and physical activity. In humans, declines in skeletal muscle [3H]ouabain binding site content have been reported with various diseases including muscular dystrophy (Desnuelle et al., 1982), McArdles disease (Haller et al., 1998), and liver cirrhosis (Aagaard et al., 2002), as well as injury and associated inactivity (Leivseth and Reikerås, 1994; Perry et al., 2015). In contrast, physical training increases muscle [3H]ouabain binding site content and is also typically associated with improved exercise performance (McKenna et al., 1996; Clausen, 2013). Given this malleability in muscle NKA and the link with muscular performance, any effects of age on the NKA are of considerable interest.

# AGE ASSOCIATED ALTERATIONS TO NKA WHOLE MUSCLE CONTENT MEASURED VIA [3H]OUABAIN BINDING

The widely accepted method for quantification of the total number of functional NKA in skeletal muscle is through measurement of the [3H]ouabain binding site content (Clausen, 2003a). The procedure is performed on small pieces of whole muscle samples (typically between 10 and 20 mg) and is based on the high affinity binding of cardiac glycosides to the α subunit of the NKA, with a stoichiometry of 1:1 (Hansen, 1984). By incubation of muscle samples in tritiated ouabain and counting of β particles via liquid scintillation, it is possible to quantify the NKA content in molar units, typically expressed as NKA in pmol. g wet weight−1 (Hansen and Clausen, 1988). In human skeletal muscle, the standard [3H]ouabain binding assay detects each of the three α isoforms (Wang et al., 2001) and thus this is a measure of NKA content; thus in the article these terms can be read interchangeably. This contrasts rat muscle, where only the α2 isoform is detected at the concentration of ouabain used, due to its much higher affinity than the other α isoforms (Hansen, 2001), although this is clearly the most dominantly expressed of the α isoforms (Hansen, 2001). Thus, when referring to rat skeletal muscle the [3H]ouabain binding site content is referred rather than the term NKA content which is incorrect.

Few studies have investigated age effects on skeletal muscle [ 3H]ouabain binding site content. From the limited available literature, rodent muscle shows a clear age-dependent effect, although by far the greatest changes are a large upregulation that early occur early in life. In rats, soleus muscle [3H]ouabain binding site content increased four-fold from birth up until 1 year of age, which was then followed by a 50–70% decline over the following 2–20 months (Kjeldsen et al., 1984). In rats, soleus muscle exhibited a 58% decrease in [3H]ouabain binding site content from rats aged between 28 and 85 days (Kjeldsen et al., 1982). Mice showed less prominent changes in muscle [3H]ouabain binding site content than rats. From the first week of life [3H]ouabain binding site content increased from around 300 to 800 pmol.g wet weight−1 by the 4th week of life, this was followed by a 25% decline which plateaued over the subsequent 4 weeks (Kjeldsen et al., 1984).

There are no lifespan time-course studies investigating chronological changes in human muscle NKA. Several studies utilizing a cross-sectional design have compared muscle NKA content in young healthy participants with a mean age of ∼24 years vs. older adults with a mean age of ∼68 years (Klitgaard and Clausen, 1989; McKenna et al., 2012; Wyckelsma et al., 2016). The earliest study reported a non-significant (14% lower) difference in NKA content in older compared to the younger adults (Klitgaard and Clausen, 1989). Two recent studies from our group reported no difference in NKA content between young and older adults (McKenna et al., 2012; Wyckelsma et al., 2016). These studies are therefore consistent in finding no significant difference in [3H]ouabain binding site content between young

population data have not been included in this figure. \*Different to 68 years (no

exercise), + different to young (active), p < 0.05.

and older adults. However, an interesting observation from another recent study was that NKA content was 26% lower in older adults aged between 69 and 81 years compared to those aged from 55 to 68 years (Perry et al., 2013). This suggested that an age associated reduction in muscle NKA content in humans might only be apparent at more advanced ages. To investigate this possible age effect on NKA content further, we analyzed all of the [3H]ouabain binding site content data in aging-related research collected in healthy, older participants in our laboratory over the past 4 years (McKenna et al., 2012; Perry et al., 2013; Wyckelsma et al., 2016). The first comparison revealed no difference in NKA content between younger adults aged 18–30 years (n = 32) compared to those aged from 55 to 76 years (n = 25; 348.3 ± 82.2 vs. 361.8 ± 59.0 pmol.g wet weight−1 , Young vs. Old, respectively, p = 0.53), confirming the above conclusion (**Figure 2**). Comparison between adults after dividing older adults in subgroups based on decades lived, from 18 to 30 (n = 32), 55–59 (n = 3), 60–69 (n = 11), and 70– 76 years of age (n = 11), also revealed no differences with age (**Figure 2**). This suggests that the decline in [3H]ouabain binding reported previously in the oldest age category (Perry et al., 2013) may be related to their chronic physical activity levels rather than their age. Older participants in one of the studies reported similar physical activity levels to the healthy young controls, despite the intensities of activities likely differing considerably between the groups (Wyckelsma et al., 2016). This may suggest that preserving some level of physical activity might be the important factor in the maintenance of skeletal muscle NKA with age in humans. Thus, whilst studies in rat and murine muscle suggest that aging is associated with a moderate decline in [3H]ouabain binding site content, there is no evidence that this occurs in human skeletal muscle. However, since we have no analyses of muscle [3H]ouabain binding in healthy adults greater than 80 years, we cannot exclude the possibility that a decline may occur beyond this age.

## AGE ASSOCIATED ALTERATIONS TO NKA ISOFORMS

Despite the [3H]ouabain binding site content being the gold standard for measuring NKA content in human muscle, this method (using standard concentration of ouabain) cannot differentiate between the individual α isoforms. Hence researchers are unable to identify which of the three specific α isoforms might have changed with a particular intervention, condition or with aging. Given the possibility of differing roles and fiber-type specificity of the individual isoforms, it is therefore necessary to also investigate the isoform abundances in muscle. This has typically been conducted via western blotting, although one study utilized immunohistochemistry (IHC) to analyze differences in NKA isoforms in aged rat muscle (Zhang et al., 2006). The advantages of IHC are that it allows detection of the cellular localization of specific isoforms and can provide an indication of the fiber-type specificity of a protein. However, disadvantages include the inability to quantify the abundance of protein and the problem of non-specific binding of antibodies.

Few studies have investigated the impacts of aging on NKA isoforms in rat muscle, and the use of different ages of rats and analytical techniques (e.g., western blotting vs. IHC) makes direct comparisons between studies difficult; it is therefore not surprising that findings are inconsistent (Sun et al., 1999; Ng et al., 2003; Zhang et al., 2006). Nonetheless, these studies have identified that aging in rats is associated with skeletal muscle NKA isoform changes that comprise an increased α1, either no change or a decline in α2, an increase or no change in β1, decreased β2, and an increases β3, as summarized in **Table 1**. These findings from advanced aged in rodent muscle suggests there may be more NKA α1β1 heterodimers present in aged skeletal muscle (Sun et al., 1999; Ng et al., 2003). The α1β1 has a higher Na+ affinity in resting muscle (Crambert et al., 2000), and it has been hypothesized that aged rat muscle may

FIGURE 2 | Muscle [3H]ouabain binding does not change with age in human skeletal muscle. Muscle [3H]ouabain binding site content collated from data collected on healthy young and healthy older adults from the McKenna research group between 2012 and 2016. (A) Shows the data combined into two discrete age groups and analyzed by unpaired t-test (p = 0.53). (B) Shows all data plotted into relevant decades of life analyzed by one-way ANOVA (p = 0.30); the mean of each group is also shown as a horizontal line.

TABLE 1 | Effects of age on NKA isoform abundances in skeletal muscle.

Reference 1, Sun et al., 1999; 2, Ng et al., 2003; 3, Zhang et al., 2006; 4, McKenna et al., 2012; 5, Wyckelsma et al., 2016.

Age, mean ± SD. Symbols: ↓, denotes decrease; –, no change; ↑, increase, data in parentheses denotes % difference between groups. ↑ \* not quantitative but increased compared to young ↓ \* not quantitative but decreased compared to young. NR not reported, NS not significant.

Muscles: EDL, Extensor Digitorum Longus; RG, Red Gastrocnemius; WG, White Gastrocnemius; VL, Vastus Lateralis.

have increased K+ and Na+ fluxes with aging, due to reduced amount of caveolin-3 located in the transverse tubular system of aged mice (Barrientos et al., 2015). This is consistent with the important housekeeping role of the α1 isoform in Na+/K+ exchange. From the above findings and also the reduction of α2β2 in aged isoforms in rat, it seems reasonable to suggest in aged rats, that the increased abundance of the α1β1 isoforms may be compensatory for these losses. It was also shown in red gastrocnemius muscle, that α1 was less phosphorylated in aged rats (Zhang and Ng, 2007), although the functional implications of this are not yet understood.

Research into the NKA isoform abundances with age in humans is sparse. The initial study by our group investigated NKA isoform abundance in human vastus lateralis muscle homogenates and found a 24% decrease in α2 and 23% decrease in β3 isoform abundances in older compared to young adults, without change in the other NKA isoforms (McKenna et al., 2012). As there was no difference in [3H]ouabain binding site content, it was suggested that the decrease in α2 abundance might reflect an increased fiber membrane density due to the smaller fibers in aged muscle (McKenna et al., 2012). A later study revealed a decrease with age in the abundance of several proteins typically used as housekeeping proteins, including glyceraldehyde-3-phosphate dehydrogenase (GAPDH; Vigelsø et al., 2015). The GAPDH protein was used in the above study to normalize NKA western blots (McKenna et al., 2012). We therefore re-examined the effects of age on NKA isoform abundances in muscle homogenates, but normalized against total protein rather than GAPDH and also introduced a calibration curve for normalization (Murphy and Lamb, 2013; Wyckelsma et al., 2016). With this improved method in our subsequent study, we found no difference in the abundance in the α2 in aged muscle, which was consistent with the lack of difference in [3H]ouabain binding site content. We also observed no differences in other NKA isoforms, apart from a large (∼250%) increase in β3 isoform abundance (Wyckelsma et al., 2016). This suggests that aging may have quite different effects on the abundance of NKA isoforms in rat and human skeletal muscle.

Since human skeletal muscle comprises a mixed muscle fiber type population, analysis of protein changes in a muscle homogenate might mask changes with aging that occur in different muscle fiber types. We therefore also undertook analysis of NKA isoforms from single fiber segments from biopsies Wyckelsma and McKenna Aging and Muscle Na+, K+-ATPase

obtained from elderly humans (Wyckelsma et al., 2016). This study firstly showed a lack of fiber-type specificity for any of the six NKA isoforms, with isoforms similarly abundant in Type I and II fibers (Wyckelsma et al., 2016). When isoform abundance was compared between young and older adults in a given fiber type, a number of differences were identified. Compared to young adults, the NKA α3 and β2 isoforms were both lower by ∼47% and ∼85%, respectively, in Type II fibers of the older adults, whilst there was a ∼71% greater abundance of α1 in Type I fibers in aged muscle compared to young. The NKA β3 was greater in aged muscle in both Type I (∼96%) and II (∼285%) fibers, reflective of β3 changes seen in the whole muscle homogenate (Wyckelsma et al., 2016).

The greater α1 and β3 isoforms, and lesser β2 isoforms in aged human single fiber segments, showed some similarities with rodent studies (see **Table 1**). However, no age-associated decreases in the α2 or β1 have been reported in studies with humans, contrary to some animal studies (Sun et al., 1999). Direct comparison between these studies is difficult due to varying methodological techniques, as well as is the different ages of mice studied. Previous studies utilized mice aged 29 (Ng et al., 2003), and 30 months (Sun et al., 1999; Zhang et al., 2006). Since 13.8 days for a rat is approximately equivalent to 1 year for a human, mice aged 29–30 months would be equivalent to a human aged 75–80 years (Sengupta, 2013). In human physiological research it is quite difficult to recruit older participants of quite advanced age that are in good health, have limited ongoing pharmacological treatments, as well as are willing to undergo invasive procedures, such as muscle sampling. Furthermore, those that do volunteer may already tend to be reasonably active, which might in itself influence expression of NKA α isoforms (Perry et al., 2013).

In human muscle, the increase in the abundance of α1 which was commonly seen in rodent has been observed, but no measures of phosphorylation have been investigated. Interestingly all studies with rodents report decreased β2 in white and red gastrocnemius muscle (Sun et al., 1999; Ng et al., 2003; Zhang et al., 2006) and also with humans in Type II fibers (Wyckelsma et al., 2016). The role of β2 is unknown in skeletal muscle. It is important to explore the role the β2 isoform plays in NKA enzymatic activity and excitability, especially in Type II fibers, since Type II fibers undergo a loss of specific force in aged compared to compared to young adults (Lamboley et al., 2015). Whilst FXYD1 phosphorylation has been measured in human muscle (Thomassen et al., 2013, 2016), and increased with exercise in aged rats (Reis et al., 2005), the effects of aging have not yet been investigated in aged humans.

#### CHRONIC REGULATION OF NKA WITH EXERCISE TRAINING IN THE AGED

In young adults, physical activity is known to upregulate the NKA content in skeletal muscle (McKenna et al., 1996; Clausen, 2003b), but little is known about training effects on muscle NKA in the aged. The study by Klitgaard and Clausen (1989), compared [3H]ouabain binding site content in healthy but sedentary older and younger adults with three cohorts of active older adults, who had undertaken 12–17 years of regular physical training. The active groups participated in either running, swimming or resistance training and had considerably greater [3H]ouabain binding site content than the sedentary older adults (**Figure 1**). Furthermore, it was found that the resistance trained older adults had a greater [3H]ouabain site content compared to untrained young controls (Klitgaard and Clausen, 1989).

In rats, exercise training further increased the α1 isoform, which was already upregulated with age, as measured by western blotting and appeared to reverse the age associated alterations of the NKA β3 isoform (Ng et al., 2003) (**Table 2**). Additional training adaptations included increases in the α2 and β1 in rat EDL, red and white gastrocnemius muscle (Ng et al., 2003). Another study in aged rats found upregulation of FXYD1 in EDL, red and white gastrocnemius muscle with exercise training (Reis et al., 2005) (**Table 2**). There were also tendencies for the co-immunoprecipitation of FXYD1 to the α1 which tended to decrease with training and the α2 tended to increase the co-immunoprecipitation of FXYD1 with training (Reis et al., 2005). However, this study was conducted in only 3 rats, which they suggested may have resulted in a Type II error. The effects of longitudinal training in older adults is yet to be published.

1, Ng et al., 2003; 2, Reis et al., 2005. Blank space indicated not measured, ↑ increased ↓ decreased – no change.

Muscles: EDL, Extensor Digitorum Longus; RG, Red Gastrocnemius; WG, White Gastrocnemius; VL, Vastus Lateralis.

#### PERSPECTIVES

The review highlights the lack of research in aging and NKA regulation in skeletal muscle, with many aspects of NKA regulation still required to be explored. It further demonstrates that changes that might be observed with age in rodent muscles cannot be anticipated to also occur in humans and the reasons for this are unclear. This highlights the importance of conducting studies in human muscle. Such research is particularly important given the vital role of NKA in regulation muscle excitability and function. The use of skeletal muscle knockout mice provide excellent models to determine function of the specific NKA isoforms in skeletal muscle. Future research could consider the effects of aging utilizing these knockout mice to investigate muscle NKA, contractility and function, and including with α3, and β1−3

# REFERENCES

isoform knock out models. Until the roles of the NKA isoforms in skeletal muscle are more fully understood, the functional translation of findings in rodent to human muscle physiology can only be speculative. Further research should in addition explore the acute activation of the skeletal muscle NKA in aged muscle and its regulation, as well as the impacts on muscle Em, excitability and the link to function during repeated contractions. Finally, any future work with NKA regulation in aged human skeletal muscle should include measures in single fiber segments to ensure that any fiber-type specific responses are not masked.

#### AUTHOR CONTRIBUTIONS

VW and MM drafted, edited and approved the final version of the manuscript.

**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 Wyckelsma and McKenna. 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.

# P4-ATPases as Phospholipid Flippases—Structure, Function, and Enigmas

Jens P. Andersen1 , Anna L. Vestergaard1 , Stine A. Mikkelsen1 , Louise S. Mogensen1 , Madhavan Chalat 2 and Robert S. Molday 2 \*

*1 Department of Biomedicine, Aarhus University, Aarhus, Denmark, 2 Department of Biochemistry and Molecular Biology, University of British Columbia, Vancouver, BC, Canada*

P4-ATPases comprise a family of P-type ATPases that actively transport or flip phospholipids across cell membranes. This generates and maintains membrane lipid asymmetry, a property essential for a wide variety of cellular processes such as vesicle budding and trafficking, cell signaling, blood coagulation, apoptosis, bile and cholesterol homeostasis, and neuronal cell survival. Some P4-ATPases transport phosphatidylserine and phosphatidylethanolamine across the plasma membrane or intracellular membranes whereas other P4-ATPases are specific for phosphatidylcholine. The importance of P4-ATPases is highlighted by the finding that genetic defects in two P4-ATPases ATP8A2 and ATP8B1 are associated with severe human disorders. Recent studies have provided insight into how P4-ATPases translocate phospholipids across membranes. P4-ATPases form a phosphorylated intermediate at the aspartate of the P-type ATPase signature sequence, and dephosphorylation is activated by the lipid substrate being flipped from the exoplasmic to the cytoplasmic leaflet similar to the activation of dephosphorylation of Na+/K+-ATPase by exoplasmic K+. How the phospholipid is translocated can be understood in terms of a peripheral hydrophobic gate pathway between transmembrane helices M1, M3, M4, and M6. This pathway, which partially overlaps with the suggested pathway for migration of Ca2+ in the opposite direction in the Ca2+-ATPase, is wider than the latter, thereby accommodating the phospholipid head group. The head group is propelled along against its concentration gradient with the hydrocarbon chains projecting out into the lipid phase by movement of an isoleucine located at the position corresponding to an ion binding glutamate in the Ca2+- and Na+/K+-ATPases. Hence, the P4-ATPase mechanism is quite similar to the mechanism of these ion pumps, where the glutamate translocates the ions by moving like a pump rod. The accessory subunit CDC50 may be located in close association with the exoplasmic entrance of the suggested pathway, and possibly promotes the binding of the lipid substrate. This review focuses on properties of mammalian and yeast P4-ATPases for which most mechanistic insight is available. However, the structure, function and enigmas associated with mammalian and yeast P4-ATPases most likely extend to P4-ATPases of plants and other organisms.

Keywords: P-type ATPases, P4-ATPases, ATP8A2, CDC50, flippases, membrane asymmetry, phospholipid transport

#### Edited by:

*Olga Vagin, University of California, Los Angeles, USA*

#### Reviewed by:

*Mikhail Y. Golovko, University of North Dakota, USA Emilia Lecuona, Northwestern University, USA*

> \*Correspondence: *Robert S. Molday<EMAIL_ADDRESS>
#### Specialty section:

*This article was submitted to Membrane Physiology and Membrane Biophysics, a section of the journal Frontiers in Physiology*

> Received: *21 February 2016* Accepted: *20 June 2016* Published: *08 July 2016*

#### Citation:

*Andersen JP, Vestergaard AL, Mikkelsen SA, Mogensen LS, Chalat M and Molday RS (2016) P4-ATPases as Phospholipid Flippases—Structure, Function, and Enigmas. Front. Physiol. 7:275. doi: 10.3389/fphys.2016.00275*

# INTRODUCTION

Non-random distribution of lipids across the lipid bilayer is a characteristic feature of most eukaryotic cell membranes. This asymmetrical distribution is most evident in the plasma membrane where phosphatidylcholine (PC), sphingomyelin, and glycolipids are enriched on the external or exoplasmic leaflet of the membranes and phosphatidylserine (PS), phosphatidylethanolamine (PE), and phosphatidylinositol (PI) are primarily confined to the cytoplasmic leaflet (Zachowski, 1993; Lenoir et al., 2007). Cholesterol, a major lipid of the plasma membrane, can move freely between the two leaflets, but is likely to be more concentrated on the exoplasmic leaflet due to its affinity for sphingomyelin and glycosphingolipids. Membranes of other subcellular organelles including the trans-Golgi network (TGN), secretory vesicles, and endosomes show a similar asymmetrical distribution of lipids. Membrane lipid asymmetry plays a crucial role in numerous cellular processes that take place at the plasma membrane and intracellular membranes including cell and organelle shape determination and dynamics, vesicle budding and trafficking, membrane protein regulation, membrane stability and impermeability, cell signaling, neurite extension, blood coagulation, apoptosis, fertilization, and bile and cholesterol homeostasis, among others.

The amphipathic nature of phospholipids impedes their transverse movement or flip-flop across the hydrophobic lipid bilayer. The half time for phospholipid flip-flop in protein-free liposomes typically exceeds several hours. However, the movement of lipids across biological membranes is greatly accelerated by membrane proteins generally called flippases (Pomorski and Menon, 2006). Three families of proteins, scramblases, ATP-binding cassette (ABC) transporters, and P4-ATPases, have been implicated in the transverse movement of lipids across membranes. Scramblases are energyindependent, bidirectional transporters that dissipate membrane asymmetry (Williamson, 2015). Ca2+-activated scramblases play an important role in exposing PS on the surface of cells to initiate such cellular processes as blood coagulation and apoptosis. Some scramblases such as the G-protein coupled receptor rhodopsin are constitutively active thereby randomizing transmembrane lipid distribution in specialized membranes such as photoreceptor discs (Menon et al., 2011). ABC transporters and P4-ATPases use the energy from ATP hydrolysis to transport specific lipids across membranes against their concentration gradient, thereby establishing and maintaining lipid asymmetry in biological membranes (Coleman et al., 2013; Lopez-Marques et al., 2014). Most ABC transporters translocate lipids from the cytoplasmic to the exoplasmic leaflet of membranes and are often called floppases. Some mammalian ABC transporters, however, can transport phospholipids in the opposite direction (Quazi and Molday, 2013). P4-ATPases are a subfamily of P-type ATPases that transport specific phospholipids from the exoplasmic to the cytoplasmic leaflet of membranes to generate and maintain membrane lipid asymmetry (Tang et al., 1996; Coleman et al., 2009; Zhou and Graham, 2009).

A number of excellent reviews have focused on the molecular properties of lipid flippases and their role in cellular processes (Sebastian et al., 2012; Coleman et al., 2013; Lopez-Marques et al., 2013, 2014; Hankins et al., 2015; Williamson, 2015; Montigny et al., 2016). In this review we discuss recent studies on the lipid substrate specificity and transport mechanism of P4-ATPases and their role in cell physiology and disease with emphasis on mammalian and yeast P4-ATPases. We also present an overview of recent site-directed mutagenesis and molecular modeling studies that provide insight into the possible pathways used by P4-ATPases to translocate phospholipids across membranes. Finally, we discuss similarities in the structure and transport mechanisms of P4-ATPase lipid pumps and P2-ATPase ion pumps.

# P-TYPE-ATPases

P-type ATPases constitute a family of membrane proteins that utilize the energy from ATP hydrolysis to transport ions and lipids across biological membranes (Palmgren and Nissen, 2011). These pumps are found in prokaryotes, archaea, and eukaryotes and are distinguished from other ATP-dependent transporters by the presence of a conserved aspartic acid residue that undergoes transient phosphorylation during the ATP cycle (Pedersen and Carafoli, 1987). On the basis of phylogenetic analysis, P-type ATPases have been organized into five main classes (P1-P5 ATPases) (Palmgren and Axelsen, 1998). The first three classes have been further divided into subclasses based on sequence similarity and ion transport specificity. P1A-ATPases are only found in some bacteria and transport K+; P1B-ATPases transport heavy metal ions such as Ag+, Zn2+, Cd2+, and Cu2+; P2A ATPases (sarco(endo)plasmic reticulum Ca2+-ATPase (SERCA) and secretory pathway Ca2+-ATPase (SPCA)) and P2B ATPases (plasma membrane Ca2+-ATPase (PMCA)) transport Ca2+ and Mn2+; P2C-ATPases transport monovalent ions and include Na+/K+-ATPases and H+/K+-ATPases; P2D-ATPases are found in fungi and transport Na+; P3A-ATPases transport H+; and P3B ATPases transport Mg2+. P4-ATPases are unique in that they transport or flip phospholipids across membranes. The substrates for P5-ATPases have yet to be determined, but they are predicted to play an important role in the endosomal-lysosomal system and have been implicated in Parkinson's disease (Schultheis et al., 2004; Cohen et al., 2013; De La Hera et al., 2013).

#### P4-ATPases

P4**-**ATPases are only found in eukaryotes. The human genome encodes 14 P4-ATPases that are organized into five classes with each class having multiple members (**Table 1**, **Figure 1A**): Class 1a (ATP8A1, ATP8A2); Class 1b (ATP8B1, ATP8B2, ATP8B3, ATP8B4); Class 2 (ATP9A, ATP9B); Class 5 (ATP10A, ATP10B, ATP10D); and Class 6 (ATP11A, ATP11B, ATP11C) (Paulusma and Elferink, 2010; Van Der Mark et al., 2013). In contrast, yeast (Saccharomyces cerevisiae) contains five P4-ATPases (Drs2p, Neo1p, Dnf1p, Dnf2p, Dnf3P).

All P4-ATPases consist of a large polypeptide with a molecular mass of approximately 120 kDa which is composed of four main domains (**Figure 1B**). Based on amino acid sequence

#### TABLE 1 | Mammalian P4 -ATPases.

alignment (**Figure 2**), P4-ATPases possess three cytoplasmic domains involved in the ATPase catalytic cycle similar to Ca2+-ATPase and Na+,K+-ATPase: the nucleotide or N-domain binds ATP; the phosphorylation or P-domain contains the aspartic acid (D) residue within the conserved DKTGT motif that undergoes transient phosphorylation; and the actuator or A-domain has the DGET (TGES in Ca2+-ATPase and Na+/K+-ATPase) motif that facilitates the dephosphorylation of the phosphorylated intermediate (Lenoir et al., 2009; Coleman et al., 2012). The membrane or M-domain serves as the pathway for translocation of lipid substrates across cell membranes and is predicted to contain 10 transmembrane segments (M1-M10, **Figure 2**) analogous to the Ca2+-ATPase and the Na+/K+- ATPase (Maclennan et al., 1985; Toyoshima et al., 2000; Morth et al., 2007). Relatively short segments join the transmembrane segments on the exoplasmic side of the membrane, whereas large domains and segments connect some of the transmembrane segments on the cytoplasmic side (**Figures 1**, **2**). Recent studies suggest that transmembrane segments M1–M6 form the principal unit for transport of phospholipids across membranes with M7–M10 playing a supporting role (Baldridge and Graham, 2012; Vestergaard et al., 2014). P4-ATPases also contain regulatory domains that modulate the transport activity and targeting domains that participate in the trafficking of the P4-ATPases to their preferred subcellular membranes. These domains are present, at least in part, along the cytoplasmic C-terminal and N-terminal segments of P4-ATPases, but to date are poorly characterized for mammalian P4-ATPases (Takatsu et al., 2011; Zhou et al., 2013).

# CDC50–ACCESSORY SUBUNIT

P4-ATPases with the exception of ATP9A and ATP9B assemble with an accessory or β-subunit to form a heteromeric complex, most likely a heterodimer. The β-subunit belongs to the CDC50/LEM3 family of evolutionary conserved proteins present in all eukaryotes (Katoh and Katoh, 2004; Saito et al., 2004). There are three CDC50 proteins (CDC50A, CDC50B, and CDC50C, also known as TMEM30A, TMEM30B, and TMEM30C) in humans and three in yeast (Cdc50p, Lem3p, Crf1p). Since there are more P4-ATPases (α-subunits) than CDC50 proteins (βsubunits), many P4-ATPases associate with the same CDC50 protein to form the heteromeric complex (Bryde et al., 2010).

The interaction of mammalian P4-ATPases with specific CDC50 proteins has been studied primarily by heterologous protein co-expression and co-immunoprecipitation (Paulusma et al., 2008; Bryde et al., 2010; Van Der Velden et al., 2010b; Coleman and Molday, 2011; Takatsu et al., 2011, 2014; Naito et al., 2015). These studies indicate that most mammalian P4- ATPases use CDC50A as their β-subunit. These include ATP8A1, ATP8A2, ATP11A, ATP11B, ATP11C, ATP10A, ATP10B, and ATP10D. Several P4-ATPases including ATP8B1, ATP8B2, and ATP8B4 can associate with either CDC50A or CDC50B. The physiological consequence of this dual specificity, however, remains to be determined. CDC50C is predominantly expressed in testes (Osada et al., 2007), but to date it has not been shown to associate with any P4-ATPase. Although, heterologous overexpression provides useful information on the interaction of P4-ATPases with specific CDC50 proteins, the presence of such complexes needs to be confirmed in physiologically relevant cells and tissues. In one study ATP8A2 was found to interact with CDC50A in photoreceptors in support of heterologous cell expression and immunoprecipitation studies (Coleman et al., 2009; Coleman and Molday, 2011).

CDC50 proteins are relatively small membrane glycoproteins containing approximately 350 amino acids. They consist of two transmembrane segments joined by a large exoplasmic domain with two or more N-linked oligosaccharide chains that contribute to the stability of the protein (Coleman and Molday, 2011; Garcia-Sanchez et al., 2014; **Figure 1B**). CDC50 proteins typically migrate on SDS gels as a broad band with an approximate molecular mass of 50 kDa due to the highly heterogeneous nature of the oligosaccharide chains. The exoplasmic domain also contains four conserved cysteine residues that stabilize the structure through the formation of two disulfide bonds (Puts et al., 2012). Relatively, short N-terminal and C-terminal segments are exposed on the cytoplasmic side of the membrane.

Heterologous expression studies point to the crucial role of CDC50 in protein folding and formation of a functionally active P4-ATPase complex (Paulusma et al., 2008; Bryde et al., 2010; Van Der Velden et al., 2010b; Coleman and Molday, 2011). In the absence of CDC50 expression, P4-ATPases fail to exit the ER and are incapable of undergoing phosphorylation or phospholipiddependent ATP hydrolysis (Lopez-Marques et al., 2010; Coleman and Molday, 2011; Puts et al., 2012). Likewise, in the absence of P4-ATPase expression, CDC50A is retained in the ER of cells. However, co-expression of P4-ATPases and CDC50 results in a P4-ATPase-CDC50 complex that can exit the ER and function as an active phospholipid flippase (Coleman and Molday, 2011). CDC50 proteins do not play a direct role in the targeting of P4- ATPases to their preferred subcellular location once they exit the ER in multicellular organisms (Lopez-Marques et al., 2010).

Several studies suggest that CDC50 proteins participate in ATP-dependent phospholipid transport. The affinity of yeast Drs2p for Cdc50p fluctuates during the catalytic reaction cycle with the highest affinity observed for Drs2p in the E2P state with a bound phospholipid substrate (Lenoir et al., 2009). Phosphorylation of Drs2p, ATP8B1, and ATP8B2 are dependent on interaction with their CDC50 subunit (Lenoir et al., 2009; Bryde et al., 2010). Studies using mammalian CDC50A/CDC50B chimeric proteins further indicate that both the transmembrane and exoplasmic domains of CDC50A are necessary for the binding CDC50A to ATP8A2 and the formation of an active complex (Coleman and Molday, 2011). The Nterminal cytoplasmic domain of CDC50A also modulates the kinetics of ATP8A2 ATPase activity.

Certain mutations in the transmembrane segments of ATP8A2 decrease glycosylation of CDC50 without significantly affecting the functional properties of the complex and seem to define a groove between M1, M3, and M4 of ATP8A2 as a possible location of CDC50 interactions (Vestergaard et al., 2015; see later text in relation to **Figure 6**). Altogether, the available information indicates that CDC50 accessory proteins contact P4-ATPases at multiple sites to promote correct protein folding, exit of the P4- ATPase complex from the ER, and formation of a functionally active flippase complex. Hence, CDC50 proteins appear to play a role reminiscent of the β-subunits of Na+/K+-ATPases (Puts and Holthuis, 2009). Because E2P is the state, in which the P4-ATPase is supposed to bind its phospholipid substrate from the exoplasmic membrane leaflet (see reaction cycle in **Figure 3B**), the preferential interaction of the CDC50 protein with E2P (Lenoir et al., 2009) is consistent with participation of CDC50 in substrate loading analogous to the role of the β-subunit of Na+/K+-ATPase in K+ binding (Shinoda et al., 2009). Nevertheless, CDC50 and the β-subunit of Na+/K+- ATPase differ with respect to the number of transmembrane segments (two for CDC50 and only one for the Na+/K+-ATPase

#### FIGURE 1 | Continued