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Imaging adenosine triphosphate (ATP).

Abstract. Adenosine triphosphate (ATP) is a universal mediator of metabolism and signaling across unicellular and multicellular species. There is a fundamental interdependence between the dynamics of ATP and the physiology that occurs inside and outside the cell. Characterizing and understanding ATP dynamics provide valuable mechanistic insight into processes that range from neurotransmission to the chemotaxis of immune cells. Therefore, we require the methodology to interrogate both temporal and spatial components of ATP dynamics from the subcellular to the organismal levels in live specimens. Over the last several decades, a number of molecular probes that are specific to ATP have been developed. These probes have been combined with imaging approaches, particularly optical microscopy, to enable qualitative and quantitative detection of this critical molecule. In this review, we survey current examples of technologies available for visualizing ATP in living cells, and identify areas where new tools and approaches are needed to expand our capabilities.

Introduction

Significant efforts have been made over the last several decades to directly visualize adenosine triphosphate (ATP) in living systems. ATP is a molecule at the center of metabolism and signaling both inside and outside the living cell, and its universal importance in biology reaches well beyond its most familiar role as an energy metabolite.

In energy transduction. ATP hydrolysis provides a thermodynamic driving force for cellular chemistry (Westheimer, 1987; Kamerlin et al., 2013). It remains ambiguous why ATP evolved such a central and universal role (Plattner and Verkhratsky, 2016), but it is clear that energy-dependent reactions are critical for numerous cellular processes: from maintenance of neuronal membrane potential (Magistretti and Allaman, 2015), to organelle transport (Zala et al., 2013), to nucleocytosolic translocation (Dzeja et al., 2002). In fact, hydrolysis of other nucleotides, such as guanosine triphosphate (GTP) and other phosphate metabolites, can also provide a driving force for specific, energy-dependent processes. Despite the energetic contribution of these alternatives, ATP still plays a central role through the action of nucleotide diphosphate kinases and enzymes that create a phosphotransfer network involved in the distribution of bioenergetics (Shugar, 1996; Dzeja and Terzic, 2003).

Beyond energy transduction, ATP plays a central role in signaling. ATP serves as a phosphate-group donor for substrate activation in metabolic reactions and as the coenzyme for a large number of kinases. In the regulation of protein function. ATP can serve as both a phosphate- and an adenylyl-group donor for post-translational modifications. ATP is also required for the biosynthesis of cyclic adenosine monophosphate (cAMP), which is a critical second messenger in signal transduction. Furthermore. ATP itself can serve as a bona fide signaling ligand for ATP-sensitive or purinergic ionotropic and G-protein coupled receptors. With ATP's vast repertoire of biological functions, its spatial and temporal dynamics broadly affect both intracellular and extracellular processes that determine normal physiology, as well as pathological states. We briefly review examples demonstrating the importance of ATP in brain energy metabolism and inflammation, and survey a number of ATP-detection and imaging methods that are currently available.

Intracellular ATP in metabolism and signaling: brain energy metabolism

In the healthy brain. ATP-consuming processes that are involved in electrochemical signaling impose significant energy demands, which are reflected in the high rates of cerebral glucose and oxygen consumption (Harris et al., 2012; Howarth et al., 2012; Magistretti and Allaman, 2015). The dynamics of brain energy metabolism are complex, in part because ATP production must respond to these energy demands in an activity-dependent manner. For example, neuronal excitability relies on sufficient energy production to maintain neuronal ion gradients and membrane potentials in the face of continuous action potential generation and signaling in the brain. ATP production also must support ancillary processes associated with neurotransmission, such as biosynthesis of neurotransmitters, vesicle loading, and axonal transport, to name just a few (Howarth et al., 2012). Many open questions remain concerning the precise mechanisms by which fuel utilization and ATP production are coordinated in the brain, both within cells and between cells. For example, while ATP production is typically viewed as requiring efficient mitochondrial respiration in neurons, aerobic glycolysis occurs under physiological conditions in certain regions of the brain, such as the parietal cortex and prefrontal cortex, especially during development (Vaishnavi et al., 2010; Goyal et al., 2014). Observation of regional aerobic glycolysis may be related to the compartmentalization of metabolism between neurons and glia. It has been hypothesized that astrocytes support neuronal activity by performing aerobic glycolysis and shuttling lactate to neurons for mitochondrial respiration and ATP generation (Belanger et al., 2011). Furthermore, the complex morphology of dendritic arbors and elongated axons requires local ATP generation for synaptic function, which may be provided by motile glycolytic machinery or mitochondria (Zala et al., 2013; Rangaraju et al., 2014).

Because of the high numbers and activities of ATP-consuming processes in the brain, bioenergetics fundamentally influence cognition and behavior. As a result, the physiology underlying activity-dependent ATP production is closely linked to--and ultimately dictates--the utility of methods such as [.sup.18]F-fluorodeoxyglucose positron emission tomography, magnetic resonance imaging (MRI) of cerebral metabolic rates, and blood oxygen level-dependent, functional MRI of cerebral blood flow (Raichle and Mintun. 2006). These methods have, in turn, demonstrated the vulnerability of the brain to metabolic insult. Bioenergetic deficits occur in aging, injury, and neurological diseases such as Alzheimer's and Parkinson's diseases (Johnson et al., 2012; Surmeier et al., 2012). In brain ischemia and stroke, for example, attenuated cerebral blood flow causes loss of glucose and oxygen availability, intracellular ATP depletion, and collapse of ion gradients within minutes (Dreier and Reiffurth, 2015). In addition, ischemic injury results in the release of ATP to the pericellular space, initiating an extracellular cascade of events, including purinergic signaling, chemotaxis of immune cells, and inflammation (Pedata et al., 2015).

Extracellular ATP in metabolism and signaling: inflammation

Leakage of intracellular ATP into the pericellular space can result from disruption of the plasma membrane, which may be precipitated by mechanical damage or necrotic cell death. In addition to this unregulated release, cells can promote ATP secretion through expression of protein channels that connect the cytoplasm to the extracellular space, as has been observed in cells undergoing apoptosis and in activated inflammatory immune cells (Fig. 1). In both cases, the rapid egress of ATP is driven by the gradient from high intracellular ATP concentration to low extracellular ATP concentration. If extracellular ATP levels become significantly elevated, activation of membrane-bound nucleotide receptors on neighboring cells incites a strong proinflammatory response (Idzko et al., 2014). In this context, extracellular ATP acts as a damage-associated molecular pattern (DAMP) molecule and is interpreted by immune cells as a danger signal (Eltzschig et al., 2012).

The receptors activated by ATP (Fig. 1), which are broadly expressed on immune and other cells, are members of the P2 class of receptors for extracellular nucleotides and are classified into two families, P2Y and P2X (Burnstock and Boeynaems, 2014). The P2Y family of receptors are G-protein-coupled receptors (GPCRs), eight members of which have been identified to date. P2X receptors are ligand-gated ion channels. The P2 receptors vary in their responsiveness to a range of nucleotides; P2Y receptors are optimally responsive to endogenous nucleotides such as ATP, adenosine diphosphate (ADP). uridine-5'-triphosphate (UTP), among others, depending on the receptor subtype and species. On the other hand, ATP is the preferred endogenous agonist for all subtypes of P2X receptors.

The P2[X.sub.7] receptor (P2[X.sub.7]R) is the most studied member of the ionotropic P2X receptor family. Activation of P2[X.sub.7]R by ATP is an important proinflammatory signal for a number of immune cells, including macrophages, microglia, and dendritic cells (Di Virgilio, 2007). ATP binding to P2[X.sub.7]R causes channel opening and membrane depolarization via sodium ([Na.sup.+]) and calcium ion ([Ca.sup.2+] ) influx, and potassium ion ([K.sup.+]) efflux. P2[X.sub.7]R activates the NLRP3 inflammasome, which is a multiprotein assembly that enables the caspase-1 catalyzed release of the proinflammatory cytokines, Interleukin-1b (IL-1b) and IL-18. In addition, the P2[X.sub.7] receptor undergoes a conformational change that widens the channel to allow passage of larger molecules, including ATP. However, the mechanism and role of channel widening are debated, and it is not necessary for NLRP3 activation; a decrease in intracellular potassium is necessary and sufficient (Munoz-Planillo et al., 2013). P2[X.sub.7]R activation and the resulting IL-1b signaling have been shown to be important in bacterial pathogen clearance by macrophages (Coutinho-Silva and Ojcius, 2012) and in cancer antigen presentation to lymphocytes by dendritic cells (Ghiringhelli et al., 2009). However, when dysregulated or activated inappropriately, this pathway contributes to a number of inflammatory disorders, including inflammatory bowel disease (Kurashima et al., 2012) and asthma (Miiller et al., 2011).

From the metabotropic P2Y receptor family, P2[Y.sub.2]R is an important mediator of inflammation, and is active in the recruitment of immune cells to sites of infection and wound healing (Elliott et al., 2009). ATP released from damaged or infected cells that bind P2[Y.sub.2]R on leukocytes acts as a chemoattractant to draw in neutrophils and macrophages, which can then phagocytose the ailing cells and help resolve the inflammation. Interestingly, neutrophils use directional., autocrine ATP signaling as a mechanism to amplify their migration response to chemoattractants. Specifically, Chen and coworkers (2006) reported that stimulation with the chemoattractant. N-formyl-Met-Leu-Phe (FMLP), causes human neutrophils to release ATP that is localized near the stimulus. The released ATP then binds P2[Y.sub.2]Rs expressed on the neutrophil cell surface and promotes chemotaxis along the FMLP gradient. Neutrophils from mice deficient in P2[Y.sub.2]R showed significantly impaired chemotaxis.

The proinflammatory signaling of extracellular ATP is terminated by nucleotidases active in the interstitial fluid and the membrane-bound ectonucleotidases, CD39 and CD73 (Fig. 1) (Idzko et al., 2014). CD39 catalyzes the conversion of ATP first to ADP and then to AMP; it can also accept ADP as a substrate for conversion to AMP. Subsequently, CD73 converts AMP to adenosine, and thus a decrease in extracellular ATP results in an increase in extracellular adenosine. Extracellular adenosine can engage PI receptors, of which there are four subtypes: [A.sub.1], [A.sub.2A], [A.sub.2B], and [A.sub.3]. Whereas ATP signaling generally upregulates the immune response, signaling at adenosine receptors is associated with the dampening and resolution of inflammation. Adenosine signaling is terminated by cellular uptake of adenosine through equilibrative nucleoside transporters 1 and 2 (ENT1, ENT2). Following cellular uptake, adenosine is metabolized to either AMP by adenosine kinase or to inosine by adenosine deaminase.

Examples from brain energetics and inflammation illustrate why the "how, when, and where" of ATP production and consumption clearly impact physiology. These examples also highlight the importance of studying the spatial and temporal dynamics of ATP. Because ATP levels can respond rapidly and locally to physiological changes, realtime imaging in live specimens is essential for 1) fully capturing how changes in ATP levels are distributed throughout cells and tissues, and 2) providing experimental access to dissect the mechanisms and machinery responsible for the roles of ATP in metabolism and signaling.

Methods for Detection and Imaging of ATP

A number of well-established and newly developed methods can measure ATP, though there are far fewer methods that can image ATP in live specimens. Chemical and physical methods ranging from liquid chromatography to mass spectrometry (Khlyntseva et al., 2009) can offer superb specificity. For example, extracellular ATP has been measured using enzyme-coated platinum microelectrodes (Kueng et al., 2004; Llaudet et al., 2005) or optical fibers (Wang et al., 2013). However, these methods typically lack spatiotemporal resolution or compatibility with live specimens. Chemical and physical methods that involve deproteinization to measure intracellular concentrations also release bound forms of adenine nucleotides (Harris et al., 1973). This type of sample preparation results in measurements of total rather than free ATP, and A DP concentrations that may skew estimates of equilibrium and energy status (Veech et al., 1979; Morikofer-Zwez and Walter, 1989; Tantama et al., 2013). As an alternative to instrumental analysis, molecular probes specific to free ATP offer the opportunity for real-time analysis, with greater resolution or compatibility with live specimens or both. Thus, we present examples of both non-imaging and imaging methods for detecting ATP in order to contrast their advantages.

A number of non-imaging approaches to ATP measurement have been developed. For example, in an electrophysiological approach, the ligand sensitivity of P2X receptors can be exploited because, as discussed, they are ATP-gated cation channels and thus produce ATP-dependent currents. When P2X receptors are expressed in the plasma membrane of PC12 cells (Praetorius and Leipziger, 2009) or HEK-293 cells (Hayashi et al., 2004), patch-clamp electrophysiology in outside-out or whole-cell mode can be used to detect ATP in proximity to the electrode-immobilized membrane or cell. This technique has been used to study ATP release from a variety of cells; for example, Hazama et al. (1998) measured ATP concentrations released from pancreatic [beta]-cells in real time by measuring P2X current amplitudes via whole-cell patch-clamp. Also using an electrophysiological approach, nucleotide-calibrated measurements of ATP-sensitive potassium ([K.sub.ATP]) channel currents have been employed to calculate submembrane concentrations of ATP and to test for local gradients in COSm6 monkey kidney cells and in Xenopus oocytes (Gribble et al., 2000). In a contrasting, non-imaging spectroscopic approach. Vancraenenbroeck and Webb recently published a sensor based on malonyl-coenzyme A synthetase, an enzyme that undergoes a conformational change upon ATP binding. Two molecules of tetramethylrhodamine are conjugated to the synthetase in a manner such that ATP binding causes a substantial increase in fluorescence intensity, with high selectivity for ATP and micromolar sensitivity (Vancraenenbroeck and Webb. 2015). These and other non-imaging methods provide important means of quantifying ATP and. in many cases, are ideal for assaying cells or extracts with high sensitivity and high throughput. However, non-imaging methods cannot provide the same high level of spatial and time-resolved information content achieved with imaging methods.

Several methods of imaging ATP have been developed over the last several decades (Fig. 2) (Table 1). While the ideal approach, in live specimens, would be quantitative and compatible with non-invasive, probe-free visualization of ATP, with high spatial and temporal resolution, a single method that satisfies these criteria does not yet exist. Magnetic resonance spectroscopy techniques that exploit phosphorous magnetization transfer can non-invasively quantify ATP in vivo, but practical considerations, such as long acquisition times per voxel and the limited availability of the instrumentation, have restricted its applicability to imaging (Du et al., 2008; Chaumeil et al., 2009; Befroy et al., 2012). Instead, methods of visualizing ATP rely primarily on optical microscopy paired with molecular probes, which provide signal readouts specific to ATP. These molecular probes have been developed using a variety of physical formats, from small organic indicators to nanoparticles, and they use both indirect and direct ATP detection mechanisms.

Magnesium Green, for example, is a magnesium-sensitive, small, organic fluorophore that can be used to indirectly detect ATP hydrolysis (Leyssens et al., 1996). The majority of intracellular ATP is complexed with divalent magnesium ions; however. ADP has a lower affinity for magnesium ions than ATP. Thus, hydrolysis of MgATP causes an increase in free magnesium ion concentration and subsequent increase in Magnesium Green fluorescence. By invoking binding equilibrium. Magnesium Green has been used in non-imaging studies to determine the ATP-ADP exchange rate through the mitochondrial adenine nucleotide transporter in isolated mitochondria (Chinopoulos et al., 2009) and permeabilized cells (Kawamata et al., 2010). In an imaging study using fluorescence confocal microscopy and isolated hair cells loaded with Magnesium Green. Shin et al. (2007) indirectly visualized a higher contribution of creatine kinase activity to ATP generation in hair bundles than in their soma. While sensitive to ATP hydrolysis, Magnesium Green has the disadvantage of imperfect specificity because it has moderate affinity for calcium. On the other hand. Magnesium Green is more useful than other magnesium indicators because its fluorescence can be excited with illumination in visible range, reducing phototoxicity compared to indicators such as Mag-Fura-2 and MagIndo-1, which require ultraviolet (UV) excitation. However, Magnesium Green shows a simple increase in fluorescence intensity upon binding magnesium ions without a ratiometric shift in excitation or emission peaks. Thus, unlike ratiometric probes, the Magnesium Green signal also depends on dye concentration, making it a challenge to use in quantitative studies. Loading efficiency and photobleaching can cause variations in signal that are unrelated to changes in ATP hydrolysis (Leyssens et al., 1996).

To directly visualize ATP pools, fluorescent analogues can be used as tracers once they are loaded into the tissue, cell, or organelle of interest. Analogues can be synthesized by conjugating fluorescent groups such as a methylanthraniloyl (mant) or a coumarin (deac) to the nucleobase, ribose, or phosphate groups of ATP. The analogues, mant-ATP and deac-ATP, have been used extensively to study kinase and adenosine triphosphatase (ATPase) activities in solution (Fili and Toseland, 2014). For example, mant-ATP has been used to monitor vesicular ATP release from dopaminergic neurons (Ho et al., 2015), and deac-ATP was instrumental in a study of the kinetics of myosin Va movement on actin. using total internal reflection fluorescence (TIRF) microscopy (Sakamoto et al., 2008). Analogues of ATP modified with Cy3 or BODIPY fluorescent dyes, which shift excitation bands into the visible range to reduce phototoxicity, are also available.

Additional synthetic ATP analogues have been developed to detect ATP hydrolysis. These analogues exploit Forstertype resonance energy transfer (FRET) between a donor fluorophore covalently attached to a [gamma]-phosphate group and an acceptor fluorophore linked to the base or ribose (Hardt et al., 2013). Cleavage of the phosphodiester bond allows the donor and acceptor to diffuse away from one another. causing a drastic loss of FRET and an increase in donor fluorescence. Hacker et al. (2013) have used such an analogue to monitor ATP consumption in real time during ubiquitin activation by UBA1, a human El enzyme. Similarly, a FRET-based ATP analogue with the organic fluorophore Sulfo-Cy3 dye linked to the y-phosphate and a quencher linked to the ribose C2 position was used by Gutierrez Acosta et al. (2014) to study acetone degradation in cell extract of Desulfococcus biacutus. While these FRET-based ATP analogues have not yet been used for imaging, in principle they could be used in similar fashion to mant-ATP. It is important to recognize, with these ATP analogues, that the fluorescent dye group can be of equivalent or greater mass than ATP itself. Therefore, the covalent modification could change the behavior of the analogue in unpredictable ways. Functional assays are critical to validate the use of such analogues.

Small-molecule sensors for ATP as an analyte have also been reported. For example, quinacrine is a fluorescent acridine derivative that stains peptide-bound ATP found in high concentrations in intracellular granules (Irvin and Irvin, 1954: Bodin and Burnstock, 2001). In this manner. quinacrine has been used to image vesicular ATP release in endothelial and epithelial cells (Bodin and Burnstock. 2001: Feranchak et al., 2010; Akopova et al., 2012). Alternatively, Pak et al. (2015) developed an imidazolium-based ratiometric sensor for ATP with a pyrene excimer clamp. When ATP binds to this sensor, it forms a pyrene-adenine-pyrene sandwich by [PI]-[PI] stacking. Formation of the complex results in an increase in pyrene emission at 375 nm and a decrease in emission at 487 nm. This pyrene-based sensor was used in HeLa cells to monitor the decrease in ATP levels upon addition of the ATP synthase inhibitor, oligomycin, and upon hydrolysis of ATP to ADP by apyrase (Pak et al., 2015).

Aptamers are single-stranded DNA or RNA oligonucleotides that can be easily modified to attain high-affinity and specificity for their targets. Aptamers are used extensively to study small-molecule metabolites (Paige et al., 2012; Feng et al., 2014), and are analyzed by a variety of methods, such as fluorescence spectroscopy (Sun et al., 2010; Park et al., 2015b; Wang et al., 2015; Song et al., 2016), electrochemistry (Mukherjee et al., 2015; Zhao et al., 2015). surface plasmon resonance (Park et al., 2015a), and colorimetry (Huo et al., 2016). Aptamers can be engineered to detect ATP in the nanomolar to millimolar ranges; however, cell permeability and degradation issues limit their use in live-cell imaging (Wang et al., 2014). To overcome this problem, nanoparticles have been used to deliver and protect aptamers from degradation by DNases and RNases in cells while modifying their fluorescent properties. For example, Qiang et al. (2015) reported a polydopamine, nanosphere-linked aptamer hybrid that protects the aptamer and quenches its fluorescence. The addition of ATP releases the aptamer, resulting in an increase in fluorescence. The aptamer is highly selective and sensitive, detecting ATP in the 0.01-2-mmol [1.sup.-1] range. Changes in ATP concentration could be measured in HeLa cells upon treatment with oligomycin or [Ca.sup.2+]. Similarly, nanoparticles have been used to construct nanoflares (Zheng et al., 2009) to image ATP. Aptamers bound to gold nanoparticles are hybridized with fluorescent DNA strands that are quenched by proximity to the nanoparticle. Binding of ATP to the aptamers releases the fluorescent strands, and the increased fluorescence can be used to quantitate ATP in live cells (Zheng et al., 2009; Torabi and Lu, 2014). One drawback of these biosensors is that the aptamers have been engineered for adenine selectivity, which can make it challenging to distinguish between adenine derivatives (Ozalp et al., 2010). To overcome this disadvantage. Sassanfar and Szostak (1993) synthesized RNA aptamers specific to ATP, and Ozalp et al. (2010) developed DNA aptamers selective for ATP (apparent [K.sub.D]: 3.2 mmol [1.sup.-1]) and (ADP apparent [K.sub.D]: 4.4 mmol [1.sup.-1]).

While both small organic indicators and aptamer-based biosensors have found utility in imaging studies, they pose a challenge to sample preparation because they require cell penetration or cell loading of the exogenous reagent. In simple model systems such as monolayer cell cultures, introduction of these ATP imaging reagents typically can be achieved by sustained incubations, electroporation, or microinjection. However, these preparatory requirements limit their application in many cell types and complex or thick tissues. In contrast to these technologies, genetically encoded indicators are protein-based reagents that are in part or wholly encoded by an appropriate gene sequence. Thus. genetically encoded imaging reagents offer compatibility with a wide variety of specimens when the appropriate gene transfer or expression method is available, such as transfection reagents, viral transduction, and tissue-specific expression in transgenic mice. Recent examples of genetically encoded ATP imaging reagents have utilized luciferases and fluorescent proteins.

Luciferases are used to produce bioluminescence in order to visualize cellular activities and to track cell populations in vivo. Firefly luciferase and its exogenously supplied substrate, luciferin, can be used to measure ATP because the chemiluminescent reaction is ATP-dependent. Adenylation of luciferin by luciferase activates the substrate for conversion to oxyluciferin and thus the resulting luminescence is proportional to ATP concentration (Manfredi et al., 2002; Lundin, 2014). Current commercial and academic efforts have produced a variety of luciferases that have been engineered and codon-optimized from different species for bioluminescence imaging in live samples (Thorne et al., 2010; Hall et al., 2012). But the ATP-dependent firefly and beetle luciferases are most commonly used to measure ATP (Manfredi et al., 2002; Lundin, 2014). Branchini et al. (2015) have developed a luciferase with higher activity and quantum yield by fusing the N-terminal domain of the Photinus pyralis luciferase joined to the C-terminal domain of the Luciola italica luciferase. The chimeric luciferase was further optimized to show threefold higher sensitivity in live cells than the commercial Luc2 variant (Branchini et al., 2015). Mutagenesis also has produced luciferase variants with red-shifted luminescence (Branchini et al., 2010). While these emission variants typically exhibit lower luminescence yields, they offer the advantage of lesser tissue absorbance of red luminescence, making them good reporters for in vivo imaging (Liang et al., 2012). There are important caveats to quantitation of ATP by luciferase. For example, optimization is critical because of the inherent dependence on oxygen concentration and the often underappreciated complication of product inhibition, as well as inhibition by cellular factors or pharmacological agents (Leitao and Esteves da Silva, 2010; Lundin. 2014).

Despite its difficulties with absolute quantitation, luciferase has been used to monitor ATP changes in a variety of cells, such as HEK-293 cells, cardiomyocytes, and neurons (Brovko, 2010). Bell et al. (2007) used wild-type luciferase inside the cell to monitor differences in intracellular ATP responses in mitochondria and the cytosol of cardiomyocytes after stimulation. Free cytosolic and membrane-tethered luciferase also have been used to study the possibility of compartmentalization of submembrane ATP and control of [K.sub.ATP] channel activity in pancreatic [beta]-cells and hypothalamic neurons (Kennedy et al., 1999; Ainscow et al., 2002). In the detection of intracellular ATP, luciferase is introduced into mammalian cells by gene transfection, microinjection, or viral vectors; however, Lee et al. (2012) developed luciferase fused to a protein transduction domain to facilitate direct transport of the luciferase protein into the cell. Even with genetic encoding, the exogenous luciferin substrate must be supplied and must penetrate tissues and cells. While this has proven to be achievable in live cells and animals, there is still the possibility of variable substrate access and cell-type-specific toxicity that must be taken into account (Rangaraju et al., 2014). In the detection of extracellular ATP, luciferase also has been used to study ATP release by tethering the enzyme to the extracellular face of the plasma membrane via conjugation to primary Immunoglobulin G (IgG) antibodies that can bind to surface antigens via streptavidin-biotin tags and via glycophosphatidyIinositol (GPI) lipid anchors (pmeLuc) (Praetorius and Leipziger, 2009). For example, HEK-293 cells that were stably transfected with pmeLuc detected micromolar levels of ATP in the tumor microenvironment, while extracellular ATP was undetectable in healthy tissues (Pellegatti et al., 2008). The luciferase-luciferin system can offer a low-background, low-toxicity method of monitoring ATP; however, low luminescence limits its applicability in real-time ATP imaging. Bioluminescence imaging typically requires long exposure times, which limit spatiotemporal resolution: in some applications, it can require specialized equipment (Bell et al., 2007; Brovko, 2010). For example, Furuya et al. (2014) improved temporal resolution to 100 ms by using a cooled electron-multiplying charge-coupled device (EMCCD) camera together with an image intensifier to compensate for low photon emission rates. They used it along with low-magnification, high numerical aperture objectives to monitor ATP release from a single cell with 10-nmol [1.sup.-1]-detection sensitivity. Furthermore, interference from the sample matrix can occur due to inhibitory levels of various anions and salts, ion channel inhibitors, and P2 receptor antagonists (Praetorius and Leipziger, 2009).

An alternative to luciferase-based detection is the use of fluorescent protein-based ATP sensors. Fluorescent, protein-based sensors do not require the addition of an exogenous luciferin substrate, and they offer ease of manipulation at the DNA level and subsequent ease of expression in cells and in vivo. For example, as an alternative to the electrophysiological approach (Gribble et al., 2000). ATP-level changes in HEK-293 cells have been visualized by imaging the ATP-dependent, conformational change of [K.sub.ATP] channels fused to an ECFP-EYFP cyan and yellow fluorescent protein FRET pair (Tsuboi et al., 2004). Imamura et al. (2009) developed a family of sensors named the "ATeams," which are also based on Forster-type resonance energy transfer (FRET) between a donor fluorescent protein and an acceptor fluorescent protein. In the original ATeams, the [epsilon] subunit from the [F.sub.0][F.sub.1]-ATP synthase is linked to the CFP and YFP FRET pair. The [epsilon] subunit is a 14-kDa protein subunit that is composed of a N-terminal beta-barrel and two C-terminal helices. It undergoes a large conformational change upon binding of ATP, which is the mechanism by which the e regulates [F.sub.0][F.sub.1] ATPase activity based on intracellular ATP levels. The ATeam sensors exhibit apparent dissociation constants, ranging from 7.4 [micro]mol [1.sup.-1] to 3.3 mmol [1.sup.-1], with at least 10- to 100-fold greater selectivity for ATP over ADP, and response kinetics on the timescale of seconds.

The ATeam sensors have been used to study ATP changes in bacterial cells, neurons, and a number of different cell types in different species (Imamura et al., 2009; Toloe et al., 2014). In unicellular organisms. Maglica et al. (2015) used ATeam sensors to monitor antibiotic-induced cell death by single-cell tracking of ATP levels in Mycobacterium smegmatis, demonstrating its potential as an important technology in drug discovery for antibiotic screening and mechanism of action imaging assays. In multicellular organisms. Ozawa et al. (2015) found that glycolysis-dependent ATP production was necessary for formation of lamellipodia in podocytes. In contrast, the cortical subcellular region of podocytes produces ATP by both glycolysis and oxidative phosphorylation in mitochondria. These metabolic studies demonstrate how the ATeam sensors can provide important insight into podocyte cell biology, potentially advancing our understanding of the role of metabolic dysfunction in chronic kidney diseases (Ozawa et al., 2015).

When imaging fluorescent ATP sensors that excite and emit in the 400-500-nm spectral range, it is important to consider metabolism-dependent changes in autofluorescence background. For example, autofluorescence of flavins such as flavin adenine dinucleotide (FAD) have long been used to visualize metabolic changes in live samples, and continue to be used as a "label-free imaging" option (Quinn et al., 2013: Jahn et al., 2015). This background could convolute fluorescent signals when sensors are being used at low expression levels, but at moderate sensor expression this is less likely because of the lower extinction coefficient (~10,000 mol [1.sup.-1] c[m.sup.-1]) and fluorescence quantum yield (< 0.01 to 0.06) of FAD (Valle et al., 2012). To mitigate problems with spectral overlap with FAD autofluorescence and to generate spectral diversity for multi-sensor imaging, Nakano et al. (2011) replaced the CFP and YFP in the original ATeam sensors with a different FRET pair. They used green (GFP: cpl73-mEGFP) and orange (OFP; mKO[KAPPA]) fluorescent proteins to generate a red-shifted ATeam named GO-ATeam. This allowed them to image [Ca.sup.2+]-level changes, using [Ca.sup.2+] sensors that are excited with UV illumination, simultaneously with ATP levels in HeLa cells. GO-ATeam is also more stable to acidification, an important advantage because metabolic stress can cause a decrease in intracellular pH (Nakano et al., 2011). Subsequently. Rueda et al. (2015) used GO-ATeam to image ATP depletion in the presence and absence of [Ca.sup.2+] upon NMDA exposure in neurons.

Zadran et al. (2013) developed an ATP sensor, using a particular type of fluorescent protein FRET pair that can exhibit enhanced acceptor fluorescence. In their approach, a mutated variant of the e subunit of Bacillus subtilis [F.sub.0][F.sub.1]-ATP synthase is coupled to GFP and YFP. Both GFP and YFP are excited at the same wavelength, and ATP binding results in enhanced YFP acceptor fluorescence signal as a result of increased FRET. Zadran's sensor was shown to detect as low as 10 nmol [1.sup.-1] ATP with high specificity (Zadran et al., 2013). Subsequently, this sensor was used to monitor ATP flux in tumors and during transition to metastatic behavior (Zadran et al., 2014).

Yaginuma et al. (2014) recently developed the QUEEN (quantitative evaluator of cellular energy) family of sensors, which are related to the ATeam sensors but with a different architecture, one that uses a circularly permuted fluorescent protein (cpFP). In the QUEEN architecture, a cpFP is inserted between two [alpha] helices of the [F.sub.0][F.sub.1]-ATP synthase e subunit with linkers. Two variants were designed with apparent affinities of 7 [micro]mol [1.sup.-1] and 2 mmol [1.sup.-1], called QUEEN-7[micro]. and QUEEN-2m, respectively (Yaginuma et al., 2014). Using these sensors, they quantified ATP distribution in individual E. coli cells.

Using a circular permutation strategy, Tantama et al. (2013) reported an ATP-to-ADP ratio sensor, PercevalHR, which is an improved version of the original Perceval (Berg et al., 2009). The PII family protein, GlnKl, which can bind MgATP and ADP with high affinity, was modified with a cpFP insertion within a loop at the nucleotide binding site. MgATP binding causes a conformational change that alters the chromophore environment of the cpFP. As a result of the protein engineering, the sensor has two peaks in its excitation spectra, at approximately 420 nm and 500 nm. MgATP binding increases fluorescence intensity with 500-nm excitation. while ADP binding increases fluorescence intensity with 420-nm excitation. These nucleotide-dependent spectral features enable excitation ratiometric imaging, which offers the significant advantage of concentration independence that normalizes the signal for protein expression and reduces artifacts from photobleaching. The fast association and dissociation rates, as well as the high dynamic range of the sensor at physiologically relevant ATP-to-ADP ratio ranges, make PercevalHR useful for studying energy metabolism in cells (Tantama et al., 2013). Although the Perceval sensors exhibit some pH sensitivity, the ATP response can be deconvoluted from changes in sample pH, using a pH sensor and experimental calibration (Tantama et al., 2011: Tantama and Yellen, 2014). Using Perceval to measure ATP-to-ADP ratios in neurons. Zala et al. (2013) showed that mitochondrial trafficking is dependent on mitochondrial ATP but not glycolysis. With PercevalHR, Rueda et al. (2015) studied NMDA stimulation-dependent decreases in ATP-to-ADP ratio in neurons deficient in the SCaMC-3 mitochondrial., calcium-dependent MgATP-phosphate exchanger.

Finally, luciferase and fluorescent protein technologies have been combined in ATP sensors that exploit bioluminescence resonance energy transfer (BRET). These BRET sensors have been developed to improve brightness and red-shifted, luciferase-based imaging (Chu et al., 2016). Saito et al. (2012) developed an ATP sensor, Nano-lantern (ATP1), in which a split Renilla luciferase (RlucS) was modified with the [epsilon] subunit of the [F.sub.0][F.sub.1]-ATP synthase and Venus. ATP binding results in complementation and reconstitution of the active RlucS, which can efficiently transfer energy to Venus. It has an apparent ATP affinity of 0.3 mmol [1.sup.-1] and is useful for imaging in tissues with high autofluorescence. Conveniently, RlucS does not consume any ATP in its chemiluminescent reaction, simplifying the detection mechanism and interpretation. Using this sensor. Saito et al. (2012) visualized ATP increases in the mesophyll of leaf cells after light irradiation. Borghei and Hall (2014) used firefly luciferase to develop a red-shifted ATP sensor. They generated a fusion protein with firefly luciferase (X5) and the fluorescent protein, mCherry. Although the low quantum yield of mCherry (0.22) results in a weak signal., they detected an ATP-dependent increase in emission at 600 nm by mCherry (Borghei and Hall. 2014). Presumably, replacement of mCherry with a brighter red fluorescent protein would improve signal strength. Importantly. Rangaraju et al. (2014) used a synaptically targeted, engineered luciferase-mCherry sensor (Syn-ATP) to achieve ratiometric measurements of activity-dependent ATP consumption and production, although the Syn-ATP ratiometric signal was not obtained through BRET.

A wide variety of ATP detection and imaging reagents that can be used to visualize both intracellular energy metabolism and extracellular purinergic signaling are available. The choice of technologies depends on the biological process under study, the specimen format, and any limitation of the imaging instrumentation. While we have presented several examples, it is beyond the scope of this review to evaluate all detection and imaging technologies here, and other examples have been reviewed elsewhere (Khlynsteva et al., 2009 and Praetorius and Leipziger. 2009).

Discussion and Conclusions

While a number of ATP imaging technologies are available, there are still gaps in our ability to fully probe ATP metabolism and signaling. Quantitative imaging could be improved by engineering sensors with 1) a variety of ATP affinity ranges, 2) faster ATP binding and response kinetics. 3) higher brightness and contrast ratios, 4) selectivity for ATP hydrolysis products, and 5) spectral color variation. The engineering of sensors with different affinity ranges is important because ATP concentrations vary widely: from nanomolar extracellular levels in some tissues, to millimolar concentrations in cytosolic pools. Likewise, developing sensors that have faster kinetics will be helpful for capturing ATP concentration changes that possibly occur on the millisecond timescale, such as during the initial phases of vesicular ATP release. Improving the brightness of fluorescent sensors and the contrast ratio between unbound and ATP-bound states will improve signal-to-noise ratios and quantitation, and even spatial and temporal resolution in practice. Engineering sensors with selectivity for ATP hydrolysis products will expand the toolbox to include ADP. AMP, and adenosine as measurable analytes. Similarly, those sensors that offer spectral color variation will enable simultaneous imaging of multiple sensors, providing a means to directly correlate ATP with, for example, [Ca.sup.2+] signaling and kinase activities. Finally, engineering spectral variants with luminescence and fluorescence in the far-red and infrared spectral ranges will ultimately enable, in live animals, ATP imaging with specificity and subcellular resolution in timescales from subseconds to lifetime.

With current technologies and these future improvements. there is great potential to study ATP with a systems biology perspective of energy metabolism and purinergic signaling. Continued use of these methods to image activity-dependent bioenergetics will improve our understanding of the mechanisms of neurotransmission (Tantama et al., 2013; Rangaraju et al., 2014). Bioenergetic deficits have been linked to a number of aging-related neurodegenerative diseases such as Huntington's and Parkinson's diseases, and imaging approaches will aid the study of neurodegenerative mechanisms (Surmeier et al., 2012: Zala et al., 2013: Rangaraju et al., 2014). Across cell types, these technologies can probe metabolic and mitochondrial function at the single-cell level. This is becoming important for understanding diseases such as cancer, in which links to metabolism are increasingly being found (Mayers and Vander Heiden, 2015). In both healthy and disease-state biology, live-cell imaging is becoming an integral approach, and our growing ability to quantitatively visualize ATP dynamics will allow us to link phenomenology and mechanism.

Acknowledgments

MT acknowledges support from the Showalter Foundation, the Purdue Research Foundation, and National Institutes of Health grants no. NS092010 and no. EY026425.

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MEGHA RAJENDRAN (1), ERIC DANE (2), JASON CONLEY (1), AND MATHEW TANTAMA (1,*)

(1) Department of Chemistry, Purdue University, 560 Oval Drive, Box 68. West Lafayette. Indiana 47907: and (2) Koch Institute for Integrative Cancer Research. Massachusetts Institute of Technology, 77 Massachusetts Avenue. 76-211, Cambridge, Massachusetts 02139

Received 19 February 2016; accepted 19 July 2016.

(*) To whom correspondence should he addressed. E-mail: mtantama[R] purdue.edu

Abbreviations: BRET, bioluminescence resonance energy transfer: cAMP, cyclic adenosine monophosphate; CFP, cyan fluorescent protein: cpFP, circularly permuted fluorescent protein: Cy, cyanine: DAMP, damage-associated molecular pattern: deac, diethylaminocoumarin: EMCCD, electron-multiplying charge-coupled device; ENT, equilibrative nucleoside transporter: FMLP. N-formyl-Met-Leu-Phe peptide: FRET. Forster-type resonance energy transfer: mant. (2'/3'-O-(N-Methlanthraniloyl); GTP, guanosine triphosphate; NMDA, N-methyl-D-aspartate; TIRF, total internal reflection fluorescence.

Table 1
Examples of tools for adenosine triphosphate (ATP) visualization

      Technology                 Detection mechanisms

Quinacrine (a)             Fluorescent dye that binds
                            peptide-bound ATP found in

                            intracellular granules;
                            intensiometric (*)
Mant-ATP (b)               Fluorescent ATP analogue that can be
                            used as a tracer

                            of ATP pools: intensiometric
ATP Aptamer nanoflare (c)  ATP binding to DNA aptamers bound to gold
                            nanoparticles causes release of
                            Cy5-conjugated

                            "reporter" strand, whose fluorescence
                            "turns on"
                            upon release; intensiometric
Luciferase (d)             Chemiluminescent enzyme metabolizes
                            its substrate
                            luciferin using ATP, resulting in
                            ATP-dependent
                            luminescence; intensity-based signal
Syn-ATP (e)                Luciferase-mCherry bioluminescence
                            resonance energy


                            transfer (BRET) sensor targeted to neuronal

                            synapses; both luminescence and red
                            fluorescence;
                            ratiometric
ATeam (f)                  ATP binding causes an increase in
                            Forster resonance


                            energy transfer (FRET) between a CFP
                            and a YFP;

                            ratiometric

QUEEN (8)                  ATP binding causes a change in the
                            excitation

                            spectrum of a circularly permuted green
                            fluorescent

                            protein (cpEGFP): ratiometric
PercevalHR (h)             ATP binding causes a change in the
                            excitation

                            spectrum of a circularly permuted yellow
                            fluorescent

                            protein (cpVenus); ratiometrie

      Technology                        Imaging parameters

Quinacrine (a)
                           Fluorescence; [[lambda].sub.ex], 420-488 nm;
                           [[lambda].sub.cm], 490-510 nm


Mant-ATP (b)
                           Fluorescence: [[lambda].sub.ex], 356 nm;
                           [[lambda].sub.cm], 428-448 nm

ATP Aptamer nanoflare (c)  Fluorescence (dye-dependent):

                           Cy5: [[lambda].sub.ex], 649 nm;
                           [[lambda].sub.cm], 666 nm



Luciferase (d)
                           Chemiluminescence (substrate-dependent):

                           D-luciferin: [[lambda].sub.em], 560 nm

Syn-ATP (e)
                           Chemiluminescence D-luciferin:
                           [[lambda].sub.e
                           560 nm
                           Fluorescence: [[lambda].sub.ex], 587 nm;
                           [[lambda].sub.em], 610 nm



ATeam (f)
                           CFP donor fluorescence: [[lambda].sub.ex],
                           435 nm; [[lambda].sub.em], 475 nm

                           YFP acceptor fluorescence:
                           [[lambda].sub.ex].
                           515 nm; [[lambda].sub.em], 527 nm
                           FRET: [[lambda].sub.ex], 435 nm;
                           [[lambda].sub.em], 527 nm
QUEEN (8)
                           A-Band fluorescence: [[lambda].sub.ex],
                           400 nm; [[lambda].sub.em], 513 nm

                           B-Band fluorescence: [[lambda].sub.ex].
                           494 nm; [[lambda].sub.em]. 513 nm

PercevalHR (h)
                           A-Band fluorescence: [[lambda].sub.ex],
                           420 nm; [[lambda].sub.em], 515 nm

                           B-Band fluorescence: [[lambda].sub.ex].
                           500 nm; [[lambda].sub.em], 515 nm

(*) Intensiometric sensors exhibit a change in intensity without a
drastic change in spectrum: for luminescence, a single emission
wavelength is monitored.
and for fluorescence, a single excitation wavelength
([[lambda].sub.ex]) and emission wavelength ([[lambda].sub.em])
pair are monitored. Intensiometric signals depend not only on ATP
but also can be affected by dye concentration, expression level,
and photobleaching. Ratiometrie signals obtain two readouts
that are divided to provide
a normalized response. Ratiometrie signals are advantageous
because they normalize for dye concentration and expression
level, and reduce signal drift
from photobleaching.
A-Band fluorescence, excitation of the protonated chromophore
of the fluorescent protein; B-Band fluorescence, excitation
of the deprotonated
chromophore form of the fluorescent protein: CFP, cyan fluorescent
protein; Cy5, cyanine dye; YFP, yellow fluorescent protein.
(a) (Akopova et al., 2012): (b) (Ho et al., 2015); (c)
(Zheng et al., 2009); (d) (Manfredi et al., 2002); "(Rangaraju et al.,
2014); (f) (Imamura et al., 2009); (g) (Yaginuma et al., 2014); (h)
(Tantama et al., 2013).


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Author:Rajendran, Megha; Dane, Eric; Conley, Jason; Tantama, Mathew
Publication:The Biological Bulletin
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Date:Aug 1, 2016
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