55S. Masino and D. Boison (eds.), Adenosine: A Key Link between Metabolism and Brain Activity, DOI 10.1007/978-1-4614-3903-5_3, © Springer Science+Business Media New York 2013

Abstract The brain is one of the most metabolically active organs in the body, but because it has a limited capacity to store bioenergetic molecules it requires a con-tinuous supply of oxygen and energy. In order to meet the high metabolic needs of the brain, the blood–brain barrier selectively expresses speci fi c transport systems including glucose transporters and monocarboxylic acid transporters that transport lactate and ketone bodies. As a universal energy source, adenosine triphosphate (ATP) drives biological reactions essential for brain functions, and loss of such cel-lular energy results in profound abnormalities in brain function. The high-energy phosphate bonds of ATP are rather labile and thus energy inherent in ATP is readily released when ATP is hydrolyzed sequentially to ADP, AMP, and fi nally adenosine. Although each of these molecules serves different functions and can activate differ-ent signaling pathways, our focus here is on adenosine and brain energy metabo-lism. Under basal conditions, brain levels of adenosine are nearly 10,000-fold lower than ATP. Therefore, unnoticeable and possibly physiologically irrelevant decreases in ATP levels can result in dramatic and physiologically relevant rises in adenosine levels. As brain energy levels drop, adenosine levels rise to adjust brain energy sup-ply and to retaliate against an external stimulus that would otherwise cause exces-sive ATP breakdown. These actions of adenosine are mediated by adenosine receptors located on target cells including neurons, glial cells, and brain endothelial cells. A critical issue in studying brain bioenergetics is the precise and accurate measurement of levels of ATP and its metabolites including adenosine. Because these molecules can be degraded rapidly, it is challenging to make such measure-ments. Essential components in the correct assessment of brain energetics should include justifying carefully the methodology used and putting the data in the con-text of what is already known about brain energy metabolism.

X. Chen • L. Hui • J. D. Geiger (*) Department of Pharmacology, Physiology and Therapeutics , School of Medicine and Health Sciences, University of North Dakota , 501 N. Columbia Road , Grand Forks , ND 58202 , USA e-mail: jonathan.geiger@med.und.edu

Chapter 3 Adenosine and Energy Metabolism—Relationship to Brain Bioenergetics

Xuesong Chen , Liang Hui , and Jonathan D. Geiger

56 X. Chen et al.

Keywords Adenosine • Adenylate energy charge • Blood–brain barrier • Glucose • Glycogen • Ketone bodies • High-energy focused microwave irradiation

3.1 Overview of Sources and Requirements for Brain Bioenergetics

The brain is a complicated and active organ that requires an almost constant supply of energy. While the brain comprises only about 2 % of the body weight, it uses approximately 20 % of total body energy consumption; and that is during a resting state (Rolfe and Brown 1997 ) ! Roughly one-half of the brain’s energetic require-ments are used to maintain ionic gradients across neuronal plasma membranes and the vast majority of this is used by Na + /K + -ATPase, which pumps K + into and Na + out of neuronal cells: a process with keen requirements for adenosine triphosphate (ATP), a universal and ubiquitous source of cellular energy (Attwell and Laughlin 2001 ) . The brain contains the largest density of Na + /K + -ATPase molecules in the body (~11 pmol/g tissue); by comparison the concentration of this enzyme is about 20 times lower in heart, another highly active organ (Clausen 1998 ) . Other than Na + /K + -ATPase, about one-half of the brain’s energy consumption is used for trans-port of neurotransmitters and other essential molecules across cell membranes, for transport of substances within cells, and for synthesis of the building blocks of cells including proteins, lipids, and carbohydrates (Attwell and Laughlin 2001 ; McKenna et al. 2006 ) .

Although metabolically active, the brain has a very limited capacity to store bioenergetic molecules. These energetic stores include ATP, glycogen, ketone bod-ies, and in some cases lipid-based molecules. Essential for physiological regulation of brain function is the continuous supply of oxygen and energy-yielding nutrients and of course all of this originates from blood. Thus, it is easy to understand why even a very brief interruption (seconds) in blood fl ow and/or absence of oxygen results in dramatic losses of cellular energy, loss of consciousness, and profound abnormalities in brain function up to and including increased susceptibility to insult-induced neuronal cell death.

Further complicating access of energy substrates from blood into brain is the presence of the blood–brain barrier (BBB), a physical and metabolic barrier that permits selective passage of cells and substances into brain (Abbott et al. 2010 ) . While water and oxygen can readily diffuse across the BBB, other larger particles cannot cross the BBB freely. The restrictive nature of the BBB is due primarily to tight junctions formed between adjacent endothelial cells, a lack of pinocytosis by endothelial cells, tightly coupled end-feet of astrocytes, and less so by the presence of an underlying continuous basement membrane (Abbott et al. 2010 ) . Functionally, these restrictions provide a structural and functional barrier capable of preventing blood-borne toxins and cells from entering brain. However, although the BBB works to protect the brain microenvironment it does limit the entry of energy-yielding nutrients and in doing so the BBB increases the risk of the brain experiencing

573 Adenosine and Energy Metabolism—Relationship to Brain Bioenergetics

insult-induced bioenergetic crises. In order to meet the high metabolic needs of the brain, the BBB selectively expresses speci fi c transport systems (Spector 2009 ) : the highly glycosylated 55 kDa form of the glucose transporter (GLUT1) that transports blood glucose into brain parenchyma (Ito et al. 2011 ; Vannucci et al. 1997 ) and the monocarboxylic acid transporter (MCT1) that transports monocarboxylic acids including lactate and the ketone bodies acetoacetate and b -hydroxybutyrate (Ito et al. 2011 ; Vannucci and Simpson 2003 ) .

During early stages of development the brain is more fl exible in terms of using various substrates for energy. However, adult brain relies primarily on blood glucose. Indeed, it has been estimated that brain uses about 25 % of total body glucose and 20 % of respired oxygen (Magistretti et al. 1995 ) . Once blood glucose is transported into brain parenchyma via the insulin-insensitive GLUT1 transporters, glucose is taken up by astrocytes via the 45 kDa form of GLUT1, by neurons via the high-capacity GLUT3 transporter, and by microglia via GLUT5. Most of the glucose that enters mitochondria is metabolized to pyruvate and then converted to acetyl-CoA before entering the Kreb’s tricarboxylic acid cycle to produce ATP; only about 13 % of glycolytic pyruvate is converted to lactate. In addition to energy production, glucose is also used for amino acid and neurotransmitter biosynthesis, glycogen synthesis, lipid biosynthesis via the pentose phosphate shunt, maintenance of reduced glutathi-one, and incorporation in glycoproteins and glycolipids (McKenna et al. 2006 ) .

In developing brain, during periods of prolonged fasting when blood glucose levels are low, or during the ingestion of high-fat diets that are extremely restricted in terms of available carbohydrates (the so-called ketogenic diet that mimics fasting/starvation)—ketone bodies provide a key alternative fuel for the brain capable of providing 60–70 % of the energy need of the brain (Owen et al. 1967 ) . Consisting of acetoacetate, b -hydroxybutyrate, and acetone, ketone bodies are formed in hepatic mitochondria as by-products of fatty acid metabolism. Acetoacetate and b -hydroxybutyrate are transported to the brain through MCT1 expressed in the BBB. Once inside brain, these ketone bodies are taken up by neurons via the high-af fi nity MCT2 transporters, and by glia cells via MCT1 and MCT4 transporters (Bergersen et al. 2002 ; Ra fi ki et al. 2003 ; Vannucci and Simpson 2003 ) . In addition to ketone bodies coming from systemic circulation, astrocytes have been shown to produce ketone bodies from fatty acids and leucine (Auestad et al. 1991 ; Bixel and Hamprecht 1995 ; Guzman and Blazquez 2001 ; Guzman and Blazquez 2004 ; Lopes-Cardozo et al. 1986 ) . Ketone body production by astrocytes could contribute signi fi cantly to brain bioenergetics because astrocytes can outnumber neurons 9:1 and can occupy at least 50 % of the cerebral volume (White and Venkatesh 2011 ) .

In normal adults, blood levels of ketone bodies are very low and cerebral utilization of ketones is negligible. However, when glucose stores are exhausted and the rate of gluconeogenesis is insuf fi cient to meet the needs for glucose by brain, blood levels of ketone bodies rise to low mM concentrations as a result of the rapid fat catabo-lism (Levitsky et al. 1977 ; Owen et al. 1967 ) and the uptake and utilization of ketone bodies increase dramatically (Hasselbalch et al. 1994 ; Leino et al. 2001 ) . Through the actions of d - b -hydroxybutyrate dehydrogenase, acetoacetate-succinyl-CoA transferase and acetoacetyl-CoA-thiolase ketone bodies in brain can be converted

58 X. Chen et al.

into acetyl-CoA, which then can be used to produce ATP, lipids, and/or proteins (Patel et al. 1975 ) . In contrast to glucose, ketone bodies can bypass glycolysis and enter directly into mitochondria where they are oxidized to acetyl-CoA. Thus, ketone bodies are more ef fi cient in producing ATP than is glucose and can stimulate mitochondrial biogenesis via the upregulation of genes encoding energy metabo-lism and mitochondrial enzymes (Kashiwaya et al. 1994 ; Sato et al. 1995 ; Veech et al. 2001 ) . In addition, metabolism of ketone bodies produces less reactive oxygen species than does glucose (Mobbs et al. 2007 ) . Thus, the utilization of ketone bodies as an alternative energy fuel could exert bene fi cial effects to brain (Cunnane et al. 2011 ; LaManna et al. 2009 ) .

In addition to ketone bodies, lactate can be also used for energy in brain (Itoh et al. 2003 ; Smith et al. 2003 ) . Like ketone bodies, lactate from systemic circulation is transported to brain at the BBB via MCT1. Once inside brain, lactate is uptaken via MCTs and is then oxidized to produce ATP. In addition to peripherally derived lactate, astrocytes have been implicated as lactate sources inside the brain where they are thought to participate in supplying lactate to neurons especially during peri-ods of enhanced synaptic activity (Chih and Roberts 2003 ; Kasischke et al. 2004 ) .

Glycogen, primarily stored in astrocytes, is the largest energy reserve in brain. When brain glucose levels are very low, glycogenolysis in astrocytes can provide neurons the necessary glucosyl units in the form of lactate to maintain ATP synthe-sis (Choi and Gruetter 2003 ) . However, glycogen levels in brain are very low (3 mmol/kg), about 100-fold lower than in liver, and this amount of glycogen can only sustain brain energy for a very short period of time (Brown 2004 ; Cruz and Dienel 2002 ; Nelson et al. 1968 ) . As such, a continuous supply of glucose and oxy-gen is necessary to sustain neuronal activity.

For decades, it has been suggested that brain does not use fatty acids as a source of fuel, even though about 60 % of solid matter in the brain is fat. This view is sup-ported by observations that fatty acids that are attached to lipoproteins cannot cross the BBB and brain has no triacylglycerol stores. This point of view might be true for neurons, because enzyme activity for fatty acid oxidization is very low (Yang et al. 1987 ) . However, recent studies on brain energy metabolism have revealed a more intricate picture of fatty acid metabolism in brain and have highlighted the impor-tant role of astrocytes. Indeed, certain fatty acids (e.g., arachidonic acid and doco-sahexaenoic acid) can be rapidly taken up by brain and oxidized for energy production by astrocytes that express enzymes for fatty acid oxidization. In support of this, octanoate, a medium chain fatty acid, was shown to contribute signi fi cantly to total brain oxidative energy production (Ebert et al. 2003 ) .

3.2 Adenosine and ATP Metabolism

ATP, consisting of three phosphate groups bound to adenosine, is a universal energy source that drives a huge number of biological reactions essential for life and a variety of cellular functions. ATP is integrally involved in neuronal and glial

593 Adenosine and Energy Metabolism—Relationship to Brain Bioenergetics

metabolism—notably the maintenance of transmembrane ion gradients, cAMP formation, kinase-mediated phosphorylation, vesicular storage of neurotransmit-ters, and glutamate metabolism. The high-energy phosphate bonds of ATP are rather labile and thus energy inherent in ATP is readily released when ATP is hydro-lyzed sequentially to ADP, AMP, and fi nally adenosine. Although each of these molecules serves different functions and can activate different signaling pathways, they do in fl uence directly and indirectly cell activity and brain energy homeostasis. Our focus here is on adenosine and brain energy metabolism.

Under basal conditions, brain levels of adenosine (~100 nM) are nearly 10,000-fold lower than ATP (Delaney and Geiger 1996 ; Pazzagli et al. 1993 ; Van Wylen et al. 1986 ) . Therefore, unnoticeable decreases in ATP levels can result in dramatic rises in adenosine levels. In general, adenosine acts to inhibit neuronal activity and in so doing allows target cells like neurons and astrocytes to adjust their energy demands to match better energy supply—this is the basic concept behind adenosine being referred to as a “retaliatory metabolite,” for example, retaliating against exter-nal stimuli and insults capable of causing excessive ATP breakdown (Meghji and Newby 1990 ; Newby et al. 1990 ) .

In brain, de novo synthesis of adenine nucleotides is minimal and for adenosine is absent; purine salvage is absolutely key to the maintenance of cellular levels of these purines. For adenosine, its levels are regulated enzymatically by two enzymes that catalyze its synthesis, 5 ¢ -nucleotidases and S -adenosylhomocysteine hydrolase, and four enzymes that catalyze its removal, S -adenosylhomocysteine hydrolase, 5 ¢ -nucleotidases, adenosine deaminase, and adenosine kinase. In addi-tion to these enzymes, a whole family of equilibrative and concentrative (sodium-dependent) nucleoside transporters are involved in the regulation of intra- and extracellular levels of adenosine (King et al. 2006 ; Latini and Pedata 2001 ; Peng et al. 2005 ) . Equilibrative nucleoside transporters are bidirectional and they facili-tate not only the movement of adenosine into cells when its extracellular concen-tration exceeds its intracellular concentration, but also the ef fl ux of adenosine from cells when its intracellular concentration is increased. Under conditions when Na + electrochemical gradients are reversed, concentrative nucleoside trans-porters facilitate adenosine release from cells. It is important to note that extracel-lular and intracellular levels of adenosine are usually maintained in the same range because most cells possess highly ef fi cient equilibrative nucleoside transporters (Cass et al. 1999 ) .

In addition to neurons releasing ATP as a neurotransmitter, astrocytes too have been identi fi ed as a major source of adenosine formation (Pascual et al. 2005 ) . Adenosine generated from astrocytes originates from vesicular release of ATP (Pascual et al. 2005 ) , direct release of ATP through hemichannels (Iglesias et al. 2009 ; Kang et al. 2008 ) , and adenosine released via nucleoside transporters (King et al. 2006 ; Latini and Pedata 2001 ; Peng et al. 2005 ) and chloride channels (Darby et al. 2003 ) . Astrocytes may also play an important role in enzyme-linked regulation of adenosine levels because adenosine kinase is mainly expressed in astrocytes and is known to control salvage of adenosine to 5 ¢ -AMP (Boison et al. 2010 ; Studer et al. 2006 ) . The intracellular astrocyte-speci fi c enzyme adenosine kinase plays an

60 X. Chen et al.

important role in re-uptaking extracellular (Boison 2008 ; Boison et al. 2010 ) , and the removal of adenosine is predominantly regulated by adenosine kinase via conversion into 5 ¢ -AMP (Boison 2006 ; Pak et al. 1994 ) .

As such, astrocytes are uniquely situated to have a major in fl uence on adenosine levels as they are related to brain bioenergetics. Astrocytes contact synapses and coordinate synaptic networks, and astrocytes release ATP and its subsequent degra-dation into adenosine has a major regulatory function in setting a global adenosine-mediated inhibitory tone within a neuronal network. On the other hand, end feet of astrocytes are an integral part of the BBB, brain endothelial cells express both ade-nosine A

1 and A

2A receptors, and activation of these receptors modulates the perme-

ability of the BBB (Carman et al. 2011 ) . Thus adenosine could have a major role in maintaining BBB integrity, regulating blood fl ow, and modulating the entry of energy-yielding nutrients into the brain (O’Regan 2005 ) .

Adenosine signaling fi rst starts with it binding to speci fi c cell surface G-protein-coupled receptors (Benarroch 2008 ; Boison et al. 2010 ; Fredholm et al. 2005 ) . There are four subtypes of adenosine receptors: A

1 , A

2A , A

2B , and A

3 receptors. The

A 1 and A

3 receptors couple to the G

i family of G proteins and inhibit cAMP forma-

tion; A 2A

receptors couple to members of the G s family, whereas A

2B receptors cou-

ple to many G proteins including G s , G

q , and G

12 . Adenosine is approximately

equipotent on A 1 and A

2A receptors, with A

1 receptors having the highest af fi nity,

whereas A 3 and A

2B receptors require much higher agonist concentrations. Although

all four types of adenosine receptors are expressed in brain, the levels of A 1 and A

2A

receptors are much higher than those of A 2B

and A 3 receptors. Moreover, manipula-

tion of A 2B

and A 3 receptors has relatively modest impact on brain function. Thus,

the impact of adenosine on brain function mostly depends mainly on the actions of A

1 and A

2A receptors.

The actions of adenosine depend on the cellular and subcellular location of ade-nosine receptors. In neurons, adenosine A

1 receptors are located presynaptically,

postsynaptically, and nonsynaptically. Activation of presynaptic A 1 receptors inhib-

its excitatory synaptic transmission by inhibiting N-type and P/Q-type calcium channels (Ambrosio et al. 1997 ; Gundl fi nger et al. 2007 ) . Postsynaptic A

1 receptors

can in fl uence the responsiveness to excitatory stimuli by inhibiting NMDA recep-tors. Nonsynaptic A

1 receptors can stimulate potassium channels leading to neu-

ronal hyperpolarization. Although A 2A

receptors are mostly expressed at neuronal synapses, the activation of A

2A receptors appears to have limited impact on synaptic

transmission but plays a crucial role in controlling synaptic plasticity (Canas et al. 2009 ; Gomes et al. 2011 ; Mills et al. 2011 ) . All four types of adenosine receptors are expressed on brain glial cells, and adenosine can modulate glia function in many different ways including energy metabolism, reactive gliosis, glutamate uptake, and immune responses (Boison et al. 2010 ; Dare et al. 2007 ; Gomes et al. 2011 ) . A

2A

receptors are highly expressed in endothelial cells of brain capillaries, where they play an important role in controlling brain microvascular function (Mills et al. 2011 ; O’Regan 2005 ; Pelligrino et al. 2010 ) .

613 Adenosine and Energy Metabolism—Relationship to Brain Bioenergetics

3.3 Adenosine and Brain Adenylate Energy Charge

There are many different ways to assess brain energy status including adenylate energy charge (EC). Adenylate energy charge is a measure of the phosphorylating power of the adenylate pool, equal to one-half the average number of phosphate anhydride bonds per adenosine moiety present in the pool. It is de fi ned in terms of the concentrations of AMP, ADP, and ATP in the pool and expressed by the equa-tion EC = ([ATP] + ½ [ADP])/([ATP] + [ADP] + [AMP]). The maximum (mostly theoretical) EC value is 1.0, but practically speaking under physiological conditions measured brain adenylate energy charges range to 0.95, meaning that about 95 % of the adenylates are in the form of ATP (Atkinson and Walton 1967 ) . As cellular energy levels drop, ATP levels decrease, and ADP and AMP levels increase; the result is a decrease in EC.

Maintaining brain EC and ATP levels is critical for proper neuronal function, and compromised brain energy levels have detrimental effects on neuronal function, as occur in a variety of acute and chronic neurological and neuropsychiatric disorders such as brain ischemia, sleep deprivation, epilepsy, Huntington’s disease, Parkinson’s disease, Alzheimer’s disease, depression, bipolar disorder, and schizophrenia (Gomes et al. 2011 ) . Because basal brain adenosine levels are nearly 10,000-fold lower than ATP levels, even negligible drops in ATP levels have the potential to produce dramatic rises in adenosine levels. As such, elevated levels of adenosine are observed in all aforementioned neurological and psychological disorders. Although such a rise in adenosine levels could affect the functions of neuron, glia, and endothelial cells in many different ways, here we focus on effects of adenosine on brain bioenergetics. We found that adenosine levels increase as brain energy levels decrease following a variety of excitatory stimuli (Shepel et al. 2005 ) . However, this relationship was not stoichiometric, but rather occurred in three distinct phases, each de fi ned by a linear relationship with a unique slope.

In phase 1, when there is no signi fi cant external excitatory stimulation, brain adenosine levels increase only moderately while brain adenylate energy levels decrease only modestly within a physiological range. For example, moderate increases in neuronal activity can increase the release of glutamate that then can trigger the release of ATP from astrocytes and the rapid formation of adenosine. Such moderate increases in adenosine will preferentially activate adenosine A

1

receptors, inhibit voltage-gated calcium channels, inhibit excitatory synaptic trans-mission, inhibit NMDA receptors, and stimulate potassium channels. As such, neu-ronal activity will be dampened and ATP levels will be conserved.

In phase 2, when there is signi fi cant external excitatory stimulation, brain ade-nosine levels increase rapidly as brain adenylate energy charges drop below the threshold of the physiological range. For instance, when brain energy levels and ATP levels are compromised, as might occur during brain ischemia, sleep depriva-tion, and seizures, a cascade of harmful excitotoxic events may be initiated. During excitotoxicity, released glutamate at synapses perpetuates neuronal depolarization

62 X. Chen et al.

through activation of AMPA and kainate receptors, alters intracellular calcium signaling through activation of NMDA receptors, and limits its removal by revers-ing glutamate uptake. Under such conditions, not only ATP expenditure is increased, but the production of ATP can be also impaired as a result of calcium overload in mitochondria. As such, adenosine levels rise more rapidly and broadly, not just at synaptic terminals but also in glial cells and at the BBB. Unlike in phase 1 where adenosine A

1 receptors at synapses are preferentially activated, in phase 2

as occurs under pathological conditions adenosine A 2A

receptors are upregulated and may mediate the actions of adenosine. At the BBB, activation of A

2A and pos-

sibly A 2B

receptors induces vasodilation, which increases cerebral blood fl ow and brings more oxygen and glucose to the activated brain regions. Activation of ade-nosine A

2A receptors can also increase GLUT1 expression at the brain capillary

endothelial cell and thus promote glucose transport into brain as a fuel (Takagi et al. 1998 ) . In addition to its effects on glutamate clearance, controlling neu-rotrophic factors, and neuro-in fl ammatory effects, short-term activation of ade-nosine A

2A induces glycogenolysis in astroctyes (Sorg and Magistretti 1991 ) and

long-term activation of adenosine A 2B

receptors promotes glycogen synthesis in astrocytes (Allaman et al. 2003 ) , thus affecting brain energy status. Activation of neuronal adenosine A

2A receptors promotes glycolysis and mitochondria metabo-

lism (Hammer et al. 2001 ) , thus increasing ATP production. In contrast to adenos-ine A

1 receptors, activation of neuronal A

2A receptors increases glutamate release

(Marchi et al. 2002 ) and activates NMDA receptors (Rebola et al. 2008 ) , as occurs when neurons are highly activated. As a consequence, neuronal ATP expenditure might increase as a result of activation of neuronal adenosine A

2A receptors. Such

an activation of neuronal adenosine A 2A

receptors could trigger plastic changes in excitatory synapses, de fi ning the threshold for induction of plastic changes in excitatory synapses (Cunha et al. 2008 ) .

In phase 3, when brain is exposed to drastic insults such as global ischemia, brain adenosine levels increase dramatically as brain adenylate energy charge drops below 0.5, a critical level between life and death (Atkinson and Walton 1967 ) . Indeed, drastic insults such as global ischemia and ischemia associated with decapitation reduce brain adenylate energy charge levels to the 0.2–0.3 range (Delaney and Geiger 1996 ; Onodera et al. 1986 ; Verhaegen et al. 1995 ) . Under such conditions, widespread activation of NMDA receptors could facilitate a cascade of excitotoxic events including intracellular calcium dysregulation, free radical production, and inhibition of mitochondrial function, which inhibits energy production, resulting in rapid adenosine formation. Such drastic rises in levels of adenosine may not over-come the loss of brain energy levels, but rather activate low-af fi nity adenosine A

3

receptors and mobilize tissue repair mechanisms including apoptosis and astrocyte proliferation (Abbracchio et al. 1997 ; von Lubitz et al. 1999 ) .

In short, whenever brain energy levels drop adenosine levels rise to adjust brain energy supply and to retaliate against an external stimulus that would cause exces-sive ATP breakdown through activation of different adenosine receptors located on target cells including neurons, glial cells, and brain endothelial cells.

633 Adenosine and Energy Metabolism—Relationship to Brain Bioenergetics

3.4 Dif fi culty in Accurately and Precisely Measuring Levels of ATP and Adenosine

Precisely and accurately measuring levels of ATP and its metabolites including adenosine in discrete brain regions is a critical issue in studying brain bioenergetics. The main challenge in making such measurements is that even brief periods of hypoxia/ischemia cause rapid degradation of ATP and precipitous increases in lev-els of adenosine. Therefore, in order to measure ATP precisely and accurately, brain samples must be obtained either from well-ventilated anesthetized or conscious ani-mals, or from postmortem brains where the enzymes responsible for production and metabolism of adenosine have been instantaneously and completely inactivated.

Studies dating back to the 1950s demonstrated the rapid loss of ATP following decapitation. Kratzing and Narayanaswami (Kratzing and Narayanaswami 1953 ) obtained small pieces of the cortex of young guinea pigs by either killing the ani-mals and dissecting out tissue as quickly as possible or by rapidly freezing the brain with liquid air. Values from the rapidly frozen brains were around 3.0 mmol/kg whereas those from the brain slices were 0.73 mmol/kg. In order to minimize such a loss in ATP various freezing methods were employed and it was found that most of the decrease in ATP levels occurred in the fi rst minute following the ultimate hypoxic/ischemic insult: decapitation. In comparing ATP values from mouse brains rapidly dissected (20 s) and then frozen in liquid nitrogen versus brains obtained from mouse heads that were dropped immediately into liquid nitrogen the levels of ATP were 0.38 mmol/kg for the dissected brains and 1.4 mmol/kg for the rapidly frozen brains (Mandel and Harth 1961 ) . When the heads of decapitated mice were dropped into liquid Freon 12 at intervals ranging from immediately to 10 min the highest values of ATP obtained were 2.5 mmol/kg and were one-half that value with an interval delay of about 6 s and reached a low rather constant level with an interval delay of about 1 min (Lowry et al. 1964 ) . In attempting to obviate these rather dra-matic postmortem decreases in ATP, a method to freeze the brain in situ was devised. Using this method, liquid nitrogen was poured directly onto the skulls of anesthe-tized rats that had received implanted thermocouples so that the freeze front could be timed and the cessation of blood fl ow could be measured (Ponten et al. 1973 ) . At a depth of only 3 mm, blood fl ow continued for 5 to 10 s at which time the tissue had reached 20 °C and it took another 5–6 s for the tissue to freeze. Thus, one would infer that decapitating animals directly into liquid nitrogen would involve 10–15 s of ischemia even at the most super fi cial layers of the brain. Another way of inacti-vating enzymes responsible for adenine nucleotide breakdown and adenosine pro-duction is the “freeze-blowing” technique (Veech et al. 1973 ) , in which high-pressure air is blown into one side of the cranial cavity via a sharp hollow steel probe and the supratentorial brain expelled via another tube on the contralateral side onto a liquid N

2 -cooled aluminum plate, thus immediately halting metabolic reactions at de fi ned

time points on the order of seconds after an intervention. The above-mentioned freezing techniques, while to varying degrees are successful at providing accurate measures of purine levels, do not allow for the precise and accurate measurement for ATP and endogenous adenosine levels in discrete brain regions.

64 X. Chen et al.

An innovative way to measure brain energetics precisely and accurately is instantaneously inactivating the catabolic enzymes with high-energy focused micro-wave irradiation (Delaney and Geiger 1996 ) . Using this technique, brain tissue tem-perature is brought up to an enzyme-inactivating temperature of about 85 °C within a few seconds depending on the size of the animal and the power setting of the head-focused high-energy microwave. They found that a power level of 10 kW heated rat brain tissue to this temperature in 1 s, 6 kW in 2 s, and 3.5 kW in 3 s. The highest irradiation power level used (10 kW for 1.25 s) maintained the brain adenylate energy charge closest to the theoretical maximal value of 1.0. The levels of ATP increased and levels of adenosine decreased as irradiation power increased. In addition, they sacri fi ced rats and extracted tissues using techniques such as decapitation, immer-sion of the decapitated head in liquid nitrogen, and in situ freezing of the brain tissue with liquid nitrogen. The decapitation into liquid nitrogen yielded values around 0.3 mmol/kg; the decapitation followed by dissection and then freezing yielded values around 0.06 mmol/kg; and the brains that were exposed to 10 kW microwave radia-tion yielded values around 3.5 mmol/kg. Because the data reported by Delaney and Geiger were reported in mg/protein, the data showing here were translated, assuming that the protein content of the tissue was 12 % (Sagar et al. 1987 ) .

Even such an extreme method as high-energy focused microwave irradiation creates nonuniform heating throughout the brain with the basal structures of brain heating up before the dorsal structures. As a result, purinergic enzymes would not be denatured as quickly in areas of brain that do not reach 85 °C and therefore ATP levels would be lower and adenosine levels would be higher. Another drawback of the high-energy focused microwave irradiation method is its inability to differenti-ate intracellular and extracellular levels of ATP and its metabolites, because brain regions are homogenized and analyzed as a whole.

Alternatively, levels of ATP and its metabolites can be measured in vivo in well-ventilated anesthetized or conscious animals. The fi rst of such measurements are the brain microdialysis and cortical cup methods, which allow for direct measurement of ATP and adenosine from brain interstitial fl uid (Lutz and Kabler 1997 ; Melani et al. 2005 ; Pazzagli et al. 1993 ; Van Wylen et al. 1986 ) . Such methods can also inform as to the temporal sequence of release events and whether ATP is released before adenosine or vice versa. The disadvantages of such techniques are the tissue trauma caused by the implantation of the dialysis cannula and poor temporal resolu-tion in detecting ATP and adenosine levels. The development of adenosine and ATP biosensors allows for direct and real-time measurements under physiological condi-tion, which provide a higher temporal and spatial resolution in detection of adenos-ine and ATP levels (Dale et al. 2000 ; Frenguelli et al. 2003 ; Frenguelli et al. 2007 ; Llaudet et al. 2003, 2005 ) . However, purine levels measured by those biosensors are not true concentrations of adenosine and ATP, but rather relative values.

In addition to direct measurement of ATP and its metabolites to assess brain energy status, it is also critical to measure accurately and precisely the brain energy reserve, glycogen. Glycogen can be visually identi fi ed in brain sections using a cytochemical staining method that relies on the interaction of glycogen with osmium tetroxide (Cataldo and Broadwell 1986 ; Koizumi 1974 ; Wender et al. 2000 ) or the

653 Adenosine and Energy Metabolism—Relationship to Brain Bioenergetics

use of periodic acid–Schiff’s base (Kong et al. 2002 ) . The absolute glycogen content can be measured using quantitative methods, which have traditionally been carried out using a biochemical assay on postmortem tissue. However, glycogen degrades extremely rapidly, and such degradation must be halted in order to measure glyco-gen content accurately and precisely. Similar freezing methods as those used to inactivate enzymes responsible for adenine nucleotide breakdown and adenosine production have been used for quantitative glycogen measurement. Such methods include immersion of whole animals or decapitated heads in liquid nitrogen (Passonneau and Lauderdale 1974 ) or Freon-12 (King et al. 1967 ) , freezing brains in situ using liquid nitrogen poured directly on the skull (Cruz and Dienel 2002 ) , and rapidly removing brain tissue at 0 °C followed by immediate freezing (Wender et al. 2000 ) . The high-energy focused microwave irradiation, which denatures enzymes responsible for glycogen metabolism preserving glycogen content at the time of irradiation, has also been used (Sagar et al. 1987 ) . Another critical step in accurate measurement of glycogen is to minimize animal handling prior to assay (Cruz and Dienel 2002 ) . The most accurate method for measuring glycogen is deb-ranching glycogen into glucose followed by standard biochemical assay of glucose (Passonneau and Lauderdale 1974 ) , with glycogen being expressed as glucosyl units, as the debranching enzymes yield one glucose molecule per glucosyl unit of the glycogen molecule. The manner in which glycogen content is expressed can be confusing, since glycogen is often expressed as glucosyl units either relative to a normalized protein concentration (pmol glycogen/ m g protein) or relative to the wet or dry weight of the tissue ( m mol glycogen/g wet tissue). Assuming that wet brain tissue contains about 12 % protein content (Sagar et al. 1987 ) , and that dry brain tissue contains about half the protein of wet brain tissue, a simple calculation could allow interconversion.

Alternatively, glycogen content can be estimated in vivo using a noninvasive nuclear magnetic resonance (NMR) method, which detects the 13 C glucose incorpo-ration into the brain glycogen. In such a method, 13 C-labeled glucose is injected systemically and the labeled glucose is slowly incorporated into the brain glycogen pool producing a detectable glycogen signal that can be continuously monitored (Choi et al. 1999 ) . The advantage of this noninvasive technology is that it could be used in humans to record glycogen measurements under both pathological and physiological conditions. However, the observed increase in 13 C signal over time might not necessarily be re fl ecting changes in glycogen concentration alone, but also an increased incorporation of the 13 C label. A new approach was recently devel-oped that allows the measurement of in vivo brain glycogen concentration more accurately (Morgenthaler et al. 2008 ) . In this method, the in fl uence of glycogen turnover, a potential confounding factor in the 13 C NMR, was minimized by pre-labeling the animals and achieving a near steady-state isotopic enrichment using long-term [1- 13 C]-glucose administration.

In summary, it is not an easy task to measure brain energy status accurately and precisely. Essential components in assessing correctly brain energetics should include justifying carefully the methodology used and putting the data in the con-text of what is already known about brain energy metabolism.

66 X. Chen et al.

Acknowledgments The work currently conducted in our laboratories is supported by 2P20RR0017699 from the NCRR, a component of the NIH, and by R01NS069597.

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