Chapter 1 Cellular ATP

Chapter 1

Cellular ATP DAVID A. HARRIS

Introduction Structure of ATP Chemical Bonds and Conformation What Makes ATP a Good Energy Source? Other Features ofthe ATP Molecule Measurement of Cellular ATP The Freeze-Clamp Technique The Magnetic Resonance Technique Adenine Nucleotide Concentrations Within Cells Spatial Distribution Uses of ATP Contraction of Actomyosin Ion Pumping ATP in Biosyntheses ATP as Phosphate Donor ATP as Charge Neutralizer ATP and Messenger Molecules Structural Role of ATP Reactions Involving Exchange of High Energy Phosphates Creatine Kinase Adenylate Kinase Synthesis of ATP Substrate Level Phosphorylation Oxidative Phosphorylation

Principles of Medical Biology, Volume 4 Cell Chemistry and Physiology: Part II, pages 1-47 Copyright © 1996 by JAI Press Inc. All rights of reproduction in any form reserved. ISBN: 1-55938-806-4

2 3 3 3 6 7 7 8 10 10 13 13 16 20 22 23 24 25 25 26 27 28 28 31

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Control of ATP Levels ATP Levels Are Closely Maintained In Vivo Is ATP a Regulator? Control of Anaerobic ATP Production Controlof Aerobic ATP Production Pathological Disturbances of ATP Levels Malignant Hyperthermia (Malignant Hyperpyrexia) Ischemia Summary

35 35 35 36 40 43 43 43 44

INTRODUCTION In living cells, a variety of processes yield energy. In man, these are typically oxidations (of glucose, amino acids, and fatty acids), although some energy is also produced by the anaerobic breakdown of glucose to lactate (anaerobic glycolysis). The amount of energy yielded in the these processes is very variable—complete oxidation of palmitic acid yields 9,500 kJ/mol, while the conversion of glucose to lactate yields about 170 kJ/mol. Conversely, a variety of processes in the living cell require energy. These include the biosynthesis of small molecules (e.g., glucose from pyruvate) and of large biopolymers (e.g., proteins from amino acids), the transport of molecules and ions, and the performance of mechanical work (e.g., muscle contraction). Because of variety both in the chemical nature of catabolic processes, and in their energy yield, these reactions cannot be used directly to drive the variety of energy-requiring processes. In essence, energy released in catabolic processes is trapped in units of 50-60 kJ/mol, by synthesizing ATP from ADP+Pi, and used in these units in biosyntheses, ion pumping, mechanical processes, etc. Enzymes involved in the latter processes are thus adapted to accept ATP as a convenient and common unit of exchange between themselves and the variety of energy-yielding processes; ATP is sometimes known as the energy currency of the cell. ATP is a short-term store of energy within the cell; the cell content of ATP turns over about once every second. The other short-term energy store in cells is the transmembrane ion gradient, in particular the Na"*" gradient across the plasma membrane and the H"^ gradient across the mitochondrial membrane. The amount of energy stored in these gradients (comprising contributionsfromboth concentration and voltage gradients) is about 15-20 kJ/mol, i.e., 15-20 kJ is released when one mol of ions moves downhill. Thus, energy in ion gradients is stored in smaller units than it is in ATP. However, this seems convenient for most transport processes in animal cells; the plasma membrane Na^ gradient can be used as an energy source for accumulation of glucose and amino acids from the blood. Thus, ion gradients can serve as an energy source in some biological processes. Compared to ATP, however, they are far less versatile in their application. As noted above, the unit of energy stored per ion in a gradient is 15-20 kJ/mol, about one third that per mol ATP. More importantly, this energy is not portable. A gradient can drive processes only at the membrane across which it is located; it cannot power

Cellular ATP

3

the bulk of chemical reactions in the cell, which occur in free solution. Thus, ATP is pre-eminent as a diffusible energy source for biochemical processes.

STRUCTURE OF ATP Chemical Bonds and Conformation The chemical structure of ATP is shown in Figure la. The molecule consists of three notional parts, the purine base adenine, the pentose sugar ribose, and a chain of three linked phosphate groups. The same adenine-ribose-phosphate structure is found in nucleic acids (RNA), and thus the molecule belongs to the class of nucleotides; other aspects of nomenclature are indicated on the figure. In solution, ATP can adopt a variety of conformations, in particular due to rotation about the base-sugar bond (a in Figure la) and to variations in orientation of the phosphate groups. A preferred conformation is the extended form (Figure lb), with the base and to the sugar ring and the phosphate groups extended; in this form it binds to many proteins. Introduction of a bulky group into the five membered ring of the purine (e.g., in 8-bromo ATP) tends to favor the syn conformer, which binds less well to proteins. Normal cytoplasmic conditions are around pH 7.1, with p[Mg^"^]totai * 2.3. This means that >90% of cytoplasmic ATP exists as the fully ionized MgATP^~ complex. All enzymes that use ATP utilize the MgATP^~ complex rather than free ATP, with the exception of the mitochondrial ATP<->ADP exchanger, which utilizes the small amount of ATP"^ in equilibrium with the MgATP^" complex. This ensures that Mg^"*" levels inside mitochondria can be maintained independently of ATP synthesis rates. Under some physiological conditions (e.g., heavy exercise), intracellular pH may fall and MgATP^~+ H"^ MgATP(H)~ equilibrium may shift in favor of the protonated form. This may affect the availability of cellular ATP, although the magnitude of such effects are as yet unknown. What Makes ATP a Good Energy Source? In regard to energy transfer, the critical part of the ATP molecule is its phosphate tail, and, in particular, its two terminal phosphate groups (P and y phosphates). Each is linked to the neighboring phosphate by an acid anhydride link. (Note that phosphate is simply the ionized form of phosphoric acid.) Since acid anhydrides are (thermodynamically) unstable in water, they can serve as a source of energy. Quantitatively, we consider the hydrolysis ATP + H2O-^ ADP + Pj.* Conventionally, Mg^"^ ions and H"^ ions, which are buffered in the cell, are omitted from this equation. Thus, Pj indicates the prevailing ionization state of inorganic phosphate (HPO4") and ATP indicates the complex MgATP^".

PHOSPHATES

OH

OH

Figure 1, The structure of ATP. (a) Chemical structure of the MgATP complex, showing nomenclatures used. Rotation around bond (a) converts antl and syn conformers, (b) Conformation of ATP bound to an enzyme (aspartate transcarbomoylase). Note that (i) the planes of the adenine and ribose rings are at right angles; (ii) the adenine and ribose rings lie syn to each other; and (lii) the phosphate chain is extended.

Cellular ATP

5

The change in free energy (energy available for work) is given by AG = AG°' + RT ln[ADP][Pi]/[ATP]. AG°' is a term reflecting the chemical nature of the compound under consideration; for ATP hydrolysis it is around —30 kJ/mol, larger in magnitude than the value for phosphate esters (e.g., sugar phosphates) of-15 kJ/mol. This is due to the anhydride nature of the bond hydrolyzed, which allows increased resonance stabilization (electron derealization) and increased hydration in the products ADP + Pj, stabilizing them relative to the reactants ATP and H2O. In the cell, ADP levels are kept low such that the ATP/ADP ratio « 200 in the cytoplasm. This means that the actual free energy yielded on ATP hydrolysis in vivo (AG in the above equation) is larger in magnitude than AG°', due to the contribution from the second term in the equation. The free energy released on hydrolysis of intracellular ATP, often written as the phosphorylation potential AGp, is typically -55-60 kJ/mol ATP (Veech et al, 1979): AGp = -30 + RT In [Pi]/100 = -^0 kJ/mol at 37 °C and typical cellular free [Pj] = 1 mM.* ATP is one of a number of cellular phosphates with a highly negative free energy of hydrolysis. It is convenient to designate ATP and compounds with similar AG°' values for hydrolysis (e.g., GTP, UTP, etc.) or greater in magnitude (creatine phosphate, 1,3 diphosphoglycerate, phosphoenol pyruvate) as high energy phosphates; they can all, without fiirther energy input, generate ATP, which can then be used to drive cellular processes. The sum of concentrations of all these high energy phosphates is an indication of the energy status of a cell. The second essential feature of ATP as a temporary biological energy store is its kinetic stability. It is obvious that kinetic stability must be a feature of an energy store: there is no point in producing a high energy compound which hydrolyzes rapidly before it can be used. However, the concept of a thermodynamically unstable and kinetically stable compound might appear patadoxical. It can be understood by consideration of an analogous system, a mixture of hydrogen and oxygen. Such a mixture is kinetically stable: it could stand for thousands of years at room temperature with no noticeable change. However, given a catalyst, or a spark, it changes chemically with the release of large amounts of energy. Similarly, left to itself, ATP is stable in solution for several days, but, given a suitable (enzyme) catalyst, it hydrolyzes to yield large amounts of energy. The kinetic stability of ATP (as compared to, for example, acetic anhydride in water) is chemically due to the high (negative) charge density around the phosphate groups, which discourages the approach of nucleophiles.

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Other Features of the ATP Molecule

As noted above, ATP is only one of a series of high energy phosphates found within cells (Table 1). It is, however, by far the most versatile, being formed in the bulk of energy-yielding reactions (mitochondrial oxidations) and used in most energy-requiring processes. Although occasionally other compounds may serve directly as an energy source (e.g., phosphoenol pyruvate drives some bacterial transport systems; GTP drives several steps in protein synthesis), such compounds are usually used to generate ATP. The favored role of ATP may be rationalized in several ways: 1. Since it contains rwo acid anhydride links, ATP may hydrolyze either to ADP + Pj or AMP + 2Pi. Thus, the occasional reaction requiring a driving energy of more than 50-60 kJ/mol can be driven by splitting both anhydride bonds. (This is not possible for phosphocreatine, 1,3 diphosphoglycerate, etc.) 2. The adenine ring plays no part in the chemistry or energetics of ATP function; adenosine triphosphate, however, is used much more widely than GTP, CTP, etc. This may be an accident of history, with adenine appearing, by chance, early in prebiotic evolution. However, it is interesting that adenine appears in the structure of a variety of other coenzymes (NAD,

Table 1. Standard Free Energy of Hydrolysis for Biochemical Compounds Compound

AC^' (kj/mol)

phosphoeno/pyruvate ATP (-^ AMP + 2Pi) 1,3 diphosphoglycerate phosphocreatine fatty acyl Coenzyme A (-> fatty acid + CoA) amino acyl tRNA (-• amino acid + tRNA) ATP GTP, UTP, CTP PPi

-61 -58 -49 -43 -35 -35 -31 -31 -28

glucose-6-phosphate AMP glycerol-1 -phosphate

-14 -9.6 -9.2

Note:

Except where indicated, the reaction considered is X - P + H2O - » X + Pi. Compounds above the line are designated high energy compounds in biochemistry (see text). Note that the actual free energy change for hydrolysis of these compounds in vivo (e.g., AGp for ATP hydrolysis) is normally greater than the change under standard conditions, AC°', given here.

Cellular ATP

7

FAD, coenzyme A) where again it plays no part in the reaction. Perhaps it provides a particularly favorable recognition site for enzymes. 3. ATP is an acid anhydride. A major requirement for energy in macromolecule biosynthesis is in driving condensation reactions (removal of water) in an aqueous environment. Formation of a peptide bond, for example, is a dehydration: R-COOH + NH2-R' ^ RCO-NH-R' + H2O. The anhydride nature of ATP allows it to be a good dehydrating agent even, given a suitable reaction mechanism (see below), in an aqueous environment. 4. ATP serves as a source of phosphate groups in biochemical reactions. For example, glucose, on entering the cell, is phosphorylated to glucose-6-phosphate, giving it a negative charge which helps to retain it within a compartment bounded by the (lipophilic) cell membrane. Many metabolic pathways (e.g., glycolysis, histidine biosynthesis) utilize phosphorylated intermediates in this way to limit diffusion out of the cell. A contrasting example is the phosphorylation, by ATP, of enzymes such as glycogen phosphorylase which are switched on (or off) by this process. In both these cases, the important feature of ATP is not its tendency to transfer phosphate to water (high negative free energy of hydrolysis) but its tendency to phosphorylate other hydroxyl groups (high phosphate transfer potential). The energetic role of ATP in these phosphorylation reactions is to ensure the reaction is driven to completion; the loss in free energy in generating a phosphate ester in place of an anhydride is dissipated as heat.

MEASUREMENT OF CELLULAR ATP The Freeze-Clamp Technique Classically, measurement of ATP levels within cells and tissues has involved (a) the rapid arrest of metabolism and of enzyme activity in the tissue; (b) extraction of ATP from the tissue (without destroying it); and (c) assay of its concentration by enzymatic procedures or by high performance liquid chromatography (HPLC). Since the energy status of a tissue is also dependent on ADP, AMP, and Pj concentrations, these are generally measured with ATP in a single extract. This technique is highly sensitive; using firefly luciferase (bioluminescent assay) the ATP content of only a few hundred cells can be measured. This is useful when biopsy material is being studied. In a typical procedure, the tissue is perfused with an oxygenated buffer/salt solution and manipulated (e.g., electrically stimulated, treated with a drug) as desired. The tissue is then rapidly frozen, by crushing it between two flat aluminum

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plates at liquid nitrogen temperatures, to arrest metabolism. The frozen, powdered tissue is deproteinized with perchloric acid (to remove enzymes) and the soluble extract (containing the tissue metabolites) neutralized. ATP, ADP, etc. are separated by HPLC and detected by ultraviolet absorption. This approach has some disadvantages. Since ATP turns over in the cell within one second, the tissue must be maintained under physiological conditions (oxygenated, neutral pH) until metabolism can be instantaneously stopped. Organ preparations such as heart and muscle from small animals can be perfused, both outside and inside the animal. Muscle biopsies from humans (and in particular post-mortem tissue), in contrast, will not accurately reflect ATP levels in vivo. Secondly, this approach is invasive. It requires removal and destruction of the tissue under investigation which, aside from the obvious clinical problems, means that ATP levels cannot be followed over time in a single tissue. In experiments where time dependent changes are to be followed, multiple tissue samples (and statistical methods of analysis) are required. The Magnetic Resonance Technique The magnetic resonance (NMR) technique utilizes the ability of phosphorus (^^P) nuclei, when placed in a high magnetic field, to absorb radio waves. The wavelength (frequency) absorbed depends on the chemical environment of the nucleus; Pj, phosphocreatine (PCr), and the three phosphorus atoms in ATP will each absorb radiation (shown by peaks on an NMR spectrum) at slightly different wavelengths. The intensity of absorption (peak area) is proportional to the amount of material present which absorbs at that wavelength; thus, from the corresponding peak areas, the amounts of P,, PCr, and ATP in a sample can be quantitatively assessed (Radda, 1986). Since tissues are transparent to magnetic fields and to radio waves, this technique can be used to measure phosphates within the body, i.e., this technique is noninvasive. An arrangement for measuring metabolites within a human arm is shown in Figure 2a. Measurement is clearly made under physiological conditions, without having to freeze metabolism. Furthermore, since spectra can be taken within a few seconds, and the tissue is not altered in the process, the levels of ATP, etc. can be followed in time. Figure 2a shows, in fact, an arrangement for measuring levels of phosphate metabolites within arm muscle, and Figure 2b shows variations in these metabolites, during and after exercise. The main problem with the NMR method is its relatively low sensitivity. It requires gram quantities of tissue, and metabolite concentrations within the tissue of 1 mM or above. Thus, although it will detect Pj, ATP, and PCr, the technique is not sensitive enough to measure ADP or AMP levels, which typically lie below 100 iLiM.

Cellular ATP ^,

V V ^\ V

^^^^ ^f magnet

1

—)

a

blood pressure cuff (ZOOrnm Hg)

PCr,

RECOVERY

pH704 REST Figure 2. NMR measurement of ATP in human organs, (a) Device for exercising human arm in bore of NMR magnet, (b) NMR spectra of phosphate metabolites in human arm during and after anaerobic exercise. Note that nearly all the signal derives from muscle metabolites. During exercise, PCr levels are seen to fall, and ?, levels to rise, while [ATP] is hardly affected. Numbers denote intracellular muscle pH, which also falls due to lactic acid production.

ADP levels in muscle or brain may be calculated from NMR data, assuming creatine kinase to be at equilibrium in the cell, from the equilibrium relationship: Keq = [ATP][Creatine] / [ADP][PCr]. In these calculations, the value of Kgq is a known constant (66 at pH 7.1, 37 °C, etc.) and [ATP], [PCr] are measured by the NMR experiment. The concentration of free creatine must be measured enzymatically after extraction of the tissue (as above); normally it is measured as total (Cr and PCr) creatine at the end of the experiment. Note, how^ever, that this calculation is possible only for those tissues (muscle, brain) which contain creatine kinase. In other tissues, [ADP] must be measured by the freeze-clamp procedure.

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Adenine Nucleotide Concentrations Within Cells Typical values for ATP, ADP, AMP, Pj, and PCr concentrations in heart muscle, measured by each of the above techniques (Veech et al, 1979; Balaban et al., 1986) are given in Table 2. ATP levels at 8 mM are quite high relative to other metabolites (glucose-6-phosphate at 0.5 mM, citrate at 0.1 mM), reflecting the importance of this metabolite in a variety of metabolic processes. Similar ATP levels are observed in many tissues of the body. There is a clear discrepancy in Table 2 between the levels of ADP measured enzymatically (1.4 mM) and the levels calculated from the PCr/ATP equilibrium (0.04 mM). This reflects the fact that the freeze-clamp method extracts total ADP from the tissue, while the equilibrium calculation considers only that part of ADP in equilibrium with ADP and PCr, i.e., ADP that is free in solution. These figures differ, therefore, because most cellular ADP is bound to protein (largely actin) within cells. Since it is free ADP which is a substrate for ATP synthesis—which participates in the equation for the phosphorylation potential, AGp, and which regulates enzymes—^it is the calculated value which is taken as an indicator of cellular energy status. The ratio [ATP]/[ADP]free within these cells is 200 and AGp = -60 kJ/mol ATP hydrolyzed. Again both values are typical not only in heart but in a variety of tissues. Finally, despite the relatively high concentrations of ATP and PCr (together making up about 5% of the dry mass of the heart), their role as an energy store can only be short term. A rat heart uses about 2% of its high energy phosphate per beat; at this rate ATP would last about 3 seconds and PCr about 9 seconds more. Thus ATP generation, from metabolic fuels, must be rapid and continuous in heart as in all other tissues. Spatial Distribution One drawback to both techniques as described above is that they provide only an average value of nucleotide concentration across the tissue. This will obscure

Table 2. Adenine Nucleotide Levels in Rat Heart NMR measurements freeze-clamp methods

ATP

ADP

8 mM^ 8 mM

0.04 mM^ 1.4 mM

ASAP

n.d."^ 0.1 mM

Pi

0.5 mM 2-8 mM

PCr 23 mM 25 mM

Notes: ^Absolute estimation by freeze clamp methods. P,, PCr determined by peak areas relative to ATP peaks. ^Calculated from creatine kinase equilibrium (see text). ''Undetectably low by NMR. Measured values overestimate free phosphate due to some destruction of high-energy phosphates and contamination with extracellular phosphate.

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any differences between cell types within the tissue, between different compartments (e.g., mitochondria and cytoplasm) within the cell, or (particularly in the case of Pj) between the intracellular and extracellular fluids. In many cases, this is unimportant. Adenine nucleotides are present in significant concentrations only within cells, so amounts in the perfusing medium can be ignored. Similarly, mitochondria occupy only a small fraction of cell volume, and thus contain only a small fraction of its metabolites; the figures in Table 2 represent, to a close approximation, cytoplasmic concentrations. Where more precise data are required, the above techniques must be modified as outlined below. Fractionation in Non-Aqueous Solvents In a technique pioneered by Hassinen and coworkers (Kauppinen et al., 1980), a tissue after freeze clamping has the cell water replaced by organic solvents (e.g., heptane/CCU). This prevents both enzyme function and exchange of metabolites between cell compartments. Cells are then fractionated into mitochondria, nuclei, etc., by homogenization and centrifugation—still in organic solvents—^and only then are aqueous extracts made for nucleotide assay. Rapid Cell Lysis/Centrifugal Fractionation This technique (Siess and Wieland, 1976) is suitable for cultured cells in suspension (e.g., hepatocytes). At the time of measurement, cells are mixed with a small amount of digitonin, which ruptures the plasma membrane. A sample is immediately added to the upper aqueous layer of a micro centrifuge tube; this layer is separated from a lower aqueous phase by a layer of (inert) silicone oil (Figure 3). The tubes are then centrifuged, and the unlysed mitochondria spin through the oil into the lower phase (normally perchloric acid), while the cytoplasmic contents remain in the upper aqueous phase. The aqueous phases can then be separately assayed for mitochondrial and cytoplasmic nucleotides. These two methods demonstrate that, in a variety of tissues, the ATP/ADP ratio within mitochondria is around 1:1, much lower than in the cytoplasm, where this ratio is about 200:1 (Table 2). Since ATP is made inside the mitochondrion and exported into the cytoplasm, the relatively low levels of mitochondrial ATP were unexpected. Their explanation lies in the energy dependent system for ATP export/ADP import across the mitochondrial membrane, which continually expels ATP in exchange for ADP (see below). Magnetic Resonance Tomography By using, in effect, a point source of radiofrequency radiation (a surface coil) in the NMR experiment, and a rotating receiver for data collection, ^^P-NMR spectra

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cytoplasm silicone oil acid mitochondrial pellet

Figure 3. Rapid lysis/centrifugation technique for investigation of metabolite compartmentation. For explanation, see text. can be compiled for various depths within the body (typically 1—5 mm slices). By this method, (analogous to the use of X-ray tomography to map tissue density within the body), differences in the levels of PCr and ATP between cells can be mapped (Radda, 1992). As an example. Figure 4 shows a series of stacked plots, showing the levels of phosphorus metabolites at various depths within the human thorax. The changing

sChest wall muscle Surface coil phantom 2.3DPG

PDE

N

i

l

Blood Heart -• Skeletal muscle

Surface coil

Figure 4. Spatial resolution of phosphate metabolites in the human thorax. Phosphate metabolites were measured in 1 mm slices through the thorax, using ^^P-NMR tomography. The body surface is marked by the surface coil phantom. Muscle ATP is indicated by the three ridges on the right (see Figure 2). The two peaks in the ridge due to PCr mark the chest muscle and the heart muscle; the PCr/ATP ratio of skeletal muscle is seen to be higher than that of heart muscle. Note the left rearmost peak of 2,3 diphosphoglycerate (2,3 DPC); this is an allosteric regulator of hemoglobin, occurring in the heart ventricular blood. The ridge labeled PDE is due to phosphodiesters.

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intensity of the PCr peaks identifies the chest wall (skeletal muscle) and the heart, comparing these with the ATP peaks (to the right on the diagram), we see that skeletal muscle has a higher PCr/ATP ratio than heart muscle. At the very back of the diagram, on the left, we see the heart cavity, marked by a high concentration of 2,3-diphosphoglycerate (bound to hemoglobin in the blood). In a similar, but more precise, study in canine heart (Robitaille et al., 1990), it has been shown that ATP levels in heart muscle remain constant from the outside to the inside of the heart; PCr levels, in contrast, fall significantly towards the endocardial side. This indicates a gradient in PCr and ADP levels from the subepicardial cardiomyocytes (high PCr, low ADP) to the subendothelial cells (low PCr, high ADP). This reflects subtle differences in energy metabolism across the myocardium.

USES OF ATP Contraction of Actomyosin The actomyosin system is designed to convert chemical energy (from ATP hydrolysis) directly into mechanical work. In skeletal and heart muscle, the cells (fibers) are packed with a dense, semicrystalline array of actomyosin, and this protein is responsible for up to 70% of ATP consumption in contracting muscle. (A remaining 20% is consumed in ion pumping; see below.) In other tissues, actomyosin filaments may also contribute to cell motion, e.g., in phagocytes and fibroblasts, but the filaments are organized only locally within the cell, and contribute much less to overall cellular energy consumption. The operation of actomyosin is described in the cross bridge model. In this model, ATP drives the release of the myosin head from one subunit in the actin filament. This is followed by a conformational change in the myosin head such that it rebinds to a different subunit of actin, and a tension generating step in which the myosin returns to its original conformation attached to this new site along the actin filament. ATP and the Energetics of Muscle Contraction It is instructive to consider how ATP powers this overall process, because this also serves as a paradigm for harnessing ATP hydrolysis to drive processes such as ion pumping (see below). It is tempting to imagine that cleavage of the bond between p and y phosphate groups directly energizes proteins in some way, since hydrolysis of this bond in free solution yields energy. This is, however, to reckon without the ability of enzymes to juggle the energy of intermediates in reaction pathways.

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DAVID A. HARRIS

r\

T ATP

AGp

ADP^Pi

*P.^

Figure 5. Energy release during ATP hydrolysis, (a) Energy release during hydrolysis of ATP in the absence of an enzyme, (b) Energy release during hydrolysis by myosin (E). Energy is released as ATP binds tightly to the enzyme. The enzyme not only decreases activation energy AG*, but shifts energy release from the expected, bond splitting step (shown dotted) to ATP binding (solid line). Note that the total energy release (AGp) is identical in each case, 'indicates an unstable intermediate species.

Thus, in myosin, hydrolysis of ATP involves four steps, not only (a) cleavage of the ADP-Pj bond, but also (b) formation of an ATP-myosin complex, (c) release of Pj, and (d) release of ADP from the myosin-ADP-Pj complex. The first law of thermodynamics tells us that AG for ATP hydrolysis is fixed (-60 kJ/mol) under cellular conditions; that is to say, when ATP is hydrolyzed to ADP + P,—with or without enzyme—60 kJ/mol must be liberated. It does not, however, tell us at which of these four steps energy is liberated. In the case of myosin, the affinity of the enzyme for ATP is so high (10^ x stronger than for ADP), that nearly all its energy is liberated when ATP binds to myosin before bond cleavage occurs. This is shown schematically in Figure 5. We have thus established the pattern of energy release from ATP during catalysis by myosin. How does this help us understand the contraction mechanism? The answer to this lies in the strength of the actin/myosin interaction. If muscle ATP is depleted (e.g., by stimulation in the presence of metabolic inhibitors), the muscle becomes rigid (tetanus). This is because all the myosin heads are fixed in their complex with actin. Thus, the actin-myosin complex, in the absence of ATP, is a strong one—it is energetically favored. When ATP binds to myosin, sufficient energy is released to dissociate the actin-myosin complex and initiate the cross bridge cycle (Figure 6a). Thus myosin exchanges a favorable interaction with actin for another with ATP; this is brought about mechanistically by the binding of ATP inducing a conformational change in myosin which distorts the actin binding site. Subsequent chemical changes in the myosin-ATP complex (ATP cleavage. Pi release, and ADP release) simply act as triggers for conformational changes in

Cellular ATP

15

myosin conformation (tilting the head, rebinding to actin, and realignment of the myosin molecule) as shown in Figure 6a; tension development occurs at the final stage in the scheme (Hibberd and Trentham, 1986). Energy changes in the system are shown in Figure 6b. ATP binding to myosin liberates energy which is used to dissociate the actin/myosin complex. Dissociated actin and myosin thus constitutes an energized system. Recombination of actin and

hydrolysis

M

^sj^•^'''P

Au-Mro^ A+MC2)

...*/-.N AM*C3)

tension I• C I •I • I

30 k>

'

AM (1)

1

AM (4)

Figure 6. Mechanism of ATP use in muscle contraction, (a) The cross-bridge model for muscle contraction, incorporating ATP binding and hydrolysis steps. Note that each chemical change (ATP binding, bond splitting, ADP, and Pj release) produces a (kinetically) unstable conformation (*), which relaxes in the next stage of the cycle. The numbers (1), (2), etc. represent different conformational states (see text, and Figure 6b). T represents bound ATP, and D represents bound (ADP + Pj). (b) Energetics of the cross-bridge model. Energy is transferred from ATP (solid line) to actin (A) + myosin (M) by dissociation of actomyosin (AM) (dashed line), and on to tension development (dotted line) after reformation of the AM complex. For simplicity, the dissociations of Pj and ADP are shown as a single step.

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D A V I D A . HARRIS

myosin, permitted after ATP hydrolysis and conformational changes in the myosin head, thus releases this energy which is used in tension development. Ion Pumping

Most small molecules and ions are moved across cell membranes by a variety of porters (symports, antiports, etc.) using energy stored in transmembrane ion gradients (see above). Examples include the gut glucose/Na"*" symport and the Na'^/Ca^'^ antiport at the plasma membrane, and the Pj/H"^ symport and Ca^"*" uniport at the mitochondrial membrane. Some transport systems, however, are powered directly by ATP hydrolysis. These are known as primary transport systems because, typically, they build up the ion gradients (e.g., Na^ across the plasma membrane) which drive the secondary transport systems described above. Examples of such ATP-driven pumps are shown in Figure 7. NrfA

drugs

>NQ

&

P-type

85 ABC-type

fi

F-typg

tt

V-type

Figure 7. ATP driven pumps in eukaryotic cells. The pump orientation is indicated by the position of the ATP binding domain(s) (shown circular); e.g., cytoplasmic-facing for P,V and ABC type pumps. The direction of pumping is indicated by the arrows. H"^ ions are pumped where no specification is given; the plasma membrane Na"^ pump also pumps K^ ions inwards. Mito = mitochondrion; SR = sarcoplasmic reticulum; PM = plasma membrane; SV = storage vesicle (e.g., chromaffin granule, synaptic vesicle); EV = endocytotic vesicle.

Cellular ATP

17

In gross structure, all of these pumps appear similar; they comprise a transmembrane region surmounted by a large aqueous domain facing the cytoplasm. At the molecular level, however, they fall into four distinct families. P'Type Pumps In terms of net energy consumption, P-type pumps are the major ATP-driven transport systems in mammals. They include the ubiquitous NaVK"*" pump which maintains Na"^ and K"^ gradients (and thus the steady state membrane potential) across the plasma membrane. This pump is responsible for up to 40% of all ATP utilization in the brain, rather less in other tissues. Other examples include the Ca^^ pump of sarcoplasmic reticulum (responsible for up to 20% of ATP utilization in active muscle), and the gastric H^ pump, which acidifies the stomach lumen. P-type pumps contain a long polypeptide chain (about 1,000 amino acids) which traverses the membrane up to 10 times. The polypeptide contains its ATP binding site on the large extramembrane domain (between transmembrane helices IV and V) and ion binding sites (ionophores) within the membrane domain (MacLennan, 1990). In some cases, notably the Na'*'/K'^ ATPase, a second smaller polypeptide (P) is present but its function is unknown. P-type pumps are unique in carrying out ATP hydrolysis in two stages, the first involving phosphorylation of the enzyme on an aspartic acid residue (asp 351). The mechanism is thus (i) E + ATP -> E-P + ADP

(ii) E-P + H2O -^ E + Pj

This may be useful in partitioning the energy of ATP hydrolysis between movements of different ions (see below). V-Type Pumps V-type pumps pump only protons (H"^); they are responsible for the acidification of intracellular compartments. An example is the chromaffin granule ATPase, which occurs in the epinephrine-storage vesicles (chromaffin granules) of the adrenal medulla. H^ pumped into these vesicles allows them to trap catecholamines as the charged protonated form which cannot cross the vesicle membrane. Acidification of endocytotic vesicles, synaptic (acetylcholine storage) vesicles, and lysosomes involves related pumps. V-type ATPases contain upwards of 10 separate polypeptide chains, and are separable into a soluble headpiece (containing the ATP binding site) and a transmembrane sector (containing the H"^ channel) (Nelson, 1992). In these, as in the remaining pumps in this section, ATP hydrolysis occurs by direct attack of water on ATP; no phosphorylated enzyme occurs.

18

DAVID A. HARRIS

ATP Binding Cassette (ABC) Pumps ABC pumps typically do not pump cations and, in fact, may have rather wide specificities (Hyde et al, 1990). The best known example in humans is the multidrug resistance (MDR) protein (also known as the P-glycoprotein) which . pumps large organic molecules out of cells. By pumping compounds such as doxyrubicin, vincristine, etc., out of tumor cells, it can be responsible for the low sensitivity of these cells to cytotoxic agents. A similar protein is responsible for chloroquine resistance of the malarial parasite. Other examples include the chloride channel protein defective in cystic fibrosis (CFTR)—^where the compound pumped is still unknown—and the peptide transporter (TAP1-TAP2) involved in antigen presentation in lymphocytes. ABC pumps comprise a dimer of ATP binding domains (outside the membrane) bound to a dimer of transmembrane domains, each comprising six transmembrane helices. Interestingly, the polypeptide organization of these proteins may vary; TAP1-TAP2 contains two polypeptides, each containing one transmembrane and one ATP binding domain, while in the multidrug resistance protein the two are fused in a single polypeptide containing all four domains (1,280 amino acids). F-Type ATPases The F-type ATPases are ATP-driven H"*" pumps. They show similarities in structure to the V-type ATPases, in that they are separable into a soluble headpiece (5-6 different polypeptides) and a transmembrane H^ channel (3-5 polypeptides) (Senior, 1988). However, there are characteristic structural differences, the most obvious occurring in the proton channel; F-type pumps employ a peptide about 80 amino acids long (in 10-12 copies) as a H^ carrier, while in V-type pumps the equivalent peptide is twice as long. More important, however, is the difference in function. In animals, the only F-type pump, which is found in mitochondria, does not act as an ATP-driven H"*" pump but in reverse, as the H"^-driven ATP synthase responsible for all oxidative ATP synthesis. This is dealt with further below. ATP and the Energetics of Ion Pumping In ATP-driven pumps, ATP, which binds to the cytoplasmic domain, cannot interact directly with the ion being pumped, which passes through the membrane sector. Energetic coupling, therefore, is indirect. Ion movement occurs via an alternating access model; the ion binding site is exposed to one side of the membrane in one conformation (EO and to the other side in the other (E2) (Figure 8a). These ideas can be combined with those used to derive the above model for actomyosin function. Critical features are: (a) energy release from ATP is associ-

K.Ei.ATP

J
i2

,., ,

Na.Ei ATP

NaE^-^P

NaJ2± E2.ATP*

/

\EI,ATP

,

„x

/\E2-P

Na in

•

\

E2

Figure 8. Mechanism of ATP use in the NaVK"^ pump, (a) Alternating access model for ATP driven transport of ions. The inward facing (Na"^ binding) form of the enzyme is designated conventionally as E^, and the outv^ard facing (K"^ binding) form as E2. Note that each chemical step (ATP binding, protein phosphorylation) produces a kinetically unstable species (*) which relaxes in the next stage of the cycle (c.f. Figure 6). T represents (non-covalently) bound ATP, P represents covalently bound phosphate, (b) Energetics of .the alternating access model. Energy is transferred from ATP (solid line) to uphill K"^ movement (decrease in K"^ affinity) (dashed line), and from an acyl phosphate to uphill Na"^ movement (decrease in Na"^ affinity) by enzyme conformational changes. Note that energy is used to change ion binding affinity, rather than to change protein conformation per se. For simplicity, the number of ions (2K% 3Na"^) moved in each conformational change is not shown.

19

20

DAVID A. HARRIS

ated with changes in ATP binding to the protein, not with chemical changes; (b) ATP binding provides energy for a decrease in binding affinity of a second hgand (actin, above); and (c) this energy is transmitted via a protein conformational change. Applying these principles to operation of the NaVK"^ pump gives the model in Figure 8b. The analogy is strongest on the top row. K"^ binds very tightly to E2, so it can be picked up at the low concentration outside the cell (low [K]out)- To release it at high [K]in, binding must be weakened, and this is accomplished by binding ATP tightly. Since ATP binds tightly only to Ei, the enzyme changes into the Ei conformation, consequently changing the orientation of its binding site. Coupling between energy release and ion movement thus occurs via conformational changes in the protein. In the second part of the cycle, a similar process occurs. Na"^ binds tightly to El while the aspartyl phosphate residue is destabilized (Ej-P); as the conformation of the protein changes, the aspartyl phosphate is stabilized (E2-P), and the binding energy released loosens Na"^ binding (Na"^ released at high [Na]out)As noted above, the chemical processes (EiATP ~> E p P ; E2P -> E2) are not accompanied by large energy changes and serve simply as triggers for the next stage in the cycle. Similar models can be derived for all types of ATP driven pumps discussed above. ATP in Biosyntheses

In tissues carrying out large amounts of biosynthesis (e.g., liver, exocrine tissues, rapidly dividing cells), a considerable fraction of cell ATP can be devoted to driving NH2 I

NH2 ATP

I

R - C - H ""v

I

. R-C-H

"^^

C= 0 0 AMINO ACID

I

PPi

'

V

vR-C-H

"^^^

I

C= 0 AMP O-p-O-Qdenosine 0* AMINO ACYL ADENYLATE

CH3

CH3

(c H2)n CoA ppj

C=0

c=0 O-tRNA

AMINO ACYL tRNA

CH3

(CH2)nATP C=0

NH2 tRNA

(c Hzln AMP

^=0

i

' Q

'

0"

O-P-O-Qdenosine

0-S-CoA

0" FATTY ACID

FATTY ACYL ADENYLATE

FATTY ACYL Coenzyme A

Figure 9. Role of ATP in generating coenzyme-bound intermediates for macromolecule biosynthesis.

Cellular ATP

21

biosynthetic processes. In the islets of Langerhans, for example, about 50% of ATP is used to drive biosyntheses. The role of ATP in biosynthesis is to activate a molecule for further reaction. Two classes of reaction can be distinguished, ATP as a Dehydrating Agent

As noted above, macromolecules are assembled by reacting component monomers with the elimination of water (condensation). ATP is an acid anhydride; it can therefore, in principle, drive condensation reactions. It must, however, specifically remove that water formed in the reaction, in the presence of a large excess of cell water. This requires the direct reaction of ATP with one of the reacting species. Typically, ATP will activate organic acids (amino acids, fatty acids), forming an adenylated intermediate (mixed acid anhydride). This reaction would be barely favorable if Pj were released; however, PPj is released, and this is rapidly hydrolyzed by cellular pyrophosphatase, ensuring complete reaction. O

II (i) R-COO" + ATP -> R-CO-P-adenosine + PPj (ii) PPi+H20 -> 2?^ I O"

The adenylated intermediate is too unstable to release into free solution, and so the activated acid is transferred to a carrier molecule, tRNA in the case of amino acids and coenzyme A for fatty acids (Figure 9). However, the carboxylate group is still in an anhydrous state, and these species will react with the acceptor molecule (R-NH2 for amino acids, glycerol(-OH) for fatty acids) without further energy input. In the synthesis of polysaccharides, the dehydrating agent is a different nucleoside triphosphate, UTP. This replaces, in an analogous reaction, a sugar hydroxyl (alcohol) group forming the UDP-sugar compound, which can transfer the activated sugar on to the -OH of a growing polysaccharide chain. Activation of Leaving Groups

A number of biochemical reactions involve the displacement of a hydroxyl group by another group (e.g., an amino group). However, the -OH group is chemically a very poor leaving group, i.e., it shows a strong tendency to remain attached to its neighboring carbon atom. Phosphorylation of the hydroxyl group, using ATP, creates a much better leaving group -P04~, and thus makes this group susceptible to attack. Examples occur in the synthesis of glutamine from glutamate, and of CTP from UTP (Figure 10).

22

DAVID A. HARRIS

(CHzlzATP H-C-NH?

(CH2)2

0\t^,

AOP H-C-NHo

(CH2)2 Pj H-C-NH2

I

I

I

COO"

coo"

COO"

y -OUITAMTL FBOSPHATE

nKIDIME TBIFIiOSPiUTE

Figure 10.

0-P-O' 1 \.s-

NH2

I R

I R

6-I>BaSI>B0 VSIDZNE TRIPHOSPHATE

Role of ATP in activating hydroxyl groups to nucleophilic attack.

ATP as Phosphate Donor As ATP has a high phosphate transfer potential, it can transfer its terminal phosphate to an alcohol (OH) group, forming a phosphate ester, in a downhill (thermodynamically favorable) reaction. In contrast to the reactions in the previous section, the resultant compound is not especially reactive; the primary reason for such phosphate transfers is to confer negative charge onto the recipient molecule. Phosphorylation of Sugars The archetype of this class of reaction is the phosphorylation of glucose by hexokinase. This is a downhill reaction—the equilibrium is well over towards glucose-6-phosphate—^which ensures that, within (non-liver) cells, free glucose levels are kept low. The glucose-6-phosphate formed, which is negatively charged, does not readily cross the cell membrane and is thus retained within the cell.* This class of reaction rarely makes significant demands on the cellular ATP content. However, it can do so in the pathological condition of fructose intolerance. The normal pathway of fructose metabolism, which occurs in the liver, requires In liver cells, glucose entry is so fast relative to phosphorylation that free glucose does build up; this allows the liver cell to "sense" blood glucose levels, and is associated with its role in maintaining blood glucose by taking up or releasing glucose.

Cellular ATP

23

two novel enzymes, fructokinase (producing fructose 1 phosphate) and fructose 1 phosphate aldolase (which cleaves fructose 1 phosphate into 3 carbon sugars). In hereditary fructose intolerance, the aldolase is absent, and continued fructose intake will cause a build up in the liver of fructose 1 phosphate with accompanying depletion of cell phosphate and ATP. Phosphorylation of Proteins Protein kinases will transfer phosphate from ATP onto specific hydroxyl residues (serine, threonine, or tyrosine) within proteins. This is commonly associated with enzyme activation (e.g., glycogen phosphorylase, plasma membrane L-type Ca^"^ channel) or inactivation (glycogen synthase, pyruvate dehydrogenase). Thus these phosphorylations play a regulatory role. The high phosphate transfer potential of ATP again ensures that reaction can be virtually complete, i.e., nearly all enzyme molecules are in one form or the other. The sensitivity of the system is, therefore, high compared to allosteric regulation (which is based on reversible, non-covalent, binding equilibria). The role of phosphate in regulation is based largely on its charge. In glycogen phosphorylase, phosphorylation of serine 14 allows this N-terminal region to bind electrostatically to a cationic hole in the protein, triggering a conformational change at the (distant) active site (Browner and Fletterick, 1992). Phosphorylation of membrane proteins may be less precise in its effects; simply changing their surface negative charge may allow aggregation of membrane proteins (e.g., insulin receptors) or cause disaggregation (e.g., in the chloroplast membrane in green plants). ATP as Charge Neutralizer ATP is commonly found in intracellular storage granules. For example, the chromaffin granules of the adrenal medulla contain ATP levels of about 100 mM, 15 times higher than cytoplasmic levels. This ATP is metabolically inert, and seems to exist in a complex with epinephrine such that the positive charge on the catecholamine is neutralized by the negative charge on ATP"^". (Epinephrine, with one positive charge, can reach concentrations of up to 400 mM inside the granules.) Serotonin (platelets), insulin (pancreatic (3 cells), and acetylcholine (synaptic) storage granules also contain ATP. This ATP is released into the blood on exocytosis, along with the hormone. Here it is rapidly hydrolyzed to adenosine. This, too, has some endocrine action in causing the relaxation of vascular smooth muscle, increasing local blood flow, and thus aiding hormone delivery to the target tissues.

24

DAVID A. HARRIS

ATP and Messenger Molecules Generation ofcAMP In a reaction catalyzed by adenylyl cyclase, ATP is converted to 3'5'-cyclic AMP (cAMP). This compound is a ubiquitous signal molecule, generally indicating a stress situation: in both Escherichia coli and man, for example, cAMP is produced in response to nutrient limitation (starvation). In mammals, adenylyl cyclase is a membrane-bound enzyme which is activated in response to a variety of hormone receptors in the cell membrane, notably those for epinephrine and glucagon. cAMP is a second messenger for these hormones, and activates protein kinase A within cells. The reaction producing cAMP is shown in Figure 11. Due to strain in the ring formed, cAMP is (like ATP) thermodynamically unstable. The formation of cAMP from ATP is thus favored only by the hydrolysis of PPj by cellular pyrophosphatase (roughly equivalent to the hydrolysis of one high energy bond). The hydrolysis of cAMP to AMP (roughly equivalent to hydrolysis of the second high energy bond) is catalyzed by phosphodiesterase. The presence in a cell of two enzymes capable of the net uncoupled hydrolysis of ATP is a potential hazard. It can be supported, however, because the maximal capacity of adenylyl cyclase is low (and its activity is generally suppressed even further) such that cAMP is maintained at a basal steady state level of around ICT^ M, five orders of magnitude lower than ATP. When adenylyl cyclase is activated, a new steady state is established with c AMP at about 1 Qr^ M (the phosphodiesterase simply responding passively to increased cAMP levels). Thus ATP provides a nearly infinite pool of potential cAMP: cAMP levels can be changed 10-100-fold with a loss of less than 0.1% of total cell ATP. The maintenance of a large pool, and a low steady state value, of messenger molecules is an essential feature of signaling in biological systems, because it allows a rapid, many-fold change in messenger concentration. The same features, with a rather different organization, occur in Ca^"^ second messenger systems; intracellular [Ca^"*^] is normally around 10"^ M, but can be rapidly increased, in

Qdeninei

0

0

0

laden ine|

K'

^^

\

r "o

ppi

^ — 1 I OH 3 ^ 0 - P = r C / 0 molecule are Figure 11, Formation of cyclic AMP. The 3' and 5' positions of the indicated. PPj produced is rapidly hydrolyzed /n wVo(see above).

Cellular ATP

25

response to a hormone signal, to 10"^ M by transiently opening channels to a large pool (10~-^ M) of intra- or extracellular Ca^^. ATP Dependent IC Channels Besides ion channels controlled by ATP dependent phosphorylations (e.g., plasma membrane L-type Ca^^ channels), cells in a number of tissues contain a plasma membrane K"*" channel which is inhibited by the non-covalent binding of ATP. This channel normally mediates K^ efflux (leading to hyperpolarization), and can be demonstrated as an ATP-sensitive channel in patch clamp experiments. However, since I50 for blocking the channel is only 10-50 |LIM, some 100-fold lower than cellular ATP levels even during ATP depletion, the channel would be expected always to be closed in normal cells. Its role in normal cell function is thus unclear. One interesting suggestion is that this ATP-dependent K"^ channel might trigger insulin release in pancreatic P cells (Ashcroft and Rorsman, 1989). These cells respond to a rise in blood glucose in the range 3—10 mM by increasing their metabolism of glucose, (and thus ATP generation). In this model, then, as blood glucose rises, ATP levels should rise, promoting closure of this K"^ channel. Th6 cell thus depolarizes due to a net cation (Na"*") influx, and insulin release is triggered. This model cannot be regarded as proven, due to discrepancies between the measured changes in ATP concentration in p cells and those predicted for this model to operate. However, it remains an attractive model for the coupling of blood glucose concentration to insulin release. Structural Role of ATP The adenine moiety of ATP is used as part of cellular structures (e.g., coenzymes, RNA, DNA). The amount of ATP consumed in this way will depend upon the biosynthetic activity of the tissue. However, only the roles of ATP as an energy/phosphate source are discussed further in this article.

REACTIONS INVOLVING EXCHANGE OF HIGH ENERGY PHOSPHATES Two important reactions of ATP do not result in a net loss of phosphoanhydride bonds. These are the creatine kinase and adenylate kinase reactions. Creatine kinase: Cr + ATP <^ PCr + ADP Adenylate kinase: AMP + ATP <-> ADP + ADP In contrast to the reactions described above, these enzymes are primarily involved in maintenance of ATP homeostasis rather than in providing energy for some

26

DAVID A. HARRIS lfH2

^H2 C=NH

C=NH

N-CH3

NH

I

CH2

CH2

COO"

CH2 COO*

creatine

Figure

12.

p-guanidino propionate

Structure o f c r e a t i n e a n d its a n a l o g u e , p - g u a n i d i n o p r o p i o n a t e . T h e

p o s i t i o n o f p h o s p h o r y l a t i o n , t o f o r m PCr is s h o w n (*).

particular cell function. Both enzymes are highly active and freely reversible in vivo. Creatine Kinase

Phosphocreatine represents a metabolic dead end; its only reaction is with ADP, and, in the creatine kinase reaction, it acts as an ATP buffer. This reaction occurs in all muscle and neural tissue, with each tissue having its own isoenzyme of creatine kinase. Other tissues (such as liver and kidney) lack creatine kinase and PCr. The structure of creatine is shown in Figure 12. PCr concentration in heart cells is typically about 25 mM, which is 3—4 times the concentration of ATP. Thus, as ATP is used, if ADP tends to rise, it can be rephosphorylated using PCr as phosphate donor. This is a rapid mechanism of

mitochondrion

sarcoplasm

PCr

ADP.

C^: Cr

ATP

Figure 13, Transfer of high-energy phosphates by the phosphocreatine shuttle. CK^^ and CKs represent the mitochondrial and sarcoplasmic isoenzymes of creatine kinase, respectively. Free Pj is omitted for clarity.

Cellular ATP

27

regenerating ATP (buffering ATP concentration) in short-term energy deficiency (see Figure 2b). However, PCr stores cannot last more than 15-30 seconds in the absence of other energy input; PCr seems more likely to function, for example, in smoothing out ATP variation between heart beats than as a significant energy store in the tissue. A second possible role of PCr is in shuttling ADP to and from mitochondria, especially in heart (Jacobus, 1985). This proposal is based on the idea that ADP, present at very low concentration (10-30 juM), may not be able to diffuse from its site of formation on the actomyosin filament to the mitochondria fast enough to allow rapid oxidative phosphorylation. With the PCr shuttle, ATP made inside the mitochondrion emerges into the intramembrane space where it transfers its y phosphate directly to creatine. The resulting PCr moves across the cytoplasm to phosphorylate ADP near actomyosin, for use in muscle contraction, and the resulting creatine diffuses back to the mitochondrion (Figure 13). ADP is formed at the mitochondrion in this process, but can be rephosphorylated rapidly since it does not have to diffuse within the fiber, but simply to reenter the mitochondrion. The only diffusing species are Cr and PCr, present at 100-1,000 x [ADP] concentration. While such a mechanism may well operate, experiments in which the heart's creatine is depleted by treatment with the analogue P-guanidino propionate (Figure 12) (Zweier et al., 1991) with little or no effect on cell function, suggest that it is unlikely to contribute significantly to energy transfer in the heart cell. Adenylate Kinase This enzyme is present at high levels in all tissues. It could clearly act as an ATP buffer if ADP levels were to build up, but the levels of ADP in vivo (about 1% of ATP levels) make this unlikely. It is more likely to be useful in the rephosphorylation of AMP, generated in reactions like amino acid activation (above). AMP itself cannot be used in oxidative phosphorylation, but adenylate kinase will phosphorylate it to ADP, which can. Perhaps a more important role of this enzyme is in signaling the energy status of the cell to regulatory enzymes. In cells, the concentrations of ATP, ADP, and AMP are widely different, with [ATP] at about 5 mM, and free [ADP] and [AMP] at 0.01-0.05 mM (see Table 2). Hydrolysis of a small amount of ATP (say, 0.1 mM) by myosin, for example, will decrease ATP levels by a very small fraction (2%). However, this will tend to increase ADP concentration by the same amount, which is a much larger fraction (300-1,000%) of total ADP. Since adenylate kinase maintains ATP, ADP, and AMP at equilibrium, this change will be transmitted to AMP, whose concentration will also increase by a large fraction. Thus, a small percentage fall in ATP concentration is accompanied by a large percentage rise in [AMP]. AMP, therefore, is a useful regulatory molecule in the cell; changes in

28

DAVID A. HARRIS

[AMP] are amplified versions of changes in [ATP], and enzymes responsive to [AMP] as a regulator are very sensitive to changes in cellular [ATP]. Such enzymes are discussed further below.

SYNTHESIS OF ATP ATP synthesis in animals occurs in one of two modes, one in which oxygen is used (oxidative phosphorylation), which occurs in mitochondria, and one in which oxygen is not involved (substrate level phosphorylation), which occurs, largely, in the cytoplasm. In elementary textbooks, substrate level phosphorylation is often referred to as being less efficient, since it yields only 2 mols ATP per mol glucose used (as compared to 36 mols/mol glucose in oxidative phosphorylation); in mammals, however, this term is misleading since lactate, produced in substrate level phosphorylation in one tissue, can be fiilly oxidized in another with no loss of ATP to the organism. On the contrary, the ability of some tissues to carry out substrate level phosphorylation allows them to be better adapted to their fiinction, and thus increases the efficiency of the organism as a whole. Substrate Level Phosphorylation Substrate level phosphorylation in animals takes place during glycolysis. The pathway for glycolysis is shown in Figure 14. ATP synthesis typically involves two steps: (a) the generation of a high energy phosphate bond; and (b) the transfer of this phosphate to ADP. It occurs twice in the breakdown of each molecule of triose phosphate. The regulation of this pathway is discussed later. In glycolysis, energy is yielded via an internal oxidation-reduction reaction; the aldehyde group of a sugar is oxidized to an acid, and an alcohol group reduced to a methyl group. This is possible only with a carbohydrate energy source. Thus, the fuels for substrate level phosphorylation in vivo must be (blood) glucose or (stored) glycogen; fatty acids and amino acids cannot be used to generate ATP without the involvement of oxygen. Tissues that generate ATP mainly by this route are not, typically, anaerobic. Rather, they lack (sufficient) functional mitochondria to meet their energy demand. They include: 1. Erythrocytes: their (low) energy demand is met by uptake of glucose from the blood, and diffusion of lactate into it. They contain no mitochondria. 2. Lens of the eye: energy requirements are met as for erythrocytes. Here, the presence of mitochondria would impair function, as they are small particles (light scattering) and colored (due to heme). 3. Kidney medulla: in contrast to the kidney cortex, the-medulla, which contains the loops of Henle, is poorly vascularized. This may be due to the osmotic problems of capillaries in such a hypertonic space. Thus oxygen

EXTRACELLULAR FLUID adrenaline

glycogen glycogen

^

GiP

<

phosphorylase

ATP + AMP <

>2ADP

- > ADP + ?\

1,3-diphosphoglycerate

t

(b)

ATP
2-phosphoglycerate

(a)

phosphoe«o/pyruvate ATP

A

(b)

pyruvate . 4 — ^ lactatejn

<—•

lactatCot

figure 14. Glycolysis in skeletal muscle. Stages at which a high energy bond is generated are indicated by V / X / ^ , and designated (a) (see text). Phosphate transfer to ATP occurs in the following step (b). Only key regulatory enzymes are named. Molecules signaling an actual or potential increased ATP utilization are shown in bold. The role of kinases (protein kinase A, phosphorylase kinase) in the actions of cAMP and Ca^"^ on phosphorylase is omitted for clarity. 29

DAVID A. HARRIS

30

supply is low, despite a significant demand for ATP (for pumping ions), and energy derives from glycolysis. Fast twitch (anaerobic) muscle (Type lib fibers): these are responsible for short intense bursts of activity (e.g., sprinting), and are poorly vascularized, but contain large glycogen stores (increased by training). Some mitochondria are present, providing ATP for the cell at rest. On activity, internal glycogen stores are broken down to lactate; entry of glucose and oxygen from the blood would be too slow to fiiel the bursts of high activity, especially since contraction may squeeze blood out of the muscle. Note that internal glycogen stores are limited as increasing them increases muscle mass. Typically, they can maintain ATP levels for 20-30 seconds. Tumor cells: many tumors (e.g., Morris hepatoma, Ehrlich ascites tumor) contain apparently normal mitochondria, but convert a large fraction of glucose taken up to lactate. This may be partly due to limitations in oxygen diffusion to the center of solid tumors. However, it may also reflect the demand by stimulated biosynthetic pathways for glycolytic intermediates in these cells. Lymphocytes and macrophages, similarly, obtain significant amounts of ATP by converting glucose to lactate, despite the presence of functional mitochondria (Ardawi and Newsholme, 1985). Again, this is

fatty acids pyruvate fatty acids

pyruvate L j r dehydrogenase I

^'^^^ Ca^*

acetyl CoA

FADH2 MITOCHONDRIAL MATRIX

FADHj

Figure 15. Oxidative ATP generation in heart muscle. Only key regulatory enzymes are named. Regulatory molecules are shown in bold.

Cellular ATP

31

linked to a requirement for rapid biosynthesis (in response to immune stimulation) in these cells. Oxidative Phosphorylation Oxidative phosphorylation occurs inside mitochondria. It can be divided notionally into two stages, the generation of reduced cofactors (NADH and reduced flavin), and the oxidation of these cofactors by oxygen, coupled to ATP formation. Reduced cofactors are generated by the soluble enzymes of the mitochondrial matrix, notably those of the Krebs cycle and those catalyzing (3 oxidation of fatty acids. These processes are summarized in Figure 15. Typically, oxidation of a-oxo acids or hydroxy compounds yields NADH, which is a soluble cofactor in the mitochondrial matrix; oxidation of-CH2-CH2- groups (e.g., by succinate dehydrogenase, fatty acyl dehydrogenase) yields FADH2 which is bound within mitochondrial inner membrane complexes. Electrons from these cofactors are transferred between the electron transfer complexes of the mitochondrial membrane, ultimately reducing oxygen to water. In this process, protons (H"^) are pumped out of the mitochondria, building up an electrochemical H^ gradient as an intermediate energy store. This is used by another transmembrane complex, the FiFoATPase, for ATP synthesis (see below). Fuels for Oxidative Phosphorylation Compared to substrate level phosphorylation—^where the only available fiiel is glucose (free or in glycogen)—oxidative phosphorylation can be fueled by sugars, fatty acids, or amino acids. It is thus a particularly versatile (and efficient) mode of ATP production. Different oxidative tissues prefer different fuels for ATP production, and the preferred fuel may vary with nutritional status of the organism, the whole producing an integrated, adaptable system for energy production. Examples are: 1. Brain. The brain is one of the few tissues whose ATP production (in the fed state) is fueled largely by glucose oxidation. Fats and fatty acids are inaccessible to the brain because of the blood-brain barrier. In starvation, ketone bodies (partially oxidized fatty acids, produced by the liver) can fuel the brain. 2. Heart. Heart muscle, in contrast, prefers to oxidize triglyceride (from circulating lipoproteins) and lactate in the fed state. Circulating fatty acids and ketone bodies are oxidized in starvation. 3. Skeletal muscle (Types I, Ila). Like heart muscle, skeletal muscle shows a preference for lipid substrates, in particular triglycerides (fed state) and fatty acids/ketone bodies in prolonged exercise or starvation. Skeletal muscle

32

DAVID A. HARRIS does not oxidize lactate due to the low affinity of the muscle isoenzyme of lactate dehydrogenase for this substrate, relative to that of the heart enzyme. 4. Liver. Since the liver is an important center of nitrogen metabolism, its major fuel in the fed state is a-oxo acids, derived from the deamination of dietary amino acids. In starvation, it will oxidize fatty acids released from adipose tissue. 5. Gut. Perhaps surprisingly, small intestine obtains about 60% of its ATP in the fed state from the oxidation of the single amino acid, glutamine (Watford et al., 1979). (This is twice transaminated/deaminated to yield a-oxoglutarate, which is oxidized in the tricarboxylic acid cycle). Oxidation of glucose and ketone bodies make up the remaining 40%.

Mechanism of the Mitochondrial ATP Synthase The mitochondrial ATP synthase utilizes energy stored in a transmembrane H"^ gradient, transducing this into the phosphoanhydride bond of ATP. Current views hold that 3¥t are transferred across the mitochondrial inner membrane for each ATP made by the synthase. Coupling between H"^ movement and ATP synthesis is indirect, i.e., the protons moved do not interact with the ATP made, the two processes communicating solely via changes in protein conformation. The resulting mechanism is essentially the reverse of that described above for ATP-driven pumps, as indicated in Figure 16a. In essence, the synthase has a very high affinity for ATP, such that ADP + P, spontaneously form ATP on the enzyme surface; energy is required to displace ATP from the enzyme. This is supplied by binding H"^ from a solution of high electrochemical potential, inducing a conformational change that displaces ATP. The ATP synthase has a bipartite structure, with the H^ channel lying on one set of polypeptides (FQ, transmembrane fragment) and the ADP binding site on another (Fj, extrinsic fragment) (see Figure 7). Its mechanism is also complicated by the presence of not one but three ADP binding sites which operate alternately. Thus, binding of ADP + Pj, ATP formation and ATP release involve three different conformations of the catalytic site of Fj (designated Loose, Tight, and Open). In the holo-enzyme, each of the three catalytic sites pass through each of these conformations, but at 120° they are out of phase (see Figure 16b) (Boyer, 1987). The reason for this complicated arrangement may be to ensure that ATP does not simply rebind to the enzyme after its expulsion by H"^; any potential ATP binding sites have been previously filled by ADP + Pj (see Figure 16a). The asymmetric structure of Fi at any instant is thought to be due to the preferential interaction of one particular conformation of the catalytic subunits with the (stalk) polypeptides which link the membrane (FQ) and catalytic (Fi) fragments. The ATP synthase generates ATP within the mitochondrion, while most of it is needed in the cytoplasm. ATP is exported from the mitochondrion via the adenine

3H' E.ATP

30 k> ^mol

L A D ^ E.ATP ^^^

'. 3H- in

Figure 16. Mechanism of the mitochondrial ATP synthase, (a) Energetics. ATP is formed without energy input on the enzyme surface (solid line), due to the high affinity of the enzyme for ATP. For synthesis of free ATP, H"" ions moving downhill (dotted line) change the enzyme conformation, decreasing ATP affinity, (b) Alternating site mechanism. The three active sites can each exist in three different conformations: t = tight ATP binding; o = open, unable to bind nucleotide; and I = loose, in which ADP and Pj can exchange rapidly (dotted arrows) with the solution. The central mass represents the polypeptides which link Fi with the proton channel (not shown); this associates with the o conformation of active site only. Protons passing through the channel displace these polypeptides from one active site to the next (counterclockwise in this diagram), and the active site conformations thus change in sequence.

33

34

DAVID A. HARRIS

nucleotide translocase, which exchanges internal ATP for cytoplasmic ADP. This uniquely utilizes free ATP rather than the MgATP complex; if MgATP were exported, the mitochondria would lose internal Mg^"*" and charge balance would be upset. Hence, ATP is exported as ATP"*^ and ADP imported as ADP-^~. This process is energetically favored because the interior of the mitochondrion is negative relative to the outside, as a result of pumping protons (H"^) outwards. In principle, the two transporters, the ATP <-> ADP translocase and the ?J¥t symport (which imports Pj into mitochondria) can be considered as a coordinated system in which ATP"*" is exported, while ADP-^ + P?~ (substrates for ATP synthesis) are taken up at the cost of moving 1H"*" down its electrochemical gradient. This requirement for energy for ATP export explains how the cytoplasmic ATP/ADP ratio can exceed the mitochondrial ATP/ADP ratio by a factor of about 100 (see Table 2); ATP is actively expelled and ADP pumped inwards. P/O Ratios The P/O ratio is defined as: P/O = mols ATP made/atom O consumed. This will depend upon the substrate oxidized: NADH, which is a strong reducing agent (E^'=-0.3V), will yield more ATP than succinate (E^'= OV). In mechanistic terms, this is reflected in the ability of NADH/UQ oxidoreductase complex to pump H"*", while succinate/UQ oxidoreductase cannot. This ratio is clearly an important parameter in quantitating cellular energy metabolism. This being so, it is surprising that its value is not known with certainty. Consensus values are P/O = 2.5 for NADH oxidation and P/O = 1.5 for succinate oxidation (Ferguson, 1986). Since, in mitochondria, 3H"^ are used by the ATP synthase, and IH"*" by the translocase in synthesizing 1 mol (cytoplasmic) ATP, this suggests that 10 H"^ are pumped out per mol NADH oxidized. Problems in determining this ratio have been both practical and conceptual. Practical problems center on the metabolic cost of transport; different membrane preparations will show different P/O ratios, depending on whether the synthase is internal (right side out), or external (inside out) where the need for ADP and ATP transport is abolished. The conceptual difficulties arise from a historical tendency to expect this value to be a whole number, in various mechanistic models for ATP synthesis. However, since we now know that several H"^ are required to make 1 ATP, non-integral values for the P/O ratio no longer raise any conceptual problems. Various chemicals known as uncouplers (e.g., 2,4 dinitrophenol, picric acid) can decrease the P/O ratio by increasing the permeability of mitochondrial membranes to H^. In this case, protons leak across the membrane, bypassing the ATP synthase and producing heat. These chemicals, used in explosives manufacture, were responsible for weight loss and tissue wasting among explosives workers early in this

Cellular ATP

35

century. A similar syndrome is observed in a rare genetic disease of mitochondria, Luft's disease, where the membrane again is abnormally leaky to H"^ (probably due to malfunctioning Ca^"^ transport).

CONTROL OF ATP LEVELS ATP Levels Are Closely Maintained In Vivo

Since ATP participates in a wide variety of metabolic processes, it is hardly surprising that its levels are tightly controlled. Nonetheless, the degree of control observed is remarkable; variations in work rate of up to 10-fold in heart, and even more in skeletal muscle, produce no detectable (<5%) change in ATP concentration. In other tissues too, cytoplasmic ATP levels hardly change over the physiological range of energy utilization. In very severe exercise, ATP levels in skeletal muscle may fall by 50-60%, but this is accompanied by a loss of total adenine nucleotide from the tissue. The level of cellular [ATP] depends on the balance between synthesis and utilization. However, utilization depends on the needs of the cell, and ultimately the organism, the energy-requiring processes (movement, biosynthesis, etc.). Thus ATP levels must be controlled, under physiological conditions, by varying the rate of ATP synthesis. Since cellular ATP turns over about once every second, such control must be both precise and rapid. In addition, processes that require ATP are buffered against changes in its concentration. This is because enzymes that use ATP (e.g., actomyosin, hexokinase, NaVK"^ ATPase, etc.) have K^ values around 0.1-0.2 mM, which means that they are saturated at physiological [ATP] of 4-8 mM; even a 50% change in ATP concentration would hardly affect their ATP utilization rate. Muscle fatigue, for example, is accompanied by a fall in ATP levels to 2-3 mM, but the loss of muscle function in fatigue is due to a rise in intracellular [H"^] (pH falls from 7 to 6.5) rather than ATP depletion (see Figure 2). ATP levels, therefore, seem to be regulated even more precisely than one might expect. It is possible that K^ values measured in vitro may be misleading, and that apparent K^ values in vivo may be closer to physiological [ATP] (if, for example, cellular ADP competes strongly with ATP). Even so, it seems that metabolic processes are doubly insulated against a tendency of [ATP] to fall: ATP synthesis rates will be increased by regulatory mechanisms, and, in addition, enzymes are relatively insensitive to decreases in [ATP] around physiological concentrations. Is ATP a Regulator?

It is tempting to imagine ATP itself as a metabolic regulator. It is conceivable, for example, that ATP might inhibit key enzymes in glycolysis and oxidative

36

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phosphorylation; use of ATP would cause its concentration to fall, relieving this inhibition and speeding up ATP synthesis. Indeed, the regulatory enzyme phosphofructokinase (PFK) is inhibited by ATP levels of around 1-2 mM in vitro. The drawback to this simple model is the very constancy of cellular [ATP]. It is difficult to see how a drop in [ATP] of around 5% could allosterically increase the activity of an enzyme by up to 100-fold, and yet this change in glycolytic flux can occur in skeletal muscle during a transition from rest to exercise. In other words, no known system would be sufficiently sensitive for changes in ATP levels directly to cause the observed changes in flux. ATP itself is not a physiological regulator molecule. Mammals have adopted three mechanisms to circumvent this problem. 1. ADP and, in particular, AMP are used to signal changes in ATP concentration. Small percentage changes in [ATP] lead to large percentage changes in both [ADP] and [AMP] levels, due to the large difference in absolute concentration of these nucleotides and the operation of adenylate kinase (above). Thus small changes in [ATP] are amplified into large changes in concentration of these effectors. 2. The sensitivity of an ATP yielding pathway to such regulators is further amplified by the operation of substrate cycles. In a substrate cycle, two enzymes catalyze opposing reactions simultaneously so the net flux through the cycle is small; a small increase in the activity of one enzyme and a corresponding decrease in the activity of the other will, however, produce a large change in net flux (Figure 17). Substrate cycles consume ATP (1 mol per turn of the cycle), as both forward and reverse reactions must be downhill. The fact that such cycles operate in ATP generating pathways attest to the physiological importance of tightly controlling ATP generation; it is even worth paying for with existing ATP. 3. Variations in ATP demand may be signaled by indicators other than adenine nucleotides. In heart and skeletal muscle, for example, mean cytoplasmic [Ca^"*"] rises with increasing work rate due to its role in the contractile process. This can be used as a signal of ATP demand, speeding up ATP production. Extracellular regulators, such as epinephrine, signal an imminent rise in ATP demand rather than an actual one. Such regulators, via cAMP as second messenger, can also stimulate ATP generating pathways, in this case prior to an actual increase in utilization. Control of Anaerobic ATP Production Control in skeletal muscle type lib (white muscle) is taken as the archetype of such regulation, which occurs in the glycolytic pathway (oxidative phosphorylation cannot operate under anaerobiosis). Such muscle (e.g., human quadriceps) is

Cellular ATP

37 ATP , " CPF 100---^110

fructose

fructose

6-phosphQte

1,6-bisphosphQte

1--^20
>

J'j^--»90 (fBPas§ Figure 17. A substrate cycle in the amplification of a metabolic response. The enzymes depicted are phosphofructokinase (PFK) and fructose 1,6, bisphosphatase (FBPase) (see Figure 14). The figures shown indicate rates (arbitrary units). Initially, the forward rate is 100 and the reverse rate 99; the flux emerging through the pathway (J) is 1 unit. A 10% rise in PFK activity (100 -> 110), and a 10% fall in FBPase (100 -^ 90) leads to a 2,000% change in onward flux.

capable of very intense but short lived bursts of activity, during which glycolytic rates can increase up to 200 x over the rate observed at rest. Control by Nucleotide Levels Crossover analysis has demonstrated that phosphofructokinase (PFK) is the key regulatory enzyme in glycolysis. This enzyme is activated by AMP in vivo (Figure 14). It also participates in a substrate cycle with fructose 1,6 bisphosphatase (FBPase), which hydrolyzes fructose 1,6 bisphosphate to fructose 6 phosphate (F6P), opposing the action of PFK (Figure 17). In muscle, where no gluconeogenesis occurs, the only role of FBPase is to participate in this substrate cycle. Thus, as muscle is stimulated to contract, ATP is hydrolyzed to ADP. ATP levels are buffered by PCr and barely fall; by virtue of adenylate kinase, AMP levels rise significantly. PFK is thus stimulated and glycolysis speeds up. The sensitivity of the system is maximized by using AMP rather than ATP as a regulator (see above). In addition, PFK shows allosteric cooperativity (i.e., a sigmoidal response to AMP) such that a doubling of AMP concentration, in the physiological range, can give a 5—10-fold increase in active enzyme. Furthermore, due to the reverse action of FBPase in the substrate cycle, doubling the activity of PFK can give a 50-fold change in net flux through the pathway. Each of these mechanisms, therefore, amplifies the sensitivity of the pathway to the tiny fall in ATP levels, the rate of ATP synthesis rises sharply, and [ATP] is maintained almost constant. Control, however, cannot be exerted solely at PFK. Unless the supply of hexose phosphate to PFK is maintained, activating it would simply lead to a drastic fall in

38

DAVID A. HARRIS

F6P (and G6P) level, PFK would be deprived of its substrate, and its activity would fall. In rapidly contracting muscle, blood supply is restricted, and with it the source of glucose (and oxygen). In the short term, therefore (sprinting for 10-20 seconds), the source of hexose phosphate is glycogen stored within the muscle. Glycogen phosphorylase is, therefore, the other key enzyme regulated in glycolysis in muscle (Figure 14). Control at the nucleotide level is exerted, again, by AMP which cooperatively activates this enzyme. In addition, this enzyme is inhibited by glucose-6- phosphate. The stimulatory effect of AMP is thus amplified by the reduction in G6P inhibition as PFK is stimulated and sugar phosphates used up. Unlike PFK, glycogen phosphorylase is saturated with its substrate glycogen, and is thus unaffected by even quite large falls in glycogen concentration. When activated (via an increase in Vmax) phosphorylase can maintain a high level of hexose phosphate production until glycogen is nearly exhausted; it catalyzes the flux generating step of the pathway. In severe exercise, muscle AMP levels mayrisehigh enough for this nucleotide to undergo deamination to inosine monophosphate (IMP), producing NH3. The reason for this is uncertain; it may remove AMP so that adenylate kinase activity is not impaired and ADP build up (which would inhibit actomyosin) is prevented. NH3 also stimulates PFK and thus promotes ATP synthesis. However, prolonged activity at this level will deplete the cell of adenine nucleotides and thus cannot be sustained. Up to 20% of adenine nucleotides can be lost in this way; they are regenerated (the lost NH3 being replaced from aspartate) when the muscle returns to rest. Control by Other Agents In sustained contraction, cytoplasmic [Ca^"^]risesfromaround 0.3 |iM to 1.5 \xM. After stimulation by epinephrine (e.g., in anticipation of exercise) cAMP levels in the cell are raised between roughly the same concentrations. The effect of either is to stimulate glycogen phosphorylase via phosphorylation, which is mediated by phosphorylase kinase. Phosphorylation converts phosphorylase from an AMP-dependent form (phosphorylase b) to a form active independently of AMP (phosphorylase a), thus increasing the supply of sugar phosphates to PFK even without significant changes in nucleotide levels. An increased F6P level will stimulate PFK activity, both because F6P is a substrate and the enzyme is normally unsaturated and because F6P is an allosteric activator of PFK. Ca^"^ ions act directly on phosphorylase kinase, whose smallest subunit is calmodulin. This binds Ca^"^ as levels rise and the enzyme is activated. cAMP acts indirectly via protein kinase A; it is the phosphorylation of phosphorylase kinase by this enzyme which activates it. In both these cases magnitude amplification occurs because activation of glycogen phosphorylase involves phosphorylation by

Cellular ATP

39

a controlling enzyme, a few molecules of regulator (e.g., of epinephrine at the cell membrane) can activate a large number of enzyme molecules. Both these effects will stimulate glycolytic ATP synthesis in conditions of increased ATP utilization (or potential utilization) without requiring changes in nucleotide levels, and are thus useful ATP homeostatic mechanisms. However, they are not essential for normal function of muscle; I-strain mice, which lack phosphorylase kinase, can exercise normally. Presumably, in this condition, AMP serves adequately as a sole regulator molecule. A more subtle effect of epinephrine is that it will stimulate muscle FBPase in addition to PFK, possibly via the regulator fructose 2,6 bisphosphate. In other words, this hormone will increase the rate of substrate cycling in muscle. While this in itself will not lead directly to increased ATP production, it will increase the sensitivity of the pathway to regulators such as AMP. In other words, in epinephrine-stimulated muscle, a given rise in AMP will increase glycolytic flux more than in unstimulated muscle. Restoration of Blood Supply

Severe exercise can continue only for 1-2 minutes, since after this time muscle glycogen supplies are exhausted, and the pH of the muscle has fallen sufficiently to cause fatigue. For more prolonged exercise, the blood supply must be restored to remove H"*" ions and provide new substrate (glucose). Moderate exercise, in type lib muscle, can be driven for much longer under these circumstances. Indeed, patients lacking glycogen phosphorylase (McArdle's syndrome) cannot use muscle glycogen; however, they can tolerate moderate exercise entirely using blood glucose as a substrate. As might be expected, glucose entry into muscle is stimulated in exercise. However, it is not yet known how the glucose transporter senses a change in cellular energy demand. It is known that the transporter involved—^the exercise dependent glucose transporter—^is distinct from the insulin-dependent glucose transporter (glut-4) in the cell membrane (Barnard and Youngren, 1992), and is independently stimulated during exercise. It is possible that [Ca^"^] is the internal signal for this stimulation. Defects in Glycolysis

Because some tissues use glycolysis to provide all their ATP, a total absence of a glycolytic enzyme in man would be fatal. In some conditions, however, a partial deficiency is observed, leading to lowered ATP levels in such tissues. An example occurs in a common form of hemolytic anemia, in which pyruvate kinase levels in erythrocytes are reduced to 20% of normal. The erythrocytes lose their shape and lyse readily. In these cells, ATP levels fall and the erythrocyte is unable to maintain

40

DAVID A. HARRIS

its cytoskeleton (hence loss of shape) or Na"^ extrusion via the NaVK"^ pump (hence osmotic lysis). Control of Aerobic ATP Production Control in heart muscle is taken as the archetype of such regulation. Since the fuels used are typically triglyceride and lactate (in the fed state), major control points lie in the mitochondrial oxidative phosphorylation system. The work rate of heart can vary by up to 10-fold from minimal to maximal activity, by changes in heart rate and force of contraction (inotropy). Control by Nucleotide Levels Any discussion of the control of mitochondrial ATP synthesis must consider the fact that ATP utilization occurs in the cytoplasm while synthesis occurs inside the mitochondrion. Thus, some mechanism must exist for signaling changes in energy demand between these different cellular compartments. At the nucleotide level, the metabolic signal appears to be ADP (AMP, the cytoplasmic signal, cannot enter mitochondria). Two mitochondrial enzymes respond directly to ADP levels: the adenine nucleotide translocase and the ATP synthase (above). Both of these use ADP as substrate, and a rise in ADP levels will cause an increase in their activity, and an increase in ATP synthesis rate. (Note that these enzymes are not formally regulated by ADP; they are simply responding to its availability as a substrate.) Which of these enzymes might be limiting in vivo has been a matter of some debate. The answer probably depends on the tissue under consideration. In heart, the capacity of the translocase is high (Doussiere et al., 1984), and mitochondrial ADP levels may simply reflect cytoplasmic levels. Changes in ADP concentration thus affect overall flux by increasing ATP synthase turnover. In liver, the translocase activity is lower and ADP entry may limit access to the synthase. On these considerations, and on the basis of studies with mitochondria in vitro, it was suggested that mitochondrial ATP synthesis rates were simply controlled by varying the levels of substrate, cytoplasmic ADP. Such a model requires that, in vivo, cytoplasmic ADP levels vary around K^^^ for ATP synthesis (ca 30 |LIM) over a sufficient range (5-10-fold) to account for the observed rate changes. In skeletal muscle (human arm) this may well approximate to the true situation; NMR measurements of ATP synthesis and cytoplasmic [ADP] levels show a suitable relationship between ATP synthetic flux and increases in [ADP] with increasing levels of exercise. In heart, however, this is not the case. Over a wide range of heart work rates, both free [ATP] and [ADP] levels (the latter calculated from ATP/PCr ratios, above) remain effectively constant. Thus, again, we find that nucleotide levels are

Cellular ATP

41

very precisely controlled, and that changes in their concentrations are too small to be responsible for observed changes in flux. Control by Ca^^ Levels A second problem with the above model is that substrate ADP controls only the ATP synthase enzyme. This could not sustain an increased rate of ATP synthesis alone, since the proton gradient would soon fall due to a lack of oxidizable substrate. Thus, control of the ATP synthase must be integrated with control of NADH generating reactions: the Krebs cycle dehydrogenases. The messenger which communicates the need for ATP in the cytoplasm to these enzymes is Ca^"^ (Figure 15); other putative messengers such as AMP and cAMP, which control glycolysis, cannot enter the mitochondrion. Ca^"^ enters the mitochondrion through an electrogenic uniport, i.e., it uses the transmembrane potential (inside negative) as a source of energy. It can leave via an electroneutral 2Na"*"/Ca^'^ antiporter. The relative affinities of these systems lead to intramitochondrial [Ca^"^] reaching a steady state at levels (1-10 JLIM) 3-10 x higher than cytoplasmic [Ca^"*"] (0.3-1.5 |LIM). Importantly, however, changes in intramitochondrial [Ca^^] will reflect changes in cytoplasmic [Ca^"*"], and this ion is thus a signal of cytoplasmic contractile activity (energy demand) (Denton and McCormack, 1990). Four mitochondrial enzymes are known to be stimulated by physiological levels of [Ca^"*"]. Ca^"*" stimulates the phosphatase which dephosphorylates (activates) the pyruvate dehydrogenase complex; oxidation of cytoplasmic pyruvate (from lactate and, less commonly, glucose) is stimulated. Ca^"*" allosterically activates isocitrate dehydrogenase and a-oxoglutarate dehydrogenase, two key regulatory enzymes in the Krebs cycle; this stimulates NADH production from all substrates, notably fatty acids. And Ca^"^ activates the ATP synthase, probably by displacing a regulator protein which normally inhibits its activity (Figure 15) (Harris and Das, 1991). To sum up, increased contractile activity in heart leads to an increased mean cytoplasmic [Ca^"*"], leading to a rise in intramitochondrial [Ca^"^]. (The relatively low rate of the mitochondrial transporters smooth out the cyclic variations in cytoplasmic [Ca^^ occurring at each beat cycle.) Oxidation of pyruvate is stimulated, as is oxidation of acetyl CoA derived from fatty acid oxidation, and NADH levels rise. NADH dehydrogenase, which is normally not saturated with NADH, thus increases in activity and oxidation rates rise. Simultaneously, the ATP synthase is switched on and ADP is converted more rapidly to ATP. Thus, a new steady state arises, with increased flux through the respiratory chain and ATP synthase, with a limited change (commonly a small decrease) in the intermediate proton gradient. Cytoplasmic ATP levels are maintained, without any significant change in cytoplasmic ADP or AMP levels.

42

DAVID A. HARRIS

Evidence for the operation of this system comes from studies with ruthenium red, an inhibitor of mitochondrial Ca^"*" uptake (Unitt et al., 1989). Treatment of perfused heart, or cultured heart cells, with this compound inhibits activation of the dehydrogenases and of the ATP synthase which normally occurs with increased work rate, showing the importance of Ca^^ as an intracellular messenger. Furthermore, while untreated cells can maintain their ATP levels constant irrespective of work load, cells in which mitochondrial Ca^"^ uptake is blocked with ruthenium red suffer a drop in [ATP] at high work rates, showing the importance of this mechanism in maintaining normal energy balance in the heart. The effects of Ca^"*" above are discussed largely in terms of Ca^"*" released from the sarcoplasmic reticulum during contraction. However, oxidative phosphorylation can also be stimulated by positive inotropic agents (e.g., epinephrine). In this case, (cAMP-stimulated) protein kinase A phosphorylates an L-type Ca^"*" channel in the cell membrane, promoting (transient) Ca^"^ entry and raising cytoplasmic [Ca^"^]. This then acts on intramitochondrial processes as described above. Defects of Aerobic A TP Production Enzyme deficiencies in Krebs cycle enzymes generally lead to widespread tissue damage (and death), because of the central role of this pathway in a variety of metabolic pathways (including ATP generation). Deficiencies of pyruvate dehydrogenase are known, and generally lead to death in infancy; here, however, damage is localized in the brain because of the dependence of this tissue on glucose oxidation, which requires this enzyme. Tissues such as muscle and liver can utilize alternative fuels. Deficiencies in respiratory chain components also occur in mitochondrial membranes. Indeed, they appear relatively common compared to other genetic disorders. Such defects lead to functional problems in actively oxidizing tissues (i.e., to neuropathies and myopathies). However, location of the damage and severity of the symptoms vary greatly between sufferers; some show chronic tissue damage, while others are able to generate ATP adequately except under conditions of severe stress. This is because many such defects (NADH dehydrogenase, cytochrome oxidase deficiency, etc.) result from lesions on mitochondrial DNA (mDNA) (Harding, 1991). Since one cell contains several autonomously replicating mitochondria (each containing several copies of mDNA), normal and mutant mDNA may be present in the same cell. The severity and location of the functional lesion will then depend on the proportion of mutant mDNA in cells of a given tissue; this in turn will reflect how early in the lineage of the cell type the mutation arose. Less severe changes are observed in conditions (such as hypertension or alcoholic cardiomyopathy) where Ca^"*'-dependent regulation of the ATP synthase is defective. In such cases, ATP levels in the heart cannot be precisely maintained as work rate increases. Thus, ATP levels fall (ca. 20%) with a consequent rise in

Cellular ATP

43

[ADP]. Nonetheless, a new steady state is achieved. It seems that, in the absence of functioning regulation by [Ca^"^], ATP synthesis rates in heart can be increased through substrate control by ADP, as observed in skeletal muscle (above).

PATHOLOGICAL DISTURBANCES OF ATP LEVELS We have seen above how defects in the enzymes of the glycolytic pathway, or the pathway of oxidative phosphorylation, cause severe pathological changes in ATP levels, since ATP utilization continues in the absence of normal ATP synthesis rates. Other conditions exist where an imbalance exists between ATP synthesis and utilization rates, with consequent abnormalities in ATP levels. Malignant Hyperthermia (Malignant Hyperpyrexia)

This condition is characterized by a massive increase in aerobic and anaerobic ATP generation induced by the administration of volatile anesthetics (e.g., halothane). Metabolic activity is so high that body temperature rises (up to 44 °C in extreme cases) and severe lactic acidosis results. The primary defect in this syndrome lies in the Ca^"^ channel of sarcoplasmic reticulum in skeletal muscle. Halothane prolongs opening of this channel, and cytoplasmic [Ca^"^] rises sharply (Heffron, 1988). This activates the actomyosin ATPase and ATP is rapidly hydrolyzed, with consequent heat production. In addition, the Ca^^ pumps of the sarcoplasmic reticulum and plasma membrane are stimulated in an attempt to remove the excess Ca^"^, leading to further ATP hydrolysis. These rises in Ca^"^, ADP, and AMP will stimulate glycolysis and oxidative phosphorylation as described above. Oxidation, in fact, does not reach its maximal capacity, either because pyruvate transport is too slow or because of the effect of high temperature on the mitochondrial membranes; thus oxidation rates rise, typically only 3-4 times. Glycolytic rates are stimulated much more greatly, and considerable lactate is produced even by aerobic muscle. Nonetheless, ATP levels cannot be maintained, and over a 15-30 minute period, [PCr] and [ATP] levels fall below the levels needed to maintain cell viability. Malignant hyperthermia is not a particularly rare genetic disease (incidence ca. 1/12,000) and, while it has little effect on physiological function in everyday life, is a major problem in anesthesiology, particularly as no simple diagnostic test for the condition exists. Ischemia

Most cells can maintain their ATP levels during anoxia; cultured heart cells, for example, retain normal ATP levels for more than 30 minutes in the absence of oxygen. This shows that cells can switch smoothly between glycolysis and oxida-

44

DAVID A. HARRIS

tive phosphorylation as sources of ATP. If the blood supply to heart muscle is interrupted in vivo, however, consequences are much more severe. ATP levels rapidly fall by some 20%, and are maintained at this level. More importantly, however, are the changes that occur when blood supply is restored; if it has been interrupted for more than 20 minutes, reperfusion leads, surprisingly, to further ATP depletion and commonly cell death. This deleterious effect of reintroducing oxygen is known as the oxygen paradox. Since the heart is unable to regenerate new cells, this process leads to permanent damage of some area of the heart—a. myocardial infarct. Similar effects are observed in the brain tissue (stroke). The difference between anoxia and interruption of blood supply lies in the build up, in the latter, of the products of anaerobic metabolism, particularly H"^. Reoxygenation from pH 7.4 has no deleterious effects; reoxygenation (and restoring the tissue to pH 7.4) from pH 6-6.5 causes injury. This is associated with a massive influx of Ca^"*"fromthe blood, probably via the Na"^/Ca^'*" exchanger in the cell membrane (Tani, 1990). It is this abnormal Ca^"*^ load that increases energy utilization (via Ca^"*" pumps and actomyosin stimulation); in addition, it is sufficiently high to overload the mitochondria (possibly precipitating Pj internally) and to inhibit oxidative phosphorylation. Thus, ATP utilization increases, ATP production falls, and ATP levels drop precipitately. It is still unclear why reoxygenation/pH change of heart cells leads to this pathological inflow of Ca^"^. Agents such as free radicals (generated in partial oxidations) and activated phospholipases may damage the cell membrane under these conditions, but it is still uncertain whether the magnitude of such effects is sufficient to explain the dramatic increase in Ca^"^ influx on reperfusion.

SUMMARY ATP is a kinetically stable molecule with a high free energy of hydrolysis/high phosphate transfer potential. This means it can act as a common unit of exchange of energy between a variety of highly exergonic catabolic processes and energy requiring reactions within the aqueous medium of a cell. The chemical nature of the ATP molecule means that it can drive a wide variety of such reactions, including movement of ions and proteins, dehydrations (in macromolecule biosyntheses), activation of small molecules, and imparting a negative charge to sugars and proteins. The relative contributions of these processes to the energy demand of a cell depends on the tissue; muscle expends some 70% of its ATP turnover on movements of actomyosin, brain about 40% of its ATP on Na^ transport, and exocrine cells about 50% on biosyntheses. The molecular basis for the coupling of ATP hydrolysis to non-chemical processes (e.g., muscle contraction, ion pumping) is not precisely known. A model is presented in which the transducing enzyme can manipulate the stages in the release of energy from ATP to link binding energy changes to conformational changes. This rationale unifies existing schemes for the function of these enzymes,

Cellular ATP

45

and also for the functioning of the mitochondrial ATP synthase, which couples transmembrane proton flow to ATP synthesis. In humans, the bulk of ATP synthesis occurs via oxidative phosphorylation in mitochondria, although the fuel oxidized is dependent on tissue. In some tissues (and in rapidly growing tumors), however, considerable amounts of ATP are made by glycolytic conversion of glucose to lactate. This reflects an adaptation of these tissues to specific functions; for example, sustained contraction (and hence restricted O2 supply) in white muscle or a high requirement for biosynthetic precursors (e.g., in lymphocytes). ATP levels inside cells are maintained very precisely (around 8 mM) under all physiological conditions by increasing the rate of ATP synthesis to match demand. In contrast, rates of ATP synthesis can vary greatly (5-100-fold) with the energy demands of the tissue. Together, these facts indicate that ATP itself cannot regulate its own synthesis. Cytoplasmic (glycolytic) ATP synthesis is regulated internally by AMP, changes in whose concentration amplify the changes in [ATP]; other regulators are Ca^"*" and cAMP, which signal an actual or potential increased work rate by the tissue. Mitochondrial (oxidative) ATP synthesis is regulated by cytoplasmic ADP (entering via the adenine nucleotide translocase) and/or by cytoplasmic Ca^"^ (entering via the Ca^"^ uniport), depending on the tissue. Prolonged ATP depletion leads to cell death, largely due to the development of ionic and osmotic imbalance. Such depletion occurs in a variety of clinical conditions. There may be deficiencies in the enzymes of ATP production (e.g., pyruvate kinase, pyruvate dehydrogenase, mitochondrial dehydrogenases) or conditions which lead to abnormal ATP utilization (e.g., fructose intolerance, malignant hyperpyrexia, ischemia/reperfusion). The resulting symptoms vary considerably, depending on the tissue most susceptible to the defect.

ACKNOWLEDGMENTS I thank the Wellcome Trust, and the British Heart Foundation for financial support. I am grateful to Professor G.K. Radda for supplying Figures 2 and 4, and to Dr. J. Clarke and Dr. A.M. Das for their help and encouragement.

REFERENCES Ardawi, M.S.M. & Newsholme, E.A. (1985). Metabolism in lymphocytes and its importance in the immune response. Essays Biochem. 21, 1-44. Ashcroft, F.M. & Rorsman, P. (1989). Electrophysiology of the pancreatic p cell. Prog. Biophys. Mol. Biol. 54, 87-143. Balaban, R.S., Kantor, H.L., Katz, L.A., & Briggs, R.W. (1986). Relation between work and phosphate metabolites in the in vivo paced mammalian heart. Science 232, 1121-1124. Boyer, P.D. (1987). The unusual enzymology of the ATP synthase. Biochemistry 26, 8503-8507. Browner, M.F. & Fletterick, M.F. (1992). Phosphorylase; a biological transducer. Trends. Biochem. Sci. 17,66-72.

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Barnard, R.J. & Youngren, J.F. (1992). Regulation of glucose transport in skeletal muscle. FASEB J. 6,3238-3244. Denton, R.M. & McCormack, J.G. (1990). Ca^"*" as a second messenger within the mitochondria of the heart and other tissues. Ann. Rev. Physiol. 52,451-^66. Doussiere, J., Ligett, E., Brandolin, G., & Vignais, P.V. (1984). Control of oxidative phosphorylation in rat heart mitochondria: Role of the adenine nucleotide carrier. Biochim. Biophys. Acta 766, 492-500. Ferguson, S.J. (1986). The ups and down of P/0 ratios. Trends Biochem. Sci. 11, 351-352. Harding, A.E. (1991). Neurological disease and mitochondria. Trends Neurosci. 14, 132-138. Harris, D.A. & Das, A.M. (1991). Control of mitochondrial ATP synthesis in the heart. Biochem. J. 280,561-573. Heffron, J.J.A. (1988). Malignant hyperthermia—biochemical aspects of the acute episode. Brit. J. Anaesthesia 60, 274-278. Hibberd, M.G. & Trentham, D.R. (1986). Relationships between chemical and mechanical events during muscular contraction. Ann. Rev. Biophys. Biophys. Chem. 15, 119-161. Hyde, S.C, Emsley, P., Hartshorn, M.J., Mimmack, M.M., Gileadi, U., Pearce, S.R., Gallagher, M.P., Gill, D.R., Hubbard, R.E., & Higgins, C.F. (1990). Structural model of ATP binding proteins associated with cystic fibrosis, multidrug resistance and bacterial transport. Nature 346, 362—365. Jacobus, W.E. (1985). Respiratory control and the integration of heart high energy phosphate metabolism by mitochondrial creatine kinase. Ann. Rev. Physiol. 47, 707—725. Kauppinen, R.A., Hiltunen, J.K., & Hassinen, I.E. (1980). Subcellular distribution of phosphagens in the perfused rat heart. FEBS Lett. 112, 273-276. MacLennan, D.H. (1990). Molecular tools to elucidate problems in excitation/contraction coupling. Biophys. J. 58, 1355-1365. Nelson, N. (1992). Organellar proton ATPases. Curr. Opinion Cell Biol. 4, 654-660. Radda, G.K. (1986). The use of nmr spectroscopy for the understanding of disease. Science 233, 640-645. Radda, G.K. (1992). Control, bioenergetics and adaptation in health and disease: Non-invasive biochemistry from nmr. FASEB J. 6, 3032-3038. Robitaille, P.M., Merkle, H., Lew, B., Path, G., Hendrich, K., Lindstrom, P., From, A.H.L., Garwood, M., Bache, R.J., & Ugurbil, K. (1990). Transmural high energy phosphate distribution in the normal canine myocardium as studied with spatially localised ^'P nmr spectroscopy. Mag. Reson. Med. 16,91-116. Seiss, E.A. & Wieland, O.H. (1976). Phosphorylation state of cytosolic and mitochondrial adenine nucleotides and of pyruvate dehydrogenase in isolated rat liver cells. Biochem. J. 156,91—102. Senior, A.E. (1988). ATP synthesis by oxidative phosphorylation. Physiol. Rev. 68, 177-231. Tani, M. (1990). Mechanisms of Ca^"*" overload in reperfused ischemic myocardium. Ann. Rev. Physiol. 52, 54S-559. Unitt, J.F., McCormack, J.G., Reid, D., Maclachlan, L.K., & England, P.J. (1989). Direct evidence for a role of intramitochondrial Ca^"^ in the regulation of oxidative phosphorylation in stimulated rat heart. Biochem. J. 262,293-301. Veech, R.L., Lawson, J.W.R., Cornell, N.W., & Krebs, H.A. (1979). Cytosolic phosphorylation potential. J. Biol. Chem. 254, 6538-6547. Watford, M., Lund, P., & Krebs, H.A. (1979). Isolation and metabolic characteristics of rat and chicken enterocytes. Biochem. J. 178, 589-596. Zweier, J.L., Jacobus, W.E., Korecky, B., & Brandejs-Barry (1991). Bioenergetic consequences of cardiac phosphocreatine depletion induced by creatine analogue feeding. J. Biol. Chem. 266, 20296-20304.

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RECOMMENDED READINGS Books Newsholme. E.A. & Leech, A.R. (1983). Biochemistry for Medical Students. John Wiley, New York. Devlin, T.M. (1992). Textbook of Biochemistry with Clinical Correlations, 3rd ed. John Wiley, New York. Harris, D.A. (1995). Bioenergetics at a Glance. Blackwell Science, Oxford.

Articles Capaldi, R.A. (1988). Mitochondrial myopathies and respiratory chain proteins. Trends Biochem. Sci. 13, 144-147. Clausen, T., van Hardeveld, C.V., & Everts, M.E. (1991). Significance of cation transport in control of energy metabolism and thermogenesis. Physiol. Rev. 71, 733-774. Erecinska, M., Bryla, J., Michalik, M., Meglasson, M.D., & Nelson, D. (1992). Energy metabolism in the islets of Langerhans. Biochim. Biophys. Acta 1101, 273-295. Hanson, R.W. (1989). The role of ATP in metabolism. Biochem. Educ. 17, 86-92. Pedersen, P.L. & Carafoli, E. (1987). Ion motive ATPases. I. Ubiquity, properties and significance to cell function. Trends Biochem. Sci. 12, 146-150. Pendersen, P.L. & Carafoli, E. (1987). Ion motive ATPases. II. Energy coupling and work output. Trends Biochem. Sci. 12, 186-189. Tanford, C. (1983). Mechanism of free energy coupling in active transport. Ann. Rev. Biochem. 52, 379-409.