``What stronger Breastplate than a heart untainted!''

Shakespeare (1623)

King Henry VI, Part II: III:ii

2.1 Introduction

The delivery of oxygen, water, nutrients, and hormones to tissues, the removal of wastes, and the regulation of temperature in vertebrates is accomplished by the circulatory system of which the central component is the heart. Though certain morphological and structural differences exist across species, the function of the heart is essentially the same: to pump blood through the circulatory branches through periodic, rhythmic contraction. Here, a brief overview of the processes and principles involved in the physiology of the heart is presented along with a review of techniques for studying these phenomena. Finally, a discussion of the effects of hypoxia on the heart is presented and motivation for the study of Chrysemys picta bellii is explained.

2.2 Overview of the Cardiac System

The flow of fluid through the circulatory system is driven by the rhythmic pumping action of the heart. In mammals, reptiles, and amphibians, the heart consists of essentially two pumps in series which sit side by side and contract together. One side accepts deoxygenated blood which returns from the body and pumps it to the lungs. The other side accepts oxygenated blood from the lungs and pumps it through the aorta(s) to the body tissues. Each pump consists of two chambers. A thin-walled, elastic atrium accepts blood at low pressure from veins and pumps it into a thicker-walled chamber, the ventricle. The ventricle then contracts and expels blood through arteries. In mammals, the two pumps are completely separate consisting of a right atrium and ventricle and a left atrium and ventricle. In reptiles and amphibians, however, only a single ventricle exists allowing oxygenated and deoxygenated blood to mix to some extent. The degree of mixing is limited by the presence of a septum which partially divides the chamber. The extent of the septum varies across species. It is generally accepted that increasing to complete division of the right and left circulation is an evolutionary adaptation which allows for a greater sustained metabolic rate in higher vertebrates [23].

Figure 2.1 shows a typical reptilian heart from the lizard Lacerta virdis [24]. The figure clearly illustrates the two atria and single ventricle present in reptilian hearts as well as the important vessels. The primary morphological differences between this lizard heart and the heart of Chrysemys picta bellii is that in the latter, the auricles are more dorsally oriented with most of the major vessels attaching at the dorsal aspect of the heart. A schematic circulatory diagram of the same heart is presented in Figure 2.2. For Chrysemys, the ductus arteriosus (da) which connects the left pulmonary artery and the aorta still exists [24].

Abbreviations for Figures 2.1 and 2.2:

ao, dorsal aorta. av, left auriculo-ventricular valve. bisa, base of left systeinic arch (with valve). brsa, base of right systemic arch (with valve), da, ductus arteriosus (probably nQn-functionat in tacerta). dc, ductus caroticus. dch, dorsal chamber. dcha, part of dorsal chamber (cavum arteriosum) on left of muscular ridge. dchv, part of dorsal chamber (cavum venosum) on right of muscular ridge. eca, external carotid artery. fp, foramen of Panizza. ica, internal carotid artery. lau, left auricle. ice, left common carotid artery (carotid arch). lpa, left pulmonary artery. lsa, left systemic (aortic) arch. lv, left ventricle. mr, muscular ridge or secondary septum. ola , opening of left auricle. ora, opening of right auricle. put, pulmonary trunk. pv, pulmonary valve. pvc, pulmonary vein. rau, right auricle. rec, right common carotid artery (arch). rpa, right pulmonary arteryrsa, rsa right systemic (aortic) arch. rv, right ventricle. sbc, subclavian artery. sv , opening of sinus venosus. V, ventricle. vch, ventral chamber. vs, ventricular septum.

Figure 2.2: Schematic Representation of the Great Vessels in the Reptilian Circulatory System. From A. Bellairs: Life of Reptiles . vol. 1.

2.3 Cardiac Mechanics

The heart pumps blood in a cyclic pattern which is divided into two phases. During the resting or diastolic phase, the heart muscle is relaxed and allows the flow of blood into its chambers. During the contractile or systolic phase, the heart muscle is contracting and forcing blood out of the chambers and into vessels. Atria contract first in systole and force blood into the ventricles. When the ventricle(s) begins to contract, blood flow from the atria (auricles) ceases and blood is expelled through the great arterial vessels. Contraction of the ventricles occurs by the combined contraction of many cardiomyocytes (individual cardiac muscle cells) each of which contains many bundles of myofibrils made up of chains of sarcomeres [17]. The primary structure responsible for this contraction is the sarcomere. A sarcomere consists of bundles of two types of myofibrils (actin and myosin) which overlap in regular ways to form visibly distinguishable bands. A typical cardiac sarcomere is illustrated in Figure 2.3. One complete period of such an overlap is a sarcomere. When electrical depolarization of the muscle cell takes place, the actin and myosin filaments interact and cause the sarcomere to physically shorten in length. The chemically driven shortening of many sarcomeres provides the force involved in contraction. Several factors including the amount of overlap (Starling's Law) affect the available force of this contraction and will not be discussed in detail here [17].

Figure 2.3: A Typical Cardiac Muscle Sarcomere . The sarcomere consists of isotropic (I) bands consisting of actin filaments only, Z lines where these filaments are cross-linked, H bands consisting of myosin filaments only, anisotropic (A) bands where the actin and myosin filaments overlap, and an M band where myosin filaments are cross-linked. A myofibril consists of a chain of many sarcomeres.

Contraction of the sarcomeres is initiated when the cell containing them becomes electrically depolarized. Through special connections between muscle cells called intercalated disks, this wave of depolarization is allowed to spread from cell to cell causing a quick contraction of the entire heart muscle. In addition, special electrically conductive fibers help to distribute this wave so that the heart contracts efficiently and uniformly. It is this wave of electrical activity in the heart that is the basis of the electrocardiogram (ECG).

2.4 Excitation Contraction Coupling

The contraction of cardiac muscle is a process which is driven by chemical energy. However, the process is initiated by electrical activity. Thus, some mechanism must exist whereby electrical excitations are coupled to chemical reactions and concomitant force development and work production. It is free ionized calcium which plays a central role in this link, and the precise maintenance of calcium levels throughout the cardiac cycle is required for proper contractile function. Here, a description of the role of free ionized calcium in cardiac contraction will be presented.

Intracellular cytosolic free Ca²⁺ concentrations are low while the cells are in their diastolic non-contractile state, typically on the order of 0.1 mM (1 M = 1 mole/liter) [17]. Upon depolarization of the cardiomyocyte, intracellular Ca²⁺ levels rise appreciably (to as much as 1-10 mM). In addition to balancing a net efflux of potassium (K⁺) out of the cell and thus sustaining the membrane depolarization, rising calcium in the cell causes an intracellular membrane structure called the sarcoplasmic reticulum to release its stores of calcium (calcium-activated calcium-release). Though the amount of calcium which enters the cell from the extracellular space is not sufficient to cause myofibril contraction, it triggers the release of sarcoplasmic Ca²⁺ stores and the concentration of Ca²⁺ is then high enough to induce contraction.

Free Ca²⁺ in the cytosol binds to the exposed protein troponin-C which is complexed with tropomyosin and wrapped around the actin (thin) filaments of the sarcomeres. This Ca²⁺ -troponin-C complex then influences tropomyosin and induces a conformational change in the latter. When tropomyosin changes form, it recedes from cross-bridge binding sites on the actin filament. These exposed sites quickly form a strong bond with the globular heads on the cross-bridge arms of neighboring myosin filaments. Upon binding to actin, the myosin heads release bound ADP and P_i molecules which changes the head's energy state and causes a conformational change in myosin and a bending of the head forward. This change is the basis of the contractile force of the actin-myosin fiber complex.

When the head cocks forward, it alters the myosin head's affinity for ATP which, if available, will bind to the myosin head. When the head binds ATP, however, it loses its affinity for the actin filament, and the actin-myosin bond is broken. Once free of actin, the myosin head partially hydrolyzes the ATP into bound ADP and P_i. This free-energy change in the molecule causes it to once again undergo a conformational change back to the initial unbent state with a renewed affinity for actin. The cycle repeats with the filaments moving 10 nm or so each time until Ca²⁺ levels fall and tropomyosin once again blocks actin cross-bridge binding sites. The contraction of cardiac muscle is depicted schematically in Figure 2.4 below.

Figure 2.4: Excitation Contraction Coupling. A) With no calcium present, tropomyosin blocks affinity binding sites on the actin filament. B) The conformation of tropomyosin is changed when free calcium complexes with it through the mediator protein troponin C; the actin binding sites are exposed. The head on the myosin molecule binds to the actin filament and releases bound ATP and Pi causing a concomitant conformational change and relative movement of the actin and myosin fibers. C) After movement, myosin is able to bind and hydrolyze ATP which causes the actin-myosin bond to break and myosin to return to its original configuration. Cyclic binding and movement of actin and myosin ensues so long as calcium is present and ATP is available. D) When calcium is extruded, the filaments return to their inactive state.

The influx of calcium into the cell from the extracellular space is precipitated by a depolarization of the cell membrane. The resting cell membrane potential is about -90 mV. At this voltage, Na⁺ channels and Ca²⁺ channels are closed and large concentration gradients exist between the extra and intracellular compartments for both of these ions. Ca²⁺ bound to weakly negative sites in a mucopolysaccharide coat covering the cell membrane called the glycocalyx is in close equilibrium with the interstitial fluid and is the source of Ca²⁺ for calcium-ion channels. Conversely, intracellular K⁺ concentration is high relative to the extracellular and it is K ⁺ that is primarily responsible for the relative membrane potential. As neighboring cells depolarize, their potential is communicated to each other via intercalations which allow communication between the cytosol of adjacent cells. Partial depolarization of the cell causes fast voltage-activated Na⁺ channels to open and for sodium to flood into the cell. This large positive influx further depolarizes the cell and causes the large sharp spike in the beginning of the action potential (AP). When the cell depolarizes, other slower voltage-activated channels for both K⁺ and Ca ²⁺ are opened. Ca²⁺ channels are opened through phosphorylation by a cyclic-AMP (cAMP)-dependent protein kinase [17]. An initial efflux of K⁺ out of the cell causes a small repolarization dip in the AP, but within 10 ns or so, this dip is offset by a flux of Ca²⁺ (and thus positive charge) into the cell. Ca²⁺ enters the cell through two distinct sets of channels: T-type and L-type. The T-type or transient channels become active when the membrane potential climbs to about -70 mV. As their name suggest, however, they become inactivated after 100-200 ms and Ca²⁺ current through them ceases. These channels are more common in skeletal muscle cells than in cardiac muscle cells, but they still exist and are unaffected by commonly used Ca²⁺-channel blocking drugs [17].

L-type or long lasting channels do not activate until the membrane potential reaches approximately -10 mV or so. These channels become deactivated much more slowly and thus allow Ca²⁺ current to flow into the cell for several hundred ms after the beginning of the AP. This long lasting activation sustains the plateau phase of the action potential and delays repolarization. Unlike the T-Type, Ca ²⁺ L-type channels do respond to channel blocking drugs like verapamil, nifedipine, and diltiazem, and this action is the basis for certain hypertension therapies [17].

Once flux of Ca²⁺ into the cell from the extracellular compartment has been initiated, rising levels of cytosolic Ca ²⁺ cause the sarcoplasmic reticulum in the cell to release its stores of calcium and the concentration of Ca²⁺ rises sharply. The sarcoplasmic reticulum (SR) is a closed space within the cell consisting of a network of small diameter tubules in close proximity to the myofibrils. The SR has ATP driven Ca²⁺ pumps which drive calcium into the SR space and out of the cytosol when activated. Inside the SR, Ca²⁺ cations are bound to a protein complex called calsequestrin which can bind 40 or so Ca²⁺ cations. This serves to decrease the effective concentration of Ca²⁺ inside the cell and reduce the concentration gradient against which

calcium must be pumped. This is important since, in order to assure relaxation, the SR must be able to pump about 50 nmol Ca²⁺ /200 ms per gram of heart muscle. Not all of the SR's Ca ²⁺ comes from the cytosol, but rather from the extracellular fluid via invaginations in the cell membrane called T-tubules. The SR can communicate with the T-tubules and take up some calcium from the ECF. Release of Ca²⁺ from the SR is triggered by a rise in intracellular calcium through membrane channels and also by depolarization of the SR reticular membrane.

During diastole Ca²⁺ influx stops and the SR is no longer stimulated to release calcium. In fact, a cAMP-dependent protein kinase phosphorylates a protein known as phospholamban which stimulates the SR ATP-driven Ca²⁺ pump to take up calcium again. Simultaneously, troponin-I is phosphorylated and inhibits binding of Ca²⁺ by troponin-C which then causes tropomyosin to return to its normal position guarding actin-myosin cross-linking sites. Calcium is also actively driven out of the cell by an ATP-driven pump which extrudes Ca²⁺ and by a 3Na ⁺/1Ca²⁺ exchanger driven by the Na⁺ concentration gradient which is established by the ATP-driven 3Na⁺/2K ⁺ pump when sodium and potassium channels are closed. Figure 2.5 shows a schematic of calcium transients and the mechanisms responsible for them during the cardiac cycle. The duration of the ensuing calcium transient is approximately 300 ms or so [17].

Figure 2.5: Control of [Ca²⁺ ] Transients throughout the Cardiac Cycle. Initial opening of Ca channels allows calcium-activated calcium release of large Ca stores from the sarcoplasmic reticulum (SR) which is sufficient to initiate contraction. Ca is extruded from the cell by ATP-ase pumping accross the sarcolemma as well as into the SR. Gradient driven Na-Ca exchangers also help to extrude calcium. The net result is a transient in intracellular [Ca ²⁺] which lasts approximately 300 ms.

2.5 Cardiac Metabolism

The heart is an important user of energy in the mammalian body. This is exemplified by the fact that although it comprises only about 1% of total body weight, it is responsible for using about 10% of metabolic O ₂ [25]. Energy expenditures in the heart are generally divided into three categories: basal metabolism, activation energy, and physical work. Basal metabolism includes energy expenditures in the heart necessary to maintain basic cellular function but not directly related to excitation-contraction coupling or pumping. Activation energy is used to refer to energy spent to depolarize the cell, and generate the calcium transients and other events required for excitation-contraction coupling. Physical work refers to actual shortening of the muscle fibers against load and the work done in pumping, W = PDV where P is pressure, and DV is volumetric change.

2.5.1 Basal Metabolism

The resting cell requires energy for: 1) the maintenance of appropriate ion concentrations via membrane-bound ion pumps, 2) synthesis of proteins in the cell, 3) maintenance of pH balance, and 4) elimination of waste products. For the purposes of this discussion we will primarily investigate mammalian tissue under normoxic conditions. For rat papillary muscles, the basal rate of heat production is about 10 mW/g = 10 mJ/sec/g [26]. Typical cytosolic ATP hydrolysis yields about 56 kJ/mol in heat and ~ 30 kJ/mol in available free-energy (which can be used to do chemical work). This means that the cellular ATP requirement for basal metabolism is on the order of 178 nmol/sec/g of cardiac tissue [26].

The cell membrane is subject to constant slow leaks of Na⁺ , K⁺, and Ca²⁺ . In order to maintain normal concentrations of these ions, active ATP-ase pumps are located in the cell membrane. The Na⁺ -K⁺-ATPase pump extrudes 2 moles of sodium for every 3 moles of potassium brought into the cell. This pumping establishes an electrochemical gradient as well as concentration gradients. Based on known rates of Na⁺ ion leakage across the cell membrane, the energy required for this pump to maintain normal cationic concentration levels has been estimated to be anywhere from about 10%-25% of the total basal metabolic heat generation rate [26]. Ca ²⁺-ATPase pumps also contribute to the energy requirement of the cell and it is interesting to note however that even with large variations in extracellular calcium concentrations, basal metabolic rate does not change appreciably [25, 26].

Protein synthesis has been estimated to account for as much as 10% (1 mW/g) of the basal metabolic heat production in heart muscle [26, 27]. In fact, it is protein synthesis that is suspected to contribute primarily to interspecies variations in basal metabolic rate [26, 27]. It has also been found that cardiac antigenic properties (an indication of surface receptors and markers found on the cell surface) change as an adaptation to hypoxia indicating that protein synthesis is coupled in an important way to oxidative energy state [25, 26].

An interesting feature of cardiac muscle is that its basal metabolic rate is 5 or so times higher than that of skeletal muscle. An interesting hypothesis explains this difference on the basis of mitochondrial density [25, 26, 27]. Skeletal muscles, while similar in some ways to cardiac muscle, are designed to provide short-term bursts of mechanical energy. This generally depletes PCr stores which must be slowly rebuilt during periods of rest. Cardiac muscle does not have the luxury of rest, however, and must constantly maintain a PCr pool. The recovery heat is related to the immediate replenishment of energy stores after a contraction. In order to facilitate this constant ability to do work, cardiomyocytes are extremely rich in mitochondria as compared to skeletal muscle. This rich supply of mitochondria also leads to an excess of protons derived from the oxidation of fuel molecules. Since the ATP:O ratio of oxidative phosphorylation is 3 and the ATP:H ⁺ ratio of proton pumping is 2, 12 moles of protons are pumped for each mole of O₂ consumed. Since the energetic equivalent of O₂ is 448 kJ/mol, proton pumping could account for (based on assumptions about rat mitochondria density and proton generation) as much 6 mW/g of basal metabolic rate [26].

2.5.2 Activation Metabolism

As stated earlier, activation metabolism in cardiac muscle has an initial heat and a recovery heat associated with it. Activation metabolism (measured with no load) increases as heart contractility is modified via agents such as catecholamines [17]. Since such agents serve to increase peak systolic calcium concentration, this indicates that a significant portion of the active metabolism is used to remove contractile Ca²⁺ during contraction relaxation and to rebuild the energy stores depleted in doing so. This in fact accounts for the majority of the metabolic expenditure above the basal metabolic rate with a small amount of energy being lost in friction as fibers slide past each other. The total heat during activation metabolism, though variable across species, is on the order of 30 mW/g (for rabbit papillary muscle) and varies with body size and pre-load tension. This is a suprabasal metabolic rate of 20 W/g. [26]

2.5.3 Mechanical Work

Mechanical work is the most important effect of cardiac energy metabolism. It is estimated that the mechanical efficiency of the heart is on the order of 40-50% which means that of the energy allotted for contractile work generation, 40% is converted to mechanical work (blood pumping.) [25, 26]. One can estimate this requirement for humans by assuming a pumping rate of 5L/min over an average mean arterial pressure of 100 mmHg. Then d/dt[PDV] = 1.4 mJ/sec/g (assuming an 800g heart). Thus, the actual contractile work energy requirement is probably around 3.5 mW/g. [25]

2.5.4 Summary of Cardiac Metabolism

Armed with the information above, an energy budget can be constructed for the heart. This budget is of course crude because it is based on many assumptions, data from several species, and on speculative quantities from several sources. None the less, we can summarize the above information for mammals:

· Total cardiac energy consumption is about 33.5 mW/g.

· Basal energy consumption is about 10 mW/g. Of this, approximately 10-25% (we'll say 20%) goes to membrane bound ion pumps. Another 60% may go to proton pumping due to large numbers of mitochondria, and the remaining 20% will be allocated to protein synthesis.

· Activation energy consumption is about 20 mW/g above the basal rate and almost all of it is expended on resequestration of Ca²⁺ and the rebuilding of energy stores to do so.

· Normal cardiac work loads in humans probably require ~ 1.4 mW/g. If the conversion of chemical to mechanical energy is 40% efficient, then this translates to ~ 3.5 mW/g.

· Approximately 80-90% of the energy in glucose is extracted into forms useful for cellular processes.

Figure 2.6 summarizes the above information graphically.

It is generally accepted that for reptiles, total energy consumption per cardiac mass is approximately one third that of mammals. Thus, the total cardiac energy consumption for Chrysemys is probably on the order of 11 mW/g. The relative proportions of energy expenditure should however be similar.

Figure 2.6: Energy Budget of the Heart. While total energy expenditures vary between mammals and reptiles, the relative proportions should be similar for any vertebrate.

2.6 Measurements of Cardiac Function

Techniques for measuring and investigating cardiac function and performance are numerous. Of particular interest in this study are those which allow the quantification of metabolic responses to the condition of hypoxia. A brief review of several techniques will be presented here. Of particular interest are methods for assessing metabolic energy state and ion concentrations since 60% of the cardiac energy is spent on the maintenance of calcium homeostasis.

2.6.1 Gross Function

The gross function of the heart can be measured non-invasively via several techniques. Auscultation can diagnose specifics of anatomical functions such as valve openings and closings, stenoses, ventricular thickening, and heart rate among others [28]. Echocardiography allows for measurement of the range of motion of specific regions of the heart during the cardiac cycle [29]. The electrocardiogram tracks the depolarization and repolarization of the heart muscle itself [30]. Here, problems with the specialized electrical conduction pathway of the heart can be seen. Finally, imaging techniques such as fast magnetic resonance imaging (fMRI) [31] and ultrasound imaging [29] allow for direct visualization of the heart during its rhythmic cycle.

2.6.2 Metabolic Function

Measurement of cellular metabolism in the heart can be made in several ways. ³¹P NMR can be used to measure the relative concentrations of energy stores of PCr and ATP in the cell [3]. Direct calorimetry can be used to measure the heat produced by the cell [25, 26]. This measured DH is then related to DE, the energy expended by the heart and can be used to explore the relative energy expenditures of different portions of cardiac metabolism. Finally, tissue O₂ consumption (indirect calorimetry) can be used as a measure of cardiac metabolic expenditure [25, 26]. Since the primary source of cellular energy is ATP and ATP is generated primarily through oxidative phosphorylation under aerobic normoxic cellular conditions, then the utilization of O₂ can be indirectly related to the production of ATP which occurs in the ratio 6O₂:36ATP (for one glucose molecule).

Measurement of basal metabolism can be carried out in either perfused, K ⁺ arrested hearts or heart tissue. The results of several studies across mammalian species suggest that the basal metabolic rate is as high as 25-30% that of the active metabolic rate [26]. This is in stark contrast to striated skeletal muscle where basal metabolism is only 2-3% of active metabolism [26]. Basal metabolic rate increases with pre-load stress applied to the tissue, but it is relatively insensitive to temperature. Energy substrates presented to the cells can also influence basal metabolic rate.

Activation metabolic rate can also be measured either by O ₂ consumption or heat production [25, 26]. In order to measure only the activation metabolism, muscle preparations are generally pre-shortened so that they do not contract and no work is done. In a functioning (depolarizing) cell then, the active metabolism is the rate of metabolism above the basal metabolism of the arrested heart. Activation metabolism measured by heat typically has two phases: initial heat and recovery heat. Initial heat is the heat generated by the actual depolarization and calcium transient in the heart, and recovery heat is the energy expended in rebuilding energy stores used [25, 26]. This is in contrast to skeletal muscle where recovery heat is usually not seen and is indicative of the different conditions the two muscle types operate under [26].

The use of ³¹P NMR spectroscopy as a tool for investigating cardiac metabolism is particularly attractive since the non-invasive nature of this technique allows experiments to be performed on the complete, working, and in some cases in vivo heart. Wasser et al. have used magnetic resonance spectroscopy to explore the relative levels of PCr, b-ATP, and P_i and pH in the isolated, perfused heart of Chrysemys picta bellii during ischemia and subsequent reperfusion [32] and during graded extracellular lactic acidosis [33]. Using magnetization transfer techniques, Bittl et al. have used ³¹ P NMR spectroscopy in order to characterize the reaction kinetics of the conversion of PCr to ATP in a variety of tissues including, brain, heart, and skeletal muscle of the rat [34]. A study of the effects of local ischemia on the PCr/ATP ratio in the right ventricle of the pig heart was performed by Schwartz et al. using ³¹P NMR spectroscopy and a specially designed butterfly coil intended to localize spectral data to the ventricular wall [35]. Phosphate metabolism of rat hearts during low flow ischemia was also explored by Camacho et al. who correlated their results to separate studies in which intracellular calcium was measured fluorescently [16]. A review of NMR techniques for assessing cardiac function may also be found in [3].

2.6.3 Ion Concentration

Precise control of ion concentrations in the cardiac cell is important for maintenance of membrane potential, excitation contraction coupling, the function and regulation of cellular enzymes, and as second messengers for many control mechanisms. Ion concentrations can be measured in a number of ways. Ions which are NMR sensitive may be measured by quantitative spectroscopy. These include ¹H, ²³ Na, and ⁴³Ca. In the in vivo case, however, these ions are often present in such low concentrations that good measurements are hard to make with NMR. Additionally, NMR is not, in general sensitive to whether the ion is either inside our outside of the cell, and interference of proton signal from water is a problem with ¹H spectroscopy. A more common way to measure ion concentrations (for both NMR and non NMR sensitive nuclei) is via a specific indicator. An indicator is a compound which bonds reversibly and weakly with the ion in solution, and whose molecular properties differ for the bound and unbound case. Typically indicators are large organic molecules with specific responses to a stimulus. These responses are altered in the ion-bound and unbound states and thus differences in the response of the indicator can be used to determine the ratio of bound to unbound indicator. This ratio can in turn be used to deduce the concentration of the particular ion. Some examples of ion concentration indicators include indo-1 which changes its fluorescence emission wavelength when bound to Ca²⁺, fura-2 whose excitation wavelength is altered upon binding of Ca²⁺ , and TF-BAPTA which exhibits a shift in one of its ¹⁹F NMR peaks with increased Ca²⁺ binding [36].

Examples of in vivo measurement of cardiac ion regulation are numerous. London et al. discuss the synthesis and performance of both TF-BAPTA (a ¹⁹F NMR sensitive indicator) and a high dissociation constant (K_D) analogue of the fluorescent calcium indicator fura-2 for the measurement of intracellular calcium levels [36]. A brief overview of other calcium studies includes work by Backx et al. who used microinjection of fura-2 salt into rat cardiac trabeculae [37], Hirano et al. who used fura-2 loaded guinea pig ventricular myocytes to investigate modulation of L-type calcium channels [38], Satoh et al. who utilized SBFI and fluo-3 to investigate the regulation of Na⁺ and Ca²⁺ in guinea ping cardiomyocytes [39], Murphy et al. who have used the ¹⁹F NMR spectroscopy sensitive dye TF-BAPTA to measure regulation of cytosolic free calcium in the perfused rat heart [40], Chen et al. who used fura-2/AM to investigate the effects of electrically gated calcium channels on intracellular calcium concentration in an isolated, perfused rabbit heart [41], Ikenouchi et al. who have used the fluorescent dye indo-1 to investigate the effects of angiotensin II on intracellular calcium [42], Sollot et al. who investigated a novel technique for loading the free acid form of indo-1 into isolated cardiomyocytes [43], Laskey et al. who used fluorescence imaging microscopy and fura-2/AM to investigate intracellular calcium regulation in the endothelium of rabbit cardiac valves [44], and Kargacin and Kargacin who explored cardiac sarcoplasmic reticulum pump kinetics using fura-2/AM [45]. Field et al. utilized simultaneous ³¹P NMR and fura-2/AM fluorescence to measure intracellular pH and calcium concentration respectively in an ischemic, isolated rat heart [12]. Also, as mentioned above, Camacho et al. used the fluorescent dye indo-1 in an attempt to correlate intracellular calcium regulation to NMR measured phosphate metabolism during ischemia in an isolated rat heart [16]. Finally, Wasser et al. have demonstrated measurement of intracellular calcium concentration in the isolated cardiomyocytes of Chrysemys picta bellii during prolonged anoxia and acidosis [18].

2.6.4 Mechanical Function

The mechanical work produced by the heart can be measured in a number of ways. In vitro measurements can be made on the isolated heart by artificially perfusing the heart in the Langendorff mode whereby the right atrium is fitted to a cannula through which perfusate is pumped by external means which allows cardiac output to be controlled [12, 25, 26, 27]. The heart is typically supported by this tube in a larger perfusate filled tube in which the height of the fluid can be used to control the mean working pressure of the heart. By introducing a balloon catheter and pressure transducer into the ventricle, cardiac performance can be monitored by measuring isovolumetric pressure development [12, 26]. Alternatively, the heart may be fitted with both input and output cannulas and remaining vessels ligated as has been done by Wasser et al. [21]. This allows for closed-loop circulatory measures to be made on a working heart including flow and via a pressure transducer, developed ventricular pressure.

A procedure for assessing mechanical performance of the heart in vivo has been reported by Farrell et al. using the species turtle Chrysemys scripta [46]. Briefly, they cannulated the right anterior vena cava, ligated other vessels on the input side, cannulated four different output vessels, and ligated the others. With pressure transducers at the tips of the cannulas, atrial and ventricular pressures as well as volumetric flow rates could be monitored in an in situ working heart under a variety of temperature and oxygenation conditions [46].