Printer Friendly

Reliable, responsive pacemaking and pattern generation with minimal cell numbers: the crustacean cardiac ganglion.

A Physiologically Responsive, Reliable Heart Pacemaker and Muscle Activator Built From Nine Neurons

The crustacean cardiac ganglion (CG) is composed of from 6 to 16 neurons, 9 in most decapods, that autonomously provide rhythmically recurring barrages of action potentials to activate the heart muscle. In Malacostraca, the heart is neurogenic and in adults dependent for its beating on the impulses from the ganglion. The CG, consisting of the neurons and their processes, wrapped in glial and connective tissue, forms an elongated, discrete branching trunk in or on the heart. It can be dissected from the heart and will continue to show spontaneous, rhythmical bursting. As an accessible and robust in vitro preparation, the CG joins a list of crustacean preparations that have provided insights into fundamental neurophysiological mechanisms, in this case the mechanisms by which small neuronal networks can generate rhythmical, patterned output (reviews: Wiens, 1982; Marder and Calabrese, 1996). Possibly the most important contribution was the demonstration that individual neurons are endowed with an intrinsic burst -organizing mechanism that results in a patterned output to any appropriate excitatory drive, and the detailed analysis of the ionic mechanisms involved. Further, the CG demonstrated that interconnections among a small number of neurons with such a capability can ensure coordinated, patterned, rhythmic, highly fault-tolerant output from the ensemble. Patterned or bursting impulses are, of course, the essential effective activator of responses of other neurons or muscles or secretory cells. The recognition of patterning mechanisms intrinsic to individual neurons has simplified the analysis of neoronal network pattern generation, freeing it from seeking reliance on properties emergent from a network.

This review seeks to provide a reader with an overview of the studies on the CG in the context of crustacean heart function, with emphasis on the electrophysiological studies of the isolated CG. The ionic mechanisms observed have proven broadly applicable to explaining intrinsic burst generation or pattern generation by neurons in both invertebrate and vertebrate nervous systems (see further below). As early investigators noted (Welsh and Maynard, 1951), the CG has the essential properties of a brain: apparently autonomous spontaneity and the ability to sense relevant environmental changes (degree of heart filling, hormones) and integrate them with intrinsic pattern-generating properties to provide an appropriately adjusted motor output.

The unique suitability of the CG for analyzing cellular mechanisms for pattern generation and rhythmicity derives from particularities of crustacean neuronal functional anatomy, specifically the segregation of the major impulse-generating mechanism (voltage-dependent [Na.sup.+] conductance) to axons so that the soma and initial axon segment are not actively invaded by impulses ("spikes"). In the more elongate CG of lobsters, the well-separated distribution of the neuronal somata makes possible the physical separation of the impulse-generating axon or axons from the more subtle electrical responses of the non-impulse-generating soma, initial axon segments, and associated collaterals. One of these is a graded, regenerative, [Ca.sup.2+]-mediated response to depolarization, lasting 200 ms or longer, which provides drive for generation of the burst of impulses by the axon and hence is referred to as a driver potential (DP). As reviewed below, the characteristics of DPs, which, in isolated CG, are initiated in resp onse to spontaneously occurring pacemaker depolarization or to synaptic excitation, provide a basis for interpreting many of the responses of intact hearts as well as the isolated CG to both physiological and experimentally imposed modulatory influences (see also Cooke, 1988, 2002).


Morphological observations

Alexandrowicz (1932) provided an anatomical description of the innervation of decapod hearts that was based on vital methylene blue staining; this description has provided a structural underpinning for many subsequent observations of function (Fig. 1). Similar studies of the heart of stomatopods (1934) and of an isopod (1952) followed (see also Suzuki, 1934). Alexandrowicz described three neuronal elements innervating decapod hearts: the intrinsic neurons (cardiac ganglion); the extrinsic fibers "" one inhibitory and two acceleratory axons-arriving "" three segmental nerves on each side from thoracic ga"" (combined into the dorsal nerves in decapods) to rea"" the heart and CG; and nerves innervating the suspensory ""nents, ostia, and arterial valves (Fig. 1B, C, D). He called attention to the neurogenic nature of the crustacean ""s, reporting the immediate cessation of contractions "" nerves between the ganglion and muscle were cut. He "" observed that the most anterior extrinsic nerve pair in ""atopods was i nhibitory and the sequentially more pos"" two pairs were excitatory, and correctly surmised tha"" these have similar roles in decapods (Wiersma and "" 1942, crayfish; Smith, 1947, crabs; Maynard, 1953, "". Alexandrowicz provided important additional observations relevant to regulation of crustacean hearts: the ""ical description of the pericardial organs (POs) of cra"" their macruran homolog, the ligamental plexuses. T"" are neurohemal structures that release neurohormones "" cardioregulator and other functions into hemolymph ""ing to the heart (Alexandrowicz, 1953; Alexandrowicz and Carlisle, 1953).

Comparison of the circulatory systems cross the orders of Crustacea supports a consensus the evolution from a pulsatile dorsal vessel toward a compa"" heart accompanies more active lifestyles to provide for "" efficient circulation (Wilkens, 1999; for a comprehensive review, see Maynard, 1960). In the primitive bran""ods (e.g. Daphnia sp., Triops longicaudatus), no neu"" have been found in or on the heart, and the heartbeat is thus myogenic (Yamagishi et al., 1997). In the CC"" the stomatopod Squilla (Alexandrowicz, 1934), 14 or 15 neurons are distributed along a ganglionic trunk that "" on the external dorsal surface of the heart and can be "" long as 8 cm. In decapods, the CG lies on the inner "" wall of the heart (Fig. 1A, C). Although crayfish have 16 ""nsic neurons, in most decapods examined the number has been reduced to 9. There is a clear distinction in size and ""tion. The most posterior four neurons are smaller and having axonal terminations within neuropil in the gangli"" trunk, are therefore inter neurons. The more anterior fi"" neurons are larger and, because they provide axons that leave the ganglion to innervate heart muscle fibers, are mo""urons (Fig. 1B, E). Each of the neurons has dendritic "" collateral processes extending out of the ganglionic trunk to ""ify onto nearby muscle fibers that are responsive to stretch. In lobsters the somata are widely spaced along a linear "" e.g., Panulirus, Fig. 1C) or Y-shaped (e.g., Homarus, Fig. 1A) ganglionic trunk spanning nearly a centimeter in a 0.5-kg animal; in crabs of edible size the neurons are usually compacted into anterior and posterior clusters separated by several millimeters of ganglionic trunk (Figs. 1E, 8D).

Electrophysiological anatomy

One of the first conclusions to come from electrophysiological recording from lobster CG (Welsh and Maynard, 1951; Maynard, 1955; see also Matsui, 1955) was that not only the rhythmicity but also the pattern of impulses within the bursts of activity is extraordinarily stable (Figs. 2C, E; 6A, B; 9E, F). In Homarus americanus, for example, output of isolated CG consists of a 200- to 300-ms burst of tightly grouped efferent impulses that recur spontaneously at rates similar to observed heartbeat rates (50-60/min for lobsters). Further confirmation of the consistency of burst patterning came with analysis of the patterning by Hartline (1967), who used an array of five or more pairs of extracellular electrodes placed along the trunk and major nerves of the Homarus ganglion to identify each impulse with its axon by mapping the site of impulse initiation and its conduction route (Fig. 2A, B, D). It was clear that a particular one of the posterior cells (usually 8 or 7, numbering cells from anterior to posterior) co nsistently fired first in a burst, and that the firing of large cells commenced with the arrival of the first small-cell impulse, propagated along its anterior-traveling axon, at neuropil in which synapses on large-cell collateral processes occur. Thus the most posterior large cell (5) fired first, followed successively by more-anterior cells. The sites of impulse initiation observed in such studies confirmed conclusions reached from intracellular recording from the large cells (early work reviewed by Hagiwara, 1961) that impulses do not invade the somata, but rather are initiated at a site that can be more than a millimeter distant along the axon. Cells having more than one axon--for example, Cell 3 situated at the junction of the Y in Homarus--initiate impulses in each axon independently (Fig. 2D). As mentioned above, within each burst, the patterning of impulses of each axon remains highly constant, each unit showing repetitive firing (Fig. 2C). In Homarus, the first 3 to 4 impulses of large-cell axons occ ur at high frequency (90-120/s) and then continue for 3 or more at a slower rate (10-20/s, Hartline, 1967); small-cell axons fire as many as 15 impulses starting at rates of -80/s and declining during the burst toward 20-30/s. A similar analysis showing minor differences in detail from Homarus is available for Panulirus interruptus (Friesen, 1975a, b).

Studies of functional anatomy in crab CG (Tazaki and Cooke, 1979a, 1983a, in Portunus sanguinolentus; Fort and Miller, 2001, in Callinectes sapidus) show an important variation on this organization, namely that large-cell axons, rather than each firing in a consistent individual pattern within each burst, show synchronization of their impulse firing (Fig. 6A). Synchronization also occurs among the rostral neurons of the Squilla ganglion (Watanabe and Takeda, 1963).

Axons of the posterior four small cells in the lobster and crab CG remain within the ganglionic trunk and provide excitatory, chemically mediated synaptic input to the large cells. This input initiates burst activity of the large cells, and thus the small cells are considered pacemaker interneurons or premotor neurons. The large-cell axons, while providing synaptic input to each other and perhaps also to the small cells, produce the bursts of motor impulses responsible for heart muscle contraction. Synchronization of ganglionic activity is not only mediated by synaptic drive, but the general excitability of the network is shared among all of the neurons by means of electrotonic coupling capable of passing slow changes in potential. Synaptic and electrotonic interactions are discussed below.

Two further properties of the individual CG neurons are also critical: each has stretch-sensitive dendrites ramifying into heart muscle that account for the ability of intact hearts to adjust heart rate and strength of beating to the degree of filling of the heart; and each has the intrinsic ability to produce a patterned burst of impulses in response to a simple stimulus.

An Intrinsic Burst-Forming Mechanism: the Driver Potential

Possibly first discussed as an intrinsic potential by Watanabe (1958) in lobster CG, and further described in studies of the Squilla (stomatopod) CG (Watanabe et al., 1967a, b), driver potentials are relatively slow, sustained, regenerative depolarizations that may arise from a gradual pacemaker potential or be evoked by a depolarization, such as an excitatory synaptic potential (or an applied depolarizing stimulus) (Fig. 3). They provide the depolarizing drive for initiating repetitive impulses at an axonal "trigger zone." Because DPs arise in the soma and proximal axon or axons -- regions lacking impulse-generating conductances and, particularly in lobster CG, physically well separated from spiking axon -- they can be studied in relative isolation. Their properties account for much of the collective behavior of the network, including rhythmicity, reciprocity between burst rate and duration, and phase resetting in response to imposed extra stimuli.

Driver potentials arise in the non-spiking soma and proximal axon

Direct evidence for the localization of DPs is provided by simultaneous intracellular recording from a neuronal soma and its axon at a distance of several millimeters (Fig. 3A): the soma recording shows a sustained, slow depolarization (i.e., the DP) with attenuated sharp deflections that are synchronous with the overshooting impulses arising from a flat baseline recorded from the axon (Watanabe et al., 1967b, Squilla oratorio; Tazaki, 1970, Eriocheir japonicus; Tazaki, 1973, Panulirus japonicus; Tazaki and Cooke, 1983a, Portunus sanguinolentus). The localization of DPs to the soma and non-impulse-supporting initial axon was also shown in Homarus by intracellular recording during or after ligaturing at distances between 200 [micro]m and more than a millimeter from the soma (Tazaki and Cooke, 1983b) (Fig. 3B). For more distant ligatures, electrotonically decremented impulses, as recorded from the soma, were superimposed on the DP, indicating that the ligatured segment included an axonal trigger zone. It is wor th noting that more complex deflections, suggestive of synaptic potentials, were sometimes present in recordings from ganglion segments that include a single soma. This suggests the possibility that the processes of other neurons present in such a segment can contribute synaptic input. Any rapid deflections disappeared with only minimal change in the form of the underlying DP when tetrodotoxin (TTX) was added to the perfusing saline (Tazaki, 197la). Treatment of ganglia with TTX made it possible to observe DPs in the absence of any impulsemediated activity by simultaneous intracellular recording in up to three large cells (Tazaki and Cooke, 1979b, in Portunus) (Fig. 3D, two cells). Stimulation with a depolarizing current pulse in any one cell simultaneously initiated DPs in all; the amplitudes and form differed slightly, but remained characteristic in each cell, hence indicating that the DP represented an active response of each neuron, but one that was brought to threshold by the spread of the depolarizing s timulus via pervasive electrotonic coupling among all neurons of the ganglion.

The form of their driver potential shapes the pattern of impulses of each neuron

Large cells. The DPs as recorded intracellularly from large cells of crabs or lobsters show little difference in form whether examined in TTX or after isolation of a soma by ligaturing. If the neuron is relatively undamaged (as evidenced by high input resistance and a resting potential ~-50 mV), depolarization to a threshold (~-45 mV) initiates a regenerative response requiring 10 or more ms to reach a maximum ~20 mV depolarized from resting potential (Fig. 3D). The depolarization shows a rounded peak with a gradually declining shoulder, followed by a more rapid repolarization that gives way to hyperpolarizing after-potentials. These have a relatively rapidly decaying phase lasting up to 1 s, followed by a slowly decaying phase lasting tens of seconds. Although they are regenerative, DPs are not all-or-none responses; rather their size is related to the amplitude and rate of rise of the depolarizing stimulus. More importantly, threshold and amplitude are related to the rate of repetition: given a constant sti mulus, threshold becomes lower and amplitude and duration become larger with increasing time since the previous response. A maximal response requires a pause of more than 10 s. It will be obvious that at typical heart rates of 50-60/min, DPs are not at maximal amplitude but are evoked in a range over which changes in heart rate will result in decreased or increased amplitudes (Fig. 5A). Possible cellular mechanisms governing these relations are discussed below.

Small cells. An intracellular recording from a small cell, simultaneously with a large cell, in a Portunus CG treated with TTX (Tazaki and Cooke, 1983c) reveals clear differences in the form of DPs of large and small cells (Fig. 3C). The DP in the small cell is lower in amplitude but has a long (~400 ms), sustained, slowly declining plateau. The initiation of the small-cell DP by depolarizing current also causes a DP in the anterior large cell; this has the larger but less sustained form described above. The electrotonically spread continuing plateau of the small-cell DP is apparent in the large-cell recording. As mentioned previously, the bursts of the small cells are longer and show a well-sustained frequency of firing. Thus, the differing form of the DPs of small and large cells accounts for the differing pattern of impulses produced during bursts by the axons of small and large cells (Fig. 6A).

The ability of slow potentials such as DPs to generate trains of impulses from the axonal trigger zone implies that the ionic mechanisms involved in impulse initiation are not subject to rapid inactivation by depolarization. Intracellular recordings from axons, as mentioned above, show typical overshooting, all-or-none impulses with a rapid rise and fall. When intracellular penetrations were attempted, damage to the target neuron was often signaled by appearance in an extracellular recording of repetitive firing not organized into the coordinated bursting of the remaining cells of the ganglion, confirming the ability of axons to respond to sustained depolarization with minimal adaptation.

Voltage-Clamp Analyses of Ionic Currents Giving Rise to Driver Potentials

Characterization of four kinds of current in ligatured somata

The characteristics and ionic conductances responsible for DPs have been examined in most detail in the Homarus CG (Tazaki and Cooke, 1983c, 1986, 1990), in which the separation of the neurons permitted the isolation of ganglion segments with a single large-cell soma by ligaturing the ganglionic trunk. Studies with two-electrode voltage clamping show that DP characteristics involve the interplay between an inward [Ca.sup.2+]-mediated current ([I.sub.Ca]) and three outward [K.sup.+]-mediated currents: a transient current ([I.sub.A]), a slowly-inactivating [K.sup.+] current ([I.sub.K]), and a Ca-dependent [K.sup.+] current ([I.sub.KCa]) (Tazaki and Cooke, 1986, 1990). Space-clamp could be ensured in the Homarus ganglion by ligaturing. Without detailing the evidence, a brief summary of the conclusions from voltage-clamp analyses follows (see also Figs. 4 and 5 and legends). The depolarizing current that generates DPs arises from voltage-gated increased conductance to [Ca.sup.2+] and the resulting inward [Ca.sup. 2+] current ([I.sub.Ca]) (Figs. 4A, 5B). The amplitude of the DP is determined by the extent of [Ca.sup.2+] channels available for activation. An examination of the effect of the holding potential ([V.sub.h]) of the large-cell on the amplitude of [I.sub.Ca] shows a maximum for a potential (-60 mV) close to the most hyperpolarized value observed during the afterpotential following a burst. Peak current is reduced to half or less at more hyperpolarized [V.sub.h]. The amplitude of the DP is normally limited by the nearly simultaneous development, in response to depolarization, of the [I.sub.k] conductance. If [I.sub.k] is blocked with tetraethyl ammonium (TEA), [I.sub.Ca] can produce overshooting potentials (Fig. 4A). Repolarization, and thus the duration of DPs, is the result of the interplay of [I.sub.k]' inactivation of [] primarily from intracellular accumulation of [Ca.sup.2+] (Fig. 5B, D), and development of the [Ca.sup.2+] -dependent [I.sub.kCa].

The voltage-clamping data are consistent with the major role of [Ca.sup.2+] in mediating the DP depolarization (Figs. 4A, 5). However, a possible role of [Na.sup.+] in contributing to inward current remains unclear, with conflicting observations of the effects of its reduction or removal from the saline (Tazaki and Cooke, 1979c; Berlind, 1985, 1993). The differences may be attributable to the manner of substitution for [Na.sup.+] and to other differences in the experimental conditions.

The more rapidly decaying phase of the hyperpolarizing afterpotential following a CG driver potential (Fig. 3D) represents the deactivation of [I.sup.K] with repolarization, while the slowly declining phase of the afterpotential probably reflects the sequestering Or extrusion of [Ca.sup.2+] to reduce [I.sub.Kca]. A refractory period for initiation of DPs is imposed by a combination of the inactivation of [I.sub.Ca] by intracellular accumulation of [Ca.sup.2+] as well as the increased [K.sup.+] conductance from the [Ca.sup.2+] -activation of [I.sub.KCa]. When [Na.sup.+]-mediated impulses are present (in non-TTX-treated preparations), a hyperpolarizing current from activation of electrogenic ion transport also contributes to the afterhyperpolarization (Livengood and Kusano, 1972; Livengood, 1983). With time, the threshold for response to a depolarizing stimulus decreases (1) due to progressive reduction of the ability to activate transient [K.sup.+] conductance ([I.sup.A]) with decline of the afterhyperpolariza tion or pacemaker depolarization (or both); and (2) with progressing [Ca.sup.2+] extrusion or sequestration, reduction of the [I.sub.Kca] conductance and of the [Ca.sup.2+]- mediated inactivation of [I.sub.Ca] due to previous activity (Fig.5B).

Driver potentials and similar intrinsic patterning potentials have proven to be a more labile response than generation of action potentials. This is easily explained by the relatively smaller current densities involved and the closely balanced interplay of conductances that govern their form. In crustaceans, for example, [I.sub.Ca] observed under voltage-clamping conditions can be inactivated by raised intracellular [Ca.sup.2+]. Thus it is not surprising that crustacean DP generation fails if neural damage causes [Ca.sup.2+] entry from the saline, decreases input resistance, depolarizes the cell membrane (thereby increasing [I.sub.K]), and increases leakage conductance. That DPs represent a delicate balance in the interplay of inward and outward currents is confirmed by observing the spontaneous initiation of DPs in previously quiescent preparations and the augmentation of their amplitude and duration after addition of TEA to inhibit [I.sub.K] (Tazaki and Cooke, 1979c; Berlind, 1993) (see below for further di scussion of spontaneity).

Changes of DP amplitude and duration with repetition rate are determined by characteristics of [I.sub.Ca]

Analysis of the pharmacologically isolated [Ca.sup.2+] current in ligatured somata suggests that its characteristics dominate the determination of DP amplitude and duration during repetitive activation. Figure 5 combines observations (from Tazaki and Cooke, 1990) on DP amplitude vs. DP repetition rate (A), and rates of recovery of [I.sub.Ca] for the second of two stimuli (B). These follow closely parallel time courses; the slightly faster recovery of [I.sub.Ca] studied with a pair of pulses is explained by a cumulative inactivation of [I.sub.Ca] that occurs with repetitive activation. The role of intracellular [Ca.sup.2+] accumulation as the main agent of inactivation is evidenced by the lack of inward current inactivation when [Sr.sup.2+] substitutes for [Ca.sup.2+] or the soma is injected with EGTA (Fig.5B).

The accumulation of intracellular [Ca.sup.2+] also activates [I.sub.Kca]. This current is of small magnitude relative to [I.sub.Ca] and [I.sub.K], and it influences primarily the rate of repolarization after a DP, or of a pacemaker depolarization during the equivalent of interburst intervals (Tazaki and Cooke, 1986).

Rhythmicity of Cardiac Ganglion Electrical Activity

The importance for survival of the animal of a reliable, rhythmic heartbeat cannot be exaggerated. The isolated cardiac ganglion, as the pacemaker and activator of heart contraction, has proven capable of sustaining robust, rhythmical motomeuron bursting when challenged with a variety of insults and perturbations.

Behavior of the cardiac ganglion as an oscillator

A number of studies demonstrate that small sustained currents passed into a large CG soma can alter the coordinated burst rate of the entire ganglion, while brief pulses of hyperpolarizing or depolarizing current can reset the bursting phase (Fig. 6A) (Watanabe and Bullock, 1960; Watanabe et al., 1967b, in Squilla; Tazaki, 1972, in Eriocheir; Mayeri, 1973a, b, in Homarus; Matsui et al., 1977, in Panulirus; Tazaki and Cooke, 1979a, and Benson, 1980, in Portunus). The coordinated response of the entire ganglion to such perturbation of a single neuron is ensured by the combination of electrotonic and probably reciprocal excitatory synaptic interactions among all the cells. The characteristics of [I.sub.Ca] that determine the threshold and amplitude were discussed above and help to explain the reciprocity observed between burst rate (or interburst interval) and duration.

Autonomous rhythmicity: stretch responsiveness and pacemaker potentials

The basis of the spontaneity exhibited by isolated CG preparations is considered in this section. A question arises from the observation that an intact heart, if not still suspended by its elastic ligaments or stretched by internal perfusion, rapidly becomes quiescent. In situ, expansion and filling of the heart, mediated by the suspensory ligaments, probably aided under some conditions by contraction of alary muscles, stimulates heart contraction. The response to stretch or filling is undoubtedly mediated by the dendritic and collateral processes of the CG cells that ramify on heart muscle near the ganglion. Alexandrowicz (1932) pointed out that the terminations of these processes differed from the more peripheral neuromuscular junctions and noted their responsiveness to stretch. For an isolated heart, the plots of heart rate vs. perfusion pressure and the extent of contraction vs. perfusion pressure can be superimposed, and they increase along a hyperbolic curve (Maynard, 1960; see also Kuramoto and Ebara, 1984a, 1985). A possible role of hypoxia in governing heart rate has been proposed (Wilkens, 1993). Although it is likely that stretch depolarizes the neuronal processes, as is well documented for crustacean stretch receptors (e.g., Eyzaguirre and Kuffler, 1955), direct recordings of CG neuron responses to stretch of heart muscle appear not to have been obtained. In decapod hearts opened ventrally to expose the CG (Fig. 1A), contractions of the quiescent heart muscle can be elicited by probing sites in which CG dendrites are embedded (Alexandrowicz, 1932; Cooke, 1962; Hartline, 1967). Electrophysiological monitoring of the CG confirmed the absence of activity in the quiescent Homarus heart (Hartline, 1967).

The CG, after removal from the heart, exhibits the spontaneous, continuous, rhythmically recurring bursts of impulse activity described above. Intracellular recordings from the small, posterior neurons have consistently shown a slowly depolarizing pacemaker potential that develops after the post-burst hyperpolarization and leads to the initiation of the next burst (Fig. 6A) (Tameyasu, 1976; Tazaki and Cooke, 1979a, 1983c). In Homarus, a pacemaker potential is not usually seen in undamaged large cells, the DP and burst being initiated in them by synaptic driving mediated by impulses of the small-cell axons.

The question raised by the contrast in behavior of unstretched hearts and isolated ganglia is whether stretchinduced depolarization of dendrites and collateral processes normally induces bursting activity of the ganglion, while in isolated ganglia, the pacemaker potentials that functionally replace this stretch response in fact represent an injury current. It is possible that differences in the prominence of pacemaker potentials in recordings from large cells, even in the same species on the same equipment in the same laboratory (e.g., Tazaki and Cooke, 1979a; Benson, 1980; Berlind, 1982), represent differences in techniques of ganglion isolation or of intracellular electrode penetration. During the hyperpolarization following bursts and the interburst interval, the gradual reduction in conductance to [K.sup.+] or in hyperpolarizing electrogenic active transport (Livengood and Kusano, 1972; Livengood, 1983) can lead to net depolarization as the balance of currents unmasks an outward injury or "leak" current m ediated by nonspecific ionic conductances. The greater prominence of pacemaker potentials in small cells, and thus their role in initiating bursting, may be related merely to their smaller size. Observations thus far available provide no evidence for the participation of a hyperpolarization-activated cationic current, such as [I.sup.h] associated with pacemaker depolarization in several vertebrate pattern-generating neurons (reviews: Kaupp and Seifert, 2001; Santaro and Tibbs, 1999). Such a conductance would be expected to cause "sag" in the voltage responses to hyperpolarizing current. The opposite--increasing hyperpolarization indicating a decrease in conductance--occurs, as seen in Figure 4A.

Experiments in which segments of the ganglion that include single large cells are isolated have shown that such cells "spontaneously" produce rhythmical DPs (with superimposed impulse trains, if a trigger zone remains) when a pacemaker (injury?) current provides depolarizing drive (e.g., Connor, 1969; Berlind, 1982; Tazaki and Cooke, 1983a, c). Cell 3 of Homarus has more often shown recurrent DPs after ligaturing than have Cells 1 or 2 (Fig. 6B), perhaps because it has three axons and requires three ligatures rather than one, which results in greater injury current. In cells that do not show a pacemaker (or injury) current, rhythmical activity appears if a sustained depolarizing current is passed into the neuron, or, often, if TEA is added to reduce residual [K.sup.+] current (Berlind, 1982). Recently, rhythmic "bursting" has been recorded from large cells isolated into primary culture from a crab (Carcinus maenas) CG (Saver et al., 1999) (Fig. 6C). The observations on reduced preparations demonstrate that ea ch of the large cells, at least, is intrinsically capable of generating rhythmic, patterned bursts given a nonspecific excitatory drive (e.g., depolarization by injury current).

Synaptic Interactions

Excitatory chemically-mediated synapses

Studies such as those described earlier (Electrophysiological anatomy) confirmed that impulses of the posterior small-cell axons, which do not exit the ganglion, initiate activity of the large motor neurons and can thus be considered to have a pacemaker function (see also Mayeri, 1973a). Impulses of specific small-cell axons can be correlated with excitatory postsynaptic potentials (EPSPs) recorded intracellularly from large-cell somata (Fig. 6A) (Hagiwara and Bullock, 1957; Hartline and Cooke, 1969; Connor, 1969; Tazaki, 1971b; Friesen, 1975a, b, c; Tameyasu, 1987). In spiny lobsters, the most anterior of the small cells (Cell 6) provides a large initial EPSP, followed by one or more of reduced amplitude (antifacilitation); after a pause equivalent to the interburst interval, the next EPSP is augmented (Friesen, 1975c; Tameyasu, 1987). Synaptic interaction between large cells has been described in Homarus (Hartline, 1979). The observation that the bursts from small cells increase in rate and shorten when lar ge-cell impulses (but not DPs) are eliminated by TTX applied anteriorly demonstrates an influence of large-cell activity on the small cells (Berlind, 1989). Synaptic potentials have not been observable in the intracellular recordings available from small cells. It may be that synapses occur at too great an electrotonic distance from the soma to be recorded. The possibility that non-impulse-mediated transmission occurs, as shown in the stomatogastric ganglion (Graubard et al., 1983), has not been explored. Impulse-mediated EPSPs have been recorded in the large CG cells of several crab species (review: Hagiwara, 1961; Tazaki, 1967; Tazaki and Cooke, 1979a, 1983c; Berlind, 1982).

In the CG, as in crustacean ganglia generally, synaptic interactions occur in complex neuropils formed by collateral processes and branches from axons (Alexandrowicz, 1932, 1934, 1952; Obsawn, 1972; Aizu, 1975; Hawkins and Howse, 1978; Mirolli et at., 1987; Morganelli and Sherman, 1987). By injecting procion rubine and horseradish peroxidase (markers that can be distinguished by electron microscopy) into two of the large cells, Mirolli et at. (1987) demonstrated that synapses occur in all combinations among small and large cells in neuropil of the crab (Portunus sanguinolentus) ganglion (Fig. 8A). Electron microscopy of neuropil in Homarus showed symmetrical synapses (Morganelli and Sherman, 1987). In both studies, all synapses among CG cells had clear-cored vesicles, similar to those reported by others at excitatory neuromuscular synapses (Atwood, 1976). Synaptic contacts are also contributed within the neuropils by the inhibitory and excitatory extrinsic regulator fibers. These have morphologically differen t vesicles and, in turn, differ in morphology from those of the intrinsic cell synapses.

The identity of the synaptic transmitter or transmitters among the CG neurons remains under discussion. The small-cell synaptic effect on large cells is rapid and therefore must be mediated by a fast, ionotropic transmitter-receptor interaction. Current evidence now strongly supports glutamate as the intraganglionic transmitter. Applied glutamate depolarizes large CG neurons of Homarus by increasing conductance to monovalent cations (Cooke, 1966). Under voltage clamp (but not space clamp), glutamate increased conductance and produced an inward current that had a reversal potential of ~- 15 mV and was dependent nearly equally on [K.sup.+] and [Na.sup.+]. Quisqualate was an even more potent agonist than glutamate, while L-aspartate was as potent as glutamate (Hashemzadeh-Gargari and Freschi, 1992).

Immunolabeling for glutamate-like reactivity in Panulirus argus (Fig. 7A, B) has shown a compound presumed to be glutamate in each of the CG cells and their axons. It has also revealed extensive terminations of the small cells in neuropil and in networks surrounding the large-cell somata and proximal neurites (Delgado et at., 2001). Similar termination of labeled processes in the posterior of the ganglion near small cells is not reported. The demonstration of glutamate-like immunoreactivity in the axons and their terminations on the heart muscle supports the electrophysiological evidence for glutamate as a transmitter at the neuromuscular junctions (Benson, 1981). Additional evidence is the detection, by high-pressure liquid chromatography, of glutamate in extracted CG of the isopod Bathynomus doederleini and, in lower amounts, in heart muscle (Yazawa et at., 1998). Modulation of CG activity by the extrinsic fibers and neurohormones is further discussed below.

Electrotonic coupling

An important element ensuring the coordinated bursting of the motor axons from the CG is the presence of electrotonic coupling among all of the neurons (Watanabe, 1958; Hagiwara et at., 1959; Watanabe and Takeda, 1963; for small cells in crab CG, see Tazaki and Cooke, 1979a). The coupling passes slowly changing potentials effectively, but not rapidly changing ones such as impulses (the case of synchronization of large-cell impulse firing in the crabs Portunus and Callinectes was mentioned above). The coupling results in the effective spread of both pacemaker and driver potentials (or of imposed current) among the nine neurons, helping to ensure coordination of their bursting activity (Figs. 3C, D; 6A). Although electrophysiological recording demonstrates effective electrotonic coupling between large and small cells, dye coupling has not been demonstrated. Lucifer yellow injection into any of the large cells in Portunus resulted in the appearance of dye in all of the other large cells, but not in any of the sm all cells (Fig. 8D) (Tazaki and Cooke, 1983a). Similarly, injection of neurobiotin into any of the large cells of the Panulirus CG resulted in its appearance in all large cells, but not in any small cells (Delgado et al., 2001). A search for images suggesting gap or tight junctions in neuropil has failed to detect them (e.g., Mirolli et at., 1987, in Portunus; Morganelli and Sherman, 1987, Homarus). Areas of membrane close apposition are observed among fine processes in neuropil and may serve as electrotonic junctions (Fig. 8A). The location of electrotonic connections on the small collateral processes would account for the low-pass filtering of electrotonic transmission observed electrophysiologically. These studies found that axo-axonic close appositions occur between the small-cell axons (Fig. 8B, C). Such appositions have also been described in CO of Panulirus (Ohsawa, 1972) and Squilla (Irisawa and Hama, 1965; Watanabe et at., 1967a). In all of these species, the axo-axonic close appositions occur only b etween axons of small cells or pacemaker neurons.

Mechanisms for Modulation of Heart Function in Response to Physiological Demands

Crustacean heart muscle is similar to other crustacean muscle in being striated and innervated by multiple distributed boutons of the motor axons of most or all of the five CO motorneurons (Fig. 7B) (Anderson and Smith, 1971; Kuramoto and Kuwasawa, 1980). Heart muscle fibers are electronically coupled, end-to-end, unlike other muscle in the animal (e.g., Anderson and Cooke, 1971). Neural activation of heart muscle shares most of the characteristics of neuromuscular physiology observed in other crustacean muscles: repetitive impulses produce EPSPs that facilitate extensively, and there is spatial as well as temporal summation of responses to impulses arriving from different motorneurons (van der Kloot, 1970; Anderson and Cooke, 1971; Kuramoto and Kuwasawa, 1980; Benson, 1981; Florey and Rathmayer, 1990; Brown, 1964a, b; for a summary and references to recording from crustacean and other arthropod hearts, see table 1 in Anderson and Cooke, 1971). The strength of contraction depends mainly on the degree to which the facilitated and summed EPSPs produce a sustained depolarization of the muscle (Orkand, 1962). Hence, the heart rate and strength of heart contractions are exquisitely sensitive to the rate, intraburst frequency, and also the patterning of impulses of each of the CG motorneurons. The effects of modulators on heart function can be largely explained by their effects on CG output.

Extrinsic fibers from the CNS

Numerous studies of semi-intact preparations have demonstrated the effects of electrically stimulating the extrinsic fibers to the heart (see Maynard, 1961, for references to early studies). Implanted electrodes and heart monitoring in lobsters (Homarus) have shown that the extrinsic nerves modify heart rate appropriately in relation to behavioral demands. For example, exercising on a treadmill accelerated the heart rate; bradycardia accompanied startle responses (Guirguis and Wilkens, 1995). In crayfish and lobsters the inhibitor and the two accelerator extrinsic nerves can be stimulated separately before they combine to enter the heart to reach the CG as the dorsal nerve (Fig. 1B, C). If the regulatory nerves are stimulated together, inhibition predominates.

Responses to stimulation of inhibitory fibers. The effectiveness of stimulation of the inhibitor depends on the rate of stimulation over a range of ~3-40 stimuli per second (review: Maynard, 1961; Cooke, 1962, in Homarus), with rapid, complete arrest for tens of seconds observed for rates of 20/s or higher. Inhibition is generally followed by a rebound acceleration of heart rate.

Stimulation of the extrinsic regulator nerve during intracellular recording from large CG neuronal somata produces postsynaptic potentials following, one for one, the stimuli to the axon, and repetitive stimulation causes the expected slowing of bursting (Fig. 7G). Inhibitory stimulation tends to move the membrane potential toward the most hyperpolarized levels observable during spontaneous bursting (Otani and Bullock, 1959; Shimahara, 1969a; Matsui et al., 1973; Watanabe et al., 1968, in Squilla). As first seen at the neuromuscular junction (Fatt and Katz, 1953), the reversal potential for the inhibitory conductance is close to the prevailing membrane potential and involves primarily an increase in conductance to [Cl.sup.-] (Shimahara, 1969a). As for peripheral inhibition, [gamma]-amino butyric acid (GABA) is a candidate for the inhibitory transmitter. In support of the earlier study (Shimahara, 1969a), voltage-clamping of large cells of Homarus indicated that GABA application increases conductance to [ p.-] (Kerrison and Freschi, 1992).

A study of GABA-like immunoreactivity in Panulirus CG (Delgado et al., 2001) found labeling in a single fiber entering the ganglion in the dorsal nerve from each side. These formed extensive processes in neuropil throughout the ganglion, including a pericellular network around large-cell somata (Fig. 7C, D). Processes also were seen to ramify into the heart together with the large-cell dendrites presumed to be mediating stretch sensitivity. This morphology strikingly matches the description by Alexandrowicz (1932) of the System I fibers that he proposed were inhibitory (Figs. ID, 7F).

In electron microscopy studies (Morganelli and Sherman, 1987, Homarus; Mirolli et al., 1987, Portunus), synapses that were found extensively in neuropils throughout the CG had terminals with vesicles similar to those associated with inhibitory synapses on crustacean muscle (Atwood, 1976). These anatomical observations provide a structural basis for the observation that inhibitory regulatory fibers are more effective than excitatory ones (Maynard, 1961), as well as for the greater prominence of postsynaptic effects of inhibition as observed from CG somata recording.

Responses to stimulation of accelerator fibers. In his 1932 study, Alexandrowicz described a pair of smaller fibers ("System II") accompanying the larger fiber ("System I," now confirmed as inhibitory) in the dorsal nerve (Fig. 1B) bringing extrinsic innervation to the CG. He suggested that these represented the acceleratory inputs. They show sparse terminations in the ganglion and send processes into the cardiac muscle.

Effects of stimulation of the accelerator fibers have been difficult to characterize because the effectiveness of stimulation appears to fatigue rapidly, especially for stimulation of the more posterior of the two fibers. Bouts of stimulation resulted in slowly developing depolarization recorded in large CG neurons of Panulirus (Fig. 7H) (Shimahara, 1969b; see also Terzuolo and Bullock, 1958). Similar responses were seen in Squilla (Watanabe et al., 1969) and in the isopod Ligia exotica (Sakurai and Yamagishi, 1998).

A recent advance toward resolving the nature of the transmitters for the accelerator fibers is the observation of immunoreactivity to an antibody raised against tyrosine hydroxylase in one of the two fibers reaching the CG from the CNS on each side in Callinectes (Fig. 7E) (Fort and Miller, 2001). Tyrosine hydroxylase catalyzes the addition of a hydroxyl to tyrosine to form dopamine. Together with earlier observations of the presence of a catecholamine in CG tissue (Ocorr and Berlind, 1983) and the excitatory effects of dopamine on the CG (see further below), the observations implicate dopamine as the transmitter for one of the accelerator fibers. Which of the two fibers shows the immunoreactivity is still unresolved.

Among many agents that have acceleratory effects on the CG, possibly the strongest candidate transmitter for the second fiber is acetylcholine (Freschi and Livengood, 1989; Sullivan and Miller, 1990; see further below). 5-Hydroxytryptamine (5HT) has been eliminated as a transmitter, at least for the more anterior of the accelerator fibers in Homarus. Response to 5HT could be blocked by previous application of D-lysergic acid diethylamide (LSD) without blocking the effects of nerve stimulation (Cooke, 1966). Responses to stimulation of the more posterior accelerator nerve were weak or absent, and hence no conclusion about inhibition by LSD was possible. Glutamate is unlikely, as no glutamate-like immunoreactivity was detected in the dorsal nerves of Panulirus (Delgado et al., 2001).

Neurohormonal modulation

Neurohormones of the pericardial organs. Alexandrowicz (1953) recognized that the webbings of neural tissue spanning the openings of the branchial sinuses in crabs were neurohemal structures and named them pericardial organs (POs). Given their location in the path of hemolymph returning from the gills and about to enter the heart, functions in modulating heart and circulatory performance were anticipated and soon established (Alexandrowicz and Carlisle, 1953). Homologous structures of lobsters are the ligamental nerve plexuses, now referred to as POs (Alexandrowicz, 1932, 1953; reviews: Cooke and Sullivan, 1982; Chaigneau, 1983). Neurohormones found in POs include the amines 5HT, dopamine (predominant in crabs), and octopamine (predominant in lobsters, Sullivan et al., 1977). Peptides that have been identified include proctolin (Sullivan, 1979), crustacean cardioactive peptide (Stangier et al., 1987; review, Dircksen, 1994) in crabs, and one or more FMRF-amide-related peptides (Trimmer et al., 1987; Mercier e t al., 1993). Most of these are also found elsewhere in the nervous system but have effects on central pattern generators and at neuromuscular junctions at concentrations consistent with their role as circulating neurohormones.

Neurohormonal effects on the heart. It would not be possible here to review the extensive literature from at least 80 years of experiments on the effects of tissue extracts, putative neurotransmitters, hormones, and pharmacological agents on crustacean hearts. More recent studies examining effects of P0 extracts and hormones on nearly intact and in sit hearts suggest that each has subtly differing sites and modes of action in altering heart performance (Florey and Rathmeyer, 1978; Kuramoto and Ebara, 1984a, b, 1988, 1991; Krajniak, 1991; Yazawa and Kuwasawa, 1992; Mercier and Russenes, 1992; Wilkens and Mercier, 1993; Wilkens et al., 1996; Saver and Wilkens, 1998; Saver et al., 1998). Hormones may have differential actions on the valves governing hemolymph distribution to different parts of the animal via the several arteries (Kuramoto et al., 1992), on alary muscles contributing to distension and refilling of the heart, on tension development by heart muscle, and on the neuromuscular junctions as well as on the CG. A broad generalization that is increasingly supported as further experiments are done is that each of the neurohormones orchestrates a subtly different coordinated response of the circulatory, respiratory, and probably other systems to homeostatic demands--as, for example, to anoxia, osmotic stress, trauma, and the like. In general, the P0 neurohormones produce slowly developing and long-lasting increases in heart rate and strength of contraction.

Neurohormonal effects on the cardiac ganglion. Many, but clearly not all, of the effects of P0 neurohormones on intact heart preparations can be accounted for by the effects observable on the efferent output from isolated CG. These hormones act on isolated CG preparations, generally increasing burst rate and usually increasing the frequency and number of impulses as well as the duration of the bursts. In an effort to distinguish sites and mechanisms of action within the CG, a localized droplet of hormone-containing saline was applied to a lobster CG held on an array of electrodes in oil (Fig. 2B) (Cooke and Hartline, 1975). Defining excitation of a neuron as an increase in the average firing frequency of one of its axons, the study found that sensitivity to neurohormones was greatest at the impulse-initiating zone and confined to the nonspiking, integrative, DP-generating, initial axon segment of each of the neurons; the somata themselves were not sensitive.

The droplet technique and the limited region of cell sensitivity made it possible to observe how neurohormone stimulation of a single cell altered the performance of the otherwise undisturbed cardiac ganglion (see Fig. 2 and legend). Stimulation of individual axons that more than doubled their average firing rate was accommodated into the bursting pattern by correspondingly much greater increases in the rate at which those axons fired within the burst (as in Fig. 2C). Analysis of the impulses of individual axons revealed that those not exposed to the hormone in their sensitive region retained a relatively unchanged average firing frequency. The overall burst rate was altered only if an application changed the average firing frequency of the acting pacemaker (small-cell) axon. The robustness of the integrating ganglion as a rhythmic heartbeat activator was emphasized by these observations: despite major perturbation of individual axon frequency, the rhythmical output of tightly grouped motor impulses was never disrupted.

The difference in effects of the neurohormones when the substances are applied regionally to primarily act on sensitive regions of small cells or large cells adds additional evidence for differences in the intrinsic properties of these cells and, particularly, in the nature of their DPs (proctolin: Miller and Sullivan, 1981; Sullivan and Miller, 1984; octopamine: Benson, 1984; dopamine: Miller et al., 1984; Berlind, 1998 [see Fig. 9E], 2001a, b; 5HT: Kuramoto and Ebara, 1988; Kuramoto and Yamagishi, 1990; Berlind, 1998). An example is provided by observations on the effects of proctolin applied to the isolated Homarus CG (Miller and Sullivan, 1981; Sullivan and Miller, 1984). Application to small cells increased the duration and firing frequency of small-cell axons, implying an underlying increase in the duration and amplitude of the small-cell DPs. This effect was rapid (5-10 s), and it could be rapidly reversed. Application of proctolin limited to large-cell sensitive regions (Fig. 9A) caused a slowly devel oping increase in burst frequency accompanying a depolarization of up to 8 mV. By contrast with responses of the small cells, the response to a brief pulse of proctolin developed over minutes and lasted for tens of minutes.

For proctolin, there are observations suggesting its membrane-level mode of action. When proctolin was applied to Homarus large cells isolated by ligaturing and TTX treatment (Fig. 9C), the resulting depolarization could induce rhythmic driver potentials in previously quiescent cells (Sullivan and Miller, 1984). Tests of the cell input resistance (Fig. 9B) showed that proctolin increased resistance, and ionic substitution experiments indicated that this represented primarily a decrease in residual [K.sup.+] conductance. Removal of [Na.sup.+] decreased the magnitude of proctolin depolarization, which these authors interpreted as confirming that depolarization normally resulted from increased effectiveness of a "leak" conductance, for which [Na.sup.+] was the dominant ion in depolarizing the membrane. Similar observations (Fig. 9C) were made by Freschi (1989), but interpreted as representing activation of a [Na.sup.+] current.

The actions of crustacean cardioactive peptide and the FMRFamide-related peptides on isolated cardiac ganglia seem not to have been reported.

Are there intrinsic neuromodulators in the cardiac ganglion? There appear to be neurohumoral modulatory effects that arise within the CG and that supplement the actions of the fast-acting synapses. This possibility is raised by observations of the effects of acetylcholine, a catecholamine, and nitric oxide (NO) and by indications that these agents are present in the ganglia. Involvement of ACh is implied by the staining of neuropil showing the presence of acetylcholinesterase (E. Maynard, 19713; the ability of CG tissue to synthesize ACh (Sullivan and Miller, 1990); and sensitivity of the ganglia to ACh, with evidence for both nicotinic and muscarmnic receptors (Freschi and Livengood, 1989). Responses of the CG to perfused ACh closely resemble those described above for proctolin (Freschi and Livengood, 1989; Sullivan and Miller, 1990). Synthesis of catecholamines by the CG and catecholamine fluorescence in large cells of Homarus have been reported (Ocorr and Berlind, 1983). However, tyrosine-hydroxylase-like immunoreactivity was not observed in cells of the Callinectes CG (Fig. 7E) (Fort and Miller, 2001). Large, dense-cored granules are present together with the predominant clear-cored vesicles in some neuropil synaptic profiles, suggesting that a catecholamine could be co-localized with another transmitter (Morganelli and Sherman, 1987). The CG is responsive to low concentrations of dopamine and octopamine, as mentioned above. The complexities of the responses to pharmacological agonists and antagonists of monoamine receptors of vertebrates when tested on the Homarus CG leave open the possibility of intrinsic neuromodulators (Berlind, 2001a, b). Recently, nitric oxide synthase has been localized in all nine neurons of a crab (Cancer productus) CG (Labenia et al., 1998). This finding may suggest a role for amines and NO in modulating CG bursting activity via metabotropic receptors rather than via ionotropic receptors such as those of the fast excitatory synapses. A further question is whether the extrinsic regul atory fibers play a role in governing such neuromodulation.

Cellular mechanisms of neuronal modulation. Effects of the neurohormonal modulators have in common that they develop over tens of seconds and may last for tens of minutes in response to brief exposure. This time-course indicates a mode of action involving second-messenger-mediated modulation of ionic conductances. In support of this type of action, pericardial organ extracts were found to increase cAMP levels in Homarus cardiac ganglia, and pharmacological manipulations that increased cAMP mimicked the effect of PO extracts (Lemos and Berlind, 1981), Manipulations expected to increase NO levels increased cGMP levels and inhibited activity of the CG, while decreasing NO synthase activity decreased cGMP levels and increased burst frequency (Labenia et al., 1998). The localization of neuronal regions sensitive to the neurohormonal modulators on the proximal axon segments (Cooke and Hartline, 1975) is consistent with the suggestion that these modulate conductances controlling the pacemaker (or leak) potentials, D Ps, or both.

The Relation of Driver Potentials to Other Intrinsic Potentials

Plateau potentials

The recognition from study of the lobster CG (Watanabe, 1958) that the capability for patterned or bursting impulse activity is an intrinsic property of an individual neuron has now been extended to include a number of types of vertebrate neurons as well as neurons from nearly all major invertebrate groups. A sampling of the studies describing intrinsic OP-like properties underlying patterning include--besides other crustacean systems (see below)--insects (Hancox and Pitman, 1991); molluscs (Kramer and Zucker, 1985; Hurwitz and Susswein, 1996; Perrins and Weiss, 1998); annelids (Arbas and Calabrese, 1987); and vertebrates (reviews, Cooke and Stuenkel, 1985; Llinas, 1988; Kiehn and Eken, 1998). The list of vertebrate neurons includes Purkinje cells (Llinas and Sugimori, 1980); thalmic neurons (Deschenes et al., 1982; Llinas and Jahnsen, 1982); hypothalamic neurons (Legendre et al., 1982); neurons responsible for lamprey swimming (Grillner et al., 1991); neurons involved in respiratory rhythm generation (Reklin g and Feldman, 1998); motorneurons (Hounsgaard and Kiehn, 1989, review; Hultborn, 1999); and subthalamic nucleus neurons (Beurrier et al., 1999). The sustaining relevance of the studies on the CG lies in the remarkable similarity of the underlying ionic mechanisms observed in a major proportion of the pattern-forming neurons that have been analyzed.

Neurons of the decapod stomatogastric ganglion display an important variant of the intrinsic properties seen in the DPs of the CG neurons. Certain of the stomatogastric neurons exhibit plateau potentials (e.g., Russell and Hartline, 1978, 1982; Dickinson and Nagy, 1983; Harris-Warrick et at., 1992a; review, Hartline and Graubard, 1992; for voltage-clamp data see Golowasch and Marder, 1992; Zhang and Harris-Warrick, 1995). The term reflects their well-sustained, stable depolarized level (see Hartline, 1997, for diagnostic characteristics). A major embellishment is the control of the neuron's capability to produce plateau potentials by neurotransmitters, neuromodulators, or neurohormones (Russell and Hartline, 1984; Nusbaum et al., 2001). Such modification provides different output patterning (i.e., command of different behaviors) by the ganglion (reviews in Harris-Warrick et al., 1992b). Modulation is also found in other crustacean pattern generators (e.g., the crab ventilatory system, DiCaprio, 1997) and in v ertebrate motorneurons (Hounsgaard and Kiehn, 1989; review, Hultborn, 1999) and subthalamic nucleus neurons (Beurrier et al., 1999). A further embellishment on the stereotyped OP is the possibility of controlling plateau duration. Plateau potentials, like DPs (Tazaki and Cooke, 1979b), can be, and often are, terminated by hyperpolarizing current such as produced by an inhibitory synaptic potential. As with DPs, plateau potentials initiate and pattern the action potentials produced by the neuron.

Reviewing common characteristics detailed for the DPs of the CG above that have been associated with the presence of intrinsic pattern-forming capability as found in neurons from a diversity of animals and neurons, they include the following:

* Appropriate depolarization (e.g., a pacemaker, synaptic potential, or imposed depolarizing current) initiates a regenerative, slowly rising, sustained (tenths of seconds to tens of seconds), depolarizing (to above impulse threshold) potential, in a majority, resulting from activation of a slowly inactivating or non-inactivating [I.sub.Ca] Application of TTX to silence impulse initiation as well as impulse-mediated synaptic input reveals the form of these potentials. However, OP- or plateau-like potentials involving [Na.sup.+]-inward current rather than, or as well as, [I.sub.Ca] have been described ([for CG, see above], e.g., Angstadt and Choo, 1996, leech Retzius neurons; Kim and McCormick, 1998, ferret perigeniculate neurons; Su et al., 2001, rat hippocampal CA1 pyramidal cells).

* The amplitude and duration of the depolarization is shaped by the suite of [K.sup.+] conductances present and by the inactivation mechanisms of the inward currents. These include both voltage-and [Ca.sup.2+]-dependent components. The depolarization can be terminated by hyperpolarizing current (from inhibitory synaptic input or imposed current).

* The pattern-forming potentials often exhibit a relatively refractory period that may include an afterhyperpolarization. The afterpolarization may activate currents (see below) contributing toward a slow pacemaker depolarization that in turn initiates the intrinsic potential, thus contributing to rhythmic pattern generation.


Intrinsic pattern-forming characteristics combined with a pacemaking mechanism or source of sustained depolarizing drive result in rhythmic bursting or pattern formation, as exhibited in the isolated CG. One of the ionic mechanisms giving rise to pacemaker potentials is hyperpolarization-activated cationic current, now generally referred to as [I.sub.h], which has been implicated in many of the vertebrate neurons studied (reviews: Santoro and Tibbs, 1999; Luthi and McCormick, 1998; Kaupp and Seifert, 2001). As mentioned above, [I.sub.h] has not been observed in the CG motorneurons. It has been reported in a crab stomatogastric motorneuron (Kiehn and Harris-Warrick, 1992). [I.sub.h], is modulated by agents that alter cyclic nucleotides such as 5HT and other amines and peptides, suggesting a mechanism by which neuromodulators influence rhythmic pattern generators.

Another current that may contribute to pacemaking is slowly inactivating or persistent [Na.sup.+] current ([I.sub.Nap]) (e.g., Hsiao et al., 1998; Brumberg et al., 2000). By contrast, in subthalamic neurons, activation of [I.sub.NaP] changes burst firing to single-spike activity (Beurrier et al., 2000). Activation or enhancement of an [I.sub.Nap]-like current has been proposed as a mechanism of action of neuromodulators of the crustacean CG, as discussed above (Freschi, 1989; Freschi and Livengood, 1989).

Recapitulation and Conclusions

Economy in neuronal cell numbers is achieved in arthropods by a cellular architecture that permits each cell to serve several functions. Thus, CG cells are stretch receptors, integrative interneurons, and, in the case of the large cells, motorneurons as well; they may also be endowed with the capability for spontaneity and pacemaking. The sensory and integrative functions occur in a region--the soma and initial axon segment and its collateral processes--segregated from the impulse-propagating distal axon (Fig. 10). A trigger zone converts the integrated signal to conducted, all-or-none, [Na.sup.+]-mediated impulses whose frequency and pattern reflect the form of the integrated depolarizing potential generated in the initial axon segment. The presence of a combination of ionic conductances in the integrating zone that give rise to the regenerative but graded driver potential (or in other ganglia, plateau potential) enables the neuron to produce a specific form of patterned impulses that can be largely independ ent of the excitatory input. A combination of three factors-a voltage-gated [Ca.sup.2+] conductance that is inactivated by the resulting entry of [Ca.sup.2+] fast and slowly inactivating voltage-dependent [K.sup.+] conductances, and finally a [K.sup.+] conductance initiated by the rise of internal [Ca.sup.2+] --imposes a refractory period. The depth and duration of this refractory period reflects the level of previous activity. The removal or sequestering of [Ca.sup.2+] reduces [K.sup.+] conductance, allowing increased influence of depolarizing currents (stretch, pacemaker, leak, or synaptic), and removes [I.sub.Ca] inactivation, thus reducing threshold for the DP. Together the behaviors of these ionic conductances account for rhythmic recurrence of DPs and bursting, given general excitatory drive. The conductances also shape the amplitude and duration of DPs as a function of the rate of recurrence; they account for the reciprocity between burst rate and duration, and for the advanced or delayed phasing of the burst cycle when perturbed.

The CG shows remarkable reliability in providing rhythmic bursts of motor impulses in the face of a variety of imposed experimental perturbations and insults, and theoreticians have used it to model fault-tolerant networking (e.g., Sivan et al., 1999). In the CG, reliability surely arises from the intrinsic ability for rhythmic burst formation of each neuron combined with the extensive interconnection among all the neurons that ensures coordination of their activity. Electrotonic coupling links all cells of the ganglion and passes slowly changing potentials such as stretch-induced or pacemaker potentials and DPs, and these can continue to recur synchronously and rhythmically when impulse propagation has been eliminated with TTX. Each of the cells provides fast, chemically mediated excitatory synaptic input to, probably, all other cells. While these synapses are known to be impulse-mediated, the possibility that some are also active in the absence of impulses has not been examined.

Although the similarity among the nine neurons (in the case of most decapods) perhaps provides reliability by means of redundancy, each exhibits consistent differences in the detailed pattern of its impulse bursts. In the isolated CO, small cells have prominent pacemaker depolarizations and a DP of low amplitude and long duration. This form of DP thus generates a long burst of impulses at relatively steady frequency which mediates a sustained barrage of excitatory input to the motorneurons. Large cells exhibit a larger amplitude and a shorter DP, and they produce a burst having an initial high frequency of impulse firing, efficacious in rapidly depolarizing and initiating contraction in the heart muscle fibers. Other more subtle differences among the CG neurons are also documented. Additional differences become evident in considering the modulation of ganglion output by the several neurohormones secreted from the pericardial organs. Each neurohormone produces a different change of CG output and, further, has differing effects on small and large cells. The anatomical segregation of function within neurons is again observed with the restriction of responsiveness to neurohormones to the nonspiking initial axon segment.

The apparent simplicity of the CG has proven somewhat illusory. The above review will make obvious that there are many unresolved questions. To list a few:

* How is stretch (heart filling) transduced and integrated to influence the activity of the CG neurons? The existence and importance of stretch receptiveness is well documented and presumably provides the ability of the CG to monitor and adjust its activity to the results of its actions as well as to other external influences and thus function as an autonomous control system.

* Are there specific currents providing depolarizing pacemaker drive?

* Does non-impulse-mediated synaptic chemical transmission occur among CG neurons as, for example, in the stomatogastric ganglion (Graubard et al., 1983)?

* Where and how do putative neuromodulators intrinsic to the CG such as ACh and NO act?

* What is the identity of the transmitter or transmitters released from the other extrinsic regulator fiber?

* Which conductances are involved in CG modulation by the various known and proposed neurohormones or neuromodulators?

* Which possible second messenger systems participate in modulation of CG activity, how extensive are they, and what roles do they play?

New immunoreagents invite further studies exploiting the resolving capabilities of confocal microscopy for localization of receptors and enzyme systems in the CG. The availability of voltage-clamping analyses of the ionic currents, analyses of impulse-mediated synaptic interactions, and dye-fills that provide anatomical detail invite application of modeling, as in the case of the stomatogastric ganglion, to discover missing components in the available information. Demonstration that it is feasible to isolate CG neurons in primary culture and that they retain their ability to produce rhythmical bursts and respond to neurohormones (Saver et a!., 1999) opens the possibility for examining some of these questions free of the complexities presented by the interactions among the neurons in even this small, "simple" ganglion.

A candidate for the most significant contribution to neurobiology of studies on crustacean cardiac ganglia would be the unambiguous demonstration that a single neuron can be endowed with the intrinsic capability of providing an output of distinctively patterned impulses in response to nonpatterned, general excitation or single stimuli. Detailed analyses of the electrophysiological underpinning of this capability are important components of the demonstration. These observations have suggested new, much simpler explanations for the generation of complex neuronal patterns, including mechanisms by which neurohormonal and other modulators can alter these patterns and the behaviors they direct.








Professor John H. Welsh requires acknowledgement as a generative force in proposing physiological studies of crustacean cardiac ganglia and their neurohumoral modulation to receptive students (Donald Maynard, D. K. Hartline, and the author). Work in the author's laboratory reflects the intellectual input, technical skill, and hard work of many collaborators: D. K. Hartline, A. Berlind, M. E. Anderson, K. Tazaki, J. Benson, M. W. Miller, M. Mirolli, S. Talbot, R. E. Sullivan. Martha W. Goldstone, B. Haylett, and S. Grau provided valuable technical assistance. The work was supported by grants from the National Science Foundation (GB4315, GB8201, BS81-07289) and the National Institutes of Health (NS-11808), and by grants to the Bekesy Laboratory of Neurobiology by the Ida Russell Cades Fund of the University of Hawaii Foundation.

T. Fort and M. W. Miller kindly provided an unpublished figure (Fig. 7E). I thank D. K. Hartline for suggestions on the manuscript; C. Kosaki and H. Roop for assistance in preparing the reference list, and E. Garcia for help with the figures.

Permission to reproduce the figures was kindly provided by the authors and the journal copyright holders.

Received 2 July 2001; accepted 11 January 2002.

Literature Cited

Aizu, S. 1975. Fine structure of cardiac ganglion trunk in prawn, Penaeus japonicus Bates. Tissue Cell 7: 433-452.

Alexandrowicz, J. S. 1932. Innervation of the heart of the Crustacca. I. Decapoda. Q. J. Microsc. Sci. 75: 182-249.

Alexandrowicz, J. S. 1934. The innervation of the heart of Crustacea. II. Stomatopoda. Q. J. Microsc. Sci. 76: 511-548.

Alexandrowicz, J. S. 1952. Innervation of the heart of Ligia oceanica. J. Mar. Biol. Assoc. UK 31: 85-96.

Alexandrowicz, J. S. 1953. Nervous organs in the pericardial cavity of the decapod Crustacea. J. Mar. Biol. Assoc. UK 31: 563-580.

Alexandrowicz, J. S., and D. B. Carlisle. 1953. Some experiments on the function of the pericardial organs in Crustacea. J. Mar. Biol. Assoc. UK 32: 175-192.

Anderson, M. E., and I. M. Cooke. 1971. Neural activation of the heart of the lobster Homarus americanus. J. Exp. Biol. 55: 449-468.

Anderson, M. E., and D. S. Smith. 1971. Electrophysiological and structural studies on the heart muscle of the lobster Homarus americanus. Tissue Cell 3: 191-205.

Angstadt, J. D., and J. J. Choo. 1996. Sodium-dependent plateau potentials in cultured Retzius cells of the medicinal leech. J. Neurophysiol. 76: 1491-1502.

Arbas, E. A., and R. L. Calabrese. 1987. Ionic conductances underlying the activity of interneurons that control heartbeat in the medicinal leech. J. Neurosci. 7: 3945-3952.

Atwood, H. L. 1976. Organization and synaptic physiology of crustacean neuromuscular systems. Prog. Neurobiol. 7: 291-391.

Benson, J. A. 1980. Burst reset and frequency control of the neuronal oscillators in the cardiac ganglion of the crab, Portunus sanguinolentus. J. Exp. Biol. 87: 285-313.

Benson, J. A. 1981. Synaptic and regenerative responses of cardiac muscle fibers in the crab, Portunus sanguinolentus. J. Camp. Physiol. A 143: 349-356.

Benson, J. A. 1984. Octopamine alters rhythmic activity in the isolated cardiac ganglion of the crab, Portunus sanguinolentus. Neurosci. Lett. 44: 59-64.

Benson, J. A., and I. M. Cooke. 1984. Driver potentials and the organization of rhythmic bursting in crustacean ganglia. Trends Neurosci. 7: 85-91.

Berlind, A. 1982. Spontaneous and repetitive driver potentials in crab cardiac ganglion neurons. J. Comp. Physiol. A 149: 263-276.

Berlind, A. 1985. Endogenous burst-organizing potentials in two classes of neurons in the lobster cardiac ganglion respond differently to alterations in divalent ion concentration. J. Comp. Physiol. A 157: 845-856.

Berlind, A. 1989. Feedback from motor neurones to pacemaker neurones in lobster cardiac ganglion contributes to regulation of burst frequency. J. Exp. Biol. 141: 277-294.

Berlind, A. 1993. Heterogeneity of motorneuron driver potential properties along the anterior-posterior axis of the lobster cardiac ganglion. Brain Res. 609: 51-58.

Berlind, A. 1998. Dopamine and 5-hydroxytryptamine actions on the cardiac ganglion of the lobster, Homarus americanus. J. Camp. Physiol. A 182: 363-376.

Berlind, A. 2001a. Monoamine pharmacology of the lobster cardiac ganglion. Camp. Biochem. Physiol. C 128: 377-390.

Berlind, A. 2001b. Effects of haloperidol and phentolamine on the crustacean cardiac ganglion. Camp. Biochem. Physiol. C 130: 85-95.

Beurrier, C., P. Congar, B. Bioulac, and C. Hammond. 1999. Subthalamic nucleus neurons switch from single-spike activity to burst-firing mode. J. Neurosci. 19: 599-609.

Beurrier, C., B. Bioulac, and C. Hammond. 2000. Slowly inactivating sodium current ([I.sub.NaP]) underlies single-spike activity in rat subthalamic neurons. J. Neurophysiol. 83: 1951-1957.

Brown, H. F. 1964a. Electrophysiological investigations of the heart of Squilla mantis. I. The ganglionic nerve trunk. J. Exp. Biol. 41: 689-700.

Brown, H. F. 1964b. Electrophysiological investigations of the heart of Squilla mantis. II. The heart muscle. J. Exp. Biol. 41: 701-722.

Brumberg, J. C., L. G. Nowak, and D. A. McCormick. 2000. Ionic mechanisms underlying repetitive high-frequency burst firing in supragranular cortical neurons. J. Neurosci. 20: 4829-4843.

Chalgneau, J. 1983. Neurohemal organs in Crustacea. Pp. 53-89 in Neurohemal Organs of Arthropods: Their Development, Evolution, Structures, and Functions, A. P. Gupta, ed. Charles C. Thomas, Springfield, IL.

Connor, J. A. 1969. Burst activity and cellular interaction in the pacemaker ganglion of the lobster heart. J. Exp. Bial. 50: 275-295.

Cooke, I. M. 1962. The neurohumoral regulation of the crustacean heart. Ph.D. dissertation, Harvard University.

Cooke, I. M. 1966. The sites of action of pericardial organ extract and 5-hydroxytryptamine in the decapod crustacean heart. Am. Zool. 6: 107-121.

Cooke, I. M. 1988. Studies on the crustacean cardiac ganglion. Camp. Biochem. Physiol. C 91: 205-218.

Cooke I. M. 2002. Physiology of the crustacean cardiac ganglion. In The Crustacean Nervous System, Vol. 2, K. Wiese, ed. Springer-Verlag, Berlin. In press.

Cooke, I. M., and D. K. Hartline. 1975. Neurohormonal alteration of integrative properties of the cardiac ganglion of the lobster Homarus americanus, J. Exp. Biol. 63: 33-52.

Cooke, I. M., and E. L. Stuenkel. 1985. Electrophysiology of invertebrate neurosecretory cells. Pp. 115-164 in The Electrophysiology of the

Secretory Cell, A. M. Poisner and J. M. Trifaro, eds. Elsevier, Amsterdam.

Cooke, I. M., and R. E. Sullivan. 1982. Hormones and neurosecretion. Pp. 205-290 in The Biology of Crustacea, Vol. 3, D. Bliss, H. Atwood, and D. Sandeman, eds. Academic Press, New York.

Delgado, J., E. Oyola, and M. W. Miller. 2001. Localization of GABA-and glutamate-like immunoreactivity in the cardiac ganglion of the lobster Panulirus argus. J. Neurocytol. 29: 605-619.

Deschenes, M., J. P. Roy, and M. Steriade. 1982. Thalamic bursting mechanisms: an inward slow current revealed by membrane hyperpolarization, Brain Res. 239: 289-293.

DiCaprio, R. A. 1997. Plateau potentials in motor neurons in the ventilatory system of the crab. J. Exp. Biol. 200: 1725-1736.

Dickinson, P. S., and F. Nagy. 1983. Control of a central pattern generator by an identified modulatory interneurone in Crustacea. II. Induction and modification of plateau properties in pyloric neurones. J. Exp. Biol. 105: 59-82.

Dircksen, H. 1994. Distribution and physiology of crustacean cardioactive peptide in arthropods. Pp. 139-148 in Perspectives in Comparative Endocrinology. National Research Council of Canada, Ottowa.

Eyzaguirre, C., and S. W. Kuffler. 1955. Processes of excitation in the dendrites and in the soma of single isolated sensory nerve cells of the lobster and crayfish. J. Gen. Physiol. 39: 87-119.

Fatt, P., and B. Katz. 1953. The effect of inhibitory nerve impulses on a crustacean muscle fibre. J. Physiol. (Lond.) 121: 374-389.

Florey, E., and M. Rathmayer. 1978. The effects of octopamine and other amines on the heart and on neuromuscular transmission in decapod crustaceans: further evidence for a role as neurohormone. Comp. Biochem, Physiol, C 61: 229-237.

Florey, E., and M. Rathmayer. 1990. Facilitation and potentiation of transmitter release at neuromuscular synapses in the heart of Squilla mantis: functional and theoretical implications. Pp. 330-337 in Frontiers in Crustacean Neurobiology, Advances in Life Sciences, K. Wiese, W.-D. Krenz, J. Tautz, H. Reichert, and B. Mulloney, eds. Birkhauser Verlag, Basel.

Fort, T. J., and M. W. Miller. 2001. Functional organization of the cardiac system of the blue crab Callinectes sapidus: gabaergic and catecholaminergic regulatory fibers. Soc. Neurosci. Abstr. 27: 2500.

Freschi, J. E. 1989. Proctolin activates a slow, voltage-dependent sodium current in motoneurons of the lobster cardiac ganglion. Neurosci. Lett. 106: 105-111.

Freschi, J. E., and D. R. Livengood. 1989. Membrane current underlying muscarinic cholinergic excitation of motoneurons in lobster cardiac ganglion. J. Neurophysiol. 62: 984-1995.

Friesen, W. O. 1975a. Physiological anatomy and burst pattern in the cardiac ganglion of the spiny lobster Panulirus interruptus. J. Comp. Physiol. 101: 173-189.

Friesen, W. O. 1975b. Synaptic interactions in the cardiac ganglion of the spiny lobster Panulirus interruptus. J. Comp. Physiol. 101: 191-205.

Friesen, W. O. 1975c. Antifacilitation and facilitation in the cardiac ganglion of the spiny lobster Panulirus interruptus. J. Comp. Physiol. 101: 207-224.

Golowasch, J., and E. Marder. 1992. Ionic currents of the lateral pyloric neuron of the stomatogastric ganglion of the crab. J. Neurophysiol. 67: 318-331.

Graubard, K., J. A. Raper, and D. K. Hartline. 1983. Graded synaptic transmission between identified spiking neurons. J. Neurophysiol. 50:508-521.

Grillner, S., P. Wallen, and L. Brodin. 1991. Neuronal network generating locomotor behavior in lamprey: circuitry, transmitters, membrane properties, and simulation. Annu. Rev. Neurosci. 14: 169-199.

Guirguis, M. S., and J. L. Wilkens. 1995. The role of the cardioregulatory nerves in mediating heart rate responses to locomotion, reduced stroke volume, and neurohormones in Homarus americanus. Biol. Bull. 188: 179-185.

Hagiwara, S. 1961. Nervous activities of the heart in Crustacca. Ergeb. Biol. 24: 287-311.

Hagiwara, S., and T. H. Bullock. 1957. Intracellular potentials in pacemaker and integrative neurons of the lobster cardiac ganglion. J. Cell. Comp. Physiol. 50: 25-47.

Hagiwara, S., A. Watanabe, and N. Saito. 1959. Potential changes in syncytial neurons of lobster cardiac ganglion. J. Neurophysiol 22:554-572.

Hancox, J. C., and R. C. Pitman. 1991. Plateau potentials drive axonal impulse bursts in insect motoneurons. Proc. R. Soc. Lond. B244: 33-38.

Harris-Warrick, R. M., E. Marder, A. I. Selverston, and M. Moulins, eds. 1992a. Dynamic Biological Networks: The Stomatogastric Nervous System. MIT Press, Cambridge, MA. 328 pp.

Harris-Warrick, R. M., E. Nagy, and M. P. Nusbaum. 1992b. Neuromodulation of stomatogastric networks by identified neurons and transmitters. Pp. 87-138 in Dynamic Biological Networks: The Stomatogastric Nervous System, R. M. Harris-Warrick, E. Marder, A. I. Selverston, and M. Moulins, eds. MIT Press, Cambridge, MA.

Hartline, D. K. 1967. Impulse identification and axon mapping of the nine neurons in the cardiac ganglion of the lobster, Homarus americanus. J. Exp. Biol. 47: 327-340.

Hartline, D. K. 1979. Integrative neurophysiology of the lobster cardiac ganglion. Am. Zool. 19: 53-65.

Hartline, D. K. 1997. Plateau potential. Pp. 1656-1657 in The Encyclopedia of Neuroscience, G. Adelman and B. H. Smith, eds. Elsevier Science, Amsterdam.

Hartline, D. K., and I. M. Cooke. 1969. Postsynaptic membrane response predicted from presynaptic input pattern in lobster cardiac ganglion. Science 164: 1080-1082.

Hartline, D. K., and K. Graubard. 1992. Cellular and synaptic properties in the crustacean stomatogastric nervous system. Pp. 31-86 in Dynamic Biological Networks: The Stomatogastric Nervous System, R. M. Harris-Warrick, E. Marder, A. I. Selverston, and M. Moulins, eds. MIT Press, Cambridge, MA.

Hashemzadeh-Gargari, H., and J. Freschi. 1992. The effects of glutamate agonists on voltage-clamped motoneurons of the lobster cardiac ganglion. J. Exp. Biol. 169: 53-63.

Hawkins, W. E., and H. D. Howse, 1978. A light and electron microscopic study of the cardiac ganglion of the blue crab Callinectes sapidus Rathbun. Trans. Am. Microsc, Soc. 97: 363-380.

Hounsgaard, J., and O. Kiehu. 1989. Serotonin-induced bistability of turtle motoneurons caused by a nifedipine-sensitive calcium plateau potential. J. Physiol. (Lond.) 414: 265-282.

Hsiao, C. F., C. A. Del Negro, P. R. Trueblood, and S. H. Chandler. 1998. Ionic basis for serotonin-induced bistable membrane properties in guinea pig trigeminal motoneurons. J. Neurophysiol, 79: 2847-2856.

Hultborn, H. 1999. Plateau potentials and their role in regulating motoneuronal firing. Prog. Brain Res. 123: 39-48.

Hurwitz, I., and A. J. Susswein. 1996. B64, a newly identified central pattern generator element producing a phase switch from protraction to retraction in buccal motor programs of Aplysia californica, J. Neurophysiol. 75: 1327-1344.

Irisawa, A., and K. Hama. 1965. Contact of adjacent nerve fibers in the cardiac nerve of mantis shrimp. Jpn. J. Physiol. 15: 323-330.

Kaupp, U. B., and R. Seifert, 2001. Molecular diversity of pacemaker ion channels. Annu. Rev. Physiol. 63: 235-257.

Kerrison, J., and J. E. Freschi. 1992. The effects of [gamma]-aminobutyric acid on voltage-clamped motoneurons of the lobster cardiac ganglion. Comp. Biochem. Physiol. C 101: 227-233.

Kiehn, O., and T. Eken. 1998. Functional role of plateau potentials in vertebrate motor neurons. Curr. Opin. Neurobiol. 8: 746-752.

Kiehn, O., and R. M. Harris-Warrick. 1992. 5-HT modulation of hyperpolarization-activated inward current and calcium-dependent outward current in a crustacean motor neuron. J. Neurophysiol. 68: 496-508.

Kim, U., and D. A. McCormick. 1998. Functional and ionic properties of a slow afterhyperpolarization in ferret perigeniculate neurons in vitro. J. Neurophysiol. 80: 1222-1235.

Krajniak, K. G. 1991. The identification and structure-activity relations of a cardioactivc FMRFamide-related peptide from the blue crab, Callinectes sapidus. Peptides 12: 1295-1302.

Kramer, R. H., and R. S. Zucker. 1985. Calcium-dependent inward current in Aplysia bursting pacemaker neurones. J. Physiol. (Lond.) 363: 107-130.

Kuramoto, T., and A. Ebara. 1984a. Effects of perfusion pressure on the isolated heart of the lobster, Panulirus japonicus. J. Exp. Biol. 109: 121-140.

Kuramoto, T., and A. Ebara. 1984b. Neurohormonal modulation of the cardiac outflow through the cardioarterial valve in the lobster. J. Exp. Biol. 111: 123-130.

Kuramoto, T., and A. Ebara. 1985. Effects of perfusion pressure on the bursting neurones in the intact or segmented cardiac ganglion of the lobster, Panulirus japonicus. J. Neurosci. Res. 13: 569-580.

Kuramoto, T., and A. Ebara. 1988. Combined effects of 5-hydroxytryptamine and filling pressure on the isolated heart of the lobster, Panulirus japonicus. J. Comp. Physiol. B 158: 403-412.

Kuramoto, T., and A. Ebara. 1991. Combined effects of octopamine and filling pressure on the isolated heart of the lobster, Panulirus japonicus. J. Comp. Physiol. B 161: 339-347.

Kuramoto, T., and K. Kuwasawa. 1980. Ganglionic activation of the myocardium of the lobster, Panulirus japonicus. J. Comp. Physiol. 139: 67-76.

Kuramoto, T., and H. Yamagishi. 1990. Physiological anatomy, burst formation, and burst frequency of the cardiac ganglion of crustaceans. Physiol. Zool. 63: 102-116.

Kuramoto, T., E. Hirose, and M. Tani. 1992. Neuromuscular transmission and hormonal modulation in the cardioarterial valve of the lobster, Homarus americanus. Pp. 62-69 in Phylogenetic Models in Functional Coupling of the CNS and the Cardiovascular System, R. B. Hill, K. Kuwasawa, B. R. McMahon, and T. Kuramoto, eds. Karger, Basel.

Labenia, J., N. L. Scholz, M. F. Goy, and K. Graubard. 1998. NO/cGMP modulates the crustacean cardiac ganglion. Soc. Neurosci. Abstr. 24: 360.

Legendre, P., I. M. Cooke, and J. D. Vincent. 1982. Regenerative responses of long duration recorded intracellularly from dispersed cell cultures of fetal mouse hypothalamus. J. Neurophysiol. 48: 1121-1141.

Lemos, J. R., and A. Berlind. 1981. Cyclic adenosine monophosphate mediation of peptide neurohormone effects on the lobster cardiac ganglion. J. Exp. Biol. 90: 307-326.

Livengood, D. R. 1983. Coupling ratio of the Na-K pump in the lobster cardiac ganglion. J. Gen. Physiol. 82: 853-874.

Livengood, D. R., and K. Kusano. 1972. Evidence for an electrogenic sodium pump in follower cells of the lobster cardiac ganglion. J. Neurophysiol. 35: 170-186.

Llinas, R. R. 1988. The intrinsic electrophysiological properties of mammalian neurons: insights into central nervous system function. Science 242: 1654-1664.

Llinas, R., and H. Jahnsen. 1982. Electrophysiology of mammalian thalamic neurones in vitro. Nature 297: 406-408.

Llinas, R., and M. Sugimori. 1980. Electrophysiological properties of in vitro Purkinje cell dendrites in mammalian cerebellar slices. J. Physiol. (Lond.) 305: 197-213.

Luthi, A., and D. A. McCormick. 1998. H-Current: properties of a neuronal and network pacemaker. Neuron 21: 9-12.

Marder, E., and R. L. Calabrese. 1996. Principles of rhythmic motor pattern generation. Physiol. Rev. 76: 687-717.

Matsui, K. 1955. Spontaneous discharges of the isolated ganglionic trunk of the lobster heart (Panulirus japonicus). Sci. Rep. Tokyo Kyoiku Daigaku B 7: 257-268.

Matsui, K., N. Ai, and K. Kuwasawa. 1973. Spontaneous inhibitory post-synaptic potentials in the cardiac ganglion preparation of the lobster Panulirus japonicus. Comp. Biochem. Physiol. A 44: 953-965.

Matsui, K., K. Kuwasawa, and T. Kuramoto. 1977. Periodic bursts in large cell preparations of the lobster cardiac ganglion (Panulirus japonicus). Comp. Biochem. Physiol. A 56: 313-324.

Mayeri, E. M. 1973a. Functional organization of the cardiac ganglion of the lobster, Homarus americanus. J. Gen. Physiol. 62: 448-472.

Mayeri, E. M. 1973b. A relaxation oscillator description of the burst generating mechanism in the cardiac ganglion of the lobster, Homarus americanus. J. Gen. Physiol. 62: 473-488.

Maynard, D. M. 1953. Activity in a crustacean ganglion. I. Cardioinhibition and acceleration in Panulirus argus. Biol. Bull. 104: 156-170.

Maynard, D. M. 1955. Activity in a crustacean ganglion II. Pattern and interaction in burst formation. Biol. Bull. 109: 420-436.

Maynard, D. M. 1960. Circulation and heart function. Pp. 161-226 in The Physiology of Crustacea, T. H. Waterman, ed. Academic Press, New York.

Maynard, D. M. 1961. Cardiac inhibition in decapod Crustaces. Pp. 144-178 in Nervous Inhibition, E. Florey, ed. Pergamon Press, New York.

Maynard, E. 1971. Microscopic localization of cholinesterases in the nervous systems of lobsters, Panulirus argus and Homarus americanus. Tissue Cell 3: 215-230.

Mercier, A. J., and R. T. Russenes. 1992. Modulation of crayfish hearts by FMRFamide-related peptides. Biol. Bull. 182: 333-340.

Mercier, A. J., I. Orchard, V. TeBrugge, and M. Skerrett. 1993. Isolation of two FMRFamide-related peptides from crayfish pericardial organs. Peptides 14: 137-143.

Miller, M. W., and R. E. Sullivan. 1981. Some effects of proctolin on the cardiac ganglion of the Maine lobster, Homarus americanus (Milne Edwards). J. Neurobiol. 12: 629-639.

Miller, M. W., J. A. Benson, and A. Berlind. 1984. Excitatory effects of dopamine on the cardiac ganglia of the crabs Portunus sanguinolentus and Podophthalmus vigil. J. Exp. Biol. 108: 97-118.

Mirolli, M., I. M. Cooke, S. R. Talbott, and M. W. Miller. 1987. Structure and localization of synaptic complexes in the cardiac ganglion of a portunid crab. J. Neurocytol. 16: 115-130.

Morganelli, P. M., and R. G. Sherman. 1987. Nerve terminals and synapses in the cardiac ganglion of the adult lobster Homarus americanus. J. Morphol. 191: 177-191.

Nusbaum, M. P., D. M. Blitz, A. M. Swensen, D. Wood, and E. Marder. 2001. The roles of co-transmission in neural network modulation. Trends Neurosci. 24: 146-154.

Ocorr, K. A., and A. Berlind. 1983. The identification and localization of a catecholamine in the motor neurons of the lobster cardiac ganglion. J. Neurobiol. 14: 51-59.

Ohsawa, K. 1972. Morphological organization and fine structures of the cardiac ganglion of the lobster, Panulirus japonicus. Sci. Rep. Tokyo Kyoiku Daigaku B 15: 1-24.

Orkand, R. K. 1962. The relation between membrane potential and contraction in single crayfish muscle fibres. J. Physiol. (Lond.) 161: 143-159.

Otani, T., and T. H. Bullock. 1959. Effects of presetting the membrane potential of the soma of spontaneous and integrating ganglion cells. Physiol. Zool. 32: 104-114.

Perrins, R., and K. R. Weiss. 1998. Compartmentalization of information processing in an Aplysia feeding circuit interneuron through membrane properties and synaptic interactions. J. Neurosci. 18: 3977-3989.

Rekling, J. C., and J. L. Feldman. 1998. Prebotzinger complex and pacemaker neurons: hypothesized site and kernel for respiratory rhythm generation. Annu. Rev. Physiol. 60: 385-405.

Russell, D. F., and D. K. Hartline. 1978. Bursting neural networks: a reexamination. Science 200: 453-456.

Russell, D. F., and D. K. Hartline. 1982. Slow active potentials and bursting motor patterns in pyloric network of the lobsters, Ponulirus interruptus. J. Neurophysiol. 48: 914-937.

Russell, D. F., and D. K. Hartline. 1984. Synaptic regulation of properties and burst oscillations of neurons in the gastric mill system of spiny lobster, Panulirus interruptus. J. Neurophysiol. 52: 54-73.

Sakurai, A., and H. Yamagishi. 1998. Identification of two cardioacceleratory neurons in the isopod crustacean, Ligia exotica and their effects on cardiac ganglion cells. J. Comp. Physiol. A 182: 145-152.

Santoro, B., and G. R. Tibbs. 1999. The HCN gene family: molecular basis of the hyperpolarization-activation pacemaker channels. Ann. NY Acad. Sci. 868: 741-764.

Saver, M. A., and J. L. Wilkens. 1998. Comparison of the effects of live hormones on intact and open heart cardiac ganglionic output and myocardial contractility in the shore crab Carcinus maenas. Comp. Biochem. Physiol. A 120: 301-310.

Saver, M. A., J. L. Wilkens, and C. N. Airriess. 1998. Proctolin affects the activity of the cardiac ganglion, myocardium, and cardioarterial valves in Carcinus maenas hearts. J. Comp. Physiol. B 168: 473-482.

Saver, M. A., J. L. Wilkins, and N. I. Syed. 1999. In situ and in vitro identification and characterization of cardiac ganglion neurons in the crab, Carcinus maenas. J. Neurophysiol. 81: 2964-2976.

Shimahara, T. 1969a. The inhibitory post-synpatic potential in the cardiac ganglion cell of the lobster, Panulirus japonicus. Sci. Rep. Tokyo Kyoiku Daigaku B 14: 9-26.

Shimahara, T. 1969b. The effect of the acceleratory nerve on the electrical activity of the lobster cardiac ganglion. Zool. Mag. 78: 351-355.

Sivan, E., H. Parnas, and D. Dolev. 1999. Fault tolerance in the cardiac ganglion of the lobster. Biol. Cybern. 81: 11-23.

Smith, R. I. 1947. The action of electrical stimulation and of certain drugs on cardiac nerves of the crab, Cancer irroratus. Biol. Bull. 93: 72-88.

Stangier, J., C. Hilbich, K. Beyreuther, and R. Keller. 1987. A novel cardioactive peptide (CCAP) from pericardial organs of the shore crab Carcinus maenas. Proc. Natl, Acad. Sci. USA 84: 575-579.

Su, H., G. Alroy, E. D. Kirson, and Y. Yaari. 2001. Extracellular calcium modulates persistent sodium current-dependent burst-firing in hippocampal pyramidal neurons. J. Neurosci. 21: 4173-4182.

Sullivan, R. E. 1979. A proctolin-like peptide in crab pericardial organs. J. Exp. Zool. 210: 543-552.

Sullivan, R. E., and M. W. Miller. 1984. Dual effects of proctolin on the rhythmic burst activity of the cardiac ganglion. J. Neurobiol. 15: 173-196.

Sullivan, R. E., and N. W. Miller. 1990. Cholinergic activation of the lobster cardiac ganglion. J. Neurobiol. 21: 639-650.

Sullivan, R. E., B. J. Friend, and D. L. Barker. 1977. Structure and function of spiny lobster ligamental nerve plexuses: evidence for synthesis, storage and secretion of biogenic amines. J. Neurobiol. 8: 581-605.

Suzuki, 5. 1934. Ganglion cells in the heart of Ligia exotica (Roux). Sci. Repts. Tohoku Imp. Univ. Fourth Ser. 9: 214-218. [Cited in Maynard, 1960.]

Tameyasu, T. 1976. Intracellular potentials in the small cells and cellular interaction in the cardiac ganglion of the lobster, Panulirus japonicus. Comp. Biochem. Physiol. 54A: 191-196.

Tameyasu, T. 1987. The mechanism of the burst formation in the cardiac ganglion of the lobster (Panulirus japonicus): a re-examination. J. Comp. Physiol. A 161: 389-398.

Tazaki, K. 1967. Intracellular potential changes in the cardiac ganglion cell of the crab, Eriocheir joponicus. Sci. Rep. Tokyo Kyoiku Daigaku B12: 191-210.

Tazaki, K. 1970. Slow potential changes during the burst in the cardiac ganglion of the crab, Eriocheir japonicus. Annor. Zool. Jpn. 43: 63-69.

Tazaki, K. 1971a. The effects of tetrodotoxin on the slow potential and spikes in the cardiac ganglion of a crab, Eriocheir joponicus. Jpn. J. Physiol. 21: 529 -536.

Tazaki, K. 1971b. Small synaptic potentials in burst activity of large neurons in the lobster cardiac ganglion. Jpn. J. Physiol. 21: 645-658.

Tazaki, K. 1972. The burst activity of different cell regions and intercellular co-ordination in the cardiac ganglion of the crab, Eriocheir japonicus. J. Exp. Biol. 57: 713-726.

Tazaki, K. 1973. Impulse activity and pattern of large and small neurones in the cardiac ganglion of the lobster, Panulirus japonicus. J. Exp. Biol. 58: 473-486.

Tazaki, K., and I. M. Cooke. 1979a. Spontaneous electrical activity and interaction of large and small cells in the cardiac ganglion of the crab, Portunus sanguinolentus/ J. Neurophysiol. 42: 975-999.

Tazaki, K., and I. M. Cooke. 1979b. Isolation and characterization of slow, depolarizing responses of cardiac ganglion neurons in the crab, Portunus Portunus sanguinolentus. J. Neurophysiol. 42: 1000-1021.

Tazaki, K., and I. M. Cooke. 1979c. The ionic bases of slow depolarizing responses of cardiac ganglion neurons in the crab, Portunus sanguinolentus. J. Neurophysiol. 42: 1022-1047.

Tazaki, K., and I. M. Cooke. 1983a. Topographical localization of function in the cardiac ganglion of the crab, Portunus sanguinolentus. J. Comp. Physiol. A 151: 311-328.

Tazaki, K., and I. M. Cooke. 1983b. Separation of neuronal sites of driver potential and impulse generation by ligaturing in the cardiac ganglion of the lobster, Homarus americanus. J. Comp. Physiol. A 151: 329-346.

Tazaki, K., and I. M. Cooke. 1983c. Neuronal mechanisms underlying rhythmic bursts in crustacean cardiac ganglia. Symp. Soc. Exp. Biol. 37: 129 -157.

Tazaki, K., and I. M. Cooke. 1986. Currents under voltage clamp of burst-forming neurons of the cardiac ganglion of the lobster (Homnarus americanus). J. Neurophysiol. 56: 1739-1762.

Tazaki, K., and I. M. Cooke. 1990. Characterization of Ca current underlying burst formation in lobster cardiac ganglion motorneurons. J. Neurophysiol. 63: 370-384.

Terzuolo, C. A., and T. H. Bullock. 1958. Acceleration and inhibition in crustacean ganglion cells. Arch. Ital. Biol. 96: 117-134.

Trimmer, B. A., L.A. Kobierski, and E. A. Kravitz. 1987. Purification and characterization of FMRFamide-like immunoreactive substances from the lobster nervous system: isolation and sequence analysis of two closely related peptides. J. Comp. Neurol. 266: 16-26.

van der Kloot, W. 1970. The electrophysiology of muscle fibers in the hearts of decapod crustaceans. J. Exp. Zool. 174: 367-380.

Watanabe, A. 1958. The interaction of electrical activity among neurons of lobster cardiac ganglion. .Jpn. J. Physiol. 8: 305-318.

Watanabe, A., and T. H. Bullock. 1960. Modulation of activity of one neuron by subthreshold slow potentials in another in lobster cardiac ganglion. J. Gen. Physiol. 43: 103 1-1045.

Watanabe, A., and K. Takeda. 1963. The spread of excitation among neurons in the heart ganglion of the stomatopod, Squilla oratorio, J. Gen. Physiol. 46: 773.

Watanabe, A., S. Obara, T. Akiyama, and K. Yumoto 1967a. Electrical properties of the pacemaker neurons in the heart ganglion of a stomatopod, Squilla oratorio. J. Gen. Physiol. 50: 813-838.

Watanabe, A., S. Obana, and T. Akiyama. 1967b. Pacemaker potentials for the periodic burst discharge in the heart ganglion of a stomatopod, Squilla oratorio. J. Gen. Physiol. 50: 839-862.

Watanabe, A., S. Obara, and T. Akiyama. 1968. Inhibitory synapses on pacemaker neurons in the heart ganglion of a stomatopod, Squilla oratoria. J. Gen. Physiol. 52: 908-924.

Watanabe, A., S. Obara, and T. Akiyama. 1969. Acceleratory synapses on pacemaker neurons in the heart ganglion of a stomatopod, Squilla oratorio. J. Gen. Physiol. 54: 212-231.

Welsh, J. H., and D. M. Maynard. 1951. Electrical activity of a simple ganglion. Fedr. Proc. 10: 145.

Wiens, T. J. 1982. Small systems of neurons: control of rhythmic and reflex activities. Pp. 193-240 in The Biology of Crustacea, Vol. 4, D. Bliss, D. Sandeman, and H. Atwood, eds. Academic Press, New York.

Wiersma, C. A., and E. Novitski. 1942. The mechanism of the nervous regulation of the crayfish heart. J. Exp. Biol. 19: 255-265.

Wilkens, J. L. 1993. Re-evaluation of the stretch sensitivity hypothesis of crustacean hearts: Hypoxia, not lack of stretch, causes reduction in heart rate of isolated hearts. J. Exp. Biol. 176: 223-232.

Wilkens, J. L. 1999. Evolution of the cardiovascular system in Crustacea. Am, Zool. 39: 199-214.

Wilkens, J. L., and A. J. Mercier. 1993. Peptidergic modulation of cardiac performance in isolated hearts from the shore crab, Carcinus maenas. Physiol Zool. 66: 237-256.

Wilkens, J. L., T. Kuramoto, and B. R. McMahon. 1996. The effects of six pericardial hormones and hypoxia on the semi-isolated heart and sternal arterial valve of the lobster Homarus americanus. Comp. Biochem. Physiol. C 114: 57-65.

Yamagishi, H., H. Ando, and T. Makioka. 1997. Myogenic heartbeat in the primitive crustacean Triops longicaudatus. Biol. Bull. 193: 350-358.

Yazawa, T., and K. Kuwasawa. 1992. Intrinsic and extrinsic neural and neurohumoral control of the decapod heart. Experientia 48: 834-840.

Yazawa, T., K. Tanaka, M. Yasumatsu, M. Otokawa, Y. Aihara, K. Ohsuga, and K. Kuwasawa. 1998. A pharmacological and HPLC analysis of the excitatory transmitter of the cardiac ganglion in the heart of the isopod crustacean Bathynomus doederleini. Can. J. Physiol. Pharmacol. 76: 599-604.

Zhang, B., and R. M. Harris-Warrick. 1995. Calcium-dependent plateau potentials in a crab stomatogastric ganglion motor neuron. I. Calcium current and its modulation by serotonin. J. Neurophysiol. 74: 1929-1937.

Abbreviations: CG, cardiac ganglion; DP, driver potential; PO, pericardial organ.

COPYRIGHT 2002 University of Chicago Press
No portion of this article can be reproduced without the express written permission from the copyright holder.
Copyright 2002 Gale, Cengage Learning. All rights reserved.

Article Details
Printer friendly Cite/link Email Feedback
Author:Cooke, Ian M.
Publication:The Biological Bulletin
Geographic Code:1USA
Date:Apr 1, 2002
Previous Article:Evolution of cannabinoid receptors in vertebrates: identification of a [CB.sub.2] gene in the puffer fish Fugu rubripes.
Next Article:Timing is everything: the effects of putative dopamine antagonists on metamorphosis vary with larval age and experimental suration in the prosobranch...

Terms of use | Privacy policy | Copyright © 2020 Farlex, Inc. | Feedback | For webmasters