The Distribution of Nadph-Diaphorase-Labelled Interneurons and the Role of Nitric Oxide in the Swimming System of Xenopus Laevis Larvae

The labile and highly diffusible diatomic free radical gas nitric oxide (NO) is now known to be widely distributed and to play a diversity of roles in both vertebrates (for a review, see Moncada and Palmer, 1991) and invertebrates (for a review, see Vincent, 1995). Within the central nervous system (CNS), NO is recognised as an important, albeit unconventional, chemical messenger that is intimately involved with the regulation of synaptic function in a variety of brain regions (for reviews, see Schuman and Madison, 1994; Vincent, 1994). The location of nitrergic neurons across a broad phylogenetic range has been studied using histological and/or immunocytochemical markers for nitric oxide synthases (NOS), the synthetic enzymes that convert L-arginine to NO. One simple and reliable staining method involves the nicotinamide adenine dinucleotide phosphate (NADPH) diaphorase histochemical reaction, because the enzyme responsible for this reaction is NOS (Hope et al., 1991). This technique has revealed highly localised staining in particular clusters of neurons and their processes in various locations that extend to virtually all areas of the brain (for a review, see Vincent and Hope, 1992). Since this picture differs among species and can also alter at different developmental stages in the same species, it seems likely that NO has highly specific behavioural roles, many of which are either poorly defined or not yet discovered.

In particular, nothing is known about the possible roles of NO in the control of the spinal rhythm-generating networks responsible for locomotion in vertebrates. For this reason, we have chosen to study the distribution of nitrergic neurons and their role in a very simple vertebrate locomotor system, that controlling swimming in newly hatched Xenopus laevis tadpoles. Swimming in these animals at the time of hatching (stage 37/8; Nieuwkoop and Faber, 1956) involves the coordinated contractions of segmented myotomal muscles that are distributed along each side of the body. The muscles on the left and right sides contract in strict alternation during swimming, while contracting in a head-to-tail sequence down each side (Kahn et al., 1982). This cycle of muscle contraction creates back-propagating waves of body curvature that generate reactive thrust against the water column and lead to the forward propulsion of the organism. Fictive swimming activity involves the ventral root discharge appropriate to drive this swimming rhythm in immobilised tadpoles and can be generated by a network of neurons located entirely within the spinal cord, often called the swim central pattern generator. At these early stages in development, the spinal cord contains as few as eight types of differentiated neuron (Roberts and Clarke, 1982), of which three are known to be involved in the generation of rhythmic swimming activity (for a review, see Roberts, 1990). After the first day of larval life (stage 42; Nieuwkoop and Faber, 1956), the swimming system is dramatically transformed to produce a pattern in which motor neurons that used to fire a single spike per cycle at stage 37/8 (Soffe, 1990) now discharge a burst of action potentials in each cycle of swimming (Sillar et al., 1991, 1992a,b). This transition is dependent upon certain brainstem neurons with descending axonal projections to the spinal cord (Sillar et al., 1995a,b). These raphe neurons modulate the output of the spinal locomotor circuitry via a range of serotonin-dependent actions (Sillar et al., 1993) including changes in the strength of synaptic connections in the spinal cord (McDearmid et al., 1997) and in the electrical properties of spinal neurons (Scrymgeour-Wedderburn et al., 1997; Reith and Sillar, 1998).

In this paper, we further describe the possible contribution of descending projection neurons to the control of swimming and present evidence that one such system involves the release of NO. We first examine the distribution of NO-synthesising cells in the CNS using NADPH-diaphorase histochemistry in cryostat-sectioned material and in wholemount preparations of the excised CNS. Next, we explore the effects of NO on the rhythmic motor pattern for swimming in paralysed tadpoles by manipulating the level of exogenous NO and the activity of endogenous NOS. We discuss these results in the broader context of descending modulatory control systems in the vertebrate brainstem and argue that NO is used by brainstem neurons as a cotransmitter that slows swimming through potentiation of inhibitory mechanisms. These results have not been published previously except in abstract form (McLean and Sillar, 1999).

Xenopus laevis larvae (stage 42; pre-feeding) were obtained by induced breeding from an adult laboratory colony and staged according to Nieuwkoop and Faber (1956). For wholemount histochemistry, animals were deeply anaesthetised in tricaine methanesulphonate (MS-222; 0.01–0.1 %) and fixed at room temperature (20–22 °C) for 2–4 h in ice-cold 4 % paraformaldehyde. The animals were then rinsed in 30 % sucrose in 0.1 mol l⁻¹ phosphate buffer (PB; pH 7.4) and pinned through the notocord with finely etched tungsten pins to a Sylgard (Dow-Corning) platform within a Perspex dissecting bath filled with PB or phosphate-buffered saline (PBS). Using fine needles, the skin was removed, and the CNS was then carefully dissected out to the level of myotomes 4–6 (as numbered from the otic capsule). Immediately after dissection, the CNS was transferred to 30 % sucrose/PB and stored for up to 24 h. The CNS was then pretreated in 0.3 % Triton X-100/PB to aid penetration of reagents. Subsequently, each CNS was transferred to the incubation medium [0.1 mol l⁻¹ PB containing 1 mg ml⁻¹ β-nicotinamide adenine dinucleotide phosphate (reduced form, NADPH), 0.1 mg ml⁻¹ Nitroblue Tetrazolium (NBT) and 0.3 % Triton X-100] for NADPH-diaphorase histochemistry. Incubations were carried out at 37 °C in a humid chamber for 1–3 h. All reagents were purchased from Sigma (UK).

The enzyme reaction was stopped by transferring the preparations to a fresh PB solution. The stained preparations were then dehydrated in a graded alcohol series (50, 70, 90, 95 and 100 %), cleared in methylbenzoate and xylene, and mounted in DPX on cavity slides. The resulting preparations were studied and catalogued using a Zeiss Axiolab microscope equipped with a camera. Photographs were taken with an Olympus OM-4 Ti camera using Fuji 200 ASA film, and tracings of the resulting photographs were made on a Jessop light box. Drawings and cell counts were made with a camera lucida attachment. The results derive from 20 processed stage 42 central nervous systems.

For transverse sectioning, 12 animals were anaesthetised in MS-222, scored along the dorsal fin to aid fixative penetration and immersed in ice-cold 4 % paraformaldehyde for 2–4 h at room temperature. To remove aldehydes, tadpoles were transferred to 30 % sucrose/PB until they sank, and they were then embedded in either rat brain or rat liver that had also been fixed in 4 % paraformaldehyde. Tissues were then frozen in a cryostat (Leica Jung Frigocut 2800E; −18 °C) in a supporting cryomatrix and sectioned at 5–25 μm. The cryostat-cut tissues were thaw-mounted directly on poly-L-lysine-coated slides and allowed to dry at room temperature for 1–2 h prior to histochemical processing.

The protocol for the NADPH-diaphorase histochemical reaction was similar to that described above for wholemounts. Briefly, the sections were incubated at 37 °C for 2–3 h in the staining solution described above. The slides were rinsed in PB and distilled water and then allowed to dry overnight at room temperature. The slides were then cleared in xylene and coverslipped in DPX. The resulting slides were studied and catalogued using the equipment described above.

All experiments were performed on Xenopus laevis tadpoles at larval stage 42. Animals were immobilised in 10 μmol l⁻¹ α-bungarotoxin and secured in a chamber with recirculating frog Ringer (ionic composition in mmol l⁻¹: NaCl, 120; KCl, 2.5; CaCl₂, 2; MgCl₂, 1; NaHCO₃, 2.5; Hepes, 10, pH 7.4). The experimental chamber was gravity-fed from a 10 ml header tank via a stock bottle containing 100 ml of saline. L-NAME, D-NAME, L-NNA, N-acetylpenicillamine (NAP) (Sigma, UK) and S-nitroso-N-acetylpenicillamine (SNAP) (Chemistry Department, University of St Andrews) were first dissolved in distilled water (L-NAME and D-NAME) or dimethyl sulphoxide (DMSO) [L-NNA, NAP and SNAP; <0.01 %] and then added to the stock bottle to achieve the desired final bath concentration. The SNAP was NO-depleted by keeping it at room temperature under light for 24 h before use. Animals were secured on their sides using finely etched tungsten pins (through the notocord) to the Sylgard (Dow Corning) surface of a rotatable Perspex platform housed within the experimental chamber; one pin was placed rostrally, at the level of the otic capsule, and another caudally, approximately half-way along the length of the body. The flank skin on the left side was carefully removed to the level of the otic capsule using finely etched tungsten needles to reveal the underlying myotomal swimming muscles (see schematic diagram in Fig. 3). In experiments using NOS inhibitors, the first three myotomes on the left side were removed to facilitate drug access (see schematic diagram in Fig. 4). Fictive motor patterns appropriate to drive swimming behaviour were recorded in clefts between myotomal swimming muscles, as described previously (Sillar et al., 1991). In all experiments, episodes of fictive swimming were evoked by a brief 1 ms current pulse applied to the tail skin via a glass suction electrode. Data were recorded, displayed conventionally and stored on videotape using a PCM adaptor (Medical Systems Corporation). Hard copy recordings were made off-line using a thermal plotter (Graphtec). Data were digitised with a CED 1401 and analysed off-line using Spike 2 software. The data presented here derive from experiments on 61 stage 42 larvae.

In cross sections of intact tadpoles, positive staining was located in structures external to the CNS, the detailed description of which is beyond the scope of the present paper. Briefly, however, labelled areas included the walls of blood vessels and various cells in the retina and the skin (data not illustrated). These tissues are known to contain NOS in a variety of species (Salter et al., 1991), and our observations therefore support the conclusion that the positive staining described here corresponds with NO-producing cells.

Within the CNS, intense labelling was found in three discrete bilaterally symmetrical clusters of neurons located in the brainstem (Fig. 1A,B). Only a few cells were labelled in the forebrain, and no cell bodies were stained in the spinal cord. The most numerous and most caudal population extended from the junction between the spinal cord and hindbrain rostrally for 200–300 μm and consisted of approximately 150 neurons (70–80 neurons per side; Fig. 1Ci). A cross section of the CNS in this region is shown in Fig. 1Bi. The majority of neurons in this group had cell bodies located ventrally and close to the neurocoel, extending ventrolaterally to mid-dorsal positions. At more rostral locations, this cluster split into two subgroups with the second, smaller population occupying a more dorsal position. All the neurons in the caudal hindbrain cluster had unipolar somata with usually a single, but sometimes a double, neurite extending laterally into the marginal zone of the brainstem, where an extensive ramification of dendritic processes occurred. Many of the neurons in this cluster had contralaterally projecting processes that could be detected as they coursed beneath the neurocoel (Fig. 1Bi). Beyond the rostral extent of this population there followed a region of the brainstem in which no staining of cell bodies occurred. The next most rostral cluster occurred approximately 200 μm from the hindbrain–midbrain boundary and extended caudally for approximately 100 μm. These neurons, numbering approximately 80 (approximately 40 neurons per side; Fig. 1Cii), were positioned ventromedially, with neurites extending from the unipolar somata laterally into the margins of the rostral brainstem (Fig. 1Bii). No evidence was found for crossing projections in neurons belonging to this cluster. The most rostral brainstem cluster was located very close to the midbrain–forebrain boundary and numbered approximately 40 somata (approximately 20 per side; Fig. 1Ciii). These tightly packed cells occupied a more dorsal and lateral region of the brainstem (Fig. 1Biii). Their distinctive processes were unilateral and consisted of very long primary neurites that projected ventrally, where they terminated in a plexus of dendritic processes in the ventrolateral marginal zone.

The topology of these three primary clusters of putative nitrergic brainstem neurons was clearest in wholemount preparations of the excised intact CNS (Figs 1A, 2). From a lateral perspective, the location of the three groups in the dorso-ventral plane is readily apparent (Fig. 1A). From the dorsal aspect, it is clear that each cluster is bilaterally symmetrical (Figs 1A, 2A), that the most caudal cluster represents the most numerous population and that the cells of this group give rise to processes that cross the ventral commissure beneath the neurocoel (Fig. 2B). The wholemount preparations also allow a description of the axonal projections of these nitrergic neurons in the brainstem. The caudal group emanated axons that projected caudally in the marginal zone and reached only the rostral-most segments of the spinal cord (Fig. 2C). Of course, these axons were only detected by the presence of NADPH reaction product, and it is entirely possible that the axons of neurons in this cluster were both more numerous and more extensive in their innervation of the spinal cord (see Discussion). Rostrally projecting nitrergic axons also emanated from this cluster and could clearly be seen (Fig. 2D) as they coursed within the marginal zones between the cell groups.

The motor pattern during swimming provides four measurable features: (i) burst duration, which is the time for which a recorded ventral root motor pool is active in each cycle; (ii) cycle period, defined as the interval between consecutive bursts and measured from the onset of a burst in one cycle to the onset of the burst in the next cycle; (iii) rostrocaudal delay, the delay between consecutive bursts as the motor activity progresses down the body; and (iv) episode duration, which is the time for which the animal swims in response to a brief sensory stimulus (see Materials and methods). S-nitroso-N-acetylpenicillamine (SNAP), a stable analogue of endogenous S-nitroso compounds, was used to increase levels of NO in vivo with a view to assessing its effects on these four parameters of swimming. The bath application of 100–500 μmol l⁻¹ SNAP produced a significant (t-test, P<0.05) and reversible decrease in episode duration in approximately 90 % (13 of 14) of the experiments (Fig. 3A,B). There was also a profound lengthening of cycle period in approximately 80 % (11 of 14) of experiments in the presence of SNAP (Fig. 3C,D). There was, however, no consistent effect observed on either burst duration or rostrocaudal delay. To investigate whether the effect was specific to NO, experiments were conducted with NO-depleted SNAP, the inactive isomer of SNAP, N-acetylpenicillamine (NAP), and its vehicle DMSO. Bath application of NO-depleted SNAP (50–100 μmol l⁻¹), NAP dissolved in DMSO (100–500 μmol l⁻¹) or DMSO alone (0.01–1 %) produced no significant reduction in episode duration (t-test, P>0.05). The effect on cycle period in each case was highly variable. NO-depleted SNAP decreased cycle period in 40 % (2 of 5) of experiments, increased cycle period in 25 % (1 of 5) of experiments and had no significant effect in 40 % (2 of 5) of experiments. In approximately 27 % (3 of 11) of experiments, NAP decreased cycle period, in approximately 36 % (4 of 11) it increased it, and in approximately 27 % (3 of 11) there was no significant effect (t-test, P>0.05). DMSO decreased cycle period in 40 % (4 of 10) of experiments, increased it in 30 % (3 of 10) and had no significant effect in 30 % (3 of 10; t-test, P>0.05).

A response to the exogenous application of a NO donor cannot alone prove that endogenous NO plays a role in the tadpole nervous system. To demonstrate this, NOS inhibitors can be used to determine whether a reduction in endogenous NO produces any change in fictive swimming activity. To this end, a series of experiments was conducted using both broad-spectrum (L-NAME) and more selective neuronal (L-NNA) NOS inhibitors. The bath application of either L-NAME or L-NNA (0.5–5 mmol l⁻¹) resulted in a significant (t-test, P<0.05) increase in episode duration in approximately 65 % (7 of 11) of L-NAME treatments and approximately 55 % (5 of 9) of L-NNA treatments (Fig. 4A,B). This was also associated with a decrease in cycle period in approximately 70 % (8 of 11) of L-NAME treatments and approximately 75 % (7 of 9) of L-NNA treatments (Fig. 4C,D). The most consistent effects on episode duration occurred when control episodes were relatively short. To control for the specificity of NOS inhibition, experiments using the inactive form of L-NAME, D-NAME (5–10 mmol l⁻¹), were conducted. There was no significant increase in episode duration (t-test, P>0.05), and the effect on cycle period was inconsistent, with 50 % (3 of 6) increasing, approximately 16 % (1 of 6) decreasing and approximately 33 % (2 of 6) showing no significant effect (t-test, P>0.05). As one would expect, therefore, the inhibition of endogenous NOS produced precisely the opposite effects to those of exogenously applied SNAP on most parameters of swimming.

The main conclusion of this series of anatomical and electrophysiological experiments is that the free radical gas NO is generated by brainstem neurons and exerts a profound influence upon the network of spinal neurons responsible for rhythmic swimming activity. We believe this to be the first demonstration of a role for nitrergic transmission in the function of a rhythmic locomotor network.

The pattern of staining obtained using NADPH diaphorase histochemistry, at least at this early stage of development, is restricted to specific groups of neurons located in the brainstem. There is strong precedence to suggest that this staining procedure labels neurons that synthesise NO using one or more isoforms of NOS (Hope et al., 1991). However, confirmation that the NADPH-diaphorase-positive cells in X. laevis express a particular isoform of NOS will require further immunocytochemical studies. Nevertheless, the present study has revealed labelling in precisely the same regions in which NOS is normally found, for example in the walls of blood vessels (not illustrated). Moreover, a range of vertebrate neurons in similar locations in the CNS to those we have described in the tadpole brainstem have been shown to be nitrergic (Schober et al., 1994; Bruning and Mayer, 1996; Vincent and Kimura, 1992).

The structure and location of the neurons that stained in X. laevis larvae CNS indicate that they belong to particular subsets of brainstem neurons that may have been described previously and that may co-localise other transmitters. For example, by far the largest population is in the hindbrain, and at least some of these neurons showed positive labelling in axons that extended to the rostral spinal cord. Presumably, therefore, these neurons are reticulospinal. A variety of neuron clusters in this region have been described using both standard anatomical tracing methods, such as horseradish peroxidase labelling and immunocytochemistry using antibodies against different neurotransmitters (Roberts and Alford, 1986; Roberts et al., 1987). These include the most rostral members of a population of descending excitatory interneurons that also extend along most of the length of the spinal cord. These interneurons have ipsilaterally projecting axons and are known to generate the excitatory drive for swimming by activating glutamate receptors on spinal motor and interneurons (Roberts and Alford, 1986). Commissural interneurons, which are glycinergic, provide the reciprocal mid-cycle inhibition during swimming (Dale et al., 1986). Like the descending interneurons, the commissural interneurons also extend rostrally from the spinal cord into the caudal hindbrain. However, neither of these two populations was labelled in the spinal cord using the NADPH-diaphorase technique, and it seems unlikely that a subpopulation would selectively express NOS only in the caudal hindbrain. Mid-hindbrain reticulospinal (mhr) interneurons also occur in the region of NADPH-diaphorase staining described in the present paper. They have been identified in X. laevis embryos using antibodies directed against γ-aminobutyric acid (GABA) (Roberts et al., 1987) and they participate in a descending GABAergic ‘stopping’ response (Boothby and Roberts, 1992). The mhr neurons are activated following stimulation of the rostral cement gland and release GABA in the spinal cord to terminate swimming when the embryo encounters an obstacle in its path. A proportion of mhr neurons have decussating axons, like some of the NADPH-diaphorase-positive neurons we have described (Figs 1Bi, 3B). Other hindbrain reticulospinal neurons are likely to have excitatory functions in the spinal cord. However, NO often co-localises with inhibitory transmitters (Vincent, 1995) and has been reported to evoke GABA release (Ohkuma et al., 1995). While we have yet to determine the transmitter phenotype of the reticular NADPH neurons, it seems plausible that they might use GABA or perhaps glutamate as a fast descending transmitter and NO as a cotransmitter.

The next most rostral cluster of NADPH-diaphorase-positive neurons is located ventrally, close to the mid-brain/hindbrain boundary and extends caudally for approximately 150 μm. This is an area of the brainstem that substantially overlaps a region that is known to be occupied by serotonergic neurons of the raphe nucleus (van Mier et al., 1986; Sillar et al., 1995b). The serotonergic neurons and the NADPH-positive neurons are remarkably similar in overall morphology; the cell bodies are arranged as a column that extends dorsally from the ventral commissure close to the boundary of the neurocoel (Fig. 1Bii). Primary neurites then project ventro-medially to the lateral margins of the brainstem, where they branch into a dense plexus of dendritic processes. Although requiring confirmation, our present proposal, based on the location of this group and their general anatomical features, is that they correspond to serotonergic neurons of the raphe nucleus. Since many raphe neurons have axonal projections that innervate the spinal cord, it is conceivable that NO released from these neurons is somehow involved in the NO-induced modulation of swimming that we have described. However, the raphespinal projections in X. laevis larvae are responsible for enhancing the intensity and duration of rhythmic motor bursts during fictive swimming, the opposite effect to that produced by exogenous NO.

The most rostrally located cluster of NADPH-diaphorase-positive neurons is more difficult to correlate with previously described groups of brainstem neurons. The neurons are positioned more dorsally and laterally than the other groups, very close to the boundary between the mid-and the hindbrain. They are fewer in number (approximately 20 cell bodies per side) and have a very distinctive anatomy. These neurons are similar in morphology to the catecholaminergic neurons of the nucleus tractus solitarii identified in the larval stage of the amphibian Pleurodeles waltlii (Gonzalez et al., 1995). Whether these diaphorase-positive neurons are also tyrosine-hydroxylase-immunoreactive in X. laevis has yet to be confirmed.

We have provided the first evidence for intrinsic regulation of vertebrate locomotor rhythm generation by NO. This evidence is based on the fact that the NO donor SNAP and the NOS inhibitors L-NAME and L-NNA strongly affect the rhythmic motor output for swimming. While it is possible that the manipulations of NO influence rhythmic output indirectly, via effects on the cardiovascular system, we think this unlikely. X. laevis larvae have a rudimentary vascular system at this early stage of development. While blood vessels did display NADPH-diaphorase staining, there were none detected within or close enough to the CNS to act as the source of NO. Our data instead suggest that endogenous release of NO from one or more subsets of the NADPH-diaphorase-positive brainstem neurons exerts modulatory control over the spinal locomotor circuitry. We have argued above that the most likely group is the most numerous and most caudally located cluster of reticular neurons. In principle at least, NO released from brainstem neurons could affect spinal rhythm generation by acting directly on spinal neurons (or their synaptic interconnections) or by altering the descending drive to the spinal swimming circuitry. At present, we cannot distinguish between these two possibilities, but both seem likely. Since the main target for NO is soluble guanylate cyclase, we presume that the NO-induced modulation of swimming is accomplished by cGMP-dependent facilitation of synaptic transmission, in keeping with the results of previous studies on the central nervous effects of NO (for a review, see Schuman and Madison, 1994). Increased levels of extracellular NO following application of SNAP lead to a reduction in the duration of swimming episodes and a diminution in the intensity and frequency of swimming, consistent with either an increase in inhibitory synaptic processes or a decrease in the excitatory drive for swimming. Of these two non-mutually exclusive possibilities, we favour the former, that NO slows swimming by increasing inhibitory synaptic transmission. This is because inhibitory synapses in the swimming system appear to be much more variable and open to modulation than excitatory synapses. For example, the amine noradrenaline has a very similar effect to NO upon swimming in producing slow, weak activity. This is accomplished by the presynaptic facilitation of glycinergic transmission (McDearmid et al., 1997), although noradrenaline can also facilitate GABAergic transmission (McDearmid, 1998; J. McDearmid and K. T. Sillar, unpublished observations). The GABA synapses in X. laevis tadpoles are also presynaptically facilitated by a neuroactive steroid (5β3α; Reith and Sillar, 1997), which again weakens and slows down the swimming rhythm. In future experiments, we intend to test the hypothesis that NO affects swimming by facilitating descending GABAergic inhibition at sites in the brainstem.

We thank John Simmers and Simon Merrywest for critically reading the manuscript. We also thank Mrs A.-M. Woolston, Miss J. Inglis and Miss G. Morgan for their contributions to the initial stages of this project. This work was supported by the BBSRC (UK) and The Wellcome Trust. D.L.M. is an earmarked BBSRC research student.