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• W1574030158 abstract "Key points Rhythmic activity is a feature of many regions of the CNS, but surprisingly, no precise pathway for the initiation of locomotion is yet known for any vertebrate. Using a well-proven, simple vertebrate system, the hatchling Xenopus tadpole, we report here our discovery of a detailed neuron-by-neuron pathway for initiating locomotor activity on one side of the CNS. We describe a small population of brainstem neurons (trigeminal interneurons) that are directly excited by trigeminal sensory neurons when the head skin is touched. These neurons amplify brief sensory signals and relay excitation to an electrically coupled population of hindbrain reticulospinal neurons (descending interneurons), whose firing initiates swimming locomotion. We believe that our discovery of this primitive, direct pathway, which appears simpler than initiation pathways so far defined in invertebrates, is of evolutionary interest and raises the important possibility of equivalent pathways in more complex vertebrates including mammals. Abstract While we understand how stimuli evoke sudden, ballistic escape responses, like fish fast-starts, a precise pathway from sensory stimulation to the initiation of rhythmic locomotion has not been defined for any vertebrate. We have now asked how head skin stimuli evoke swimming in hatchling frog tadpoles. Whole-cell recordings and dye filling revealed a nucleus of ∼20 trigeminal interneurons (tINs) in the hindbrain, at the level of the auditory nerve, with long, ipsilateral, descending axons. Stimulation of touch-sensitive trigeminal afferents with receptive fields anywhere on the head evoked large, monosynaptic EPSPs (∼5–20 mV) in tINs, at mixed AMPAR/NMDAR synapses. Following stimuli sufficient to elicit swimming, tINs fired up to six spikes, starting 4–8 ms after the stimulus. Paired whole-cell recordings showed that tINs produce small (∼2–6 mV), monosynaptic, glutamatergic EPSPs in the hindbrain reticulospinal neurons (descending interneurons, dINs) that drive swimming. Modelling suggested that summation of EPSPs from 18–24 tINs can make 20–50% of dINs fire. We conclude that: brief activity in a few sensory afferents is amplified by recruitment of many tINs; these relay summating excitation to hindbrain reticulospinal dINs; dIN firing then initiates activity for swimming on the stimulated side. During fictive swimming, tINs are depolarised and receive rhythmic inhibition but do not fire. Our recordings demonstrate a neuron-by-neuron pathway from head skin afferents to the reticulospinal neurons and motoneurons that drive locomotion in a vertebrate. This direct pathway, which has an important amplifier function, implies a simple origin for the complex routes to initiate locomotion in higher vertebrates. Rhythmic activity is a feature of many regions of the CNS and there is a huge experimental and modelling literature about how rhythms are generated. Considerably less attention has been paid to the question of how episodic rhythms are normally turned on by sensory stimuli. While we understand how stimuli evoke sudden, ballistic escape responses (fish Mauthner-cell: Korn & Faber, 2005; crayfish giants: Edwards et al. 1999), the precise neuron-by-neuron pathway from sensory stimulation to the initiation of rhythmic locomotion has not been described in any vertebrate. The widely accepted hypothesis, arising particularly from brain stimulation experiments on decerebrate cats in the 1970s (Orlovsky, 1970; reviewed in Orlovsky et al. 1999), is that vertebrate locomotion begins, either spontaneously or following sensory stimulation, when descending brainstem reticulospinal pathways become active and turn on spinal central pattern generators (CPGs). These descending brainstem pathways are activated by mesencephalic and diencephalic locomotor regions (Jordan et al. 2008). The prevailing view is that the basal ganglia exert tonic inhibition on the locomotor regions and that goal-oriented locomotion can be selected by input from regions such as the cortex or via the thalamus through disinhibition of this suppression by the basal ganglia (Grillner et al. 2008). Evidence continues to accumulate suggesting that some of these regions have counterparts in all vertebrates (Stephenson-Jones et al. 2011). The organisation and operation of these systems in mammals are highly complex and remain poorly understood. However, precision in defining motor regions, centres and neuronal elements of locomotion-initiating pathways increases significantly as we move to simpler vertebrate species (lamprey: Dubuc et al. 2008; zebrafish: Kyriakatos et al. 2011). One way to initiate locomotion in vertebrates is by head stimulation. The decerebrate mammal locomotor system can be activated by electrical stimulation of the trigeminal nerve or its peripheral receptive field (cat: Aoki & Mori, 1981; Noga et al. 1988; Beresovskii & Bayev, 1991; rat: Vinay et al. 1995). In the lamprey, mechanical stimulation of the head elicits swimming locomotion, and indirect evidence has revealed possible ‘relay’ neurons placed between trigeminal sensory afferents and the descending, reticulospinal neurons which activate swimming (Viana Di Prisco et al. 2005). We study another simple vertebrate with a well-defined motor system, the hatchling frog tadpole, where head skin touch can initiate swimming (Boothby & Roberts, 1995). This paper will define a simple, primitive, sensory pathway from the head skin that excites the descending reticulospinal neurons driving locomotion. To uncover neuronal pathways initiating rhythmic activity following head skin stimulation, the hatchling Xenopus tadpole offers a unique advantage: the excitatory reticulospinal neurons (dINs) which fire on every cycle and drive firing in all other neurons active during swimming have been characterised both anatomically and physiologically (Li et al. 2006, 2010; Soffe et al. 2009). Initiation of swimming requires the activation of these neurons. Another advantage is that the properties of trigeminal sensory neurons innervating the head skin, and firing briefly to touch, have also been described (Roberts, 1980). Such touch stimuli to the head often evoke swimming where the first flexion is towards the stimulated side (Boothby & Roberts, 1995) and occurs within some 20–30 ms. Here we investigate how brief firing in trigeminal touch afferents leads to the excitation of the reticulospinal dINs that drive spinal cord swim neurons and generate the swimming rhythm. We have characterised the interneurons in a previously unknown trigeminal sensory pathway nucleus. This allows us to propose a pathway by which a head skin stimulus can initiate activity for swimming on the same side as the stimulus. We trace this pathway, step-by-step, from the trigeminal touch-sensitive neurons, through the trigeminal relay nucleus neurons, to the driver reticulospinal interneurons, and finally to the motoneurons which produce the first flexion in swimming. Procedures for obtaining developmental stage 37/38 (Nieuwkoop & Faber, 1956) hatchling Xenopus laevis (Daudin) tadpoles comply with UK Home Office regulations. All unregulated experiments on the tadpoles have been approved following local ethical committee review. Tadpoles from a captive breeding colony (13 h:11 h light regime) were briefly anaesthetised in 0.1% MS-222 (3-aminobenzoic acid ester). The dorsal fin was cut open to allow direct access for immobilisation using 10 μmα-bungarotoxin in saline (115 mm NaCl, 3 mm KCl, 2 mm CaCl2, 2.4 mm NaHCO3, 1 mm MgCl2, 10 mm Hepes adjusted to pH 7.4 with NaOH) for 20–30 min. The animals were then pinned to a rotatable Sylgard-coated platform with tungsten wire pins through the notochord. The hindbrain was exposed and its roof removed using a finely etched tungsten pin and fine forceps. Parts of the inner surface of the hindbrain were removed to reveal neuron somata. The animals were then repinned in a small recording chamber (∼1 ml) with saline perfusion, and neuron somata could be seen using a ×40 water immersion lens on an upright Nikon (Tokyo, Japan) E600FN microscope using LED illumination (cf. Safronov et al. 2007). Drops of pharmacological antagonists were added to a 100 μl chamber upstream to the record-ing chamber. The concentrations for 1,2,3,4-tetrahydro-6-nitro-2,3-dioxo-benzo[f]quinoxaline-7-sulfonamide (NBQX) used in this study were 5 μm, 10 μm or 50 μm. NBQX was obtained from Tocris Cookson (Bristol, UK), horseradish peroxidase (HRP) from Boehringer Ingelheim (Bracknell, UK) and all other chemicals from Sigma (Poole, UK). Experiments were performed at 18–22°C. Fictive swimming was initiated by stimulation of the head skin on the right side with either a touch stimulus or with a brief (0.1 ms) current pulse using a fire-polished glass suction electrode with a tip opening of 40–60 μm filled with saline. This excites the peripheral processes of sensory neurons which enter the brain via the trigeminal nerve (Roberts, 1980). To record fictive swimming activity in immobilised tadpoles, similar glass suction electrodes were placed on the right side at the intermyotomal cleft of myotomes 2/3 or 3/4. Extracellular hindbrain and trigeminal unit recordings were made with a similar electrode placed either on the inside of the hindbrain opened at the level of the 2nd to 4th rhombomere or on the trigeminal ganglion. The signals were amplified by a differential amplifier (gain: 1000, SOBS, University of Bristol, UK) and filtered (low: 30 Hz, high: 1 kHz). Whole-cell current clamp recordings were made with an Axoclamp 2B (Axon Instruments Inc., Union City, CA, USA) in bridge mode, filtered (at 30 kHz) and digitised (sampling rate: 10 or 20 kHz, ADC resolution: 16 bit) using a CED Power1401mkII interface (Cambridge Electronic Design Ltd, Cambridge, UK) and Signal 3 and 4 software (CED). Patch pipettes were filled with 0.1% neurobiotin in intracellular solution (100 mm potassium gluconate, 2 mm MgCl2, 10 mm EGTA, 10 mm HEPES, 3 mm Na2ATP, 0.5 mm NaGTP adjusted to pH 7.3 with KOH) and had resistances of 5–15 MΩ. The liquid junction potential of this solution was measured as +11 mV, but for better comparison with previous data, the values have not been corrected for this. Neuron anatomy and connections were checked with a standard avidin–biotin technique using diaminobenzidine as chromogen (see Li et al. 2001). Briefly, after experiments the animals were left in the saline for 20–25 min allowing the dye to diffuse throughout the whole cell. The animals were fixed in 2% glutaraldehyde for at least 2 h, washed in phosphate-buffered saline (PBS, 120 mm NaCl in 0.1 m phosphate buffer) and again in PBS containing 1% Triton X-100 before labelling with ExtrAvidin (1:200) for 3 h. The animals were then washed again in 0.1 m PBS, the neurons stained in 0.8% DAB in phosphate buffer, and then washed again in buffer containing 0.03% H2O2 for 5 min each. After washing in tap water, the brain and spinal cord were exposed and the specimen dehydrated in an ascending alcohol series and cleared in methyl benzoate. Specimens were mounted ventral side down between coverslips with DePeX on a reversible aluminium microscope slide. Once mounted, the hindbrain lay opened along the dorsal midline like a book. Neurons were observed using a ×100 oil immersion lens and traced using a drawing tube. Photographs of the brain were obtained using a ×20 objective lens and a CCD camera (DeltaPix DP 200, Maalov, Denmark) and arranged in Adobe Photoshop (Adobe Systems Inc., San Jose, CA, USA). Measurements were made from the scale drawings and corrected for shrinkage during processing (×1.28). The position of the recorded cell bodies was measured during recording using a micrometer connected to the microscope. It is possible that cell bodies could move and change shape slightly during recording and processing. To identify and measure the whole population of tINs, their somata were labelled retrogradely from their axons in the spinal cord. For this the animals were immobilised, mounted on a rotatable Sylgard platform and the spinal cord exposed. The left half of the spinal cord was severed just caudal to the hindbrain to avoid contralateral labelling. A vaseline well was built around the spinal cord approximately at the middle of the animal, filled with intracellular solution containing 0.1% neurobiotin and the spinal cord cut to allow the dye to enter the axons. The neurobiotin solution was removed after 30–45 min and the animal fixed and processed as above. In a few additional examples, HRP was made up in concentrated aqueous solution and dried onto the tip of an etched tungsten needle. One side of the spinal cord was then crushed between a clean needle placed on the midline and an HRP-coated needle placed on the lateral surface of the cord. After 2 min the embryo was transferred to 75% physiological saline that was further diluted as the wounds healed, fixed after 4–6 h and processed as above. Transverse sections of the hindbrain were obtained by embedding the processed animals in wax and cutting 10 μm-thick slices on a microtome. Data analysis was performed using routines purpose-written for Minitab. The latencies of postsynaptic potentials (EPSPs and IPSPs) or the first action potential were measured as the time between the peak of the presynaptic action potential or stimulus and the onset of the PSP (or the peak of the postsynaptic action potential) and averaged for each cell/connection type. Additionally for EPSPs the 10–90% rise time, the time to peak and the width at half-peak-amplitude were measured. In patch-pipette recordings from pairs of neurons, current-induced spikes in one cell caused a small cross-talk artefact in the other (see Li et al. 2002). These artefacts were removed from some recordings in which they obscured the onset of small EPSPs evoked by current-induced spikes. To do this, either averaged records of EPSP failures (artefact alone) or smoothed, differentiated spike traces were subtracted from EPSP records (EPSP plus artefact). The recorded traces, photographs and drawings were imported into and the figures finally arranged using Corel Draw (Corel Corp., Ottawa, ON, Canada). Statistical tests used are stated in the Results. The outcome of tests was regarded as significant where P < 0.05. All measurements are expressed as mean ± standard deviation (SD). Briefly touching or stroking the head of the tadpole initiates the swimming rhythm (at 10–25 Hz; Roberts, 1990) and will excite trigeminal mechanosensory neurons innervating the head skin (Roberts, 1980). Counts of axon numbers in the ophthalmic and maxilliary trigeminal nerves and measures of the receptive field areas of individual sensory neurons suggest that there are 50–80 touch-sensitive trigeminal afferents on each side of the head (Hayes & Roberts, 1983). The axons of these neurons project from the trigeminal ganglia to descend as a compact tract, some 20 μm in dorso-ventral extent, through the ipsilateral hindbrain. Some axons reach as far as the rostral spinal cord. To search for groups of sensory interneurons which might be contacted by these descending trigeminal axons, we used neurobiotin to retrogradely label hindbrain neurons projecting into the right side of the spinal cord in eight tadpoles. We also examined earlier material retrogradely labelled with HRP. Many ipsi- and contralateral neurons were seen, including a consistent ipsilateral group of 14–23 neurons in a mid-dorsoventral position lying in the region of cranial nerve VIII in rhombomeres 2–4 (Fig. 1A). Retrograde filling and extracellular recordings of possible trigeminal interneurons in the hindbrain and trigeminal afferents A, diagram of the CNS in dorsal view to show the main regions, place where left spinal cord was cut and site of neurobiotin application (BF). Aa, inset (photograph) shows a cluster of neuron somata in the hindbrain (tIN, ellipse) revealed by backfilling their axons from the ipsilateral spinal cord (rostral left). Ab, photograph of the same region in a different animal in lateral view (rostral left, dorsal up). Ac, composite drawing of 11 transverse sections of the hindbrain around the otic capsule (dorsal up) showing the labelled neurons on the right side (ellipse). BF, direction of backfill; fb, forebrain; hb, hindbrain; mb, midbrain; m, muscles; oc, otic capsule; sc, spinal cord; tg, trigeminal ganglion, r1–8, rhombomeres. B, diagram of set-up to stimulate head skin and record from hindbrain neurons. Ba, record shows firing to head skin stimulation (stroke with hair at arrow). Bb, unit in Ba shows no firing to tail skin stimulation. C, diagram for head skin stimulation and trigeminal neuron recording. Ca, trigeminal ganglion recordings show a single unit at threshold in response to a short electrical stimulus (arrowhead). Cb, a stronger stimulus evokes firing in more units. Asterisks mark the stimulus artefacts. Extracellular recordings were therefore made from the right side of the hindbrain caudal to the trigeminal nerve entry in rhombomeres 2–4 in three tadpoles to search for activity during natural sensory stimulation of the head skin. Strokes with a fine hair (10 μm diameter) evoked unit firing (Fig. 1Ba). These units fired to strokes anywhere on the same side of the head and, unlike the primary afferent neurons (Roberts, 1980), did not have local receptive fields. In some cases activity in these units was followed by swimming. They did not fire to strokes elsewhere on the body or tail (Fig. 1Bb), which were also sometimes sufficient to initiate swimming. When making whole-cell recordings from brain neurons, electrical stimulation of the skin has to be used as adjacent mechanical stimulation dislodges the electrodes. We therefore made extracellular recordings from trigeminal neurons in the ophthalmic ganglion of six tadpoles to test what response they gave to a 0.1 ms current pulse given to the head skin with a small electrode placed rostral and dorsal to the eye. As the stimulus increased to around the threshold to elicit swimming, a clear single unit appeared at 3.4 ± 0.5 ms (SDs for individual units ranged from 0.1 to 0.3, mean 0.2); with further increase, firing occurred earlier and other units appeared (between 2.9 ± 0.6 ms and 6.5 ± 1.5 ms); no units fired later than 9.3 ms or multiply (Fig. 1C). These recordings therefore suggest that a 0.1 ms current pulse to the head skin, of sufficient intensity to elicit swimming, excites the free nerve endings of a small number of sensory neurons, each of which only fires a single action potential. Since extracellular recordings had shown activity in the hindbrain caudal to the trigeminal nerve in response to head skin stimulation, we used whole-cell recordings to locate neurons in this region firing in response to a 0.1 ms current pulse to the skin on the same side of the head. Such neurons might form the first link in a pathway for the initiation of swimming (Fig. 2). We made whole-cell recordings from 37 neurons with somata in the 2nd to 4th rhombomeres. These neurons were silent at rest but were excited to fire by head skin stimulation. We will first define the anatomy revealed by neurobiotin-filling of this previously unknown type of trigeminal descending interneuron (tIN), excited by head skin stimulation. Anatomy of trigeminal interneurons (tINs) and responses to head skin stimulation A, diagram of the preparation viewed dorsally showing the head skin stimulating electrode (stim skin) on the right side, a trigeminal sensory neuron projecting into the brain and the area where tINs were recorded (purple ellipse; record tIN). Traces show tIN responses to skin stimuli (arrowhead) of increasing strength from black = no response, red = EPSP alone, blue = spike, to green = multiple or earlier spikes. Inset shows the morphology of the tIN viewed from the inside of the right side of the hindbrain (rostral left) with dendrites (d) and descending axon (a). The position of the soma (s) is shown in grey. B–H, further examples to show the range of tIN anatomy and responses; E includes a photograph of the filled tIN. Arrows in C, E, F, G and H mark IPSPs (see text). The 37 recorded and neurobiotin-filled tINs showed a consistent set of morphological characteristics: (a) somata (diameter: 10–20 μm) are mainly unipolar and form a group in the hindbrain (range: 150–330 μm from mid-hindbrain border and 30–150 μm from midline; Fig. 3A, see also Fig. 1A); (b) dendrites range from simple (Fig. 2B) or compact (Fig. 2F and H) to relatively extensive (Fig. 2E, D and G) and extend into the region where they could receive en passant synaptic contacts from the descending axons of trigeminal sensory neurons (Hayes & Roberts, 1983); (c) a long ipsilateral descending axon projects either directly from the soma or from primary processes and initially is directed medio-ventrally before descending into the spinal cord (length: 1.75 ± 0.48 mm; Fig. 3B). Many axon lengths may be underestimates as staining faded and there was no distinct end-bulb (N= 24) or axons could not be traced to the end because the spinal cord was damaged during processing (N= 10). The lengths of the three axons with a clear end-bulb and thus filled to the end were 2.0, 2.0 and 2.7 mm. The axons had uneven diameters, were relatively thick for the tadpole (up to ∼1 μm), and sometimes had short side branches (Fig. 2B, F and G). tIN soma positions and axon lengths A, positions of the somata of recorded tINs (in dotted outline in rhombomeres 2–4) and dINs (close to midline in rhombomeres 3–7). B, individual tIN soma positions (filled circles) and descending axons. Some axons were broken (capped lines) or faded out but 3 were completely filled (bold lines). In all whole cell experiments examining responses of tINs to head skin stimulation, we also recorded ventral root activity so that we could determine the threshold stimulus current required to initiate swimming (threshold = 100%). All recorded tINs, after stimulation of the head skin at swimming threshold, received a large, short latency EPSP followed by a spike. The effect of stimulus intensity was analysed in 21 of these. The intensity to evoke an EPSP without a spike was 86 ± 8% of the swim threshold, and when the stimulus was at 94 ± 6%, all tINs fired a spike. Spiking in individual tINs could thus occur without subsequent swimming (18/21); however, in only 4/21 tIN recordings did swimming occasionally start without a spike in the recorded tIN. Spikes in tINs reached their peak 5.4 ± 1.6 ms (N= 34 tINs) after the stimulus. Increasing the stimulus strength made tIN spiking more reliable; the first spike could become earlier (by up to 2 ms) and multiple firing often occurred (starting from 5 ± 7% above swim threshold). With further increase of the stimulus, the burst became longer (up to 6 spikes, instantaneous frequency >250 Hz), but 10/34 tINs only ever fired once. The excitatory responses of tINs could also be followed by IPSPs (arrows, Fig. 2). The large EPSPs produced in tINs by trigeminal afferents were analysed when weaker head skin stimuli did not evoke a spike (Fig. 4B). For each of 23 tINs, 5–15 EPSPs were measured: latencies were 4.4 ± 0.5 ms, amplitudes 14.2 ± 5.4 mV, rise times from 10 to 90% amplitude 2.5 ± 1.0 ms, time to peak 8.7 ± 1.8 ms after the stimulus, and duration at 50% amplitude 17.8 ± 10.8 ms. In 13 tINs where the stimulus current was held constant, the standard deviations of latencies of five EPSPs in each tIN ranged from 0.1 to 0.3 ms (mean 0.2). The tIN EPSP latencies are therefore only 1 ms longer than the latencies of afferent spikes recorded from the trigeminal ganglion (see above) and have equally small standard deviations. We conclude that tINs are directly, monosynaptically excited at synapses from trigeminal sensory axons. Effects of NBQX on EPSPs in tINs evoked by head skin stimulation A, diagram of the preparation in dorsal view showing the position of the stimulating and recording electrodes. B, a current pulse to the head skin (open arrowhead) leads to a tIN EPSP with a short delay. C, a stronger stimulus (black arrowhead) leads to spiking. D, NBQX perfusion (50 μm) blocks the spike and reveals a smaller slow rise and fall EPSP. E, after 20 min washing, the same stimulus evokes spiking again. Overlay of three traces each, dotted lines show resting membrane potential. F, histogram showing the tIN EPSP amplitude before (black bars) and after NBQX perfusion (open bars). Data are from 5 EPSPs from 15 tINs for each treatment, bin size: 0.5 mV. To investigate the receptors activated during tIN EPSPs we bath applied the glutamate AMPA receptor (AMPAR) antagonist NBQX (5–50 μm). A rapidly rising, large first component was blocked by NBQX (N= 18; Fig. 4D) and is therefore AMPAR mediated. A second, smaller component was only visible in NBQX, had the same latency (4.1 ± 0.6 ms; Mann–Whitney test: U= 129, n1,2= 23,15, P= 0.1983), but was smaller (peak amplitude: 4.7 ± 1.7 mV; U= 5, n1,2= 23,15, P < 0.0001), slower rising (time to peak 16.6 ± 7.3 ms; 10–90% amplitude in 10.4 ± 6.4 ms; U= 21, n1,2= 23,15, P < 0.0001) and longer lasting (width at 50% amplitude: 54.4 ± 27.5 ms; U= 26, n1,2= 23,15, P < 0.0001; N= 15 tINs) and was therefore probably NMDAR mediated. On the basis of our extracellular recordings, we suspected that individual trigeminal afferents would contact multiple tINs and conversely that individual tINs would receive excitatory synapses from many head skin afferents. To get direct evidence for the first of these, we made simultaneous recordings from six pairs of tINs. When a skin stimulus just reached the level to evoke an EPSP in one tIN (presumed to be the threshold level to activate a single trigeminal afferent; see Fig. 1Ca), an EPSP always appeared in the second tIN of the pair. Since different tINs had slight differences in spike threshold, the same weak stimulus could elicit a spike in only one despite eliciting an EPSP in both. As well as showing that single afferents do apparently excite multiple tINs, these paired recordings also showed that tINs do not excite or inhibit each other and show no evidence of electrical coupling. Lastly, excitation of multiple tINs by each sensory afferent was indicated by the observation that we could always find several tINs responding to stimuli at a single site in individual animals. We next investigated the range of afferent input to individual tINs. Trigeminal sensory afferents have rather localised receptive fields in the head skin (area: ∼0.015 mm2, diameter: ∼200 μm, Roberts, 1980). To test whether tINs also have local receptive areas, one stimulating electrode was placed on the skin rostral and dorsal to the eye in the area innervated by the ophthalmic branch of the trigeminal nerve, and another was placed caudal and ventral to the eye in the area innervated by the maxillary branch, at a spacing of >500 μm. EPSPs and spikes were evoked by just supra-threshold stimuli at either location in four recorded tINs showing that they do not have local receptive fields but receive input from widespread head skin afferents. In contrast, tINs did not spike in response to current pulse stimuli given to the trunk or tail skin on the same side (N= 18), or to stimuli at any location on the other side of the animal (N= 19). They were also not activated when stronger stimuli at the same locations evoked a skin impulse which propagates from cell to cell through the skin and can start swimming (N= 7; Roberts, 1969). Finally, tINs did not fire as a result of dimming the illumination which excites the pineal eye and can also start swimming (N= 35; Foster & Roberts, 1982). We therefore conclude: that tINs respond only to head skin stimulation on the same side; that individual trigeminal afferents contact multiple tINs; and that individual tINs receive synapses from widespread trigeminal afferents. As well as being excited by head skin stimulation, tINs can also receive early inhibition. IPSPs were seen with or without firing in an individual tIN (Figs 2 and 5) and with or without subsequent swimming activity, but never without previous EPSPs. Inhibition may be obscured by tIN firing, but clearly occurred at short latency following a stimulus in cases where the tIN did not fire (mean latency of earliest IPSPs: 9.7 ± 3.0 ms, minimum: 4.5 ms; N= 27 tINs). These short latency IPSPs suggest there may be some direct connections from trigeminal afferents onto currently unknown inhibitory trigeminal interneurons. Such inhibition could limit the occurrence or duration of tIN firing following a head skin stimulus. Inhibition of tINs after head skin stimulation A tIN receives IPSPs (arrows) following head skin stimulation (arrowhead). IPSPs also occur following a weaker stimulus (open arrowhead), insufficient to evoke a spike. These IPSPs occur earlier and are not masked by a spike. Cellular parameters were measured for 27 tINs. Resting membrane potentials were −55 ± 6 mV and input resistances 459 ± 173 MΩ (measured from the I–V regression lines fitted to voltage responses to injected positive and negative current steps around the resting potential; Fig. 6A and B). Measured relative to 0 mV, action potential peak was 22 ± 5 mV, firing threshold estimated from the inflexion in the upstroke of the spike was −16 ± 4 mV, and spike width at half-amplitude was 1.2 ± 0.3 mV. Spikes had a pronounced after-hyperpolarisation (trough: −39 ± 6 mV at 2.5 ± 0.5 ms after spike peak; Fig. 6C). When depolarising current (200 ms) was injected just above the threshold to elicit firing, tINs fired one or a few spikes. When the current was increased, all tINs fired repetitively and spiking frequency increased up to around 100–200 Hz (N= 27 tINs; Fig. 6D–F). During current-induced repetitive firing, instantaneous frequency dropped only slightly, spike amplitude decreased and spike width inc" @default.
• W1574030158 created "2016-06-24" @default.
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• W1574030158 date "2012-04-11" @default.
• W1574030158 modified "2023-09-27" @default.
• W1574030158 title "The role of a trigeminal sensory nucleus in the initiation of locomotion" @default.
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• W1574030158 doi "https://doi.org/10.1113/jphysiol.2012.227934" @default.
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