A. C. T. Takakura and T. S. Moreira contributed equally to this study.
Peripheral chemoreceptor inputs to retrotrapezoid nucleus (RTN) CO2-sensitive neurons in rats
Article first published online: 5 APR 2006
The Journal of Physiology
Volume 572, Issue 2, pages 503–523, April 2006
How to Cite
Takakura, A. C. T., Moreira, T. S., Colombari, E., West, G. H., Stornetta, R. L. and Guyenet, P. G. (2006), Peripheral chemoreceptor inputs to retrotrapezoid nucleus (RTN) CO2-sensitive neurons in rats. The Journal of Physiology, 572: 503–523. doi: 10.1113/jphysiol.2005.103788
- Issue published online: 5 APR 2006
- Article first published online: 5 APR 2006
- (Received 16 December 2005; accepted after revision 31 January 2006; first published online 2 February 2006)
The rat retrotrapezoid nucleus (RTN) contains pH-sensitive neurons that are putative central chemoreceptors. Here, we examined whether these neurons respond to peripheral chemoreceptor stimulation and whether the input is direct from the solitary tract nucleus (NTS) or indirect via the respiratory network. A dense neuronal projection from commissural NTS (commNTS) to RTN was revealed using the anterograde tracer biotinylated dextran amine (BDA). Within RTN, 51% of BDA-labelled axonal varicosities contained detectable levels of vesicular glutamate transporter-2 (VGLUT2) but only 5% contained glutamic acid decarboxylase-67 (GAD67). Awake rats were exposed to hypoxia (n= 6) or normoxia (n= 5) 1 week after injection of the retrograde tracer cholera toxin B (CTB) into RTN. Hypoxia-activated neurons were identified by the presence of Fos-immunoreactive nuclei. CommNTS neurons immunoreactive for both Fos and CTB were found only in hypoxia-treated rats. VGLUT2 mRNA was detected in 92 ± 13% of these neurons whereas only 12 ± 9% contained GAD67 mRNA. In urethane–chloralose-anaesthetized rats, bilateral inhibition of the RTN with muscimol eliminated the phrenic nerve discharge (PND) at rest, during hyperoxic hypercapnia (10% CO2), and during peripheral chemoreceptor stimulation (hypoxia and/or i.v. sodium cyanide, NaCN). RTN CO2-activated neurons were recorded extracellularly in anaesthetized intact or vagotomized rats. These neurons were strongly activated by hypoxia (10–15% O2; 30 s) or by NaCN. Hypoxia and NaCN were ineffective in rats with carotid chemoreceptor denervation. Bilateral injection of muscimol into the ventral respiratory column 1.5 mm caudal to RTN eliminated PND and the respiratory modulation of RTN neurons. Muscimol did not change the threshold and sensitivity of RTN neurons to hyperoxic hypercapnia nor their activation by peripheral chemoreceptor stimulation. In conclusion, RTN neurons respond to brain P presumably via their intrinsic chemosensitivity and to carotid chemoreceptor activation via a direct glutamatergic pathway from commNTS that bypasses the respiratory network. RTN neurons probably contribute a portion of the chemical drive to breathe.
The chemical drive to breathe relies on central chemoreceptors that detect brain extracellular fluid P via pH, and on carotid body chemoreceptors that respond to arterial P in a P- and glucose-dependent manner (Scheid et al. 2001; Feldman et al. 2003; Richerson, 2004; Putnam et al. 2004; Prabhakar & Peng, 2004; Bin-Jaliah et al. 2004). The way in which central and peripheral chemoreceptor information is integrated at the cellular and network levels is unclear except for the fact that the process occurs within the pontomedullary region (Feldman et al. 2003). The question is made even more complex by current uncertainties regarding the very nature of central chemoreceptors.
Although substances released from glia or non-neuronal cells have repeatedly been invoked to account for the sensitivity of brainstem neurons to extracellular fluid P (Erlichman et al. 1998; Gourine et al. 2005; Guyenet et al. 2005b), the dominant theory is that neuronal pH-sensitive conductances underlie central chemoreception (Loeschcke, 1982; Putnam et al. 2004; Richerson et al. 2005; Guyenet et al. 2005b). The proportion of brainstem neurons that respond to pH or Pin vitro varies from 15% to more than 90% depending on the brain region, the preparation and the criteria that is applied to define the cells as chemosensitive (Dean et al. 1989; Kawai et al. 1996; Richerson et al. 2001; Mulkey et al. 2004; Ritucci et al. 2005). In light of this evidence, central chemosensitivity could be viewed as an emergent property of the respiratory network that cannot be assigned to any particular component of the system. Yet, other evidence suggests that central chemoreception does rely on specialized neurons that drive a respiratory motor pattern generator that has no or inadequate sensitivity to pH on its own (Nattie, 2001; Feldman et al. 2003). Congenital central hypoventilation syndrome, a disease in which patients have absent or greatly diminished central hypercapnic ventilatory chemosensitivity, argues in favour of the existence of central neurons specialized for chemoreception (Spengler et al. 2001; Gaultier & Gallego, 2005). The highly differentiated responsiveness of brainstem neurons to hypercapnia in vivo provides further evidence (Guyenet et al. 2005b).
The retrotrapezoid nucleus (RTN) contains very superficial propriobulbar neurons that have properties consistent with such specialized chemoreceptors (Smith et al. 1989; Ellenberger & Feldman, 1990; Nattie et al. 1991; Cream et al. 2002; Mulkey et al. 2004; Ritucci et al. 2005; Guyenet et al. 2005a). Their response to P is far greater than that of surrounding cells, in vivo and in slices (Mulkey et al. 2004; Ritucci et al. 2005; Guyenet et al. 2005a). These CO2-responsive cells are glutamatergic and have axonal projections that are anatomically appropriate to be driving the respiratory network (Mulkey et al. 2004).
The present study is designed to test whether RTN is an important site of integration between central and peripheral chemoreception. Our working hypothesis is that the intrinsic response of RTN neurons to pH is enhanced by excitatory inputs from peripheral chemoreceptors and that the resulting activity of these neurons could be encoding some of the chemical drive to breathe.
The results demonstrate that RTN neurons receive a strong excitatory input from carotid chemoreceptor afferents that bypasses the respiratory network and probably involves a single intervening glutamatergic neuron located in the nucleus of the solitary tract (NTS). Based on this and prior evidence, we conclude that RTN neurons have the capability of detecting brain P directly and that they also respond to arterial blood gas composition via very direct neural inputs from peripheral chemoreceptors. RTN neurons could therefore be a source of integrated chemical drive to some aspect of the respiratory circuitry.
The experiments were performed on a total of 64 male Sprague-Dawley rats (Taconic; Germantown, NY, USA) weighing 250–350 g. Fifty rats were used in electrophysiological experiments, the rest for anatomy. Procedures were in accordance with NIH Animal Care and Use Guidelines and were approved by the University of Virginia's Animal Care and Use Committee.
Surgery and anaesthesia
General anaesthesia was induced with 5% halothane in 100% oxygen. The rats received a tracheostomy and artificial ventilation with 1.4–1.5% halothane in 100% oxygen was maintained throughout surgery. All rats were subjected to the following previously described surgical procedures: femoral artery cannulation for arterial pressure (AP) measurement, bladder cannulation to ease urination, femoral vein cannulation for administration of fluids and drugs, removal of the occipital plate to insert a recording electrode into the medulla oblongata via a dorsal transcerebellar approach, and skin incision over the lower jaw for placement of a bipolar stimulating electrode next to the mandibular branch of the facial nerve (Guyenet et al. 2005a). The phrenic nerve was accessed by a dorsolateral approach after retraction of the right shoulder blade. A bilateral vagotomy in the neck was performed in 11 rats. Six intact rats were additionally subjected to bilateral carotid body denervation.
Upon completion of surgical procedures, halothane was replaced by a mixture of urethane (0.5 g kg−1) and α-chloralose (60 mg kg−1) slowly administered i.v. All rats were ventilated with 100% oxygen throughout the experiment except during the hypoxia protocols. The O2 concentration of the breathing mixture was monitored with a P-sensitive electrode located at the intake of the ventilator. Rectal temperature (maintained at 37°C) and end-tidal CO2 were monitored throughout the experiment with a capnometer (Columbus Instruments, Ohio, USA) that was calibrated twice per experiment against a calibrated CO2–N2 mix. This instrument provided a reading of < 0.1% CO2 during inspiration in animals breathing 100% oxygen and an asymptotic, nearly horizontal reading during expiration. We previously showed that the capnometer readings closely approximate arterial P (Guyenet et al. 2005a). After injection of the intravenous anaesthetic mixture, the adequacy of anaesthesia was monitored during a 20 min stabilization period by testing for absence of withdrawal response, lack of BP change and lack of change in PND rate or amplitude to firm toe pinch. After these criteria were satisfied, the muscle relaxant pancuronium was administered at the initial dose of 1 mg kg−1i.v. and the adequacy of anaesthesia was thereafter gauged solely by the lack of increase in BP and PND rate or amplitude to firm toe pinch. Approximately hourly supplements of one-third of the initial dose of chloralose–urethane were needed to satisfy these criteria during the course of the recording period (3–4 h).
In vivo recordings of physiological variables and neuronal activity
Arterial pressure (AP), the mass discharge of the phrenic nerve (PND), tracheal CO2 and single units were recorded as previously described (Guyenet et al. 2005a). Before searching for RTN neurons, ventilation was adjusted to lower end-expiratory CO2 to 4% at steady-state (60–80 cycles s−1; tidal volume 1.2–1.4 ml (100 g)−1). These conditions were selected because 4% end-expiratory CO2 was below the firing threshold of both RTN units and the PND. Variable amounts of pure CO2 were then added to the breathing mixture to adjust end-expiratory CO2 to the desired level. When searching for RTN units, end-expiratory CO2 was set at 6.5–7% in order to insure that both PND and the CO2-sensitive neurons of RTN were active. RTN neurons were recorded exclusively under the caudal end of the facial motor nucleus in a region that matches the prior definition of RTN in rats (Cream et al. 2002; Weston et al. 2004; Guyenet et al. 2005a). As in prior work, the caudal and ventral boundaries of the facial motor nucleus were identified in each rat by the large (up to 5 mV) negative antidromic field potential generated in the facial motor nucleus by stimulating the mandibular branch of the facial nerve (for details see Brown & Guyenet, 1985). RTN CO2-activated neurons were encountered between 200 and 350 μm below the lower edge of the facial motor nucleus, 1.6–1.9 mm lateral to the midline and from 100 μm caudal to 400 μm rostral to the caudal end of the facial field potential (Mulkey et al. 2004; Guyenet et al. 2005a). Prior single neuron labelling experiments have indicated that this region lies between coronal planes Bregma −11.7 and Bregma −11.2 mm of the Paxinos and Watson atlas (Paxinos & Watson, 1998; Mulkey et al. 2004; Guyenet et al. 2005a). Most recordings were made on the left side of the brain. The RTN also contains presympathetic barosensitive neurons located on average dorso-medial to the CO2-sensitive neurons (Mulkey et al. 2004). These neurons cannot be silenced by hypocapnia and their discharge rate increases by at most 80% between 4 and 10% end-expiratory CO2. These cells were ignored in the present study.
All analog data (end-expiratory CO2, PND, unit activity, AP) were stored on a microcomputer via a micro1401 digitizer from Cambridge Electronics Design (CED, Cambridge, UK) and were processed off-line using version 5 of the Spike 2 software (CED). Processing included action potential discrimination and binning, neuronal discharge rate measurement, and PND ‘integration’ (iPND) consisting of rectification and smoothing (τ, 0.015 s). Neural minute × volume (mvPND, a measure of the total phrenic nerve discharge per unit of time) was determined by averaging iPND over 50 s (vagotomized rats) or during 20 respiratory cycles (rats with intact vagus nerves) and normalizing the result by assigning a value of 0 to the dependent variable recorded at low levels of end-expiratory CO2 (below threshold) and a value of 1 at the highest level of P investigated (between 9.5 and 10%). The CED software was also used for acquisition of peri-event histograms of neuronal activity and peri-event averages of iPND or tracheal CO2. The peri-event histograms of neuronal single-unit activity were triggered either on iPND or on the tracheal CO2 trace and represented the summation of at least 100 respiratory cycles (350–800 action potentials per histogram).
The steady-state relationship between RTN neuronal activity and end-expiratory CO2 was obtained by stepping the inspired CO2 level to various values for a minimum of 3 min and up to 5 min. The mean discharge rate of the neuron was measured during the last 30 s of each step at which time end-expiratory CO2 and the discharge of the neuron appeared to have reached equilibrium. End-expiratory CO2 was measured by averaging the maximum values recorded from 10 consecutive breaths at the midpoint of the time interval sampled.
Stimulation of carotid chemoreceptor was done with bolus injections of NaCN (50 μg kg−1, i.v.) or by switching the breathing mixture from 100% O2 to 10–15% O2 balanced with N2 for 30 s using an electronic valve. Evidence that the hypoxic stimulus activated neurons via stimulation of carotid chemoreceptors was obtained by demonstrating that denervation of these receptors eliminated the excitatory effect of the hypoxic stimulus on PND and the activity of hypoxia-responsive RTN neurons.
Muscimol (Sigma Chemicals Co.; 1.75 mm in sterile saline pH 7.4) was pressure injected (30 nl in 5 s) bilaterally through single-barrelled glass pipettes (20 μm tip diameter). The muscimol solution contained a 5% dilution of fluorescent latex microbeads (Lumafluor, New City, NY, USA) for later histological identification of the injection sites (Guyenet et al. 1990). These glass pipettes also allowed recordings of field potentials and multiunit brain activity, properties that were used to direct the electrode tip to the desired sites. Injections into RTN were thus guided by recording the facial field potential and were placed 200 μm below the lower edge of the field, 1.6–1.9 mm lateral to the midline and 200–300 μm rostral to the caudal end of the field. Injections into the midline raphe were done bilaterally 300 μm lateral to the midline at the same coronal level and depth as RTN. Injections into the rostral ventral respiratory group (rVRG) were guided by locating inspiratory-related multiunit activity. The target region was found 400 μm rostral to the calamus scriptorius, 1.8 mm lateral to midline and 1.9–2.2 mm below the dorsal surface of the brainstem using electrodes angled 20 deg forward. The electrophysiological recordings were made on one side only and the second injection was placed 1–2 min later in the symmetric brain location based on the stereotaxic coordinates of the first one. The classification of respiratory neurons was based on the timing of their discharge in relation to that of the phrenic nerve using accepted nomenclature (Feldman & McCrimmon, 1999).
Tracer injections were made while the rats were anaesthetized with a mixture of ketamine (75 mg kg−1), xylazine (5 mg kg−1), and acepromazine (1 mg kg−1) administered i.m. Surgery used standard aseptic methods, and after surgery, the rats were treated with the antibiotic ampicillin (100 mg kg−1) and the analgesic ketorolac (0.6 mg kg−1, s.c.).
A group of seven rats received iontophoretic injections of the anterograde tracer biotinylated dextran amine (BDA-lysine fixable, MW 10000; 10% w/v in 10 mm phosphate buffer, pH 7.4; Molecular Probes) into the commissural part of the nucleus of the solitary tract (commNTS) (20 μm tip diameter glass pipettes; 2 μA positive current pulses, 5 s duration every 10 s for 10 min). These injections were made 0.4 mm caudal to the calamus scriptorius, in the midline and 0.3–0.5 mm below the dorsal surface of the brainstem. These rats were allowed to survive 7–10 days following which they were anaesthetized with pentobarbital (60 mg kg−1, i.p.) and perfused transcardially with fixative as described below.
Another group of 11 rats received an iontophoretic injection of cholera toxin B (1% CTB in 0.2 m phosphate buffer, pH 7.35; List Biological Laboratories, Campbell, CA, USA; 20 μm tip diameter glass pipettes; 2 μA positive current pulses, 5 s duration every 10 s for 10 min) into the left RTN in order to retrogradely label commNTS neurons that innervate RTN. These injections were made below the facial motor nucleus under electrophysiological guidance as described above (200 μm below the ventral boundary of the facial nucleus, 1.6–2.0 mm lateral to the midline, between Bregma −11.6 and −11.2 mm). Seven to ten days following the CTB deposits, six of the rats were exposed for 3 h to a hypoxic breathing mixture (8% O2, balanced with N2) in a small flow-through environmental chamber. The rest of the rats were exposed to room air under the same conditions. Except in one case, the experiments were run in pairs consisting of one rat exposed to hypoxia and the other to normoxia. The pair of animals were anaesthetized with pentobarbital and perfusion fixed immediately after the 3 h of exposure to hypoxia or normoxia.
The rats were deeply anaesthetized with 60 mg kg−1 pentobarbital i.p. then injected with heparin (500 units, intracardially) and finally perfused through the ascending aorta with 150 ml of phosphate-buffered saline (pH 7.4) followed by 4% phosphate-buffered (0.1 m; pH 7.4) paraformaldehyde (Electron Microscopy Sciences, Fort Washington, PA, USA). The brain was removed and stored in the perfusion fixative for 24–48 h at 4°C. Series of coronal sections (30 μm) from the brain were cut using a vibrating microtome and stored in cryoprotectant solution (20% glycerol plus 30% ethylene glycol in 50 mm phosphate buffer, pH 7.4) at −20°C for up to 2 weeks awaiting histological processing.
All histochemical procedures were done using free-floating sections according to previously described protocols. BDA was detected using either streptavidin-Cy3 (1: 200; 60 min; Jackson ImmunoResearch Laboratories, West Grove, PA, USA) or avidin–Alexa 488 (1: 200; 60 min; Molecular Probes) (Stocker et al. 2006). Vesicular glutamate transporter 2 (VGLUT2) and glutamic acid decarboxylase 67 (GAD67) were detected by immunofluorescence using a guinea-pig anti-VGLUT2 antibody (AB 5907; Chemicon International, Temecula, CA, USA; dilution 1: 2500 dilution) or a rabbit anti-GAD67 antibody (AB5992; Chemicon International; 1: 2500 dilution; 24–48 h incubation in Tris-buffered saline with 10% horse serum and 0.1% Triton X-100) (Stocker et al. 2006). The VGLUT2 antibody was revealed with Alexa-488-tagged goat anti-guinea-pig IgG (1: 200, Molecular Probes; 60 min). The GAD67 antibody was revealed with Alexa 488-tagged goat anti-rabbit IgG (1: 200, Molecular Probes; 60 min).
Fos immunoreactivity was detected using a rabbit anti-c-Fos antibody (sc-52, Santa Cruz Biotechnology, Santa Cruz, CA, USA; 1: 1500) followed by a Cy-3-conjugated goat anti-rabbit IgG (Jackson; 1: 200). CTB was detected with a goat anti-CTB antibody (List Biological Laboratories, Campbell, CA, USA; 1: 2000) followed by Alexa 488-conjugated donkey anti-goat IgG (Molecular Probes; 1: 250).
GAD67 mRNA was detected using a 3.2 kb digoxigenin-labelled cRNA probe exactly as previously described (Stornetta & Guyenet, 1999). VGLUT2 mRNA was detected using a 3.4 kb probe, also according to previously described methods (Stornetta et al. 2003a). Digoxigenin was revealed with a sheep polyclonal anti-digoxigenin antibody conjugated to alkaline phosphatase (Roche Molecular Biochemicals, Indianapolis, IN, USA) and alkaline phosphatase was reacted with nitro-blue tetrazolium (NBT) and 5-bromo-4-chloro-3-indolyl-phosphate, 4-toluidine salt (BCIP). Previous testing has established the specificity of our probes (Stornetta et al. 2002). Absence of labelling in facial, hypoglossal and nucleus ambiguus motor neurons was taken as the quality standard since these cells are the most prone to exhibit non-specific NBT/BCIP reaction product under suboptimal conditions. Probe hybridization was always carried out before any immunohistochemistry, i.e. before detection of Fos immunoreactivity and or CTB. Following hybridization, all primary antibodies were applied, then the alkaline phosphatase colourimetric reaction was performed and, finally, fluorescent secondary antibodies were applied. Finally, the sections were mounted in sequential rostrocaudal order onto slides, dried and covered with Vectashield (Vector Laboratories, Burlingame, CA, USA). Coverslips were affixed with nail polish.
No labelling was observed when the primary antibodies were omitted. The VGLUT2 antibody was raised against a peptide which, when preincubated with the primary antibody, eliminated immunoreactivity (Stocker et al. 2006). This antibody labelled terminals only. The GAD67 antibody also labelled nerve terminals exclusively. The antigen used to raise this antibody was derived from a cloned feline DNA expressed in E. coli and has been characterized previously (Kaufman et al. 1991). The anti-digoxigenin antibody produced no reaction product in the absence of digoxigenin-labelled RNA probes. Riboprobe immunolabelling was confined to neuronal cell bodies exclusive of nuclei.
Cell mapping, cell counting and imaging
A conventional multifunction microscope (brightfield, darkfield and epifluorescence) was used for all observations except when indicated. The computer-assisted mapping technique made use of a motor-driven microscope stage controlled by the Neurolucida software and has been described in detail previously (Stornetta & Guyenet, 1999). The Neurolucida files were exported to NeuroExplorer software (MicroBrightfield, Colchester, VT, USA) to count the various types of neuronal profiles within a defined area of the reticular formation.
Section alignment between brains was done relative to a reference section. To align sections around RTN level, the most caudal section containing an identifiable cluster of facial motor neurons was identified in each brain and assigned the level 11.6 mm caudal to Bregma (Bregma −11.6 mm) according to the atlas of Paxinos & Watson (1998). Levels rostral or caudal to this reference section were determined by adding a distance corresponding to the interval between sections multiplied by the number of intervening sections. The same method was also used to identify the Bregma level of the muscimol injections targeted to the rVRG. When the object of the experiment was to locate neurons or injection sites at commissural NTS level, the reference section used to align all others was the one closest to mid-area postrema level (Bregma −13.8 mm).
To count the number of BDA-labelled varicosities located in RTN following tracer injection into commNTS, a 200 μm-wide rectangle extending from the medial edge of the trigeminal tract to the lateral edge of the pyramidal tract was placed tangentially to the ventral medullary surface. BDA-labelled axonal varicosities were counted within this rectangle at four levels of the RTN.
The Neurolucida files were exported to the Canvas 9 software drawing program for final modifications. Photographs were taken with a 12-bit colour CCD camera (CoolSnap, Roper Scientific, Tuscon, AZ, USA; resolution 1392 pixels × 1042 pixels).
An Olympus IX81 DSU spinning disk confocal microscope (Olympus America, Inc., Melville, NY, USA) was used to test for colocalization of BDA and either VGLUT2 or GAD67 immunoreactivity in axonal varicosities located in the RTN region. Images were captured with a SensiCam QE 12-bit CCD camera (resolution 1376 pixels × 1040 pixels, Cooke Corp., Auburn Hills, MI, USA). IPLab software (Scanalytics, Rockville, MD, USA) was used for merging of colour channels in photographs of dual labelling experiments.
The neuroanatomical nomenclature is after Paxinos & Watson (1998).
Statistical analysis was done with Sigma Stat version 3.0 (Jandel Corporation, Point Richmond, CA, USA). Data are reported as means ± standard error of the mean. A t test, (paired or unpaired) and one- or two-way parametric ANOVA followed by the Newman-Keul multiple comparisons test were used as appropriate. Significance was set at P < 0.05.
Hypoxia activates commNTS glutamatergic neurons that innervate RTN
Carotid chemoreceptor afferents terminate predominantly in commNTS. To test whether commNTS innervates RTN, the anterograde tracer BDA was injected by iontophoresis into commNTS in seven rats. Five BDA injections out of seven were correctly placed (Fig. 1A). In each of these five cases, the BDA injection was centred at the level of the calamus scriptorius and labelled neurons were confined to the commNTS. One week after injection, numerous BDA-labelled axonal varicosities or terminals (putative synapses) were present in the RTN, defined here as the region where the cell bodies and dendrites of previously identified CO2-activated neurons are located. Figure 1B is a computer-assisted plot of the BDA-labelled varicosities that were detected in a representative coronal section located close to Bregma −11.4 mm (200 μm rostral to the caudal end of the facial motor nucleus). The plotting was limited to the ventral third of the brain. BDA-labelled varicosities were found throughout the ventral medulla albeit at variable densities. As seen in the inset, these putative synapses were especially numerous under the medial half of the facial motor nucleus in very close proximity to the ventral medullary surface. The number of BDA-labelled axonal varicosities present within RTN (defined by the rectangle shown in Fig. 1B; see Methods for details) were counted in four equidistant sections per rat and the resulting distribution histogram is shown in Fig. 1C. The number of BDA-labelled varicosities within the defined area was greatest (P < 0.05 by repeated measure ANOVA) close to the caudal end of the facial motor nucleus and dropped considerably caudal to that level (Fig. 1C). The area with the maximum density of varicosities approximates the region where we find the greatest concentration of CO2-responsive neurons.
The next experiments were designed to test whether the projection from commNTS to RTN neurons is predominantly glutamatergic or GABAergic. Sections from the same five brains were reacted for simultaneous detection of BDA and either VGLUT2 or GAD67 immunoreactivity and the material was examined by confocal microscopy. We focused our observations on the marginal layer of RTN, a readily identified 50 μm-thick region of the ventral medulla that lies closest to the ventral medullary surface under the spinocerebellar tract. This choice was made because the marginal layer contains an especially dense input from commNTS (Fig. 1B) and because the CO2-activated neurons of RTN have profuse dendrites within the marginal layer regardless of whether their cell bodies reside within or dorsal to this region (Mulkey et al. 2004; Weston et al. 2004). The BDA-labelled varicosities present within this region of RTN were rarely immunoreactive for GAD67 but they were commonly VGLUT2-immunoreactive (Fig. 2). An estimate of the percentage of excitatory (VGLUT2-ir) versus inhibitory (GAD67-ir) BDA-labelled varicosities was obtained by sampling the marginal layer with a confocal microscope in each of the five rats. Four equidistant sections between Bregma −11.3 and Bregma −12.2 mm were selected. Within this region an average of 51% of the total number of BDA-labelled varicosities sampled were immunoreactive for VGLUT2 (243 varicosities) whereas only 5% of 212 sampled varicosities were GAD67-ir. The proportion of BDA-positive varicosities containing VGLUT2- versus GAD67-immunoreactivity was approximately the same at the four levels investigated (Fig. 1D). Each level also contained numerous BDA-labelled axonal varicosities in which no other immunoreactivity could be detected.
The next series of experiments was designed to test whether the glutamatergic projection from commNTS to RTN includes neurons that are activated by systemic hypoxia. CommNTS with projections to RTN were prelabelled with CTB 1 week before the rats were exposed to hypoxia (8% O2, 6 rats) or to normal air (5 rats). Fos immunoreactivity was used to identify commNTS neurons that were activated by hypoxia. CommNTS neurons were classified as glutamatergic or GABAergic based on whether they contained VGLUT2 mRNA or GAD67 mRNA.
The CTB injections, placed with electrophysiological guidance under the caudal end of the facial motor nucleus, were centred 100–300 μm dorsal to the ventral medullary surface 2–400 μm rostral to the caudal end of the facial motor nucleus. Figure 3A depicts the location of the CTB injections in the six rats that were exposed to hypoxia. Fos immunoreactivity was absent in the NTS of the five control rats exposed to room air and these rats were not examined further. One series of 30 μm-thick coronal sections (180 μm apart) was selected from the brain of each hypoxia-treated rat and reacted for simultaneous detection of Fos immunoreactivity, VGLUT2 mRNA and CTB immunoreactivity. An adjacent series of sections from the same six brains was reacted for detection of Fos immunoreactivity, GAD67 mRNA and CTB immunoreactivity. As illustrated in Fig. 4, neurons immunoreactive for both Fos and CTB commonly contained VGLUT2 mRNA (Fig. 4A–C) whereas they typically did not contain GAD67 mRNA (Fig. 4D–F). For quantification purposes, the coronal sections located at mid-area postrema level and 180, 360, 540 and 720 μm caudal to that plane were selected and commNTS neurons containing pertinent combinations of Fos immunoreactivity, CTB immunoreactivity and VGLUT2 mRNA were mapped and counted. CommNTS neurons with the same marker combination were summed across the five coronal sections in each rat and the single resulting number was averaged across the six rats. As shown in Fig. 3B and C, the vast majority of the commNTS neurons that were immunoreactive for both Fos and CTB contained VGLUT2 mRNA (92 ± 13%) whereas only a small proportion of the same class of neurons (Fos- and CTB-positive) contained GAD67 mRNA (12 ± 9%; Fig. 3D).
Carotid chemoreceptor stimulation activates RTN neurons
As in prior work, RTN neurons were defined by their location (2–350 μm below the inferior margin of the antidromic facial field potential) and by the fact that they were silent below a threshold level of CO2 and increasingly active at higher levels of CO2. An example of one cell recorded in a rat with intact vagus nerves is shown in Fig. 5A. As under halothane anaesthesia, RTN cells became respiratory modulated only at higher levels of central respiratory drive (Fig. 5A, insets) but their firing probability did not reach zero at any period of the cycle even at 10% end-expiratory CO2, the highest level examined (Fig. 5B). The particular cell shown in Fig. 5B displayed one of several respiratory patterns that we previously observed in halothane-anaesthetized rats (Guyenet et al. 2005a). This particular pattern was interpreted previously as the result of inhibitory volleys during post-inspiration and late expiration (Guyenet et al. 2005a). The full range of respiratory patterns previously identified under halothane was present under chloralose–urethane anaesthesia including the common pattern consisting of early inspiratory and post-inspiratory inhibition, an example of which is shown in Fig. 9B1. However, the respiratory modulation of RTN units was generally less pronounced under chloralose–urethane anaesthesia than under halothane judging by the fact that in a high proportion of the neurons (46%; 13 of 28 sampled neurons) less than 15% difference was observed between the presumed nadir and apex of the PND-triggered histograms even when the CO2 level was above 9%.
At steady state, the discharge rate of RTN neurons was a linear function of end-expiratory CO2 at first and then exhibited incomplete saturation at higher levels of CO2 (Figs 5C and 6). PND also exhibited a somewhat curvilinear relationship to CO2 (Figs 5C and 6) although the saturation was less marked than under halothane. On average, the discharge rate of RTN units increased by 2.2 ± 0.4 Hz for each 1% increase in end-expiratory CO2 during the linear portion of the relationship and reached a mean level close to 10 Hz at 10% end-expiratory CO2 (Fig. 6B). The CO2 threshold of RTN neurons was not statistically different in vagotomized versus intact rats (Fig. 6B) unlike that of the PND, which was significantly lower in vagotomized rats (Fig. 6). Consequently, the CO2 threshold of RTN neurons was lower than that of PND in intact rats (mean threshold of 41 RTN neurons in 23 rats: 5.1 ± 0.2%; mean PND threshold: 6.4 ± 0.3%; P < 0.05) whereas these threshold values were closer in vagotomized rats (mean CO2 threshold of 20 RTN neurons in 11 rats: 5.0 ± 0.3%; mean PND threshold: 5.3 ± 0.5%; P > 0.05) (Fig. 6).
Stimulation of carotid chemoreceptors with brief periods of hypoxia (30 s of 10–13% O2) or intravenous injection of NaCN (50 μg kg−1) was performed in 16 rats with intact vagus nerves, 10 of which had intact carotid nerves and the rest were denervated bilaterally. In rats with intact peripheral chemoreceptors, hypoxia or cyanide increased PND activity and activated every RTN neuron sampled (hypoxia and NaCN: 41 neurons; Fig. 7A and C). During these tests, the CO2 level was set slightly above the PND threshold, i.e. close to 6.5% CO2. The stimulatory effects of hypoxia and cyanide on both PND and RTN neurons was absent in rats with carotid body denervation (Fig. 7B and D). The sensitivity of RTN neurons to CO2 was unaffected by carotid body denervation (control rats: 2.3 ± 0.3 Hz per 1% rise in end-expiratory CO2; chemodenervated rats: 2.2 ± 0.4 Hz; n.s.). On average, carotid body denervation had no effect on the CO2 threshold measured under steady-state conditions (control rats: 4.7 ± 0.3%; chemodenervated rats: 5.2 ± 0.2%; n.s.).
To determine more precisely how peripheral chemoreceptor inputs and central chemosensitivity are integrated at RTN level, we measured the steady-state relationship between RTN neuron discharge rate and end-expiratory CO2 during long exposures to three levels of oxygen (100%, 15% and 21% O2 balanced with N2; 20 min per O2 level). These measurements were made sequentially in the order indicated in six cells from five rats with intact vagus nerves (Fig. 8). The effects of more severe hypoxia could not be tested because excessive hypotension occurred with long duration exposure to hypoxia below 15% O2. In the presence of 15% O2, the CO2 threshold of RTN neurons was significantly reduced (hypoxia: 4.5 ± 0.6% CO2; hyperoxia: 5.3 ± 0.3% CO2; P < 0.05) but the sensitivity of the cells to CO2 was not changed (hypoxia: 3.3 ± 1.1 Hz per 1% rise in end-expiratory CO2; hyperoxia: 2.3 ± 0.5; n.s.; Fig. 8). ‘Normoxia’ (21% O2) had no effect on the CO2 threshold of RTN neurons (normoxia: 5.2 ± 0.4% CO2; hyperoxia: 5.3 ± 0.3% CO2; n.s.) nor on their CO2 sensitivity (normoxia: 2.1 ± 0.2 Hz per 1% rise in end-expiratory CO2; hyperoxia: 2.3 ± 0.5 Hz; NS; Fig. 8).
Injection of muscimol in the rostral ventral respiratory group does not change the sensitivity of RTN neurons to central or peripheral chemoreceptor stimulation
The anatomical experiments described in the first paragraph of the Results section suggested that the activation of RTN neurons by peripheral chemoreceptor stimulation could be due to a direct excitatory input from commNTS. Our prior work on RTN neurons is consistent with the notion that their response to CO2 under hyperoxia is due to their intrinsic chemosensitivity (Mulkey et al. 2004; Guyenet et al. 2005a). If both hypotheses are correct, the response of RTN neurons to hyperoxic hypercapnia and to peripheral chemoreceptor stimulation should persist after inactivation of the central respiratory pattern generator. The GABAA receptor agonist muscimol was selected for this purpose. In six vagotomized rats, bilateral injections of muscimol (1.75 mm, 30 nl per side) were placed under electrophysiological guidance into a region of the VRG from where strong inspiratory-related multiunit activity could be recorded. Muscimol injection into this region eliminated the PND for up to 2 h (Fig. 9A and B). Upon histological examination, the centres of the injection sites were found close to 1.5 mm posterior to the caudal end of the facial motor nucleus (Fig. 10C). By our previous estimates, these sites correspond to the rVRG (Stornetta et al. 2003b). The fluorescent microbeads extended 270 ± 15 μm on each side of the injection centre and therefore muscimol, which probably diffuses more freely that microbeads, is likely to have also reached the respiratory neurons located in the pre-Bötzinger complex.
One or two RTN neurons were fully characterized in each of the six rats before muscimol injection. Attempts to hold the same neuron before and after muscimol injection into the ventrolateral medulla were not successful because of the mechanical disturbance created by moving the muscimol injection pipette. The example shown in Fig. 9 illustrates an experiment in which an RTN neuron was characterized before muscimol and a second RTN neuron was found just after the second muscimol injection in the immediate vicinity of the first one. As shown in Fig. 9A, muscimol eliminated PND but the RTN neuron recorded after muscimol responded to hyperoxic hypercapnia and to peripheral chemoreceptor stimulation in the same way as the cell recorded before muscimol. Consistent with the loss of central respiratory network activity (i.e. PND), the CO2-activated neuron recorded after muscimol injection had no detectable respiratory-like modulation, in other words the cell discharged much more regularly (Fig. 9B versus C).
On average, the properties of RTN neurons recorded before muscimol (9 neurons in 6 rats) were the same as those recorded after muscimol before any recovery of PND occurred (1 neuron per rat in the same 6 rats) (Fig. 10A and B). Specifically, under hyperoxia, the CO2 threshold and the CO2 sensitivity of the cells were the same (Fig. 10A) and the responses of the cells to hypoxia or cyanide were also indistinguishable (Fig. 10B). The only difference was that every neuron recorded after muscimol lacked respiratory modulation and discharged much more regularly (Table 1).
|Inter-spike interval (mean)||Inter-spike interval (s.d.)||Number of neurons|
|Before muscimol||0.115 ± 0.009||0.052 ± 0.08||9|
|After muscimol||0.096 ± 0.008||0.022 ± 0.03||6|
|P (paired t test)||n.s.||< 0.05|
Bilateral injections of muscimol into RTN eliminate PND at rest and during chemoreceptor stimulation
RTN is believed to contribute an important source of excitatory drive to the respiratory network under anaesthesia. The final experiments were designed to determine whether bilateral inhibition of RTN is capable of suppressing the stimulatory effect of both central and peripheral chemoreceptor activation on PND. Muscimol (1.75 mm, 30 nl) was injected bilaterally under electrophysiological guidance 200 μm below the facial motor nucleus and 200 μm rostral to the caudal end of this nucleus to target the region that contains the highest density of CO2-sensitive RTN neurons according to our prior experience (Mulkey et al. 2004; Guyenet et al. 2005a). The centre of the injection sites were in the desired location as shown by histological mapping of fluorescent microbeads included in the injectate (Fig. 11C). The beads were found to have spread approximately 260 μm on each side of the injection centre.
As illustrated in Fig. 11A, muscimol eliminated PND and no activity, even tonic, could be restored by combined central (10% CO2) and peripheral chemoreceptor stimulation (10–13% O2 or i.v. injection of NaCN). Complete recovery from the effect of muscimol occurred within 2 h (Fig. 11B). Muscimol produced the same complete PND inhibition in each of eight rats. These muscimol injections also reduced blood pressure from 121 ± 5 to 97 ± 6 mmHg (P < 0.01), consistent with muscimol having inhibited some of the blood pressure-regulating neurons that reside in the ventrolateral medulla at and caudal to RTN level.
Bilateral injections of the same amount of muscimol into the midline raphe at the same coronal plane as RTN (two 30 nl injections; 6 rats) produced no effect on resting mvPND measured at an end-expiratory CO2 of 7–8% (96 ± 12% of pre-drug mvPND after muscimol; n.s.). Muscimol injection into the raphe had no effect on blood pressure (from 122 ± 5 to 123 ± 5 mmHg; n.s.).
The present study indicates that RTN neurons respond both to central P and to signals from carotid chemoreceptors. The pathway between carotid chemoreceptor afferents and RTN neurons is excitatory and may involve a single intervening glutamatergic neuron located within commNTS (Fig. 12). We conclude that RTN neurons integrate central and peripheral chemoreceptor information and may drive diaphragmatic activity and/or other aspects of the cardiorespiratory network.
Properties of RTN neurons under chloralose–urethane anaesthesia
RTN neurons recorded under chloralose–urethane anaesthesia had similar properties as under halothane, the anaesthetic used in our previous experiments. The sensitivity of RTN neurons to CO2 under hyperoxic conditions presumably reflects their intrinsic response to extracellular pH (Mulkey et al. 2004; Guyenet et al. 2005a). This CO2 sensitivity was the same under both anaesthetics (2.2 Hz per 1% change in end-expiratory CO2). This fact is noteworthy since halothane opens TWIK-related acid-sensitive channels (TASK), which are expressed by many brainstem respiratory neurons and are candidate molecular substrates of central chemosensitivity (Bayliss et al. 2001; Washburn et al. 2003). The present results do not exclude a contribution of TASK channels to central chemosensitivity but they demonstrate that 1% halothane does not affect the pH sensitivity of RTN neurons any more than chloralose–urethane anaesthesia used at a dose that produces a comparable depth of anaesthesia.
There were a few minor differences between the two anaesthetics, however. The CO2 threshold of RTN neurons under hyperoxia was slightly higher under chloralose–urethane (5%versus 4.2%) and the relationship between RTN neuron activity and CO2 exhibited a somewhat less pronounced saturation at high levels of hypercapnia. PND had a similar CO2 threshold as under halothane but, like RTN neurons, PND also exhibited a less pronounced saturation at high levels of hypercapnia. As shown previously, the intrinsic response of RTN neurons to P is linear and the saturation of their discharge rate at high levels of CO2 is caused by inhibitory inputs from the CPG that increase in intensity along with the central inspiratory drive (Guyenet et al. 2005a). This input is somewhat weaker under chloralose–urethane as suggested by a generally more modest central respiratory modulation of RTN neurons than under halothane. A weaker input should produce a more linear relationship between RTN activity and CO2, as was observed.
RTN neurons receive oligo-synaptic excitatory inputs from carotid chemoreceptors
All RTN neurons were vigorously activated by brief periods of hypoxia and by intravenous cyanide. Since these responses were observed only in rats with intact carotid chemoreceptors, they depended entirely on afferent inputs from these organs. These results are congruent with prior experimentation in rats and cats (Bodineau et al. 2000b; Mulkey et al. 2004).
Carotid chemoreceptor afferents primarily innervate commNTS (Blessing et al. 1999). Most of the commNTS neurons that are activated by carotid body stimulation are not respiratory modulated and therefore are probably not part of the respiratory pattern generator (Koshiya & Guyenet, 1996; Paton et al. 2001). Many of these carotid body-responsive neurons project to the ventrolateral medulla where one of their targets has long been assumed to be the blood pressure-regulating C1 neurons (Aicher et al. 1996; Koshiya & Guyenet, 1996; Paton et al. 2001). According to the present study, the commNTS neurons that are activated by carotid body stimulation innervate the RTN neurons and this projection is predominantly glutamatergic. These NTS neurons could also innervate other ventrolateral medullary neurons including the blood pressure-regulating C1 neurons (Aicher et al. 1996; Sun & Reis, 1996) but this issue is not addressed by the present experiments.
The presumption that commNTS neurons innervate the CO2-sensitive neurons is based on the observation that many BDA-labelled axonal varicosities were found in the marginal layer of RTN where the CO2-sensitive cells have extensive dendrites (Mulkey et al. 2004; Guyenet et al. 2005a). The Fos-expression experiments also indicated that the projection from commNTS to RTN contains many neurons that are activated by hypoxia. We assume that the vast majority of the NTS neurons that express Fos following hypoxia respond to activation of the carotid bodies rather than to secondary effects of hypoxia on blood pressure. In any event, the present results are consistent with prior electrophysiological evidence that commNTS cells with carotid chemoreceptor input innervate the rostral ventrolateral medulla in rats and the RTN region of cats (Koshiya & Guyenet, 1996; Bodineau et al. 2000a).
In addition, the present study shows that the hypoxia-activated pathway between commNTS and RTN is predominantly glutamatergic since an average of 92% of these neurons contained detectable levels of VGLUT2 mRNA. A smaller percentage of BDA-labelled varicosities were found to contain VGLUT2 immunoreactivity in the anterograde tracing experiments (51%) but this method may underestimate the proportion of terminals that are glutamatergic for two reasons. First, dual labelling for BDA and VGLUT2 may not reveal each marker equally, especially in the depth of the tissue because of incomplete antibody penetration. It is also conceivable that a portion of the BDA-labelled structures that look like axonal varicosities may lack VGLUT2 immunoreactivity simply because they are not synapses. This second interpretation is less likely because, where investigated, the vast majority of BDA axonal varicosities seen at the light microscopic level correspond to synaptic sites seen by electron microscopy of thin sections (Kincaid et al. 1998). Therefore, it is more instructive to contrast the high percentage of putative VGLUT2-immunoreactive synapses identified in the projection from commNTS to RTN (around 51% of the BDA-labelled varicosities) with the low percentage of terminals identified as GAD67-immunoreactive (5%). In summary, the vast majority of hypoxia-sensitive commNTS neurons with RTN projections are glutamatergic but a small fraction of this pathway may be inhibitory since around 12% of neurons containing Fos and CTB also contained GAD67 mRNA in rats exposed to hypoxia. This conclusion is also compatible with the fact that RTN contained a small number of BDA-labelled terminals that were immunoreactive for GAD67 after injection of the tracer into commNTS. This inhibitory projection may not target the CO2-activated neurons of RTN but may contact more medially located neurons that regulate sympathetic tone to the skin and the brown adipose tissue, outflows that are inhibited by hypoxia (Madden & Morrison, 2005).
The present results do not prove that the hypoxia-activated glutamatergic neurons of commNTS that innervate RTN are second-order neurons, i.e. receive monosynaptic inputs from carotid chemoreceptor afferents. However, this possibility is likely given that, in slices, low-jitter, presumably monosynaptic, EPSCs can be elicited by stimulating the solitary tract in most of the NTS neurons that project towards the ventrolateral medulla (Bailey et al. 2004).
In summary, the present study demonstrates that the RTN region receives glutamatergic inputs from commNTS neurons that are activated by peripheral chemoreceptor stimulation. These glutamatergic neurons could well be the sole central neurons interposed between carotid chemoreceptor afferents and the chemosensitive neurons of RTN though the existence of interneurons within the NTS or the RTN is not ruled out by the present evidence. In any case, the main pathway between carotid chemoreceptor afferents and RTN neurons almost certainly bypasses the respiratory rhythm and pattern-generating network since injections of muscimol into the ventral respiratory column caudal to RTN eliminated PND but did not affect the response of RTN neurons to hypoxia or cyanide.
RTN neurons respond both to brain extracellular fluid P and to blood gas composition as detected by peripheral chemoreceptors
At present, the theory that a pH-sensitive resting potassium conductance underlies the chemosensitivity of RTN neurons accounts satisfactorily for prior observations in slices (Mulkey et al. 2004; Putnam et al. 2004). However, ATP release, possibly from nearby non-neuronal cells, may also contribute to the CO2 responsiveness of RTN neurons or other nearby chemoreceptors in vivo (Gourine et al. 2005). Whether it is intrinsic, paracrine or both, the response of RTN neurons to brain Pin vivo does not seem secondary to the activation of the respiratory network, at least under anaesthesia. This notion is supported by prior evidence that the response of RTN neurons to hyperoxic hypercapnia is unaffected by high concentrations of a glutamate receptor antagonist in vivo and that RTN neurons have a comparable response to pH in coronal slices (Mulkey et al. 2004; Guyenet et al. 2005a). Consistent with this interpretation, in the present study the response of RTN neurons to hyperoxic hypercapnia was unaffected by inhibiting the ventral respiratory column with muscimol at sites caudal to the RTN. The sole effect of muscimol was to regularize the discharge of RTN neurons as expected from the loss of the respiratory-phasic inputs that these neurons receive from the central respiratory pattern generator (Guyenet et al. 2005a).
Steady-state peripheral chemoreceptor stimulation increased the discharge rate of RTN neurons by a fixed amount regardless of the level of end-expiratory P. In other words, extracellular fluid pH and peripheral chemoreceptor stimulation seem to have roughly additive effects on RTN neuron activity, at least at the mild level of hypoxia that could be reliably investigated without causing hypotension (15% O2). Interestingly, the effect of peripheral chemoreceptor stimulation and the central action of CO2 on PND are also typically additive under anaesthesia (Nattie et al. 1991). At RTN level, this summation had the effect of lowering the end-expiratory CO2 threshold of the neurons, i.e. the CO2 level at which they started to discharge. If the molecular substrate of central chemosensitivity is nothing more than a set of pH-sensitive neuronal conductances, it is logical to expect that the CO2 response of central chemoreceptor neurons would vary according to the synaptic inputs that they receive. However, to our knowledge, RTN neurons are the first documented example where this convergence involves an excitatory input from peripheral chemoreceptors.
In summary, under anaesthesia, RTN neurons detect brain extracellular fluid pH, probably directly, and they encode this variable in a manner that depends on the strength of the input that they simultaneously receive from carotid chemoreceptors. The CO2 threshold of the cells is therefore not fixed but dependent on the synaptic inputs that these cells receive.
RTN and the chemical drive to breathe
Injections of muscimol into RTN eliminated PND at rest and during activation of central and peripheral chemoreceptors. These injections were centred below the caudal pole of the facial motor nucleus where the presumed RTN chemoreceptor neurons are most concentrated based on present and prior recordings (Mulkey et al. 2004; Guyenet et al. 2005a,b). At face value, these results are congruent with many other lines of evidence which suggest that RTN is a chemosensitive region that drives inspiration (Nattie et al. 1991; Feldman et al. 2003; Onimaru & Homma, 2003) and with anatomical evidence that some RTN neurons may be directly antecedent to phrenic premotor neurons (Dobbins & Feldman, 1994). The contribution of the RTN region to breathing is not limited to the anaesthetized state. Even unilateral lesion of this bilateral structure produces notable chronic deficits in the hypercapnic ventilatory response of awake rats (−39%; Akilesh et al. 1997). The fact that muscimol also eliminated the effect of carotid chemoreceptor stimulation on PND could also be viewed as evidence that RTN encodes the chemical drive to breathe, at least under anaesthesia.
Although the macrophysiological data (RTN lesions, stimulations, etc.) are generally consistent with the electrophysiological characteristics and projection pattern of RTN neurons, both lines of supporting evidence have inherent limitations. First, regarding the electrophysiological evidence, the discharge and projection pattern of RTN neurons provide necessary but not sufficient evidence of their role in respiratory control since the exact targets of these neurons are still unknown. Second, the exact boundaries of the tissue affected by procedures like muscimol injection, ventral medullary surface cooling, and chemical or electrolytic lesions are difficult to assess accurately (Millhorn, 1986; Fukuda et al. 1993; Nattie & Li, 1994; Forster et al. 1995).
The fluorescent microbeads that were co-injected with muscimol migrated up to 300 μm from the injection centre. Respiratory and other ventrolateral medullary formation neurons in rat typically have dendrites that spread no farther than 200 μm from their cell bodies in the rostrocaudal direction but exceptions (up to 500 μm spread) are not uncommon (Pilowsky et al. 1990; Schreihofer & Guyenet, 1997). Therefore it is conceivable that neurons located from 500 μm up to 800 μm caudal to the centre of the injection sites (3–600 μm caudal to the posterior edge of the facial motor nucleus) could have been inhibited to various degrees by injecting muscimol into RTN. Our estimates of the effective spread of muscimol are much smaller than the 1.7–2 mm chemical spread measured autoradiographically by Edeline et al. (2002). This difference can be explained in part by the fact that we administered smaller volumes and eight times less drug than the minimum dose administered by these authors (30 versus 50 nl, 50 versus 440 pmol). In addition, chemical spread and effective spread are loosely related since the first depends on the sensitivity of the detection method whereas the second depends on a threshold concentration being achieved for meaningful receptor occupancy. Our estimate of the effective spread (at most 800 μm from the centre and probably much less) is consistent with the fact that injections into the raphe 1.5 mm lateral to the centre of RTN produced no respiratory effect at all. They are also consistent with the fact that muscimol injection into the rVRG did not decrease blood pressure. Decreases of 50–60 mmHg would have been expected if the drug had inhibited a significant fraction of the blood-pressure regulating neurons whose cell bodies are largely confined to the Bötzinger level of the ventral respiratory column therefore well within 1 mm of the rVRG (Kanjhan et al. 1995; Schreihofer et al. 1999b). Judging the spread of muscimol from these physiological criteria, injections placed into RTN could have inhibited respiratory neurons located at the Bötzinger level of the VRC but most probably not caudal to that level. The Bötzinger region contains expiratory augmenting neurons and other respiratory neurons (including pre-inspiratory neurons identified in vitro) that could play an essential role in generating inspiratory activity (Onimaru et al. 1992; Kanjhan et al. 1995; Sun & Reis, 1996; Schreihofer et al. 1999a; Mellen et al. 2003). In short, we cannot exclude that muscimol injection into RTN could have eliminated PND in part or even in totality by silencing neurons located caudal to the CO2-activated neurons that are the focus of the present study. Consequently, alternate functions of RTN neurons should also be considered, particularly the possibility that these neurons could be regulating selected autonomic efferents or respiratory outflows other than to the diaphragm.
Some experiments suggest that the RTN region may preferentially regulate the activity of expiratory muscles (Forster et al. 1995; Janczewski & Feldman, 2006). The theory that RTN neurons are part of an expiratory rhythm generator (Janczewski & Feldman, 2006) is not at odds with the present observations. Since the activity of expiratory muscles is increased by central and peripheral chemoreceptor stimulation, RTN neurons could be a, or the, source of chemical drive for this part of the respiratory network. The small respiratory modulation of RTN neurons that we observed could be a consequence of the anaesthesia. On the other hand, the greater sensitivity of expiratory than inspiratory muscle activity to RTN inhibition (Forster et al. 1995; Onimaru & Homma, 2003) does not exclude the possibility that RTN could drive both outflows; the differential susceptibility of these outflows to RTN inhibition could simply be due to a higher threshold for activation of the downstream expiratory network. This interpretation is consistent with the fact that inspiratory activity is much more strongly inhibited by RTN lesions in models in which inspiratory drive is reduced by cold or anaesthesia (Nattie & Li, 2000; and present results) than in awake animals in which synaptic activity would certainly be more robust (Li & Nattie, 1997). It is also consistent with the fact that RTN acidification, which is less likely to affect neurons distant from RTN than muscimol, does increase inspiratory activity (Li & Nattie, 1997). Finally, the possibility that RTN neurons drive numerous targets is also consistent with the variety of central respiratory patterns exhibited by these neurons (Guyenet et al. 2005a). In fact, based on their location, anatomical connections or central respiratory discharge pattern, RTN neurons could also be regulating other respiratory efferents (airway muscles) or a selection of sympathetic (e.g. to heart, kidney or muscles) or parasympathetic outflows (e.g. to tracheal muscles) (Millhorn & Eldridge, 1986; Perez Fontan & Velloff, 1997; Guyenet et al. 2005a).
Raphe and breathing
The midline raphe contains serotonergic neurons and perhaps other neurons that regulate breathing and change the response of the respiratory network to CO2 (Hodges et al. 2004; Nattie et al. 2004; Richerson et al. 2005). Under our experimental conditions, muscimol injection into the midline raphe at the same rostrocaudal level as RTN produced no effect on either PND or blood pressure. Interestingly, muscimol injection into approximately the same region of the medullary raphe in awake rats does not reduce the effect of hypercapnia on ventilation either (Taylor et al. 2005). Although this portion of the medullary raphe contains very large numbers of serotonergic neurons (Stornetta et al. 2005), these particular serotonergic neurons may not regulate breathing. Alternatively, the raphe neurons that are relevant to breathing could have been inactive in our experiments because of the anaesthesia, or the fraction of the raphe that was impaired by muscimol could have been too small to produce detectable effects on breathing (Hodges et al. 2004). The first alternative is more likely because muscimol injection into the rostral medullary raphe of awake rats does produce effects on respiration (Taylor et al. 2005). However, these effects are difficult to reconcile with the notion that the rostral medullary raphe contains respiratory chemoreceptors since muscimol potentiates the increase in ventilation produced by hypercapnia (Taylor et al. 2005). Furthermore, injection of dl-homocysteic acid into the same region of the raphe produces apnoea and powerful inhibition of the PND under anaesthesia (Verner et al. 2004), which also suggests that this region of the raphe inhibits breathing when it is activated. In any event, our data indicate that the midline raphe at RVL level does not contribute detectably to central chemosensitivity under our experimental conditions.
Conclusion: RTN as common central chemoreceptor pathway
Millhorn & Eldridge (1986) have suggested the possibility that afferents from carotid and medullary chemoreceptors may converge on neurons situated upstream of the respiratory centre pattern generators. The dysfunction of a medullary centre that integrates central and peripheral chemoreceptor inputs provides the most plausible explanation of the spectrum of respiratory deficits experienced by central congenital hypoventilation syndrome (CCHS) patients (Shea et al. 1993). The present experiments indicate that RTN neurons have the physiological properties and anatomical location that are required of such a chemoreceptor integrating centre. Based on our prior work, we postulate that the so-called central chemoreceptor input to RTN neurons may be nothing more than their intrinsic sensitivity to pH. The present work does not exclude the possibility that integration between intrinsic chemosensitivity and inputs from peripheral chemoreceptors could also be occurring at other brainstem sites, including in regions that are capable of influencing the respiratory network.
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