Vol. 283, Issue 4, R843-R852, October 2002
Chemical activation of C1-C2 spinal
neurons modulates activity of thoracic respiratory interneurons in
rats
C.
Qin,
J.
P.
Farber,
M. J.
Chandler, and
R. D.
Foreman
Department of Physiology, University of Oklahoma Health
Sciences Center, Oklahoma City, Oklahoma 73190
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ABSTRACT |
Discharge patterns of thoracic dorsal
horn neurons are influenced by chemical activation of cell bodies in
cervical spinal segments C1-C2.
The present aim was to examine whether such activation would
specifically affect thoracic respiratory interneurons (TRINs) of the
deep dorsal horn and intermediate zone in pentobarbital sodium-anesthetized, paralyzed, artificially ventilated rats. We also
characterized discharge patterns and pathways of TRIN activation in
rats. A total of 77 cells were classified as TRINs by location,
continued burst activity related to phrenic discharge when the
respirator was stopped, and lack of antidromic response from selected
pathways. A variety of respiration-phased discharge patterns was
documented whose pathways were interrupted by ipsilateral C1 transection. Glutamate pledgets (1 M, 1 min) on the
dorsal surface of the spinal cord inhibited 22/49, excited 15/49, or excited/inhibited 3/49 tested cells. Incidence of responses did not
depend on whether the phase of TRIN discharge was inspiratory, expiratory, or biphasic. Phrenic nerve activity was unaffected by
chemical activation of C1-C2 in this
preparation. Besides supraspinal input, TRIN activity may be influenced
by upper cervical modulatory pathways.
propriospinal; glutamate; cervical spinal cord; thoracic spinal
cord; respiration.
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INTRODUCTION |
PREVIOUS
EVIDENCE REVEALS that neurons in the upper cervical spinal cord
can serve as a potential filter, processor, and integrator of sensory
information between visceral organs separated by distance and function
(6). For example, chemical activation of upper cervical
spinal neurons with glutamate primarily reduces spontaneous activity
and responses of thoracic spinal neurons to splanchnic nerve
stimulation (22) as well as to chemical stimulation of cardiac afferents or esophageal distension (7). Similar
stimulation also reduces responses of lumbosacral spinal cells to
noxious colorectal distension (32). These effects most
likely descend in propriospinal pathways, because supraspinal
structures are not required.
Thoracic respiratory interneurons (TRINs) have been investigated in
cats (15, 16, 27). These neurons receive supraspinal respiratory drive, and many project to the contralateral ventral horn.
Some TRINs are likely to transmit central respiratory drive to
motoneurons and are hypothesized to coordinate respiratory movements
and other motor activities bilaterally (15, 16, 27).
However, to our knowledge, propriospinal sources of inputs to TRINs
have not been studied systemically. Furthermore, firing patterns of
TRINs have not been examined in rats.
The present study was designed to reveal the functional propriospinal
connections of upper cervical spinal neurons to TRINs. Results of this
study showed that firing patterns of rat TRINs receiving ipsilateral
supraspinal respiratory drive were similar to those in cats.
Furthermore, chemical activation of upper cervical spinal neurons with
glutamate modulated activity of TRINs. A preliminary report has been
published as an abstract (33).
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METHODS |
Experiments were performed on 36 male Sprague-Dawley rats
weighing 320-460 g (Charles River, Boston, MA), which were
initially anesthetized with a bolus injection of pentobarbital sodium
(60 mg/kg ip). Supplementary pentobarbital sodium (10-15
mg · kg
1 · h1 iv) was
administered through a catheter placed in the left jugular vein. The
right carotid artery was cannulated to monitor arterial blood pressure
continuously. After paralysis with pancuronium bromide (0.4 mg/kg ip),
artificial ventilation with room air was administered by a
volume-controlled pump (50-60 strokes/min, 3-5 ml stroke
volume). Supplemental doses of pancuronium bromide (0.2 mg · kg1 · h1 ip) were
administered as necessary to maintain muscle relaxation throughout
experiments. Arterial blood pressure and pupil diameter were monitored
closely to determine the anesthesia level. Average blood pressure was
80-100 mmHg, and pupils were constricted during experiments.
Additionally, arterial blood pressure was unresponsive to noxious
pinching of the feet or tip of tail. Rectal temperature was kept
between 37 and 38°C by using a servo-controlled heating blanket and
overhead infrared lamps. The Institutional Animal Care and Use
Committee of the University of Oklahoma Health Sciences Center approved
the protocols performed in this study.
The experimental set-up is shown in Fig.
1A. The left phrenic nerve
crossing the brachial plexus in the neck was exposed, desheathed, and
crushed caudally. A bipolar platinum hook electrode was placed around
the phrenic nerve to monitor inspiratory drive from the brain stem. To
ensure an adequate phrenic signal under prolonged pentobarbital sodium
anesthesia, CO2 (0-4%) was added to the inspired air.
To exclude recording of sympathetic preganglionic neurons in the upper
thoracic spinal cord, a pair of platinum stimulating electrodes was
wrapped around the left cervical sympathetic trunk after crushing it
rostrally (8, 11). Stimulation parameters were set at
20-30 V, 1 Hz, 0.2 ms to activate sympathetic preganglionic neurons antidromically. To exclude neurons projecting to the
cerebellum, bipolar stainless steel electrodes (4-5 mm apart) were
placed across the midline of the cerebellum, just below the surface
(4, 12). Stimulation parameters were 1-2 mA, 1 Hz,
0.2 ms. To activate descending propriospinal pathways from
C1-C2 spinal neurons, glutamate (1.0 M)
was absorbed onto filter paper pledgets (2 × 2 mm) and placed on
the surface of the C1-C2 spinal cord
(28, 32). This chemical stimulus was used because it
activates cell bodies and does not affect axons passing through these
segments (9). Pledgets remained on the spinal cord for 1 min while examining responses of TRINs. A recovery period of 20-30
min was allowed between glutamate applications. Pledgets with normal
(0.9%) or hypertonic saline (1.0 M) were applied as controls onto
C1-C2 segments before glutamate. In a few
cases, effects of glutamate on TRINs were examined before and after the
superficial dorsal layer of C1-C2 spinal
cord was damaged with heat coagulation.
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Fig. 1.
Experimental set-up and identification of thoracic respiratory
interneurons (TRINs). A: experimental set-up. Ba
and Bb: absence of antidromic activation from
electrical stimulation of cervical sympathetic trunk (20-30 V, 0.2 ms, 10 Hz) and cerebellum (1-2 mA, 0.2 ms, 10 Hz). Arrows mark the
artifacts of antidromic stimulation. Top traces are moving
average of phrenic nerve activity (0.04 s per bin), and
bottom traces are discharges of a single thoracic spinal
neuron. Insets ba and bb: expanded recordings of
phrenic nerve and cell activity. C: respiratory firing of a
TRIN continued when ventilator was stopped, which suggests that this
neuron was driven by central respiratory activity. D
and E: cell activity disappeared or became mainly tonic
when the ventilator was stopped, indicating that these neurons were
driven by peripheral proprioceptive inputs. Top traces are
integrated phrenic nerve activity (0.04 s per bin). Middle
traces are discharges of phrenic nerve. Bottom traces are
discharges of a single thoracic spinal neuron.
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After laminectomies were performed to expose the third thoracic
(T3) spinal segment for recording cells and
C1-C2 segments for application of
glutamate and/or transection of the spinal cord, the rat was mounted in
a stereotaxic headholder and stabilized with vertebral clamps. Dura
mater was carefully cut, and the spinal cord was covered with warm agar
(3-4% in saline) to improve recording stability. Carbon-filament
glass microelectrodes were used for recording extracellular potentials
of single T3 spinal neurons within 0- to 1.4-mm depth from
the dorsal surface and at 0.5-2.0 mm lateral to midline. The
microelectrode was mounted on a micromanipulator at an angle of
70-80° from the vertical in the caudorostral axis. This angled
approach made penetration easier and reduced dimpling of the cord
surface during electrode insertion. To identify TRINs, the following
criteria were used: 1) absence of antidromic activation from
cervical sympathetic trunk and cerebellum (Fig. 1B);
2) no decrease in respiration-related firing pattern
of spinal neurons when the ventilator was stopped for 5-10 s (Fig.
1C). Figure 1, D and E, shows two
examples of neurons whose major respiratory input was associated with
movement of the chest. These neurons were excluded from further study.
All neural signals were monitored online with the Spike 3 data-acquisition system (CED). An increase or decrease in baseline cell
activity >20% during a stimulus was considered an excitatory or
inhibitory response, respectively. Latency to responses, mean maximal
responses, and duration of responses using rate histograms (1 s per
bin) were measured when glutamate was applied to
C1-C2. Also, peak frequency of respiratory burst activity as well as firing level of tonic discharge was measured
using rate histograms (0.1 s per bin) of 10 breaths for either control
activity or maximal response to a stimulus. Expanded traces of cell
activity for illustrations used rate histograms of 0.04 s per bin.
Moving average (0.04 s per bin) of rectified phrenic nerve activity was
used to assess any changes in phrenic nerve activity or breathing rate
accompanying the various manipulations.
To interrupt descending respiratory drive from supraspinal structures
to TRINs, contralateral and sequential ipsilateral spinal transections
with a scalpel blade were made at the C1 segment in eight
animals. In three other cases, ipsilateral and sequential contralateral
spinal transections were made at the C1 segment. Phrenic
nerve activity and cell activity were compared before and after
transection of the spinal cord.
After the study of a spinal neuron was completed, an electrolytic
lesion (50-µA direct current, 20 s) was made at most recording sites. At the end of experiments, animals were killed with an overdose
of pentobarbital sodium, and the thoracic spinal cords were removed and
placed in 10% buffered formalin solution. Frozen sections (55-60
µm) were examined, and laminae containing lesions were identified
(20). Segmental locations of spinal transections were also
confirmed. Statistical comparisons were made using Student's paired or
unpaired t-test and chi-square analysis. Statistical significance was established as P < 0.05, and data are
presented as means ± SE.
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RESULTS |
A total of 77 neurons that showed a firing pattern related to the
central respiratory rhythm were identified as TRINs. Firing rates of
TRINs covered a wide range, from two or three well-spaced impulses per
respiratory cycle to peak frequency >150 impulses/s. In
relation to phrenic nerve discharges, three firing patterns of TRINs
were classified as 1) inspiratory (I) neurons with maximal discharge during phrenic nerve activity; 2) expiratory (E)
neurons with maximal discharges during phrenic silence; and
3) biphasic (B) neurons that did not fit into the above
categories. Lesions of recording sites of 38 TRINs were histologically
identified in the spinal cord (Fig. 2).
Most of the lesion sites were in laminae V and VII of the spinal cord
gray matter. No regional difference in responses of TRINs to glutamate
at C1-C2 was found.
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Fig. 2.
Lesion sites of TRINs in representative third thoracic
(T3) segment of the rat spinal cord. A:
inspiratory TRINs. B: expiratory TRINs. C:
biphasic TRINs.  , TRINs inhibited by glutamate at
C1-C2;  , TRINs excited by
glutamate at C1-C2;  ,
TRINs excited/inhibited by glutamate at
C1-C2;  , TRINs not
responsive to glutamate at C1-C2;
 , TRINs not tested for glutamate at
C1-C2. D: laminae of
T3 segment based on Molander et al. (20). I-V,
VII-IX, laminae; CC, column of Clarke; IL, intermediolateral
nucleus; IM, intermedial nucleus; Liss, Lissauer's tract; LSN, lateral
spinal nucleus; Pyr, pyramidal tract.
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Characteristics of TRINs.
Inspiratory and expiratory firing peaks of inspiratory TRINs averaged
over 10 cycles in 100-ms bins were 66.6 ± 5.8 and 6.5 ± 1.8 impulses/s (n = 50), respectively. Inspiratory TRINs
were further classified as I all, I early, I late, or I tonic according to the starting time and duration of their discharges in the
respiratory cycle (Fig. 3,
Aa-Ad).
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Fig. 3.
Discharge patterns and classification of TRINs.
Aa-Ad: inspiratory (I)-related TRINs and subgroups.
Ba-Bc: expiratory (E)-related TRINs and subgroups.
C: biphasic TRINs and subgroups. PS, phase-spanning.
D: comparison of phasic activity of inspiratory, expiratory,
and biphasic TRINs, and phrenic nerve activity. Activity labeled on
y-axis represents different units of measure for phrenic
nerve activity and spinal neuronal activity. B/M, breaths/min; PMA,
peak moving average of phrenic activity in arbitrary units.
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Examples of expiratory neurons are shown in Fig. 3,
Ba-Bc. Inspiratory firing levels of
expiratory TRINs (1.5 ± 0.5 impulses/s, n = 15)
were significantly lower than those of inspiratory TRINs, whereas their
expiratory firing peaks (18.2 ± 1.5 impulses/s) were
significantly higher than those of inspiratory TRINs (Fig. 3D). Similar to the inspiratory neurons, expiratory TRINs
were also divided into three subcategories: E all, E early, and E late according to the starting time and duration of discharges per respiratory cycle.
Discharges of biphasic TRINs exhibited two cycles during inspiratory
and expiratory phases (Fig. 3Ca). The phase-spanning I-E
TRINs had a postinspiratory discharge of lesser frequency after the
higher-frequency inspiratory discharges (Fig. 3Cb). The
phase-spanning E-I TRINs showed a preinspiratory onset of discharges
during the expiratory phase followed by a further increase in activity
with inspiration (Fig. 3Cc). A summation of all biphasic TRINs showed that the inspiratory firing peaks (23.4 ± 4.9 impulses/s, n = 12) were significantly lower than those
of inspiratory TRINs, but higher than those of expiratory TRINs (Fig.
3D). Expiratory firing peaks of biphasic TRINs (10.6 ± 2.5 impulses/s) were significantly lower than those of expiratory TRINs
(Fig. 3D).
Effects of spinal transection.
To determine descending pathways of central respiratory drive onto
TRINs, activity of TRINs was examined before and after the spinal cord
was transected at the C1 segment in eight animals. Contralateral transection at C1 did not significantly
change either phrenic nerve activity or phasic discharges in all TRINs
examined (Table 1). Sequential
ipsilateral transection at C1, however, completely
abolished phrenic nerve activity and respiratory discharges in 6/8
TRINs. An example of this response is shown in Fig.
4A. Respiratory discharge
pattern of two TRINs became tonic after spinal ipsilateral transection
when phrenic nerve activity in these animals was absent (Fig.
4B). When ipsilateral transection at C1 was made
first in three other animals, phrenic nerve activity and discharges of
TRINs were abolished, and Fig. 4C shows an example. These
results showed that supraspinal respiratory activity drives TRINs under
the present conditions by exclusively ipsilateral descending pathways
within the spinal cord.
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Fig. 4.
Effects of spinal transection at C1 segment
on phrenic nerve activity and spontaneous activity of TRINs.
Aa-Ac: respiratory activity of a TRIN was not affected
by contralateral C1 transection and disappeared after
sequential transection of ipsilateral C1 segment.
Ba-Bc: respiratory firing pattern of a TRIN became
tonic after sequential transection of ipsilateral C1
segment. Traces at right of Ba-Bc
show single action potentials of this TRIN before and after spinal
transections. Ca-Cc: initial ipsilateral transection at
C1 abolished activity of the phrenic nerve and a TRIN.
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Responses to glutamate at C1-C2.
Glutamate pledgets applied to the dorsal surface of the
C1-C2 segments changed activity of 40/49
(82%) TRINs. Excitatory and inhibitory responses to glutamate at
C1-C2 among neurons in the inspiratory, expiratory, and biphasic groups are shown in Table 2. No statistical difference among these
three groups was found in the incidence of excitation or inhibition by
glutamate. Excitation or inhibition of TRINs could change the
categories of cell discharge as presented in Fig. 3. For those cells
excited, one biphasic cell and 4 I (all and late) cells became I tonic,
and one E early cell became E tonic. Twelve cells were inhibited
strongly enough to eliminate all respiratory modulation, one I tonic
cell became I late, and three I all cells became I late.
No statistical difference in responses of TRINs to normal and
hypertonic saline at C1-C2 was observed
(Table 3). Therefore, responses of TRINs
to the two solutions were calculated together as saline control in
subsequent figures. Despite the lack of group differences, in
individual experiments hypertonic saline did elicit effects, but these
were considerably smaller than those induced by glutamate. Figure
5, A and B, shows
examples of hypertonic saline controls and excitatory or inhibitory
responses of TRINs to glutamate on C1-C2
segments.
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Table 3.
Comparison of effects of normal saline (0.9%) and hypertonic saline
(1.0 M) at C1-C2 on activity of TRINs
responding to glutamate at C1-C2
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Fig. 5.
Effects of hypertonic saline (1 M) and glutamate (Glu) at
C1-C2 on TRINs and a comparison of
responses of TRINs to glutamate at C1-C2
before and after superficial layer of
C1-C2 segments was damaged with heat
coagulation. A and A': effects of hypertonic
saline and glutamate at C1-C2 on a TRIN
inhibited by glutamate at C1-C2.
B and B': effects of hypertonic saline and
glutamate at C1-C2 on a TRIN excited by
glutamate at C1-C2. C and
C': effects of damaging the superficial layer of
C1-C2 spinal segments with heat
coagulation on the inhibitory response of a TRIN to glutamate at
C1-C2. D: comparison of spinal
histological sections of intact and damaged superficial layer of
C2 spinal segments. imp/s, Impulses/s.
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The discharge rate (1 s per bin) of 22/40 TRINs decreased with
glutamate at C1-C2 (Table
4). Saline at
C1-C2 did not change activity of these
TRINs (14.2 ± 2.4 vs. 14.5 ± 2.5 impulses/s). An example of
a TRIN inhibited by glutamate at C1-C2 and
the accompanying phrenic nerve discharge is shown in Fig.
6, A and B. In
contrast to responses of TRINs to saline at
C1-C2, both inspiratory and expiratory
phasic activity of TRINs significantly decreased with glutamate at
C1-C2 (Fig. 6, C and
D), whereas phrenic nerve activity did not change before and
during glutamate at C1-C2.
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Fig. 6.
Inhibitory responses of TRINs to glutamate at
C1-C2. Traces from top to
bottom are integration of phrenic nerve activity (0.04 s per
bin), discharges of phrenic nerve, cell discharges, and rate histogram
of cell discharges. A: effects of normal saline at
C1-C2 on activity of a TRIN. B:
glutamate at C1-C2 decreased activity of
this TRIN. Ba-Bc: expanded recording of phrenic nerve
and cell activity. Rate histograms of cell activity in A and
B were performed using 1 s per bin while expanded
traces (Ba-Bc) used 0.04 s per bin. C
and D: summary for effects of saline (normal or
hypertonic) or glutamate at C1-C2 on
phrenic nerve activity and phasic activity of TRINs. Activity labeled
on y-axis represents different units of measure for phrenic
nerve activity and spinal neuronal activity.
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Glutamate at C1-C2 increased the discharge rate
(1 s per bin) of 15/40 TRINs (Table 4), which was compared with effects
of saline at C1-C2 segments on these TRINs
(14.1 ± 3.1 vs. 15.8 ± 3.9 impulses/s). Examples of saline
control and an excitatory response of a TRIN to glutamate at
C1-C2 and the accompanying phrenic nerve
discharge are shown in Fig. 7,
A and B. Compared with effects of saline at
C1-C2 on phasic activity of TRINs,
glutamate at C1-C2 significantly increased
both inspiratory and expiratory firing peaks of TRINs (Fig. 7,
C and D). Phrenic nerve activity did not change
with glutamate or saline applied to C1-C2
segments (Fig. 7, C and D). Additionally, three
TRINs exhibited excitatory-inhibitory responses to glutamate at
C1-C2, and characteristics of these responses are shown in Table 4.
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Fig. 7.
Excitatory responses of TRINs to glutamate at
C1-C2. A: effects of normal
saline at C1-C2 on activity of a TRIN.
B: glutamate at C1-C2 increased
activity of this TRIN. Ba-Bc: expanded recording of
phrenic nerve and cell activity. C and D: summary
for effects of saline (normal or hypertonic) or glutamate at
C1-C2 on phrenic nerve activity and phasic
activity of TRINs. Activity labeled on y-axis represents
different units of measure for phrenic nerve activity and spinal
neuronal activity.
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To evaluate whether superficial neurons might be responsible for
effects on TRINs, heat coagulation was applied to damage the
superficial layer of the C1-C2 segment in
three animals. Superficial damage abolished the initial inhibitory
effects of C1-C2 glutamate on one TRIN and
the excitatory effects on two TRINs. Figure 5, C and
C', shows an example of a TRIN inhibited by glutamate at C1-C2 before and after damaging
superficial layers of C1-C2 segments; a
histological comparison of a normal spinal cord with the lesion is
shown in Fig. 5, D and D'. In the illustrated
example, some inhibition developed over time. This effect was not
observed for the other two cells.
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DISCUSSION |
The major finding of this study was that chemical activation of
upper cervical spinal neurons influenced the activity of most (82%)
TRINs; 55% of responsive neurons were inhibited, 38% were excited,
and the remainder were excited/inhibited by glutamate at
C1-C2 segments. Results also showed that
firing patterns of rat TRINs were generally similar to those occurring
in cats (15); supraspinal respiratory activity drove TRINs
by ipsilateral descending pathways within the spinal cord.
Identification of TRINs.
Four criteria were used for identification of TRINs in the present
study. First, rhythmic activity of thoracic spinal neurons was retained
even after artificial ventilation was turned off. This excluded spinal
neurons receiving proprioceptive inputs from the muscle and joint
receptors during respiratory movement. Second, TRINs were not
antidromically activated by electrical stimulation of the cervical
sympathetic trunk. This excluded thoracic sympathetic preganglionic
neurons receiving central respiratory drive (8). Third,
antidromic activation from stimulation of the cerebellum was absent in
thoracic spinal neurons with rhythmic activity. This test was done
because dorsal spinocerebellar tract neurons in the thoracic spinal
cord can convey information from central respiratory generator
(29) as well as peripheral events in the chest wall during
respiratory movement (12). Fourth, recording depths from
the dorsal surface at which the units were found (mean 971.4 ± 19.5 µm) were histologically located in deeper dorsal laminae and
intermediate zone of spinal gray matter (Fig. 2). Our electrodes were
not driven deeply enough to penetrate motor nuclei. Kirkwood et al.
(15) reported that locations of the respiratory
interneurons in the ventral horn clearly overlap with those of the motoneurons.
We attempted to avoid recording from motor neurons by limiting
electrode penetration to the intermediate zone of spinal cord. A few
superficial penetrations of the ventral horn were above the nuclei
containing motor neurons (24). We prepared animals for
antidromic activation of thoracic respiratory motor neurons by
electrical stimulation of intercostal nerves after cutting dorsal
roots. However, two methodological factors made antidromic identification difficult. First, electrical stimulation of intercostal nerves usually evoked a large field potential that was considerably larger than the amplitudes of neuronal spikes. Second, sites of nerve
stimulation were very close to the spinal cord (7-10 mm); therefore, antidromic responses of spinal motoneurons would be expected
to occur at very short latencies (typically <1 ms). Thus neuronal
spikes could not be distinguished from the large field potentials
produced by intercostal nerve stimulation. A similar issue was noted by
Kirkwood et al. (15) in the study of cats. While we cannot
exclude the possibility that distal dendrites of a motor neuron were
recorded in a few cases, it would be more likely that recordings were
obtained from the dendritic fields closer to the cell bodies of the
numerous interneurons.
Firing patterns of TRINs.
The firing patterns and proportions of TRINs in the present experiments
appear to be generally similar to a previous study in cats
(15). They reported that thoracic respiratory interneurons (T3-T9) in cats were classified as
inspiratory (64%) and expiratory (25%) neurons according to phrenic
nerve activity. The remaining neurons were considered as
postinspiratory or as neurons that did not fit into the other
categories. In the present study, 65% of TRINs were classified as
inspiratory, 19% as expiratory, and the remaining 16% had biphasic
firing patterns. Inspiratory and expiratory TRINs were further divided
into four subgroups, i.e., all, early, late, and tonic, according to
the onset of their discharges in the respiratory cycle. These subgroups
are in general consistent with the classification for respiratory
interneurons in cats, although the "all" subgroup has not been
mentioned (15). A comparison of the two classifications
shows that postinspiratory neurons and "I early" neurons in the
study of Kirkwood et al. (15) were similar to "E
early" and "I all" neurons in the present study, respectively.
Levels of phasic activity among groups of TRINs were different in the
present study. In particular, peak activity of inspiratory TRINs was
significantly greater than those of the other categories.
Central respiratory drive to TRINs.
Descending bulbospinal respiratory neurons have limited direct inputs
to intercostal motoneurons (1, 14, 19, 31), thus
implicating a pool of thoracic interneurons as an intermediary of this
pathway. In the present study, spinal cord hemisection demonstrated
that TRINs received central respiratory drive by an ipsilateral
descending pathway within the spinal cord. No obvious contralateral
respiratory activation of TRINs was observed.
In rats, phrenic motoneurons within the
C3-C6 level of the cervical spinal cord
are connected monosynaptically with bulbospinal respiratory pathways
that relay central respiratory drive (5, 10, 31).
Ipsilateral spinal transection also abolished ipsilateral phrenic nerve
activity in all cases in this study. This result is consistent with
previous studies (21, 25, 34), although others have found
bilateral projections (3, 23, 31).
Effect of C1-C2 activation on TRINs.
Glutamate applied to the surface of the spinal cord has been used to
activate cell bodies in the cervical spinal cord in previous studies
(26, 28, 32) because it does not affect axons of passage
(9). Chemical activation of upper cervical spinal neurons with glutamate primarily inhibits thoracic spinal neurons responsive to
splanchnic stimulation (22) and to chemical stimulation of cardiac afferents or noxious esophageal distension
(7). This procedure also reduces excitatory responses of
lumbosacral spinal cells to noxious colorectal distension
(32). This propriospinal descending inhibition does not
require supraspinal structures because it still occurs after spinal
transection at rostral C1 segment in rats. In the present
study, glutamate applied to the C1-C2
segment inhibited 45% and excited 31% of TRINs. In contrast, C1-C2 glutamate reduces spontaneous
activity and/or excitatory response of about 3/4 of nonrespiratory
upper thoracic neurons responding to either noxious cardiac or
esophageal stimuli (7). This suggests a more complex
modulation for respiratory interneurons.
The location of C1-C2 cell bodies affected
by glutamate is unclear. One possibility is that glutamate at
C1-C2 directly affected activity of upper
cervical inspiratory neurons in laminae V and VII, which then modulated
activity of TRINs by propriospinal descending pathways. In
cats, the majority of upper cervical inspiratory neurons have long
descending axonal projections toward the region of thoracic motoneurons
(T3-T5) (2, 13, 17). In rats,
similar projections of these inspiratory neurons from the upper
cervical spinal cord may also be present (18, 30).
However, such a possibility was thought to be unlikely. First,
glutamate at C1-C2 produced effects at
such short latencies (1.5 ± 0.3 s for excitation and
2.4 ± 0.6 s for inhibition) that cells in laminae V and VII were unlikely to be the targets of glutamate. Second, damaging the
superficial layer of the upper cervical spinal cord with heat coagulation eliminated the effects of
C1-C2 glutamate on thoracic TRINs. This
could mean that superficial neurons have descending projections to
thoracic spinal segments. However, another possibility might be that
glutamate at the surface of C1-C2 segment
excited superficial neurons, and these neurons, in turn, activated
deeper neurons with descending propriospinal projection to the thoracic spinal cord. Furthermore, our data do not exclude the possibility that
over time glutamate could produce effects from deeper layers. Such a
pathway would be consistent with the slowly developing effect after
superficial lesion illustrated in Fig. 5C', but the residual
response in that instance may simply be due to the reduced pool of
superficial cells.
Phrenic nerve activity.
Glutamate on C1-C2 did not significantly
affect central respiratory drive monitored by phrenic nerve activity.
Phrenic motoneurons within C3-C6 levels of
the cervical spinal cord of rats are connected monsynaptically with
bulbospinal respiratory neuronal groups that generate and relay central
respiratory drive (5, 10, 30). Connections between upper
cervical inspiratory neurons and spinal phrenic nuclei are considered
to be weak, although some monosynaptic and paucisynaptic connections to
phrenic motoneurons exist (30). Furthermore, our previous
study and those of others showed that upper cervical inspiratory
neurons are located in deeper spinal laminae and the intermediate zone
of gray matter (17, 18, 35), so direct effects of their
activation would be unlikely in the initial response.
| |
ACKNOWLEDGEMENTS |
The authors thank Dr. C. J. Jou and D. Holston for excellent
technical assistance.
| |
FOOTNOTES |
This work was supported by National Institute of Neurological Disorders
and Stroke Grant NS-35471.
Address for reprint requests and other correspondence: C. Qin, Dept. of Physiology, Univ. of Oklahoma Health Sciences Center, P.O. Box 26901, Oklahoma City, OK 73190 (E-mail:
chao-qin{at}ouhsc.edu).
The costs of publication of this
article were defrayed in part by the
payment of page charges. The article
must therefore be hereby marked
"advertisement"
in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
June 20, 2002;10.1152/ajpregu.00054.2002
Received 9 January 2002; accepted in final form 18 June 2002.
| |
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