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Dalton Cardiovascular Research Center, Department of Veterinary Biomedical Sciences, College of Veterinary Medicine, University of Missouri, Columbia, Missouri 65211
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ABSTRACT |
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Top
Abstract
Introduction
Methods
Results
Discussion
References
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The purpose of this study was to evaluate the
use of the fluorescent membrane label FM1-43 as a measure of
synaptic terminal exocytosis during stimulation of labeled aortic
baroreceptor and unlabeled nodose ganglia neurons. Activation of the
nerve terminals with electrical stimulation or depolarization with 90 mM KCl in the presence of 2.0 µM FM1-43 resulted in bright,
punctate staining of synaptic boutons. Additional depolarization in the
absence of dye resulted in destaining with a time course that was
consistent and repeatable in multiple boutons within a given terminal.
Destaining was dependent on calcium influx and was blocked by bath
application of 100 µM CdCl2.
Whole cell patch-clamp studies have reported that
depolarization-induced calcium influx in aortic baroreceptor cell
bodies is predominantly caused by the activation of 
-conotoxin GVIA
(-CgTx)-sensitive N-type calcium channels. In addition, these N-type
channels have been shown to be inhibited by activation of metabotropic
glutamate receptors. In the present study, exocytosis in aortic
baroreceptor terminals was not affected by bath application of 5 µM
nifedipine and only partially inhibited by bath application of 2.0 µM
-CgTx. However, depolarization-induced exocytosis was significantly
inhibited by bath application of 200 µM L-AP4, a type III
metabotropic glutamate receptor agonist. Results from this study
suggest that 1) FM1-43 can be
used to measure synaptic vesicle exocytosis in baroreceptor neurons;
2) the N-type calcium channel may
not be involved in the initial phase of vesicle exocytosis; and
3) activation of L-AP4-sensitive
metabotropic glutamate receptors inhibits 90 mM KCl-induced vesicle
release.
nodose neurons; metabotropic glutamate receptors
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INTRODUCTION |
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Top
Abstract
Introduction
Methods
Results
Discussion
References
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BARORECEPTOR AFFERENTS containing feedback information concerning the level of blood pressure ascend centrally and synapse on medullary neurons (13, 14) within the nucleus of the solitary tract (NTS), where their combined information is integrated. Activity- and time-dependent integration of this baroreceptor information is the first step in the normal regulation of cardiovascular reflex function. Alteration of synaptic transmission between the baroreceptor afferents and NTS neurons has been suggested to contribute to inappropriate reflex control of sympathetic outflow.
Although high-frequency stimulation of baroreceptor afferents has been shown to decrease evoked NTS excitatory potentials (2, 17, 18) and results in a rightward shift in the baroreceptor reflex (6, 11), the cellular mechanisms underlying activity-dependent alterations in the signal transduction between baroreceptor afferents and NTS neurons are unknown. One potential mechanism, which may be important in the modulation of baroreceptor afferent synaptic efficacy, is an alteration in transmitter release from the presynaptic terminal. It has been suggested that the maintenance of synaptic transmission over a wide range of frequency stimulations requires that nerve terminals maintain pools of synaptic vesicles in reserve that can be recruited to active zones during periods of intense activity (8, 19). Thus the mechanisms governing synaptic turnover at the baroreceptor terminals may serve as a common site for the regulation of baroreflex function.
Unfortunately, measurements of synaptic vesicle endocytosis and exocytosis from baroreceptor neurons are not technically possible in vivo. The study of the process of exocytosis and endocytosis in other cell types has been most directly approached by measuring changes in membrane capacitance that are associated with the turnover of synaptic vesicles. These studies have been restricted to relatively large neuroendocrine cells and retinal bipolar cells because of the inaccessibility of most small neuronal synaptic terminals to these types of measurements. An optical technique developed by Betz and colleagues (4, 5) allows for a functional assay of synaptic vesicle exocytosis and recycling by quantitative fluorescence imaging of styryl dyes such as FM1-43 [N-(3-(triethylammonium)propyl)-4-(4-dibutylaminostyryl pyridinium, dibromide]. This dye labels the membranes of recycling synaptic vesicles and has been used in a number of different cell types including motor nerve terminals, hippocampal neurons in culture, pituitary cells, and retinal bipolar cells to monitor exocytosis and endocytosis (4, 21-24). In the present study, FM1-43 was used to monitor vesicle exocytosis and recycling in aortic baroreceptor neurons and unlabeled nodose neurons. Here we have described the time course of exocytosis during both electrically evoked action potential stimuli and high-potassium depolarization. In addition, the modulation of aortic baroreceptor exocytosis by selective calcium-channel toxins and the metabotropic glutamate receptor (mGluR) agonist L-AP4 was evaluated.
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METHODS |
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Top
Abstract
Introduction
Methods
Results
Discussion
References
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Neuronal culture and preparation. Methods for labeling and dissociation of nodose ganglia neurons from 3-wk-old Sprague-Dawley rats were similar to those described previously (10, 15). Briefly, the aortic baroreceptor neurons were identified with the use of the diffusible fluorescent dye DiI (Molecular Probes). Two-week-old Sprague-Dawley rats were anesthetized with Metaphane and artificially ventilated. From a midline incision, the right aortic depressor nerve (ADN) was isolated from the surrounding tissue. A small piece of Parafilm was placed underneath the nerve. With the use of a glass micropipette, small amounts of dye were placed along the length of the isolated nerve. To prevent any dye from spilling out to the adjacent tissue, the area was covered with Wacker-Silicone gel, which was allowed to dry before surgical closure. The left aortic nerve was then exposed and similarly labeled. Seven days after the labeling procedure, the nodose ganglia neurons were isolated and placed in cold DMEM + F12, with 5% serum, 0.1% serum extender (GIBCO), and 1% penicillin-streptomycin (DMEF12+). The tissue was then incubated for 30 min at 37°C in an Earle's balanced salt solution containing 5 mg/ml trypsin, 12 µM cysteine, 0.5 mM EDTA, and 1.5 mM CaCl2 and then gently triturated in a trypsin-free solution with serially smaller pipettes until most of the tissue was dissociated. Dissociated cells were then rinsed in DMEF12+ and finally plated on poly-D-lysine-coated coverslips and maintained in DMEF12+ with 8 ng/ml nerve growth factor. Neuronal cultures were used 7-14 days after plating.
Neuronal activation: optical measurements. For optical recordings, coverslips were transferred to a low-volume (0.5 ml) constant-flow recording chamber. Bath solution consisted of (in mM) 138 NaCl, 5.0 KCl, 2.0 CaCl2, 1.0 MgCl2, 10 HEPES, and 10 glucose, pH 7.4. For neurons selectively stimulated through a patch pipette, the pipette solution contained (in mM) 145 K-Asp, 1.0 MgCl2, 2.0 CaCl2, 2.0 EGTA, and 10 glucose. After successful formation of a high-resistance seal, intracellular access using standard whole cell patch techniques was obtained by rupturing the membrane under the pipette with additional suction (9). The configuration was then switched to current clamp, and no holding current was applied. Action potentials were generated by applying 10-ms, 200-nA pulse at 1, 5, 10, or 20 Hz for 20 s. In some experiments, neurons were stimulated by placement of a bipolar stimulating electrode near the neuron of interest. In these experiments, neurons were activated by 10-µA, 2.0-ms, 20-Hz pulses for 15, 30, or 60 s. Finally, some neurons were depolarized by the application of 90 mM KCl to the bath solution.
Loading of the terminal boutons with the fluorescent styryl membrane probe FM1-43 was similar to methods described by others (4, 21-24). Terminals were loaded with FM1-43 (2.0 µM) in the following manner. 1) For experiments using current-clamp stimulation and bipolar electrode stimulation, FM1-43 was added to the bath solution and neurons were stimulated at 20 Hz for 60 s. Neurons then remained in the FM1-43 solution for an additional 120 s to allow for maximum endocytosis of synaptic vesicles (23-25). 2) For experiments using 90 mM KCl to depolarize the neurons, 2.0 µM FM1-43 was added to a 90 mM KCl solution. This solution was then applied to the neurons for 120 s. After both loading procedures, the cells were washed for 5 min before we began the destaining experimental protocol. Quantitative fluorescence measurements were made using a cooled charge-coupled device camera (Photometrics SynSys, Tucson, AZ) and Axon Workbench software (Axon Instruments). Synaptic boutons were viewed with an Olympus IL-2 epifluorescence microscope (40× oil-immersion lens, 1.3 NA) and illuminated with a 75-W xenon lamp. Fluorescence imaging was achieved with 475- to 490-nm excitation and 525- to 550-nm emission filters. Digital images were acquired every 20-30 s and stored on a Gateway Pentium computer. Functional synapses were identified as brightly stained punctae that decreased in fluorescence intensity during stimulation. Regions of interest were generally 1.5 µm2 and were selected to include the largest number of individual fluorescent spots while including minimal background area. This region was digitally marked, and the pixel intensities within the region were then averaged together to obtain a measure of the fluorescent intensity of an individual bouton. Changes in intensity of the region of interest were acquired before, during, and after stimulation. Background images were acquired from dark regions not on the synaptic terminal. The background region was digitally marked, and the average pixel intensity was subtracted from each image. On average, 8-12 regions were selected from each field for study. Data were stored and statistically analyzed using Microsoft Excel. Data are presented as means ± SE with significant differences calculated using ANOVA and Student's t-test.Patch-clamp techniques.
Calcium currents were recorded using whole cell patch-clamp techniques
(9) with polished glass electrodes (1- to 3-M
resistance). The
reference electrode was an Ag-AgCl plug immersed in a 150 mM KCl agar
bridge that was placed in the bath. Recordings were made at room
temperature using an Axopatch 1D patch-clamp amplifier and filtered at
3 kHz using a four-pole Bessel filter. Currents were digitized on-line
at 10 kHz and stored for analysis. Currents were analyzed using the
Axograph data analysis program (Axon Instruments). The bath solution
contained (in mM) 140.0 tetraethylammonium chloride, 5.0 4-aminopyridine, 15.0 glucose, 10.0 HEPES, and 2.0 CaCl2, pH 7.38. The pipette
solution contained (in mM) 124.0 CsCl, 11.0 EGTA, 1.0 CaCl2, 2.0 MgCl2, and 10.0 HEPES, pH 7.3.
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RESULTS |
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Top
Abstract
Introduction
Methods
Results
Discussion
References
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The fluorescent dye FM1-43 was used to label synaptic vesicle membranes in an activity-dependent manner. Synaptic vesicle exocytosis was measured in both DiI-labeled aortic baroreceptor neurons and unlabeled nodose neurons. Figure 1 shows FM1-43 staining and destaining in a labeled ADN neuron. Figure 1A is a phase-bright micrograph of a nodose ganglia neuron. Figure 1B is from the same field but under fluorescent illumination with a rhodamine filter. The DiI-labeled ADN neuron is easily identified by the presence of DiI in the soma. Stimulation of the baroreceptor neuron in the presence of 2.0 µM FM1-43 resulted in the labeling of vesicle membranes (Fig. 1C). With the use of a patch pipette, the neuron was stimulated at 20 Hz (200 nA) for 1 min in the presence of 2.0 µM FM1-43 in the bath solution and was then rinsed for 5 min. The fluorescent punctae are a result of this staining procedure and correspond to recently retrieved synaptic vesicle membrane (4, 21-24). Arrows indicate staining in the terminals. The staining in the soma is similar to what has been recently reported in dorsal root ganglia neurons (12) and may reflect somatic vesicle turnover. Figure 1D is the same field as in Fig. 1C, with the area of interest enlarged. In the absence of FM1-43 and after a 5-min wash period, further stimulation at 1.0 Hz for 60 s resulted in a 45 ± 12% decrease in fluorescence intensity. Increasing the stimulation frequency to 10.0 Hz for an additional 60 s resulted in the nearly complete release of the trapped fluorescence (83 ± 6% decrease, n = 8 regions, P < 0.01; Fig. 1E). The areas that do not destain are considered to be nonspecific background staining. The staining and destaining of the bouton regions is repeatable. Figure 1F shows subsequent staining and destaining of 10 bouton regions in a labeled aortic baroreceptor neuron during a 10-Hz, 60-s stimulation via patch-clamp pipette. Plotted are two subsequent trials, each preceded by the staining with 2.0 µM FM1-43 and a 5-min wash period. The time course and extent of destaining are the same for the two trials. Similar results were obtained from four other cultures.
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The decrease in FM1-43 fluorescence and the corresponding exocytosis have been reported to be dependent on the influx of extracellular calcium (4). To determine whether the decrease in FM1-43 fluorescent intensity in the present study required calcium influx, the effect of the calcium-channel blocker CdCl2 on vesicle exocytosis was determined. Figure 2A shows the staining and destaining of an aortic baroreceptor terminal. In the presence of 2.0 µM FM1-43 without stimulation, no dye is visible in the terminal (Fig. 2A, top). After a 20-Hz stimulation with a bipolar electrode for 1 min in the presence of 2.0 µM FM1-43, followed by a 5-min FM1-43-free wash, fluorescent boutons appeared along the terminal (Fig. 2A, middle). Additional stimulation at 20 Hz for 30 s resulted in the release of FM1-43 from the boutons and a decrease in fluorescent intensity (Fig. 2A, bottom). Figure 2B, top, shows the individual responses from eight different regions over time. The bottom panel of Fig. 2B shows the averaged responses of these eight regions in the absence (control, 90 ± 4% decrease in fluorescence) and presence (CdCl2, 12 ± 2% decrease in fluorescence) of 100 µM CdCl2. Application of 100 µM CdCl2 inhibited the stimulus-evoked destaining. Similar results were obtained in six other cultures.
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Exocytosis can also be induced by high-potassium depolarization. In Fig. 2C, nodose ganglia terminals were loaded with FM1-43 by exchanging the normal bath solution with a solution containing 2.0 µM FM1-43 and 90 mM KCl for 2 min. The bath was then washed in normal solution in the absence of FM1-43 for 5 min. Destaining was then induced by replacing the normal solution with an FM1-43-free 90 mM KCl solution. Figure 2C shows data from the same terminal, which was stained and destained with FM1-43 in three separate trials. Depolarization in the presence of FM1-43 followed by a 5-min wash resulted in punctate fluorescent staining along the length of the process that, when repeated, reappeared in the same regions. Figure 2D illustrates repeatability of the time course of the decrease in fluorescent intensity over time after depolarization with 90 mM KCl. These results are similar to that reported in other preparations and show that, within a given terminal, depolarization-induced staining and destaining of synaptic vesicles with FM1-43 can be repeated in a consistent manner (4, 21-24).
In aortic baroreceptor neuronal cell bodies, the channel responsible
for ~70% of the total calcium current has been shown to be a N-type,
-conotoxin GVIA (-CgTx)-sensitive calcium channel (10, 15).
Figure 3A
illustrates the effect of the L-type-channel blocker nifedipine and the
N-type-channel blocker -CgTx on evoked calcium currents from an
aortic baroreceptor cell body. After a step depolarization to 0 mV from
a 
80 mV holding potential, the evoked calcium current was only
slightly inhibited by 5.0 µM nifedipine. However, application of 2.0 µM -CgTx inhibited the peak calcium current by 73 ± 2%
(similar results were obtained in 12 other cells). To examine the role
of the L- and N-type calcium channels in vesicle exocytosis
from aortic baroreceptor neurons, we examined the effects of nifedipine
(5.0 µM) and -CgTx (2.0 µM) on 90 mM KCl depolarization-induced
exocytosis. The top left panels in Fig.
3B show the effect of 5 µM
nifedipine on 90 mM KCl-induced synaptic vesicle exocytosis. After
staining of the terminal regions (control), the terminal was
depolarized by 90 mM KCl in the absence of FM1-43. The same field
was restained with FM1-43 and then depolarized again in the
presence of 5.0 µM nifedipine. Application of nifedipine had no
significant effect on the rate or extent of vesicle exocytosis (Fig.
3B,
bottom
left, 24 regions in 3 cultures). The
top right panels in Fig. 3B show the
effect of 2.0 µM -CgTx on 90 mM KCl-induced synaptic vesicle exocytosis. After the staining of the terminal regions (control), the
terminal was depolarized by 90 mM KCl in the absence of FM1-43. The same field was stained again with FM1-43 and then depolarized again in the presence of 2.0 µM -CgTx. Application of -CgTx had
no significant effect on the initial rate or extent of vesicle exocytosis recorded in the first 60 s of destaining (Fig.
3B, bottom
right). However, -CgTx did
significantly inhibit the slow phase of destaining that occurred after
60 s of depolarization (at 120 s, control = 85 ± 9% decrease in
fluorescence, -CgTx = 56 ± 5% decrease in fluorescence;
P < 0.05). Similar
results were obtained from eight other cultures.
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Activation of mGluRs has been hypothesized to inhibit neurotransmission in a number of different preparations (3, 7, 20). Previous studies from our laboratory (10) have reported that activation of mGluRs attenuates N-type Ca2+ channels recorded from nodose neuron cell bodies. The present study examined the action of the mGluR agonist L-AP4 on vesicle exocytosis and FM1-43 destaining. Figure 3, C and D, illustrates the effects of L-AP4 (200 µM) on depolarization-induced destaining. Terminals were stained with FM1-43 by replacing the normal bath solution with a solution containing 2.0 µM FM1-43 and 90 mM KCl for 2 min. The bath was then washed in normal solution in the absence of FM1-43 for 5 min. In the control run, destaining was induced by replacing the normal solution with an FM1-43-free 90 mM KCl solution. Depolarization resulted in a rapid decrease in fluorescent intensity as shown previously. In the experimental run, the terminals were stained with FM1-43 and then washed in normal solution containing 200 µM L-AP4 for 3 min. Destaining was then induced by replacing the normal solution with an FM1-43 free, 90 mM KCl solution and containing L-AP4 (200 µM). The addition of L-AP4 markedly inhibited the depolarization-induced exocytosis (control, 89 ± 3% decrease in fluorescence; L-AP4, 9 ± 6% decrease in fluorescence). Similar results were obtained in 7 of 11 cells tested.
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DISCUSSION |
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Top
Abstract
Introduction
Methods
Results
Discussion
References
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There are three major findings from this study. The first aim of this study was to determine whether the styryl dye FM1-43 could be used to stain synaptic vesicles in aortic baroreceptor neurons. The results from this study demonstrate that in these neurons synaptic vesicles can be stained with FM1-43 either by electrical stimulation of action potentials or by high-potassium-induced depolarization. This fluorescent staining decreased in a time-dependent manner on additional stimulation in the absence of the dye. This pattern of staining and destaining is repeatable within a given terminal field. The styryl dye FM1-43 has been shown to label recycling synaptic membranes in a number of different preparations (4, 21-24). However, the present study is the first to report on the use of FM1-43 to monitor synaptic vesicle release in visceral afferent neurons of the nodose ganglia.
The second aim of this study was to determine whether N-type calcium
channels are responsible for synaptic vesicle exocytosis in aortic
baroreceptor terminals. Previous studies have demonstrated that there
is heterogeneity in the region of distribution of voltage-gated calcium
channels in nodose ganglia neurons (16). Somatic calcium currents in
nodose neurons have been shown to be predominantly caused by the
activation of -CgTx-sensitive N-type calcium channels (10, 15).
Likewise, visceral afferent activation of NTS neurons has been shown to
possess an N-type, -CgTx-sensitive component (1). However,
peripheral terminal baroreceptor activation of baroreceptor afferents
has been shown to be insensitive to -CgTx blockade. Results from the
present study suggest that in aortic baroreceptor neurons, KCl
depolarization-induced synaptic vesicle exocytosis that occurs within
the first 60 s is independent of the activation of -CgTx-sensitive
calcium channels. The reasons for the discrepancy between the effects
of -CgTx on vesicle exocytosis as measured in isolated neurons in
the present study and the effects of -CgTx on solitary tract-evoked
NTS excitatory postsynaptic potential in the slice preparation (1) are
unknown. However, there are a number of factors that could
contribute to this discrepancy. In our experiments we used global,
constant depolarization with 90 mM KCl to maximally activate exocytosis
mechanisms. It is likely that the population of
Ca2+ channels activated by the
clamping of the membrane potential with 90 mM KCl is a different
population than those Ca2+
channels activated during the action potential-evoked exocytosis in the
slice. It might be expected that under the 90 mM KCl conditions, the
calcium that enters the cell through the -CgTx insensitive channels
is more than enough to trigger exocytosis. Importantly, -CgTx was
effective at attenuating the exocytosis that continued after the first
minute of depolarization. The significance of the ability of -CgTx
to attenuate this later stage of exocytosis but not the initial stage
is not known. It may suggest a heterogeneity in the population of
Ca2+ channels involved in vesicle
exocytosis in baroreceptor terminals. An -CgTx-insensitive channel
may be responsible for initial exocytosis, and the -CgTx-sensitive
channel may play a more prominent role during prolonged depolarization.
Alternatively, the residual -CgTx-insensitive current may be enough
to release the docked vesicle pool, and the N-type channels may be
essential for refilling of the readily releasable synaptic vesicle
pool.
The third preliminary finding from this study was that vesicle
exocytosis in baroreceptor terminals can be inhibited by application of
the mGluR agonist L-AP4. Previously, we have shown that N-type, -CgTx-sensitive Ca2+ currents
recorded from nodose ganglia cell bodies are inhibited by activation of
mGluR (10). However, because -CgTx was found to have little effect
on the initial rate of exocytosis in this study, it seems reasonable to
hypothesize that the ability of L-AP4 to inhibit vesicle exocytosis may
not involve modulation of a -CgTx-sensitive
Ca2+ channel. Additional studies
will be needed to determine the nature of the mGluR involved and the
mechanisms underlying the inhibition by L-AP4 of vesicle exocytosis in
baroreceptor terminals.
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ACKNOWLEDGEMENTS |
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The authors thank Kathy Lindsley for expert technical assistance. We are also grateful to Dr. Kevin Gillis for suggestions and comments on the manuscript.
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FOOTNOTES |
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This work was supported by National Heart, Lung, and Blood Institute Grants HL-50304, HL-03620, and HL-59676 to M. Hay and HL-54669 to E. M. Hasser.
Address for reprint requests: M. Hay, Dalton Cardiovascular Research Center, Research Park, Univ. of Missouri, Columbia, MO 65211.
Received 18 November 1997; accepted in final form 21 April 1998.
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Abstract
Introduction
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Discussion
References
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C. M. Foley, H. W. Vogl, P. J. Mueller, M. Hay, and E. M. Hasser Cardiovascular response to group I metabotropic glutamate receptor activation in NTS Am J Physiol Regulatory Integrative Comp Physiol, May 1, 1999; 276(5): R1469 - R1478. [Abstract] [Full Text] [PDF]
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C. M. Foley, J. A. Moffitt, M. Hay, and E. M. Hasser Glutamate in the nucleus of the solitary tract activates both ionotropic and metabotropic glutamate receptors Am J Physiol Regulatory Integrative Comp Physiol, December 1, 1998; 275(6): R1858 - R1866. [Abstract] [Full Text] [PDF]
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