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The Journal of Neuroscience, December 15, 1998, 18(24):10619-10628
Noradrenergic Excitation of Magnocellular Neurons in the Rat
Hypothalamic Paraventricular Nucleus via Intranuclear Glutamatergic
Circuits
Shabrine S.
Daftary1,
Cherif
Boudaba2,
Kriszta
Szabó2, and
Jeffrey G.
Tasker2, 3
1 Molecular and Cellular Biology Program,
3 Neuroscience Training Program, and
2 Department of Cell and Molecular Biology, Tulane
University, New Orleans, Louisiana 70118
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ABSTRACT |
Noradrenergic projections to the hypothalamus play a critical role
in the afferent control of oxytocin and vasopressin release. Recent
evidence for intrahypothalamic glutamatergic circuits prompted us to
test the hypothesis that the excitatory effect of noradrenergic inputs
on oxytocin and vasopressin release is mediated in part by local
glutamatergic interneurons. The voltage response to norepinephrine (30-300 µM) was tested with whole-cell recordings in
putative magnocellular neurons of the paraventricular nucleus (PVN) in hypothalamic slices (400 µm). Norepinephrine elicited an
 1 receptor-mediated direct depolarization in 23% of the
magnocellular neurons tested; however, the most prominent response,
seen in 42% of the magnocellular neurons, was an 1
receptor-mediated increase in the frequency of EPSPs. The
norepinephrine-induced increase in EPSPs was blocked by tetrodotoxin
and by ionotropic glutamate receptor antagonists, suggesting that
norepinephrine excited presynaptic glutamate neurons to cause an
increase in spike-mediated transmitter release. The increase in EPSPs
also was observed in a surgically isolated PVN preparation (64% of
cells) and with microdrop applications of norepinephrine (1 mM, 33% of cells) and glutamate (0.5-1 mM,
28%) in the PVN, indicating that the norepinephrine-sensitive
presynaptic glutamate neurons are located within the PVN. Biocytin
injection and subsequent immunohistochemical labeling revealed that
both oxytocin and vasopressin neurons responded to norepinephrine. Our
data indicate that magnocellular neurons of the PVN receive excitatory
inputs from intranuclear glutamatergic neurons that express
1-adrenoreceptors. These glutamatergic interneurons may serve as an excitatory relay in the afferent noradrenergic control of
oxytocin and vasopressin release under certain physiological conditions.
Key words:
hypothalamus; paraventricular nucleus; oxytocin; vasopressin; norepinephrine; adrenoreceptors; glutamate
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INTRODUCTION |
Magnocellular neurons of the
hypothalamic paraventricular nucleus (PVN) and supraoptic nucleus (SON)
are stimulated under certain physiological conditions to generate
bursts of action potentials. Although intrinsic membrane ionic
conductances capable of sustaining bursting activity have been
characterized in magnocellular neurons (Bourque and Renaud, 1991 ;
Legendre and Poulain, 1992), the synaptic mechanisms responsible for
triggering bursts and for coordinating the bursting activity in
response to specific sensory stimuli are less well understood.
Several lines of evidence suggest that noradrenergic inputs play a
critical stimulatory role in the release of oxytocin and vasopressin
under conditions of increased hormone demand. Oxytocinergic and
vasopressinergic magnocellular neurons of the PVN and SON are contacted
directly by noradrenergic synapses (Ginsberg et al., 1994; Michaloudi
et al., 1997), and stimulation of the brainstem A1/A2 noradrenergic
cell groups activates putative magnocellular neurons and causes
oxytocin and vasopressin release (Day et al., 1984; Tanaka et al.,
1985; Day and Sibbald, 1988; Kim et al., 1989). This effect is blocked
by 6-hydroxydopamine lesion of the hypothalamic noradrenergic
projections (Day et al., 1984). Intracerebroventricular or local
hypothalamic injection of norepinephrine or 1-receptor agonists results in the release of oxytocin (Bridges et al., 1976; Tribollet et al., 1978) and vasopressin (Benetos et al., 1986; Willoughby et al., 1987). Similarly, 1-receptor agonists
applied to hypothalamic explants causes depolarization and spike
generation in SON magnocellular neurons and leads to an increase in
oxytocin and vasopressin release (Armstrong et al., 1986; Randle
et al., 1986a,b).
These noradrenergic afferents may be part of the ascending sensory
pathways responsible for selectively activating the oxytocin neurons
during parturition and reflex milk ejection and the vasopressin neurons
during hemorrhage. During parturition, the norepinephrine concentration
in the SON rises prior to and in parallel with increases in blood
levels of oxytocin (Herbison et al., 1997). Blockade of noradrenergic
inputs to the hypothalamus with 6-hydroxydopamine lesion or with
1-adrenoreceptor antagonists inhibits the reflex release
of oxytocin associated with milk ejection (Tribollet et al., 1978;
Clarke et al., 1979; Crowley et al., 1987). Similarly, reflex
vasopressin release in response to unloading of arterial baroreceptors
is accompanied by a rise in the concentration of norepinephrine in the
PVN (Van Huysse and Bealer, 1991), which is caused by the activation of
A1 noradrenergic projections (Day and Renaud, 1984).
Several studies suggest a possible role of glutamate in the triggering
mechanism for oxytocin release. Glutamate levels in the SON have been
found to rise abruptly just before parturition and to decline before
delivery is terminated (Herbison et al., 1997), and
intracerebroventricular injection of a glutamate receptor antagonist
blocks the suckling-induced release of oxytocin (Parker and Crowley,
1993a). We recently found evidence for excitatory synaptic inputs to
magnocellular neurons of the PVN and SON from intrahypothalamic
glutamate neurons (Boudaba et al., 1997). The current study was
conducted to determine whether the excitatory effect on oxytocin and
vasopressin release of noradrenergic inputs is mediated by local
glutamatergic circuits.
A preliminary account of this data has been published previously in
abstract form (Daftary et al., 1996).
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MATERIALS AND METHODS |
Slice preparation. Male Sprague Dawley rats (50-150
gm; Charles River, Wilmington, MA) were deeply anesthetized with
pentobarbitol sodium (50 mg/kg body weight) and decapitated. The brain
was quickly and gently removed from the cranial cavity and immersed in
cooled (1-2°C), oxygenated (100% O2) artificial
CSF (aCSF). The composition of the aCSF was (in mM):
140 NaCl, 3 KCl, 1.3 MgSO4, 1.4 NaH2PO4, 2.4 CaCl2,
11 glucose, and 5 HEPES; pH was adjusted to 7.2-7.4 with NaOH.
The hypothalamus was blocked with a razor, and 400 µm hypothalamic
slices were sectioned in the coronal plane using a vibrating microtome
(World Precision Instruments, Sarasota, FL). Two slices containing the
PVN were identified, and a single slice was transferred immediately to
a ramp-style, interface recording chamber where it was perfused with
humidified, oxygenated aCSF maintained at 32-34°C and allowed to
equilibrate for at least 1 hr before recordings were started. The
second slice was stored submerged in a holding chamber in oxygenated
aCSF at room temperature until it was used. In some experiments, the
PVN was surgically isolated from the rest of the slice under a
dissecting microscope using a scalpel.
Electrophysiological methods. Sharp microelectrodes were
made from microfilament glass capillaries [1.0 mm outer diameter (o.d.), 0.6 mm inner diameter (i.d); World Precision Instruments], and
patch pipettes were pulled from borosilicate glass (1.65 mm o.d., 1.2 mm i.d.; KG-33; Garner Glass, Claremont, CA) using a Flaming-Brown P-97
micropipette puller (Sutter Instruments, Novato, CA). Sharp electrodes
were filled with 2 M potassium acetate. Patch pipettes were
filled with a solution containing (in mM): 120 potassium
gluconate, 10 HEPES, 1 NaCl, 1 CaCl2, 1 MgCl2, 2 Mg-ATP, 0.3 Na-GTP, and 10 EGTA; pH
was adjusted to 7.2-7.4 with KOH. The osmolarity of the patch solution
was made hyperosmotic (290-310 M /l) with 20 mM
D-sorbitol to reduce series resistance.
The slice was transilluminated in the recording chamber, and the
recording electrode was positioned in the magnocellular PVN under
visual guidance using a dissecting microscope. The electrode was
lowered through the slice by 2- to 4-µm-steps with a piezoelectric microdrive (Nanostepper, Adams & List, Westbury, NY). Recordings were
performed in current-clamp mode using an Axoclamp 2A amplifier (Axon
Instruments, Foster City, CA) and were monitored continuously on a
digital storage oscilloscope (Hitachi, Tokyo, Japan). Data were
converted to digital video format (Neurocorder, Neurodata Instruments,
New York, NY) and stored on videotape for off-line analysis. Episodes
of spontaneous EPSPs (30-60 sec) 1 min before and 8-9 min into
norepinephrine application were amplified 10×, filtered at 1 kHz with
a filter/amplifier (Cygnus Technology, Delaware Water Gap, PA), and
digitized at 2-4 kHz with a TL-1 interface and the pClamp 6.1 suite of
software (Axon Instruments). EPSP responses to 100 µM
norepinephrine were analyzed for changes in frequency and amplitude
with the Datapac program (Run Technologies, Laguna Hills, CA).
Cumulative probability distributions of EPSP amplitude and
instantaneous frequency were generated using an in-house program and
compared with the Kolmogorov-Smirnov test. Population means were
compared with the Student's paired t test and the Wilcoxon
signed rank test for frequency and amplitude values, respectively.
Probability values <0.05 were considered significant. Means are
expressed as ± SE.
Drug application. Norepinephrine, adrenoreceptor
antagonists, tetrodotoxin (TTX), glutamate and glutamate receptor
antagonists were dissolved in aCSF and either bath-applied, or, where
indicated, norepinephrine and glutamate microdrops were applied by
pressure on the surface of the slice using a picospritzer (General
Valve, Fairfield, NJ). Norepinephrine (Sigma, St. Louis, MO) was
bath-applied for 5-15 min at concentrations ranging from 30 µM to 300 µM. Norepinephrine (1 mM) and glutamate (0.5-1 mM) microdrops were
applied under visual control at one or more sites in the PVN using a
patch pipette with a broken tip positioned with a micromanipulator
(Newport Corporation, Irvine, CA). Janus green (0.1%) was added to the norepinephrine and glutamate microdrop solutions to monitor visually the spread of the drops. Adrenoreceptor antagonists included the 1-adrenoreceptor antagonist prazosin hydrochloride (10 µM) and the  -adrenoreceptor antagonist propranolol
hydrochloride (10 µM) (Research Biochemicals
International, Natick, MA); glutamate receptor antagonists included the
NMDA receptor antagonist D,L-2-amino-5-phosphonovalerate (AP5) (100 µM) and the non-NMDA receptor antagonist
5,6-dinitroquinoxaline-2, 3-dione (DNQX; 50 µM) (Tocris
Cookson, Ballwin, MO). Adrenoreceptor and glutamate receptor
antagonists were bath-applied for 15 min before the reapplication of
norepinephrine. Tetrodotoxin (1.5-3 µM) (Sigma) was used
to block voltage-gated sodium channels and spike-mediated transmitter
release. Stock solutions (10 mM) of prazosin hydrochloride
and propranolol hydrochloride were prepared in aCSF and stored in the
dark at  20°C until use.
Biocytin histology and peptide immunohistochemistry.
Biocytin was added to microelectrodes (1%) and to patch pipettes
(0.3-0.5%) as an intracellular marker. The biocytin leaked into the
recorded cells during patch recordings, or in the case of
microelectrode recordings was iontophoresed intracellularly at the end
of experiments by passing negative current pulses (250 pA, 250 msec,
2 Hz) for 5-10 min.
After experiments, slices were removed from the recording chamber and
fixed overnight in 4% paraformaldehyde in 0.1 M PBS at
4°C. They were then sectioned on a freezing microtome at 20-25 µm,
and the biocytin-injected cells were labeled by incubating the
sections for 4 hr in streptavidin-conjugated
7-amino-4-methyl-coumarin-3-acetic acid (AMCA; Molecular Probes,
Eugene, OR). The AMCA was diluted 1:300 in 0.1 M PBS
containing 0.5% Triton X-100. Sections were scanned under a
fluorescence microscope using a UV/420K filter combination to detect
the presence of biocytin-filled, AMCA-labeled neurons.
Sections containing the AMCA-labeled cells were placed in 2% normal
sheep serum in 0.1 M PBS for 15 min. To determine whether the stained cells were oxytocin or vasopressin magnocellular neurons, we used a mixture of a rabbit polyclonal antibody to oxytocin (VA-10)
and a mouse monoclonal antibody to vasopressin-associated neurophysin
(PS41) on the same section. Both antibodies were kindly provided by Dr.
H. Gainer (National Institutes of Health, Bethesda, MD) (Ben-Barak et
al., 1985; Altstein et al., 1988). The polyclonal oxytocin antibody
(1:2000) and the monoclonal vasopressin-associated neurophysin antibody
(1:200) were applied together for 36 hr at 4°C in 0.1 M
PBS + 1% normal sheep serum and 0.2% sodium azide. After treatment
with the primary antibodies, sections were rinsed with 0.1 M PBS, incubated for 1 hr in a mixture of anti-rabbit IgG
conjugated to fluorescein isothiocyanate (FITC, 1:100; Vector Labs,
Burlingame, CA) and anti-mouse IgG conjugated to rhodamine (1:100;
Jackson ImmunoResearch Labs, West Grove, PA), and rinsed again in 0.1 M PBS. They then were mounted, coverslipped, and examined
under 450-490 nm excitation/515 nm barrier filters to detect the
FITC-labeled oxytocin neurons and 515-560 nm excitation/580 nm barrier
filters to see the rhodamine-labeled vasopressin neurons. Recorded
cells were positively identified as either oxytocinergic or
vasopressinergic only if they labeled positive for one of the two
antibodies and negative for the other. No cells were found to be
positively labeled for both antibodies, confirming the specificity of
the antibodies. We have tested the polyclonal oxytocin antibody (VA-10)
for specificity using preabsorption controls (Boudaba et al.,
1996).
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RESULTS |
Putative magnocellular neurons of the PVN were distinguished from
putative parvocellular neurons during recordings based on specific
electrophysiological properties (Hoffman et al., 1991; Tasker and
Dudek, 1991). In particular, magnocellular neurons have a prominent
transient outward rectification, a relatively short membrane time
constant, and linear current-voltage relations.
A total of 136 putative magnocellular neurons were recorded in this
study: 122 cells were tested for their response to norepinephrine and
14 cells for their response to glutamate microdrops. Thirteen of the
cells were recorded with sharp electrodes to control for washout of the
norepinephrine signal; these cells showed the same responses to
norepinephrine as the rest of the cells that were recorded with patch
electrodes. The neurons recorded with sharp electrodes had a mean
membrane potential of 54 ± 2 mV (SE), input resistance of
215 ± 24 M, and action potential amplitude of 65 ± 2 mV
(threshold-peak). The cells recorded with patch electrodes had a mean
membrane potential of 66 ± 1 mV (corrected for a 11 mV
junction potential), input resistance of 871 ± 65 M, and
action potential amplitude of 66 ± 1 mV (n = 58).
The patch solution was weakly hyperosmotic, which resulted in a
hyperpolarized resting potential. Application of norepinephrine caused
a direct depolarization or an increase in EPSPs, or both, in the
majority (60%) of magnocellular neurons recorded.
Direct effect of norepinephrine
Norepinephrine was considered to have a presumptive direct,
postsynaptic effect if it resulted in a sustained change in membrane potential of at least 3 mV that was reversed with washout. Bath application of norepinephrine at concentrations of 30-300
µM elicited a reversible depolarization (7.27 ± 0.6 mV) in 18 of 90 (20%) putative magnocellular neurons tested (Fig.
1A). The depolarization occurred within ~3-7 min of norepinephrine introduction into the recording chamber and was accompanied by a decrease in input resistance (15 ± 2%) in 4 of 16 cells tested. The response was not blocked by TTX (1.5-3 µM, n = 3), suggesting
that it was not mediated by an increase in spike-mediated transmitter
release. It was blocked or reduced by the
1-adrenoreceptor antagonist prazosin (10 µM, n = 4) but was not affected by the
-adrenoreceptor antagonist propranolol (10 µM,
n = 3) (data not shown). The direct response to
norepinephrine was qualitatively similar to that described in
magnocellular neurons of the supraoptic nucleus (Randle et al., 1986a)
and was not investigated further in this study.
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Figure 1.
Norepinephrine responses of PVN magnocellular
neurons. A, Direct depolarization elicited by
norepinephrine. A putative magnocellular neuron responded to bath
application of norepinephrine (300 µM) with a 10 mV
depolarization; no apparent change in input resistance was seen (data
not shown). This cell had a resting membrane potential of 54 mV.
Positive spikes are action potentials, which were truncated by digital
filtering. Bar indicates the duration of the norepinephrine
application. B, Increased EPSPs in norepinephrine. Bath
application of norepinephrine (100 µM) elicited an
increase in EPSPs (Norepinephrine) with a latency of
~8 min in a putative magnocellular neuron recorded at resting
potential. The response reversed after 20 min in normal aCSF
(Wash). This cell had a resting membrane potential of
60 mV. Bottom traces are expanded recordings of the
periods in the top traces indicated by the
bars. C, Cumulative probability plots of
the EPSP frequency and amplitude distributions in a representative
neuron in control aCSF ( ) and during norepinephrine application
( ). There is a significant shift toward higher instantaneous
frequencies and larger amplitudes of the EPSPs in norepinephrine
(p < 0.01; n = 5;
Kolmogorov-Smirnov test). D, Changes in mean frequency
and amplitude of EPSPs in norepinephrine. Mean frequencies and
amplitudes were calculated in control medium and in 100 µM norepinephrine. These values were averaged, and the
differences were expressed as percentage increase in norepinephrine.
There was a 147% average increase in the mean frequency
(n = 22) and a 53% average increase in the mean
amplitude (n = 10) of EPSPs in
norepinephrine.
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Norepinephrine activation of presynaptic glutamate neurons
Bath application of norepinephrine (30-300 µM)
caused a large increase in EPSPs in 38 of 90 (42%) putative
magnocellular neurons tested in the whole slice (Fig.
1B). An increase in EPSPs was indicated by a
qualitatively detectable rise in the frequency (i.e., of at least
10-20%) of positive-going synaptic potentials recorded at resting
membrane potential. The increase in EPSPs occurred ~7-8 min after
introduction of the norepinephrine into the recording chamber, was
maintained throughout the application, and reversed within 10-33 min
of washout of the norepinephrine. Neurons responding to 100 µM norepinephrine were selected for EPSP frequency and
amplitude analysis. Norepinephrine caused a significant increase in
both the frequency (p < 0.001;
n = 22; Student's paired t test) and the
amplitude (p < 0.05; n = 10; Wilcoxon signed rank test) of spontaneous EPSPs collected in 30-60 sec
episodes. This was seen in individual cells as a significant shift in
the cumulative EPSP frequency and amplitude distributions (p < 0.01; n = 5;
Kolmogorov-Smirnov test) (Fig. 1C). The mean percentage
increases in EPSP frequency (n = 22) and amplitude (n = 10) were 145 and 52%, respectively (Fig.
1D). The marked rise in the frequency of EPSPs
suggests that norepinephrine increases the probability of transmitter
release from presynaptic excitatory neurons. The moderate increase in
EPSP amplitude suggests that it may also modulate the postsynaptic
responsiveness of the magnocellular neurons, although this effect is
less robust. No apparent desensitization of the synaptic response was
observed during the norepinephrine application or with a second
application of norepinephrine (n = 3). Of the 38 cells
that showed an increase in EPSPs, eight cells also depolarized in
response to norepinephrine.
The norepinephrine-induced increase in EPSPs was blocked by the
1-adrenoreceptor antagonist prazosin hydrochloride (10 µM) in six of six cells tested (Fig.
2). The -receptor antagonist propranolol hydrochloride (10 µM) failed to block the
EPSPs elicited by norepinephrine (n = 5). Thus,
norepinephrine appears to activate presynaptic excitatory neurons by
acting at 1-receptors.
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Figure 2.
The norepinephrine-induced increase in EPSPs is
mediated by 1-receptor activation. Bath application of
norepinephrine (100 µM) caused a reversible increase in
the frequency of EPSPs (Norepinephrine) in a putative
magnocellular neuron. A second norepinephrine application in the
presence of the 1-receptor antagonist, prazosin
hydrochloride (10 µM; Norepinephrine in
Prazosin), failed to elicit the increase in EPSPs, suggesting
that the response was mediated by the activation of
1-adrenoreceptors. This cell had a resting membrane
potential of 64 mV. The cell was subsequently found to be
immunopositive for oxytocin and immunonegative for vasopressin, which
is shown in Figure 7.
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To test whether norepinephrine caused an increase in EPSPs by acting at
presynaptic neurons, we applied TTX (1.5-3 µM) to block
spike-mediated transmitter release. Bath application of TTX blocked
completely the norepinephrine-evoked increase in EPSPs in eight of
eight cells tested (Fig. 3). This
indicated that the increased EPSPs were caused by an increase in
spike-evoked transmitter release and suggested that norepinephrine was
acting at receptors at the somatic/dendritic region of presynaptic
excitatory neurons. In the cells that responded with an increase in
EPSPs, norepinephrine had no apparent effect on the depolarizing
voltage responses to positive current pulses (n = 4),
suggesting that norepinephrine was not acting postsynaptically to
amplify spontaneous EPSPs by enhancing or attenuating voltage-gated
currents.
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Figure 3.
The norepinephrine-induced increase in EPSPs is
caused by activation of local presynaptic excitatory neurons.
Norepinephrine (100 µM) elicited a reversible increase in
EPSPs (Norepinephrine) in a putative magnocellular
neuron recorded at a resting membrane potential of 55 mV. The
norepinephrine-induced EPSPs were blocked by TTX (Norepinephrine
in TTX), suggesting that they were caused by the
activation of presynaptic neuronal somata/dendrites, resulting in an
increase in spike-mediated release of excitatory neurotransmitter. That
the presynaptic neurons were intact in the slice suggests that they
were cells with short axonal projections located in relative proximity
to the recorded cell.
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That norepinephrine application led to an increase in the frequency of
fast EPSPs suggested that it was acting to stimulate presynaptic
glutamate neurons, causing an increase in glutamate release and
activation of postsynaptic ionotropic glutamate receptors. We tested
this hypothesis by bath applying the NMDA and non-NMDA receptor
antagonists AP5 and DNQX, respectively, to block ionotropic glutamate
receptors. In cells that had shown a norepinephrine-evoked increase in
EPSPs, AP5 and DNQX blocked completely the synaptic response to
norepinephrine in six of six cells tested (Fig.
4). Thus, the presynaptic neurons that
responded to norepinephrine with a spike-mediated increase in
transmitter release onto PVN magnocellular neurons were glutamatergic,
and the synaptic responses were caused by glutamate activation of
ionotropic glutamate receptors.
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Figure 4.
The norepinephrine-induced increase in EPSPs is
mediated by presynaptic glutamate neurons. Norepinephrine (100 µM) caused an increase in the frequency of EPSPs in a
magnocellular neuron. The response reversed after 21 min in wash (data
not shown). The EPSPs elicited by norepinephrine were completely
blocked by the NMDA and non-NMDA receptor antagonists AP5 (100 µM) and DNQX (50 µM), suggesting that they
were mediated by the release of glutamate and the activation of
ionotropic glutamate receptors. This cell had a resting membrane
potential of 63 mV.
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Intranuclear localization of the presynaptic glutamate neurons
The TTX experiments suggested that the presynaptic glutamate
neurons were present and intact in our slices, because the
norepinephrine was probably acting at the somatic/dendritic regions of
the presynaptic cells to cause action potential generation. This, along
with the relatively high percentage of magnocellular neurons (42%) in
our slices that responded to norepinephrine with an increase in EPSPs, indicated that the presynaptic glutamate neurons were in close proximity to the magnocellular neurons, possibly inside the PVN.
We conducted three experiments to test this hypothesis. The first
experiment tested the effects of norepinephrine on the incidence of
EPSPs in PVN magnocellular neurons in a slice preparation in which the
PVN was surgically isolated by cutting away and removing the rest of
the slice. In this preparation, bath application of norepinephrine
caused an increase in the frequency of EPSPs in 7 of 11 (64%) putative
magnocellular neurons tested (Fig. 5). Next, glutamate (0.5-1 mM) was applied as microdrops
directly into the PVN to stimulate neurons focally within the PVN
without activating axons of passage (Christian and Dudek, 1988).
Glutamate microstimulation led to an increase in EPSPs in 4 of 14 (28%) putative magnocellular neurons tested (data not shown),
suggesting the presence of intranuclear excitatory circuits. Finally,
to determine whether these intranuclear excitatory circuits were the
same circuits as those activated by bath application of norepinephrine, we applied microdrops of norepinephrine (1 mM) directly
into the PVN. Norepinephrine microdrops also elicited an increase in
EPSPs in 7 of 21 (33%) putative magnocellular neurons tested (Fig.
6). Four of the seven (57%) neurons also
exhibited a depolarization in response to norepinephrine. The results
of these experiments all point to the presence of glutamate neurons
within or very closely apposed to the PVN that are excited by
norepinephrine and that send intranuclear projections to magnocellular
neurons. Stimulation of these local glutamate interneurons via
1-receptor activation results in an increase in
excitatory synaptic input to magnocellular neurons.
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Figure 5.
Norepinephrine-evoked increase in EPSPs in an
isolated PVN preparation. A putative magnocellular neuron was recorded
in a slice preparation in which the PVN had been surgically isolated
from the rest of the slice (inset). Bath application of
norepinephrine (100 µM) elicited a robust increase in the
frequency of EPSPs (Norepinephrine), and the response
reversed after 20 min in regular aCSF (Wash). This cell
had a resting membrane potential of 65 mV. PVN,
Paraventricular nucleus; Fx, fornix.
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Figure 6.
Norepinephrine microdrops in the PVN evoked an
increase in EPSPs. Norepinephrine microdrop application in the PVN
(inset) elicited a robust increase in EPSPs recorded in
a putative magnocellular neuron. This effect was accompanied by a
depolarization of the cell from its resting membrane potential of 64
to 55 mV. Bottom traces are expanded recordings of the
periods in the top traces indicated by the
bars. Top time calibration applies to top trace; bottom
time calibration applies to expanded traces. NE,
Norepinephrine; PVN, paraventricular nucleus;
Fx, fornix; 3V, third ventricle.
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Immunohistochemical identification of magnocellular neurons
A total of 20 putative magnocellular neurons were recovered after
biocytin labeling and oxytocin/vasopressin immunohistochemical processing with antibodies to oxytocin and vasopressin neurophysin. Only those cells that were immunopositive for one and negative for the
other of the two antibodies were counted. Of the eight cells that
responded to norepinephrine with a depolarization, five were
immunopositive for oxytocin and immunonegative for vasopressin, and
three were immunopositive for vasopressin and immunonegative for
oxytocin. Of the 12 cells that responded to norepinephrine with an
increase in EPSPs, three were immunopositive for vasopressin and
immunonegative for oxytocin (Fig. 7A), and nine were
immunopositive for oxytocin and immunonegative for vasopressin (Fig.
7B). These results suggest that both oxytocin and
vasopressin magnocellular neurons express functional
1-adrenoreceptors and receive intranuclear excitatory
synaptic inputs from glutamate neurons located within the PVN.
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Figure 7.
Immunohistochemical double-labeling of
norepinephrine-responsive oxytocin and vasopressin magnocellular
neurons in the PVN. Magnocellular neurons that responded to
norepinephrine with an increase in EPSPs were injected with biocytin
and immunohistochemically processed with antibodies to oxytocin and
vasopressin-associated neurophysin. A, A
biocytin-injected, AMCA-labeled cell (A1,
arrow) was visualized under the AMCA filter combination
(Biocytin). The same section was visualized under
rhodamine filters to detect the vasopressinergic neurons
(Vasopressin) and under FITC filters to see the
oxytocinergic neurons (Oxytocin). This cell was
immunopositive for vasopressin (A2,
arrow) and immunonegative for oxytocin
(A3, arrow), indicating that it was a
vasopressinergic magnocellular neuron. B, Another
magnocellular neuron that responded to norepinephrine with an increase
in EPSPs (recordings shown in Fig. 2) and was labeled with biocytin
(B1, arrow) was immunonegative for
vasopressin (B2, arrow) and
immunopositive for oxytocin (B3, arrow),
indicating that it was an oxytocinergic magnocellular neuron. The
FITC-oxytocin label bleeds through the UV filters used to visualize
the biocytin-AMCA label but is readily distinguished from the AMCA
label by its intensity and its color (seen as gray scale
here). 200× magnification.
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DISCUSSION |
Despite the apparent prominent role of norepinephrine in the
control of oxytocin and vasopressin neuronal activity and hormone release, little is known about the mechanisms or the locus of the
excitatory actions of norepinephrine in the PVN. This is the first
study to characterize the physiological actions of norepinephrine in
identified oxytocin and vasopressin neurons. Electrophysiological distinction between magnocellular and parvocellular neurons combined with intracellular dye injection and post hoc
immunohistochemical double-labeling with selective antibodies for
oxytocin and vasopressin-associated neurophysin provided a reliable
means of identifying PVN magnocellular neurons.
Norepinephrine excited PVN magnocellular neurons either directly by
membrane depolarization or indirectly via an increase in EPSPs. The
depolarizing effect of norepinephrine was mediated by
1-receptor activation and was similar qualitatively to
that described in magnocellular neurons of the SON (Randle et al., 1986a), and it was therefore not characterized in further detail in
this study. The norepinephrine-induced increase in synaptic activity in
magnocellular neurons, on the other hand, has not been reported
previously and was the primary focus of this study.
Several observations indicate that the increase in EPSPs was caused
mainly by a presynaptic action of norepinephrine. The norepinephrine-induced synaptic response was TTX-sensitive and was
characterized by a robust increase in the frequency of EPSPs, suggestive of an enhanced probability of transmitter release. The
majority of the cells that responded to norepinephrine with an increase
in EPSPs (79%) did not show any change in resting membrane potential
or input resistance, indicating that norepinephrine was not acting on
postsynaptic conductances active at resting potential. Similarly,
norepinephrine had no effect on depolarizing voltage responses to
positive current pulses in these cells, suggesting that it was not
acting on postsynaptic voltage-gated conductances. However, the
norepinephrine-induced increase in EPSP amplitudes suggests that
norepinephrine may also have a postsynaptic modulatory effect on
ionotropic glutamate receptor-mediated currents in these cells, but
these actions of norepinephrine appear less robust than the presynaptic actions.
The norepinephrine-induced increase in EPSPs was most likely due to the
activation of receptors located in the somatic/dendritic region of
presynaptic neurons. The response was blocked completely by TTX and was
therefore dependent on spike generation, presumably at the initial
segment of the axon. That the response was not mediated by modulation
of spike-dependent conductances at presynaptic terminals or preterminal
axons (Lena et al., 1993) is suggested by our experiments involving
glutamate microdrops, whose excitatory actions should be restricted to
the somatic/dendritic membrane (Christian and Dudek, 1988; Schrader and
Tasker, 1997). Glutamate microdrops elicited EPSPs in a proportion of
PVN cells comparable to the proportion of cells that responded to
norepinephrine microdrops (28 vs 33%, respectively). If the glutamate
and norepinephrine microdrops acted on the same presynaptic cells,
which seems a likely possibility, then the norepinephrine, like the
glutamate, was probably acting at presynaptic somatic/dendritic receptors.
Our data suggest that norepinephrine activated presynaptic glutamate
interneurons located within, or in very close proximity to, the PVN.
The fast kinetics of the EPSPs and their sensitivity to ionotropic
glutamate receptor antagonists indicate that they were mediated by
glutamate release. The glutamatergic synaptic inputs originated in
neurons located inside the PVN or very close to the periphery of the
nucleus because both norepinephrine microdrops applied within the PVN
and bath application of norepinephrine in a surgically isolated PVN
elicited EPSPs. The percentage of cells responding to norepinephrine
with augmented EPSPs increased in the isolated PVN preparation, from
~40 to >60%, which could be explained by excitatory actions of
norepinephrine on perinuclear inhibitory neurons (Boudaba et al., 1996)
that innervate the presynaptic glutamate neurons in the PVN. Severing
these projections with surgical isolation of the PVN could lead to
disinhibition of the PVN glutamate interneurons.
Twenty of the norepinephrine-responsive magnocellular neurons were
identified immunohistochemically as oxytocinergic or vasopressinergic (i.e., labeled positive for one peptide and negative for the other). Approximately 63% of the neurons that responded to norepinephrine with
a direct depolarization (five of eight) and 75% of the neurons that
responded with an increase in EPSPs (9 of 12) were oxytocinergic, whereas the remaining cells, 37% and 25%, respectively, were
vasopressinergic. It is not possible from these data to ascertain
whether this represents a preferential responsiveness of oxytocin
neurons to norepinephrine inputs or a sampling bias in our recordings.
However, it is clear that both oxytocin and vasopressin neurons of the
PVN express functional 1-adrenoreceptors and receive
local excitatory synaptic inputs from
1-adrenoreceptor-expressing glutamate neurons in the
PVN. A schematic diagram of a proposed model circuit is presented in
Figure 8.
View larger version (15K):
[in this window]
[in a new window]
|
Figure 8.
Model of the regulation of magnocellular neurons
by norepinephrine. Norepinephrine projections, probably from the A1/A2
noradrenergic cell groups in the medulla, activate glutamatergic
interneurons (GLU) within the PVN that send
intranuclear excitatory projections to oxytocinergic and
vasopressinergic magnocellular neurons (OXY/VP).
Norepinephrine acts via 1-adrenoreceptors to excite the
glutamate interneurons, which leads to an increase in ionotropic
receptor-mediated EPSPs in oxytocin and vasopressin neurons. PVN
oxytocin and vasopressin neurons also receive direct noradrenergic
inputs and express 1-adrenoreceptors. 3V,
Third ventricle.
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|
The intranuclear origin of the glutamatergic inputs to magnocellular
neurons raises the question of the identity of the presynaptic cells.
There are several possibilities, including a separate population of
glutamatergic interneurons, a subtype of PVN parvocellular neuron that
co-expresses glutamate, and other magnocellular neurons that co-express
glutamate along with vasopressin or oxytocin. In the course of this
study, we recorded the responses of both putative magnocellular and
putative parvocellular neurons to norepinephrine, and interestingly,
very few putative parvocellular neurons (~2%) were depolarized by
norepinephrine (Daftary et al., 1996) [note that all nonmagnocellular
neurons located in the PVN were classified as parvocellular neurons;
Hoffman et al. (1991)]. In contrast, 23% of the putative
magnocellular neurons showed a relatively robust depolarization
(7.27 ± 0.6 mV) in response to norepinephrine, as described
above. This depolarization was strong enough in some of the neurons to
generate a train of action potentials (Fig. 1A),
which if the cells co-released glutamate would induce a robust increase
in EPSPs postsynaptically, similar to the synaptic response to
norepinephrine reported here. Although more experiments are necessary
to determine the identity of the presynaptic neurons, our current
working hypothesis is that the glutamatergic inputs to oxytocin and
vasopressin neurons arise either from other magnocellular neurons or
from a separate, sparsely distributed population of glutamatergic
interneurons in the PVN.
Indirect noradrenergic activation of oxytocin neurons via glutamate
interneurons could account for seemingly paradoxical findings from both
anatomic and physiological studies. Although there is compelling
evidence that noradrenergic afferents and norepinephrine play a
critical role in the control of oxytocin and vasopressin release
(Tribollet et al., 1978; Clarke et al., 1979; Randle et al., 1986b;
Crowley and Armstrong, 1992), several immunohistochemical studies have
suggested that noradrenergic inputs are less densely concentrated in
intranuclear regions occupied by oxytocin neurons than those in which
vasopressin neurons predominate (McNeill and Sladek, 1980; Swanson et
al., 1981; Hornby and Piekut, 1987; Cunningham and Sawchenko, 1988;
Ginsberg et al., 1994; although see Michaloudi et al., 1997). Our
current findings provide a physiological correlate to recent
observations of noradrenergic synapses directly on both oxytocin and
vasopressin neuronal somata (Michaloudi et al., 1997), but they also
suggest that many of the oxytocin and vasopressin neurons in the PVN
receive an indirect input from noradrenergic afferents by way of
glutamatergic relay cells. This is consistent with the facilitatory
interaction between adrenoreceptor and glutamate-receptor mechanisms on
oxytocin release described in the lactating rat (Parker and Crowley,
1993b). These data also are in line with the observation that
norepinephrine levels in the SON and PVN show a prolonged increase
before and during the parturition-associated release of oxytocin,
whereas glutamate levels rise sharply just before oxytocin release and
subside rapidly (Herbison et al., 1997). Taken together, these findings
suggest that noradrenergic afferents may be activated by sensory inputs
in a tonic or slow phasic manner and that after a latency caused by an
as-yet-unknown gating mechanism at the level of the PVN (and SON?),
glutamate interneurons are stimulated to fire abruptly to trigger
oxytocin neuron activation and bolus release of oxytocin. Parker and
Crowley (1993b) found that the increase in oxytocin release caused by norepinephrine application in the SON in vivo was blocked by
an AMPA receptor antagonist, but also that the glutamate-induced increase in oxytocin release was attenuated by an adrenoreceptor antagonist. This suggests that the situation is probably more complicated than the simple monosynaptic and disynaptic PVN circuits presented in Figure 8.
Although the role of the intranuclear glutamate circuits in the control
of oxytocin and vasopressin release is not yet known, their presence
provides a significant potential mechanism for the generation and
coordination of the patterned electrical activity seen in oxytocin and
vasopressin neurons. A recent report showed that local glutamatergic
circuits in hypothalamic slice cultures are capable of driving bursting
activity in individual oxytocin neurons (Jourdain et al., 1998). In
addition to serving as a pattern generator, a small group of glutamate
interneurons could provide synchronizing inputs to oxytocin neurons in
the PVN, and if these glutamatergic projections extended to the other
magnocellular nuclei, they could drive the synchronous bursting
activity characteristic of these neurons during the milk ejection
reflex and parturition. Future studies will need to determine whether
similar intranuclear glutamatergic circuits are present in lactating
female rats and whether glutamatergic projections serve to interconnect
the magnocellular nuclei.
| |
FOOTNOTES |
Received July 16, 1998; revised Sept. 8, 1998; accepted Sept. 22, 1998.
This work was funded by National Institute of Neurological Disorders
and Stroke Grant NS31187. S.S.D. was partially supported by a
predoctoral fellowship from the Louisiana American Heart Association.
We thank Dr. H. Gainer for supplying oxytocin and vasopressin
antibodies, Dr. A. Fancsik for his help with the computer analyses, and
Drs. Shi Di and Andrei Belousov for their critical reading of this manuscript.
S.S.D. and C.B. contributed equally to this study
Correspondence should be addressed to Jeffrey Tasker, Department of
Cell and Molecular Biology, Tulane University, New Orleans, LA 70118.
| |
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Top
Abstract
Introduction
