Department of Neurology and Neurosurgery, Montreal Neurological
Institute, McGill University, Montreal, Quebec H3A 2B4, Canada
 |
INTRODUCTION |
The thalamus receives a
massive input from the neocortex via corticothalamic fibers. The number
of corticothalamic fibers is one order of magnitude larger than the
number of thalamocortical axons, and cortical afferents to the thalamus
largely outnumber the sensory input from peripheral receptors
(Guillery 1969
; Sherman and Guillery
1996
; Steriade et al. 1997
). In the
somatosensory system the main type of corticothalamic fiber originates
in layer VI and projects to the ventrobasal thalamus (ventral posterior medial and lateral nuclei), leaving a fiber collateral in the reticular
nucleus (nRt) (Zhang and Deschenes 1997
).
Corticothalamic pathways have been proposed to play a role in modifying
the size, strength, and selectivity of thalamocortical receptive fields and/or to establish cortico-cortical communication via the thalamus (Contreras et al. 1996
; Diamond 1995
;
Ergenzinger et al. 1998
; Guillery 1995
;
Krupa et al. 1999
; Sillito et al. 1994
;
Weinberger 1995
; Yuan et al. 1985
). Short
periods of repetitive stimulation of corticothalamic fibers at
frequencies above 2 Hz produces strong synaptic facilitation or
short-term potentiation in vitro (Castro-Alamancos and
Calcagnotto 1999
; McCormick and von Krosigk
1992
; Scharfman et al. 1990
; Turner and
Salt 1998
; von Krosigk et al. 1999
) and in vivo
(Deschenes and Hu 1990
; Frigyesi 1972
;
Lindstrom and Wrobel 1990
; Mishima
1992
; Steriade 1999
; Steriade and
Timofeev 1997
; Steriade and Wyzinski
1972
; Tsumoto et al. 1978
). A longer period of repetitive stimulation at 10 Hz induces long-term
potentiation, while stimulation at 1 Hz induces long-term depression
(Castro-Alamancos and Calcagnotto 1999
). Thus
corticothalamic synapses possess mechanisms to modify the effectiveness
with which the neocortex can influence the thalamus. In addition to
activity-dependent changes, corticothalamic synapses may be subject to
neuromodulator-dependent changes (Gil et al. 1997
;
Isaacson et al. 1993
; Miller 1998
;
Scanziani et al. 1997
; Thompson et al.
1993
; Wu and Saggau 1997
). Several
neuromodulatory systems from the brain stem and basal forebrain
innervate the thalamus. In fact, neuromodulatory synapses from the
brain stem and corticothalamic synapses may provide the two main
synaptic inputs to the thalamus (Erisir et al. 1997
).
Cholinergic and noradrenergic fibers project densely to the ventrobasal
thalamus and nRt (Asanuma 1997
; Steriade
et al. 1997
). Neurons from these neuromodulatory systems
discharge vigorously during behavioral activation (Aston-Jones et al. 1991
; Buzsaki et al. 1988
), producing
effects on the firing properties of thalamic neurons that have been
extensively described (McCormick 1992
;
Steriade et al. 1997
). In contrast, their effects on
synaptic transmission in the thalamus are less understood.
 |
METHODS |
In vitro methods
Thalamocortical slices were prepared from adult (
7 wk) BALB/C
mice as previously described (Agmon and Connors 1991
;
Castro-Alamancos and Calcagnotto 1999
). Slices were cut
in ice-cold buffer using a vibratome and kept in a holding chamber for
a least 1 h. Experiments were performed in an interface chamber at
32°C. The slices were perfused constantly (1-1.5 ml/min) with
artificial cerebrospinal fluid (ACSF) containing (in mM) 126 NaCl, 3 KCl, 1.25 NaH2Po4, 26 NaHCO3, 1.3 MgSO4
7H2O, 10 dextrose, and 2.5 CaCl2 2H2O. The ACSF was
bubbled with 95% O2-5%
CO2. Synaptic responses were evoked using a
concentric stimulating electrode placed in the thalamic radiation
unless otherwise indicated. The stimulus consisted of a 200-µs pulse
of <50 µA. Field recordings from the ventrobasal thalamus were made
using a low-impedance pipette (~0.5 M
) filled with ACSF containing
400 µM bicuculline methobromide (BMI) unless otherwise indicated.
Intracellular recordings were performed using sharp electrodes (60-80
M
) filled with Cs+-acetate (1 M) and QX-314
(50 mM) to suppress K+ and
Na+ currents and postsynaptic
GABAB responses. In some experiments, the
GABAB receptor antagonist CGP35348 (100 µM) was
also included in the bath. GABAA receptors were
blocked using bath application (10-20 µM) or local application of
BMI from the extracellular recording electrode or bath application of
picrotoxin (10-20 µM). The test stimulus was delivered at 0.05 Hz
and was either single or a pair with a 50-ms interstimulus interval to
evaluate paired-pulse facilitation or four stimuli delivered at
different frequencies to perform a spectrum analysis. Input resistance
was measured by applying a 50- to 100-ms negative current pulse (0.3 nA). Group data are expressed as means ± SD. For single
experiments, which represent typical examples, every response at 0.05 Hz is displayed. To ensure that activity in cortical circuits did not
feed back to the thalamus, we severed all connections between thalamus
and neocortex with a cut just below the cortical white matter
(Castro-Alamancos and Calcagnotto 1999
). Drugs were
tested using bath application for 5-10 min unless otherwise indicated.
They were prepared fresh and protected from light and from oxidation
(40 µM ascorbic acid in the ACSF) as required.
In vivo methods
Adult Spague-Dawley rats (300 g) were anesthetized with urethan
(1.5 g/kg ip) and placed in a stereoaxic frame. All skin incisions and
frame contacts with the skin were injected with lidocaine (2%). A
unilateral craniotomy extended over a large area of the parietal
cortex. Small incisions were made in the dura as necessary, and the
cortical surface was covered with ACSF. Body temperature was
automatically maintained constant with a heating pad. The level of
anesthesia was monitored with field recordings and limb-withdrawal reflexes and kept constant at about stage III/3 using supplemental doses of urethan (Friedberg et al. 1999
). Electrodes
were inserted with stereotaxic procedures (all coordinates are given in
mm, in reference to bregma and the dura) (Paxinos and Watson
1992
). Coordinates for the ventrobasal recording electrode were
anterior-posterior =
3.5, lateral = 3, and depth = 5-6. Coordinates for the thalamic radiation stimulating electrode were
anterior-posterior =
2, lateral = 4, and depth = 3-4.
Coordinates for the brain stem reticular formation stimulating
electrode were anterior-posterior =
9, lateral = 0.7, and
depth = 5-6. This stimulating electrode was placed close to the
laterodorsal tegmentum and locus coeruleus to activate both cholinergic
and noradrenergic fibers innervating the thalamus. Stimulation of the
thalamic radiation consisted of four 200-µs pulses of <200 µA
delivered at 0.1, 0.5, 1, 2, 5, 10, 20, or 40 Hz. Stimulation of the
brain stem reticular formation consisted of 200-µs pulses of <300
µA delivered at 100 Hz for 1 s. A microdialysis probe was placed
in the thalamus 0.5-1 mm medial from the thalamic recording electrode
as previously described (Castro-Alamancos 1999
). This
allowed infusing drugs into the thalamus during recordings.
6-Cyano-7-nitroquinoxaline-2,3-dione disodium (CNQX) was applied at 200 µM in the ACSF. The muscarinic antagonist scopolamine and the
-adrenergic antagonist phentolamine were applied at 500 µM in the
ACSF. The Animal Care Committee of McGill University approved protocols
for all experiments.
 |
RESULTS |
Field and intracellular potentials were recorded from neurons of
the ventrobasal thalamus in brain slices of adult mice. Orthodromic stimuli applied to the thalamic radiation evoked a negative potential in field recordings from populations of neurons and an excitatory postsynaptic potential (EPSP) in intracellular recordings from individual neurons of the ventrobasal thalamus (Fig.
1). Some field potential recordings also
display a fiber volley immediately before the synaptic negativity (Fig.
1, arrows). The fiber volley follows high-frequency stimulation and is
abolished by tetrodotoxin, but not by glutamate receptor antagonists
(Castro-Alamancos and Calcagnotto 1999
). The field and
intracellular EPSPs reflect a monosynaptic excitatory connection
between corticothalamic fibers and neurons in the ventrobasal thalamus
(Castro-Alamancos and Calcagnotto 1999
). We explored the
effects of acetylcholine and norepinephrine on corticothalamic EPSPs.
To reduce the postsynaptic effects of these neuromodulators,
Na+ and K+ currents were
suppressed with the intracellular recording solution, and, to assure
that the effects were not mediated by inhibition, we blocked
GABAA and postsynaptic
GABAB receptors. Thus while most postsynaptic
actions of the neuromodulators will be blocked in the
intracellular EPSPs, the extracellular field EPSPs will represent a combination of presynaptic and postsynaptic effects. The
suppression of Na+ and K+
currents was indicated by the following observations. First, the
membrane potential of neurons recorded using Cs+
and QX-314 (
51 ± 6 mV, mean ± SD) was significantly more
depolarized than that of neurons recorded using
K+-acetate (
73 ± 2 mV). Further
depolarization from these values was impeded by application of negative
current. Second, the input resistance of neurons recorded using
Cs+ and QX-314 (100-150 M
) is greater than in
thalamic neurons recorded using K+-acetate
(40-60 M
). Third, action potentials were completely abolished, and
strong positive current pulses evoked only large depolarizing plateau
potentials. Finally, application of the neuromodulators was performed
after extensive diffusion of the intracellular solution by waiting long
periods after impalement (more than 2 h for the neurons shown in
Fig. 1). Under these conditions, application of either acetylcholine
(1-10 mM) or norepinephrine (10-100 µM) caused corticothalamic
EPSPs to depress as reflected in the extracellular and intracellular
evoked response (n = 3-5; Fig. 1). Corticothalamic EPSPs returned to their baseline amplitude after wash out of the drugs.
The depression of corticothalamic EPSPs was not accompanied by a change
in the field potential fiber volley, nor was it followed by a change in
input resistance or baseline membrane potential (Fig. 1). To further
eliminate the contribution of inhibition via the nRt, we produced a cut
that excised this nucleus from the ventrobasal thalamus in several
experiments. This assured that the norepinephrine was not depolarizing
nRt cell bodies (McCormick 1992
) to increase GABA
release in the ventrobasal thalamus and depress corticothalamic EPSPs
via presynaptic GABAB receptors. In this case
corticothalamic stimulation was delivered at the nRt-ventrobasal
thalamus border. To further block GABAB
receptors, in some experiments we also applied CGP35348 (100 µM) in
the bath. These manipulations did not interfere with the depression
exerted by acetylcholine or norepinephrine (n = 3 per
drug).

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Fig. 1.
Acetylcholine and norepinephrine depress corticothalamic
excitatory postsynaptic potentials (EPSPs) in vitro.
The effects of acetylcholine (left) and of norepinephrine
(right) on the amplitude of the intracellular EPSP, the
amplitude of the field EPSP, the input resistance and membrane
potential of the neurons, and on the fiber volley (arrow) are
subsequently displayed. The traces represent intracellular and field
EPSPs recorded in the ventrobasal thalamus in response to stimulation
of the thalamic radiation before and during application of
acetylcholine and norepinephrine. Application of acetylcholine
(ACh, 10 mM) and norepinephrine (NE, 100 µM) reversibly depressed
corticothalamic EPSPs, with no significant effects on the neuron's
input resistance, membrane potential, or the fiber volley. The
bottom graphs represent the average effects of these drugs
on 3-5 experiments per group as compared with baseline responses
(* P < 0.01, t-test). Intracellular
recordings were performed with QX-314 and Cs+ in
the pipette. Bicuculline methobromide (BMI) was bath applied.
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To test via which receptors acetylcholine (muscarinic or nicotinic) and
norepinephrine (
- or
-adrenergic) depress corticothalamic EPSPs,
we used receptor agonists to mimic the effects of these neuromodulators. We also used receptor antagonists to block the effects
of the neuromodulators. Application of a cholinergic receptor agonist
(carbachol; 5 µM, n = 7) or a muscarinic receptor
agonist (muscarine; 10 µM, n = 5) depressed
corticothalamic field EPSPs (Fig. 2).
Nicotine (10 µM, n = 5) or dimethylphenylpiperazinium (10 µM, n = 5), which activate nicotinic receptors,
had no significant effects. Application of a muscarinic receptor
antagonist (scopolamine; 10 µM, n = 5) abolished the
depression of corticothalamic field EPSPs induced by acetylcholine (10 mM). This indicates that acetylcholine depresses corticothalamic
synapses by activating muscarinic receptors. Application of an
2-adrenergic receptor agonist (clonidine,
10-40 µM, n = 5) also depressed corticothalamic
field EPSPs (Fig. 2). In contrast, an
1-adrenergic receptor agonist (phenylephrine; 5-50 µM, n = 5) and a
-adrenergic receptor
agonist (isoproterenol; 10-50 µM, n = 6) were
ineffective. Application of an
-adrenergic receptor
antagonist (phentolamine; 100 µM, n = 5) blocked the depression of corticothalamic field EPSPs exerted by norepinephrine. This indicates that norepinephrine depresses corticothalamic synapses by activating
2-adrenergic receptors.

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Fig. 2.
Activation of muscarinic and 2-adrenergic receptors
depress corticothalamic EPSPs in vitro. The effects of cholinergic
(left) and of noradrenergic (right)
receptor agonists and antagonists on field EPSPs are displayed
subsequently. A: the traces correspond to field
responses recorded before and during application of the drug in the
experiment displayed below. The cholinergic agonist (carbachol, 5 µM)
depressed corticothalamic responses. Nicotine (10 µM) did not
significantly affect corticothalamic responses. Application of a
muscarinic antagonist (scopolamine, 10 µM) blocked the effects of
acetylcholine (ACh, 10 mM) on corticothalamic responses. The
2-adrenergic agonist (clonidine, 40 µM) depressed
corticothalamic responses. The -adrenergic agonist (isoproterenol,
50 µM) did not significantly affect corticothalamic responses.
Application of an -adrenergic antagonist (phentolamine, 100 µM)
blocked the effects of norepinephrine (NE, 100 µM) on corticothalamic
responses. B: average effects of the drugs on 5-7
experiments per group as compared with baseline responses
(* P < 0.01, t-test). The
left panel displays the effects of acetylcholine (10 mM), carbachol (5 µM), nicotine (10 µM), and acetylcholine in the
presence of scopolamine (ACh + Scop; 10 mM + 10 µM). The right
panel displays the effects of norepinephrine (NE, 100 µM),
clonidine (40 µM), phenylephrine (50 µM), isoproterenol (50 µM),
and norepinephrine in the presence of phentolamine (NE + Phent; 100 µM + 100 µM). BMI was in the recording pipette.
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Although acetylcholine and norepinephrine depress corticothalamic
EPSPs, it is possible that their characteristic frequency-dependent facilitation increases. This could happen if corticothalamic depression is due to a decrease in neurotransmitter release probability. Synapses
with a low release probability display stronger facilitation than
synapses with a high release probability (Debanne et al. 1996
; Dobrunz and Stevens 1997
;
Fisher et al. 1997
; Manabe et al. 1993
;
Zucker 1989
). Indeed, we found that bath
application of acetylcholine or norepinephrine enhanced facilitation
(Fig. 3). This was observed in the field
(n = 5-7 per drug) and intracellular EPSPs
(n = 3-5 per drug), was not followed by a significant
change in input resistance (Fig. 3A), and was reversible
(Fig. 3B). The enhancement in facilitation induced by
acetylcholine was mimicked by carbachol, but not by nicotine, and was
blocked by the muscarinic receptor antagonist, scopolamine (Fig.
3C). Also, the enhancement in facilitation induced by
norepinephrine was mimicked by clonidine, but not by phenylephrine or
isoproterenol, and was blocked by the
-adrenergic antagonist,
phentolamine.

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Fig. 3.
Acetylcholine (left) and norepinephrine
(right) enhance corticothalamic facilitation.
A: the top traces represent the typical
facilitation displayed by intracellular and field EPSPs in response to
a pair of stimuli at a 50-ms interstimulus interval (ISI). Overlaid are
responses before and during the application of acetylcholine (10 mM)
and norepinephrine (100 µM). The insets below show the
result of scaling the amplitude of the 1st (depressed) response during
the drug to the amplitude of the response before the drug. This reveals
an enhancement of facilitation. Also shown are the neuron's responses
to a DC current pulse ( 0.3 nA) delivered before and during drug
application. B: the experiments corresponding to the
extracellular traces shown in A are illustrated. The
top graph shows the amplitude of the response to the 1st
stimulus ( ) and to the 2nd stimulus ( )
delivered with a 50-ms ISI. The graph below displays ( )
the percentage of change in amplitude of the 2nd response with respect
to the 1st response (PPF). Notice that during application of
acetylcholine (ACh, 10 mM) and norepinephrine (NE, 100 µM), the
evoked responses depressed but facilitation increased.
C: the graphs represent the average effects of the drugs
on the baseline PPF (5-7 experiments per group;
* P < 0.01, t-test). The
left panel displays the effects of acetylcholine (10 mM), carbachol (5 µM), nicotine (10 µM), and acetylcholine in the
presence of scopolamine (ACh + Scop; 10 mM + 10 µM). The right
panel displays the effects of norepinephrine (NE, 100 µM),
clonidine (40 µM), phenylephrine (50 µM), isoproterenol (50 µM),
and norepinephrine in the presence of phentolamine (NE + Phent; 100 µM + 100 µM). Intracellular recordings were performed with QX-314
and Cs+ in the pipette. BMI was bath applied.
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Previous work in vivo has shown that thalamic responses can change
during distinct behavioral states or after stimulation of the reticular
formation (Pare et al. 1990
; Steriade et al. 1969
, 1986
; Timofeev et al.
1996
). Those studies found that ascending cerebellothalamic and
mamillothalamic pathways in vivo are enhanced during arousal. The
present results in vitro suggested that during arousal, when
acetylcholine and norepinephrine are released into the thalamus, the
descending corticothalamic pathway depresses. The next experiments
explored this possibility. To induce arousal we used stimulation of the
brain stem reticular formation in vivo. As previously described
(Moruzzi and Magoun 1949
; Steriade and McCarley
1990
), this stimulation produces a strong activating effect in
the thalamus consisting of the abolition of slow-wave activity and an
increased depolarization that results in high-frequency unit firing
(Fig. 4A). These effects last
for at least 10 s, which was defined as our testing period
(between 1 and 10 s after the reticular formation stimulation).
Stimulation of the thalamic radiation in anesthetized rats produced a
corticothalamic response in the ventrobasal thalamus very similar to
the one recorded in vitro (compare Figs. 3A and
4B), with its characteristic strong facilitation (Fig.
4C). As previously observed in vitro
(Castro-Alamancos and Calcagnotto 1999
), the
corticothalamic response was abolished in vivo by application of a
glutamate receptor antagonist into the thalamus (Fig. 4B).
This shows that antidromic activation does not contribute to the
synaptic field response we measure because this response is completely
abolished by CNQX. Also, antidromic excitation in the ventrobasal
thalamus cannot produce recurrent EPSPs because thalamocortical neurons
are not synaptically connected to each other. We first tested the
effects of arousal on the corticothalamic pathway by comparing
responses before and after reticular formation stimulation
(n = 7). Corticothalamic responses were strongly
depressed during the testing period (Fig. 4D). This
depression usually lasted between 20 and 60 s. Thus as predicted
from the effects of acetylcholine and norepinephrine in vitro,
stimulation of the reticular formation resulted in a strong depression
of the corticothalamic pathway.

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Fig. 4.
Stimulation of the reticular formation in vivo suppresses low-frequency
but not high-frequency corticothalamic responses. A:
typical example of the effects of stimulating the brain stem reticular
formation (RF, 100 Hz/1 s) on thalamic activity. The 3 traces
correspond to the same recording with different filter settings. The
low-pass filter displays slow oscillations, while the high-pass filter
displays multi-unit activity. RF stimulation abolished slow
oscillations and increased high-frequency neuronal discharges.
B: stimulation of the thalamic radiation in vivo induces
a corticothalamic response that is similar to those recorded in vitro.
It facilitates when stimulated at a 50-ms interstimulus interval (ISI).
The response is abolished by application of a
non-N-methyl-D-aspartate (non-NMDA)
glutamate receptor antagonist (CNQX,
6-cyano-7-nitroquinoxaline-2,3-dione) into the thalamus.
C: the amplitude of the corticothalamic responses
displayed in response to 4 stimuli delivered at 0.1, 0.5, 1, 2, 5, 10, 20, and 40 Hz are displayed as a percentage of change with respect to
the 1st response. D: the traces represent typical
examples of the effects of RF stimulation on corticothalamic responses
evoked at 0.5 Hz (above; only the 1st and 4th responses are shown) and
at 20 Hz (below). The 20-Hz train triggers facilitation, but not the
0.5-Hz train. Overlaid are the responses before and after RF
stimulation. Notice the depression of the responses at 0.5 Hz, but only
of the 1st response in the 20-Hz train. E: as shown in
C, facilitation reaches a steady state by the 3rd
response. We used the 3rd and 4th responses of the trains to perform a
spectrum analysis on the effects of RF stimulation. The average
amplitude of the 3rd and 4th responses for each frequency is displayed
as the percentage of change with respect to the control response
(before RF) at 0.1 Hz. Notice that the steady-state low-frequency
responses are abolished while the high-frequency responses are
unaffected by RF stimulation. The data correspond to the average of 4 experiments.
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Does this mean that during arousal the cortex cannot communicate with
the thalamus? One possibility is that corticothalamic responses only
depress at low frequencies, but not at high frequencies that engage
facilitation. This was suggested by the observation, in vitro, that
acetylcholine and norepinephrine enhance facilitation. Thus we compared
the effects of reticular formation stimulation on low-frequency and on
high-frequency corticothalamic activity (Fig. 4D). The
results showed that while the corticothalamic responses to
low-frequency activity were strongly depressed by stimulating the
reticular formation, the responses to high-frequency activity were not
affected. A spectrum analysis revealed that corticothalamic responses
below 5 Hz were strongly suppressed during arousal, but higher
frequency responses were unaffected. (Fig. 4E). The steady-state responses at a frequency below 5 Hz were significantly depressed by stimulation of the reticular formation (n = 4; t-test, P < 0.01). These results
indicate that during arousal corticothalamic activity is high-pass
filtered at 5 Hz. Moreover, consistent with the in vitro results,
application of both muscarinic and
-adrenergic antagonists into the
thalamus abolished the depression of corticothalamic responses induced
by stimulating the reticular formation (n = 3; Fig.
4F). This experiment also allowed dissociating the synaptic effects from the activating effects of reticular formation stimulation. Figure 5 shows the
effect of stimulating the reticular formation before (ACSF) and during
the application of both muscarinic and
-adrenergic antagonists into
the thalamus (scopolamine and phentolamine). The reticular formation
stimulation was effective under both conditions in producing thalamic
activation (Fig. 5A). This was expected because it is known
that many different neuromodulators produce activating effects in the
thalamus and that acetylcholine acting via nicotinic receptors and
norepinephrine via
-adrenergic receptors (which were not blocked)
also depolarize thalamic neurons (McCormick 1992
;
Steriade et al. 1997
). Although application of
scopolamine and phentolamine did not eliminate the thalamic activation
produced by stimulating the reticular formation, it did abolish the
depression of corticothalamic responses (Fig. 5B). This
indicates that the depressing effect on corticothalamic synapses
induced by stimulating the reticular formation is independent from the
changes in cellular excitability of thalamic activation.

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Fig. 5.
Blocking muscarinic and -adrenergic receptors in the thalamus
dissociates between the synaptic and cellular excitability effects of
stimulating the reticular formation. A: typical examples
showing similar effects of stimulating the brain stem reticular
formation (RF, 100 Hz/1 s) on thalamic activity during application of
artificial cerebrospinal fluid (ACSF; control) or during application of
scopolamine and phenolamine (500 µM; Sco + Phe) into the thalamus.
Traces are low-pass filtered. B: differential effects of
stimulating the brain stem reticular formation on corticothalamic
responses during application of ACSF (control; left) or
during application of scopolamine and phentolamine into the thalamus
(Sco + Phe; right). The traces represent typical
examples of the effects of RF stimulation on corticothalamic responses
evoked at 0.5 Hz (above; only the 1st and 4th responses are shown) and
at 20 Hz (below). The 20-Hz train triggers facilitation, but not the
0.5-Hz train. Overlaid are the responses before and after RF
stimulation in the presence (right) and absence
(left) of scopolamine and phentolamine in the thalamus.
Notice that in the presence of Sco + Phe, stimulation of the reticular
formation does not depress corticothalamic responses at 0.5 Hz or the
1st response of the train at 20 Hz. All data in this figure are from
the same experiment. C: group data
(n = 3) shown are the mean amplitudes of the
steady-state responses for a low- (0.5) and a high-frequency (20 Hz)
stimulus displayed as the percentage of the control (before RF)
response at 0.1 Hz. RF stimulation depresses selectively the
low-frequency response, and infusing scopolamine and phentolamine (500 µM) in the thalamus blocks this effect.
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We also tested the effects of reticular formation activation on
thalamic responses evoked by stimulating the medial lemniscus (Castro-Alamancos, unpublished observations) and found that these responses were either enhanced or not affected (n = 4;
not shown) as previously described (Steriade and Demetrescu
1960
; for review Singer 1977
; Steriade
and McCarley 1990
; Steriade et al. 1997
). This
suggests that the depressing effects of stimulating the reticular formation are restricted to corticothalamic synapses. Therefore the
effects in vivo on corticothalamic responses induced by stimulating the
reticular formation are likely synaptic and not a general consequence
of a change in excitability of thalamic neurons.
Finally, we tested whether activation of cholinergic or
2-adrenergic receptors in slices using
receptor agonists was sufficient to high-pass filter corticothalamic
responses in vitro. To mimic our conditions in vivo, we recorded
corticothalamic field responses in the absence of GABA receptor
antagonists. Under these conditions corticothalamic field responses
recorded in slices are similar to those recorded in vivo (compare Figs.
4D and 6A), with the characteristic strong
facilitation (Fig. 6B). The
neuromodulator agonists were applied locally (at the recording site)
using a low-impedance pipette containing the drug (at a dose 10 times higher than in the bath) dissolved in ACSF as previously described (Castro-Alamancos and Calcagnotto 1999
). Application of
either clonidine (400 µM, n = 3) or carbachol (50 µM, n = 3) resulted in a strong depression of the
corticothalamic responses (Fig. 6A). Low-frequency responses
were always affected more than high-frequency responses (Fig.
6A). This effect resembles the result obtained by
stimulating the reticular formation in vivo. As in vivo, we performed a
spectrum analysis, which revealed that low-frequency corticothalamic
responses were strongly suppressed but high-frequency responses were
unaffected by local application of clonidine or carbachol (Fig.
6C). The steady-state responses at a frequency below 10 Hz
were significantly depressed by carbachol or clonidine (n = 3; t-test, P < 0.01).
It is unlikely that we would be able to completely simulate in vitro
the exact pattern and dosage of receptor activation that corresponds to
stimulating the brain stem reticular formation in vivo. Several
differences were noted between the in vivo and in vitro experiments.
First, the GABAA receptor-dependent positivity
that follows the negative field potential EPSP, which is only observed
in the absence of BMI, showed strong frequency-dependent depression in
vitro but not in vivo. Second, the frequency-dependent facilitation
displayed in vivo and in vitro is very similar but not identical.
Facilitation in vivo tends to be stronger and begins at higher
frequencies than in vitro. This is likely due to differences in the
experimental conditions (e.g., ACSF). Surely, these factors contribute
to the differences between the frequency curves of the steady-state
responses in vivo and in vitro. Taken together, the results in vitro
and the antagonist experiments in vivo indicate that activation of muscarinic or
2-adrenergic receptors at
corticothalamic synapses during arousal result in the selective
suppression of low-frequency corticothalamic activity.

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Fig. 6.
Application of muscarinic or 2-adrenergic receptor
agonists in vitro suppresses low-frequency but not high-frequency
corticothalamic responses. A: the traces represent
typical examples of the effects of carbachol on corticothalamic
responses evoked at 0.5 Hz (above; only the 1st and 4th responses are
shown) and at 40 Hz (below). The 40-Hz train triggers facilitation, but
not the 0.5-Hz train. Overlaid are the responses before and during
carbachol. Notice the depression of the responses at 0.5 Hz, but only
of the 1st and 2nd responses in the 40-Hz train. B: the
amplitude of the corticothalamic responses displayed in response to 4 stimuli delivered at 0.1, 0.5, 1, 2, 5, 10, 20, and 40 Hz are displayed
as a percentage of change with respect to the 1st response.
C: as shown in B, facilitation reaches a
steady state by the 3rd response. We used the 3rd and 4th responses of
the trains to perform a spectrum analysis on the effects of carbachol
and clonidine. The average amplitude of the 3rd and 4th responses for
each frequency is displayed as the percentage of change with respect to
the control response (before the drug) at 0.1 Hz. Notice that the
steady-state low-frequency responses are abolished while the
high-frequency responses are unaffected by clonidine and carbachol. The
data correspond to the average of 3 experiments.
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DISCUSSION |
The present experiments, performed in vitro and in vivo, reveal
that acetylcholine and norepinephrine depress corticothalamic synapses
via muscarinic and
2-adrenergic receptors.
However, since corticothalamic depression is followed by an enhancement of facilitation, only low-frequency corticothalamic activity is suppressed during arousal, while high-frequency activity, which triggers facilitation, flows unaffected.
The results indicate that the release of acetylcholine or
norepinephrine in the thalamus depresses corticothalamic synapses by
activating muscarinic and
2-adrenergic
receptors, respectively. First, acetylcholine and norepinephrine
depress corticothalamic EPSPs measured intra- and extracellularly. This
depression occurs both in the presence or absence of GABA-mediated
inhibition. Second, cholinergic and
2-adrenergic receptor agonists, but not
nicotinic or
-adrenergic agonists, depress corticothalamic EPSPs.
Third, a muscarinic antagonist and a
-adrenergic antagonist block
the depressing effects of acetylcholine and norepinephrine on
corticothalamic EPSPs, respectively. Fourth, stimulation of the
reticular formation, which releases several neuromodulators into the
thalamus, depresses corticothalamic field EPSPs, and this depression is
abolished by the application of muscarinic and
-adrenergic
antagonists into the thalamus.
Previous work has demonstrated that corticothalamic synapses are
depressed by glutamate via type III metabotropic glutamate receptors
(Turner and Salt 1999
). The present study ads both
muscarinic and
2-adrenergic receptors to the
list of neuromodulators that regulate corticothalamic communication. By
which mechanisms do muscarinic and
2-adrenergic receptors depress corticothalamic EPSPs? A differential presynaptic modulation of low-frequency versus
high-frequency synaptic activity by muscarine and by GABA has been
previously shown in the ventral striatum (Pennartz and Lopes da
Silva 1994
) and in the auditory system (Brenowitz et al.
1998
), respectively. The present results also imply that
corticothalamic depression induced by acetylcholine and norepinephrine
is due to a presynaptic mechanism involving a decrease in the
probability of neurotransmitter release. This is suggested by the
finding that both neuromodulators produced an enhancement in
paired-pulse facilitation, which is consistent with a decrease in
neurotransmitter release probability (Manabe et al.
1993
; Zucker 1989
). One mechanism that cannot
account for the effects of the neuromodulators on facilitation is a
change in the frequency-dependent depression of disynaptic inhibitory
postsynaptic potentials (von Krosigk et al. 1999
)
because, in our experiments, inhibition was blocked. Moreover, a
postsynaptic effect of these neuromodulators consisting in a change in
cellular excitability is unlikely to explain the synaptic depression
because we significantly suppressed the postsynaptic actions of these
neuromodulators during intracellular recordings by blocking
Na+ and K+ conductances,
and this did not affect the synaptic depression. However, a
postsynaptic action of the neuromodulators at corticothalamic synapses
cannot be ruled out. Of particular interest was the dissociation in
vivo between the effects of muscarinic and
-adrenergic antagonists on corticothalamic responses and on thalamic activation. These antagonists specifically abolished the depression of low-frequency corticothalamic responses induced by stimulating the reticular formation, but not the thalamic activation. This suggests that the
well-known postsynaptic depolarizing effects produced by the release of
neuromodulators on thalamic neurons are not contributing to the
corticothalamic depression. The dissociation between thalamic activation and the depression of corticothalamic responses was also
supported by the observation that the corticothalamic depression always
outlasted the thalamic activation. Finally, the observation that only
corticothalamic, but not medial lemniscus, responses are depressed by
stimulating the reticular formation demonstrates that the effect is
input specific. This suggests that the depressing effect of the
neuromodulators is occurring at corticothalamic synapses and is not a
general consequence of a change in excitability. Thus the present
results indicate that the depression occurs at corticothalamic
synapses. Further work should clarify whether the locus of depression
is presynaptic or postsynaptic.
What is the functional significance of a decrease in corticothalamic
efficacy that is accompanied by an increase in facilitation? The
thalamocortical system undergoes dramatic functional changes between
sleep and arousal (Steriade et al. 1997
). For example, during slow-wave sleep the neocortex and thalamus are engaged in
low-frequency oscillations, while during arousal they engage in
high-frequency gamma oscillations (Steriade et al.
1993
). Gamma oscillations at 20-40 Hz coincide with the
frequencies that are effective in triggering short-term synaptic
facilitation in corticothalamic synapses. Interestingly, stimulating
the brain stem reticular formation enhances both gamma oscillations
(Steriade and Amzica 1996
; Steriade et al.
1991
) and synaptic facilitation. Thus an important role for
this corticothalamic high-pass filter during arousal is to permit the
flow of gamma oscillations between neocortex and thalamus, while
impeding low-frequency oscillations. It would serve as a gate that
allows the flow of low-frequency and gamma activity during sleep, while
allowing only gamma activity during arousal.
We thank Dr. Steriade for helpful comments.
This research was supported by the Medical Research Council of Canada,
the Natural Sciences and Engineering Council of Canada, Fonds de la
Reserche en Sante du Quebec, the Canadian Foundation for Innovation,
and the Savoy Foundation.
Address for reprint requests: M. Castro-Alamancos, Montreal
Neurological Institute, 3801 University St., Rm. WB210, Montreal, Quebec H3A 2B4, Canada (E-mail:
mcastro{at}bic.mni.mcgill.ca).