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Received 20 April 1998; accepted after revision 4 September 1998.
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ABSTRACT |
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INTRODUCTION |
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Although nitric oxide (NO) has been established as a neural messenger (Bredt & Snyder, 1994), its role in synaptic plasticity is controversial. The induction of long-term potentiation (LTP) in area CA1 of the hippocampus is facilitated by NO (Böhme et al. 1991; Schuman & Madison, 1991; Son et al. 1996). However, the contribution of NO to the induction of hippocampal LTP is dependent on various experimental conditions such as temperature and animal age (Williams et al. 1993), or stimulus patterns (Lum-Ragan & Gribkoff, 1993). In addition, there are discrepant reports regarding the stimulus intensity for induction of LTP (Lum-Ragan & Gribkoff, 1993; Haley et al. 1993). Cerebellar long-term depression (LTD) in parallel fibre (PF)-Purkinje cell synapses is also dependent on NO signalling. Although NO does not affect LTD of glutamate-induced currents recorded in cultured Purkinje cells (Linden et al. 1995), requirement of NO-cGMP signalling for induction of cerebellar LTD has been clearly demonstrated in slice preparations (Ito & Karachot, 1990; Crepel & Jaillard, 1990; Shibuki & Okada, 1991; Lev-Ram et al. 1995; Hartell, 1996). NO release from PFs has been demonstrated with electrochemical NO probes (Shibuki & Okada, 1991; Shibuki & Kimura, 1997). In accordance with these data obtained from cerebellar slices, certain types of cerebellar motor learning, for which cerebellar LTD is regarded as the cellular mechanism, are also dependent on NO signalling (Nagao & Ito, 1991; Yanagihara & Kondo, 1996).
In the neocortex, development of the primary sensory cortex relies on activity-dependent synaptic plasticity (Hubel & Wiesel, 1963; Blakemore & Cooper, 1970). The ocular dominance shift of neurones in the visual cortex following monocular deprivation is a well-known example of developmental plasticity (Wiesel & Hubel, 1963). Local injection of a nitric oxide synthase (NOS) inhibitor into the visual cortex, however, does not affect the ocular dominance shift (Reid et al. 1996; Ruthazer et al. 1996). The induction of LTP in layer II-III (LTPII-III) in the visual cortex does not depend on NO signalling (Kirkwood & Bear, 1994). However, LTP in layer V (LTPV) of the medial frontal cortex is NO dependent (Nowicky & Bindman, 1993). The apparent difference in the NO dependence of LTPII-III in the visual cortex and LTPV in the medial frontal cortex might be attributed to differences in cortical layers, cortical areas or other experimental conditions. To understand the role of NO in neocortical LTP, it is necessary to study NO dependence of LTP in different layers of the same cortical area under the same experimental conditions. Marked LTP of population spikes is observed in the rat auditory cortex (Kudoh & Shibuki, 1996). The net LTP in the auditory cortex is twice as large as that in the visual cortex (Kudoh & Shibuki, 1997). Therefore, we studied the layer specificity of the NO dependence of neocortical LTP using slices obtained from the rat auditory cortex, and found that LTPV was NO-cGMP dependent while LTPII-III was not.
The layer-specific NO dependence of LTPV in the auditory cortex suggests biased NO release in layer V. Neuronal NOS (nNOS) is the main isoform of NOS in the neocortex (Huang et al. 1993). Although the density of nNOS in the neocortex is only one third of that in the cerebellum (Huang et al. 1993), strongly NOS-positive non-pyramidal neurones are found in the neocortex (Bredt et al. 1991; Valtschanof et al. 1993). The layer-specific NO dependence of LTP suggests a biased distribution of nNOS-positive neurones in rat auditory cortex. NO release from PFs in cerebellar slices has been characterized using electrochemical NO probes (Shibuki & Kimura, 1997). However, the properties of NO release in the neocortex are unknown. We studied the distribution of nNOS-positive neurones in the auditory cortex, and characterized the NO release in the auditory cortex using electrochemical NO probes.
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METHODS |
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Slice preparations
The experiments in this study were performed according to the guidelines of Niigata University, and had the approval of the ethics committee of Niigata University. Wistar rats of both sexes (4-7 weeks old) were used. After the rats had been deeply anaesthetized with ether, they were immersed in ice-cold water, except for the nose, for 3 min to reduce the brain temperature. Immediately after decapitation, a block of brain tissue including the auditory cortex was dissected out. The location of the auditory cortex was determined to be that of area 41 of the temporal cortex (Krieg, 1964). Frontal slices (400 µm thick) of the auditory cortex were prepared from the block in an ice-cold medium using a microslicer (DTK-2000, Dosaka, Osaka, Japan). The composition of the medium, which was the same as that used in our previous studies (Kudoh & Shibuki, 1996, 1997; Shibuki & Kimura, 1997), was (mM): NaCl, 124; KCl, 5; NaH2PO4, 1·24; MgSO4, 1·3; CaCl2, 2·4; NaHCO3, 26; and glucose, 10. This medium was continuously bubbled with 95 % O2 and 5 % CO2 for at least 1 h before use. After incubation at 30°C for more than 1 h, the slices were transferred to a small recording chamber (about 0·3 ml in volume), in which they were kept submerged. The recording chamber was maintained at 30°C and was continuously perfused with the oxygenated medium at a flow rate of 1 ml min-1.
Drugs
NG-Nitro-L-arginine (NA), NG-methyl-L-arginine (MA) and 8-bromo-cGMP (Br-cGMP) were purchased from Sigma. 6-Cyano-7-nitroquinoxaline-2,3-dione (CNQX) and D-2-amino-5-phosphonovalerate (APV) were from Tocris Cookson (Bristol, UK). (±)-(E)-Ethyl-2-[(E)-hydroxyimino]-5-nitro-3-hexeneamide (NOR3) was from Dojindo Laboratories (Kumamoto, Japan), and bicuculline and phaclofen were from RBI (Natick, MA, USA). These drugs were applied to the slices by addition to the perfusion medium. Dimethyl sulphoxide (DMSO) was used as a solvent for the application of 10 µM CNQX and 5 µM NOR3 to the slices. The solution of 20 mM CNQX or 100 mM NOR3 dissolved in DMSO was mixed with the perfusion medium. DMSO alone (0·05 or 0·005 %) had no significant effect on the field potentials, LTP or NO release (data not shown). Other drugs were directly dissolved in the perfusion medium. In LTP experiments, Br-cGMP and NOR3 were applied to the slices for a few minutes before tetanic stimulation of layer IV and during a period of the stimulation. Other drugs were applied to the slices throughout LTP recording.
Electrical stimulation
Field potentials and NO release were elicited by layer IV stimulation, because layer IV stimulation is known to elicit LTPII-III efficiently (Kirkwood & Bear, 1994). Layer IV in the slices was stimulated with biphasic current pulses through the cut end of a Teflon-coated Ag wire (diameter, 75 µm) placed on the surface of layer IV at which a faint band was observed under a binocular microscope. This band probably reflects the characteristic cyto-architecture in layer IV, or the denser thalamo-cortical axons in layer IV than in layers III and V. When negative pulses not followed by positive pulses are passed through the stimulation electrode, H2 gas is generated at the metal surface of the Ag wire and is detected by a NO probe (Shibuki & Kimura, 1997). To avoid this stimulus artefact in NO measurement, the Ag wire was coated with AgCl by passing positive currents through the stimulation electrode in a NaCl solution. In addition, each negative pulse was followed by a positive pulse, the absolute amplitude of which was 5 % larger than that of the preceding negative pulse. The duration of each pulse phase was 100 µs. Under these conditions, the negative stimulus current was carried by Cl- dissociating from AgCl, avoiding electrolytic generation of H2 gas.
LTP recording and induction
Field potentials in slices were recorded through a metal electrode. This electrode was made from a piece of Ag wire (200 µm in diameter), which was electrolytically polished in 10 % KNO3 solution and was insulated with polyvinyl chloride except for the part within 60 µm from the tip. Signals were amplified 10 times with a handmade electronic circuit using an operational amplifier (OPA128LM, Burr-Brown, Tucson, AZ, USA), and were passed through a bandpass filter between 0·2 Hz and 10 kHz. The output was stored in a computer (PC-9801BA2, NEC, Tokyo, Japan) via an analog-digital converter board (ADXM-98A, Canopus, Kobe, Japan) for later analysis. We developed the BASIC programs used for the recording and analysis using the software library supplied by Canopus.
Field potentials in layer II-III or layer V were elicited by layer IV stimulation with biphasic pulses (intensity < 800 µA; duration of each pulse phase, 100 µs) in a slice. For LTP recording, the magnitude of the trans-synaptic responses in layer II-III or layer V was measured from the peak amplitude of the second negativity in the field potentials elicited by layer IV stimulation (Figs 1 and 2). Although very similar results were obtained from the measurement of the initial slopes of the second negativity (data not shown), we used the measurement of the peak amplitude, since it was less affected by the preceding antidromic activities of pyramidal neurones than was the initial slope. LTP recording was performed in the slice only when the amplitude of the trans-synaptic field potential was larger than 0·8 mV. The current intensity of test stimuli was adjusted between 200 and 500 µA to ensure that half-maximal responses were elicited, and there was no systematic difference in the intensity of test stimuli between the experiments for LTPII-III and LTPV. Test stimuli were applied to the slice at 20 or 30 s intervals, and two or three traces were averaged each minute to determine baseline responses. After stable baseline responses were recorded for at least 10 min, tetanic stimulation of layer IV was carried out. To evoke LTPII-III, 100 pulses at 100 Hz were applied to layer IV twice, at an interval of 30 s. Tetanic stimulation at the test stimulus intensity was usually insufficient to evoke marked LTPII-III (Kudoh & Shibuki, 1996). Therefore, we placed another stimulation electrode near the first electrode (distance between the tips < 100 µm) and simultaneously applied tetanic stimulus pulses that were 1·5 times the intensity (300-750 µA) and 2 times the duration (duration of each pulse phase, 200 µs) of test pulses. Tetanic stimulation at 100 Hz was not sufficient to evoke LTPV (Nowicky & Bindman, 1993). Therefore, 100 pulses at 200 Hz were applied to layer IV through the first and second stimulation electrodes 3 times at 20 s intervals to evoke LTPV. The amplitude of the trans-synaptic field potential was normalized to the averaged value of three consecutive traces recorded immediately before tetanic stimulation. The magnitude of LTP was evaluated as the averaged amplitude in the trace recorded at 30 min after tetanic stimulation and the preceding two traces, unless otherwise specified.
NO recording
NO release in the neocortical slices after layer IV stimulation was measured using electrochemical NO probes. The NO probes were fabricated as described previously (Shibuki & Kimura, 1997). Briefly, the tip of a glass pipette was polished obliquely and flame smoothed. The pipette was filled with 30 mM NaCl and 0·3 mM HCl. The opening of the pipette was sealed with a thin membrane of silicon rubber (TSE399, Toshiba, Tokyo, Japan). For the working electrode, a Teflon-coated Pt wire (metal diameter, 125 µm) was cut obliquely and the cut end was heated in a flame for a few seconds to remove the Teflon coating. The exposed Pt wire was insulated with heat-melted dental wax, except at the tip. The Pt wire was inserted into the pipette to a position where the tip protruded from the pipette end (Fig. 5). A reference electrode (Teflon-coated Ag wire) was also inserted into the pipette, and was connected to the ground. The working Pt electrode was connected to a handmade current-voltage converter using an operational amplifier (OPA128LM). The voltage of the Pt wire was kept at +0·9 V, unless otherwise specified. Each NO probe was calibrated by measuring the probe currents in response to a 30 µM NO solution, which was prepared by dissolving 36 µl NO gas in 50 ml of degassed saline in a glass syringe. We used one point calibration since the current changes in our NO probes were proportional to NO concentration (Shibuki & Kimura, 1997). The NO sensitivity of the probes was 0·5-1·4 nA (µM NO)-1. The NO probes were positioned on the surface of layer V or layer II-III of the slices (Fig. 5). NO release was elicited by repetitive stimulation of layer IV (20 Hz for 5 s) at a current intensity of 500 µA, unless otherwise specified. The typical distance between the tip of the NO probe and the stimulation electrode in layer IV was 250-300 µm. Stimulus pulse trains were applied to the slices at 2 min intervals, and five consecutive traces of changes in NO concentration were averaged to improve the signal-to-noise ratio of NO recording.
Immunohistochemistry of nNOS
After slices of the auditory cortex (400 µm thick) had been incubated at 30°C for several hours, they were fixed by immersion in ice-cold 4 % paraformaldehyde in 0·1 M sodium phosphate buffer (PBS; pH 7·4) for 12 h, followed by equilibration in 30 % sucrose in 0·1 M PBS. Subsequently, the slices were frozen, and sections (30 µm thick) were cut using a cryotome (AS620, Shandon, Runcorn, UK). The sections were immersed in PBS containing 0·1 % Triton X-100 and 6 % normal horse serum (blocking solution; Vector Laboratories, Burlingame, CA, USA) for 3 h at room temperature (21-23°C), and then were reacted with a 1 : 3000 dilution of an anti-nNOS monoclonal antibody (N-2280, Sigma) in blocking solution for 24 h at 4°C. This monoclonal antibody was produced using recombinant nNOS fragment (amino acids 1-181) as immunogen, and reacts specifically with human, goat, porcine and rat nNOS but not with endothelial or inducible NOS. After repeated washes with PBS containing 0·1 % Triton X-100, the sections were incubated with a biotinylated anti-mouse secondary antibody (Vector Laboratories) overnight at 4°C. Immunohistochemical detection was carried out with an avidin-biotin-peroxidase technique using the Vectastain ABC kit and DAB substrate kit obtained from Vector Laboratories.
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RESULTS |
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NO-independent LTP in layer II-III
The field potentials in layer II-III in the auditory cortex are composed of early and late negative waves. The former represent antidromic firing of pyramidal neurones and axonal activities while the latter correspond to trans-synaptic responses of pyramidal neurones (Kudoh & Shibuki, 1996, 1997). The trans-synaptic responses in the auditory cortex mainly reflect population spikes (Kudoh & Shibuki, 1996). Tetanic stimulation of layer IV produced LTPII-III only in the trans-synaptic responses (Fig. 1Ab and B). The magnitude of LTPII-III at 30 min after tetanic stimulation was 171 ± 9 % (mean ±
Aa, diagram showing the relative positioning of recording and stimulation electrodes in cortical layers for LTPII-III recording. WM, white matter. b, field potentials elicited by layer IV stimulation and recorded in layer II-III before and 30 min after (
NO-dependent LTP in layer V
Layer IV stimulation elicited field potentials of two negative waves in layer V, as observed in layer II-III. Only the late negative waves were blocked by 10 µM CNQX, a non-NMDA receptor antagonist (Fig. 2Ab). LTPV was selectively produced by tetanic stimulation of layer IV in the late negative waves (Fig. 2Ac and B). The magnitude of LTPV at 30 min after tetanic stimulation was 138 ± 3 % (n = 17) of the control recorded immediately before stimulation and that at 60 min after stimulation was 137 ± 4 % (n = 5). Since no significant difference was found between these values, LTPV was evaluated at 30 min after tetanic stimulation in the following experiments. Bicuculline (1 µM), a GABAA receptor antagonist, showed no significant effect on LTPV (131 ± 3 %, n = 7), as reported for LTPII-III in the auditory cortex (Kudoh & Shibuki, 1997). In contrast to LTPII-III, however, the induction of LTPV was significantly suppressed by 10 µM NA (P < 0·001, Mann-Whitney U test; Fig. 2Ad and B) or by 100 µM MA (P < 0·001). The magnitude of LTPV was 106 ± 3 % (n = 10) in the presence of 10 µM NA and 109 ± 3 % (n = 12) in the presence of 100 µM MA.
Aa, diagram showing the relative positioning of recording and stimulation electrodes in cortical layers for LTPV recording. b, field potentials elicited by layer IV stimulation and recorded in layer V before and during application of 10 µM CNQX. c, field potentials recorded in layer V before and 30 min after (
To test the specificity of the effect of NA on LTPV, NOR3, a NO donor, was added to the perfusion medium for a few minutes before tetanic stimulation and during a period of the stimulation. LTPV elicited in the presence of 5 µM NOR3 alone (140 ± 8 %, n = 5) was not significantly different from LTPV elicited in normal medium (Fig. 3Ab). However, LTPV elicited in the presence of 10 µM NA plus 5 µM NOR3 (137 ± 5 %, n = 7; Fig. 3Aa and Ab) was significantly larger than LTPV elicited in the presence of 10 µM NA alone (P < 0·001; Fig. 3Ab). These results strongly suggest that the blockade of the induction of LTPV by NA can be attributed to the inhibition of NO synthesis by NA.
Stimulation of cGMP synthesis is one of the main biological functions of NO (Bredt & Snyder, 1994). Therefore, Br-cGMP, a membrane-permeant analogue of cGMP, was applied to the slices instead of NOR3. The magnitude of LTPV elicited in the presence of 1 mM Br-cGMP was 139 ± 7 % (n = 5), and Br-cGMP alone had no significant effect on LTPV (Fig. 3B b). However, LTPV elicited in the presence of 10 µM NA plus 1 mM Br-cGMP (128 ± 5 %, n = 6; Fig. 3B a and Bb) was significantly larger than LTPV elicited in the presence of 10 µM NA alone (P < 0·001; Fig. 3B b). These results clearly indicate that the induction of LTPV, but not that of LTPII-III, is dependent on NO-cGMP signalling in the rat auditory cortex.
Aa, field potentials in layer V recorded before and 30 min after (
Cellular localization of nNOS in the auditory cortex
It is reported that nNOS is concentrated in only 1-2 % of neurones in the neocortex (Bredt et al. 1991). The layer-specific NO dependence of LTP suggests that more nNOS-positive neurones may be present in the deeper layers of the slices of the auditory cortex. Therefore, we stained the slices obtained from the auditory cortex with a monoclonal anti-nNOS antibody. As reported previously (Bredt et al. 1991), there were only a small number of nNOS-positive neurones (Fig. 4A). Their cell bodies and processes were mainly located in deep cortical layers. High magnification of the sections showed non-pyramidal neurones stained densely with the anti-nNOS antibody (Fig. 4B and C).
A, frontal section of the slices of the auditory cortex. Scale bar, 500 µm; WM, white matter. Cells indicated by arrowheads are also shown in B and C. The scale bar in B (100 µm) also applies to C.
NO release in layer V and in layer II-III
The layer-specific NO dependence of LTP and the biased distribution of nNOS-positive neurones in the auditory cortex suggest that NO release in the auditory cortex may also have a biased distribution. Therefore, we measured NO release elicited by layer IV stimulation in layer V and in layer II-III with electrochemical NO probes. Layer IV stimulation with 100 pulses at 20 Hz produced currents in the NO probes placed on the surface of layer V (Fig. 5A). The currents were augmented by application of 100 µM L-arginine (Arg), the substrate of NOS, by 46 ± 11 % (n = 5), and were almost completely blocked by application of 10 µM NA (Fig. 5A). The amplitude of the currents corresponded to an increase in NO concentration of 380 ± 14 pM (n = 55). Layer IV stimulation with 100 pulses at 200 Hz also produced currents which corresponded to 427 ± 37 pM NO (n = 3) and were completely blocked by 10 µM NA (Fig. 5B). However, the currents repeatedly elicited by 200 Hz stimulation were sometimes gradually reduced. In contrast, stimulation at 20 Hz produced more reproducible results than the stimulation at 200 Hz, and therefore stimulation at 20 Hz was used in the following experiments unless otherwise specified. The anode voltage between the working Pt wire and the reference of the NO probe drastically modulated the amplitude of the currents recorded in layer V (Fig. 5C). The dependence of the currents recorded in layer V on the anode voltage was the same as that of the currents induced by 30 µM NO (Fig. 5D). Similar currents blocked by 10 µM NA were observed in layer II-III, but the amplitude of the estimated NO increase (55 ± 8 pM, n = 5) was only one-seventh of that in layer V; this difference was significant (P < 0·001; Fig. 5E). These data indicate that layer IV stimulation elicited more NO release in layer V than in layer II-III.
A, NO release in layer V elicited by repetitive layer IV stimulation (20 Hz for 5 s). Three traces recorded before (Control) and during application of 100 µM Arg or 10 µM NA are shown superimposed. The inset shows the relative positioning of the NO probe and stimulation electrode in the cortical layers. G, reference electrode connected to ground. B, NO release in layer V elicited by repetitive stimulation with 100 pulses at 20 Hz, or at 200 Hz before and during application of 10 µM NA. The three superimposed traces were recorded in a slice. C, NO release in layer V. Three traces were recorded with a NO probe, in which the anode voltage between the working Pt wire and the reference was maintained at +0·5, +0·7 or +0·9 V. D, dependence of the NO probe currents recorded in layer V (
Properties of NO release in layer V
NO release from PFs in cerebellar slices is almost linearly dependent on the stimulus intensity of PF stimulation, and exhibits marked frequency facilitation (Shibuki & Kimura, 1997). For comparison, we studied the dependence of NO release in layer V on the intensity and frequency of layer IV stimulation (Fig. 6). The dependence of cortical NO release on stimulus intensity was almost the same as that of cerebellar NO release (Fig. 6Aa and Ab). However, the frequency facilitation of NO release was not clearly observed in the auditory cortex (Fig. 6B a and Bb).
Aa, NO release in layer V elicited by repetitive layer IV stimulation (frequency, 20 Hz; duration of pulse train, 5 s) at stimulus intensities of 100, 300 or 500 µA. b, dependence of the amplitude of the NO increase (
NO release following layer IV stimulation may be triggered by direct stimulation of NOS-positive neurones or by glutamatergic inputs to these neurones. We applied CNQX, a non-NMDA receptor antagonist, and APV, an NMDA receptor antagonist, to the slices, and studied the effects of these drugs on NO release in layer V. The amplitude of the increase in NO concentration was significantly reduced by 10 µM CNQX to 34 ± 4 % (n = 10, P < 0·002, Wilcoxon signed-rank test), and by 50 µM APV to 51 ± 5 % (n = 10, P < 0·002) of that recorded before drug application (Fig. 7). Simultaneous application of 10 µM CNQX plus 50 µM APV significantly and almost completely suppressed the NO release to 3 ± 1 % (n = 13, P < 0·0005) of that before drug application (Fig. 7). However, NO release was not modulated by either 1 µM bicuculline, a GABAA receptor antagonist, or 1·5 mM phaclofen, a GABAB receptor antagonist (Fig. 7C). These results indicate that ionotropic glutamate receptors are involved in NO release in layer V.
A, NO release in layer V recorded before (Control) and during application of 10 µM CNQX alone and 10 µM CNQX plus 50 µM APV. B, NO release recorded in a similar experiment, in which 50 µM APV alone was applied to the slices before the simultaneous application of 10 µM CNQX plus 50 µM APV. C, effects of 10 µM CNQX, 50 µM APV, 10 µM CNQX plus 50 µM APV, 1 µM bicuculline (Bic) and 1·5 mM phaclofen (Pha) on NO release in layer V. The amplitude of the NO increase was normalized to the value recorded before drug application (390 ± 20 pM, n = 30). Means and
Layer-specific NO dependence of LTP
LTPII-III of the visual cortex is not blocked by 100 µM NA (Kirkwood & Bear, 1994), while LTPV in the medial frontal cortex is blocked by 100 µM MA (Nowicky & Bindman, 1993). These apparently different results could be attributed to differences in cortical areas, cortical layers or other experimental conditions. In this study, neither NA nor MA significantly affected the magnitude of LTPII-III, while these NOS inhibitors significantly suppressed the induction of LTPV. The layer-specific effects of NOS inhibitors on the induction of LTP strongly suggest that their effects cannot be attributed to non-specific side effects of these drugs. The suppressive effect of NA on LTPV was antagonized by NOR3 and Br-cGMP, indicating that the suppression of LTPV results from that of NO-induced cGMP formation. Since LTPII-III and LTPV were elicited by tetanic simulation at 100 and 200 Hz, respectively, the difference in NO dependence of LTP in this study might be due to the difference in stimulus patterns, as reported for hippocampal LTP (Lum-Ragan & Gribkoff, 1993). However, biased distribution of nNOS-positive neurones and NO release were also found in the auditory cortex. These results strongly suggest that NO dependence of LTP in the auditory cortex is different between the cortical layers.
NO source in the auditory cortex
Small non-pyramidal NOS-positive neurones are distributed widely in the neocortex (Bredt et al. 1991; Valtschanoff et al. 1993). Quantitative analysis of NADPH diaphorase-reactive neurones, which are thought to correspond to NOS-positive neurones (Hope et al. 1991), revealed biased distribution of NADPH diaphorase-reactive neurones in deep cortical layers in the medial prefrontal cortex (Gabbott et al. 1997). It has been reported that layer V pyramidal neurones in neocortical slices become NADPH diaphorase reactive after the slicing procedure (Divac et al. 1993). Therefore, we studied the cellular localization of nNOS several hours after the slicing procedure, and found no layer V pyramidal neurone stained with an anti-nNOS antibody (Fig. 4A). However, the immunohistochemical staining in the present study demonstrated that there were more cell bodies and processes of nNOS-positive non-pyramidal neurones located in the deeper cortical layers of the auditory cortex. In accordance with these morphological data, we detected transient NO release in the neocortex with electrochemical NO probes. The amplitude of the NO increase in layer II-III was only one-seventh of that in layer V. These results strongly suggest that the NO source responsible for the induction of LTPV in the auditory cortex is small non-pyramidal neurones in deep cortical layers.
Characteristics of NO release in the neocortex
NO release from PFs in cerebellar slices has been measured with NO probes of the same type as those used in this study (Shibuki & Kimura, 1997). It is, therefore, interesting to compare the properties of NO release between the neocortex and the cerebellum. The amplitude of NO release in layer V of the neocortex was less than 10 % of that in cerebellar slices (Shibuki & Kimura, 1997). This difference might be attributed to the lower density of nNOS in the neocortex compared with that in the cerebellum (Huang et al. 1993).
Another difference in the properties of NO release is that frequency facilitation of NO release was clearly observed in the cerebellum but not in the neocortex (Fig. 6B). Activity of nNOS is modulated by Ca2+ via calmodulin (Bredt & Snyder, 1994). Therefore, the difference in the frequency facilitation suggests that the dependence of stimulus-evoked local [Ca2+] rise around nNOS molecules on the stimulus frequency may be different between the neocortex and the cerebellum. In accordance with this possibility, NO release in the neocortex was blocked by simultaneous application of CNQX and AVP, while NO release from PFs in the cerebellum is not affected by these drugs and is triggered by Ca2+ influx through voltage-gated Ca2+ channels (Shibuki & Kimura, 1997). The N-terminal of nNOS molecules has a domain which binds postsynaptic density proteins (Brenman et al. 1996). Therefore, Ca2+ influx through postsynaptic NMDA receptors is likely to efficiently trigger NO release in the neocortex. Postsynaptic non-NMDA receptors, which are also Ca2+ permeable in hippocampal non-pyramidal neurones (Isa et al. 1996), are also likely to be responsible for the NO release in the neocortex. Non-pyramidal NOS-positive neurones are GABAergic (Valtschanoff et al. 1993). However, GABA receptor antagonists, which are expected to block the GABAergic receptors on GABAergic terminals (Misgeld et al. 1995), had no clear effect on the NO release in layer V (Fig. 7C). Therefore, Ca2+ influx through voltage-gated Ca2+ channels at the axon terminals, which is important for NO release from cerebellar PFs, may not have an essential role in the NO release in layer V. These differences in the site of Ca2+ influx triggering NO release might explain the differential frequency facilitation between the neocortex and the cerebellum.
Target of NO in the neocortex
In this study, we used field potential recording for the analyses of LTP. Since the neocortical field potentials are thought to be generated mainly by the activities of pyramidal neurones (Rall, 1962), the NO-dependent LTPV is probably elicited in the synapses formed on layer V pyramidal neurones. This possibility is in accordance with a previous study which showed that monosynaptic EPSPs recorded in layer V pyramidal neurones in the medial frontal cortex exhibit NO-dependent LTP (Nowicky & Bindman, 1993). Soluble guanylate cyclase, one of the target molecules of NO signalling, is present in neocortical pyramidal neurones (Ariano et al. 1982). These results suggest that the target of NO signalling regarding the induction of NO-dependent LTPV is the synapses formed on layer V pyramidal neurones.
Physiological role of NO signalling in the auditory cortex
In this study, the induction of LTPV in the auditory cortex was NO dependent, although ocular dominance shifts in the visual cortex of young animals are insensitive to NOS inhibitors (Reid et al. 1996; Ruthazer et al. 1996). One explanation for this apparent discrepancy is that LTPV does not have an essential role in the formation of ocular dominance columns in the developing brain. We recorded LTPV in the auditory cortex of adult rats (4-7 weeks old), and no age-dependent difference was found regarding the LTP experiments. However, it is reported that LTP in young animals is different from that in adult animals (Komatsu, 1994). Therefore, NO-dependent LTPV might have a physiological role in adult animals but not in the development of neural circuitry in young animals.
Layer V pyramidal neurones in the auditory cortex project to subcortical structures such as the cochlear nucleus (Weedman & Ryugo, 1996) and the inferior colliculus (Games & Winer, 1988). Therefore, LTPV in the auditory cortex may serve to provide long-term facilitation of feedback mechanisms in auditory information flow. Due to the diffusiveness of NO, co-operative activities of many neurones are expected to be involved in the feedback mechanisms. Although establishment of a behavioural test for evaluating the function of the auditory cortex is required before investigation of the role of LTPV would be possible, the layer-specific NO dependence of LTPV in the auditory cortex may provide a good opportunity to pharmacologically isolate the function of LTPV from that of neocortical LTP of other types.
We thank Y. Tamura and N. Taga for technical assistance. This work was supported by grants from the Japanese Government, Toyota RIKEN and the Uehara Foundation.
Corresponding author
K. Shibuki: Department of Neurophysiology, Brain Research Institute, Niigata University, 1 Asahi-machi, Niigata 951-8585, Japan.
Email: shibuki{at}bri.niigata-u.ac.jp
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Figure 1. NO dependence of LTPII-III
) tetanic stimulation of layer IV. c, field potentials recorded before and 30 min after () tetanic stimulation in the presence of 10 µM NA. B, time course of LTPII-III recorded in normal medium (Control; 
) and in the presence of 10 µM NA (
). The field potential amplitudes were normalized to the averaged value of three consecutive traces recorded immediately before tetanic stimulation. Each symbol and bar represents the mean and
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Figure 2. NO dependence of LTPV
) tetanic stimulation. d, field potentials recorded before and 30 min after () tetanic stimulation in the presence of 10 µM NA. B, time course of LTPV recorded in normal medium (Control; ) and in the presence of 10 µM NA (). Each symbol and bar represents the mean and
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Figure 3. Reversal of the suppression of LTPV
) tetanic stimulation. In this experiment, 10 µM NA was present in the perfusion medium throughout the recording and 5 µM NOR3 was applied to the slices for a few minutes before tetanic stimulation and during a period of the stimulation. b, time course of LTPV elicited in the presence of 10 µM NA plus 5 µM NOR3 (). LTPV elicited in the presence of 10 µM NA alone () or 5 µM NOR3 alone () is also shown for comparison. Ba, field potentials in layer V recorded before and 30 min after () tetanic stimulation. In this experiment, 10 µM NA was present in the perfusion medium throughout the recording and 1 mM Br-cGMP was applied to the slices for a few minutes before tetanic stimulation and during a period of the stimulation. b, time course of LTPV elicited in the presence of 10 µM NA plus 1 mM Br-cGMP (), 10 µM NA alone () or 1 mM Br-cGMP alone (). In Ab and Bb, each symbol and bar represents the mean and
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Figure 4. Immunohistochemical staining of the auditory cortex with an anti-nNOS antibody
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Figure 5. NO release in the auditory cortex recorded with NO probes
) or currents recorded by the same probe induced by application of 30 µM NO () on the anode voltage. The current amplitude was normalized to that recorded at the anode voltage of +0·9 V. These data represent means and
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Figure 6. Dependence of NO release on stimulus parameters
) on the stimulus intensity. The amplitude of the NO increase was normalized to the value recorded at an intensity of 500 µA (362 ± 53 pM, n = 5). For comparison, data recorded in cerebellar slices () are also plotted (reproduced from Shibuki & Kimura, 1997). Ba, NO release in layer V elicited by repetitive layer IV stimulation (intensity, 500 µA; duration of pulse train, 5 s) at frequencies of 5, 10, 20 or 40 Hz. b, dependence of the amplitude of the NO increase () on the stimulus frequency. The amplitude of the NO increase was normalized to the value recorded at a frequency of 20 Hz (377 ± 33 pM, n = 5). Data from the cerebellum () are also shown (reproduced from Shibuki & Kimura, 1997). The abscissa is shown in logarithmic scale. In Ab and Bb, values shown are means and
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Figure 7. Block of NO release by glutamate receptor antagonists
DISCUSSION
Top
Abstract
Introduction
Methods
Results
Discussion
References
REFERENCES
Top
Abstract
Introduction
Methods
Results
Discussion
References
Acknowledgements
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