Cardiff School of Biosciences, Cardiff University and , 2 Departments of Neurobiology, Psychology and Psychiatry, University of California at Los Angeles, CA, USA
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Abstract |
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Introduction |
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Experience-dependent plasticity in the cerebral cortex is known to last longer than a period of hours. Plastic changes in cat visual cortex last for the lifetime of the animal if the deprivation is maintained to the end of the critical period (Hubel and Wiesel, 1970), and changes within the critical period take days even to begin to reverse (Mioche and Singer, 1989
; Blakemore et al., 1981
). Plastic changes in layers II/III of the somatosensory cortex do not necessarily last for the lifetime of the animal because these layers do not have a critical period. Neurones are therefore always adapting to the recent history of sensory stimulation. However, the rate of change is relatively slow in the somatosensory cortex, with changes in receptive fields taking many days to accumulate (Glazewski and Fox, 1996
). Because changes cannot accumulate in this way if plasticity decays rapidly, plasticity must last at least several days in this system, which is consistent with the finding that vibrissae regrowth for a period of 810 days after deprivation does not negate the effect of the preceding deprivation.
It is not known whether experience-dependent plasticity requires protein synthesis and, in particular, whether CREB is involved. Experiments show that CRE-mediated gene transcription is increased in the spared barrel during vibrissae deprivation (Barth et al., 1999) which could indicate a role for CREB. We have therefore studied whether CREB is necessary for plasticity in barrel cortex by looking at the effects of vibrissae deprivation on receptive field plasticity in layers II and III of mice lacking the alpha and delta isoforms of CREB. This mutation has been shown to affect hippocampal LTP and memory in mice (Bourtchuladze et al., 1994
). Since previous results have shown that plasticity mechanisms differ in young (12 months) and adult (>6 month) mice (Glazewski et al., 1996
), we have recorded from both age groups. In this paper we describe results which argue in favour of a role for CREB in neocortical experience-dependent plasticity.
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Materials and Methods |
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The CREB alpha/delta knock-out mice were maintained in a C57BL/6 genetic background and derived from C57BL/6 heterozygote (+/) males from Cold Spring Harbor (Bourtchuladze et al., 1994). The homozygous, heterozygous and wild-type animals used in the experiments described here were F2 offspring from a cross between heterozygotes (in the C57BL/6 background) with 129SV (+/+) mice from Harlan UK. The heterozygotes in the F1 generation were interbred (cousin matings) to produce the F2 generation. The F2 generation was not bred to produce experimental animals (Silva et al., 1997
).
In the adolescent age group (P2835), a total of seven undeprived (four +/+, three /+, zero /) and 16 deprived animals (four +/+, six /+, six /) were used to assess the effect of genotype on vibrissae deprivation. In the adult age group (>6 months), a total of 14 undeprived (six +/+, four /+, four /) and 16 deprived mice were used (six +/+, four +/, six /). In all experiments the experimenter was blind to the genotype. In addition, two mice of each genotype were used for the Western analysis (two +/+, two +/, two /).
Vibrissae Deprivation
The method of depriving the vibrissae has been described before (Fox, 1992) and consists of removing the vibrissa by applying slow tension to the base of the vibrissa. This method has been shown not to cause damage to the vibrissae follicle innervation (Li et al., 1995
). All the large caudal vibrissae except for D1 were removed starting from P2835 in the case of the adolescent age group and at least P183 in the case of the adult animals. Vibrissae were checked every other day for regrowth and kept deprived for 18 days. Vibrissae were allowed to regrow for 810 days before the day of recording.
Surgery and Anaesthesia
Anaesthesia was induced with metofane (Arovet AG, Switzerland) and maintained with urethane (1.5g/kg body wt). Anaesthetic depth was monitored throughout the experiment by testing reflexes and observing the spontaneous firing rate of neurones. Supplements of urethane were administered to maintain a state where the hindlimb withdrawal reflex was sluggish.
For cortical recordings, the skull was thinned 2.53.5 mm lateral to the midline and ~13 mm caudal to bregma by careful drilling. From this position it was possible to reflect a small part of the skull with a hypodermic needle and introduce the electrode through the resultant hole. The dura was left intact as the carbon fibre electrodes were able to pass through it.
Stimulation
The stimulus consisted of a 200 µm deflection of the vibrissa applied ~10 mm from the face (1° deflection) and delivered at 1 Hz. The stimulator was a lightweight glass capillary touching the vibrissa attached to a fast piezoelectric bimorph wafer. All stimulus parameters were identical to those used in previous studies (Glazewski et al., 1996).
Recording
Cortical neurones were recorded using single-barrel carbon fibre micro electodes (Armstrong-James et al., 1980). The signal was band-passed between 700 Hz and 7 kHz and spikes discriminated using a voltage window discriminator. Post-stimulus time histograms and raster plots were generated on-line and primary data stored for later analysis using Spike 2 software (CED, Cambridge, UK).
Sampling
Neurones were sampled evenly approximately every 50 µm throughout the penetration. Cells were isolated by moving the electrode to the next position and discriminating a cell using its spontaneous activity. The electrode position was then adjusted by ~1020 µm to optimize discrimination. If, when a stimulus was applied, a larger spike occurred that was more easily discriminated it was often, but not always, used for study instead.
Histology
Cytochrome Oxidase
At the end of recording from each penetration, a small focal lesion (1.0 µA, DC, 10 s tip negative) was made at a site of known depth in layer IV. The cortex was flattened, processed for cytochrome oxidase histology as described before (Wong-Riley, 1979; Fox, 1992
) and the location of each recording penetration identified within the barrel field. In this way we could identify the principal vibrissa for each recorded cell.
Immunohistochemistry
To assess the distribution of CREB-immunoreactive cells within barrel cortex, several different experimental conditions were used. In all cases, animals were anaesthetized with metofane, decapitated and then brains were dissected out into cold ACSF. In the first case (n = 4 animals; Table 1) tissue was cut into 400 µm thick slices and then fixed in 3% paraformaldehyde, 0.1 M phosphate buffer on ice for 2 h before cryoprotection with sucrose and further sectioning to 50 µm thickness. In the second case (n = 3 animals; Table 2
), brains were fixed whole under the same conditions, then subject to an overnight incubation in a light detergent (0.02% Nonidet P-40, 0.01% sodium deoxycholate) buffer solution at 37°C before cryoprotection and sectioning to 25 or 50 µm thickness. This second procedure was performed in an attempt to permeabilize the tissue fully and breakdown extracellular proteins that might impede antibody access. Floating sections were blocked in 10% heat-inactivated goat serum and incubated in a 1:1000 dilution of anti-CREB antibody (New England Biolabs, Beverly, MA) at 4°C overnight (Table 1
) or 1:330 (Table 2
) dilution of anti-CREB antibody (New England Biolabs) at 4°C overnight. Secondary antibodies (from Sigma, St Louis, MO; using alkaline-phosphatase-conjugated antibodies for quantitation, FITC-conjugated for Fig. 1
) were applied for several hours at the manufacturer's recommended dilution. [FITC-conjugated secondary antibodies (Sigma) were used at a dilution of 1:160 for 2 h at room temperature.] All washes and antibody solutions included 0.51% Triton X-100. Sections were counterstained with 0.1% propidium iodide to label all nuclei. Raw cell counts were averaged from four fields per layer per animal (Tables 1 and 2
). A different anti-CREB antibody (Upstate Biotechnology Inc., Lake Placid, NY) was also used in parallel to these experiments (Table 2
) and showed similar results to the New England Biolabs antibody.
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All data were analysed using post-stimulus time histograms (PSTHs) and latency histograms (LHs). Response magnitude to a particular vibrissa was defined as the number of spikes per stimulus occurring between 5 and 50 ms after the stimulus minus the spontaneous activity occuring during an identical time period. The modal latency was used to describe the response latency of the neurone. For a complete description see Armstrong-James and Fox (1987).
To estimate the shift to the spared vibrissa, the relative response of each cell to the principal vibrissa stimulation and spared D1 vibrissa was calculated as F = (spared)/(spared + principal), where the response to each whisker was the number of spikes produced to 50 stimuli within 550 ms of deflecting the vibrissa. The weighted vibrissae dominance index (WVDI) was then calculated for each animal based on the F-values for each cell recorded (see Glazewski et al., 1996 for description). This value is similar to a quantitative ocular dominance index (Kasamatsu et al., 1979).
To estimate expansion of the D1 domain, the position of each penetration was charted on a standard map of the barrel field and the response to D1 vibrissa stimulation averaged for the cells in layers II/III of that penetration (see Fig. 4).
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Because the surround receptive field responses were slightly larger in mutants versus wild-types it was necessary to introduce an index of the spared vibrissa's response relative to its initial response level. This was done by comparing responses to the spared vibrissa to responses to symmetrically displaced near neighbouring vibrissa. The relative change in D1 responses was then given by G = (spared vibrissa response)/(spared + surround). Like the vibrissa dominance index this value varies between 0 and 1. In normal animals it is 0.5, indicating equal responses from all surround receptive field vibrissae. For statistical tests G was calculated for each animal from the average spared and surround receptive field responses for that animal.
Western Blots
A small core of tissue was extracted from the barrel cortex of adult wild-type, heterozygote and homozygote mice using a plastic pipette tip cut down to a diameter of ~3 mm. The remaining cortex was flattened and processed for cytochrome oxidase histology to confirm whether the barrel cortex had indeed been sampled.
The tissue sample was rapidly removed and frozen in liquid nitrogen. Tissue was thawed in denaturing reducing buffer and sonicated on ice for 5 min. Equal volumes of each sample were loaded into a 12.5% denaturing polyacrylamide gel and separated by electrophoresis. Western blotting was performed as previously described (Laemmli, 1970). Anti-CREB antibody (New England Biolabs) was used at a 1:1000 dilution, and alkaline phosphatase-conjugated secondary antibody (Sigma) was used at a concentration of 1:200. Blots were developed using an alkaline phosphatase developing solution (Biorad).
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Results |
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Although CREB has been reported to be present in most cells in the brain (Herdegen et al., 1993; Blendy et al., 1996
), expression levels have not been analysed quantitatively. We therefore used immunohistochemistry to examine the distribution of CREB in the barrel cortex (Fig. 1
). Extragranular layers showed a high proportion of CREB-immunoreactive nuclei, where 53.8% of nuclei were CREB positive (see Table 1
). This contrasted with fewer CREB-positive nuclei in layer IV, where only 26.5% of total cells were immunoreactive (Table 1
). In general, the histochemical signal from layer IV nuclei was also weaker than that observed in layer II/III. The lower levels of CREB seen in layer IV relative to layers II/III (Fig. 1
) are therefore a combination of fewer CREB-positive cells and weaker immunoreactivity per cell.
To check whether the CREB antibody had sufficient access to the cells we studied a further three cases in which permeability had been further enhanced (see Materials and Methods). The results can be seen in Table 2 which show that, overall, the percentage of CREB-positive cells can be increased to 69.1% in layers II/III and 49.8% in layer IV. This implies that access of the antibody is a factor in absolute quantification of CREB immunoreactivity. However, under both conditions we consistently saw a population of cells characterized by small nuclei that were CREB immunonegative and under both conditions we saw some cells showing far less anti-CREB immunoreactivity than others in the same layer. Therefore, the general conclusions, that not all cells in barrel cortex express CREB and that fewer cells express CREB in layer IV than layers II/III, are valid for both sets of tissue treatment.
The Effect of Vibrissae Deprivation in Adult Mice
Three measures were used to assess the degree of plasticity present in the CREB knock-out mice: the absolute response magnitude of the spared vibrissa relative to deprived surround receptive field vibrissae, the vibrissae dominance shift, and the spatial map of the vibrissa domain (see Materials and Methods). Each is described in the following sections for the adult animals.
Absolute Response Magnitude in Wild-types
As shown in Figure 2, the spared vibrissae response increased 3.7-fold in barrel-columns surrounding the D1 barrel-column relative to the surround receptive field recorded in the same animals (mean spared response ± SEM = 0.93 ± 0.07 spikes per stimulus; surround response = 0.25 ± 0.08). This difference was found to be significant [F(1,10) = 48.7, P < 103]. However, there was no effect of deprivation on the principal vibrissa response nor on the surround receptive field response in homozygote or heterozygote mutants, suggesting that where plasticity occurred it was specific to the spared vibrissa (see Table 3
).
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In deprived heterozygotes, the spared D1 vibrissa response increased 3.5-fold relative to surround receptive field responses recorded in the same animals (mean spared response = 1.13 ± 0.13, mean surround response = 0.32 ± 0.06 spikes per stimulus). This difference was found to be significant [F(1,8) = 32, P < 0.0005, MANOVA]. In homozygotes, the spared vibrissa response appeared to be 1.7-fold greater than the surround receptive field responses as a result of deprivation (mean spared vibrissa response = 0.97 ± 0.21, mean surround response = 0.55 ± 0.07 spikes per stimulus) but this difference was not found to be significant [F(1,7) = 2.0, P = 0.19, MANOVA].
There was no effect of deprivation on the principal vibrissa response nor on the surround receptive field response in homozygote or heterozygote mutants, again suggesting that where plasticity occurred it was specific to the spared vibrissa (see Table 3).
Comparison of Spared and Deprived Surround Receptive Field Responses
Data shown in Figure 2 (top panel) indicate that the surround receptive field responses are larger in mutants than in wildtypes. The average surround receptive field response is close to 0.5 for mutants but approximately half that value at 0.25 spikes per stimulus for wild-type littermates. A two-way ANOVA showed that deprivation had no effect on the surround receptive field response (P = 0.55) but that genotype had a large effect (homozygote surround = 0.52, wild-type surround = 0.26; P < 0.001) with no interaction between the two (P = 0.4). This implies that the spared vibrissa response potentiates from a higher baseline level in homozygotes and heterozygotes and conversely, from a lower level in wild-types.
In order to take into account the effect of the naturally larger surround receptive field response in the mutants, a simple ratio of spared to surround response was calculated for deprived and undeprived animals. As can be seen from Figure 2 (lower panel), most values lie very close to unity for undeprived controls independent of genotype. However, while the spared vibrissa response increases 3.8-fold for wild-types and 3.5-fold for heterozygotes, it increases less for homozygotes (1.7-fold). This shows that homozygotes have a reduced capacity for receptive field potentiation compared to wild-types or heterozygotes.
We tested whether plasticity was greater or smaller in the mutants by comparing the spared vibrissa response versus the deprived surround vibrissa response across genotype [relative spared vibrissae response = (spared response)/(spared + surround); see Materials and Methods]. There was an effect of genotype on the relative spared vibrissa response [F(2,18) = 7.4, P < 0.005]. Post-hoc tests revealed that this was due to the homozygotes showing less spared vibrissa potentiation relative to their surround receptive fields than either heterozygotes or wild-types. The relative spared vibrissae response was less in homozygotes than in wild-types [t(12) = 8.2, P < 0.02] or heterozygotes [t(12) = 10.9, P < 0.01] but equal in heterozygotes and wild-types [t(12) = 0.14, P = 0.71].
Vibrissae Dominance in Wild-types
Figure 3 shows the vibrissae dominance histograms for deprived and undeprived wild-types. The shift in vibrissa dominance is clear from the increased number of cells responding to the spared D1 vibrissa after deprivation (86% versus 55% in controls; grey bars) and the increased number producing a greater response to D1 versus their own principal vibrissa (40% versus 5% in controls; bins > 0.5). Accordingly, the WVDI increased from 0.17 ± 0.04 in undeprived animals to 0.42 ± 0.05 in deprived wild-type adults and this effect was significant [t(12) = 3.56, P < 0.01].
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A similar degree of plasticity to that shown by the wild-types was apparent in the heterozygote group. The number of cells driven by D1 increased with deprivation (to 91% from 53% in controls) and the number of cells responding more to D1 than the principal vibrissa also increased (to 48% from 11%). The WVDI increased from 0.17 ± 0.06 in undeprived animals to 0.51 ± 0.03 in deprived heterozygotes, which was a significant change [t(9) = 5.9, P < 0.01].
The homozygotes showed only slightly less of a shift than the heterozygotes, with the number of cells responding to D1 increasing to 85% from 55% in controls and those responding more to D1 than the principal vibrissa increasing to 38% from 5%. The WVDI increased from 0.16 ± 0.04 in undeprived animals to 0.42 ± 0.05 in deprived homozygotes, which was significant [t(8) = 3.45, P < 0.01]. Taken together with the data on absolute response magnitude, this suggests that a similar number of cells are plastic in homozygotes and wild-types but that the degree of spared vibrissa potentiation is less in homozygotes than wild-types.
Spared Vibrissa Domain in Adults
Figure 4 summarizes the distribution of penetrations made in control (undeprived) wild-type mice with respect to the barrel field. The intensity of response evoked by deflecting the D1 vibrissa can be seen from the colour of the circles (see figure legend). In undeprived animals the D1 domain is confined mainly to the D1 vibrissa with very few penetrations containing cells firing at above 0.4 spikes per stimulus on average (Fig. 4A
). However after deprivation, the D1-responsive domain extends into neighbouring barrels, and neurones discharge as powerfully to D1 in neighbouring barrels as they do within the D1 barrel to D1 vibrissa stimulation (Fig. 4B
; note the increase in penetrations where cells responded above 0.8 spikes per stimulus). Some expansion of the spared D1 domain also occurs in homozygous animals (Fig. 4D
). However, note that in undeprived homozygotes several penetrations in barrels surrounding D1 respond at intermediate levels to D1 (Fig. 4C
). This is a reflection of the higher surround receptive field responses of alpha/delta mutants in general.
The Effect of Vibrissae Deprivation in Adolescent Mice
The three main indicators for plasticity described above for the adult animals (vibrissae dominance shift, expansion of the spared vibrissa domain and increase in the spared vibrissa response relative to the surround) all showed that plasticity was present and at normal levels in the homozygous adolescent animals. In homozygotes, the number of cells responding to the spared D1 vibrissa were high at 92%, as were the number of cells responding more powerfully to D1 at 59%. Their WVDI was also significantly different from the undeprived mutants [homozygotes = 0.61 ± 0.07, t(7) = 2.5, P < 0.01]. The vibrissae domain for the spared D1 vibrissa expanded into surrounding barrels (not shown) and the spared vibrissa potentiated compared to the surround vibrissae recorded in the same animals (Fig. 5).
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Up-regulation of Beta CREB
It has been reported that a normally minor isoform of CREB (beta) is up-regulated in the alpha/delta knock-outs (Blendy et al., 1996). Immunohistochemistry using polyclonal anti-CREB antibodies revealed intense staining in the barrel cortex of mutant homozygotes (data not shown). To verify that this histochemical signal came from a specific up-regulation of the beta CREB isoform, a small plug of tissue was extracted from the barrel cortex of wild-type, heterozygote and homozygote mice and subjected to SDSPAGE and Western blotting (Fig. 6
). A sample from the hippocampus was also run for comparison. The alpha/delta isoforms can be detected in both wild-type and heterozygote samples. The 39.5 kDa beta isoform is also recognized by the same antibody and is observed in the homozygotes sample (lane 3 for hippocampus, lane 6 for barrel cortex). These results confirm that the beta isoform is not only present but up-regulated in barrel cortex of the homozygous alpha/delta knock-out mutants.
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Discussion |
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Similarly, knock-out of the alpha and delta isoforms of CREB impairs the maintenance phase of LTP in the hippocampus (Bourtchuladze et al., 1994). Potentiation occurs in the null mutants for ~90 min before returning to baseline. In addition, it has been found that when LTP is induced with a tetanus that does not produce long-lasting LTP (i.e. it returns to baseline within ~4 h), CRE-mediated gene expression is not induced in the hippocampal CA1 region, whereas when LTP does not decrement and is maintained over 4 h, CRE-mediated gene expression is high (Impey et al., 1996
). These studies imply a role for CREB or a related transcription factor in maintaining plasticity.
Even though the CREB knock-out has a measurable effect in reducing plasticity, our findings suggest that it does not block plasticity as completely as it does in other systems. There are several reasons why this might be the case. First, only about two-thirds of the cells in layers II/III showed CREB immunoreactivity. Because only ~3040% of cells show a vibrissae dominance shift (even in wild-types), all of these cells would need to be CREB positive in order for the mutation to have a complete direct effect. Since some plasticity does occur, one possibility is that the two populations do not entirely overlap. Second, we have shown that the beta isoform is up-regulated in the alpha/delta knock-outs, and this may be sufficient to partially rescue cortical plasticity. Activator forms of CREM are another related transcription factor that can act to initiate transcription at the CRE site and which is up-regulated in the brain of alpha/ delta knock-outs (Blendy et al., 1996; see Silva et al., 1998).
A third reason for the partial effect of this knock-out is related to the way in which CREB is thought to be activated. It has been suggested that the mechanism by which CREB activates gene transcription is dependent on CREB activator and repressor forms. Transcription only proceeds when the activator function overcomes the blocking effect of repressors. Theoretically, both are phosphorylated during learning or induction of synaptic plasticity, but the activator form itself inactivates slower than the repressor form (Yin et al., 1994). This idea could explain why in CREB knock-out mice, memory induced by massed training is more affected than memory induced after spaced training (Bourtchuladze et al., 1994
; Yin et al., 1994
; Kogan et al., 1996
). Spaced training may favour the accumulation of activator forms of CREB-related genes because the inactivation of repressors is faster than the inactivation of activators. This hypothesis suggests that the timing of stimuli may be critical in revealing the phenotype of CREB mutants. Thus it is possible that the conditions of our experiment are more akin to spaced training since the deprivation lasts many days. On the other hand we do not know what events in the animal's behaviour may be analogous to training. CREB activation may only be induced when the animal is actively exploring and attending to particular stimuli and the timing interval between these events is unknown. A paradigm with more control of the timing of vibrissae plasticity may be useful in future studies on this subject.
One unexpected finding in the present study is that the surround receptive field responses are greater in the homozygous and heterozygous animals compared to wild-type littermates. This effect was present in the young and the older animals though it was more pronounced in the adults. This suggests that CREB may play some role in development of surround receptive field structure in the cortex. Previous studies have shown that, in the rat, surround receptive fields are largely produced via intracortical connection between the barrels (Armstrong-James and Fox, 1987; Armstrong-James et al., 1992
; Fox, 1994
). Intracortical connections are set up during the second postnatal week in mice (McCasland et al., 1992
), and it is possible that CREB may play some role in guiding this process.
In summary, adult but not adolescent mutants showed a distinct phenotype. The magnitude of surround receptive field responses were greater in mutants than in wild-types before vibrissa deprivation, and following vibrissae deprivation homozygote animals exhibited a smaller-fold increase in potentiation of the spared vibrissa than did wild-types. In addition, the effect of the mutation on plasticity may have been attenuated by up-regulation of a novel isoform of CREB in barrel cortex. These data are consistent with a role for CREB in neocortical plasticity.
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Notes |
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Address correspondence to Professor K. Fox, Cardiff School of Biosciences, Cardiff University, Museum Avenue, Cardiff CF1 3US, Wales, UK. Email: foxkd{at}cardiff.ac.uk.
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Notes |
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References |
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