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The Journal of Neuroscience, December 1, 2002, 22(23):10313-10323
Normal Development of Embryonic Thalamocortical Connectivity in
the Absence of Evoked Synaptic Activity
Zoltán
Molnár1, *,
Guillermina
López-Bendito1, *,
Juan
Small1,
L. Donald
Partridge3,
Colin
Blakemore2, and
Michael C.
Wilson3
1 Department of Human Anatomy and Genetics, University
of Oxford, Oxford, OX1 3QX, United Kingdom, 2 University
Laboratory of Physiology, University of Oxford, Oxford, OX1 3PT, United
Kingdom, and 3 Department of Neurosciences,
University of New Mexico Health Sciences Center, Albuquerque, New
Mexico 87131
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ABSTRACT |
This study is concerned with the role of impulse activity and
synaptic transmission in early thalamocortical development. Disruption
of the gene encoding SNAP-25, a component of the soluble N-ethylmaleimide-sensitive factor attachment protein
(SNAP) receptor complex required for regulated neuroexocytosis,
eliminates evoked but not spontaneous neurotransmitter release
(Washbourne et al., 2002 ). The Snap25 null mutant mouse
provides an opportunity to test whether synaptic activity is required
for prenatal neural development. We found that evoked release is not
needed for at least the gross formation of the embryonic forebrain,
because the major features of the diencephalon and telencephalon were normal in the null mutant mouse. However, half of the homozygous mutants showed undulation of the cortical plate, which in the most
severely affected brains was accompanied by a marked reduction of
calbindin-immunoreactive neurons. Carbocyanine dye tracing of the
thalamocortical fiber pathway revealed normal growth kinetics and
fasciculation patterns between embryonic days 17.5 and 19. As in normal
mice, mutant thalamocortical axons reach the cortex, accumulate below
the cortical plate, and then start to extend side-branches in the
subplate and deep cortical plate. Multiple carbocyanine dye placements
in the cortical convexity revealed normal overall topography of both
early thalamocortical and corticofugal projections.
Electrophysiological recordings from thalamocortical slices confirmed
that thalamic axons were capable of conducting action potentials to the
cortex. Thus, our data suggest that axonal growth and early topographic
arrangement of these fiber pathways do not rely on activity-dependent
mechanisms requiring evoked neurotransmitter release. Intercellular
communication mediated by constitutive secretion of transmitters or
growth factors, however, might play a part.
Key words:
thalamus; cortex; mouse; carbocyanine dyes; synaptogenesis; synaptic activity; SNAP-25
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INTRODUCTION |
In the adult nervous system, neural
communication is mediated primarily through chemical synapses, where
neurotransmitter release is evoked by presynaptic action potentials
(APs). Neurotransmitter secretion has also been demonstrated in growing
axons before target contact and synapse formation, suggesting that
neurotransmitters could play a role in pathway guidance and target
selection (Girod et al., 1995; Verderio et al., 1999). Patterns of
impulses and regulated transmitter release may guide the precise
topography of certain neuronal projections (Katz and Shatz, 1996).
Thalamocortical development in the mammalian brain is a highly ordered
process that is well suited for evaluating these mechanisms. Thalamic
axons pass among populations of neurons on their way to the
intermediate zone of the telencephalon (Molnár et al., 1998a),
arrive in the subplate, and extend side branches radially into the
developing cortical plate (Naegele et al., 1988; Ghosh and Shatz,
1992a; Catalano et al., 1996). Once thalamic fibers enter layer 4, the
topography becomes fairly precisely established (Agmon et al., 1993;
Krug et al., 1998). Its final refinement involves activity-dependent
strengthening of synaptic interactions (Stryker and Harris, 1986;
Molnár and Hannan, 2000; Erzurumlu and Kind, 2001), and synaptic
input to cortical neurons can influence their morphological
differentiation (Hannan et al., 2001). However, it is less clear
whether neurotransmission is involved in the early stages of formation
of the thalamocortical projection.
It has been suggested that thalamic fibers form temporary connections
on subplate neurons before advancing and forming their ultimate
synapses in cortical layer 4 (Friauf and Shatz, 1991). Early neural
activity within the subplate or lower cortical layers might stabilize
certain side-branches and thereby select the appropriate cortical
target area (Ghosh and Shatz 1992a,b; Catalano and Shatz, 1998;
Molnár et al., 2000). However, the nature of this neural activity, and whether it involves evoked release of neurotransmitter, has not been resolved.
To address these issues, we examined the development of
thalamocortical projections in the absence of evoked
neurotransmitter release in SNAP-25-deficient fetal mice. SNAP-25,
together with syntaxin-1 and vesicle-associated membrane
protein-2, forms the core soluble
N-ethylmaleimide-sensitive factor attachment protein (SNAP)
receptor (SNARE) complex, which plays an essential role in exocytotic
release of neurotransmitter (Südhof, 1995). Mice homozygous for a
Snap25 null mutation develop to term, and fetal brain
development appears superficially normal, although evoked neurotransmitter release is eliminated entirely (Washbourne et al.,
2002). SNAP-25-deficient neurons extend axons that terminate in
synapses where spontaneous AP-independent release still occurs, but APs
do not trigger neurotransmission. Thus, genetic ablation of SNAP-25
expression appears selectively to disable the vesicular processes
responsible for evoked synaptic transmission, leaving intact membrane
trafficking for axon outgrowth and exocytosis for spontaneous
neurotransmitter secretion.
If activity-dependent neurotransmitter release is necessary for the
guidance and targeting of thalamic and cortical projections, the
topography of these fibers should be disorganized in the
Snap25 null mutant brain. Thalamocortical projections might
show aberrant branching at the subplate and fail to enter the correct
region of overlying cortical plate. In fact, our results demonstrate that the absence of evoked neurotransmitter release does not grossly disrupt these processes.
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MATERIALS AND METHODS |
Animals
SNAP-25-deficient mice and littermate controls were obtained by
mating mice heterozygous for a null mutation of the Snap25 gene, with exons 5a and 5b deleted, which was backcrossed onto a
C57BL/6J background for four to five generations, as described previously (Washbourne et al., 2002). Late-stage homozygote null mutants are completely paralyzed, although the heart beats, presumably because of the presence of electrically active nodal cells and electrical cardiac conduction pathways. Embryos usually survive until
birth but die immediately after parturition, probably from respiratory
failure. This effectively limited our studies to the embryonic period.
Mice, which have a gestation period of 19-20 d, were time-mated at the
University of New Mexico (Albuquerque, NM), fetuses were collected
between embryonic day (E) 17.5 and E19 on the basis of the plug date,
which was defined as E0. Littermates were analyzed without previous
confirmation of the genotype. The mice were housed in a pathogen-free
barrier facility at the Association for Accessment and Accreditation of
Laboratory Animal Care International-approved Animal Resource
Facility at the University of New Mexico Health Sciences Center campus.
All animal procedures were performed in accordance with the guidelines
of the University of New Mexico Laboratory Care and Use Committee and
the National Institutes of Health.
For routine genotype screening, a semiquantitative PCR assay, based on
amplification of the exon 5 region, was conducted on tail DNA, in which
comparable primers for the IL-1 gene (proximal to Snap25
on mouse chromosome 2) were used as an internal control (Washbourne et
al., 2002). Occasionally, genotypes were confirmed by Southern
blotting analysis, but in the vast majority of cases the PCR assay
proved sufficient to assign the appropriate Snap25 genotype.
The genotypes of litters from heterozygote matings exhibited a
Mendelian distribution (see Table 1).
Histology, immunohistochemistry, and axonal tracing
For experiments conducted on fixed tissue, fetuses were removed
by Cesarean section under general anesthesia. The fetuses were then
decapitated, and the heads were fixed in 4% paraformaldehyde in 0.1 M phosphate buffer (PB), pH 7.4, overnight.
Nissl staining. To study cytoarchitecture, fixed brains were
sectioned coronally at 60 µm (Leica VT 1000S Vibroslicer) and stained
with 0.5% cresyl violet solution. Nissl-stained sections were examined
by light microscopy and photographed with a Leica DC 500 digital
camera. Plates were assembled using Photoshop 6.0 (Adobe).
Immunohistochemistry. Brains were studied from E17.5 and
E18.5 homozygous Snap25+/+
wild-type (WT) mice, heterozygote (HT) mice, and homozygous, Snap25 /
null mutant, "knock-out" (KO) animals (see Table 1). After
overnight fixation in 4% paraformaldehyde/0.1 M
PB, the brains were embedded in 4% agarose and sectioned coronally at
60 µm. Free-floating sections were incubated in 10% normal goat
serum (NGS) diluted in 50 mM Tris buffer (TB), pH
7.4, containing 0.9% NaCl [Tris-buffered saline (TBS)], with or
without 0.2% Triton X-100, for 1 hr. Sections were then incubated for
48 hr in affinity-purified polyclonal antibodies in TBS containing 1%
NGS: (1) anti-calbindin (Swant; 1:2000), (2) anti-calretinin (Swant;
1:2000), (3) anti-SNAP-25 (a gift from Dr. H. Hirling, University of
Lausanne; 1:500), (4) anti-L1 (Chemicon, 1:100), (5)
anti-phospho-Histone H3 (Upstate Biotechnology, 1:500), and (6) a
monoclonal antibody against reelin clone E4 (de Bergeyck et al., 1998)
(gift from Dr. A. Goffinet, University Louvain Medical School,
Brussels, Belgium).
After washes in TBS, the sections were incubated for 2 hr in
biotinylated goat anti-rabbit or anti-mouse IgG (Vector Laboratories, Burlingame, CA) diluted 1:100 in TBS containing 1% NGS. They were then
transferred into avidin-biotin-peroxidase complex (ABC kit, Vector
Laboratories) diluted 1:100 for 2 hr at room temperature. Peroxidase
enzyme activity was revealed using 3,3'-diaminobenzidine tetrahydrochloride (0.05% in TB, pH 7.4) as chromogen and 0.01% H2O2 as substrate. Sections
were rinsed, dehydrated, and mounted in Eukitt mounting media. For
control experiments, the primary antibody was replaced by 0.2% Triton
X-100 in PBS and then reacted as above. These control sections showed
no positive immunoreactivity. Photomicrographs were prepared by light
microscopy with a Leica DC 500 digital camera.
Axonal tracing from the thalamus with carbocyanine dyes.
Carbocyanine dyes were used to trace axon pathways (Godement et al., 1987) in fixed brains of littermate mice from E17.5 to E19. We examined
two litters (14 individuals altogether). For detailed methods see
Molnár et al. (1998a). The brains were removed from the skull
under a dissecting microscope, and both hemispheres were used for axon
tracing. One or more tiny individual crystals (0.1-0.3 mm diameter) of
fluorescent carbocyanine dye were inserted, under an operating
microscope, into the diencephalon (after transecting the brainstem
rostral to the colliculi to expose the posterior thalamus) or into the
surface of the neocortex, using a fine pair of forceps or a fine
stainless steel wire, under an operating microscope. The dye used for
single-dye tracing was 1,1'-dioctadecyl 3,3,3'3'-tetramethylindocarbocyanine perchlorate (DiI). In some experiments,
4-[4-dihexadecylamino)stryryl]-N-methylpyridinium iodide
(DiA) was also used (both dyes from Molecular Probes, Eugene, OR). The
depth to which the crystal was inserted into the thalamus or the cortex
was ~0.5-1 mm from the cut surface. After insertion of dye crystals,
the brains were stored in PBS, with 0.01% sodium azide to prevent
contamination, at room temperature, or at 37°C to facilitate the
diffusion of dye along axons. The incubation period ranged from 2 to 4 weeks; shorter periods were used for higher incubation temperatures.
At the end of incubation, the brains were embedded in 4% agarose (made
up in 0.9% saline), and coronal sections were cut with a Vibroslicer
(Leica VT 1000S). The sections, 50-100 µm thick, were counterstained
with bisbenzimide (Riedel-De Haen AG, Seelze-Hanover, Germany; 2.5 µg/ml in PBS) for 10 min to reveal the main cytoarchitectonic features and to confirm the presence of chromatin in back-labeled cells. The sections were coverslipped in 0.1 M PB with
glycerol (1:1), or PBS, and then sealed with nail varnish. Each series of sections was examined in a conventional fluorescence microscope, using different filters to reveal either the DiI dye or the
bisbenzimide staining. The sections were photographed with a Leica
DC500 digital camera. Numerous sections, selected in conventional
fluorescence microscopy, were subsequently examined and imaged in a
laser scanning confocal microscope (TLSM-Fluovert; Leica, Heidelberg, Germany).
Experiments on topography. In three brains from a single
litter at E18.5 (each brain representing a different genotype), we made
alternating deposits of DiI and DiA at three points 2-3 mm apart, in
either a parasagittal or a coronal row across the cortex of each
hemisphere. This was done to study the topography of both corticofugal
projections (labeled by anterograde diffusion) and thalamocortical
projections (retrogradely labeled). DiI and DiA were selected because
they are clearly distinguishable under different wavelengths of
fluorescent illumination. DiA needs slightly shorter incubation periods
(1-3 weeks at room temperature) for reliable fiber labeling.
Therefore, DiI placement was conducted 1 week before that of DiA. After
an incubation period of 2 weeks (in PBS, with 0.1% sodium azide at
room temperature), the brains were embedded in agarose and sectioned
horizontally or coronally. Each series of sections was examined in a
conventional fluorescence microscope, and selected sections were
subsequently examined and imaged in the laser scanning confocal microscope.
Electrophysiology
Fetal mice at E18.5 were obtained from time-mated females by
Cesarean section under general anesthesia, and after decapitation, the
whole fetal brain was removed and immersed in chilled oxygenated artificial CSF (ACSF) containing (in mM): NaCl 124, KCl 5, NaH2PO4 1.24, MgSO4 1.3, CaCl2 2.4, NaHCO3 26, D-glucose 10 (saturated with 95% O2 and 5% CO2).
The brains were dissected in chilled ACSF and then embedded in low
melting point agarose (4%; Invitrogen) made up in ACSF. The agar
blocks were chilled and blocked so that vertical oblique sections
(45° to coronal and sagittal planes; see Fig. 8) could be cut with a
Vibratome (Higashi et al., 1996, 2002). Sections of 400 µm thickness
were cut in low-Ca2+ ACSF containing (in
mM); NaCl 124, KCl 5, NaH2PO4 1.24, MgSO4 10, CaCl2 0.5, NaHCO3 26, D-glucose 10 (saturated
with 95% O2 and 5% CO2),
maintained at 34°C in low Ca2+ ACSF, and
then held at room temperature in normal ACSF for a minimum of 1 hr
before recording at 34°C. Of the series of sections from each brain,
one or two sections, in which bundles of thalamocortical fibers could
be followed all the way from the thalamus through the internal capsule
to the cortex were selected under a binocular microscope.
In some experiments we blocked glutamatergic synaptic transmission with
inhibitors of AMPA receptors (40 µM CNQX; Tocris) and
NMDA receptors (50 µM APV; Sigma). In some cases we
inhibited GABAA-mediated transmission with
bicuculline (20 µM; Sigma) or blocked voltage-activated
sodium channels with tetrodotoxin (TTX; 600 nM).
Borosilicate glass (A-M Systems) microelectrodes filled with 0.9% NaCl
solution (electrical resistance, 1-5 M ; tip diameter, ~3
µM) were used to record field potentials evoked in the
putative somatosensory cortex by the stimulation of the ventrobasal
(VB) complex of the thalamus. The recording microelectrode was mounted on a three-dimensional motor-driven micromanipulator (MS-314 WPI, World
Precision Instruments). Its tip was positioned roughly in the middle of
the cortical plate, at the locus of maximum response. Suprathreshold
stimulation (duration, 0.1 msec; frequency, 0.05 Hz) was elicited with
a concentric bipolar stimulating electrode (tip diameter, 200 µm).
The intensity of the stimulus was adjusted to approximately two-thirds
of that evoking the maximum response. Toward the end of each
experiment, a few previous recording positions were revisited to
confirm that there was no change in the response during the recording session.
Field potentials were averaged for five stimulation trials with the
stimulus applied every 20 sec. We used this very low rate of
stimulation to ensure that synaptic transmission, if present, would not
suffer from the exhaustion that is characteristic of immature synapses.
The electrical signals were averaged on-line using pCLAMP6 (Axon
Instruments) software and a Digidata 1200 A/D converter.
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RESULTS |
Our description is based on experiments involving
approximately equal numbers of
Snap25/
homozygous null mutant [KO] animals, HTs, and homozygous
Snap25+/+ WT specimens from the
same litters (Table 1). The appearance of
the embryonic brain structures in both WT and HT mice in the present
study conformed at every stage to the previous descriptions of brain
development in normal mice (Caviness, 1988; Molnár et al., 1998b;
Hevner et al., 2001). Moreover, we saw no gross differences in brain
morphology between WT and SNAP-25-deficient
(Snap25/)
mice.
Our study was based on Nissl staining, immunohistochemistry, axon
tracing, and electrophysiological recording, covering the period from
E17.5 to E19. For the sake of clarity, we present the results as
follows: general phenotypic characteristics; gross histological
appearance, exhibited by Nissl staining; patterns of
immunohistochemical staining for SNAP-25, calretinin, and calbindin; outgrowth of thalamocortical fibers and their arrival at the cortex, based on anterograde labeling from DiI crystals implanted in the dorsal
thalamus; global topography of corticofugal and thalamocortical projections, examined with anterograde and retrograde tracing from
multiple dye crystal placements in the cortical convexity; and
electrophysiological recordings in slice preparations.
Phenotypic appearance of the mice
Snap25/
fetuses develop until term with few overt phenotypic abnormalities
(Washbourne et al., 2002). At E17.5-19, the Snap25 homozygote null mutant mice can be recognized by their tucked, immobile
posture and their failure to respond to physical contact (Washbourne et
al., 2002). Interestingly, the heart contracts and is able to pump
blood, but homozygotes do not survive after birth, presumably because
paralysis prevents them from breathing.
Nissl staining reveals grossly normal brain development,
except for a prominent irregularity of the cortical plate in the
minority of the mutants
Examination of cresyl violet-stained sections revealed no
striking histological abnormalities within the forebrain, brainstem, or
cerebellum of heterozygous or homozygous null mutant animals compared
with control WT littermates. Importantly there were no areas of
widespread degeneration in early developing regions of brain, including
brainstem and hypothalamus, as exhibited in Munc 18-1 mutants at later
embryonic stages in which both spontaneous AP-independent and evoked
AP-dependent neurotransmitter release are totally eliminated (Verhage
et al., 2000). We paid particular attention to morphology, cellular
structure, and lamination patterns in the neocortex (Fig.
1). By E17.5, in both normal and
SNAP-25-deficient null mutant mice, the migration of true cortical
neurons through the lower part of the preplate and their accumulation
below the marginal zone had created sharp boundaries between the
regions of different cell density at the upper and lower limits of the thickening cortical plate (Fig. 1). The basic laminar pattern, to the
extent that it is evident at this age, seemed normal in the
SNAP-25-deficient brain.
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Figure 1.
Nissl staining reveals grossly normal
development of the telencephalon and diencephalon in
Snap25 null mutant mice. Only mild abnormalities in the
regularity of the cortical plate were observed in the null mutant.
Coronal sections (60 µm) were cut and stained with cresyl violet to
reveal major brain structures. The general morphology, cellular
distribution, and lamination patterns of WT, heterozygous, and null
mutant littermates were compared. A, E,
Coronal sections at different rostrocaudal levels of an E18.5 WT brain
(+/+). B, F, Higher-magnification views
of the boxed areas labeled b and
f in A and E,
respectively, to demonstrate the normal lamination of the cortex
(ctx) at the two levels. I, Detail of the
boxed area (i) in
F. J, Part of a coronal section from
another E18.5 WT brain, showing the patterning of cells of the
primordial corpus striatum (str) created by axon bundles
of the primitive internal capsule. C, G,
Coronal sections at different rostrocaudal levels in the
Snap25 null mutant mouse ( /) at E18.5.
D, H, High magnification of the areas
boxed in C and G,
respectively. Note the abnormal lamination of the cortical plate,
particularly in the upper layers and marginal zone (mz),
with distinct peaks (arrowheads) and troughs
(arrows) along the upper margin of layer 2. K, Detail of the box labeled k
in H. Note the abnormal undulation of the upper layers
of the cortical plate (arrowheads and
arrows). L, View of the cellular
patterning created by axonal bundles in the striatum in the null
mutant, comparable with that seen in the WT (J).
Scale bars: A, C, E,
G, 800 µm; B, D,
F, J, H, L,
200 µm; I, K, 100 µm.
cp, Cortical plate; hp, hippocampus;
cc, corpus callosum; se, septal eminence;
pa, pallidum; vz, ventricular zone;
iz, intermediate zone; sp, subplate;
wm, white matter.
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Deficit in cortical plate development
The only clearly aberrant feature observed in approximately half
of the mutant brains was a curious tangential undulation of the
neocortical plate, especially along its upper border with the marginal
zone. Although the pial surface was smooth, there were irregular bulges
along the top of the cortical plate (Fig. 1D,H,K), with
reciprocal variations in thickness of the marginal zone. We detected
these undulations in 5 of 11 homozygote mutant brains at
E18.5, but in none of the heterozygote (five of five) or wild-type
(four of four) animals. Of the five mutants exhibiting undulation, two
appeared markedly more severe (Fig. 1C,D). These brains also showed significant reduction in
calbindin-immunoreactive neurons in neocortex. In the two most extreme
cases (Fig. 1D), the undulating pattern extended
through the entire thickness of the cortical plate, although it was
always more exaggerated along the upper border. The least affected
mutants were almost normal in appearance (Figs.
2C,
3C,F,I).
Moreover, the overall thickness of the cerebral wall and the cortical
plate, measured at the convexity of the hemisphere at the
mid-hippocampal level, were not affected. For three mutants, one with
severe undulations, one moderate, and one without undulations, the
measurements were as follows: thickness of cerebral wall 598 ± 25 µm SD; cortical plate 214.6 ± 13 SD. For three HT animals, the
values were as follows: cerebral wall: 598 ± 44 µm SD; cortical
plate: 221 ± 13.5 µm SD. To compare the levels of
cellular proliferation, KO and HT brains were stained with an antibody
against phosphorylated histone-H3, which labels cells in
metaphase. Similar numbers of immunopositive cells were seen (HT, 21.9 per section; KO, 23.4 per section; n = 2 each genotype; 6-10 sections were counted for each animal), which is consistent with
the similarity in cortical thickness.
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Figure 2.
Expression of SNAP-25
(A-F) and L1, marker of early cortical
connectivity, is revealed by immunohistochemistry in WT (+/+), HT
(+/), and KO (/) brains. A, SNAP-25
immunoreactivity was observed in fiber bundles extending through the
corpus striatum and intermediate zone and in the upper segment of the
corpus callosum in the WT. D, An enlarged view of
box d in A. Arrows
indicate labeled fiber bundles in the intermediate zone
(iz). B, E, Similar,
although less intensive, labeling in the HT brain. C,
F, Complete lack of immunoreactivity in these regions in
KO littermates. G, H, L1 immunoreactivity
is observed in the intermediate zone, striatum, and internal capsule in
an E18.5 WT brain. Labeled axon fascicles crossed the striatum
(arrows) and turned to the intermediate zone.
I, J, No detectable differences were
observed in the KO brain. ctx, Cerebral cortex;
hp, hippocampus; vz, ventricular zone;
iz, intermediate zone. Scale bars:
A-C, 200 µm;
D-F, 100 µm; G,
I, 500 µm; H, J, 200 µm.
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Figure 3.
Expression of calretinin
(CR) (A-F), reelin
(Reln) (G-L), and
calbindin (CB) (M-R)
revealed by immunohistochemistry in WT (+/+, left
column), HT (+/, middle column), and KO brains
(/, right column).
A-C, Calretinin (CR) is
expressed in some cells of the marginal zone (mz),
cortical plate (cp), and hippocampus.
D-F, Higher-power views of the
boxed regions in A-C,
respectively, showed no obvious differences in density of calretinin
cells. Calretinin immunostaining revealed the undulations at the base
of the marginal zone of the null mutant (F,
arrows and arrowheads).
G-L, Reelin immunostain revealed equal
numbers of Cajal-Retzius cells in the cortical marginal zone of the WT
and KO (compare G, H with
J, K). Similar density of reelin
immunoreactive cells was also detected in the hippocampal fissure
(hf) of the hippocampus (compare
I, L).
M-O, Calbindin (CB) is
expressed in several different regions of the telencephalon, including
cells of the striatum (str), cortex
(ctx), and hippocampus (hp). The general
pattern was qualitatively similar in WT
(M), HT (N), and KO
(O) brains. P-R,
Higher-power views of the boxed regions in
M-O showing that the density of
calbindin-immunoreactive cells is substantially reduced in the KO
cortex. wm, White matter; vz, ventricular
zone; lv, lateral ventricle; hf,
hippocampal fissure. Scale bars: A-C,
M-O, 300 µm;
D-F, I, L,
P-R, 100 µm; G,
J, 500 µm; H, K, 50 µm.
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Immunohistochemical analysis of cortical neuron populations
SNAP-25 expression
In the normal adult mouse brain, expression of SNAP-25, which is
transported by fast axonal transport (Loewy et al., 1991; Hess
et al., 1992), is limited primarily to presynaptic terminals. However,
during development, SNAP-25 also accumulates in axons (Catsicas et al.,
1991; Oyler et al., 1991). To document the expression pattern of
SNAP-25 at these early stages of development, we performed immunohistochemistry with a polyclonal antibody anti-SNAP-25 protein in
WT, HT, and homozygote null mutant E18.5 embryos. In both WT and HT
brains, immunoreactivity was present but generally low, especially in
the HT. Within the telencephalon, the strongest labeling was found in
fibers in the intermediate zone, at the junction of the cortex and
striatum (Fig.
2A,B,D,E),
which, judged from their position, could correspond to thalamocortical
axons. As expected, there was no immunoreactivity for SNAP-25 in the homozygote null mutant brain (Fig. 2C,F).
In contrast, immunohistochemical staining for L1, a cell adhesion
molecule thought to be expressed specifically on thalamocortical and
other diencephalic axons (Godfraind et al., 1988; Fukuda et al.,
1997), demonstrated a normal pattern of labeling in the mutant (Fig.
2G-J).
Expression of calretinin, reelin, and calbindin
To investigate further for any abnormalities of cortical
development, we performed immunohistochemistry for two
Ca2+ binding proteins, calretinin and
calbindin, which label different, types of interneurons (Hendry and
Jones, 1991) as well as early populations of migratory neurons
(Parnavelas, 2000). Calretinin is expressed in some cells of the
marginal zone (Meyer et al., 1998), and reelin is expressed
specifically in Cajal-Retzius cells known to be responsible for the
early development of cortical lamination (Ogawa et al., 1995).
Calbindin is a useful marker for the majority of tangentially migrating
neurons (Parnavelas, 2000).
Similar patterns of immunoreactivity for calretinin were observed in
WT, HT, and homozygote null mutant brains at E18.5 in all telencephalic
areas (Fig. 3A-F), with no obvious
difference in the densities of calretinin cells (WT/HT, 20.9 per
section ± 0.5 SD; KO, 19.8 per section ± 0.4 SD).
Similarly, there were no significant differences in the numbers of
reelin immunoreactive cells (Fig. 3, compare G, H
with J, K) (WT/HT, 39.2 per section ± 9.6 SD; KO, 27.3 ± 8.6 SD), indicating that the generation of this early cell population was unperturbed. However, in the two null
mutants that were strikingly affected morphologically by neocortical
undulation, far fewer calbindin-immunoreactive cells were seen in the
cortex (Fig. 3M-R), suggesting a selective
reduction of tangentially migrating cells. Quantitative analysis
comparing these severely affected animals with heterozygote controls in the marginal zone and cortical plate compartments showed that this
reduction in fact was significant (marginal zone: KO, 10.5 cells per
section ± 0.1 SD compared with HT, 22.6 cells per section ± 1.2 SD, p < 0.001; cortical plate: KO, 14.5 cells per
section ± 4.9 SD compared with HT, 50.5 cells per section ± 3.7 SD, p < 0.003).
Outgrowth of thalamocortical projections
The total gestation period of mice is 19-20 d, compared with
21-22 d in the rat. Hence, although the development of the
thalamocortical fiber pathway in the normal mouse is essentially
similar to that in rat, each step occurs ~1-2 d earlier
(Molnár et al., 1998a,b). We examined the fine pattern of
thalamic axon innervations in brains from normal and null mutant E17.5
and E18.5-E19 embryos from three litters (14 individuals altogether)
with carbocyanine dye tracing. Counterstaining with bisbenzimide
revealed major anatomical features, such as the pial surface of the
cortex, the junction of layers 1 and 2, and the gray matter-white
matter boundaries. This, together with the L1 immunostaining,
provided further evidence for the grossly normal anatomical appearance
of the null mutant brain.
We placed a DiI carbocyanine dye crystal in the dorsolateral aspect of
the diencephalon (posterior dorsal thalamus) of both hemispheres to
label thalamocortical axons. In WT, HT, and Snap25 null
mutant specimens, labeled thalamocortical fibers were present, and
their outgrowth had progressed to the same extent, appropriate for the
developmental stage. At both E17.5 (Fig.
4) and E18.5-E19 (Fig.
5), the appearance of afferent axons
from the thalamus was essentially indistinguishable among KO, HT, and
WT mice. Individual thalamic axons formed an impressively ordered,
parallel array throughout their course within the telencephalon. As
they passed through the anlage of the corpus striatum, they were
organized in parallel, fasciculated bundles, with small groups of axons occasionally crossing from one bundle to another. The fibers
defasciculated and turned at the striatocortical boundary, forming a
thick band that entered the intermediate zone, running tangentially,
without dispersion, toward and into the subplate layer. The axons did not appear to cross each other extensively at any point along their
course.
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Figure 4.
Outgrowth of thalamocortical projections
revealed with DiI tracing from the dorsal thalamus in WT
(A, D, G), HT
(B, E, H), and KO
(C, F, I) brains at
E17.5. Coronal sections were photographed with two different filters to
reveal the DiI label (red) and bisbenzimide counterstain
(blue). A-C, In all three
genotypes, thalamic axons traversed the primitive internal capsule as
an organized array of fiber bundles and then defasciculated and turned
dorsally to run through the intermediate zone and into the subplate
below the cortex (ctx). Within a single litter,
individuals showed slight variation in their maturity, but there was no
consistent difference among WT, HT, and Snap25 KO brains
in the state of advancement of the thalamocortical fibers.
D-F, Higher-power views of the
boxes in A-C, showing the
indistinguishable patterns of ingrowth of thalamic axons into the
cortical plate (cp). Thalamic fibers could also be seen
extending through the lower intermediate zone (liz) and
into the subplate (sp) layer below the cortical plate.
At this stage, axons did not substantially invade the cortical plate:
the radial ingrowth of thalamocortical fibers was limited to a few side
branches arising in the intermediate zone and subplate.
G-I, Confocal microscopic
reconstructions revealed side branches of similar form and extent in
all three genotypes (arrows). These small side branches
of thalamic axons penetrated only the lowest part of the cortical
plate. Bisbenzimide counterstaining (blue in
A-F) showed major anatomical
features, such as the pial surface of the cortex, layer 1, and the gray
matter-white matter boundaries. In this particular null mutant brain
(C, F), there were no obvious
undulations in the cortical plate. Scale bar:
A-C, 200 µm;
D-F, 100 µm;
G-I, 50 µm.
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Figure 5.
Thalamocortical projections, traced with DiI, show
similar ingrowth patterns in WT (A, C,
E) and KO (B, D,
F) brains at E18.5. A-D,
Double-exposure photomicrographs of coronal sections showing the
DiI-labeled axons (red) and bisbenzimide counterstaining
(blue) in the left hemisphere. Thalamic axons exhibited
similar fasciculation patterns in the internal capsule
(ic) and cortex (ctx) of WT (+/+) and KO
(/) brains. In both genotypes, axons had started quite
substantially to invade the cortical plate (C,
D, arrows). E,
F, Confocal microscopic reconstructions revealed similar
patterns of invasion in the two genotypes. Thalamic projections
extended up to the upper third of the cortical plate and began to show
branch formation (arrows). Scale bars: A,
B, 300 µm; C, D, 200 µm; E, F, 100 µm. dt,
Dorsal thalamus; sp, subplate; liz, lower
intermediate zone; mz, marginal zone.
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At E17.5 (Fig. 4), thalamic axons had traversed the primitive internal
capsule and had accumulated within the subplate layer. In the null
mutant mice, just as in normals, there was very limited radial ingrowth
of thalamocortical fibers into the cortical plate, perhaps because the
cortical plate itself is relatively nonpermissive to ingrowth at this
age (Götz et al., 1992; Molnár and Blakemore, 1995; Tuttle
et al., 1995). Confocal microscopic reconstructions revealed a few
short side-branches, with growth cones at their tips, penetrating the
lowest part of the plate in the null mutant (Fig.
4I), just as in normal mice at this stage (Catalano
et al., 1996; Molnár et al. 1998a,b). As described previously
(Molnár et al., 1998b), we also saw a small number of axons, some
even forming tiny fasciculated bundles, running up obliquely through the entire cortical plate and entering the marginal zone.
At E18.5-E19 in WT, HT, and KO brains, there was substantially more
invasion of the cortical plate by thalamic axons and their side
branches (Fig. 5), some extending radially through virtually the entire
thickness of the plate.
Topography of embryonic corticofugal and
thalamocortical connectivity
We used multiple dye placements, in parasagittal or coronal rows
across the cortical hemisphere at E18.5, to examine the global topography of fiber pathways (Molnár et al., 1998a,b), and we documented the trajectory and distribution of the labeled fiber bundles
and backlabeled thalamic cell groups. Each crystal labeled a group of
axons, forming a discrete, fairly tight bundle, and the spatial
relationships of the separate labeled bundles was maintained throughout
their path (Fig.
6G-J). The
labeled axons followed straight or slightly curved trajectories as they
formed a fan-shaped array, converging on the primitive internal
capsule, always maintaining topography correlated with the spatial
separation of their sites of origin. As seen in Figure 6, the bundles
of axons labeled by the different dyes stayed separate from each other,
even as they passed through the constriction of the primitive internal
capsule, and they could be traced back within the diencephalon to
separate groups of backlabeled thalamic cells (Fig.
6A-F).
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Figure 6.
Tracing with DiI and DiA from multiple cortical
crystal placement sites revealed the normal topography of
thalamocortical and corticofugal projections at E18.5 in both WT
(A, C, D,
G, I) and Snap25
null mutant (B, E, F,
H, J). Two crystals of DiI
(red) were implanted in a parasagittal row in the cortex
(ctx) of the left hemisphere of WT
(A) and KO (B) brains, and
one crystal of DiA (green) was placed midway
between the DiI placements (shown schematically in top
left). Horizontal sections were counterstained with
bisbenzimide. Triple-exposure pictures were taken on a fluorescence
microscope (A, B, G,
H) or a confocal microscope
(C-F, I,
J). The trajectory and distribution of the
labeled fiber bundles and backlabeled thalamic cell groups were
documented at different horizontal levels. A,
B, Low-power images show the fiber bundles in both the
cortex (top right) and the dorsal thalamus
(dt) (bottom left) in horizontal sections
corresponding to the red box in the schematic diagram
above (rostral is to the right). Each crystal labeled a
discrete group of axons (a, a' and
c, c' labeled with DiI; b,
b' labeled with DiA). The spatial arrangement of the
separate labeled bundles was maintained throughout their path, with a
90° rotation of the array between telencephalon and diencephalon.
C, Higher-power confocal image, corresponding to the
box in A. D, Detailed
images of the box in C, showing discrete
groups of labeled thalamic cells at the tip of the axons
(arrows mark individual cells). E,
F, Confocal images, comparable in position and scale
with C and D, respectively, from other
similarly treated null mutant brains. G,
H, Low-power views of more ventral sections. The bundles
labeled by the different dyes stayed separate from each other, even as
they passed through the constriction of the primitive internal capsule
(ic). I, Higher-power confocal image of
the box in G, showing fiber bundles at
the level of the primitive internal capsule. J,
Higher-power image of fiber bundles passing through the internal
capsule from a null mutant. Scale bars: A,
B, G, H, 200 µm;
I, J, 100 µm; shown on
C: C, E, 100 µm;
D, 40 µm. hp, Hippocampus.
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A crystal placed in the cortex at E18.5 labels not only thalamocortical
axons but also corticothalamic and other corticofugal projections. In
Figure 7, corticothalamic fibers can be
seen separating from the other descending corticofugal projections,
which deviate ventrally on their way toward the cerebral peduncle. The
segregation of staining by the different dyes along the entire extent
of the fiber pathways provides clear evidence that all fiber
projections to and from the cortex maintain their initial order and
relationships along their entire course through the telencephalon and
diencephalon.
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Figure 7.
Topography of thalamic and corticofugal
connectivity at E18.5. A parasagittal row of crystals (DiI, DiA, DiI)
was implanted in the left hemisphere (schematic illustration,
top left) of WT (A, C) and
Snap25 KO (B, D) brains.
At this age, each crystal labeled both thalamocortical axons
(retrogradely) and corticofugal fibers (anterogradely).
A, B, Confocal micrographs at the level
of the primitive internal capsule, as indicated by the red
box in the schematic coronal section above (dorsal is
up). As the mixed array of corticofugal fibers enters
the diencephalon (A, B,
right), the descending corticofugal axons diverge from
the bundle of corticothalamic and thalamocortical fibers and turn
toward the cerebral peduncle (ped). Axons stained
from the three crystal placements are labeled a',
b' (DiA), and c' (DiI). C,
D, At a more caudal level, the labeled bundles entering
the cerebral peduncle have separated from those entering the dorsal
thalamus (dt). Scale bars:
A-D, 100 µm.
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Electrophysiology
We used electrophysiological methods to confirm that thalamic
axons were not only present in the cortical plate of the mutant mice
but were capable of conducting action potentials. In E18.5 whole
forebrain slices, we stimulated the VB complex of the thalamus and
recorded in the putative somatosensory cortex. These records provided
clear evidence for transmission of APs along thalamocortical axons in
both null mutant and normal mouse brains. Figure
8 shows representative examples of field
potentials recorded in the cortex of an HT mouse (in which synaptic
transmission is intact) (Washbourne et al., 2002) and a homozygous null
mutant mouse. The responses are indistinguishable except for a
difference in the form of the stimulus artifact, which was caused
simply by a difference in the stimulating polarity. Essentially
identical results were obtained in slices from a total of two WT, three
HT, and four KO animals. In every case, we were able to record field
potentials within the cortical plate after thalamic stimulation,
regardless of the stimulation charge polarity. In two of the HT and two
of the homozygote mutant slices, we applied 40 µM CNQX,
10 µM MK801, and 20 µM bicuculline to block
glutamatergic and GABAA synaptic transmission. In
each instance, as in the examples in Figure 8, the field potential was
not obviously altered and therefore was caused presumably entirely by
presynaptic activity. In one WT, two HT, and three homozygote mutant
slices, after recording a stable field potential, we applied 600 nM TTX to block Na+
channel-dependent conduction. In every instance, as in the examples in
Figure 8, the field potential was eliminated. These results indicate
that at this stage of development, thalamic axons are capable of AP
propagation, both in null mutant and normal brains, but they have yet
to establish detectable, functional synaptic connections within the
cortical plate, even in WT mice. Previously, Higashi et al. (2002)
showed, using optical recordings with voltage-sensitive dyes, that
direct thalamic stimulation can elicit sustained postsynaptic depolarizations in embryonic thalamocortical slices; however, current
source density analyses, like our field potential recordings, were
unable unequivocally to demonstrate glutamate receptor- mediated postsynaptic responses (Molnár et al., 2002). This suggests that fields generated by extracellular current flow from synaptic potentials are likely to be small and less synchronous and therefore not sufficient to demonstrate the few synaptic connects that may be made at
this time.
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Figure 8.
Electrophysiological experiments using a whole
forebrain slice preparation at E18.5 (Higashi et al., 2002) revealed
that thalamic axons conduct APs, but there is no obvious synaptic
transmission onto neurons of the cortical plate in WT, HT, or KO
brains. A, Diagram to illustrate the plane of vertical
section (45° to both coronal and sagittal planes) at which brains
were cut at 400 µm to produce slices containing the VB complex of the
thalamus, the putative somatosensory cortex, and the entire fiber
pathway in between. B, Camera lucida tracing of a
thalamocortical slice preparation showing the position of the
stimulating electrode in the thalamus (TH).
CP, Cortical plate; SP, subplate;
IZ, intermediate zone; VZ, ventricular
zone; IC, internal capsule. C,
Extracellular recordings after thalamic stimulation in slices from HT
(+/, left column) and KO (/, right
column) brains. In both genotypes, stimulation of the VB
produced an initial artifact (the polarity of which depended on the
stimulating polarity), followed by a negative-going field potential
with a peak at ~4 msec. This peak was not eliminated by applying
ionotropic blockers (40 µM CNQX, 10 µM
MK801, 20 µM bicuculline) in the bath. The field
potential was eliminated, however, by the subsequent application of 600 nM TTX. The responses are indistinguishable except for a
difference in the form of the stimulus artifact.
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DISCUSSION |
Evoked neurotransmitter release at chemical synapses is mediated
by the SNARE core complex, of which SNAP-25 is an essential component.
Null mutation of the Snap25 gene results in a complete loss
of stimulus-dependent synaptic transmission while sparing the
exocytotic mechanisms that support AP-independent spontaneous neurosecretion (Washbourne et al., 2002). Hence, the Snap25
null mutant allowed us to examine the extent to which the mouse
forebrain develops in the absence of evoked neurotransmitter release.
Abolition of evoked neurotransmission eliminates neuromuscular
function, hence preventing diaphragm contraction and respiration. This
probably accounts for the perinatal morbidity of SNAP-25-deficient
fetuses (although other more widespread deficits caused by the lack of regulated secretion might contribute). Our study was limited, then, to
prenatal development.
The lack of overt pathology in the formation of most tissues suggests
that SNAP-25-deficient fetuses develop fairly normally. The
morphological appearance of the null mutant is clearly distinct, however, from that of normal fetuses at E17.5-E19, which may be attributable primarily to abnormal neuromuscular development. Importantly, as shown previously (Washbourne et al., 2002) and demonstrated more extensively here, the loss of evoked synaptic activity does not prevent the grossly normal formation of the brain. As
evidenced by Nissl staining, cellular proliferation, migration,
differentiation, lamination, and neuronal survival seem mostly normal
in the Snap25 null mutant. In particular, the spatial
distribution and structural aspects of neurons appear unaltered, and
the main features of the diencephalon and telencephalon develop
normally. Moreover, we find no evidence of widespread neurodegeneration
in the absence of evoked neurotransmission. This contrasts with the
significant cell death found in the nsec-1/munc18-1 mutant,
which lacks both evoked and AP-independent neurotransmitter release
(Verhage et al., 2000). This strongly suggests that
stimulus-independent secretion of transmitters or growth factors is
sufficient to validate and maintain neuronal populations during development.
The only deficits that we saw in the KO mouse, at the light-microscopic
level, were the relative reduction in density of calbindin-positive neurons (but not of calretinin-positive cells) in the cortex and the
strange undulations of the cortical plate in some Snap25
null mutants.
Analysis of thalamocortical fiber projections with carbocyanine dyes
revealed normal growth kinetics, fasciculation patterns, and topography
in the E17.5-E19 null mutant brain. Thalamocortical axons reach the
cortex by their normal route, accumulate below the cortical plate, and
then begin to form side branches in the subplate and the deep cortical
plate, just as in the wild-type brain. We detected no abnormality in
the pattern or timing of their entry into the cortical plate in the
null mutant. Thus, thalamocortical axonal projection and the early
steps necessary for proper targeting proceed without evoked
neurotransmitter release. It is interesting to compare our results with
those of Harris (1984), who found roughly normal retinotopic topography
in the tectum of axolotls, which, before retinal axon outgrowth, had been parabiotically joined to TTX-secreting Californian newts. Hence
retinal axons in amphibians are also capable of finding the target in
the absence of impulse activity, although they often took unusual routes.
The apparently normal growth of thalamocortical fibers might be thought
to contradict evidence that inhibition of SNAP-25 by antisense
oligonucleotides or clostridial neurotoxins causes growth cone collapse
and inhibition of axonal growth, in vitro and in
vivo (Osen-Sand et al., 1993, 1996). However, such inhibition of
SNAP-25 expression in the chick retina affects primarily the length and
terminal differentiation of outgrowing amacrine cell processes and not
the direction of their growth or the general morphology of the neurons
(Osen-Sand et al., 1993). This may reflect functions of SNAP-25 at late
stages of elongation when it appears to be expressed mainly in axonal
growth cones (Oyler et al., 1989, 1991, Osen-Sand et al., 1993).
SNAP-25-deficient motoneurons are certainly able to successfully
innervate neuromuscular targets and generate AP-independent miniature
endplate potentials, and spontaneous postsynaptic activity can be
recorded from cortical slices and cultured hippocampal neurons. These
results demonstrate that, in general, SNAP-25 and hence evoked
neurotransmission are not required to assemble the cellular processes
for the initial stages in synaptogenesis (Washbourne et al., 2002). The
ability of SNAP-25-deficient neurons to locate their targets and
establish initial contacts, in the absence of evoked synaptic activity, provides an important insight into decision-making mechanisms during
the development of connectivity. The results also provide further
evidence that the membrane addition necessary for axonal extension
occurs independent of the neurosecretory machinery (Leoni et al., 1999;
Schoch et al., 2002; Washbourne et al., 2002).
The observation, in many mammals, that axons from the thalamus
temporarily pause or halt when they reach the cortical subplate, before
substantially invading the cortical plate, has led to the idea that
some recognition process, critical for future ingrowth, occurs during
this "waiting period" (Rakic, 1976; Shatz et al., 1990). Catalano
and Shatz (1998) have demonstrated that blockade of activity with TTX
in neonatal ferrets causes disruption of projections from lateral
geniculate nucleus to visual cortex (which develop relatively late in
ferrets), and they suggested that the recognition process during the
waiting period requires impulse activity. However, we found that in the
Snap25 null mutant mouse, thalamic axons not only elongate,
with proper spatiotemporal relationships, but also arrive, pause in the
subplate, and invade the cortical plate quite normally. This strongly
supports the probability that the formation of thalamocortical
projections and the initial ingrowth of thalamic axons to the
appropriate cortical region do not rely on AP-dependent synaptic
transmission, within the subplate or elsewhere.
It is conceivable that the results of Catalano and Shatz (1998)
represent a distinct effect of TTX beyond the elimination of APs:
possibly the impairment of postsynaptic signaling of subplate neurons
in response to spontaneous transmitter release from thalamic axons. On
the other hand, the apparent difference in results might be explained
simply by the fact that we were able to examine only the initial stages
of the targeting process. Because we were limited to observing prenatal
brain development, before the formation of functional connections in
layer 4, we cannot resolve whether synaptic transmission is needed for
thalamocortical axons to innervate their appropriate target neurons
precisely. Indeed, a host of evidence suggests that AP-dependent
neurotransmitter release is important for the refinement of topography
that occurs when these projections reach their final targets (Stryker
and Harris, 1986), as well as for the activity-dependent
differentiation of cortical architecture (Hannan et al.,
2001).
Our electrophysiological observations show that the thalamic axons
themselves are functional, in that they are able to transmit sodium
channel-dependent, TTX-sensitive APs. The similarity of field
potentials in the cortex of normal and null mutant mice also supports
the conclusion that thalamic fibers are similarly distributed within
the plate. Because glutamate receptor blockers produced no change in
cortical field potentials in any genotype, we conclude that, at E18.5,
studied with extracellular recording, there is no detectable synaptic
transmission within the cortical plate of the mouse. This issue will
need to be addressed with intracellular and optical recordings (Higashi
et al., 2002).
SNAP-25-deficient neurons are able to generate spontaneous
AP-independent neurotransmitter release (Washbourne et al., 2002). This
alone may be sufficient to mediate intercellular recognition processes
within the subplate or the cortical plate or along the route from the
thalamus. Although synaptic contacts formed in the null mutant are
unable to evoke postsynaptic APs, they may be able to communicate
sufficiently via constitutive secretion of transmitter or possibly neurotrophins.
Recent evidence suggests that the Slit family of chemorepellant
proteins plays a major role in axonal guidance of central fiber
pathways, including thalamocortical projections (Bagri et al., 2002).
Our findings (in the SNAP-25-deficient mutant) are consistent with the
notion that the secretion of Slit proteins to form repulsive gradients,
which act in conjunction with attractive cues and dictate the
trajectory of axon pathways, does not require evoked synaptic activity
but is mediated via constitutive, AP-independent mechanisms.
Further work is needed to determine whether the reduction of
calbindin-expressing neurons, which coincides with the most severe morphological deficits (undulations) in the upper cortical plate of
roughly 20% of the mutants, is caused directly by the absence of evoked transmitter release, as well as the identity of any transmitters involved. It is known that migrating cortical neurons express functional receptors (Métin et al., 2000;
López-Bendito et al., 2002), and blockade of these receptors
alters cortical migration (Behar et al., 2001). Accordingly,
corticofugal projections might provide presynaptic AP-dependent
signaling as they make intimate contact with these migrating neurons
(Métin et al., 2000; Denaxa et al., 2001). It is possible, then,
that the absence of AP-mediated transmitter release interferes somewhat
with the radial migration of excitatory cortical neurons (hence
generating the irregular lamination) as well as with the tangential
migration of calbindin cells.
Neurotransmitter secretion along growing nerve processes is clearly
evident in developing neurons (Zakharenko et al., 1999). During
development there may be a gradual transition from mechanisms dependent
only on constitutive secretion, mediated by exocytotic machinery that
does not require the SNARE complex, to later processes in which
regulated transmitter release controls the formation of mature synapses
and modulates functional synaptic plasticity.
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FOOTNOTES |
Received April 3, 2002; revised Sept. 16, 2002; accepted Sept. 23, 2002.
*
Z.M. and G.L.-B. contributed equally to this work.
This work was supported by the National Institutes of Health
(Grant MH 48989; M.C.W.), the Oxford McDonnell Centre for Cognitive Neuroscience (North American Network Grant; Z.M., M.C.W.), the Swiss
National Science Foundation (Grant 3100-56032.98; Z.M.), the European
Community (Grant QLRT-1999-30158; Z.M.), and The Wellcome Trust (Grant
063974/B/01/Z; Z.M.). We are very grateful to Rosalind Carney and
Courtney Voelker for thoughtful comments on this manuscript, James R. Mathews and Lynden Guiver for excellent technical support, and Sayuri
Nixon for genotyping. We also thank Dr. Nathalie Garin-Schwaller for
help with Figure 3G-I. Inquiries about Snap25
null mutant mice should be addressed to M.C.W.
Correspondence should be addressed to Zoltán Molnár,
Department of Human Anatomy and Genetics, University of Oxford, South Parks Road, Oxford, OX1 3QX, UK. E-mail:
zoltan.molnar{at}anat.ox.ac.uk.
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