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The Journal of Neuroscience, June 1, 1999, 19(11):4449-4461
Glutamate Acting at NMDA Receptors Stimulates Embryonic Cortical
Neuronal Migration
Toby N.
Behar1, 2,
Catherine A.
Scott1,
Carolyn L.
Greene1,
Xiling
Wen1,
Susan V.
Smith1,
Dragan
Maric1,
Qi-Ying
Liu1,
Carol A.
Colton2, and
Jeffery L.
Barker1
1 Laboratory of Neurophysiology, National Institute of
Neurological Disorders and Stroke, National Institutes of Health,
Bethesda, Maryland 20892, and 2 Department of Physiology
and Biophysics, Georgetown University School of Medicine, Georgetown
University, Washington DC 20007
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ABSTRACT |
During cortical development, embryonic neurons migrate from
germinal zones near the ventricle into the cortical plate, where they
organize into layers. Mechanisms that direct neuronal migration may
include molecules that act as chemoattractants. In rats, GABA, which
localizes near the target destination for migrating cortical neurons,
stimulates embryonic neuronal migration in vitro. In mice, glutamate is highly localized near the target destinations for
migrating cortical neurons. Glutamate-induced migration of murine
embryonic cortical cells was evaluated in cell dissociates and cortical
slice cultures. In dissociates, the chemotropic effects of glutamate
were 10-fold greater than the effects of GABA, demonstrating that for
murine cortical cells, glutamate is a more potent chemoattractant than
GABA. Thus, cortical chemoattractants appear to differ between species.
Micromolar glutamate stimulated neuronal chemotaxis that was mimicked
by µM NMDA but not by other ionotropic glutamate receptor
agonists (AMPA, kainate, quisqualate). Responding cells were primarily
derived from immature cortical regions [ventricular zone
(vz)/subventricular zone (svz)]. Bromodeoxyuridine (BrdU) pulse
labeling of cortical slices cultured in NMDA antagonists (µM MK801 or APV) revealed that antagonist exposure
blocked the migration of BrdU-positive cells from the vz/svz
into the cortical plate. PCR confirmed the presence of NMDA receptor
expression in vz/svz cells, whereas electrophysiology and
Ca2+ imaging demonstrated that vz/svz cells
exhibited physiological responses to NMDA. These studies indicate that,
in mice, glutamate may serve as a chemoattractant for neurons in the
developing cortex, signaling cells to migrate into the cortical plate
via NMDA receptor activation.
Key words:
mouse; chemotaxis; cortex; development; slice; chemoattractant
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INTRODUCTION |
During CNS development,
cortical neurons are generated in specialized germinal zones adjacent
to the ventricle, termed the neuroepithelium. After terminal mitosis,
neurons migrate from the neuroepithelium through the cell-sparse
intermediate zone (iz) into the cortical plate (cp). After entering the
cp, neuroblasts migrate through the established neuronal lamina and
settle onto the outermost layer, forming an "inside-out" gradient
of maturation, a process which is essential to cortical neuronal
lamination (Jacobson, 1991 ).
As neurons travel from the germinal zones, many of them move along the
fibers of radial glial cells (Rakic, 1972, 1990; Hatten, 1990), which
extend from the mitotic zone to the pial surface. Some neurons also
migrate along alternative pathways. Neurons in embryonic slice
preparations have been observed to move orthogonally, along nonradial
pathways that are perpendicular to the axis of glial processes
(O'Rourke et al., 1992, 1995). This suggests that, during development,
mechanisms other than direct contact with radial glial fibers may also
influence nerve cell movement and thus contribute to the structural
creation of the cortex.
Extracellular factors may provide directional cues to migrating
neuroblasts. In the developing spinal cord, embryonic motoneurons transplanted into the dorsal half of the cord migrate back to the
ventral horns, suggesting that locally mediated factors in the target
region provide positional cues to the cells (Eisen, 1991). The
positional cues may result from gradients of diffusible molecules,
which act as chemoattractants to direct the cells toward their target
destinations. Molecules present in the microenvironment of migrating
nerve cells in vivo modulate the movement of the cells
in vitro. Among these, platelet-derived growth factor,
brain-derived neurotrophic factor, and GABA promote movement of
embryonic cells acutely dissociated from the developing rat CNS
(Armstrong et al., 1990; Behar et al., 1994, 1996). Komuro and Rakic
(1993) reported that granule cell migration in postnatal rat cerebellar slice preparations was inhibited by antagonists of ionotropic glutamate
receptors, suggesting that glutamate provides motility signals to
migratory neurons. Glutamate is also a likely candidate for a
chemoattractant in the developing mouse cerebral cortex. The
Cajal-Retzius (C-R) cells, which are generated first and are located in
layer 1 of the cp, contain glutamate during the final week of gestation
(del Rio et al., 1995), when neurons are actively migrating into this
region (Jacobson, 1991). Thus, glutamate is found in an appropriate
location to serve as a chemoattractant.
We used an in vitro chemotaxis assay (Falk et al., 1980) to
examine whether glutamate influences embryonic cortical nerve cell
movement. The assay was used to measure the effects of glutamate on the
migration of acutely dissociated murine embryonic cortical cells. In
addition, we examined the effects of glutamate on neuronal migration
using organotypic slice preparations. Modulation of cortical neuroblast
migration with glutamate receptor agonists and antagonists suggests
that receptors for NMDA (NMDA-Rs) mediate the motility signals of
glutamate in a subset of cortical neurons.
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MATERIALS AND METHODS |
Dissection and dissociation of the cells
Cortical cells from C57BL/6 pups at embryonic days 13-18
(E13-E18) were used in these experiments. Dams were killed with
CO2, embryos were removed by cesarean section, and
embryonic age was determined by measurement of crown-rump length
(Schambra et al., 1992). Brains from littermates were removed and
placed into Earle's balanced salt solution (EBSS) (Life
Technologies, Grand Island, NY). After removing the meninges,
the cerebral cortices were dissected, minced, and dissociated into
single cell suspensions using a papain digest described previously
(Behar et al., 1994). Dissociated cells were resuspended in EBSS
supplemented with 16 mM glucose at a concentration of
106 cells/ml for the migration studies. For
quantitative analysis using immunocytochemistry, 2.5 × 105 cells were seeded onto 35 mm Nunc (Naperville,
IL) culture dishes precoated with poly-D-lysine
[>450,000 molecular weight (MW), 20 µg/ml] (Collaborative
Research, Bedford, MA) and allowed to adhere onto the dishes for 1 hr
at 37°C.
Preparation of ventricular zone and cp cell suspensions.
Detailed description of the separation of the two brain regions was described previously (Behar et al., 1998). Briefly, brains from littermates were removed. A McIlwaine tissue chopper (Brinkman, Westbury, NY) was used to prepare 350 µm coronal slices of the telencephalon. Slices were transferred to a dish and were
microdissected through the cortical intermediate zone, which divided
the cortex into two tissue segments designated as the cp and the
ventricular zone (vz). Tissue designated "cp" included the cp,
subplate (sp), and the upper half of the iz. Tissue designated "vz"
included the vz, svz, and the lower half of the iz. Tissue segments
from each region were enzymatically dissociated into single cell
suspensions with papain, and the cells were either immunostained or
used in the microchemotaxis assay.
Quantitative analysis of TUJ1 and nestin staining in the
vz and cp dissociates. Acutely adherent cells on culture plates
(dissociated from the vz and cp preparations) were fixed 30 min in 4%
paraformaldehyde (PF) and washed in PBS, pH 7.2. Cells were
incubated at 4°C overnight in a mixture of rabbit anti-nestin
antibody (a progenitor cell marker; Tohyama et al., 1992) and the
monoclonal TUJ1 antibody (a neuronal marker; Lee et al., 1990). The
cells were washed three times in PBS and then incubated 1 hr at 21°C
in appropriate secondary antibodies [donkey anti-rabbit FITC and rat
anti-mouse tetramethylrhodamine isothiocyanate (TRITC); 1:50;
Jackson ImmunoResearch, West Grove, PA]. PBS supplemented with 0.05%
saponin and 0.1% BSA was used for the antibody diluent. Immunolabeled
cells were examined on a Zeiss (Thornwood, NY) Axiophot microscope
equipped with epifluorescence and the appropriate filters for the
coincident visualization of fluoresceine and rhodamine. The percentage
of positively stained cells from both cp and vz regions was determined
by dividing the number of fluorescently labeled cells in a field by the
total number of cells in the same field (visualized under phase
contrast using a water immersion 40× objective). Ten random fields
were counted per dish (1000 cells per dish) for duplicate samples from seven separate experiments. Data were subjected to ANOVA to
determine variation within groups.
Chemotaxis assay
Migration of the acutely dissociated cells was measured using a
microchemotaxis assay described previously (Behar et al., 1994). Before
assembly, the filter was coated with poly-D-lysine (>450,000 MW; 30 µg/ml; Collaborative Research) to promote adhesion.
Cells were incubated in the presence of attractants for 18 hr at 37°C
with 10% CO2 and 90% air. After the incubation, the chambers were disassembled, and the cells on the upper surface of the
filter that failed to migrate were scraped off. Migrated cells on the
underside of the membrane were fixed, stained, and counted as described below.
Quantification of migration. For quantitative assays,
migrated cells were fixed 30 min in 4% PF with 0.1% glutaraldehyde in 0.1 M phosphate buffer, pH 7.2, and stained 5 min in 0.1%
cresyl violet. The membranes were mounted cell side up onto 2 × 3 inch glass slides, allowed to air dry, and covered with immersion oil. Cells were counted under bright field using oil immersion 16× or 40×
Zeiss planapo objectives on a Zeiss photomicroscope.
Only cells that passed completely through the pores and migrated out
onto the underside surface of the membrane were considered migrated
(positive) cells. Spontaneous (random) migration of cells in control
wells containing buffer only was also assessed in all assays. Each
attractant condition was run in triplicate wells, and 5-10 fields of
stained cells were counted for each well. The average number of
migrated cells per square millimeter for each attractant
condition was calculated, and the statistical significance was analyzed
using ANOVA followed by Fisher's PLSD test. Illustrations of data
pertaining to the chemotaxis assay represent individual trials unless noted.
Chemotaxis (the directed migration of cells toward regions of higher
concentrations of a chemical attractant) was distinguished from
chemokinesis (stimulation of increased random cell motility) by placing
the same concentration of attractant in both the upper and lower wells
of the chamber, thereby eliminating a chemical gradient. The number of
cells migrating under these conditions was considered a measure of chemokinesis.
Characterization of migrated cells by immunocytochemistry
In some assays, migrated cells on the underside of the filter
were fixed, stained with cresyl violet, and then analyzed for expression of neurofilament (NF) using a monoclonal anti-NF tissue culture supernatant (1:8; gift of Dr. C. Gibbs, National Institute of
Neurological Diseases and Stroke, National Institutes of Health, Bethesda, MD). The cells were washed three times and incubated in
peroxidase-conjugated secondary antisera (goat anti-mouse IgM; 1:40;
Jackson ImmunoResearch) for 1 hr at room temperature. Immunoreaction product was visualized using a diaminobenzidine (DAB) substrate. Cresyl
violet staining was used to determine the number of total cells,
whereas NF was used to indicate neurons only. Cells were counted under
bright field using a 40× planapo objective.
Viability of cells in the presence of inhibitors. The
starting population of cells, suspended in EBSS with 16 mM
glucose, was seeded onto 35 mm Nunc culture dishes precoated with
poly-D-lysine (>450,000 MW, 20 µg/ml; Collaborative
Research) at 2.5 × 105 cells per dish. Cells
were allowed to adhere onto the dishes for 1 hr at 37°C and then were
fed with either control medium (EBSS with 16 mM glucose) or
EBSS with glucose containing attractant (1 µM glutamate
or NMDA) only or attractant plus inhibitor [10 µg/ml BAPTA-AM, 10 µM (+)-5-methyl-10,11-dihydro-5H-dibenzo [a,d] cyclohepten-5,10-imine maleate (MK801), or 10 µM
R( )-2-amino-5-phosphonovaleric acid (APV)]. Cells
were incubated in the presence of the ligands for 18 hr at 37°C, and
then the viability of the cells was determined using trypan blue
exclusion. In control media (EBSS plus glucose), cell viability at 18 hr was 94.3 ± 0.4%. Viability of cells maintained in 1 µM glutamate was 96.2 ± 3.3%, whereas the
viability of cells incubated in NMDA was 95.7 ± 1.2%. In the
presence of each inhibitor, the viability equaled or surpassed that of
the controls (BAPTA-AM, 98.5 ± 0.2%; MK801, 94.1 ± 2.2%;
APV, 98.7 ± 0.9%).
Culture of embryonic cortical slices
E17 brains from littermates were removed and placed into cold
(4°C) slicing medium (in mM: 120 NaCl, 5 KCl, 1.2 KH2PO4, 14 dextrose, 26 NaHCO3, and 1.24 MgSO47H2O
plus 5 mg/ml phenol red), and 300 µm coronal slices of the
telencephalon were prepared using a vibratome. Slices were transferred
onto Millipore (Bedford, MA) filters and placed into six-well Costar
(Cambridge, MA) plates containing ice-cold slicing medium. The slices
were allowed to recover for 2 hr at 4°C and were then transferred on
the filters to plates containing growth medium (Neuralbasal medium
supplemented with B27 and glutamine; Life Technologies). At the
beginning of the incubation period, the growth medium was supplemented
with 50 µg/ml of bromodeoxyuridine (BrdU). After 18 hr,
BrdU-containing medium was replaced with growth medium lacking BrdU. To
control for anatomical gradients of cortical maturation, as well as
possible differences in cortical development among embryos of the same litter, slices from one cortical hemisphere cultured under control conditions was compared with slices of the contralateral hemisphere cut
from the same embryo and maintained in media containing antagonists at
NMDA receptors (100 µM MK801 or APV). Antagonists were
added at high micromolar levels to ensure that the ligands penetrated through the thick sections. Slices were cultured at 37°C in a mixture
of 10% CO2 and 90% air, and medium was replaced every 3 d.
BrdU immunolabeling. Two or 6 d after removing the BrdU
from the cultures, the slices were fixed for 1 hr in 4% PF and then permeabilized for 1 hr in 0.5% Triton X-100. After rinsing in PBS, the
slices were first incubated for 10 min in cold (4°C) 0.1N HCl and
then incubated for 25 min at 37°C in 1N HCl. The HCl was removed and
the pH of the slices was neutralized by adding 0.1 M
Tris-HCl buffer, pH 8.0. The slices were then rinsed two times in PBS
and incubated overnight at 4°C in the anti-BrdU antibody (diluted
1:33 in PBS) (Boehringer Mannheim, Indianapolis, IN). Immunolabeling
for BrdU was visualized using the Elite Vectastain ABC kit (Vector
Laboratories, Burlingame, CA) and DAB. After immunostaining, slices
were briefly (30 sec) counterstained with 0.05% cresyl violet for
visualization of cytoarchitectural features. Slices were examined on a
Zeiss Axiophot microscope, and the density of BrdU-labeled cells,
cortical plate thickness, and total cp cells per unit area were
measured using NIH Image analysis software. Control and experimental
data were taken from corresponding anatomical positions. In every case,
the measurement was an experimental value normalized to control and
expressed as a percent. A total of 30 measurements from six coronal
slices from three different littermates were collected and averaged for
each trial. Illustrations present the averages (± SEM) from five
trials. Image analysis data were analyzed using Student's t test.
Density of BrdU-labeled cells. Using the NIH Image software,
the pixel density of BrdU-labeled nuclei was measured within five fixed
areas of the cortical plate in the control. The average pixel density
per unit area in the control slice was compared with similar
measurements on pixel density of labeled nuclei in the same fixed area
and position on the treated contralateral slice. Each capture was
exposed to the same analysis subroutine and density slice thresholding.
Cortical plate thickness. Using the NIH Image software, the
perpendicular linear distance (in pixels) was measured from the innermost edge of cortical layer VI to the outermost edge of cortical layer I. This distance was assessed in four places in a control slice
and compared with values measured in matched positions on the
contralateral treated slice.
Packing cell density in the cp. The average number of cells
per unit area within the cortical plate of control slices was compared
with the average number of cells per the same unit area in
contralateral treated slices.
Semiquantitative PCR
A hot-start reverse transcription (RT)-PCR protocol was
used to measure gene expression. Gene-specific primers were
designed from GenBank sequences using the Oligo software (National
Biosciences, Plymouth, MA). The sequences were as follows: NR1f,
ATAGTGACAATCCACCAAGAGCC; NR1r, GTAGCTCGCCCATCATTCCGTT; NMDA2Af,
CTCCAATATGTCCAACATGAACTCC; NMDA2Ar, GTCAACATCGCTACAGTCCTTGGG; NMDA2Bf,
TCCGCCGAGAGTCCTCCGT; NMDA2Br, CTGCGTTGCCCTCGATGTT; NMDA2Df,
GAACAGCAGACCCTCCGCC; NMDA2Dr, ACCCTTGCAGCATCTCTTCTCC. Total RNA was
isolated from cortices of E17-E18 mice. RNAstat 60 (Tel-Test, Inc.,
Friendswood, TX) and its recommended protocol was used for
the RNA isolation. To confirm purity of the product RNA, absorption
ratios at 260 nm/280 nm were determined to be >1.8 for all samples.
The samples were adjusted to 200 ng/µl for reverse transcription and
PCR according to absorption at 260 nm. PCR [Perkin-Elmer (Norwalk, CT)
GeneAmp RNA PCR kit; Applied Biosystems, Foster City, CA] involved
preheating a mixture of Taq antibody (TaqStart; Clontech,
Palo Alto, CA), primers, cDNA, and PCR components to 97°C for 90 sec
before amplification. PCR cycle was 30 sec at 95°C (dissociation), 45 sec at 60°C (annealing), and 60 sec at 72°C (extension).
Amplification was within the exponential range. Control RNA
(transcribed from PAW 108 plasmid DNA; PAW 108 forward and reverse
primers; Applied Biosystems) in all reverse transcription and PCR
amplification reactions allowed for a ratiometric method for
determining gene expression levels and permitted detection of
inefficient PCR reactions.
To control for artifacts caused by DNA contamination, in some
studies, RT-PCR was performed without reverse transcriptase. In the
absence of reverse transcriptase, no bands were visualized. In
addition, RNA samples in some studies were verified to be free of DNA
contaminants as described previously (Somogyi et al., 1995).
Numbers were obtained by calculating the densitometry (NIH Image) of
PCR products separated on polyacrylamide gradient gels. The ratios of
sample bands to corresponding control bands was used as the measurement
of gene expression. For every gene, ratiometric data for each group of
triplicate reactions were averaged. PCR product identities were
confirmed by restriction enzyme digestion. The NR2A cDNA was cleaved
with AvrII enzyme, generating fragments of 435 and 160; NR2B
cDNA was cleaved with PstI, generating fragments of 149 and
310; NR1 cDNA was cleaved with ApaLI, generating fragments of 230 and 89; and NR2D cDNA was cleaved with EarI,
generating a fragment of 228.
Viability of cells in the cultured slices. The viability of
cells in the slice cultures was monitored using propidium iodide (PI),
which intercalates into double-stranded DNA and fluoresces a brilliant
orange using the FITC filters. Cultured slices were transferred from
the filters onto glass microscope slides and were completely covered
with a drop of PI (50 µg/ml; Sigma, St. Louis, MO). After a 5 min
incubation at room temperature, the sections were coverslipped and
examined immediately on a Zeiss Axiophot microscope, equipped with
epifluorescence and the appropriate filters for the visualization
fluoresceine. To induce cell death in the slices, one set of sections
was incubated at room temperature in ambient air for 4 hr and then was
transferred to glass slides, overlayed with the propidium iodide, and
examined on the microscope. Slices cultured for 3 d and acutely
labeled with PI had very few cells with labeled nuclei; however, a few
PI-positive (PI+) cells were observed scattered along the
outermost edges of the slices. No difference in cell viability was
noted between the antagonist-treated and the control slices. To ensure
that the PI was able to penetrate through the slice, one set of slice
cultures was allowed to stand in ambient air for 4 hr at room
temperature and then was exposed to PI. The slices incubated at room
temperature had an abundance of PI+ cells distributed throughout the
cerebral wall.
Ca2+ recording of vz cells by
digital videomicroscopy
Unless otherwise stated, cell recordings were performed in
normal physiological medium (NPM) composed of the following reagents (in mM): 145 NaCl, 5 KC1, 1.8 CaCl2, 0.8 MgCl2, 10 glucose, and 10 HEPES, pH 7.3 (osmolarity, 290 mOsm). Ca2+-free
recording media contained (in mM): 145 NaCl, 5 KCl, 0.8 MgCl2, 5 EGTA-NaOH, 10 glucose, and 10 HEPES, where
Ca2+ was estimated to be <10 nM. NMDA
and APV were obtained from Research Biochemicals (Natick, MA). After
dissociation, vz cells were washed in NPM and plated at a density of
3 × 104 cells/cm2 on
poly-D-lysine coated, photo-etched gridded coverslips
(Bellco Glass Inc., Vineland, NJ) preglued to 35 mm tissue culture
dishes (MatTek Corp., Ashland, MA). After being allowed to adhere for 1 hr at 37°C, the cells were loaded with 2 µM fura-2 AM
(Molecular Probes, Eugene, OR) for 1 hr at 37°C. At the end of the
incubation period, the cells were rinsed in NPM.
Fura-2-loaded cells were recorded using the Zeiss Attofluor
Ratio-Vision work station (Atto Instruments, Rockville, MD)
equipped with an Axiovert 135 inverted microscope (Zeiss, Thornwood,
NY) and an ICCD camera (Atto Instruments). The fura-2 dye was
sequentially excited at 500 msec intervals with a 100 W mercury arc
lamp filtered at 334 ± 5 and 380 ± 5 nm and the respective
emissions acquired through a 510 nm dichroic mirror and 520 nm
long-pass filter set. All filters were obtained from Chroma Technology
Corp. (Brattleboro, VT). To collect the fura-2 fluorescence data,
either square- or polygonal-shaped regions of interest (ROI) were
electronically drawn around each of up to 69 cells per recording field.
The fluorescence intensities from each ROI were digitized with a Matrox
Graphics (Dorval, Quebec, Canada) image processing board and plotted as line graphs using Attograph for Windows analysis software (Atto Instruments).
Fura-2 fluorescence emissions were converted into estimated
[Ca2+]c concentrations using the
following equation:
where KD is the
fura-Ca2+ binding constant (225 nM),
R is a ratio of fura-2 fluorescence at 334 and 380 nm,
Rmin and Rmax are values
of R in Ca2+-free and normal
[Ca2+]o medium, respectively, using
fura-2 Penta K+ salt (Molecular Probes) as a
Ca2+ indicator, and
F0/F is the
ratio of fura-2 fluorescence at 380 nm in Ca2+-free
and 1.8 mM [Ca2+]o medium.
The data were calibrated on-line using the Attofluor RatioVision
acquisition software (Atto Instruments).
Cell responses to NMDA were examined by exposing the cells to 10 µM NMDA (Research Biochemicals) in the presence or
absence of 100 µM APV. The source of
Ca2+ contribution to
[Ca2+]c was evaluated by exposing the
cells to NMDA in Ca2+-free recording media. All
media were delivered to the cells using gravity-driven perfusion at
flow rates of ~2 ml/min. All measurements were performed at
37°C.
Electrophysiology
The current responses to NMDA were examined in cells of the vz
preparation using electrophysiology. All recordings were performed at
room temperature (22-25°C) on a Nikon (Tokyo, Japan) inverted microscope. Before recording, dishes were removed from the incubator, and the culture medium was completely replaced with Tyrode's solution containing (in mM): 145 NaCl, 5.4 KCl, 1.8 CaCl2, 0.8 MgCl2, 10 glucose, and
10 HEPES-NaOH, pH 7.4 (310 mOsm). Standard patch-clamp recordings
(Hamill et al., 1981) were made with pipettes pulled in three
stages from 1.5 mm outer diameter glass capillary tubes (World
Precision Instruments, Sarasota, FL) with a computer-controlled pipette
puller (BB-CH-PC; Mecanex SA, Nyon, Switzerland). These pipettes
had a resistance of 3-5 M when filled with internal solution
composed of (in mM): 145 CsCl, 2 MgCl2,
0.1 CaCl2, 1.1 EGTA, 5 HEPES, 5 ATP (potassium
salt), and 5 phosphocreatine, pH 7.2, (290 mOsm). Whole-cell currents
were recorded with a L/M EPC-7 patch-clamp amplifier (Medical Systems
Corp., Greenvale, NY) at a gain of 5 mV/pA. Series resistance was
compensated for more than 70%. Current signals were digitized with a
Digidata 1200 (Axon Instruments, Foster City, CA) and acquired with
Axoscope 7.0 (Axon Instruments) on a Pentium-based personal computer.
Recorded cells were continuously superfused with a perfusion system
composed of a locally made perfusion controller and miniature electric solenoid valves (The Lee Co., Essex, CT) that allows fast switching (<200 msec complete solution exchange time) among different solutions (Liu et al., 1999). The perfusion rate (0.3-0.5 ml/min) was controlled by the air pressure applied to the solution reservoirs. Significance of
responses to 1 µM NMDA were evaluated using Student's
t test to compare 8300 points recorded at the holding
current with 8300 points recorded during exposure to 1 µM NMDA.
Immunostaining embryonic cortical sections
Brains from E13 or E16 mice were removed, fixed for 48 hr in 4%
PF with 0.1% glutaraldehyde, and then equilibrated in 30% sucrose.
Frozen coronal sections (12 µm) were prepared using a cryostat.
Serial sections were incubated for 48 hr at 4°C in rabbit anti-glutamate antibody (1:100; Signature Biologicals, Salt Lake City,
UT), rinsed in three changes of PBS, and then incubated 1 hr at
room temperature in TRITC-conjugated donkey anti-rabbit IgG (1:50;
Jackson ImmunoResearch). Sections were examined using 10×, 16×, and
25× objectives on an Axiophot microscope (Zeiss) equipped with
epifluorescence and the appropriate filters for the visualization of rhodamine.
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RESULTS |
Glutamate is present in the embryonic murine cortex during the
stages of neurogenesis and neuronal migration
Cryostat sections of embryonic cortex were immunostained with
anti-glutamate antibodies at two embryonic ages to identify the
distribution of glutamatergic cells and fibers during the prenatal
period. At E13, high levels of glutamate were detected in the fibers
and cells of the primordial plexiform layer (Fig. 1A). At this stage, the
neuroepithelium did not label with the anti-glutamate antibodies. By
E16, glutamate-immunoreactive elements were identified in the outer
half of the cortical plate (Fig. 1B), whereas the
cells of the subplate were not immunopositive. Within layer I, fibers
also immunolabeled with the anti-glutamate antibodies. Thus, at the
ages examined, glutamate-immunoreactive structures were detected near
the target destinations for migrating cortical neurons.
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Figure 1.
Glutamate immunoreactivity in the
embryonic murine cortex. Photomicrographs of coronal sections of the
embryonic cortex immunostained with anti-glutamate antisera.
A, At E13, glutamate-immunoreactive cells
(arrow) and processes are evident in the primordial
plexiform layer (PP). B, By E16, fibers
in the outer half of the cortical plate are highly immunoreactive for
glutamate. ne, Neuroepithelium; cp, cortical plate; sp,
subplate. Scale bar, 40 µm.
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Glutamate stimulates migration of dissociated embryonic cortical
cells in vitro
An in vitro microchemotaxis assay was used to analyze
the effects of glutamate on the migration of cortical cells acutely dissociated from embryonic (E13-E18) C57BL mice. At E17, glutamate stimulated cell motility in a dose-dependent manner (Fig.
2A). Glutamate
concentrations ranging from 5 nM to 5 µM
produced levels of migration that were significantly higher than
spontaneous random motility to buffer only (1-5
cells/mm2). Maximum migration to glutamate was seen
at 500 nM. GABA also induced significant levels of
migration compared with spontaneous values. However, fewer cells moved
in response to GABA compared with glutamate, and the effective
concentration range of GABA was more limited (Fig.
2A). At optimal concentrations (500 nM), the number of cells migrating to glutamate was 10-fold greater than the
number of cells responding to GABA. Approximately 5-20% of the
starting population migrated to glutamate during the 18 hr assay,
indicating that the amino acid stimulated motility in a subpopulation
of cells. Significant migration to glutamate was observed in
dissociates of each of the six embryonic ages analyzed; however, peak
motility to glutamate occurred at E17-E18 (Fig. 2B).
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Figure 2.
Glutamate is a more potent chemoattractant than
GABA for dissociated murine embryonic cortical cells. A,
Glutamate and GABA stimulate migration in a dose-dependent manner. At
E17, distinctive ranges of glutamate ( ) or GABA ( ) stimulate
cells to migrate. Maximum numbers of migrating cells occur at 500 nM glutamate; however, significant migration (>50
cells/mm2) is observed at glutamate concentrations
ranging between 5 nM and 5 µM. Nanomolar GABA
also stimulates motility but at lower levels. At all effective
concentrations, glutamate stimulates more migration than GABA.
B, Response by age. Glutamate (1 µM) stimulates migration from E13 onward. Peak
migratory responses to glutamate are observed at E17. C,
Characterization of migration. Glutamate (1 µM) or NMDA
stimulate both directed migration (chemotaxis) and random motility
(chemokinesis). Approximately twice as many cells migrate in the
presence of a chemical gradient ( ) than in the absence of one ( ).
Error bars indicate SEM. *p  0.01; ANOVA followed
by Fisher's PLSD test. Separate trials: A, Glutamate,
n = 6; GABA, n = 4;
B, E13, n = 3; E14,
n = 3; E15, n = 3; E16,
n = 3; E17, n = 6; E18,
n = 3; C, n = 4.
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We characterized the migratory response to glutamate in terms of the
requirement for a chemical gradient across the membrane in the
chemotaxis chamber. The number of cells undergoing gradient-dependent migration (chemotaxis) was significantly greater than the number exhibiting gradient-independent random motility (chemokinesis) (Fig.
2C).
The chemotropic effects of glutamate are mimicked by NMDA and
blocked by BAPTA-AM
To determine which type(s) of glutamate receptor mediates cell
motility, agonists of different ionotropic glutamate receptors were
assessed for their ability to stimulate migration. Only NMDA (1 µM) mimicked the effects of 1 µM glutamate,
stimulating a similar number of cortical cells to migrate (Fig.
3A). NMDA stimulated both
chemotaxis and chemokinesis; however, nearly twice as many cells
exhibited gradient-dependent migration compared with
gradient-independent random motility (Fig. 2C). AMPA,
kainate, and quisqualate failed to stimulate motility that was
significantly different from spontaneous migration in controls. The
conformational isomer, D-glutamate, also failed to induce
migration. Thus, the signal for glutamate-induced migration appears to
be mediated via NMDA-Rs. To confirm the involvement of NMDA-Rs,
migration was assessed in the presence of 1 µM glutamate and serial dilutions of the NMDA-R antagonists MK801 or APV. In the
presence of MK801 (100 nM to 100 µM), a
noncompetitive receptor antagonist (Monaghan and Wenthold, 1997),
glutamate-induced migration was reduced to 50% (Fig. 3B).
APV, a competitive receptor antagonist (Monaghan and Wenthold, 1997),
blocked glutamate-induced migration in a dose-dependent manner (Fig.
3B), providing further evidence that glutamate-induced
motility involves NMDA-Rs. High concentrations of APV (10-100
µM) inhibited glutamate-induced migration by nearly 67%;
lower levels of APV failed to block significant levels of migration.
Both chemotactic and chemokinetic responses to 1 µM glutamate or NMDA were attenuated in the presence of 10 µM either antagonist (data not shown). In the
absence of [Mg2+]o, migration
induced by lower levels of NMDA was potentiated (data not shown).
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Figure 3.
NMDA receptors mediate glutamate-induced cortical
cell migration. A, Only NMDA mimics the effects of
glutamate. E17 cells were migrated to 1 µM
L-glutamate, D-glutamate, kainate, quisqualate,
NMDA, or AMPA. Only NMDA stimulates a similar level of motility as
L-glutamate. B, MK801 or APV, antagonists at
NMDA receptors, reduce the number of cells migrating to glutamate.
Glutamate (1 µM) was mixed with serial dilutions of MK801
or APV (100 nM to 100 µM). At all
concentrations of MK801, migration to glutamate was inhibited ~50%.
APV blocked glutamate-induced migration in a dose-dependent manner.
C, Inhibition of migration by BAPTA-AM. Migration of E17
cells to 1 µM glutamate () or NMDA () is entirely
blocked in the presence of 10 µM BAPTA-AM.
D, The metabotropic receptor agonist ACPD stimulates
migration in a dose-dependent manner. Migratory responses to micromolar
concentrations of ACPD (1-100 µM) are similar to
migration induced by 1 µM glutamate (dashed
line). Lower levels of ACPD fail to stimulate significant
migration (>50 cells/mm2). *p 0.01; ANOVA followed by Fisher's PLSD test. Separate trials:
A, n = 5; B,
n = 3; C, n = 3.
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We also assessed the effect of the Ca2+-chelator
BAPTA-AM on NMDA- and glutamate-induced chemotaxis. BAPTA (10 µM) was added directly to the wells containing the cells
during the incubation period. In the presence of BAPTA-AM, cell
migration to glutamate and NMDA was completely blocked (Fig.
3C), indicating that NMDA- or glutamate-induced elevations
in cytosolic Ca2+
([Ca2+]c) are required
for chemotaxis to occur.
In these studies, NMDA was the only ionotropic glutamate receptor
agonist that mimicked the chemotropic effects of glutamate. However,
exposure to the NMDA-R antagonists resulted in only a partial block of
glutamate-induced cortical cell migration. This suggests that a
subpopulation of cells migrates to glutamate via activation of a
different class of receptor, such as the metabotropic receptors.
Therefore, we migrated E17 cortical cells to decreasing concentrations
of trans-(±)-1-amino-1,3-cyclopentanedicarboxylic acid
(ACPD), an agonist of metabotropic glutamate receptors (Monaghan and
Wenthold, 1997). E17 cortical cells migrated to ACPD in a dose-dependent manner (Fig. 3D); micromolar ACPD stimulated
a robust migratory response, whereas nanomolar concentrations failed to
induce migration that was significantly higher than spontaneous motility to buffer only.
Viability studies demonstrated that the 18 hr exposure of the
dissociated cells to the ligands used in the study did not alter cell
survival (see Materials and Methods). Thus, the observed decreases in
the number of migrating cells when BAPTA or the antagonists was present
reflects inhibition of motility responses and not death of the cell.
Glutamate stimulates chemotaxis of vz neurons
To identify the anatomical origins of the migrating cells, E17
cortices were first microdissected into ventricular zone/subventricular zone (vz) and cortical plate/subplate (cp) regions. Immunocytochemistry was used to confirm that the microdissection effectively separated differentiating neurons of the cp (TUJ1+ only) (Lee et al., 1990) from
the more immature neurons and precursors of the vz (TUJ1+/nestin+ and
TUJ1/nestin+, respectively). Immunostaining demonstrated that
the two dissociates were comprised of cells at different stages of
maturation. In the vz preparation, ~45% of the cells expressed
nestin only (Table 1), indicating that
they were precursors or radial glia (Tohyama et al., 1992). The
remaining vz cells (55.7 ± 2.4%) were TUJ1+ neurons. Many of the
vz neurons also expressed nestin, indicating that they had recently
undergone terminal mitosis. In contrast, the cp preparation contained
primarily neurons (82.7 ± 5.0% TUJ1+) (Table 1), and the
majority of TUJ1+ cp cells (72.4%) did not express nestin,
demonstrating that they were more differentiated.
The migration of cellular dissociates from each region were examined in
the microchemotaxis assay. In vitro, the dissociated vz
cells demonstrated a greater migratory response to glutamate or NMDA
than the cp cells (Fig.
4A). Moreover,
migration was limited to neuronal cells because immunostaining
demonstrated that all of the migrated vz cells expressed NF protein
(Fig. 4B).
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Figure 4.
Response by region. A, In
vitro migratory responses of vz and cp cells. Vz cells
(filled bars) exhibit greater migration to 1 µM glutamate or NMDA than cp cells (open
bars) Significant numbers of cp cells do not respond to
glutamate. Error bars indicate SEM. *p 0.01;
ANOVA followed by Fisher's PLSD test. B, Vz neurons
migrate to glutamate. Vz cells were immunolabeled for neurofilament
protein after migrating to glutamate in the chemotaxis assay. All vz
cells that migrate to glutamate in vitro express
neurofilament protein, indicating that they are neurons.
Asterisks denote 8 µm pore in the membrane. Scale bar,
20 µm. Separate trials: A, B,
n = 7.
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Vz cells express NMDA-Rs
Vz and cp tissue segments were microdissected from E17-E18
cortices and were analyzed by semiquantitative PCR for expression of
mRNA encoding NMDA-R subunit proteins. Transcripts detecting NR1 and
three NR2 subunits (NR2A, NR2B, and NR2D) were evident in the cp cells
(Fig.
5A1-D1). In
contrast, vz cells only expressed transcripts for NR1, NR2B, and NR2D
(Fig. 5A1-D1). Densitometry demonstrated
that the relative abundance of transcripts in cp cells was NR1 > NR2B > NR2D > NR2A (Fig.
5A2-D2). In the vz cells, NR1 > NR2B > NR2D (Fig.5A2-D2). Both the cp and vz cells expressed similar levels of mRNA encoding NR1 (Fig. 5A2). The results of the present PCR studies
demonstrated that cells derived from both the immature (vz) and more
differentiated (cp) regions of the cortex express mRNA encoding NMDA
receptor subunit proteins. Western blotting confirmed the presence of
NR1, NR2A, and NR2B proteins in embryonic cortical homogenates (data not shown).
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Figure 5.
Vz cells express mRNA encoding NMDA receptors
subunits. Semiquantitative PCR was used to probe E17 vz and cp
dissociates for mRNA encoding NR1 (A1,
A2), NR2A (B1,
B2), NR2B (C1,
C2), and NR2D (D1,
D2) subunits. Densitometry
(A2, B2,
C2, D2) revealed average
relative abundance of transcripts in tissue homogenates from each
region. For each region, n = 3.
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NMDA-induced physiological responses of cells in the
vz dissociate
Digital video microscopy was used to assess the effect of NMDA on
[Ca2+]c levels in vz cells preloaded
with the Ca2+ indicator dye, fura-2. NMDA evoked
either rapidly rising (31 ± 12 sec to peak) or slowly rising
(238 ± 56 sec to peak) elevations in
[Ca2+]c in 15 and 9% of the recorded
vz cells (n = 117), respectively (Fig.
6). Typically, the more rapid response
was associated with a larger increase in
[Ca2+]c (830 ± 517 nM) when compared with the increase that occurred during
the slower response (94 ± 47 nM). The former response
was predominantly observed in larger (>10 µm in diameter),
process-bearing cells. Both responses were antagonized by APV and were
dependent on extracellular Ca2+
[Ca2+]o. In some studies,
cells were fixed and labeled with TUJ1 antibody after the fura-2
recording. All of the cells that responded to NMDA with increased
[Ca2+]c immunolabeled with the TUJ1
antibody, indicating they were postmitotic neurons.
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Figure 6.
NMDA induces increases in
[Ca2+]c in vz cells. After recording
their resting [Ca2+]c in control
(CON) medium (A), the cells
were sequentially exposed to 10 µM NMDA
(B), 100 µM APV plus 10 µM NMDA, followed by 10 µM NMDA in
[Ca2+]o-free saline (see log
above the traces for the duration of each
exposure). The data show rapid (Cell #1) and slow
(Cell #2) [Ca2+]c
responses evoked by NMDA, typical of those recorded in the vz
population. Both responses were reversibly blocked by APV and
eliminated in [Ca2+]o-free
saline.
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Electrophysiology
NMDA-induced currents were recorded in cells of the vz preparation
using conventional patch-clamp recording techniques. When membrane
potential was clamped at 80 mV, low concentrations of NMDA (1 µM) induced a just-detectable inward current deflection (1.12 ± 0.01 pA), which was significantly different
(p < 0.001) from the holding current
(n = 2 of 7). Higher levels of NMDA induced larger
currents with means of 6.62 ± 0.04 pA at 10 µM NMDA
(n = 3 of 10) and 13.83 ± 0.08 pA at 50 µM NMDA (n = 5 of 10), which were
superimposed with obvious fluctuations, presumably reflecting the
underlying single-channel activity. The currents induced by 50 µM NMDA were almost completely blocked by equimolar
concentrations of the NMDA antagonist MK801 (0.16 ± 0.03 pA)
(Fig. 7).
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Figure 7.
NMDA-induced currents in vz cells.
A, At 1 µM, NMDA induces a small
(1.12 ± 0.01 pA; p < 0.001) inward current
deflection that is significantly different from the holding current. At
higher concentrations (B, C), NMDA
induces larger currents with obvious superimposed open-channel noise.
At the higher concentrations, the mean currents are 6.62 ± 0.04 pA at 10 µM (B) and 13.83 ± 0.08 pA at 50 µM (C). The current
induced by 50 µM NMDA (C) is almost
completely blocked by equimolar MK801 (0.16 ± 0.03).
A and B were recorded from the same cell
in the vz preparation; C was recorded from a different
cell in the vz dissociate. Membrane potential was clamped at 80
mV.
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Neuronal migration in cultured cortical slices is mediated
via NMDA-Rs
The studies on the dissociated cells revealed that direct
application of glutamate stimulates chemotaxis of dissociated embryonic cortical neurons. To determine whether glutamate generated by cells
within the cortex influences cell motility, we analyzed the effect of
NMDA-R antagonists on cellular migration in organotypic cortical slice
cultures. In theses studies, no exogenous glutamate was added to the
slice cultures. In the developing cortex, the proliferating cells are
located in the ventricular and subventricular zones (Jacobson, 1991).
Because proliferating cells incorporate BrdU, we exposed the cultured
slices to a pulse of BrdU and then used changes in the anatomical
distribution of BrdU-labeled cells in the slices over time as an
indication of postmitotic cell migration. Patterns of migration were
assessed by administering an 18 hr pulse of BrdU to acutely prepared
E17 slices. Culture medium was then replaced with growth media with or
without NMDA-R antagonists, and slices were cultured for 2 or 6 additional days. Slices were fixed and immunostained for BrdU, and the
distribution of labeled cells within the slices was examined. After
2 d of culture under these conditions, virtually all of the
BrdU-labeled cells in control slices were localized to the cp and sp
(Fig. 8A); the
ventricular and intermediate zones in the control slices were virtually
devoid of labeled cells (Fig. 8C). Because proliferation
occurs within the ventricular regions, these results suggest that the
cells that incorporated BrdU during the pulse migrated away from the germinal regions into the sp and cp. In contrast, contralateral slices
maintained for 48 hr in the presence of APV (100 µM) had few labeled cells in the cp (Fig. 8B). The
BrdU-positive cells in the treated slices were distributed throughout
the ventricular and intermediate zones (Fig. 8D).
Similar results were observed when slices were cultured for 6 d in
the presence of APV or 100 µM MK801 (data not shown).
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Figure 8.
Glutamate-induced migration of cells in cultured
cortical slices. A, Control slice maintained in
vitro for 48 hr after the BrdU pulse. Almost all BrdU-labeled
cells are in the cp and sp. B, The contralateral slice
treated for 48 hr with 100 µM APV has few BrdU-labeled
cells in the cp or sp. Scale bar, 80 µm. C, Low
magnification of an untreated control slice shows few labeled cells in
the vz or iz. D, The contralateral treated slice has an
abundance of labeled cells in the vz and iz. Scale bar, 80 µm.
E, Densitometry of cultured slices. Densitometry
of contralateral slices treated with with either 100 µM
APV () or MK801 (). E, Density of BrdU-labeled
nuclei in the cortical plate. At 2 and 6 d, there is a significant
reduction in the number of BrdU-labeled cells in the cp of
antagonist-treated slices compared with controls. F,
Cortical plate thickness. At 2 d, antagonist treatment results in
a minor decrease in cp thickness. By 6 d, the thickness of the
cortical plate is not significantly diminished. G,
Density of total cells in the cortical plate. At both 2 and 6 d,
antagonist treatment results in significantly fewer cells per unit area
when compared with controls. *p 0.05; Student's
t test. Separate trials: n = 5.
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Densitometry of the BrdU-labeled slices demonstrated that, in the
presence of the NMDA-R antagonists, cellular migration into the cp was
significantly attenuated (Fig. 8E). Slices maintained in APV for 48 hr showed an 80 ± 5.1% decrease in the
number of BrdU-labeled cells in the cp. Slice cultures maintained
for an extended period up to 6 d showed similar results. After
6 d of culture in APV, labeled cp cells decreased by 74 ± 10.3%. Similarly, slices maintained for 6 d in MK801 had an
85 ± 4.8% reduction of BrdU-labeled cells in the cp.
Effects of antagonists on cortical thickness in slice cultures
Because fewer BrdU-labeled cells migrated into the cp in
antagonist-treated slices, the overall thickness of the cp might be
expected to decrease. Therefore, we measured the thickness of the cp
(from the innermost edge of layer VI through outermost edge of layer I)
in treated and untreated slices. Antagonist treatment did not result in
a significant decrease in cortical plate thickness. After 48 hr of
culture, APV exposure resulted in only a 17 ± 10.3% decrease in
cp thickness (Fig. 8F). Similarly, MK801-treated
slices showed only a minor reduction of 18 ± 11.0% in the
overall thickness of the cp when compared with controls. After 6 d
of culture, treated and untreated cultures exhibited no significant
difference in cp thickness (Fig. 8F).
Effects on cell density within the cp
These studies indicated that, although antagonist treatment
resulted in a marked reduction in the number of BrdU-labeled cells in
the cp, the overall cp thickness was not markedly reduced. To determine
whether the antagonist treatment resulted in increased extracellular
space between cp cells, we measured the relative "packing" density
of total cells within the cp in control versus treated slices.
Nontreated control slices contained a greater number of cells per unit
area than treated slices (Fig. 8G). This was evident in
slices treated with antagonists for 2 and 6 d. APV treatment for
2 d decreased the number of cells per unit area by 25 ± 1.3% compared with nontreated slices. After 6 d in culture, the
number of cells per unit area in treated slices was also reduced (by
35 ± 1.8% in APV and 45 ± 2.4 in MK801) (Fig.
8G). Thus, although the overall thickness of the cp was not
markedly diminished, the packing density of cells in treated
cultures was significantly less, indicating that exposure to NMDA-R
antagonists resulted in fewer BrdU-labeled cells and an overall
decrease in the number of total cells in the developing cp.
| |
DISCUSSION |
Our studies demonstrate that murine embryonic cortical neurons
migrate in response to glutamate via mechanisms that involve NMDA
receptors. NMDA was the only ionotropic agonist that mimicked the
effects of glutamate on acutely dissociated cortical cells, stimulating
a similar number of cells to migrate. Glutamate-induced migration of
dissociated cells was inhibited by either MK801 or APV, antagonists at
NMDA receptors (Monaghan and Wenthold, 1997). Thus, in a subpopulation
of acutely dissociated murine embryonic cortical cells, NMDA receptors
appear to mediate motility responses to glutamate. The cortical slice
cultures supported the findings in the dissociated cells, providing
further evidence that NMDA-Rs are involved in migratory responses to
glutamate. Slices maintained in NMDA-R antagonists had an abundance of
BrdU+ cells in the germinal zones but lacked BrdU-labeled cells in the
cp region, suggesting that in the slices, NMDA-R antagonists inhibited
postmitotic cell migration into the cp. Together, these results suggest
that, in vivo, glutamate serves as a chemoattractant,
stimulating the migration of murine embryonic cortical neurons toward
their target destinations in the cp.
If chemoattractants provide positional cues to cells, then
they should localize near the target destinations of migrating cells.
In the embryonic mouse cortex, glutamate is highly expressed in the
outer half of the cortical plate and in the C-R cells which lie
within the primordial plexiform layer, near the target destination of
migratory neurons (del Rio et al., 1995). In the developing rat cortex,
GABA exhibits a similar distribution (Imamoto et al., 1994; Behar et
al., 1996), and GABA has been shown to be a potent chemoattractant for
rat embryonic cortical cells (Behar et al., 1996). In the present
study, glutamate was a more effective chemoattractant for murine
cortical cells than GABA. Thus, although the ligands differ, both rat
and mouse appear to use motility-signaling molecules found in the
developing cortical plate and C-R cells.
If chemoattractants play a relevant role in development, then migrating
cells should express receptors for the attractants. During the final
week of gestation, in situ hybridization studies have
revealed that cells throughout the embryonic mouse cortex express mRNA
transcripts encoding NMDA-R subunits (Feldmeyer and Cull, 1996;
Monaghan and Wenthold, 1997). Here, RT-PCR revealed that both vz and cp
dissociates express mRNA encoding NR1, an NMDA-R subunit protein that
is an essential component of NMDA receptors (Monaghan and Wenthold,
1997). In addition, transcripts for NR2B and NR2D subunits were
detected in dissociates from the two anatomical regions. Previous
studies reported that embryonic cells in the cp of the developing
cortex express functional NMDA-Rs (LoTurco et al., 1991); our findings
demonstrate that cells derived from immature cortical regions also
exhibit physiological responses to NMDA. Electrophysiology and
Ca2+ imaging studies revealed that, after
application of NMDA, cells in the vz preparation showed an increased
current and exhibited increases in
[Ca2+]c. In vitro, neurons
dissociated from immature cortical regions exhibited migratory
responses to NMDA. Together, these results demonstrate that neurons in
the vz, svz, or lower intermediate zone express functional NMDA
receptors. Thus, when cortical lamination occurs in vivo,
both the attractant and its receptors are expressed in appropriate
anatomical locations to form a physiologically relevant circuit.
In vivo, diffusible gradients of glutamate released by cells
in the superficial regions of the cortex may act as chemoattractants for newly generated postmitotic neurons in the germinal zones, directing their migration into the cortical plate. Embryonic cortical sections immunostained for glutamate demonstrated that glutamate was
most highly expressed in the cortical plate, the target region for
migrating embryonic cortical neurons. In studies on the dissociated cells, glutamate primarily stimulated directed migration of neurons from immature regions: (1) more of the vz cells migrated to glutamate or NMDA than cp cells, (2) all of the responding vz cells were neuronal, and (3) the predominant mode of motility was directed migration, or chemotaxis. Peak migration to glutamate was observed at
E17, suggesting that glutamate prominently influences the migration of
neurons destined for cortical layers II and III (Angevine and Sidman,
1961; Schmidt and Lent 1987; Polleux et al., 1997). At earlier stages
of development, migrating cells destined for the deep cortical lamina
(V and VI) may lack the appropriate receptors and/or signaling
mechanisms that mediate chemotactic responses to glutamate.
Glutamate-induced migration requires increases in
[Ca2+]c. Migration to glutamate or
NMDA was abolished in the presence of the Ca2+-chelating agent BAPTA-AM, which maintains
[Ca2+]c at nanomolar levels. These
findings are consistent with previous studies reporting that cellular
locomotion requires increases in
[Ca2+]c (Komuro and Rakic, 1992, 1996;
Hinrichsen, 1993). In this regard, NMDA-Rs could directly influence
cell motility by triggering increases in
[Ca2+]c that modulate cytoskeletal
dynamics required for cell movement. In the present study, application
of NMDA resulted in a rapid flux of current in responding vz cells,
whereas the Ca2+ imaging experiments revealed a slow
rather than immediate rise in [Ca2+]c
after NMDA exposure. The slow rise in cytosolic Ca2+
suggests that NMDA receptor activation leads to local increases in
[Ca2+]c, which in turn trigger
Ca2+ release from intracellular stores and/or
stimulate further [Ca2+]o entry via
other pathways.
NMDA-Rs appear to mediate motility signals in a cortical cell
subpopulation. In other subpopulations, metabotropic receptors may
mediate the motility signals of glutamate. NMDA-R antagonists only partially blocked migration to glutamate, suggesting that some
cells migrate to glutamate via activation of a different type of
receptor. ACPD, a metabotropic receptor agonist (Monaghan and Wenthold,
1997), also stimulated dissociated cells to migrate, indicating that
glutamate receptors coupled to GTP binding proteins can induce nerve
cell movement. In rats, GABA promotes motility of cortical cells via
multiple classes of receptors, some of which couple to GTP binding
proteins (Behar et al., 1998). A similar situation may exist in the
developing murine cortex in which multiple classes of glutamate
receptors stimulate migration in distinct subpopulations of embryonic
neurons via different intracellular signaling mechanisms.
Cortical plate cells may express a non-NMDA-R that, when activated,
arrests cell movement. Although NMDA stimulated migration in both vz
cell and cp cell dissociates, the cp cells failed to migrate to
glutamate. This demonstrates that the cp cells had the capacity to
migrate in response to NMDA-R activation; however, they failed to move
in the presence of the natural ligand. In rats, the GABA receptors that
promote cp cell motility (GABAB- and
GABAC-like) differ from those that arrest cp cell movement (GABAA) (Behar et al., 1998). A parallel mechanism
may exist in the cp cells of the mouse; whereas NMDA-Rs promote
motility, activation of a second class of glutamate receptor may
attenuate migration. In vivo, migrating neurons may acquire
a functional form of this second class of receptor soon after entering
the cp. High concentrations of glutamate encountered near the target
destinations could activate the receptor, providing a "stop" signal
to migrating cells approaching their final positions.
Slice culture studies suggest that glutamate released from cortical
cells stimulates embryonic neuronal migration toward the cp via NMDA-R
activation
Blockade of NMDA-Rs prevents BrdU+ postmitotic cells from
migrating into the cp. In untreated slice cultures, 2 d after the pulse, most BrdU+ cells were located in the cp. Only proliferating cells in ventricular regions incorporate BrdU; hence, the vz cells that
took up the label during the pulse apparently migrated into the cp
within 2 d. In contrast, antagonist-treated slices had few BrdU+
cells in the cp after 2 d; most labeled cells in these slices were
observed in the vz, svz, and iz. The propidium iodide studies confirmed
that the antagonist treatment of the slices did not affect cell
viability (see Materials and Methods). Thus, the absence of BrdU+ cells
in the cp of treated slices was not a result of cell death. Rather, the
BrdU-labeled cells remained in the immature regions and apparently
failed to migrate into the cp. Although the cp thickness was not
diminished in antagonist-treated slices, the overall cell density was
decreased. Hence, by the end of the culture period, slices maintained
in NMDA-R antagonists had fewer total cp cells. These results
demonstrate that inhibition of NMDA-Rs during corticogenesis leads to
fewer cells in the cp, suggesting that in vivo, glutamate
released from cortical cells directs the movement of postmitotic
neurons into this region.
Our studies provide evidence that NMDA-Rs mediate motility signals in
embryonic cortical neurons. These results appear to contradict earlier
findings on transgenic mice that lack the NR1 subunit, which suggested
that NMDA receptors are not involved cortical neuronal migration
(Forrest et al., 1994; Messersmith et al., 1997). In an early study on
mice lacking functional NR1 genes, gross observations of the cortex
indicated that it had developed normally, although the authors did not
conduct a careful microscopic examination of the cerebral cortex that
would reveal subtle abnormalities in cortical lamination patterns
(Forrest et al., 1994). Thus, the authors concluded that they could not exclude the possibility that NMDA receptors influence cortical nerve
cell movement. In a subsequent report, Messersmith et al. (1997)
suggested that NMDA receptors are not involved in cortical neuronal
migration (Messersmith et al., 1997); however, their study examined the
migration of cells destined for deep cortical lamina (layers V and VI).
Here, we provide evidence that glutamate-induced migration primarily
affects the cells destined for superficial cortical lamina (layers II
and III). Furthermore, transgenic mice lacking NR1 subunits have
reportedly developed compensatory mechanisms for fluxing
Ca2+, which could directly influence cell motility
(Messersmith et al., 1997). Finally, there may be a redundancy of
mechanisms involved in glutamate-induced cortical cell migration. Our
studies suggest that both ionotropic and metabotropic receptors are
involved in cortical nerve cell movement. Compensatory mechanisms,
coupled with a redundancy in glutamate receptor mechanisms that promote cortical cell movement, could account for the reportedly "normal" morphology of the cerebral cortex observed in transgenic mice that lack
NR1 subunits.
The results of the present study provide evidence that rats and mice
use different chemoattractants to promote the migration of cells
destined for cortical layers II and III. In rats, GABA stimulates
robust migration, whereas glutamate is a more potent attractant for
murine cortical cells. In each species, the appropriate attractant is
found highly expressed near the target locations for migrating cells;
rat C-R and cp cells are GABAergic (Imamoto et al., 1994; Behar et al.,
1996), whereas murine C-R and cp cells are glutamatergic (del Rio et
al., 1995). It is unknown, however, whether the converse is true; rat
cp cells may express high levels of glutamate, whereas murine cp cells
may express GABA. Double-immunolabeling studies for GABA and glutamate
would reveal whether the two molecules are detectable in the cortical
plates of each species. Furthermore, GABA was a weak attractant for
murine cells at E17; however, the molecule may stimulate robust
chemotaxis at other developmental stages. Thus, a systematic study in
rats and mice that compares the chemotropic effects of GABA and
glutamate throughout the period of cortical histogenesis is warranted.
These types of studies would resolve whether or not the two species use
different signaling molecules to regulate cell chemotaxis in the
developing cortex.
In summary, our results on the dissociated cells suggest that glutamate
promotes chemotaxis of newly differentiated neurons within the
developing embryonic mouse cortex. Pharmacological studies indicate
that NMDA-Rs mediate the motility signals of glutamate in a
subpopulation of cells. The studies on organotypic slice cultures
confirmed the pharmacology demonstrated on the dissociated cells and
provided evidence that, in vivo, glutamate released from
cortical cells acts as a chemoattractant for embryonic neurons
migrating toward the cp. Glutamate has been reported to stimulate
neuronal movement via NMDA-Rs in the developing cerebellum (Komuro and
Rakic, 1993), suggesting a widespread role for glutamate as a
chemoattractant in the developing nervous system.
| |
FOOTNOTES |
Received Jan. 6, 1999; revised March 15, 1999; accepted March 22, 1999.
Correspondence should be addressed to T. N. Behar, Building 36, Room 2C02, 36 Convent Drive, Bethesda, MD 20892-4066.
| |
REFERENCES |
-
Angevine JB,
Sidman RL
(1961)
Autoradiographic study of cell migration during histogenesis of cerebral cortex in the mouse.
Nature
192:766-768[Medline].
-
Armstrong RC,
Harvath L,
Dubois-Dalcq M
(1990)
Type 1 astrocytes and oligodendrocyte-type 2 astrocyte glial progenitors migrate toward distinct molecules.
J Neurosci Res
27:400-407[Web of Science][Medline].
-
Behar TN,
Schaffner AE,
Colton CA,
Somogyi R,
Olah Z,
Lehel C,
Barker JL
(1994)
GABA-induced chemokinesis and NGF-induced chemotaxis of embryonic spinal cord neurons.
J Neurosci
14:29-38[Abstract].
-
Behar TN,
Li YX,
Tran HT,
Ma W,
Dunlap V,
Scott C,
Barker JL
(1996)
GABA stimulates chemotaxis and chemokinesis of embryonic cortical neurons via calcium-dependent mechanisms.
J Neurosci
16:1808-1818[Abstract/Free Full Text].
-
Behar TN,
Schaffner AE,
Scott CA,
O'Connell C,
Barker JL
(1998)
Differential response of cortical plate and ventricular zone cells to GABA as a migration stimulus.
J Neurosci
18:6378-6387[Abstract/Free Full Text].
-
del Rio J,
Martinez A,
Fonseca M,
Auladell C,
Soriano E
(1995)
Glutamate-like immunoreactivity and fate of Cajal-Retzius cells in the murine cortex as identified with calretinin antibody.
Cereb Cortex
5:13-21[Abstract/Free Full Text].
-
Eisen JS
(1991)
Determination of primary motoneuron identity in developing zebrafish embryos.
Science
252:569-572[Abstract/Free Full Text].
-
Falk W,
Goodwin RH,
Leonard EJ
(1980)
A 48 well micro-chemotaxis assembly for rapid and accurate measurement of leukocyte migration.
J Immunol Methods
33:239-247[Web of Science][Medline].
-
Feldmeyer D,
Cull CS
(1996)
Functional consequences of changes in NMDA receptor subunit expression during development.
J Neurocytol
25:857-867[Web of Science][Medline].
-
Forrest D,
Yuzaki M,
Soares HD,
Ng L,
Luk DC,
Sheng M,
Stewart CL,
Morgan JI,
Connor JA,
Curran T
(1994)
Targeted disruption of NMDA receptor 1 gene abolishes NMDA response and results in neonatal death.
Neuron
13:325-338[Web of Science][Medline].
-
Hamill OP,
Marty A,
Neher E,
Sakmann B,
Sigworth FJ
(1981)
Improved patch-clamp techniques for high-resolution current recording from cells and cell-free membrane patches.
Pflügers Arch
391:85-100[Web of Science][Medline].
-
Hatten ME
(1990)
Riding the glial monorail: a common mechanism for glial-guided migration in different regions of the developing brain.
Trends Neurosci
13:179-184[Web of Science][Medline].
-
Hinrichsen RD
(1993)
Calcium and calmodulin in the control of cellular behavior and motility.
Biochim Biophys Acta
1155:277-293[Medline].
-
Imamoto K,
Karasawa N,
Isomura G,
Nagatsu I
(1994)
Cajal-Retzius neurons identified by GABA immunohistochemistry in layer I of the rat cerebral cortex.
Neurosci Res
20:101-105[Web of Science][Medline].
-
Jacobson M
(1991)
In: Developmental neurobiology, Ed 3, pp 401-448. New York: Plenum.
-
Komuro H,
Rakic P
(1992)
Selective role of N-type calcium channels in neuronal migration.
Science
257:806-809[Abstract/Free Full Text].
-
Komuro H,
Rakic P
(1993)
Modulation of neuronal migration by NMDA receptors.
Science
260:95-97[Abstract/Free Full Text].
-
Komuro H,
Rakic P
(1996)
Intracellular Ca2+ fluctuations modulate the rate of neuronal migration.
Neuron
17:275-285[Web of Science][Medline].
-
Lee MK,
Tuttle JB,
Rebhun LI,
Cleveland DW,
Frankfurter A
(1990)
The expression and posttranslational modification of a neuron-specific
 -tubulin isotype during chick embryogenesis.
Cell Motil Cytoskeleton
17:118-132[Web of Science][Medline]. -
Liu Q-Y,
Schaffner AE,
Barker JL
(1999)
Electrophysiological studies on receptors in vitro.
In: In vitro neurochemical techniques (Boulton AA,
Baker GB,
Bateson AN,
eds), pp 37-61. Totowa, NJ: Humana.
-
LoTurco J,
Blanton M,
Kriegstein A
(1991)
Initial expression and endogenous activation of NMDA channels in early neocortical development.
J Neurosci
11:792-799[Abstract].
-
Messersmith EK,
Feller MB,
Zhang H,
Shatz CJ
(1997)
Migration of neocortical neurons in the absence of functional NMDA receptors.
Mol Cell Neurosci
5:347-357.
-
Monaghan DT,
Wenthold RJ
(1997)
In: The ionotropic glutamate receptors. Totowa, NJ: Humana.
-
O'Rourke NA,
Dailey ME,
Smith SJ,
McConnell SK
(1992)
Diverse migratory pathways in the developing cerebral cortex.
Science
258:299-302[Abstract/Free Full Text].
-
O'Rourke NA,
Sullivan DP,
Kaznowski CE,
Jacobs AA,
McConnell SK
(1995)
Tangential migration of neurons in the developing cerebral cortex.
Development
121:2165-2176[Abstract].
-
Polleux F,
Dehay C,
Kennedy H
(1997)
The timetable of laminar neurogenesis contributes to the specification of cortical areas in mouse isocortex.
J Comp Neurol
385:95-116[Web of Science][Medline].
-
Rakic P
(1972)
Mode of cell migration of the superficial layers of fetal monkey neocortex.
J Comp Neurol
145:61-84[Web of Science][Medline].
-
Rakic P
(1990)
Principles of neural cell migration.
Experientia
46:882-891[Web of Science][Medline].
-
Schambra UB,
Lauder JM,
Silver J
(1992)
In: Atlas of the prenatal mouse brain. San Diego: Academic.
-
Schmidt SL,
Lent R
(1987)
Effects of prenatal irradiation on the development of cerebral cortex and corpus callosumof the mouse.
J Comp Neurol
264:193-204[Medline].
-
Somogyi R,
Wen X,
Ma W,
Barker JL
(1995)
Developmental kinetics of GAD family mRNAs parallel neurogenesis in the rat spinal cord.
J Neurosci
15:2575-2591[Abstract].
-
Tohyama T,
Lee V,
Rorke L,
Marvin M,
McKay R,
Trojanowski J
(1992)
Nestin expression in embryonic human neuroepithelium and in human neuroepithelial tumor cells.
Lab Invest
66:303-313[Web of Science][Medline].
Copyright © 1999 Society for Neuroscience 0270-6474/99/19114449-13$05.00/0
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[Abstract]
[Full Text]
[PDF]
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[Abstract]
[Full Text]
[PDF]
|
|
|
|
|
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9 - 17.
[Abstract]
[Full Text]
[PDF]
|
|
|
|
|
|
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744 - 753.
[Abstract]
[Full Text]
[PDF]
|
|
|
|
|
|
|
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PNAS,
April 25, 2001;
(2001)
91113598.
[Abstract]
[Full Text]
|
|
|
|
|
|
|
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J. Neurosci.,
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21(3):
934 - 943.
[Abstract]
[Full Text]
[PDF]
|
|
|
|
|
|
|
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J. Neurosci.,
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20(15):
5764 - 5774.
[Abstract]
[Full Text]
[PDF]
|
|
|
|
|
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J. Neurosci.,
January 15, 2000;
20(2):
696 - 708.
[Abstract]
[Full Text]
[PDF]
|
|
|
|
|
|
|
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PNAS,
May 22, 2001;
98(11):
6372 - 6377.
[Abstract]
[Full Text]
[PDF]
|
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