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The Journal of Neuroscience, July 15, 2001, 21(14):5139-5146
Abnormal Development of Dendritic Spines in
FMR1 Knock-Out Mice
Esther A.
Nimchinsky,
Adam M.
Oberlander, and
Karel
Svoboda
Howard Hughes Medical Institute, Cold Spring Harbor Laboratory,
Cold Spring Harbor, New York 11724
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ABSTRACT |
Fragile X syndrome is caused by a mutation in the
FMR1 gene leading to absence of the fragile X mental
retardation protein (FMRP). Reports that patients and adult
FMR1 knock-out mice have abnormally long dendritic
spines of increased density suggested that the disorder might involve
abnormal spine development. Because spine length, density, and motility
change dramatically in the first postnatal weeks, we analyzed these
properties in mutant mice and littermate controls at 1, 2, and 4 weeks
of age. To label neurons, a viral vector carrying the enhanced green
fluorescent protein gene was injected into the barrel cortex.
Layer V neurons were imaged on a two-photon laser scanning microscope
in fixed tissue sections. Analysis of >16,000 spines showed clear
developmental patterns. Between 1 and 4 weeks of age, spine density
increased 2.5-fold, and mean spine length decreased by 17% in normal
animals. Early during cortical synaptogenesis, pyramidal cells in
mutant mice had longer spines than controls. At 1 week, spine length was 28% greater in mutants than in controls. At 2 weeks, this difference was 10%, and at 4 weeks only 3%. Similarly, spine density was 33% greater in mutants than in controls at 1 week of age. At 2 or
4 weeks of age, differences were not detectable. The spine abnormality
was not detected in neocortical organotypic cultures. The transient
nature of the spine abnormality in the intact animal suggests that FMRP
might play a role in the normal process of dendritic spine growth in
coordination with the experience-dependent development of cortical circuits.
Key words:
fragile X; FMRP; dendritic spine; critical period; somatosensory cortex; development; mental retardation; two-photon
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INTRODUCTION |
Mental retardation is a component of
many syndromes. Some are caused by large-scale chromosomal
abnormalities; others are caused by physical, infectious, or
biochemical insults sustained early in brain development. The fragile X
syndrome is remarkable because it is a mental retardation syndrome
caused by a mutation in a single gene, FMR1. It is
characterized by a constellation of signs in addition to the cognitive
deficit, including macro-orchidism, certain facial features, and
abnormalities in attention and short-term memory (Schapiro et al.,
1995 ; de Vries et al., 1998). The mutation giving rise to the syndrome,
a CGG repeat expansion in the 5' untranslated region of the
FMR1 gene, interferes with transcription, and patients do
not have measurable amounts of the gene product, fragile X mental
retardation protein (FMRP) (Pieretti et al., 1991). This protein
appears to act as an RNA binding protein (Feng et al., 1997; Brown et
al., 1998) and is localized to neurons, and to dendrites in particular
(Devys et al., 1993; Verheij et al., 1993; Feng et al., 1997).
Interestingly, FMRP mRNA is found in dendrites, and the
expression of FMRP is increased by activation of metabotropic glutamate
receptors (Weiler et al., 1997), linking FMRP to synaptic
function. Remarkably little is known about the neuropathologic
features of this disorder. One finding reported both in patients
(Rudelli et al., 1985; Hinton et al., 1991; Irwin et al., 2001) and in
adult FMR1 knock-out mice (Comery et al., 1997) is an
abnormality in dendritic spines, which were described as being
unusually long, similar to the spines observed in cortical circuits
during development (Dailey and Smith, 1996; Fiala et al., 1998; Lendvai
et al., 2000), and of increased density. These observations have led
some to speculate that the absence of FMRP causes a defect in spine
maturation and pruning (Comery et al., 1997). This hypothesis has not,
however, been tested directly in the intact brain. Another
characteristic of spines, necessarily neglected in postmortem
morphologic studies, is their motility, which is believed, in part, to
represent postsynaptic participation in synapse formation (Dailey and
Smith, 1996). Spine motility has been shown to be sensitive to sensory
deprivation (Lendvai et al., 2000) and is greatest early in postnatal
life (Dunaevsky et al., 1999; Lendvai et al., 2000). Because an
abnormality in spine motility early in postnatal life could affect
synaptogenesis, it is of interest to determine whether the absence of
FMRP affects normal spine motility. Here we show that dendritic spines
in the intact brains of FMR1 knock-out mice are abnormal
early in postnatal life. These abnormalities in the somatosensory
cortex are most pronounced during the period of greatest synaptogenesis
in that region (White et al., 1997) and subside largely by the end of the first postnatal month. Spines in comparable neurons maintained in vitro showed no such abnormalities and were normally
motile. These observations support the notion that FMRP plays a role in synaptogenesis in the intact brain.
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MATERIALS AND METHODS |
Male FMR1 knock-out mice of the FVB strain and
wild-type (wt) littermate controls were used for the present study.
Genotypes were determined by PCR analysis of DNA extracted from tail
samples taken before perfusion or culture preparation. The primers used were the same as those outlined in the original article describing these animals (Dutch-Belgian Fragile X Consortium, 1994). All experimental protocols were conducted according to National Institutes of Health guidelines for animal research and were approved by the
Institutional Animal Care and Use Committee at Cold Spring Harbor Laboratory.
Mice (6, 13, and 27 d of age; n = 5 for each
group) were administered intracortical injections of enhanced green
fluorescent protein (EGFP) in a replication-deficient Sindbis
expression vector (Invitrogen, San Diego, CA). Briefly, mice were
anesthetized with a mixture of ketamine and xylazine (22 and 15 mg/kg,
i.m., respectively). A small burr hole was drilled in the skull
overlying the posteromedial barrel field using stereotactic coordinates
determined previously for each age on the basis of cytochrome oxidase
staining, and 50-100 nl of virion-rich supernatant was delivered using
a Picospritzer (General Valve Co, Fairfield, NJ) at a depth of 300-500
µm. Injection volume was titrated to yield sufficient numbers of well
separated neurons within the injected barrel and its neighbors (Chen et al., 2000). The number of neurons labeled with comparable injections decreased with postnatal age as described previously (Chen et al.,
2000). The skin was closed and the animal was returned to its cage.
After 24 hr, the animal was anesthetized again and perfused transcardially with cold 4% paraformaldehyde. The brain was removed and post-fixed for an additional 6-12 hr. The brain was blocked and
cut into 200-µm-thick sections in the coronal plane with a Vibratome
(Technical Products International Inc., St Louis, MO). Labeled
neurons were identified and imaged using a custom-built two-photon
laser scanning microscope (2PLSM) based on an Olympus Fluoview laser
scanning microscope (Olympus America, Inc., Melville, NY). The
light source was a Ti-Sapphire laser (Tsunami; SpectraPhysics, Mountain View, CA) running at a wavelength of ~910 nm (repetition frequency, 80 MHz; pulse length, 150 fsec). The average power delivered to the backfocal plane of the objective (40×; numerical aperature, 0.8) varied depending on the imaging depth (range, 30-150 mW). Laser power was adjusted so that additional power failed
to reveal previously undetected protrusions. Fluorescence was detected
in whole-field detection mode with a photomultiplier tube (Hamamatsu
Corp., Bridgewater, NJ). Dendritic segments of different orders were
sampled in a systematic random manner throughout the dendritic tree.
For in vitro studies, neocortical slice cultures were
prepared from 2-d-old mice (n = 5 animals for each
group) according to the method of Stoppini et al. (1991). Slices
containing the posteromedial barrel field were selected. The EGFP gene
was introduced using biolistic gene transfer (Lo et al., 1994)
(Bio-Rad, Hercules, CA) after 3 d in vitro (DIV), and
layer V neurons were imaged 2 d later (7 d from the date of
birth). Slice cultures were placed into a chamber and were continually
perfused with oxygenated artificial CSF consisting of (in
mM): 119 NaCl, 2.5 KCl, 26.2 NaHCO3, 1.0 NaHPO4, 11 glucose, 2.5 CaCl2, and 1.3 MgCl2; cultures were maintained at 32°C.
Time-lapse imaging of systematically sampled dendritic segments
consisted of collecting image stacks every 2 min for a total of 22 min,
yielding 10 time intervals per segment.
Protrusions were analyzed using custom software written in Interactive
Data Language (Research Support Instruments, Inc., Boulder, CO). Image stacks consisting of 10-50 sections spaced 1 µm
apart were generated, and protrusion lengths were obtained by measuring
the projection of the protrusion from its tip to the point at which it
meets the dendritic shaft. When the identity of a protrusion was
unclear (for instance, whether it was a single spine, or two, or an
axonal profile), the individual z images were reviewed. No
effort was made to analyze spines emerging below or above the dendrite.
The reason for this is that the resolution of the imaging system, as
with all optical systems, was lower in the z direction than
in the x and y directions. Because only the
longest spines would reliably be detected along the optical axis, the
analysis was restricted to spines that could be detected in the
x- and y-axes. For spine density measurements,
the sampling unit was the individual dendritic segment imaged. As a
measure of motility, absolute length differences between subsequent
time points were calculated; we refer to the mean difference over 10 intervals as "mean motility" (Chen et al., 2000; Lendvai et al., 2000). The "range of motility" refers to the total excursion of the
spine over the imaging time period and is the difference between the
maximum and minimum lengths of the spine. "Proportion persistent" was defined as the proportion of the total number of protrusions that
were present throughout the 22 min imaging session (i.e., with a length
that never measured 0 µm). To test for interactions among postnatal
ages and genotype and to control for variability at different sampling
levels, spine lengths and densities in mutant and control groups were
analyzed using a nested ANOVA procedure (SAS Institute, Cary, NC). This
procedure was performed on second-order apical dendrites, which
represented the largest single dataset. Length distributions were
compared using the Kolmogorov-Smirnov two-sample test. For purposes of
illustration, however, values for each genotype were pooled. Throughout
the study, injections, perfusions, sectioning, imaging, and spine
measurements were performed blind to genotype.
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RESULTS |
One day after intracortical injection of the viral vector
containing the EGFP gene, neurons were well labeled. The tissue sections were reminiscent of Golgi stains, in that a small subset of
neurons was fully labeled (Fig.
1A,B). The dendrites of
mutant animals showed no gross abnormalities when compared with
controls. Analysis was restricted to layer V pyramidal neurons in the
posterior medial barrel field without evidence of truncated apical or
basal dendritic trees. Typically four to seven neurons were analyzed in
each animal.
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Figure 1.
Two-photon laser scanning micrographs showing
morphology of and spine development in EGFP-expressing layer V
pyramidal neurons in the somatosensory cortex of wild-type
(left) and mutant (right) mice
transfected in vivo before fixation. Note the presence
of long protrusions at 1 week, particularly in the mutant mice
(arrows). Scale bars: A, B, 50 µm;
C-H, 8 µm.
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Analysis of the morphologic properties of dendritic spines in mutant
and control animals during the first postnatal month revealed clear
developmental trends. Spine length changed during this period. At 1 week, the overall mean spine length was 20% greater than at 4 weeks.
Consistent with other accounts, spine density also changed, increasing
2.5-fold between 1 and 4 weeks in normal animals. These developmental
changes were evident to a different degree in the mutant animals as
well. The differences between the genotypes depended on the postnatal
age. At 1 week, the overall mean spine length was 28% greater in the
mutant animals than in the wild-type animals (p < 0.0001; n = 1806 and 2034 spines for wild-type and
mutant animals, respectively). This difference dropped to 10% at 2 weeks (n = 3452 and 3224 spines for wild-type and
mutant animals, respectively) and only 3% at 4 weeks
(n = 2202 and 3490 spines for wild-type and mutant
animals, respectively) (Fig.
2A-C). At 1 week, the
distribution of spine lengths differed significantly between the two
genotypes (Kolmogorov-Smirnov two-sample test; p < 0.001), with mutants having fewer short spines (0.5-1.5 µm) and more
medium-to-long spines (2.5-5 µm) (Figs. 1C,D,
2D). When categorized by dendritic order, the spines
at most dendritic orders were significantly longer in the mutant
animals compared with the wild-type animals, with the exception of
spines on tertiary basal dendrites, for which there was no significant
difference between the groups (Fig. 2G). At 2 weeks, the
difference between the length distributions in the two groups was
smaller but still highly significant (p < 0.001) (Fig. 2E). These differences were accounted
for almost completely by differences localized to secondary apical and
to primary and tertiary basal dendrites (Fig. 2H). At
1 month, there were no significant differences overall either in the
length distribution or at any dendritic orders except for tertiary
basal dendrites, for which spines in the mutant animals were
significantly shorter than in the wild-type animals (Fig. 2F,I). A nested ANOVA procedure performed on
second-order apical dendrites, the largest dataset, confirmed these
findings, showing a significant interaction between age and genotype
(p < 0.01) despite considerable cell-to-cell
variability within animals.
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Figure 2.
Developmental spine length changes in
FMR1 knock-out mice and littermate controls. A,
D, G, 1 week; B, E, H, 2 weeks; C, F,
I, 4 weeks. A-C, Mean spine length in mutant
and control mice. D-F, Cumulative frequency
distribution of spine lengths. G-I, Mean spine length
at different branching levels in the dendritic tree. In
D-F, black areas indicate wild type and
white areas indicate mutant. In
G-I, black bars indicate wild type and
gray bars indicate mutant. ap1-3,
Primary through tertiary apical dendrites; bas1-3,
primary through tertiary basal dendrites. Error bars indicate SEM.
*p < 0.05; **p < 0.001;
***p < 0.0001.
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Spine density also varied developmentally with genotype. At 1 week,
mean spine density in the mutant animals was 33% higher than in the
wild-type animals (n = 168 and 177 dendritic segments for wt and mutant animals, respectively), and this difference was
evident at all sampled dendritic levels (Fig.
3A,D). At 2 and 4 weeks there
was no significant difference in spine density (2 weeks,
n = 176 and 170 dendritic segments for wt and mutant animals, respectively; 4 weeks, n = 92 and 133 dendritic segments for wt and mutant animals, respectively) (Fig.
3B,C). Interestingly, in secondary apical dendrites, spine
density was elevated at 1 week, normal at 2 weeks, and then
significantly elevated again at 4 weeks (Fig. 3D-F).
Again, a nested ANOVA procedure performed on second-order apical
dendrites confirmed a highly significant interaction between age and
genotype (p < 0.001). The changes in spine
length and density over time in normal and mutant animals are
summarized in Figure 4.
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Figure 3.
Developmental spine density changes in
FMR1 knock-out mice and littermate controls. A,
D, 1 week; B, E, 2 weeks; C, F, 4 weeks. A-C, Mean spine density in mutant and control
mice. D-F, Mean spine density at different branching
levels in the dendritic tree. Means are shown only for primary and
secondary dendrites, because the number of tertiary dendrites sampled
was too inconsistent to provide reliable estimates. Black
bars, Wild type; gray bars, mutant.
ap1-2, Primary through tertiary apical
dendrites; bas1-2, primary through tertiary
basal dendrites. Error bars indicate SEM. *p < 0.05; **p < 0.001; ***p < 0.0001.
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Figure 4.
A, C, Summary of changes
in spine length (A) and density
(C) in wild-type mice during the first postnatal
month. n = 5 animals in each group. Error bars
indicate SEM. B, D, Summary of changes in spine length
(B) and density (D) in
mutant mice during the first postnatal month. The value for each
parameter is normalized to the wild-type value. Error bars are computed
as the sum of the SEM for wild-type and mutant animals.
n = 5 animals in each group.
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To study the morphologic properties of neurons in the absence of
sensory input, neocortical slice cultures were prepared from 2-d-old
mouse pups and analyzed at 1 week after the date of birth, corresponding to the time point showing the greatest differences in the
intact brain. Neurons labeled using biolistic gene transfer with EGFP
were analyzed for spine length, density, and motility. Layer V neurons
with typical pyramidal morphology were chosen for analysis (Fig.
5A-D). The morphologies of
these neurons were somewhat different from those transfected in the
intact brain, in that apical dendrites reached layer I less frequently.
Unlike the results described for neurons labeled in vivo
(Figs. 1-3), spine lengths and densities in this preparation did not
differ significantly between the two genotypes (Fig.
6A,C). In separate
experiments, hippocampal organotypic cultures (prepared at postnatal
day 7) were transfected either biolistically or by injection of the
same Sindbis viral vector used in the experiments in the intact brain and imaged after 7 DIV. No significant difference in spine density was
detected between neurons transfected with the two methods (virus,
0.72 ± 0.07; biolistic, 0.73 ± 0.035; p > 0.05), indicating that the transfection method was not responsible for
the lack of a morphologic phenotype in vitro.
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Figure 5.
Two-photon laser scanning micrographs of
biolistically transfected layer V neurons in neocortical organotypic
cultures at 1 week from the date of birth. A, C, Wild
type. B, D, Mutant. A, B,
Low-magnification projections showing pyramidal morphology. C,
D, High-magnification projections of dendritic segments. Note
the increased presence of long protrusions in this preparation. Scale
bar: A, B, 50 µm; C, D, 4 µm.
E, Spine motility in neocortical cultures imaged using
2PLSM at 1 week from the date of birth. Micrographs taken at 2 min
intervals show typical changes in spine morphology. One protrusion
(arrow) changes shape several times during this
interval, branching, extending, and retracting. Another
(arrowhead) undergoes changes in the morphology of its
head. Others change relatively little during this interval. This
example was taken from a wild-type animal. Numbers indicate time in
minutes. Scale bar, 2 µm.
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Figure 6.
Spine length and density in neocortical
cultures analyzed 1 week from the date of birth. A, Mean
spine length; n = 1754 and 1667 spines for wt and
mutant animals, respectively. B, Spine length at
different levels in the dendritic tree. C, Mean spine
density; n = 247 and 255 dendritic segments for wt
and mutant, respectively. D, Spine density at different
levels in the dendritic tree. E-G, Spine motility in
neocortical cultures analyzed 1 week from the date of birth.
E, Mean motility per 2 min time interval.
F, Mean length range over which spines vary over a 22 min time interval. G, Proportion-persistent spines for
wild-type and mutant animals. Black, Wild type;
gray, mutant. ap1-3, Primary through
tertiary apical dendrites; bas1-3, primary through
tertiary basal dendrites. Error bars indicate SEM.
*p < 0.05; **p < 0.001;
***p < 0.0001.
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Analysis of spine motility in neocortical cultures also showed no
significant differences between mutants and controls. Although protrusions were quite motile (Figs. 5E,
6E-G), the mean motility per 2 min time interval,
the rate of turnover, and the lengths over which spines ranged were not
significantly different in the mutant animals compared with the
wild-type animals (Fig. 6E-G).
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DISCUSSION |
Histologic studies of brains of patients with fragile X syndrome
are scarce and can be summarized by noting that the brains are grossly
normal, with apparently normal neuronal densities. The only reported
abnormalities have been a decrease in synapse length (Rudelli et al.,
1985) and abnormally long and thin dendritic spines (Rudelli et al.,
1985; Hinton et al., 1991; Irwin et al., 2001). There is good reason to
look at spines for neuropathology, because several mental retardation
disorders are characterized by abnormalities in dendritic spines
(Marin-Padilla, 1972; Purpura, 1975). It is therefore of great interest
that a Golgi study of the occipital cortex of FMR1 knock-out
mice at 16 weeks found the spines there to be ~20% longer and up to
50% more densely distributed than in wild-type controls (Comery et
al., 1997).
Our present findings confirm increases in spine length and density in
the knock-out mouse. However, we measured increased spine density only
transiently at 1 week, with no detectable differences at 2 and 4 weeks.
Similarly, increases in spine length were greatest at 1 week and were
barely detectable by 1 month of postnatal life, much less pronounced
than in the 16-week-old animals described previously (Comery et al.,
1997). Several explanations for this discrepancy are possible. First,
the previous study was in the occipital cortex rather than in primary
somatosensory cortex, and dendritic maturation may differ sufficiently
in these two cortical areas to account for this difference. However,
two studies of dendritic spine maturation in the rat showed a
comparable time course of dendritic spine development in these two
areas (Wise et al., 1979; Miller, 1981). Thus, it is unlikely that
regional differences would account for the discrepancies in spine
density between the two FMR1 studies.
Second, because transfection efficiency decreases with postnatal age
(Chen et al., 2000), the possibility exists that the decrease in the
magnitude of the phenotype is attributable to a selective
decrease of labeling of only certain cell populations (for instance,
neurons with abnormal spines). Although we have no reason to expect
such an effect, we cannot conclusively rule it out in the absence of
methods, such as electron microscopy, that do not depend on the
labeling of neuronal subpopulations. However, if the labeling of
abnormal cells decreased with age, we would expect the cell-to-cell
variability in mutant animals to be greater at 1 week than at 4 weeks.
In this study, there is a very slight increase in variability between 1 and 4 weeks (results of ANOVA; data not shown).
Third, there is evidence that spine density normally decreases after 1 month postnatally (Wise et al., 1979; Miller, 1981). Thus the results
of the previous study might reflect an abnormality during a later phase
of synapse pruning. Our methods effectively preclude our resolving this
issue, because viral infection efficiency is low after 6 weeks (Chen et
al., 2000). This explanation would support a continuing role for FMRP
in the production and maintenance of appropriate synaptic connections
beyond development. In fact, although spine density was normal on
average at 4 weeks, it was elevated selectively in second-order apical
dendrites, after having been normal at this dendritic level at 2 weeks.
This change could therefore represent the beginning of a trend
continuing into adulthood and might have evolved into the difference
detected in the 16-week-old animals. Finally, methodological
differences must be considered, because it is difficult to compare the
neuronal populations labeled with the Golgi stain with those labeled
virally as in the present study, or to compare neurons analyzed with an
eyepiece reticule with those imaged using laser scanning microscopy.
The use of optical rather than electron microscopic methods presents
certain advantages, including the ability to sample very large numbers
of spines in each animal in an efficient manner. But an important
limitation of optical studies is that the fluorescence intensity, and
by extension, the detectability, of small structures such as spines is
proportional to their volume. Thus the smallest spines may go
undetected using our method. Therefore, the greater spine density
observed in mutant animals at 1 week cannot be clearly dissociated from
differences in spine shape and length observed at the same time. At 2 and 4 weeks, density is apparently normal, whereas spine length remains
abnormal. However, the large variance in the density measurements might
still obscure an abnormality in spine density in more mature tissue. It
is important to stress that our study makes no effort to describe the
absolute lengths and densities of dendritic spines; such measurements
require other methods. The purpose was to detect differences between
genotypes and among postnatal ages. Nonetheless we note the close
agreement between the spine density measurements reported here at 4 weeks and those reported previously in a study using the Golgi method in the adult animal (Comery et al., 1997). In both of these studies, it
is understood that the density values obtained are underestimates of
the actual spine density.
During the first postnatal week, dendritic spines are sparsely
distributed and somewhat longer than in the adult (Dailey et al., 1996;
Fiala et al., 1998; Lendvai et al., 2000; the present study). Within
the next 3 weeks, they increase in density and become somewhat shorter,
with fewer very long protrusions. The developmental change in spine
density has been described previously (Juraska and Fifkova, 1979;
Juraska, 1982; Petit et al., 1988), but although it is commonly
accepted that spine length decreases with postnatal age, the present
study is to our knowledge the first systematic description of this
phenomenon. The normal general pattern of development is followed in
the FMR1 knock-out mice, but the increase in density is
stunted and the decrease in length is exaggerated by the abnormal
values early in postnatal life. Nevertheless, general synaptogenesis
and remodeling of spines does take place in these animals, and although
the abnormality is not grossly disruptive, its subtlety might point to
a role for FMRP in fine adjustments of synaptic connections.
The present study also demonstrates that pyramidal neurons in
vitro that are prepared from FMR1 knock-out mice do not
have spines with abnormal length, density, or motility, despite the abnormalities in spine length and density in the intact brain. The
absence of a comparable difference in 1-week-old neocortical cultures
invites several interpretations. First, the method of transfection in
the intact brain differed from that used in vitro. However,
no difference in spine density was found between neurons transfected
in vitro with the Sindbis viral construct and those transfected biolistically. In addition, even if the labeling method differed, this could not explain the differences observed between genotypes. Second, neurons in organotypic cultures 1 week after the
date of birth might not be comparable with neurons in the intact brain
1 week after the date of birth. Similarly, it is possible that our
inability to detect a difference between mutant and wild-type animals
can be attributed to the presence or absence of some factor in the
culture medium. Although neurons in organotypic cultures mature roughly
comparably to those in the intact brain (Caesar et al., 1989; Caeser
and Schuz, 1992; Muller et al., 1993), a more subtle effect remains a
possibility. Notably, a recent study in cultured hippocampal neurons
failed to detect differences in spine length between mutant and
wild-type animals, and instead found morphologic and
electrophysiological evidence for presynaptic defects (Braun and Segal,
2000). This finding suggests the unmasking of a defect in the initial
stages of synaptogenesis, which must take place in dissociated cultured
neurons after plating but might not be detected in organotypic
cultures, in which the vast majority of connections are already present
when the culture is prepared.
The absence of an abnormal phenotype in organotypic cultures prepared
from mutant mice, together with the strong developmental dependence of
the abnormality in the intact brain, suggests that FMRP might not
function simply to maintain dendritic spines within certain limits of
density and length; its function may be coordinated with normal
patterned activity. In this context, it is significant that FMRP
expression is increased by afferent stimulation (Todd and Mack, 2000)
and that there is evidence for local translation of FMRP in response to
metabotropic glutamate receptor stimulation (Weiler and Greenough,
1993; Weiler et al., 1997). The coordination of expression with
synaptic activity is consistent with a phenotype that is most severe
during synaptogenesis, as shown in the present study. This raises the
possibility that the function served by FMRP early in life is related
to the proper establishment of synaptic connections in response to
synaptic activity. If this process is disturbed by the absence of this
protein, this disruption could be reflected at that time by an
abnormal spine density and morphology, and later by cognitive defects
resulting from improperly established connections.
To our knowledge, this is the first suggestion of a link between a
developmentally restricted structural or functional abnormality and a
mental retardation syndrome. This observation may have implications for
genetic therapeutic approaches, because it would indicate that
intervention must be administered as early as possible in brain
development to avoid long-lasting abnormalities in brain function.
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FOOTNOTES |
Received Feb. 5, 2001; revised April 12, 2001; accepted April 19, 2001.
This work was supported by the FRAXA, Mather, and Pew
Foundations and by National Institutes of Health Grants AA05518 and NS38259. We thank Barry J. Burbach and Karen Greenwood for expert technical assistance, Jodi Koblentz for PCR analysis, Drs. Edward Stern
and Gerald Latter for statistical assistance, and Drs. Patrick R. Hof,
Kristen M. Harris, Karen Zito, and Joshua Trachtenberg for helpful discussion.
Correspondence should be addressed to Esther A. Nimchinsky, Howard
Hughes Medical Institute, Cold Spring Harbor Laboratory, One Bungtown
Road, Cold Spring Harbor, NY 11724. E-mail: nimchins{at}cshl.org.
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ABSTRACT
INTRODUCTION
