 |
 Previous Article | Next Article 
The Journal of Neuroscience, November 15, 2000, 20(22):8262-8268
Regulation of Spine Calcium Dynamics by Rapid Spine Motility
Ania
Majewska,
Ayumu
Tashiro, and
Rafael
Yuste
Department of Biological Sciences, Columbia University, New York,
New York 10027
 |
ABSTRACT |
Dendritic spines receive most excitatory inputs in the CNS and
compartmentalize calcium. Spines also undergo rapid morphological changes, although the function of this motility is still unclear. We
have investigated the effect of spine movement on spine calcium dynamics with two-photon photobleaching of enhanced green fluorescent protein and calcium imaging of action potential-elicited
transients in spines from layer 2/3 pyramidal neurons in mouse visual
cortex slices. The elongation or retraction of the spine neck during spine motility alters the diffusional coupling between spine and dendrite and significantly changes calcium decay kinetics in spines. Our results demonstrate that the spine's ability to compartmentalize calcium is constantly changing.
Key words:
GFP; imaging; two photon; photobleaching; LTP; neocortex
| |
INTRODUCTION |
As first predicted by Ramón y
Cajal (1891) , dendritic spines receive most synaptic inputs in the
mammalian CNS (Gray, 1959; Harris and Kater, 1994). Spines are
separated from their parent dendrites by a thin neck and
compartmentalize calcium during synaptic stimulation (Müller and
Connor, 1991; Yuste and Denk, 1995; Yuste et al., 2000). Calcium
compartmentalization in spines is likely to be functionally important,
because calcium mediates input-specific forms of synaptic plasticity
(Lynch et al., 1983; Malenka et al., 1989). Calcium decay kinetics in
spines is controlled by diffusion of calcium across the spine
neck and active removal of calcium from the spine cytoplasm (Majewska
et al., 2000a) as well as by calcium buffers endogenous to the
spine head. Therefore, the morphology of the spine neck and the
expression and regulation of calcium pumps and buffers control the
duration of calcium transients in spines.
Spines have been shown recently to be extremely motile on the timescale
of seconds (Fischer et al., 1998; Dunaevsky et al., 1999). The function
of spine motility is still unclear. Although new filopodia and spines
can appear after electrical stimulation (Engert and Bonhoeffer, 1999;
Maletic-Savatic et al., 1999), basal motility, such as that present in
the absence of stimulation, appears resilient to major changes in the
activity of the cell (Dunaevsky et al., 1999). Both spine and
filopodial motility declines with development (Dailey and Smith, 1996;
Ziv and Smith, 1996; Dunaevsky et al., 1999) and is thought to be
related to critical periods of synaptic rearrangements (Dunaevsky et
al., 1999; Lendvai et al., 2000). Finally, volatile anesthetics block
spine motility, suggesting that rapid motility may play a global role
in brain function (Kaech et al., 1999).
The finding that calcium decays in spines are regulated by calcium
diffusion through the spine neck (Majewska et al., 2000a) suggests that spine motility could alter this diffusional coupling and
potentially modify spine calcium decay kinetics. On the other hand,
calcium pumps at the spine (Majewska et al., 2000a) and endogenous calcium buffers also control decay kinetics, and the type
and amount of motility could be too small to produce significant changes in these dynamics.
We have explored the effect of spine motility on calcium dynamics by
the use of two-photon excitation to photobleach enhanced green
fluorescent protein (EGFP) and image calcium indicators in motile
spines from layer 2/3 pyramidal neurons in slices of mouse visual
cortex. We find that the diffusion of EGFP across the spine neck is
linearly correlated with the length of the spine neck and is altered
during motility. We also find that changes in spine neck length affect
the initial fast calcium decay in spines. Our findings show that spine
motility alters calcium decay kinetics and demonstrate, for the first
time, that spine motility has an effect on spine function. We
hypothesize that, by altering spine-dendrite coupling, rapid spine
motility serves to alter constantly spine calcium dynamics and
therefore continuously changes the spine's capacity as a calcium
compartment and, presumably, changes the implementation of local
learning rules.
| |
MATERIALS AND METHODS |
Slices and electrophysiology. All experiments were
performed in accordance with the National Institutes of Health Guide
for the Care and Use of Laboratory Animals (NIH publication no. 86-23; revised 1987). Cultured slices were prepared from the primary visual
cortex of postnatal day 0 (P0)-P1 C57 mice as described (Dunaevsky et
al., 1999), incubated for 2 weeks, and biolistically transfected with
cytomegalovirus-EGFP vector (Clontech, Palo Alto, CA). After 2-4 d,
slices were imaged in artificial CSF (ACSF) that contained (in
mM): 126 NaCl, 3 KCl, 2 CaCl2, 2 MgSO4, 1.1 NaH3PO4, 26 NaHCO3, and 10 dextrose, bubbled with 95%
O2 and 5% CO2. Acute
slices were made from primary visual cortex from P12 to P16 mice as
described (Peterlin et al., 2000). Layer 2/3 pyramidal neurons in V1
were selected under differential interference contrast, and
whole-cell recordings were made with an Axoclamp 2B (Axon Instruments,
Foster City, CA) amplifier. Borosilicate pipettes with an outer
diameter of 1.5 mm and an inner diameter of 0.86 mm were used. The
intracellular solution was (in mM): 5 NaCl, 10 KCl, 10 HEPES, 135 KMeSO4, 2.5-4 Mg-ATP, 0.3 Na-GTP, and 1-5 mM calcium green-1 (Molecular Probes,
Eugene, OR); resistances were 6-7 M . Electrophysiological signals
were digitized by the use of an analog-to-digital board and Superscope
(InstruNet; GW Instruments, Somerville, MA). Action potentials were
elicited by stimulating the cell extracellularly with a patch pipette
containing ACSF and positioned close to the base of the soma.
Stimulating currents were in the range of 0.5-1 mA and were elicited
with an IsoFlex stimulator (AMPI).
Two-photon imaging and photobleaching. Imaging and
photobleaching were done by the use of a custom-made two-photon
laser-scanning microscope (Majewska et al., 2000b) consisting of
a modified confocal microscope (Fluoview; Olympus, New Hyde Park, NY)
and a titanium-sapphire laser providing 130 fsec pulses at 75 MHz at
wavelengths of 740-850 nm (Mira; Coherent, Palo Alto, CA) pumped by a
solid-state source (Verdi; Coherent). A 60×, 0.9 numerical aperture
water immersion objective (LUMPlanFl/IR1; Olympus) was used.
Fluorescence was detected by the use of a photomultiplier tube
(HC125-02; Hamamatsu) in external, whole-area detection mode,
and images were acquired and analyzed with Fluoview (Olympus) software.
A Pockels cell (model 350-50; ConOptics, Danbury, CT) was used to
create brief (~3-20 msec) pulses of high-intensity illumination for
measurements of diffusional times across the spine neck and to block
the beam during times in the scan when data were not being collected.
Spines were chosen randomly from all areas of the cell in a 200 µm
radius from the soma including spines on basal, apical, and oblique
dendrites. In the current configuration our microscope has the
resolution (1/e radius) of 0.3 µm in the radial dimension and 1 µm
in the z dimension (point spread function measured with
subresolution beads). Images of spines were acquired at the highest
digital zoom (10×), resulting in a nominal spatial resolution of 30 pixels/µm and in a time resolution of 12.5 msec/point in line-scan
mode. For each imaged spine, a z-stack at intervals of 0.1 µm was taken to allow a careful reconstruction of spine morphology,
and photobleaching or calcium measurements were performed. After 5-30
min, the procedure was repeated on the same spine. To determine spine
motility indexes, time-lapse sequences were taken every 30-60 sec. At
each time point, five to nine focal planes 0.5-1 µm apart were
scanned; these were later projected into a single image. Because spine motility is temperature dependent, experiments were performed at room
temperature to allow all data to be collected before a spine changed
shape. Control measurements were taken at 37°C and yielded
correlations that agreed with the room temperature data (n = 3 spines for calcium; n = 10 for
GFP measurements); both sets of measurements were therefore pooled for
the correlation of the changes in neck length and changes in diffusion
time constants, whereas only room temperature data were used in the
correlation of neck length and time constants.
Analysis. Fluorescence signals were analyzed with Igor
(Wavemetrics, Lake Oswego, OR) as described previously (Majewska et al., 2000a). Two to four traces were averaged for analysis of  GFP, whereas three to seven traces were
averaged for measurements of calcium decay kinetics. Fluorescence
intensity was corrected for background fluorescence measured in an area
adjacent to the structure and presented either as the change in
fluorescence divided by the initial fluorescence recorded before the
stimulus was applied ( F/F) or as total
fluorescence divided by the initial fluorescence (F/Finitial). F/F and
F/Finitial decays were fitted to single or
multiple exponentials. Spine calcium decays were mostly double exponential, and the time constant of the first fast exponential was
determined by fixing the second slow time constant to that of the
parent dendrite. No difference was observed in traces analyzed without
assuming any time constant for the second decay phase (Majewska et al.,
2000a). In some cases in which the dendrite did not undergo a
significant calcium increase, the spine appeared to be single
exponential, and the time constant of this exponential was used.
Morphological measurements, image processing, and analysis were done
with custom-written macros using NIH Image as described (Dunaevsky et
al., 1999). Images were aligned to correct for drift in the
x-y plane. For calculations of the motility index, images were thresholded to a single level throughout the entire sequence. Images were then binarized, and the spines were outlined. The motility
index (Dunaevsky et al., 1999) is a measure of the space a spine
occupies as it moves over time. It is defined as the ratio of the
difference between accumulated and smallest areas occupied by the spine
divided by the average area of the spine, when the outlines of the
spine in a time-lapse recording were superimposed digitally:
|
|
Spine neck lengths were measured from z-stack images.
In most cases the spine necks were clearly visible and could be
followed from the spine to the dendrite. In some cases the spine necks were too dim to be seen. In these cases the position of the spine neck
was inferred from the shape of the spine and was taken to start at the
point in the spine head where the morphology deviated from a spherical
shape. It is possible in these cases that the true length of the spine
neck was underestimated if the spine neck was curved. To minimize the
error caused by angled spine necks, only spines in the same
z plane as the middle of the dendrite or spines with spine
necks that were clearly visible in a single z section were analyzed.
| |
RESULTS |
Changes in spine neck length during spine motility
We first characterized the effects of spine motility on the length
of the spine neck by using a custom-built two-photon microscope to
image layer 2/3 pyramidal neurons from mouse V1 cultured slices transfected with EGFP (Fig.
1A,B). Spines on
apical, oblique, and basal dendrites were imaged in time-lapse
sequences and found to be highly motile over periods of minutes. As
described (Fischer et al., 1998; Dunaevsky et al., 1999),
morphological changes were diverse and included morphing, growth and
retraction, development of filopodia, and appearance and disappearance
of spines. We analyzed the changes in spine neck lengths, finding large
changes during imaging sequences lasting 15 min (Fig. 1C).
Spine neck changes ranged from 20 to 250% of the initial length. The
average maximal changes in spine neck length were 80 ± 15% of
the starting length in spines (mean ± SEM for all data;
n = 15 from 2 neurons). We concluded that spine neck
length is constantly changing in spines.
View larger version (73K):
[in this window]
[in a new window]
|
Figure 1.
Spine neck lengths change during spine movement.
A, Two-photon image of two EGFP-transfected layer 2/3
pyramidal neurons. The pial surface is to the top right.
B, High-magnification image of the boxed
area in A. Note how dendritic spines are clearly
visible. C. Spine necks elongate and retract as they move. Analysis of
the spine neck changes in four spines followed for a 15 min period.
Scale bars: A, 50 µm; B, 1 µm.
|
|
Diffusional coupling between spine and dendrite depends on the
spine neck length
Diffusion across the spine neck can be modeled as diffusion
through a pipe. The rate of diffusional equilibration between spine and
dendrite should depend on morphological features such as spine neck
length, spine neck diameter, and spine head volume. The equation that
governs the relationship between morphological parameters and the
timescale of diffusion through a pipe and, under our assumptions,
between spine and dendrite is:
|
(1)
|
where l is the length of the spine neck, V
is the volume of the spine head, D is the diffusion
coefficient of the diffusing molecule, and r is the radius
of the spine neck. Because long-necked spines tend to have slower
diffusion of fluorescein dextran across the spine neck than do
short-necked spines (Svoboda et al., 1996), we focused on the length of
the spine neck and measured the diffusion rates of EGFP across the neck
in spines with necks of different lengths. For these experiments, EGFP
in a spine head was bleached by the use of a brief (3-20 msec;
approximate spine exposure of 500 µsec to 1 msec) pulse of
high-intensity illumination in line-scan mode, and the fluorescence
recovery caused by the diffusion of unbleached fluorophore (Axelrod et
al., 1976) from the dendrite was measured with lower intensity
excitation (Svoboda et al., 1996; Majewska et al., 2000a) (Fig.
2). We found that the EGFP fluorescence
recovery trace was well fitted with a single exponential with an
average of 220 ± 18 msec (n = 72). In
addition, short spines tended to have faster recoveries than did
long-necked spines (Fig. 2A,B). In fact, across all
spines there was a significant correlation between spine neck length
and the EGFP fluorescent recovery time constant
(GFP; p < 0.0001; correlation
coefficient = 0.53; n = 72; Fig. 2C).
This implies that spine neck length determines the diffusional coupling
between spine and dendrite. Although it is possible that spine neck
length correlates with another parameter such as neck diameter or the
viscosity or obstruction of the spine neck, such correlations were not
found in statistical studies using EM (Harris and Stevens, 1989).
Therefore, we think it likely that the relationship between spine neck
length and EGFP diffusion is causal.
View larger version (27K):
[in this window]
[in a new window]
|
Figure 2.
Diffusional coupling between spine and dendrite
depends on the length of the spine neck. A, B,
Measurements of diffusional coupling in short
(A)- and long (B)-necked
spines. A, Top, Fluorescence recovery
curve of the spine shown in the bottom panel, whose neck
length is 0.4 µm. The fitted monoexponential curve (thin
line) shows a GFP of 68 msec. B,
Top, Similar measurement from a long-necked (1.5 µm)
spine, with a GFP of 275 msec. C,
Relation of GFP and spine neck length. There is a good
correlation between these two variables. Scale bars, 1 µm.
|
|
Diffusional coupling between spine and dendrite changes as
spines move
After establishing that spine neck length is correlated with
diffusional coupling and that spine neck changed during spine motility,
we examined whether diffusional coupling is also altered during
motility. We first determined whether the bleaching procedure altered
spine motility because exposure to high-intensity light could cause
injury to the cell. For this purpose we measured the motility of spines
on a dendritic branch before and after a small percentage of these
spines was bleached. We used the motility index (Dunaevsky et al.,
1999) (MI; see Materials and Methods) to measure spine motility.
Bleached spines did not show a significant difference in the motility
index before (1.34 ± 0.33) and after (1.34 ± 0.11) the
bleaching procedure (n = 5; Fig.
3A). In addition, spines that had not undergone the bleaching procedure but were on the
same dendritic branch also did not significantly change their motility
after photobleaching (1.37 ± 0.26 before bleaching; 1.46 ± 0.16 after bleaching; n = 6; p = 0.74;
Fig. 3B). We concluded that the bleaching did not
alter spine motility on the bleached dendrite.
View larger version (15K):
[in this window]
[in a new window]
|
Figure 3.
Diffusional coupling between spine and dendrite is
altered as spines elongate or retract. A, B, Lack of
effect of the bleaching protocol on the spine motility of bleached
(A) or unbleached
(B) spines on the same dendrite. Each
panel shows the motility index of spines before
(squares) and after (circles) the
bleaching protocol was applied. There were no significant differences
in the average motility indexes. C, Relation between the
R GFP (change in fluorescent
recovery time constant) and the RL
(change in spine neck length). Notice the strong correlation between
the two parameters (p < 0.0001;
n = 38; linear regression; correlation
coefficient = 0.73) indicating that changes in the times for
diffusion of EGFP through the spine neck are tied to the changes in
spine neck length during motility.
|
|
We then compared the fluorescence recovery of EGFP in individual spines
at different time points, after the spines showed significant changes
in neck length. Observed changes in spine neck length ranged from 0 to
250%. To measure the change in spine neck length, we used the ratio
RL, where RL
is defined as the neck length at the current time point divided by the
neck length measured at the time point directly before it (typically 15 min). Similarly,
RGFP was
defined as GFP at the current time point divided by the GFP measured at the preceding
time point. Thus, a value of 1 for either of these ratios signifies no
change in either length or GFP. In fact, we
found a strong correlation between the change in neck length
(RL) of individual spines and the change
in GFP (p < 0.0001;
correlation coefficient = 0.73; n = 38 from 26 spines; Figs. 3C, 4). Most
spines that did not change neck length also did not change
GFP (Fig. 4A). This shows that the spine's diffusional coupling to the dendrite and therefore its ability to act as a separate biochemical compartment change during
motility.
View larger version (51K):
[in this window]
[in a new window]
|
Figure 4.
Changes in EGFP diffusion during movement in
individual spines. A-C, Measurements of diffusional
coupling in spines whose necks stayed the same length
(A), shortened (B), or grew
longer (C). Middle,
Right, Images of the spine at the first time point
(middle) and at the second time point
(right). White lines trace the
length of the neck. Left, The EGFP fluorescence recovery
traces. Solid traces correspond to the
recoveries obtained at the first time point (middle),
whereas stippled traces correspond to those obtained at
the second time point (right). The change in time
constants for the spines shown is as follows: from 130 to 135 msec
(A), from 380 to 170 msec
(B), and from 260 to 480 msec
(C). Scale bars, 1 µm.
|
|
The spine calcium decay kinetics is proportional to the spine
neck length
We then wondered whether changes in spine neck length during spine
movement translated to changes in the regulation of calcium decay
kinetics in spines. To answer this, we imaged calcium dynamics in
spines from neurons in acute slices filled with calcium green, using
the calcium transient elicited by backpropagating action potentials
(APs) (Yuste and Denk, 1995) as a stereotyped delta function to monitor
the timescales of calcium decays in spines during motility.
Spine motility is actin-based (Fischer et al., 1998; Dunaevsky et al.,
1999) and is blocked by prolonged perfusion of the cell cytoplasm
during whole-cell recording (Dunaevsky et al., 1999). To avoid
"washing out" spine motility, we patched pyramidal neurons with
high concentrations of dye (a bolus injection) and withdrew the pipette
within 30-90 sec of the break-in. This bolus injection (Helmchen et
al., 1996) allowed us to fill the cell with concentrations of dye
sufficient for the discrimination of spines and the imaging of calcium
transients while circumventing the need for sustained whole-cell
recording. Indeed, spines on control neurons that were whole-cell
recorded for prolonged periods of time with 200 µM
calcium green were almost completely stationary (MI = 0.7 ± 0.1; n = 7 spines from 2 cells), whereas spines on bolus-filled cells were highly motile (MI = 1.9 ± 0.2;
n = 10 spines from 2 cells).
After filling the neurons with the bolus injection and waiting 30 min
to allow the dye to equilibrate throughout the neuron, we stimulated
the neuron with a pipette placed close to the base of the soma. AP
firing was inferred from generalized calcium increases in spines and
dendrites (Yuste and Denk, 1995). In control experiments the pipette
was not withdrawn, and we confirmed that indeed the stimulation caused
the cell to fire single APs (n = 3).
We found that in response to single APs, spines from bolus-injected
cells had double-exponential decay kinetics (Majewska et al.,
2000a) (Fig. 5A) with a
fast first decay (fast = 298 ± 20 msec;
n = 22), intrinsic to the spine (Majewska et al.,
2000a), and a second decay with a slower timescale determined by
the parent dendrite (d = 590 ± 116 msec; n = 7). As expected (Majewska et al.,
2000a), the first fast decay scaled with the length of the spine
neck (p < 0.01; correlation coefficient = 0.66; n = 16; Fig. 5B). Thus, spines with
longer necks had slower calcium decay kinetics.
View larger version (21K):
[in this window]
[in a new window]
|
Figure 5.
Correlation between changes in the spine neck
length and changes in the fast calcium decay in spines.
A, Calcium dynamics in a spine after the firing of a
backpropagating action potential through an extracellular pipette
placed near the cell soma. The action potential elicits an increase in
calcium concentration in both dendrite and spine. The calcium decay in
the dendrite (stippled line) follows single-exponential
kinetics, whereas the decay in the spine (solid line)
shows the characteristic double-exponential kinetics (Majewska et al.,
2000a). The trace is an average of six trials,
filtered with a seven-point smoothing kernel. B,
Relation between spine fast (fast time constant of
decay) and spine neck length. There is a strong correlation between
these two variables (p < 0.01; correlation
coefficient = 0.66; n = 16). C,
Relation between Rfast and
RL. Note the strong correlation
between these parameters (p < 0.001;
n = 16; correlation coefficient = 0.83)
demonstrating that changes in spine neck length during spine movement
affect the kinetics of the calcium decay in spines.
|
|
Calcium decays in spines change during spine movement
The observed correlations predicted that the changes in spine neck
length occurring during spine movement would alter the first fast
decay. Indeed, we observed that the initial fast decay kinetics of
spines was not stable over time as compared with that of their parent
dendrites. Although dendritic decay time constants fluctuated by <10%
over time, spine kinetics changed by as much as 60% (SDs for
spine-dendrite pairs over a period of 30 min were 7 ± 1% for
dendrites vs 38 ± 5% for spines; n = 9 from 4 cells). Nevertheless, other factors besides the spine neck length can control spine decay kinetics. In particular, spine decay kinetics is
also regulated by spine calcium pumps (Majewska et al., 2000a), as well as other morphological factors such as the neck diameter or
blocking of the neck by the spine apparatus (Gray, 1959), which could
potentially have a large effect on spine calcium decay kinetics.
To explore whether changes in spine neck length produce corresponding
changes in spine calcium decays, we performed sequential calcium
imaging from spines for up to 30 min and correlated their fast decay
kinetics with simultaneous measurements of their spine neck length (10 spines from 6 neurons). Most of the imaged spines retracted, and only
one spine elongated. It is possible that we preselected for longer
spine necks as these are easier to distinguish with calcium
green-filled neurons that are dimmer than GFP-transfected cells. We
found a strong correlation between the change in calcium decay kinetics
(Rfast) and
the change in the length of the spine neck
(RL; p < 0.005;
correlation coefficient = 0.83; n = 16; Fig.
5C). The linear fit through the data yielded a slope close
to 1, as expected (slope = 1.04, linear regression; see Eq. 1).
Changes in calcium decay kinetics with motility were also evident in
individual spines, with decays becoming faster as spines retracted and
becoming slower as spine necks lengthened (Fig. 6A). In a spine in
which five time points were available, the correspondence between
changes in decay kinetics and changes in neck length was also
significant (p < 0.005; n = 5;
Fig. 6B). We concluded that changes in the spine neck
length because of rapid spine motility affect calcium decay
kinetics.
View larger version (29K):
[in this window]
[in a new window]
|
Figure 6.
Changes in the fast calcium decay component during
spine movement in individual spines. A, Example of the
calcium dynamics in a spine in response to an AP. Two calcium decays
from the same spine measured at different time points
(top) are shown. The traces have been
normalized to the same amplitude for ease of comparison. The
solid line shows the decay that corresponds to the
bottom left image of the spine. The stippled
line corresponds to the bottom right image when
the spine had retracted and the spine neck had become shorter. Notice
the faster initial decay of the stippled trace. The
second slow decay phase is similar in the two spines because of the
slower and less variable dendritic decay. B, Similar
analysis from a spine that did not change its neck length. Note how the
calcium decay kinetics is very similar. C, Relation
between Rfast and
RL for a spine in which five time
points were available. The correlation is highly significant
(p < 0.005; n = 5).
Scale bars, 1 µm.
|
|
| |
DISCUSSION |
Spine motility regulates spine calcium compartmentalization
Morphological changes in spine shape in animals subjected to many
different experimental paradigms have been well documented (Valverde,
1967; Purpura, 1974; Fifkova and Anderson, 1981; Brandon and Coss,
1982), including changes in the length of the spine neck (Fifkova and
Anderson, 1981; Brandon and Coss, 1982). In 1977, actin was purified in
the postsynaptic density and was proposed to underlie spine
motility (Blomberg et al., 1977). A similar proposal was later made by
Crick, who predicted the existence of an actin-based "twitching" of
spines (Crick, 1982). Indeed, rapid motility of filopodia (Dailey and
Smith, 1996; Ziv and Smith, 1996) and spines (Fischer et al., 1998;
Dunaevsky et al., 1999) has been found recently throughout the CNS.
Spine motility changes during critical periods for monocular
deprivation in mouse area V1 (Dunaevsky et al., 1999) and can be
regulated by sensory deafferentation (Lendvai et al., 2000), and new
protrusions can grow in response to synaptic stimulation (Engert and
Bonhoeffer, 1999; Maletic-Savatic et al., 1999). This suggests that
spine motility is related to critical period plasticity. Nevertheless,
spine motility persists in the absence of synaptic stimulation
(Dunaevsky et al., 1999) and appears intrinsic to the neuron (Dunaevsky
et al., 1999). Thus, spine motility remains an orphan phenomenon
without a clearly demonstrated function.
Although some studies suggest that motility may allow spines to find
synaptic partners (Dailey and Smith, 1996; Ziv and Smith, 1996), we
show here, for the first time, that spine motility has an effect on the
spine's function as a calcium compartment: changes in spine neck
length that occur during motility produce changes in calcium decay
kinetics. This is important because the main specific function of
dendritic spines could be to compartmentalize calcium and implement
local learning rules (Müller and Connor, 1991; Yuste and Denk,
1995; Yuste et al., 2000). We concentrated on the study of spine neck
length although other morphological parameters such as spine head
volume and spine neck radius that might be altered during motility
could also have an effect on spine function. By measuring the diffusion
of EGFP across the spine neck in transfected neurons, we find that as
spines retract and shorten their spine necks, diffusion between the
spine and dendrite becomes faster. Conversely, when spines elongate,
diffusion between the two structures becomes slower. By measuring
AP-induced calcium transients in neurons injected with calcium green-1,
we determine whether calcium kinetics is also affected during motility. We show that as spines retract they maintain high calcium
concentrations for shorter periods of time as indicated by their faster
initial decays and become less able to serve as calcium compartments. Therefore, as spines move, their ability to act biochemically independently of the dendrite is altered.
Interestingly, small changes in neck length can cause large changes in
the decay kinetics of spines, especially if the initial length of the
spine neck is small. Stubby spines, which were undersampled in this
study because of their small size and proximity to the dendrite, could
experience major changes in calcium dynamics after minuscule changes in
neck length. In fact recent studies have shown that spine categories
such as stubbies, thin and mushroom spines, and filopodia are not
stable and that considerable conversions between these categories take
place (Parnass et al., 2000). This suggests that changes in decay
kinetics are more profound than those documented here, especially in
the case in which stubby spines change into thin or mushroom spines.
Comparisons with previous studies
Previously Svoboda et al. (1996) studied the diffusion of
fluorescein dextrans between spine and dendrite by bleaching the dye or
uncaging it in spines. In a statistical population, the authors found
that diffusion time increased in longer necked spines although direct
correlations between neck length and diffusion times were not
presented. Our study documents changing diffusion times as the neck
lengths change during single-spine elongation and retraction. Although
the diffusion coefficients of fluorescein dextran and EGFP in cytoplasm
are similar (Popov and Poo, 1992; Swaminathan et al., 1997), we
find diffusion times a factor of two to three larger than that reported
in the previous study, even in populations of spines with similar neck
lengths. This could be caused by differences in the preparation used or
by differences in spine geometriessuch as spine neck diameter or
spine head volume. Also, the extensive whole-cell recordings necessary
to introduce fluorescein dextran into the neurons could have reduced the neurons' viscosity.
As opposed to fluorescein dextran and EGFP diffusion, the dynamics of
calcium in neurons is subject to complicated regulation that is still
not well understood (Neher, 1998). Our study builds on our previous
work that has described heterogeneity in the calcium decay dynamics in
spines (Majewska et al., 2000a). Here we show that the static
heterogeneity described in our previous study is a dynamic process in
which spines do not maintain their calcium kinetics but change between
the various extremes that were described. There are important
methodological differences between our studies. Our first study was
performed in rat CA1, whereas this study is done in mouse V1. Also our
first study used prolonged whole-cell recordings, whereas in this work
we use brief whole-cell recordings. This method does not wash out the
cellular contents as profoundly as prolonged whole-cell recording, and
therefore we expect that most of the spine's mobile calcium buffers
remain intact. In this present study, we again find that the spine
decay kinetics is double exponential. This argues that our previous
work describing the mechanisms of the decay kinetics is not seriously
affected by the potential washout of endogenous buffers.
Other factors may regulate spine calcium decays
Spine calcium decays are shaped by many factors such as calcium
extrusion mechanisms that contribute to both phases of the decay
(Majewska et al., 2000a). Extrusion rates may be regulated in
time either by inactivation mechanisms (Wang et al., 1991) or by
increased expression, and these processes may be calcium dependent
(Guerini et al., 1999). Endogenous calcium buffers also shape calcium
kinetics by slowing down extrusion-based timescales (Tank et al., 1995)
and altering diffusional ones depending on their mobility (Gabso et
al., 1997). Endogenous buffers in pyramidal neurons have been shown to
be relatively immobile and appear to have low buffer capacities and
affinities (Helmchen et al., 1996), suggesting that they slow diffusion
and do not saturate unless trains of stimuli are presented. We find no
changes in extrusion or buffering mechanisms at the timescales we have
explored although it is possible that longer timescales or different
stimulation paradigms could reveal this regulation.
Mechanisms of changes in spine neck length
The major effect of changes in spine neck length on spine
biochemical compartmentalization draws attention to the molecular mechanisms that might control this process and their regulation. Indeed, we have found evidence recently of the involvement of the Rho
family of small GTPases in the control of the spine neck length
(Tashiro et al., 2000). The Rho family is implicated in the regulation
of the actin cytoskeleton in a variety of cell types, including neurons
(Hall, 1994; Luo et al., 1996; Threadgill et al., 1997). In our study
in hippocampal pyramidal neurons overexpression of a constitutively
active form of Rho significantly reduced the length of the spine neck.
At the same time, expression of C3 transferase, which is analogous to a
dominant-negative Rho, significantly increased the spine neck length.
Therefore, the ability of the spine to compartmentalize calcium is
directly regulated by this GTPase. It thus becomes interesting to
explore the regulation of the expression and activity of Rho in
neurons, as well as the effects of this regulation on the function of
the circuit and behavior of the animal.
Implications for synaptic function and plasticity
The modulation of calcium decay kinetics that we observe could
have a large impact on the function of the synapse. Spines appear to be
ideally built to compartmentalize calcium (Wickens, 1988; Holmes, 1990;
Koch and Zador, 1993; Yuste et al., 2000), and the regulation of
[Ca2+]i dynamics
in spines could underlie synapse-specific learning rules (Levy and
Steward, 1979; Gustafsson and Wigstrom, 1986; Lisman, 1989). The
changes in kinetics that we measure can be several fold in 30 min.
Therefore major changes in the integrated [Ca2+]i
experienced by a spine occur as a consequence of its motility. This
implies that the same paradigm that elicits a type of synaptic plasticity may not be effective in the same synapse a few minutes later. We would predict that synapses on spines are constantly changing
the time window for their calcium-dependent learning rules. This effect
would not occur on synapses made directly on the dendritic shaft,
frequent on interneurons (Peters et al., 1976). The purpose of this
fluid change in spine time constants is enigmatic, but it may provide a
highly adaptive mechanism to ensure flexible temporal integration of signals.
| |
FOOTNOTES |
Received July 18, 2000; revised Aug. 28, 2000; accepted Aug. 29, 2000.
This work was funded by the National Eye Institute Grant EY 111787 and
the Human Frontier Science Program. We thank E. Brown, A. Dunaevsky, J. Goldberg, K. Holthoff, and C. Mason for comments.
Correspondence should be addressed to Dr. Ania Majewska, Department of
Biological Sciences, Columbia University, 1212 Amsterdam Avenue, Box
2435, New York, NY 10027. E-mail: akm21{at}columbia.edu.
| |
REFERENCES |
-
Axelrod D,
Koppel D,
Schlessinger J,
Elson E,
Webb W
(1976)
Mobility measurement by analysis of fluorescence photobleaching recovery kinetics.
Biophys J
16:1055-1069[Web of Science][Medline].
-
Blomberg F,
Cohen R,
Siekevitz P
(1977)
The structure of postsynaptic densities isolated from dog cerebral cortex. II. Characterization and arrangement of some of the major proteins within the structure.
J Cell Biol
86:831-845[Abstract/Free Full Text].
-
Brandon J,
Coss R
(1982)
Rapid dendritic spine stem shortening during one-trial learning: the honeybee's first orientation flight.
Brain Res
252:51-61[Web of Science][Medline].
-
Crick F
(1982)
Do spines twitch?
Trends Neurosci
5:44-46.
-
Dailey ME,
Smith SJ
(1996)
The dynamics of dendritic structure in developing hippocampal slices.
J Neurosci
16:2983-2994[Abstract/Free Full Text].
-
Dunaevsky A,
Tashiro A,
Majewska A,
Mason CA,
Yuste R
(1999)
Developmental regulation of spine motility in mammalian CNS.
Proc Natl Acad Sci USA
96:13438-13443[Abstract/Free Full Text].
-
Engert F,
Bonhoeffer T
(1999)
Dendritic spine changes associated with hippocampal long-term synaptic plasticity.
Nature
399:66-70[Medline].
-
Fifkova E,
Anderson CL
(1981)
Stimulation-induced changes in dimensions of stalks of dendritic spines in the dentate molecular layer.
Exp Neurol
74:621-627[Web of Science][Medline].
-
Fischer M,
Kaech S,
Knutti D,
Matus A
(1998)
Rapid actin-based plasticity in dendritic spine.
Neuron
20:847-854[Web of Science][Medline].
-
Gabso M,
Neher E,
Spira ME
(1997)
Low mobility of Ca2+ buffers in axons of cultured Aplysia neurons.
Neuron
18:473-481[Web of Science][Medline].
-
Gray EG
(1959)
Electron microscopy of synaptic contacts on dendritic spines of the cerebral cortex.
Nature
183:1592-1594[Medline].
-
Guerini D,
Garcia-Martin E,
Gerber A,
Volbracht C,
Leist M,
Merino C,
Carafoli E
(1999)
The expression of plasma membrane Ca2+ pump isoforms in cerebellar granule neurons is modulated by Ca2+.
J Biol Chem
274:1667-1676[Abstract/Free Full Text].
-
Gustafsson B,
Wigstrom H
(1986)
Hippocampal long-lasting potentiation produced by pairing single volleys and brief conditioning tetani evoked in separate afferents.
J Neurosci
6:1575-1582[Abstract].
-
Hall A
(1994)
Small GTP-binding proteins and the regulation of the actin cytoskeleton.
Annu Rev Cell Biol
10:31-54[Web of Science].
-
Harris KM,
Kater SB
(1994)
Dendritic spines: cellular specializations imparting both stability and flexibility to synaptic function.
Annu Rev Neurosci
17:341-371[Web of Science][Medline].
-
Harris KM,
Stevens JK
(1989)
Dendritic spines of CA1 pyramidal cells in the rat hippocampus: serial electron microscopy with reference to their biophysical characteristics.
J Neurosci
9:2982-2997[Abstract].
-
Helmchen F,
Imoto K,
Sakmann B
(1996)
Ca2+ buffering and action potential-evoked Ca2+ signaling in dendrites of pyramidal neurons.
Biophys J
70:1069-1081[Web of Science][Medline].
-
Holmes W
(1990)
Is the function of dendritic spines to concentrate calcium?
Brain Res
519:338-342[Web of Science][Medline].
-
Kaech S,
Brinkhas H,
Matus A
(1999)
Volatile anesthetics block actin-based motility in dendritic spines.
Proc Natl Acad Sci USA
96:10433-10437[Abstract/Free Full Text].
-
Koch C,
Zador A
(1993)
The function of dendritic spinesdevices subserving biochemical rather than electrical compartmentalization.
J Neurosci
13:413-422[Web of Science][Medline].
-
Lendvai B,
Stern E,
Chen B,
Svoboda K
(2000)
Experience-dependent plasticity of dendritic spines in the developing rat barrel cortex in vivo.
Nature
404:876-881[Medline].
-
Levy WB,
Steward O
(1979)
Synapses as associative memory elements in the hippocampal formation.
Brain Res
175:233-245[Web of Science][Medline].
-
Lisman J
(1989)
A mechanism for the Hebb and anti-Hebb processes underlying learning and memory.
Proc Natl Acad Sci USA
86:9574-9578[Abstract/Free Full Text].
-
Luo L,
Hensch T,
Ackerman L,
Barbel S,
Jan L,
Jan YN
(1996)
Differential effects of the Rac GTPase on Purkinje cell axons and dendritic trunks and spines.
Nature
379:837-840[Medline].
-
Lynch G,
Larson J,
Kelso S,
Barrionuevo G,
Schottler F
(1983)
Intracellular injections of EGTA block induction of hippocampal long-term potentiation.
Nature
305:719-721[Medline].
-
Majewska A,
Brown E,
Ross J,
Yuste R
(2000a)
Mechanisms of calcium decay kinetics in hippocampal spines: role of spine calcium pumps and calcium diffusion through the spine neck in biochemical compartmentalization.
J Neurosci
20:1722-1734[Abstract/Free Full Text].
-
Majewska A, Yiu G, Yuste R (2000b) A custom-made
two-photon microscope and deconvolution system. Pflügers Arch, in
press.
-
Malenka RC,
Kauer JA,
Perkel DJ,
Nicoll RA
(1989)
The impact of postsynaptic calcium on synaptic transmissionits role in long term potentiation.
Trends Neurosci
12:444-450[Web of Science][Medline].
-
Maletic-Savatic M,
Malinow R,
Svoboda K
(1999)
Rapid dendritic morphogenesis in CA1 hippocampal dendrites induced by synaptic activity.
Science
283:1923-1927[Abstract/Free Full Text].
-
Müller W,
Connor JA
(1991)
Dendritic spines as individual neuronal compartments for synaptic Ca2+ responses.
Nature
354:73-76[Medline].
-
Neher E
(1998)
Usefulness and limitations of linear approximations to the understanding of Ca2+ signals.
Cell Calcium
24:345-357[Web of Science][Medline].
-
Parnass Z, Tashiro A, Yuste R (2000) Analysis of spine
morphological plasticity in developing hippocampal pyramidal neurons.
Hippocampus, in press.
-
Peterlin ZA,
Kosloski J,
Mao B,
Tsiola A,
Yuste R
(2000)
Optical probing of neuronal circuits with calcium indicators.
Proc Natl Acad Sci USA
97:3619-3624[Abstract/Free Full Text].
-
Peters A,
Paley SL,
Webster HdF
(1976)
In: The fine structure of the nervous system. Philadelphia: Saunders.
-
Popov SV,
Poo MM
(1992)
Diffusional transport of macromolecules in developing nerve processes.
J Neurosci
12:77-85[Abstract].
-
Purpura D
(1974)
Dendritic spine "dysgenesis" and mental retardation.
Science
186:1126-1128[Abstract/Free Full Text].
-
Ramón y Cajal S
(1891)
In: Significación fisiológica de las expansiones protoplásmicas y nerviosas de la sustancia gris. Congreso médico valenciano. June 24.
-
Svoboda K,
Tank DW,
Denk W
(1996)
Direct measurement of coupling between dendritic spines and shafts.
Science
272:716-719[Abstract].
-
Swaminathan R,
Hoang CP,
Verkman AS
(1997)
Photobleaching recovery and anisotropy decay of green fluorescent protein GFP-S65T in solution and cells: cytopastic viscosity probed by green fluorescent protein and rotational diffusion.
Biophys J
72:1900-1907[Web of Science][Medline].
-
Tank DW,
Regehr WG,
Delaney KD
(1995)
A quantitative analysis of presynaptic calcium dynamics that contribute to short-term enhancement.
J Neurosci
15:7940-7952[Abstract].
-
Tashiro A,
Minden A,
Yuste R
(2000)
Regulation of dendritic spine morphology by the Rho family of small GTPases: antagonistic roles of Rac and Rho.
Cereb Cortex
10:927-938[Abstract/Free Full Text].
-
Threadgill R,
Bobb K,
Ghosh A
(1997)
Regulation of dendritic growth and remodeling by Rho, Rac, and Cdc42.
Neuron
19:625-634[Web of Science][Medline].
-
Valverde F
(1967)
Apical dendritic spines of the visual cortex and light deprivation in the mouse.
Exp Brain Res
3:337-352[Web of Science][Medline].
-
Wang KK,
Du YS,
Diglio C,
Tsang W,
Kuo TH
(1991)
Hormone-induced phosphorylation of the plasma membrane calcium pump in cultured aortic endothelial cells.
Arch Biochem Biophys
15:103-108.
-
Wickens J
(1988)
Electrically coupled but chemically isolated synapses: dendritic spines and calcium in a rule for synaptic modification.
Prog Neurobiol
31:507-528[Web of Science][Medline].
-
Yuste R,
Denk W
(1995)
Dendritic spines as basic units of synaptic integration.
Nature
375:682-684[Medline].
-
Yuste R,
Majewska A,
Holthoff K
(2000)
From form to function: calcium compartmentalization in dendritic spines.
Nat Neurosci
3:653-659[Web of Science][Medline].
-
Ziv N,
Smith S
(1996)
Evidence for a role of dendritic filopodia in synaptogenesis and spine formation.
Neuron
17:91-102[Web of Science][Medline].
Copyright © 2000 Society for Neuroscience 0270-6474/00/20228262-07$05.00/0
This article has been cited by other articles:
|
|
|
|
 
B. N. Routh, D. Johnston, K. Harris, and R. A. Chitwood
Anatomical and Electrophysiological Comparison of CA1 Pyramidal Neurons of the Rat and Mouse
J Neurophysiol,
October 1, 2009;
102(4):
2288 - 2302.
[Abstract]
[Full Text]
[PDF]
|
|
|
|
|
|
|
J. H. Goldberg and R. Yuste
Two-Photon Calcium Imaging of Spines and Dendrites
CSH Protocols,
June 1, 2009;
2009(6):
pdb.prot5231 - pdb.prot5231.
[Abstract]
[Full Text]
|
|
|
|
|
|
|
A. Grunditz, N. Holbro, L. Tian, Y. Zuo, and T. G. Oertner
Spine Neck Plasticity Controls Postsynaptic Calcium Signals through Electrical Compartmentalization
J. Neurosci.,
December 10, 2008;
28(50):
13457 - 13466.
[Abstract]
[Full Text]
[PDF]
|
|
|
|
|
|
|
P. J. Sjostrom, E. A. Rancz, A. Roth, and M. Hausser
Dendritic Excitability and Synaptic Plasticity
Physiol Rev,
April 1, 2008;
88(2):
769 - 840.
[Abstract]
[Full Text]
[PDF]
|
|
|
|
|
|
|
T. L. Spires-Jones, M. Meyer-Luehmann, J. D. Osetek, P. B. Jones, E. A. Stern, B. J. Bacskai, and B. T. Hyman
Impaired Spine Stability Underlies Plaque-Related Spine Loss in an Alzheimer's Disease Mouse Model
Am. J. Pathol.,
October 1, 2007;
171(4):
1304 - 1311.
[Abstract]
[Full Text]
[PDF]
|
|
|
|
|
|
|
J. M. Mateos, A. Luthi, N. Savic, B. Stierli, P. Streit, B. H. Gahwiler, and R. A. McKinney
Synaptic modifications at the CA3 CA1 synapse after chronic AMPA receptor blockade in rat hippocampal slices
J. Physiol.,
May 15, 2007;
581(1):
129 - 138.
[Abstract]
[Full Text]
[PDF]
|
|
|
|
|
|
|
A. Lorincz, B. Rozsa, G. Katona, E. S. Vizi, and G. Tamas
Differential distribution of NCX1 contributes to spine-dendrite compartmentalization in CA1 pyramidal cells
PNAS,
January 16, 2007;
104(3):
1033 - 1038.
[Abstract]
[Full Text]
[PDF]
|
|
|
|
|
|
|
R. Araya, K. B. Eisenthal, and R. Yuste
Dendritic spines linearize the summation of excitatory potentials
PNAS,
December 5, 2006;
103(49):
18799 - 18804.
[Abstract]
[Full Text]
[PDF]
|
|
|
|
|
|
|
R. Araya, J. Jiang, K. B. Eisenthal, and R. Yuste
The spine neck filters membrane potentials
PNAS,
November 21, 2006;
103(47):
17961 - 17966.
[Abstract]
[Full Text]
[PDF]
|
|
|
|
|
|
|
V. Scheuss, R. Yasuda, A. Sobczyk, and K. Svoboda
Nonlinear [Ca2+] Signaling in Dendrites and Spines Caused by Activity-Dependent Depression of Ca2+ Extrusion
J. Neurosci.,
August 2, 2006;
26(31):
8183 - 8194.
[Abstract]
[Full Text]
[PDF]
|
|
|
|
|
|
|
T. Kuriu, A. Inoue, H. Bito, K. Sobue, and S. Okabe
Differential control of postsynaptic density scaffolds via actin-dependent and -independent mechanisms.
J. Neurosci.,
July 19, 2006;
26(29):
7693 - 7706.
[Abstract]
[Full Text]
[PDF]
|
|
|
|
|
|
|
S. Oray, A. Majewska, and M. Sur
Effects of Synaptic Activity on Dendritic Spine Motility of Developing Cortical Layer V Pyramidal Neurons
Cereb Cortex,
May 1, 2006;
16(5):
730 - 741.
[Abstract]
[Full Text]
[PDF]
|
|
|
|
|
|
|
A. K. Majewska, J. R. Newton, and M. Sur
Remodeling of synaptic structure in sensory cortical areas in vivo.
J. Neurosci.,
March 15, 2006;
26(11):
3021 - 3029.
[Abstract]
[Full Text]
[PDF]
|
|
|
|
|
|
|
M. Nuriya, J. Jiang, B. Nemet, K. B. Eisenthal, and R. Yuste
Imaging membrane potential in dendritic spines
PNAS,
January 17, 2006;
103(3):
786 - 790.
[Abstract]
[Full Text]
[PDF]
|
|
|
|
|
|
|
M. Zagrebelsky, A. Holz, G. Dechant, Y.-A. Barde, T. Bonhoeffer, and M. Korte
The p75 Neurotrophin Receptor Negatively Modulates Dendrite Complexity and Spine Density in Hippocampal Neurons
J. Neurosci.,
October 26, 2005;
25(43):
9989 - 9999.
[Abstract]
[Full Text]
[PDF]
|
|
|
|
|
|
|
A. Sobczyk, V. Scheuss, and K. Svoboda
NMDA Receptor Subunit-Dependent [Ca2+] Signaling in Individual Hippocampal Dendritic Spines
J. Neurosci.,
June 29, 2005;
25(26):
6037 - 6046.
[Abstract]
[Full Text]
[PDF]
|
|
|
|
|
|
|
T. L. Spires, Z. Molnar, P. C. Kind, P. M. Cordery, A. L. Upton, C. Blakemore, and A. J. Hannan
Activity-dependent Regulation of Synapse and Dendritic Spine Morphology in Developing Barrel Cortex Requires Phospholipase C-{beta}1 Signalling
Cereb Cortex,
April 1, 2005;
15(4):
385 - 393.
[Abstract]
[Full Text]
[PDF]
|
|
|
|
|
|
|
K. Okamura, H. Tanaka, Y. Yagita, Y. Saeki, A. Taguchi, Y. Hiraoka, L.-H. Zeng, D. R Colman, and N. Miki
Cadherin activity is required for activity-induced spine remodeling
J. Cell Biol.,
December 6, 2004;
167(5):
961 - 972.
[Abstract]
[Full Text]
[PDF]
|
|
|
|
|
|
|
M. Roelandse and A. Matus
Hypothermia-Associated Loss of Dendritic Spines
J. Neurosci.,
September 8, 2004;
24(36):
7843 - 7847.
[Abstract]
[Full Text]
[PDF]
|
|
|
|
|
|
|
W. Wallace and M. F. Bear
A Morphological Correlate of Synaptic Scaling in Visual Cortex
J. Neurosci.,
August 4, 2004;
24(31):
6928 - 6938.
[Abstract]
[Full Text]
[PDF]
|
|
|
|
|
|
|
D. A. Richards, V. de Paola, P. Caroni, B. H. Gahwiler, and R. A. McKinney
AMPA-receptor activation regulates the diffusion of a membrane marker in parallel with dendritic spine motility in the mouse hippocampus
J. Physiol.,
July 15, 2004;
558(2):
503 - 512.
[Abstract]
[Full Text]
[PDF]
|
|
|
|
|
|
|
A. Majewska and M. Sur
Motility of dendritic spines in visual cortex in vivo: Changes during the critical period and effects of visual deprivation
PNAS,
December 23, 2003;
100(26):
16024 - 16029.
[Abstract]
[Full Text]
[PDF]
|
|
|
|
|
|
|
H. Schmidt, K. M Stiefel, P. Racay, B. Schwaller, and J. Eilers
Mutational analysis of dendritic Ca2+ kinetics in rodent Purkinje cells: role of parvalbumin and calbindin D28k
J. Physiol.,
August 15, 2003;
551(1):
13 - 32.
[Abstract]
[Full Text]
[PDF]
|
|
|
|
|
|
|
C. Portera-Cailliau, D. T. Pan, and R. Yuste
Activity-Regulated Dynamic Behavior of Early Dendritic Protrusions: Evidence for Different Types of Dendritic Filopodia
J. Neurosci.,
August 6, 2003;
23(18):
7129 - 7142.
[Abstract]
[Full Text]
[PDF]
|
|
|
|
|
|
|
D. Tsay and R. Yuste
Role of Dendritic Spines in Action Potential Backpropagation: A Numerical Simulation Study
J Neurophysiol,
November 1, 2002;
88(5):
2834 - 2845.
[Abstract]
[Full Text]
[PDF]
|
|
|
|
|
|
|
G. W. Huntley, O. Gil, and O. Bozdagi
The Cadherin Family of Cell Adhesion Molecules: Multiple Roles in Synaptic Plasticity
Neuroscientist,
June 1, 2002;
8(3):
221 - 233.
[Abstract]
[PDF]
|
|
|
|
|
|
|
P. W. Vanderklish and G. M. Edelman
Dendritic spines elongate after stimulation of group 1 metabotropic glutamate receptors in cultured hippocampal neurons
PNAS,
January 24, 2002;
(2002)
32681099.
[Abstract]
[Full Text]
[PDF]
|
|
|
|
|
|
|
C. D. Holmgren and Y. Zilberter
Coincident Spiking Activity Induces Long-Term Changes in Inhibition of Neocortical Pyramidal Cells
J. Neurosci.,
October 15, 2001;
21(20):
8270 - 8277.
[Abstract]
[Full Text]
[PDF]
|
|
|
|
|
|
|
R. Yuste and A. Majewska
Book Review: On the Function of Dendritic Spines
Neuroscientist,
October 1, 2001;
7(5):
387 - 395.
[Abstract]
[PDF]
|
|
|
|
|
|
|
E. Korkotian and M. Segal
Regulation of Dendritic Spine Motility in Cultured Hippocampal Neurons
J. Neurosci.,
August 15, 2001;
21(16):
6115 - 6124.
[Abstract]
[Full Text]
[PDF]
|
|
|
|
|
|
|
P. W. Vanderklish and G. M. Edelman
Dendritic spines elongate after stimulation of group 1 metabotropic glutamate receptors in cultured hippocampal neurons
PNAS,
February 5, 2002;
99(3):
1639 - 1644.
[Abstract]
[Full Text]
[PDF]
|
|
|
|
|
TOP
ABSTRACT
INTRODUCTION
