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Small GTPases Rac and Rho in the Maintenance of Dendritic Spines
and Branches in Hippocampal Pyramidal Neurons
Ann Y. Nakayama,
1,2
Matthew B. Harms,
1
and Liqun Luo
1,2
1
Department of Biological Sciences and
2
Neurosciences Program, Stanford University, Stanford, California 94305-5020
The shape of dendritic trees and the density of dendritic spines
can undergo significant changes during the life of a neuron. We
report here the function of the small GTPases Rac and Rho in the
maintenance of dendritic structures. Maturing pyramidal neurons
in rat hippocampal slice culture were biolistically transfected with
dominant GTPase mutants. We found that expression of
dominant-negative Rac1 results in a progressive elimination of
dendritic spines, whereas hyperactivation of RhoA causes a
drastic simplification of dendritic branch patterns that is depen-
dent on the activity of a downstream kinase ROCK. Our results
suggest that Rac and Rho play distinct functions in regulating
dendritic spines and branches and are vital for the maintenance
and reorganization of dendritic structures in maturing neurons.
Key words: Rac; Rho; dendritic spines; biolistic transfection;
pyramidal neurons; effector domain mutants; ROCK; Y-27632;
PSD-95
Perhaps the most striking features of many mammalian neurons are
their complex dendritic trees. Dendrites are the principal site
where neurons receive, process, and integrate inputs from their
multiple presynaptic partners. These functions are primarily influ-
enced by the branching pattern of the dendritic tree (Rall, 1964). In
addition to complex dendritic arbors, some mammalian neurons,
including cerebellar Purkinje cells and cortical and hippocampal
pyramidal neurons, have dendritic specializations called spines.
These spines, which protrude from the dendritic branches, are the
primary site of excitatory synapses and may function as the basic
unit of synaptic integration (for review, see Harris and Kater, 1994;
Yuste and Tank, 1996). Both the shape of dendritic trees and the
density and shape of their spines can undergo significant changes
during the development and life of a neuron. For instance, den-
dritic branches have been shown to undergo significant remodeling
in adult superior cervical ganglion neurons in vivo (Purves and
Hadley, 1985; Purves et al., 1986). Recent studies have also shown
that dendritic filopodia and spines undergo dynamic changes in
response to synaptic activity (Engert and Bonhoeffer, 1999;
Maletic-Savatic et al., 1999; Toni et al., 1999) and neurotrophin
overexpression (Horch et al., 1999).
Several extracellular molecules have been identified as potential
regulators of dendrite and spine development (Purves et al., 1988;
Snider, 1988; Lein et al., 1995; McAllister et al., 1995, 1997; Nedivi
et al., 1998; Guo et al., 1999; Horch et al., 1999). To induce
dendritic growth, branching, or retraction, as well as the formation,
elaboration, or elimination of dendritic spines, these extracellular
factors must eventually exert their effects by signaling to the cy-
toskeleton. Actin is one of the main components of the dendritic
cytoskeleton and is highly enriched in dendritic spines (see Fischer
et al., 1998). The Rho family of small GTPases, including Rho,
Rac, and Cdc42, regulates various aspects of the actin cytoskeleton
(for review, see Hall, 1994; Van Aelst and D’Souza-Schorey, 1997),
and these GTPases are therefore good candidates for mediating
these signals. In agreement with their differential effects in fibro-
blasts (for review, see Van Aelst and D’Souza-Schorey, 1997), both
in vivo and in vitro studies have shown that these GTPases appear
to have distinct effects on different aspects of neuronal morpho-
genesis (for review, see Luo et al., 1997; Mueller, 1999).
Although many studies illustrate the roles of Rho family
GTPases in the establishment of neuronal processes, it remains
unclear whether they also function in later stages of neuronal
development or in mature neurons to regulate dendritic reorgani-
zation and dynamic changes of dendritic spines. To address these
questions, we used biolistic transfection (Arnold et al., 1994; Lo et
al., 1994) to introduce dominant mutants of Rac1 and RhoA
acutely into organotypic hippocampal slices at a stage when pyra-
midal neurons have established dendritic arbors and possess den-
dritic spines yet still express the GTPases. We found that Rac1 is
required for the maintenance of dendritic spines, whereas the
elevation of RhoA activity leads to pronounced simplification of
dendritic trees. We also identified candidate downstream effector
pathways that mediate these morphological changes. These studies
demonstrate that signaling pathways used in the early development
of neuronal processes are also used in more mature neurons for the
maintenance and reorganization of dendritic structures.
MATERIALS AND METHODS
In situ hybridizations
Sense and antisense S
35
-riboprobes were generated to the C-terminal ends
of rat Rac1 and RhoA by the following steps: 5⬘-Phosphorylated primers
were designed from published sequences to amplify base pairs 259–577 of
the mouse Rac1 ORF and base pairs 226 –551 of the rat RhoA ORF. These
regions were PCR amplified from an embryonic day 13 rat cDNA library,
and the bands were purified and ligated into pBLUESCRIPT II (SK
⫹
)
digested with EcoRV. Plasmids containing insertions of both orientations
were linearized with EcoRI and transcribed with T7 RNA polymerase to
generate the sense and antisense riboprobes.
Freshly dissected brains of postnatal day 8 (P8) Long–Evans rats were
submerged in ornithine carbamyltransferase and frozen in an ethanol and
dry ice bath and then stored at ⫺80°C. T wenty micrometer cryostat
sections were prepared, fixed, washed in 1⫻PBS, and serially dehydrated
before being returned to ⫺80°C in a desiccated, sealed box. Sections were
hybridized and processed for autoradiography according to the protocol
used by Frantz et al. (1994).
Digoxigenin (DIG)-labeled in sit u hybridizations were performed ac-
cording to the protocol used by Wright and Snider (1995).
Received Feb. 8, 2000; revised April 19, 2000; accepted April 24, 2000.
McKnight, K lingenstein, and Sloan Fellowships to L.L. supported this study. A.Y.N.
is a Howard Hughes Medical Institute predoctoral fellow. We are gratef ul to Larry
Katz for introducing us to the biolistic transfection method and to Matt Scott for use
of his equipment. We thank members of the laboratories of Sue McConnell, Stephen
Smith, and Dan Madison and in particular Jim Weimann, Ami Okada, Aparna Desai,
Jack Waters, and Eric Schaible for sharing experimental expertise essential for our
study. We also thank Marc Symons, Rong-Guo Qiu, Li-Huei Tsai, Alan Hall, Jun-ichi
Miyazaki, Morgan Sheng, Haruhiko Bito, and Shuh Narumiya for plasmids and
Yoshitomi Pharmaceuticals for the Y-27632 compound. We thank Ben Barres, Ste-
phen Smith, Li-Huei Tsai, and members of the Luo laboratory for stimulating
discussions and their comments on thi s manuscript. We thank Haruhiko Bito and Shuh
Narumiya for communicating results before publication.
A.Y.N. and M.B.H. contributed equally to this work.
Correspondence should be addressed to Dr. Liqun Luo, Department of Biological
Sciences, Stanford University, 371 Serra Mall, Herrin Labs 144A, Stanford, CA
94305-5020. E-mail: lluo@stanford.edu.
Copyright © 2000 Society for Neuroscience 0270-6474/00/205329-10$15.00/0
The Journal of Neuroscience, July 15, 2000, 20(14):5329–5338
Molecular biology
p-chicken

actin–mouse CD8. pUC–mouse CD8 (mCD8) (Liaw et al.,
1986) was digested with X hoI and BamHI, and the end nucleotides were
filled in using K lenow to give a 900 bp fragment encoding mCD8. The
p-chicken

actin (pCA)–GAP– enhanced green fluorescent protein
(EGFP) vector (Okada et al., 1999) was digested with BamHI and NotI,
and the end nucleotides were filled to generate a 5.1 kb pCA backbone.
The mCD8 fragment and the pCA backbone were ligated together, gen-
erating the 6 kb pCA–mCD8.
pCA–hRac1V12. pBS–hRac1V12 (Luo et al., 1996) was digested with
Asp718, blunted with Klenow, and cut again with NotI to yield a 600 bp
fragment encoding hRac1V12. This fragment was ligated into the 5.1 kb
pCA backbone generated by digesting pCA–GAP–EGFP with SmaI and
NotI, yielding the 5.7 kb pCA–hRac1V12.
pCA–myc–hRac1W T. pUHD–myc–hRac1WT (Qiu et al., 1995a) was
digested with EcoRI, and the resulting fragment encoding myc–hRac1WT
(600 bp) was ligated into the compatible site of pNN0
3
.pNN0
3
–myc–
hRac1WT was then digested with EcoRV and NotI, and the resulting
fragments were ligated into the pCA backbone as described above to yield
pCA–myc–hRac1WT.
pCA–myc–hRac1N17, pCA–myc–hRhoAV14, and pC A–myc–hR hoAN19.
pEXV–myc–hRac1N17, pEX V–myc –hRhoAN19, and pEXV–myc–
hRhoAV14 (Qiu et al., 1995a,b) were digested with EcoRI, and the
resulting fragments encoding myc –hRac1N17 (600 bp), myc –hRhoAN19
(1 kb), and myc–hRhoAV14 (1 kb) were ligated into the compatible site of
pBLUESCRIPT II (SK
⫹
). pBS–myc–hRac1N17, pBS –myc –hRhoAN19,
and pBS–myc–hRhoAV14 were digested with EcoRV and NotI, and the
resulting fragments were then ligated into the 5.1 kb pCA backbone
generated by digesting pCA–GAP–EGFP with SmaI and NotI. The result-
ing plasmid pCA–myc–hRac1N17 is 5.7 kb in length, whereas pCA–myc–
hRhoAV14 and pCA–myc–hRhoAN19 are 6.1 kb.
pCA–myc–hRac1L61, pCA–myc–hRac1L61K40, and pCA–myc–
hRac1L61A37. pRK5–myc–hRac1L61, pRK5–myc–hRac1L61K40, and
pRK5–myc–hRac1L61A37 (Lamarche et al., 1996; Nikolic et al., 1998)
were digested with ClaI and EcoRI. The resulting 600 bp fragments
encoding myc–hRac1L61, myc–hRac1L61K40, and myc–hRac1L61A37
were ligated into the compatible sites of pBLUESCRIPT II (SK
⫹
). These
plasmids were then digested with X hoI, blunted with K lenow, and then
digested with NotI. The resulting 600 bp fragments representing the
myc-tagged Rac1 mutants were isolated and ligated into the 5.1 kb pCA
backbone generated by digesting pCA–GAP–EGFP with SmaI and NotI.
The resulting pCA–myc –hRac1L61, pCA–myc –hRac1L61K40, and pCA–
myc–hRac1L61A37 are 5.7 kb in length.
Preparation of DNA-coated gold particles
Qiagen-Midi prepped plasmids were precipitated onto 1.6
m gold beads
(Bio-Rad, Hercules, CA) at a concentration of 2
g of each plasmid per
milligram of gold beads, according to the manufacturer’s instructions.
Briefly, DNA was precipitated onto gold beads that were subsequently
dried to the sides of plastic tubing. This tubing was chopped into 0.5 inch
fragments known as “bullets.” When a gold bead was to carry two plasmids
simultaneously (cotransfection experiments), both plasmids (2
g/mg of
gold each) were first mixed and coprecipitated using a standard ethanol
precipitation protocol before precipitation onto gold beads. For “dual
gold” experiments, in which a single bullet contains beads carrying differ-
ent DNAs, the two populations of gold beads were prepared separately and
mixed just before precipitation onto the plastic tubing.
Preparation of rat hippocampal organotypic cultures
Hippocampal slices were prepared from P8 Long–Evans rats as described
previously (Stoppini et al., 1991). Briefly, the hippocampus was dissected in
ice-cold, sterile dissection medium: 1⫻MEM (Hank’s salts, 25 mM
HEPES, without L-glutamine; from Life Technologies, Grand Island, NY)
and 100 units/ml each penicillin–streptomycin (Life Technologies). Hip-
pocampi were sliced transversely at a thickness of 400
m on a tissue
chopper (Stoelting, Wood Dale, IL) and separated from one another in
filtered and preincubated (37°C; 5% CO
2
) culture medium: 0.5⫻MEM,
0.25⫻HBSS (Life Technologies), 0.25⫻horse serum (defined, heat-inac-
tivated; from HyClone, L ogan, UT), 100 units/ml each penicillin–strepto-
mycin, and 1 mML-glutamine (Life Technologies). Slices were immediately
plated onto Millicell C M membrane inserts (Millipore, Bedford, M A) in
Petri dishes containing 1 ml of preincubated culture media. Slices were
kept under 5% CO
2
at 37°C, with media changes at 1 d in vitro (DIV), 3
DIV, and every 3 d thereafter. In experiments using Y-27632 (gift from
Yoshitomi Pharmaceuticals, Tokyo, Japan) slices were transferred to Petri
dishes containing 100
MY-27632 in 1 ml of culture media just before
biolistic transfection. Control slices were similarly transferred, but to Petri
dishes containing media supplemented with the same volume of sterile
water instead of Y-27632. Animals were treated in accordance with the
animal safety protocols of the host institution.
Biolistic transfection
To identify optimal transfection and culture conditions, slices were pre-
pared from P8 rats, transfected at various DIV, and fixed for immunocy-
tochemistry 24 hr later. The number of healthy pyramidal neurons relative
to those that were dead or dying (characterized by fragmented processes or
blebbing processes, respectively) increased significantly when transfection
occurred at 2 DIV, compared with transfection at 0 or 1 DIV. When
transfection occurred at ⬎2 DIV, the number of transfected glia increased
significantly, often obscuring the dendritic arbors we wished to quantify.
Thus, to maximize the number of healthy pyramidal cells transfected and
to minimize interfering glia, experiments throughout this study were
performed on a standard preparation: hippocampal slices prepared from
P8 rats and cultured for 2 d before transfection.
After 2 DIV, slice inserts were removed from the incubator briefly for
biolistic transfection using the Gene Gun (Bio-Rad). Gold beads contain-
ing expression plasmids were propelled from plastic tubing bullets into
slices with a rapid helium burst of 160 psi. The gold beads exited the gun
⬃3.5 cm above the slices, which were plated toward the periphery of the
insert to avoid “ground zero” of the gold blast.
Immunocytochemistry
Slices were fixed according to the protocol used by McAllister et al. (1995)
on the insert membrane for 1.5 hr in 2.5% formaldehyde and 4% sucrose
in 1⫻PBS and then soaked in 30% sucrose (w/v of water) for at least 2 hr.
After a quick freeze on dry ice, slices were thawed, rinsed in 1⫻PBS for
5 min, and incubated overnight (O/N) at 4°C in blocking solution: 10%
normal goat serum and 0.25% Triton X-100 in 0.1 Mphosphate buffer.
Blocking solution was replaced with primary antibodies diluted in blocking
solution: 1:50 mouse anti-myc (Santa Cruz Biotechnology, Santa Cruz,
CA), 1:100 rat anti-mCD8 (Caltag, Burlingame, CA), or both before
another O/N incubation at 4°C. The slices were then washed three times
for 20 min each with fresh blocking solution and incubated O/N at 4°C in
secondary antibody, diluted into blocking solution: 1:200 FITC anti-mouse
(Jackson ImmunoResearch, West Grove, PA), 1:1000 indocarbocyanine
(Cy3) anti-rat (Jackson ImmunoResearch), or both (in which the low
cross-reactivity secondaries were used). Slices were subsequently washed
three times for 15 min each in 1⫻PBS, with the first wash containing 1.5
g/ml 4⬘,6-diamidino-2-phenylindole (DAPI), and mounted in SlowFade
according to the manufacturer’s specifications (Molecular Probes, Eu-
gene, OR).
Image analysis and quantification
Transfected pyramidal neurons were identified by their typical morpholog y
as well as their cell body locations in the CA1 and CA3 pyramidal layers as
indicated by DAPI staining (see Fig. 2A). Individual neurons were imaged
using a Zeiss microscope attached to a Bio-Rad MRC -1024 scanning laser
confocal microscope. For quantification of dendritic branch segments, a
stack of confocal images (Z steps of 1
m) taken with a 16⫻objective and
comprising the entire cell were merged using NIH Image 1.62 (National
Institutes of Health, Bethesda, MD) and printed. From these printouts, the
number of dendritic segments was derived by counting the number of
dendrite branch points and dendrite terminal ends. To obtain the Sholl
profiles of dendritic arbors (Sholl, 1953), printouts were placed under a
clear sheet printed with concentric circles with increasing radii of 25
m.
To minimize the effect of cell body shape variation, we positioned the
center of the circles at the base of the apical dendrite for analysis of apical
dendrites. In addition, because CA1 and CA3 pyramidal neurons exhibited
different Sholl profiles (data not shown), all Sholl analysis was with only
CA1 neurons. The center of the circles was placed at the cell body edge,
opposite the apical dendrite when analyzing basal dendrites. The number
of dendrites crossing each concentric circle was then counted. If a branch
point fell on a line, it was counted as two crossings.
For spine quantification, images of apical dendrites were taken just distal
to the first apical branch point (25–100
m from the cell body). Basal
spines were imaged at the point of the first basal branch (25–75
m from
the cell body). Serial confocal images (Z steps of 0.5
m) were taken of
Cy3 fluorescence (detecting mCD8) with a 40⫻objective with a digital
zoom factor of three. Sections were merged using N IH Image, and the
number of spines and filopodia was tallied on the screen. Spines were
defined as a headless dendritic protrusion 1–3
m long or a headed
protrusion of any length up to 3
m. Filopodia were defined as headless
protrusions ⬎3
m but not long enough to be visible on the 16⫻printout.
The length of all dendrites in the fields used to quantify spine density was
measured using NIH Image. The average dendritic diameter for each cell
was based on three measurements (at the start, middle, and end) of every
dendritic segment within the images used to analyze dendritic spine
density.
Identical procedures for acquiring images and measurements of den-
dritic length were also used to analyze GFP-tagged postsynaptic density-95
(PSD-95:GFP) clustering with Rac1N17 expression. To count PSD-95:
GFP clusters optimally in relation to spine profiles as defined above,
images were placed into channels in Photoshop 4.0 (Adobe Systems, San
Jose, CA). GFP clusters were counted independent of location, and the
spine profiles with GFP clusters either within the spine head or just
external to the head outline were also tallied. The dendrites and their
spines were outlined using the magic wand tool in the channel correspond-
ing to the myc label, which fills the entire dendritic tree including the
spines (see Fig. 3).
Graphs throughout represent averages and SEM. Statistical comparisons
of Sholl segments were done with StatView 5.0.1 (SAS Institute, Cary,
5330 J. Neurosci., July 15, 2000, 20(14):5329–5338 Nakayama et al. •Rac and Rho Maintain Dendritic Spines and Branches
NC), and all post hoc two-tailed ttests assuming equal variances were done
using Microsoft Excel 98 (Microsoft, Seattle, WA).
RESULTS
Rac1 and RhoA are expressed in developing
hippocampal pyramidal neurons
Although Rac1 and RhoA are ubiquitously expressed (for review,
see Hall, 1994), the expression pattern of these GTPases in the
hippocampus has not been described. To identify Rac1- and RhoA-
expressing cells in the hippocampus, in situ hybridizations were
performed on coronal P8 rat brain sections, the same stage at which
our slice cultures were produced. Antisense S
35
-labeled riboprobes
were generated to the C-terminal portions of each mRNA, where
the nucleic acid sequence identities between Rac and Rho are
lowest (50% identical). As shown in Figure 1, Rac1 (Fig. 1 A) and
RhoA (Fig. 1B) antisense riboprobes labeled CA1 and CA3 pyra-
midal cell layers and the dentate granular cell layer, with the CA3
layer being the most intense. Rac and Rho transcripts are also
widely distributed in other brain areas (data not shown). The sense
riboprobes to both GTPases gave only diffuse background signal,
similar to that shown for Rac1 (Fig. 1C). In sit u hybridization using
DIG-labeled probes was performed on sections made from P10
hippocampi (equivalents to the day slices were transfected), with
similar results (data not shown). No obvious dendritic localization
of the two mRNAs was observed.
Pyramidal neurons maintain in vivo characteristics while
in slice culture
We have used particle-mediated gene transfer (biolistic transfec-
tion) (Arnold et al., 1994; Lo et al., 1994) of mCD8 cDNA, and
subsequent immunocytochemical detection of the protein, to visu-
alize the dendritic arbors and spines of pyramidal neurons in
hippocampal slice cultures. mCD8 is a cell-surface marker shown
previously to label the entire morphology and fine structures of
Drosophila neurons without toxic side effects (Lee and L uo, 1999).
As shown in Figure 2, delivery of a single gold particle carrying
mCD8 under the control of the chicken

-actin promoter resulted
in the intense labeling of not only the axon and the entire dendritic
arbor (Fig. 2B) but individual spines as well (Fig. 2C). In addition
to pyramidal neurons, many other hippocampal cell types were
readily transfected, including glia, the granule cells of the dentate
gyrus, and interneurons (Fig. 2A).
In our standard preparation, hippocampal slices were obtained
from postnatal day 8 rat pups and cultured for 2 d before transfec-
tion (see Materials and Methods). To determine the developmental
state of pyramidal neurons at the time of transfection, 1 d in vitro
cultures were transfected with mCD8 and fixed 24 hr later (2 DIV).
By this time, pyramidal neurons had already acquired their char-
acteristic dendritic-branching pattern, including well differentiated
apical and basal dendrites (Fig. 2B). Counting the number of
dendritic branch segments (see Materials and Methods) gave us a
quantitative measure of dendritic complexity and revealed that
over the first few days after transfection, there is a gradual increase
in the number of dendritic segments (Fig. 2 E). At each time point
examined, we found no statistically significant difference between
the number of dendritic segments for apical and basal dendrites or
between CA1 and CA3 neurons (data not shown).
These 2 DIV neurons also displayed numerous protrusions from
their apical and basal dendrites. Although a minority of these
protrusions were filopodial in shape (Fig. 2C,arrowheads), many
more had the well defined neck and head structure characteristic of
mature spines found in adults (Fig. 2C,arrows). To verify indepen-
dently the maturity of these dendritic protrusions in our culture, we
cotransfected GFP-tagged PSD-95 as a marker for postsynaptic
density (Arnold and Clapham, 1999). We found that PSD-95:GFP
labeling was concentrated in most of the heads of the protrusions
but was absent from longer filopodia (Fig. 2D). Using the morpho-
logical criterion described above (see Materials and Methods for
details), we quantified the density of spine-like protrusions and
filopodial protrusions. We found that spine density increased with
the time spent in culture, whereas the number of filopodial protru-
sions gradually diminished (Fig. 2 F). The spine densities found in
our cultures are similar to those reported previously for similarly
aged cultures [P12 equivalent ⫽2 spines/10
m by EM (Boyer et
al., 1998)]. In accordance with Drakew et al. (1996), we did not find
a significant difference in spine density between the apical and
basal dendrites at any time point examined (data not shown).
Additionally, we did not observe a significant difference in spine
density between CA1 and CA3 pyramidal dendrites (data not
shown). Because CA1 and CA3 neurons did not differ in either
their branch segment number or their spine density, neurons from
both areas have been combined in our quantification of branch
segments and spine densities.
Expression of dominant-negative Rac1 results in a
progressive reduction of spine density
To study the function of Rho family GTPases in the maintenance
of dendritic structures, we cotransfected pyramidal neurons in our
standard preparation with both mCD8 and dominant mutants of
the GTPases. Cotransfection was accomplished by the introduction
of gold beads coated with plasmids for both genes, each under the
control of the chicken

-actin promoter. Although analogous ex-
periments with two marker genes (mCD8 and GAP–EGFP)
showed a high cotransfection rate (⬎85%), quantitative variations
in the relative expression levels of the two constructs were ob-
served. Therefore, we independently monitored the expression of
the dominant mutants by using myc epitope-tagged GTPases, an-
alyzing only those cells in which both plasmids were strongly
expressed. As an internal control and to minimize variation among
hippocampal slices, most experiments were performed using dual
gold preparations. In these experiments, slices were transfected
with a mixed population of gold beads, some carrying the mCD8
and GTPase plasmids and some the mCD8 plasmid alone. This
enabled us to visualize both control (mCD8 alone) and experimen-
tal (mCD8 and GTPase) neurons from the same slice (see Fig. 3C).
We found that transfected wild-type Rac1, as well as all mutant
proteins, was distributed throughout the entire dendritic tree, in-
cluding dendritic spines and other fine processes (see Figs. 3A,C,
4B–D,F–H).
To test whether endogenous Rac1 is necessary for the mainte-
nance of dendritic spine and branch morphology, we cotransfected
pyramidal neurons with gold beads carrying mCD8 and myc-tagged
dominant-negative Rac1 (Rac1N17). This allele is thought to exert
its dominant-negative effect by sequestering rate-limiting GDP–
GTP nucleotide exchange factors necessary for activation of en-
dogenous Rac1 (Ridley et al., 1992). Expression of Rac1N17 re-
sulted in a significant time-dependent loss of pyramidal dendritic
Figure 1. Rac1 and RhoA are expressed in developing hippocampus. A,
Dark-field image of a coronally sectioned P8 rat hippocampus hybridized
with an antisense S
35
-riboprobe against Rac1, showing distribution of Rac1
mRNA in the dentate gyrus and CA1 and CA3 pyramidal cell layers. B,
Dark-field image of antisense riboprobe showing distribution of RhoA
mRNA. C, Dark-field image of the Rac1 sense riboprobe showing back-
ground staining. D, Bright-field image of the same hippocampus shown in
Cwith the CA1, CA3, and dentate gyrus (DG) labeled. Scale bar, 2 mm.
Nakayama et al. •Rac and Rho Maintain Dendritic Spines and Branches J. Neurosci., July 15, 2000, 20(14):5329–5338 5331
Figure 2. Development of hippocampal pyramidal neurons in
cultured slices. A–C, Representative images of transfected hip-
pocampal neurons show the developmental stage at the onset of
experiments in our standard preparation. For these images
only, transfection was performed at 1 DIV, and slices were
fixed 24 hr later. A, Low magnification is shown of a hippocam-
pal slice biolistically transfected with mCD8 (red), with brack-
ets defining the DAPI-labeled (blue) pyramidal cell layers of
CA1 and CA3. Pink labeling (from overlapping red and blue
signals) represents transfected cells, including a granule cell in
the dentate gyrus (arrowhead) and a pyramidal neuron over-
lapped by glia in the CA1 cell layer (ar row). B, A CA3 pyra-
midal neuron (composite confocal image using a 16⫻objec-
tive) is shown. The immunocytochemical detection of mCD8
reveals the structure of both apical and basal dendrites, as well
as the axon (arrow). C, The spines on the apical dendrites of the
neuron pictured in B(composite confocal image using a 100⫻
objective) are shown. Arrows point to spines with characteristic
head and neck morphology, whereas arrowheads show filopo-
dial protrusions. D, Double labeling of mCD8 and PSD-95:
GFP proteins in apical dendrites (composite confocal images
using a 40⫻objective with a digital zoom factor of 3) is shown.
mCD8 labeling is in red (D,D⬘⬘), and PSD-95:GFP labeling
(PSD-95:GFP)ising reen (D⬘,D⬘⬘). Arrows indicate spines with
a head (PSD-95:GFP positive), whereas arrowheads indicate
filopodial protrusions (PSD-95:GFP negative). E, Dendritic
branch segments gradually increase on both apical and basal
dendrites with successive days in culture (apical, n⫽10, 13, 8,
10; basal, n⫽10, 14, 7, 9 for 2, 4, 6, 9 DIV, respectively). F,
Spine density increases over successive days in culture, whereas
filopodial protrusions decrease. Spines and filopodia were
counted from stacked confocal images collected at 40⫻with a
zoom of three, from a stereotyped region of the dendrite as
defined in Materials and Methods (apical, n⫽10, 19, 17, 12;
basal, n⫽11, 14, 17, 11 for 2, 4, 6, 9 DIV, respectively). Scale
bars: A, 1 mm; B,50
m; C, D,10
m.
Figure 3. Dominant-negative Rac1 expression results in a
progressive reduction of the dendritic spine density and mild
changes in the dendritic-branching pattern. A, Transfected
myc-tagged wild-type Rac1 protein is distributed along the
dendrites and in dendritic spines ( green, anti-myc; red, anti-
mCD8; composite confocal image using 40⫻objective; digital
zoom factor of 3). B, Representative images are shown of
apical dendrites that were transfected with the marker mCD8
alone (top) or with mCD8 and myc-tagged Rac1N17 (bottom)
for 1, 2, or 3 d (composite confocal images of mCD8 staining
using 100⫻objective). C, Apical dendrites from neighboring
pyramidal neurons are shown3dafter expressing mCD8 alone
(red) or expressing both mCD8 (red) and myc-tagged Rac1N17
(green) and therefore appearing yellow (composite confocal
image using a 40⫻objective with a digital zoom factor of 3).
Although it appears that Rac1N17 dendrites have thicker den-
drites in this image, quantification of the average diameter of
dendrites within every image used to measure dendritic spine
density does not reveal any significant difference (paired ttest,
apical, p⫽0.19; Rac1N17 ⫽1.50 ⫾0.14
m; control ⫽1.22 ⫾
0.51
m; n⫽18, 9, respectively; basal, p⫽0.12; Rac1N17 ⫽
0.83 ⫾0.1
m; control ⫽1.05 ⫾0.08
m; n⫽9, 8, respective-
ly). D, Rac1N17 expression progressively reduces the number
of spines on apical and basal dendrites (for 1, 2, 3 d, apical
mCD8, n⫽16, 19, 10; apical Rac1N17, n⫽19, 11, 24; basal
mCD8, n⫽15, 14, 10; basal Rac1N17, n⫽18, 11, 17, respec-
tively). E, Quantification of dendritic branch segments after 3 d
of Rac1N17 expression is shown (n⫽17, 23, 18, 21 for apical
mCD8, apical Rac1N17, basal mCD8, basal Rac1N17, respec-
tively). F, Sholl profiles of the basal dendrites of Rac1N17-
expressing and control CA1 pyramidal neurons (3 d after transfection) illustrate the slight change in dendritic-branching pattern with Rac1N17 expression
(n⫽14, 15 for basal mCD8, Rac1N17, respectively). Post hoc t tests reveal that the only significant difference occurs at 50
m (paired ttest, **p⬍0.01).
Scale bars: A, B,5
m; C,10
m.
5332 J. Neurosci., July 15, 2000, 20(14):5329–5338 Nakayama et al. •Rac and Rho Maintain Dendritic Spines and Branches
spines from both apical and basal dendrites. Whereas neurons
expressing mCD8 alone displayed an increasing trend of spine
density with successive days after transfection, those also express-
ing Rac1N17 possessed fewer and fewer spines on both their apical
and basal dendrites (Fig. 3B—D; see also Fig. 2F).Only1dafter
transfection, the basal dendritic spine density of Rac1N17 neurons
was significantly reduced compared with that of neurons expressing
mCD8 alone (68% fewer; p⬍0.05), whereas the apical spine
density was significantly lower than that of controls2dafter
transfection (54% fewer; p⬍0.05). An example of a dual gold
experiment is shown in Figure 3C.After3dofexpression, mCD8-
transfected pyramidal dendrites (red) displayed adult-like spines,
whereas the dendrites from Rac1N17-expressing cells ( yellow be-
cause of red labeling of mCD8 and green labeling of myc-tagged
Rac1N17) were nearly devoid of spines.
To address whether the corresponding synapse number is de-
creased with the loss of the dendritic spine profile in Rac1N17-
transfected neurons, we cotransfected PSD-95:GFP with Rac1N17.
In agreement with the loss of dendritic spine density, we observed
a significant decrease in PSD-95:GFP clusters with3dofRac1N17
expression ( p⬍0.05; Rac1N17 ⫽2.14 ⫾0.35; mCD8 ⫽3.84 ⫾
0.52; units are PSD-95:GFP clusters per 10
m; n⫽9, 8,
respectively).
To determine whether Rac1 may play a role in the maintenance
of dendritic growth and branching, we quantified the number of
dendritic segments after3dofRac1N17 expression. As shown in
Figure 3E, there is no statistically significant difference between
Rac1N17 and control cells (apical, p⫽0.38; basal, p⫽0.48). As
another measure of dendritic complexity, a standard Sholl analysis
(Sholl, 1953), which counts the number of dendritic crossings at 25
m concentric circles, was performed. Although the Sholl profiles
for the apical dendrites of Rac1N17-expressing neurons and con-
trol neurons are similar (repeated measures ANOVA, p⫽0.40),
their basal profiles are slightly different (Fig. 3F; repeated measures
ANOVA, p⬍0.05). There are more branches concentrated in the
proximal regions in control neurons compared with Rac1N17 neu-
rons, which appear to have more distally distributed branches (Fig.
3F). This mild effect of Rac1N17 on the dendritic-branching pat-
tern of pyramidal neurons is in contrast to its pronounced effect on
spine density, suggesting a preferential role for endogenous Rac1
in the maintenance of dendritic spines.
Activated Rac1 disrupts normal spine morphology
To elucidate how Rac1 activity affects spine morphogenesis, we
transfected pyramidal cells with a constitutively activated form of
Rac1 that is deficient in GTP hydrolysis (hereafter referred to as
activated Rac1). Previous transgenic expression of the activated
allele Rac1V12 in Purkinje cells results in a slight simplification of
dendritic complexity, a loss of normal dendritic spines, and the
formation of supernumerary smaller protrusions that were only
detectable with EM (Luo et al., 1996). We found that pyramidal
neurons expressing Rac1L61, another activated Rac1 allele, for 1,
2, or 3 d also showed similar defects in spine development (Fig.
4B). With the exception of some distal dendrites, apical and basal
dendrites lacked normal spines. In contrast to our control neurons
(Fig. 4A), which have even dendritic branches that are regularly
punctuated by spines, Rac1L61 dendrites were irregular in thick-
ness because of the presence of regions of numerous overlapping
bumps and ruffle-like structures (Fig. 4B,F, arrowheads). Addition-
ally, numerous long and fine processes were found on the cell soma
and proximal dendritic shafts (Fig. 4, compare F,arrows,E).
Although these fine processes prevented quantification of dendritic
segments, 70% of Rac1L61-expressing neurons had an otherwise
normal dendritic-branching pattern despite their perturbed den-
dritic spines. These findings suggest that dendritic spines are more
sensitive to the hyperactivation of Rac1 than are the dendritic
branches themselves. Expression of another constitutively active
form of Rac1, Rac1V12, in hippocampal neurons gave indistin-
guishable results compared with Rac1L61 expression (data not
shown). Although normal dendritic spines are lost with the expres-
sion of either dominant-negative or constitutively active Rac1,
Rac1L61 expression resulted in a net increase in dendritic protru-
sion exemplified by an increase in filopodial-like structures and
membrane ruffling, which were not seen with Rac1N17 expression
(Fig. 3B,C).
The F37A effector domain mutant abolishes the
activated Rac1 phenotype
Rac1 binds to different effectors to activate distinct downstream
signal transduction pathways. Previous in vitro experiments have
characterized several effector domain mutants that separate Rac1’s
role in lamellipodia formation from its role in activation of the Jun
kinase cascade and nuclear signaling (Joneson et al., 1996;
Lamarche et al., 1996). The Y40K mutation abolishes the binding
of several CRIB-containing proteins [e.g., p21-activated kinase
(Pak)] and when combined with an activating Rac1 mutation blocks
the stimulation of the Jun kinase (JNK) pathway without affecting
membrane ruffling and lamellipodia formation. Conversely F37A
mutation does not bind to Rho-associated kinase (ROCK) and
prevents the induction of membrane ruffling and lamellipodia by an
activating mutation of Rac (Lamarche et al., 1996), yet it still binds
to Pak and activates the JNK pathway. These data suggest that
effectors dependent on tyrosine residue 40 for binding to activated
Rac (e.g., Pak and other CRIB-containing proteins) are required
for the activation of Jun kinase, whereas effectors dependent on
phenylalanine residue 37 (e.g., ROCK) for binding are candidate
mediators for Rac1’s regulation of lamellipodia formation.
We attempted to identify the pathway responsible for the acti-
vated Rac1 phenotype in hippocampal pyramidal neurons using
these same effector domain mutants. Rac1L61K40 expression re-
sulted in irregular, bumpy dendrites containing numerous thin
processes (Fig. 4C,G) that were indistinguishable from Rac1L61
dendrites (Fig. 4B,F ), suggesting that effectors dependent on ty-
rosine 40 for binding (e.g., Pak) are not necessary to mediate the
effects of activated Rac1. In contrast, the dendritic morphology and
spine density of pyramidal neurons expressing Rac1L61A37 (Fig.
Figure 4. Effects of activated Rac1 expression on dendritic
morphology. A–H, Representative distal ( A–D) and proximal
(E–H, just above cell bodies) apical dendrites and spines of
hippocampal pyramidal neurons, 24 hr after being transfected
with mCD8 only (A, E), mCD8 and Rac1L61 (B, F ), mCD8 and
Rac1L61K40 (C, G), or mCD8 and Rac1L61A37 (D, H ) (com-
posite confocal images using a 40⫻objective with a digital
zoom factor of 3). Red staining in all images represents anti-
mCD8 immunoreactivity, and green staining in B–D and F–H
represents anti-myc immunoreactivity. All mutant Rac proteins
were expressed at comparable levels and were distributed
throughout the entire neuron on the basis of their myc staining
(see yellow labeling in B–D,F–H because of overlapping green
signal for myc and red signal for mCD8). Asterisk s represent
mature spines with heads, ar rowheads indicate ruffle-like struc-
tures, and arrows point to long and thin filopodial-like protru-
sions. Scale bars, 10
m.
Nakayama et al. •Rac and Rho Maintain Dendritic Spines and Branches J. Neurosci., July 15, 2000, 20(14):5329–5338 5333
4D,H) resembled those of control neurons (Fig. 4A,E), implying a
requirement for effectors dependent on phenylalanine 37 to medi-
ate the activated Rac1 phenotype. ROCK is one such protein
(Joneson et al., 1996; Lamarche et al., 1996). We attempted to
mimic the Rac1L61A37 rescue by treating Rac1L61-transfected
slices with 100
MY-27632, a compound that specifically inhibits
ROCK activity without affecting other similar kinases (see Uehata
et al., 1997; Madaule et al., 1998). Although this treatment effec-
tively blocked the activated RhoA phenotype (see below), the
Rac1L61 effect persisted, suggesting that ROCK is not responsible
for mediating the effect of activated Rac1.
Expression of activated RhoA results in marked
simplification of the dendritic tree
Although expression of activated Rac1 had a mild effect on the
overall dendritic branch complexity (discounting the filopodia-like
thin processes), expression of activated RhoA (RhoAV14, analo-
gous mutation to Rac1V12) caused a drastic simplification of the
dendritic tree compared with those of neurons expressing mCD8
alone (Fig. 5A,B). This effect was evident1dafter transfection and
persisted for 2 and 3 d after transfection. Branch segment quanti-
fication (Fig. 5E) indicated a highly penetrant and significant sim-
plification of both the apical and basal dendrites as early as1dafter
transfection of RhoAV14 (ttest, p⬍0.001 for both apical and
basal). The Sholl profile indicates that both the length and branch-
ing pattern of RhoAV14-expressing neurons are affected (Fig. 5G;
repeated measures ANOVA, p⬍0.001). Interestingly, there ap-
peared to be a recovery of dendritic simplification after3dof
RhoAV14 transfection, as seen by quantification of both apical and
basal dendritic segment numbers (Fig. 5E), despite the constant
level of RhoAV14 expression as judged by myc staining (data not
shown).
Strikingly, many neurons expressing activated RhoA (80%) had
unusual structures at the end of their reduced dendrites: threads of
extremely thin processes trailing behind the shortened dendrites
(Fig. 5D) resembling certain aspects of retracting axon terminals
(for review, see Bernstein and Lichtman, 1999). These shortened
dendrites often exhibited a fattened portion that appeared to have
an increased aggregation of cytoplasm and membranous structures
on the basis of the intensity of the cytoplasmic-myc and plasma
membrane-localized CD8 labeling.
Perturbation of endogenous RhoA activity with transfection of
the dominant-negative RhoA construct (RhoAN19, analogous mu-
tation as Rac1N17) did not affect the dendritic morphology of our
pyramidal neurons (Fig. 5C,F,H ) despite their high level of expres-
sion (see Fig. 5C, inset). In addition, expression of RhoAN19 did
not significantly affect the spine density when compared with con-
trol neurons2dafter transfection ( p⫽0.56; n⫽15 and 13 for
RhoAN19 and mCD8 alone, respectively), supporting the specific-
ity of the spine reduction effect seen with dominant-negative
Rac1N17 expression.
The ROCK inhibitor blocks RhoAV14-induced
dendritic simplification
RhoA acts via many different effector molecules to regulate diverse
cellular functions, including organization of the actin cytoskeleton.
To investigate the pathway by which RhoA regulates dendritic
complexity, we tested the involvement of ROCK, because ROCK
inhibition has been shown to block Rho-induced cell rounding and
process retraction in cultured neuroblastoma cells (Hirose et al.,
1998). We applied the ROCK inhibitor Y-27632 (100
M) (Uehata
et al., 1997) in culture media at the time of transfection. We limited
our quantification to the basal dendrites of neurons2dafter
transfection because the dendritic simplification was most robust at
this time, and both apical and basal dendrites were equally affected
by RhoAV14 expression (Fig. 5E).
The treatment of slices with Y-27632 alone did not result in a
significant change in dendritic complexity (Fig. 6 B,F;p⫽0.30).
Remarkably, Y-27632 treatment completely blocked the dendritic
simplification associated with RhoAV14 expression (Fig. 6, com-
pare D, C). As is quantified in Figure 6 F, although RhoAV14
expression caused a fivefold decrease in dendritic segments, appli-
Figure 5. Expression of activated RhoA results in den-
dritic simplification. A–C, Representative images of pyra-
midal neurons that have expressed the marker mCD8
alone (A), mCD8 and myc-tagged RhoAV14 (B), or
mCD8 and myc-tagged RhoAN19 ( C) for 1 d (composite
confocal images using 16⫻objective). Insets, Myc immu-
nostaining for RhoAV14 (B) and RhoAN19 ( C). D, Tip
of an apical dendrite from a pyramidal neuron expressing
RhoAV14 for 2 d (composite confocal image using 40⫻
objective, with a digital zoom factor of 3). The soma is
toward the bottom of the image. E, Quantification of
dendritic branch segments after 1, 2, and3dofRhoAV14
expression. Both apical and basal dendrites exhibit a re-
duced number of dendritic segments (see Materials and
Methods) with RhoAV14 expression compared with that
of neurons expressing mCD8 alone (***p⬍0.001; **p⬍
0.01; for 1, 2, 3 d, apical mCD8, n⫽14, 13, 17, and
RhoAV14, n⫽25, 19, 23; basal mCD8, n⫽11, 14, 18, and
RhoAV14, n⫽25, 18, 22). F, Quantification of dendritic
branch segments after 1, 2, and 3 d of RhoAN19 expres-
sion. Neurons expressing RhoAN19 do not show a change
in dendritic branch segment number (for 1, 2, 3 d, apical,
n⫽9, 10, 7; basal, n⫽7, 11, 7, respectively). G, H, Sholl
profiles for basal dendrites of CA1 neurons2dafter
expressing mCD8 alone (n⫽11) or expressing RhoAV14
(G;n⫽16) or RhoAN19 (H;n⫽8). Scale bars: A–C and
insets,50
m; D,10
m.
5334 J. Neurosci., July 15, 2000, 20(14):5329–5338 Nakayama et al. •Rac and Rho Maintain Dendritic Spines and Branches
cation of Y-27632 restored the number of dendritic segments to the
level of our control neurons. This result suggests that ROCK is a
mediator of RhoAV14-induced dendritic simplification.
The significant dendritic simplification induced with RhoAV14
expression impeded our ability to quantify the density of dendritic
spines. On a qualitative level it appeared that expression of acti-
vated RhoA resulted in a loss of adult-like dendritic spines. Inter-
estingly, although Y-27632 alone does not affect spine density, it
also restores spine density of RhoAV14-expressing neurons close to
the control level (Fig. 6G;p⫽0.08).
ROCK activation is sufficient to induce
dendritic simplification
We next tested whether activation of ROCK is sufficient to induce
dendritic simplification. Pyramidal neurons were transfected with a
truncation mutant, ROCK⌬3, which eliminates the negative regu-
latory domain in the C terminal of the ROCK protein and has been
shown to behave as a constitutively active version of ROCK in
cultured cells (Ishizaki et al., 1997). We found that ROCK⌬3
expression resulted in the reduction of dendritic complexity to an
extent similar to that of RhoAV14 expression (Fig. 6 E,H), suggest-
ing that ROCK activation is not only necessary but also sufficient in
inducing the pruning of dendritic branches.
DISCUSSION
Using biolistic transfection to introduce dominant mutants of Rac1
and RhoA acutely in hippocampal pyramidal neurons with estab-
lished dendritic arbors and some adult-like dendritic spines, we
have shown that these GTPases play important roles in the main-
tenance and reorganization of dendritic structures. Rac1 seems to
have preferential roles in regulating spine morphogenesis, whereas
RhoA is implicated in limiting the growth of dendritic branches.
The fact that expression of analogous dominant-negative or acti-
vated mutants of these similar GTPases gave qualitatively different
phenotypes argues for the relative specificity of these perturbation
experiments.
Generation and maintenance of dendritic spines
Although dendritic spines are important sites of synaptic input and
plasticity (Harris and Kater, 1994; Yuste and Tank, 1996), little is
known about the molecular mechanisms that regulate their mor-
phogenesis (Harris, 1999). We have shown previously that trans-
genic expression of activated Rac1 in mouse Purkinje cells results
in drastic changes in dendritic spine morphology (Luo et al., 1996).
In those experiments, transgenes were expressed before the onset
of dendritic growth and remained active thereafter. Although we
concluded that hyperactivation of Rac affects the development of
dendritic spines, we could not determine whether regulation of Rac
activity is important for the dynamic changes of dendritic spines in
more mature neurons, which may contribute to the morphological
plasticity of neurons underlying learning and memory (Harris and
Kater, 1994).
In this study, we have extended the previous findings in several
new directions. First, because expression of activated Rac1 mutants
in hippocampal pyramidal neurons results in a spine perturbation
phenotype analogous to transgenic expression of activated Rac1 in
Purkinje cells, it seems that the effect of increased Rac1 activity in
dendritic spine morphogenesis is not specific to Purkinje cells. A
notable difference between the two experimental conditions, how-
ever, is the presence of prof use long filopodial-like extensions in
this study (Fig. 4) that were not observed in the previous transgenic
studies (Luo et al., 1996). This could be caused by the differences
in cell types, transgene expression level, and the relative timing of
transfection versus neuronal differentiation. For instance, in the
transgenic experiments, neurons may have adapted to the constant
high level of Rac1 activity over a long period of time, such that the
filopodial response is diminished. A second new insight gained
from the current study is that activated Rac1 expression perturbs
the maintenance of spines, because at the time of transfection these
pyramidal neurons already possess dendritic spines.
Third and more importantly, our current study finds that expres-
sion of dominant-negative Rac1 resulted in a progressive reduction
Figure 6. Function of ROCK in the maintenance of dendritic branches. A–E, Representative images of neurons2dafter transfection and 100
MY-27632
treatment are shown. Neurons were transfected with mCD8 alone (A, B) or cotransfected with RhoAV14 (C, D) or ROCK⌬3(E). E,Inset, The myc
immunostaining for ROCK⌬3 is shown. Additionally, neurons in Band Dwere treated with 100
MY-27632 at the time of transfection (composite confocal
images using 16⫻objective). F, Quantification of basal dendritic branch segment numbers of mCD8- and of mCD8 plus RhoAV14-expressing neurons with
or without Y-27632 treatment is shown. Y-27632 treatment alone does not affect dendritic segment number ( p⫽0.31; control treatment, n⫽23; Y-27632
treatment, n⫽23). Y-27632 treatment blocks RhoAV14-associated dendritic segment reduction (***p⬍0.001; RhoAV14 ⫹control, n⫽17; RhoAV14
expression ⫹Y-27632 treatment, n⫽24). G, Y-27632 application does not alter the basal dendritic spine density of neurons expressing mCD8 alone ( p⫽
0.65; n⫽29, 21 for control, Y-27632 treatment, respectively). Y-27632 treatment is capable of restoring the spine density of neurons expressing RhoAV14
close to control level ( p⫽0.08; n⫽16 for Y-27632 treatment and RhoA V14 expression). H, Activated ROCK⌬3 expression results in significant reduction
of dendritic segments compared with that in mCD8 alone ( p⬍0.001 for both apical and basal; n⫽19, 20, respectively, for ROCK⌬3). Scale bars, 50
m.
Nakayama et al. •Rac and Rho Maintain Dendritic Spines and Branches J. Neurosci., July 15, 2000, 20(14):5329–5338 5335
of spine number in both apical and basal dendrites. Taken together
with the expression of Rac1 mRNA in the hippocampal pyramidal
neurons at the time of our experiments, these results suggest that
endogenous Rac1 is used for the maintenance of spine density in
maturing neurons. Because of the slow but significant turnover of
dendritic spines at analogous developmental stages reported from
several recent live-imaging studies (Engert and Bonhoeffer, 1999;
Horch et al., 1999), we cannot distinguish between the following
two possibilities. (1) Rac1 is required for the generation of new
spines. The net spine loss over time is a result of the failure of new
spine formation while existing spines naturally turn over. (2) Rac1
is required for the maintenance of existing spine structure. Reduc-
tion in Rac activity speeds up the turnover rate of existing spines.
Future time-lapse observation of dominant-negative Rac1-
transfected neurons may help to distinguish between these two
possibilities.
Interestingly, the number of PSD-95:GFP clusters, markers for
postsynaptic density, decreases with Rac1N17 expression com-
pared with controls. Although it is not possible to conclude whether
loss of Rac1 activity causes a loss of PSD-95 clustering before spine
structure loss or vice versa, this result strongly suggests that syn-
apses are lost with concomitant loss of dendritic spines with
Rac1N17 expression. It will be interesting to determine whether
Rac1 activity directly regulates the development and maintenance
of the postsynaptic density or whether it acts on the overall integ-
rity of the dendritic spine itself.
Rac has been shown to regulate a number of biological processes
including lamellipodia formation, transcription regulation, and cell
cycle progression via activation of distinct effectors and down-
stream signaling pathways (Van Aelst and D’Souza-Schorey, 1997).
We attempted to define the downstream signal transduction path-
ways by using an activated Rac1 with specific effector domain
mutations shown previously to bind to a subset of effectors and
have a subset of biological effects (Joneson et al., 1996; Lamarche
et al., 1996). We found that the effector domain mutant shown
previously to block the ability of Rac1L61 to induce lamellipodial
formation in fibroblasts (Rac1L61A37) eliminates Rac1L61-
induced morphological changes in pyramidal neurons. The similar
properties of Rac1L61A37 on spine morphogenesis and lamellipo-
dial formation, along with the fact that Rac proteins are found
throughout the dendritic tree (Fig. 3A), suggest that Rac’s role in
spine morphogenesis is more likely via local regulation of the actin
cytoskeleton.
Generation and maintenance of dendritic branches
Previous studies have implied that the Rho GTPases have different
effects on the growth of axons and neuronal processes from neu-
ronal cell lines and primary neuronal cultures. It has been generally
thought that although Rac and Cdc42 have positive effects on
process extension, Rho has a negative effect on process outgrowth
or a positive effect on process retraction. Thus Rho activation
opposes the effect of Rac (see Kozma et al., 1997; Van Leeuwen et
al., 1997). An exception to this generalization was reported by
Threadgill et al. (1997), who showed similar effects of Rho and that
of Cdc42 and Rac on dendritic as well as axonal growth in disso-
ciated mammalian cortical neurons in culture.
We investigated the effects of Rac1 and RhoA in the regulation
of dendritic branch dynamics in neurons that have a relatively
mature and established dendritic tree (Fig. 2 B). Expression of
dominant-negative and constitutively active Rac1, although pro-
foundly affecting dendritic spines, had a relatively mild effect on the
overall dendritic tree complexity (Figs. 3, 4). In contrast, expression
of activated RhoA resulted in a drastic reduction of dendritic
branches (Fig. 5B,E,G).
Several lines of evidence suggest that the dendritic simplification
associated with RhoAV14 expression is caused by the retraction of
existing dendritic branches. First, at the time of transfection the
dendrites are more complex than after 24 hr of RhoAV14 expres-
sion (compare Figs. 2B, 5B). The dynamics of dendrite addition
and elimination (Dailey and Smith, 1996) is too slow for the simple
blockage of new dendrite formation to explain the degree of
simplification observed after1dofRhoAV14 expression. Second,
if the elimination of existing dendrites were caused by local degen-
eration of dendrites, one would expect to catch remnants of such
events. Although we occasionally observed this in all transfection
conditions when the slice culture was not healthy, we observed no
increase of such remnants above background associated with
RhoAV14-transfected neurons. Finally, in most of the RhoAV14-
expressing neurons, we observed a unique enlargement at the end
of their dendrites. It appeared that the dendrites have an aggrega-
tion of cytoplasm and plasma membrane at the tip of the simplified
dendrites. The detailed process involved in the reduction of the
dendritic branches and the identity of the enlarged end structure
remain to be characterized by live imaging and electron micros-
copy, respectively.
Several different effectors have been identified for RhoA signal-
ing to the actin cytoskeleton (for review, see Narumiya, 1996; Van
Aelst and D’Souza-Schorey, 1997). In particular, a multidomain
serine/threonine kinase ROCK has been shown to mediate RhoA’s
effect on cell rounding and process retraction from cultured neu-
roblastoma cells (Hirose et al., 1998). This RhoA-mediated retrac-
tion is likely to result from RhoA regulation, via ROCK, of myosin
light chain phosphorylation and actomyosin contractility (Jalink et
al., 1994; Kimura et al., 1996; Hirose et al., 1998). We have
extended these cell culture studies into hippocampal pyramidal
neurons in slice cultures that retain a relatively native environ-
ment. We found that treatment with the ROCK inhibitor Y-27632
(Uehata et al., 1997) effectively blocked the dendritic simplification
caused by RhoAV14 expression, indicating that ROCK is necessary
for Rho-mediated dendritic branch reduction. The fact that expres-
sion of an activated ROCK mutant mimics the effect of RhoAV14
expression further supports the importance of the RhoA–ROCK
pathway in mediating dendritic branch elimination.
Bito et al. (2000) have demonstrated recently that the RhoA–
ROCK pathway is vital for axonogenesis in cerebellar granule cells.
Despite the expression of RhoA mRNA and the highly penetrant
dendritic simplification phenotype with either activated RhoA or
ROCK, we did not detect any significant effect of inhibiting the
activity of RhoA (by expressing dominant-negative RhoAN19) or
ROCK (by application of Y-27632) on dendritic growth and
branching complexity. One explanation is that RhoAN19 is not
sufficient to block endogenous Rho activity, although similar treat-
ment has been shown to be effective in blocking axonogenesis (Bito
et al., 2000). We favor the hypothesis that the RhoA–ROCK
pathway, although intact in these neurons, is primarily inactive
during normal physiological conditions, thus allowing for the main-
tenance of the dendritic tree (and their spines). Inhibition of a
primarily inactive pathway would not result in detectable conse-
quences. The fact that inhibition of ROCK activity by itself does
not result in any significant change of dendritic complexity, but can
potently inhibit the effect of RhoA activation, strongly supports this
possibility. Such an intact yet dormant signaling pathway may be
useful in reorganizing local dendritic branches in response to local
activation of RhoA.
In a separate study, we showed that removal of RhoA activity by
null mutations of RhoA in Drosophila mushroom body neurons in
mosaic animals resulted in an overextension of dendrites. Con-
versely, expression of activated RhoA in mushroom body neurons
resulted in a significant reduction of dendritic field volume and
complexity (Lee et al., 2000). Taken together, these studies indicate
that RhoA plays an evolutionarily conserved role in limiting den-
dritic growth and complexity.
Functional significance of dendritic structural changes
in normal physiology and in mental retardation
One implication from our study is that molecules and mechanisms
used for initial dendritic morphogenesis may be reused in maturing
neurons for the continuous reorganization of neuronal cytoarchi-
tecture, possibly mediating plasticity in response to environmental
changes. We found that similar to their effect in development, Rac1
5336 J. Neurosci., July 15, 2000, 20(14):5329–5338 Nakayama et al. •Rac and Rho Maintain Dendritic Spines and Branches
had a positive effect on dendritic outgrowth (at the level of den-
dritic spines and filopodia-like extensions), although hyperactiva-
tion of Rac could lead to disruption of normal spine morphology.
On the other hand, RhoA hyperactivation had a negative effect on
the maintenance of dendritic branches. Recent findings demon-
strate that dendritic spines can undergo dynamic changes in re-
sponse to synaptic stimulation under conditions that generate long-
term potentiation (Engert and Bonhoeffer, 1999; Toni et al., 1999)
or to local increases in the neurotrophin BDN F (Horch et al.,
1999). Estrogen treatment also increases dendritic spine density in
vivo (Woolley and McEwen, 1993). Regulation of Rac GTPase
activity may be one way these extracellular factors and synaptic
activity are able to exert their effects on dendritic spines. It is
noteworthy that links between RhoA and neurotrophin receptor
p75
NTR
and NMDA have been suggested by two recent studies
(Yamashita et al., 1999; Li et al., 2000).
One consequence of how the misregulation of the Rho family of
GTPases may adversely affect the physiology of neurons comes
from the recent identification of genes responsible for nonspecific
X-linked mental retardation (for review, see Chelly, 1999). T wo of
the first three identified genes encode components of the Rho
GTPase-signaling pathways, oligophrenin (Billuart et al., 1998) and
Pak3 (Allen et al., 1998). Oligophrenin possesses a GAP domain
for Rho GTPases and has GAP activity in vitro for Rho, Rac, and
Cdc42 (Billuart et al., 1998). Loss of GAP activity would increase
the activity of Rho GTPases and would lead to disruption of spine
morphology or pruning of dendritic branches according to our
present study. Pak3 belongs to a family of p21-activated kinases
that was identified as effectors of Rac and Cdc42 (Manser et al.,
1994). Although our effector domain mutant analysis did not im-
plicate Pak as directly responsible for mediating the effect of
activated Rac1 on the disruption of spine morphology, Pak has also
been shown recently to act upstream of Rac (Obermeier et al.,
1998). Pak can also act downstream of Rac1 indirectly via associ-
ation with the p35/Cdk5 complex, whose binding to Rac1L61 is
abolished by both F37A and Y40K mutations (Nikolic et al., 1998).
Dominant-negative C dk5 has been shown to block the effects of
Rac1 in axonogenesis (Ruchhoeft et al., 1999). Taken together,
these findings suggest that the regulation of Rho GTPase-signaling
pathways is important for the generation and maintenance of
neuronal morphology that may eventually contribute to proper
mental function.
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