Abnormalities in cytoskeletal organization are a common
feature of many neurodegenerative disorders, including
Alzheimer’s disease (AD). Pathological actin in a polymerized
conformation (F-actin) has been found throughout Hirano
bodies (Galloway et al., 1987; Goldman, 1983), which are
cytoplasmic inclusions found in several neurodegenerative
diseases, including AD (Gibson and Tomlinson, 1977). Actin
is also a component of cofilin-actin rods [inclusion-like
structures described in hippocampal and cortical neurons of
post-mortem AD brains and also induced by certain chemical
or physical stresses in cultured cells (Minamide et al., 2000)],
which are thought to be associated with neurodegeneration.
The Rho family of small GTPases (Rho, Rac and Cdc42) are
regulators of F-actin polymerization (Bishop and Hall, 2000),
acting as molecular switches by cycling between an inactive
GDP-bound state and an active GTP-bound state. Rac1 and
Cdc42 promote polymerization at the leading edge, orchestrating
the formation of lamellipodia and membrane ruffles (Ridley et
al., 1992), as well as peripheral actin microspikes and filopodia
(Kozma et al., 1995; Nobes and Hall, 1995). RhoA is an
antagonist, promoting retraction of the leading edge and
assembly of stress fibers (Schmitz et al., 2000).
Rho GTPase activation at the plasma membrane depends on
the geranylgeranyl lipid residue attached to the C terminus of
the GTPase proteins (Seabra, 1998) and requires the interaction
with guanine nucleotide exchange factors (GEF) that can
stimulate the exchange of GDP for GTP, allowing the Rho-
family GTPases to bind to and activate their effector proteins.
Tiam1 is a protein that functions as a GEF for Rac1 GTPase,
both in vitro and in vivo (Habets et al., 1994). Tiam1 requires
phosphorylation at threonine residues by calcium/calmodulin
kinase II (CaMK-II) and protein kinase C (PKC) that may elicit
the activation and/or translocation of the protein to the
membrane compartment (Fleming et al., 1999; Fleming et al.,
1997). Tiam1 translocation to the plasma membrane induces
membrane ruffles and JNK stimulation mediated by Rac1,
suggesting that the control of membrane association of Tiam1
is an important factor in Rac1 activation (Michiels et al., 1997).
Deregulation of the Rho GTPase pathway is implicated in
several pathological conditions, including neurodegenerative
disorders like AD. This pathology is characterized by a
progressive loss in the number of dendritic spines, as well as
by alterations in the synaptic efficacy and damage at the
synaptic terminal (Lippa et al., 1992; Masliah, 1995). The
dynamic regulation of actin polymerization is considered the
main mechanism underlying morphological changes in
dendritic spines (Matus, 2000; Halpain, 2000). Rac1/Cdc42
Rho GTPases, which are the main regulators of F-actin
polymerization, have been implicated in the maintenance and
reorganization of dendritic structures (Nakayama et al., 2000;
Luo, 2000). Neuronal populations of AD patients showed a
considerable overlap of Rac/Cdc42 with early cytoskeletal
abnormalities, as well as Rac/Cdc42 upregulation in cases of
AD in comparison with an age-matched control (Zhu et al.,
2000). We found an increase in Rac1 immunoreactivity in
cortex sections derived from Tg2576 (Otth et al., 2003), a
transgenic mouse model expressing the Swedish mutation of
the human amyloid precursor protein that causes memory
deficits and plaque accumulation with age (Hsiao et al., 1996).
Rac1 is also an essential element of the amyloid-beta signaling
cascade that leads to the generation of ROS in astroglioma cells
(Lee et al., 2002).
In neurons, amyloid-beta (A?) peptide (the main constituent
of the neuritic plaque) destabilizes Ca2+regulation and renders
neurons more vulnerable to environmental stimuli that elevate
A number of psychiatric and neurodegenerative disorders,
such as Alzheimer’s disease, are characterized by
abnormalities in the neuronal cytoskeleton. Here, we find
that the enhancement in actin polymerization induced by
fibrillar amyloid-beta peptide (A? ?) is associated with
increased activity of Rac1/Cdc42 Rho GTPases. Rac1
upregulation involves the participation of Tiam1, a Rac
guanine-nucleotide exchange factor, where A? ? exposure
leads to Tiam1 activation by a Ca2+-dependent mechanism.
These results point to Rho GTPases as one of the targets in
A? ?-induced neurodegeneration in Alzheimer’s disease
pathology, with a role in mediating changes in the actin
Supplementary material available online at
Key words: Alzheimer’s disease, Amyloid beta, Rac1 Cdc42 Rho
GTPases, Actin polymerization, Tiam1
A? ?1-42stimulates actin polymerization in hippocampal
neurons through Rac1 and Cdc42 Rho GTPases
Ariadna Mendoza-Naranjo1,*, Christian Gonzalez-Billault1and Ricardo B. Maccioni1,2
1Laboratory of Cellular, Molecular Biology and Neuroscience, Department of Biology, Faculty of Sciences and 2Department of Neurological
Sciences, Universidad de Chile, Las Palmeras 3425, Nunoa, Santiago, Chile
*Author for correspondence (e-mail: firstname.lastname@example.org)
Accepted 30 October 2006
Journal of Cell Science 120, 279-288 Published by The Company of Biologists 2007
Journal of Cell Science
Correspondingly, it has also been seen that an increase in
induces membrane translocation and
activation of Rac, an event dependent on the activation of
conventional protein kinase C (PKC) (Price et al., 2003). In
Swiss 3T3 fibroblasts, the Ca2+chelator BAPTA-AM totally
abrogated Tiam1 phosphorylation induced by PDGF,
indicating that Ca2+is essential for Tiam1 activation (Fleming
et al., 1998). These results suggest that Ca2+is an essential
regulator of Rac/Tiam1signaling.
We have studied alterations in the actin cytoskeleton in A?-
stimulated neurons and the involvement of Rac/Cdc42 Rho
GTPases and associated regulators in these changes.
levels (Mattson etal., 1993).
A? peptide increases F-actin in hippocampal cells
The presence of structures characterized by aggregates of
polymerized actin (F-actin) has been described in AD. In order
to test whether A? presence could be related with the
formation of this kind of structures, we cultured hippocampal
cells in the presence of 10 ?M A?1-42, an accepted
neurodegeneration model for AD. Such A? concentrations
have been detected in brain tissues of AD patients and mice
transfected with human amyloid precursor protein gene (Kuo
et al., 1999; McLean et al., 1999; Kawarabayashi et al., 2001;
Funato et al., 1998), and are pathophysiologically relevant. The
F-actin content was examined by Rhodamine-phalloidin
staining. An antibody that recognizes tyrosinated tubulin
(TyrTub) was also added, as a dynamic microtubules marker.
After short periods of fibrillar peptide exposure (30 minutes to
2 hours) no visual changes in actin polymerization were
detected in neurons stained with Rhodamine-phalloidin (data
not show). Four hours after A? stimulation, a noticeable
increase of F-actin was observed in comparison to control
neurons (Fig. 1A), without changes in the TyrTub stain. This
increase in polymerized actin was reflected by an augmentation
in the formation of membrane ruffles (Fig. 1A, arrowheads),
as well as in filopodia and lamellipodia, in minor processes and
in the distal tip of the growing axon (Fig. 1A, arrows). In order
to quantify the observed increase, we measured the F-actin
content in control and A?-treated hippocampal neurons
immunostained with Rhodamine-phalloidin. This analysis
revealed a statistically significant increase of over 60% in the
amount of F-actin after A? treatment (Fig. 1B).
We also measured changes in the F-actin content after
A?1-42stimulation, using an In Vivo Actin Dynamic Assay kit.
Four hours after the treatment with A?, the same time at which
a significant increase in actin polymerization was observed
(Fig. 1A), a 40% rise in F-actin content was detected, and
maintained after 24 hours of A?-treatment (Fig. 1C). In
addition, the cell homogenate was incubated with phalloidin,
as a positive control for F-actin formation, displaying higher
amounts of F-actin than control and A?-treated cells (Fig. 1C,
lane P). To further analyze these changes in actin cytoskeleton,
morphometric parameters, such as filopodia number and
growth cone area, were measured. Fig. 2 shows that the
enhancement in F-actin polymerization after A? treatments
was accompanied by a significant increase in the size of the
growth cone area (P<0.05), as well as in the number of
filopodia (P<0.001), when compared with controls.
Activation of Rac1 and Cdc42 is responsible for
increased F-actin in A?-treated cells
Rho GTPases are key regulators of F-actin polymerization. In
Journal of Cell Science 120 (2)
Fig. 1. F-actin levels are increased in
hippocampal neurons treated with A?. (A)
Control hippocampal neurons grown on
poly-D-lysine for 4 days were stimulated
with 10 ?M of fibrillar A?1-42for 4 hours
and double stained for Tub-Tyr (green) and
F-actin filaments with Rhodamine-
phalloidin. Overlay of Tub-Tyr with
phalloidin is shown. Scale bar, 20 ?m. (B)
Hippocampal cells, untreated or stimulated
for 4 hours with 10 ?M A?1-42were
immunostained with phalloidin-TRITC and
the F-actin intensity was analyzed. The
values are expressed as the main ± s.e.m.
Data are representative for three different
experiments, with at least 12
determinations per experiment (*P<0.05).
(C) F-actin levels in control and 4-hour and
24-hour A?-stimulated neurons were
measured with Actin Polymerization Assay
kit. Phalloidin (P) was used as a positive
control. The ratio of F-actin/total actin was
determined from the blots by densitometric
measurements. Significant differences are
indicated by asterisks (*P<0.05,
***P<0.005; Student’s t-test).
Journal of Cell Science
A? induces Rac1/Cdc42 activation in neurons
order to analyze whether F-actin increased polymerization by
A? is dependent of Rac1/Cdc42 activation, PBD [p21-
activated kinase-(PAK)-binding domain] was used. The
amount of activated Rac1/Cdc42 was determined after
treatment with different doses of the amyloid peptide.
Activation of Rac was dose dependent, so that 10 ?M A?1-42,
the highest concentration tested, rendered the greatest
Rac/Cdc42-GTP levels (Fig. 3A; see also supplementary
material Fig. S1). Quantitative analysis indicated that the active
pool of Rac1 increased 1.6-fold after the addition of A? in
comparison to the untreated condition (Fig. 3A). This effect
was time-dependent with significant differences found between
the control and 4 hours, as well as between the control and 24
hours of peptide incubation (Fig. 3B). In parallel, A?1-42
increased the activation of Cdc42 also in a time- and dose-
dependent manner (see supplementary material Fig. S1).
To verify the specificity of the activation of Rac1 and Cdc42
GTPases in the signaling pathway mediated by the amyloid
peptide, the addition of A?1-42was combined with pertussis
toxin (PTX), a G-protein inhibitor affecting Rac1 and Cdc42
activation (Van Leeuwen et al., 2003). PTX partially inhibited
the increases in Rac1 and Cdc42 after A? addition (Fig. 3C,D).
Cyclin-dependent kinase 5 (Cdk5), which is deregulated in
Alzheimer brains (Alvarez et al., 1999; Patrick et al., 1999) and
probably contributes to the pathogenesis of the disease,
colocalizes with Rac1 in neuronal growth cones (Nikolic et al.,
1998). Hippocampal cells treated with A?1-42 along with
roscovitine, a Cdk5 inhibitor, exhibited the same extent of
Rac1 activation as those stimulated without the inhibitor (Fig.
To further investigate the role of Rac1 and Cdc42 in A?
induction of actin polymerization, changes in the distribution
of both proteins and their relationship with F-actin after
stimulation with the peptide were examined. A? treatments
induced actin remodeling and increased the amount of actin-
dependent structures (Fig. 4A, arrows), accompanied by
enhanced colocalization of Rac1 and Cdc42 with F-actin-rich
domains (Fig. 4A, arrowheads). The extent of A?-induced
Rac1-F-actin and Cdc42-F-actin colocalization was evaluated
in two-dimensional scatter analyses of fluorescence intensity
distribution. Fig. 4B shows that A? stimulation promoted an
increase in the colocalization of Rac1 and Cdc42 with
polymerized actin. In order to confirm this increase, we
determined the colocalization coefficient obtained from the
merged images that display the intensity and distribution of red
and green pixels. Amyloid-treated neurons stained for Rac1
and phalloidin displayed higher colocalization coefficients than
those showed for control untreated neurons (Fig. 4C). Similar
results were obtained for Cdc42 and phalloidin-labeled
Translocation of Rac1 and Cdc42 to the cell plasma
membrane is essential for activating downstream effectors. To
determine whether Rac1 and Cdc42 are recruited to the cell
membrane after A? stimulation, hippocampal cultures were
fixed after detergent extraction, which was performed under
microtubule-stabilizing conditions. This method removes
soluble proteins from the cell in a way that proteins remaining
are attributable to the polymerized fraction (Brown et al.,
1992). Samples were processed for immunofluorescence in
cells fixed after detergent
immunostaining with Rac1 or Cdc42 antibodies, along with
Rhodamine-phalloidin. Rac1 and Cdc42 were mostly localized
in the neuronal body and throughout the processes, and were
preferentially recruited to the cell membrane after A?
stimulation (Fig. 5A, green stain), where they were frequently
found colocalizing with Rhodamine-phalloidin (Fig. 5A,
In order to verify the results from the immunofluorescence
experiments, Rac1 and Cdc42 association with the membrane
fraction was analyzed, using subcellular fractionation
protocols. When hippocampal neurons were stimulated with 10
?M fibrillar A?1-42an increase in the translocation of both
proteins from the cytosolic to the membrane fraction was
observed, characterized by the presence of flotillin (Fig. 5B),
reinforcing the results of GTPases activation by amyloid
treatments. Membrane fractions were negative for NF?B,
confirming that these fractions were devoid of cytoplasmic
contamination (data not shown).
Rac1 and Cdc42 activity can be changed by mutations
leading to either constitutively active or dominant-negative
forms. To further demonstrate the involvement of these proteins
extraction, by double
Fig. 2. Increase in F-actin levels correlates with enhancement in
growth cone area and filopodia number. Hippocampal neurons
control and stimulated with A?1-42were immunostained with
phalloidin-TRITC (top panels; higher magnification of the boxed
regions is shown in the insets) and growth cone area and filopodia
number were analyzed (bottom panels). Scale bar, 20 ?m. Values are
expressed as mean ± s.d.
Journal of Cell Science
in the signaling pathway activated by A? that leads to increased
actin polymerization, Rac1 and Cdc42 were inhibited by
transiently expressing their dominant negative constructions.
For this purpose, RacT17N-GFP and Cdc42T17N-GFP were
transfected in neuronal cells stimulated with A? and in
unstimulated cells. Transfections done with RacT17N-GFP or
Cdc42T17N-GFP were able to inhibit increased F-actin (stained
with Rhodamine-phalloidin) for untreated (data not shown) as
well as for A?1-42-stimulated cells (Fig. 6A, arrow), compared
with neighboring untransfected cells (Fig. 6A, arrowheads).
This effect was not obtained by transfecting GFP alone or
constructs expressing wild-type or constitutive active forms of
Rac1 and Cdc42 (data not shown). The F-actin decrease was
then estimated by analyzing the fluorescence of Rhodamine-
phalloidin staining. A decrease in F-actin of about 10% was
obtained for control hippocampal neurons transfected with
Rac1- and Cdc42-T17N forms, with respect to untransfected
cells (Fig. 6B). For A?-treated hippocampal cells the difference
between transfected and untransfected neurons was over 50%
after the expression of the dominant negative forms of Rac1 and
Cdc42 (Fig. 6B). These results demonstrate that Rac1 and
Cdc42 are participating in the increase of actin polymerization
mediated by A?1-42.
A? mediates Rac1 activation through a mechanism
dependent on Tiam1
Tiam1 requires translocation to the plasma membrane to
induce cytoskeletal changes mediated by Rac1 (Michiels et al.,
1997). To investigate whether Tiam1 is involved in A?-induced
Rac1 stimulation, we examined the recruitment of Tiam1 from
cytosolic to plasma membrane fraction. As shown in Fig. 7A,
the membrane fraction is enriched 1.5-fold in Tiam1 after
A?1-42 treatment; this was time-dependent and there was a
significant difference with controls, 4 hours after exposure to
A?1-42. This was the same time point at which Rac1 activation
was found to be significantly increased (Fig. 3B).
Phosphorylation of Thr residues of Tiam1 is another event that
may elicit the activation and/or membrane translocation of the
protein. In order to investigate the Tiam1 activation state we
analyzed Tiam1 phosphorylation on Thr residue by using an
antibody that recognizes phospho-Thr epitopes on total
immunoprecipitated Tiam1. Fibrillar A?1-42
significant increase (1.5-fold) in the amount of Thr
phosphorylation of Tiam1 (Fig. 7B).
Ca2+signaling controls Tiam1 activation (Fleming et al.,
1998), and also regulates translocation and activation of Rac
(Price et al., 2003). We explored whether A?-induced Tiam1
Journal of Cell Science 120 (2)
Fig. 3. GTPase activities of
Rac1 and Cdc42-GTPase are
increased in neurons stimulated
with A?1-42. Hippocampal
neurons cultured for 4 days
were treated with (A) 0.1, 1 and
10 ?M A?1-42for 24 hours and
(B) with 10 ?M of peptide for
30 min, 2, 4 and 24 hours.
Active GTP-Rac1 was pulled
down using the PAK-PBD
conjugated with agarose and
then tested by immunoblotting
with anti-Rac1 monoclonal
antibody. Values were
normalized against total Rac1.
Graphs show data from four
(C,D) Hippocampal neurons
stimulated 4 hours with 10 ?M
fibrilar A?1-42were pre-treated
with pertussis toxin and
roscovitin. The GTP bound to
Rac1 (C) and to Cdc42 (D) was
determined using the
Rac/Cdc42 Activation Assay
Kit, according to
recommendations. Values were
normalized, with respect to
total Rac1 and Cdc42;
Journal of Cell Science
A? induces Rac1/Cdc42 activation in neurons
and Rac1 activation is dependent on increases in intracellular
Ca2+. Fig. 8A shows a rapid enhancement of intracellular Ca2+
in neurons exposed to fibrillar A?1-42. Such enhancement was
sensitive to the addition of BAPTA-AM, which blocked this
rise. In order to test whether Ca2+affects Tiam1 activation by
A?1-42, Tiam1-threonine phosphorylation was examined by
combining the addition of the fibrillar peptide with BAPTA-
AM. Treatments with the Ca2+chelator returned Tiam1-
phosphothreonine expression to control levels (Fig. 8B). To
confirm the Ca2+
involvement in A?-mediated Tiam1
activation, we analyzed whether BAPTA was affecting the
amount of Tiam1 recovered after Rac1 was pulled down in a
GTP-active form. A? treatments induce a higher association
ofTiam1 with active Rac1.
immunoprecipitation was significantly reduced by combining
BAPTA with A? treatments (Fig. 8C). PKC also seems to play
a key role in Tiam1 activation (Fleming et al., 1997; Buchanan
et al., 2000). A? treatments combined with the addition of
Gö6976, a PKC inhibitor, also diminished Tiam1 co-
immunoprecipitation with active Rac1 (Fig. 8C).
Calcium-dependent Rac activation involves the participation
of a conventional PKC (Price et al., 2003). With the purpose
This increased co-
of evaluating Ca2+and PKC involvement in A?-mediated Rac1
activation, Rac1-GTP levels were examined in hippocampal
cells stimulated with A?1-42, along with BAPTA or Gö6976.
The increase of Rac1 activation after A?1-42 stimuli was
inhibited when neurons were incubated with the peptide in the
presence of BAPTA-AM (Fig. 9A). Inhibition of PKC also
blocked the activation of Rac1 mediated by A?1-42(Fig. 9B).
Stress stimuli such as A?1-42, the main component of the
amyloid plaque in AD, may alter the neuronal cytoskeleton.
Here, we demonstrated that fibrillar A?1-42causes alterations in
cytoskeletal actin in hippocampal cells, represented by an
increase in the F-actin content. Alzheimer pathology is
characterized by dramatic synapse and dendritic spine loss, as
well as by damage in the synaptic terminal (Lippa et al., 1992;
Masliah, 1995), where decreased cortical synapse density
correlates with cognitive decline in patients (Terry et al., 1991;
DeKosky et al., 1996). A? has been described to affect and
accumulate in synapses (Takahashi et al., 2004). Likewise, a
study using in vivo electrophysiology in the Tg2576 mouse
model of AD showed disrupted cortical synaptic integration,
Fig. 4. Rac1 and Cdc42
display enhanced F-actin
colocalization in hippocampal
cells treated with A?.
(A) Four-day hippocampal
neurons, untreated (control)
and exposed to A?1-42fibrils
for 4 hours were double
immunostained with Rac1 or
Cdc42 and Rhodamine-
phalloidin and analyzed by
microscopy. Images in the
upper panel are overlays of
Rac1 or Cdc42 vs.
staining. Arrows indicate
increased actin protusions in
Lower panels are higher
magnification of the boxed
regions showing Rac1 and
Cdc42 overlaying with F-actin
in control conditions, as well
as the augmented
colocalization of these
proteins with F-actin
(arrowhead) after A? stimulation. Scale bar, 20 ?m. (B) Respective scatter plots of
fluorescence intensity distribution for Rac1/F-actin and Cdc42/F-actin from control
and A?-stimulated neurons. (C) Colocalization coefficients between Rac1/F-actin
and Cdc42/F-actin were evaluated using Zeiss colocalization coefficient function
software. Data are expressed as mean ± s.e.m.
Journal of Cell Science
which correlated with plaque formation (Stern et al., 2004). In
a recent report using the Tg2576 transgenic mouse model, the
same group described that spine loss is most pronounced near
amyloid plaques, indicative of focal toxicity (Spires et al.,
In this respect, the dynamic control of actin polymerization
is considered critical for synaptic regulation and spine structure
control (Matus, 2000; Halpain, 2000). Imbalance in actin
dynamics has also been established to contribute to the
formation of actin rods, leading to neurodegeneration
(Minamide et al., 2000). Rod induction occurring in neurons
exposed to stress accompanies
neurodegenerative conditions. Alzheimer brain sections
display rod-like inclusions, especially in areas surrounding the
amyloid plaque, reinforcing the idea that A? peptide may play
a role in the imbalance of actin dynamics (Maloney et al.,
The dynamic actin-rich nature of dendritic spines has
pointed to the Rho-GTPase family as a central contributor
(Matus, 2000; Hering and Sheng, 2001; Etienne-Manneville
and Hall, 2002), with Cdc42 and Rac1
regulating spine morphogenesis and synapse
formation in neurons (Irie and Yamaguchi,
2002; Zhang et al., 2003). Here, we find that
this augmented actin polymerization, as shown
by increased lamellipodia and filopodia
formation after A?1-42stimulus, is correlated
with time- and dose-dependent increases in
Rac1 and Cdc42 activity. The increased actin
transfected with either Rac1 or Cdc42
dominant-negative forms, confirming the
participation of these GTPases in the cascade
of cellular events underlying the enhancement of F-actin by
A?1-42. Rac1 and Cdc42 have also been seen upregulated in
neuronal populations of AD brains in comparison to controls
(Zhu et al., 2000), and simultaneously overlapped with early
cytoskeletal abnormalities, suggesting a physiological
connection between these two processes in the context of AD.
By contrast, a previous report demonstrated that Rac
activation inhibits the formation of both dendritic spines and
synapses (Zhang et al., 2003). In this sense, alterations of the
synaptic efficacy along with a progressive loss in the number
of dendritic spines, as have been described for AD pathology,
could be a consequence of Rac1/Cdc42 activity deregulation.
One of the mechanisms that may be acting to increase Rac1
GTPase activity, as shown in this study, is the activation of
Tiam1. Tiam1 was upregulated in both cortex and hippocampal
cells stimulated with A?1-42, reflected by increased threonine
phosphorylation and enhanced membrane localization. The
association of Tiam1 to the membrane accompanies its own
activation and regulates different Rac-mediated signaling
pathways, such as the induction of membrane ruffling through
inhibited in cells
Journal of Cell Science 120 (2)
Fig. 5. Rac1 and Cdc42 increased in the
plasma membrane after A?1-42
stimulation. (A) Four-day hippocampal
neurons, untreated (control) and exposed
to A?1-42fibrils for 4 hours, were fixed
after detergent extraction performed under
cytoskeleton-stabilizing conditions and
double immunostained with Rac1 or
Cdc42 and Rhodamine-phalloidin. Both
GTPases were recruited to the plasma
membrane and were frequently found to
colocalize with F-actin after amyloid
stimulation (arrowheads). Bar, 20 ?m.
(B) Embryonic hippocampal cells,
untreated and treated for 4 and 24 hours
with ?-amyloid, were lysed and
cytoplasmic and membrane fractions
obtained. Rho GTPases recruitment to the
membrane fraction was analyzed using
Rac1 and Cdc42 antibodies and the values
were normalized against flotillin as a
membrane protein control. Samples from
the cytoplasmic fraction were analyzed by
immunoblotting using Rac1 and Cdc42,
along with actin antibody as a loading
control. Significant differences are
indicated by asterisks (*P<0.05,
**P<0.01; Student’s t-test).
Journal of Cell Science
A? induces Rac1/Cdc42 activation in neurons
Rac1 (Michiels et al., 1997). In this
sense, we detected that treatment with
formation in hippocampal
implying that Tiam1 participates in A?-
addition, an increase
association with the active form of Rac1
(Rac-GTP) after stimulus with A?1-42
was established, confirming the direct
involvement of this GEF in Rac1
activation in the amyloigenic context.
However, our experiments do not
exclude the existence
regulators, and additional mechanisms
may also be playing a role.
Changes in Ca2+homeostasis, as
occurring after A?
responses contributing to neuronal
imbalance. Here, we demonstrate that Ca2+is involved in the
enhanced Tiam1 and Rac1 activity induced by A?. PKC may
also be acting, since PKC inhibition affects Tiam1 association
with the active form of Rac-coupling GTP, as well as Rac1
activity in hippocampal neurons stimulated with A?1-42.
Altogether these results illustrate that Tiam1 upregulation is
responsible for increased Rac1 activity in the hippocampus, by
stimulus with fibrillar A?.
Recent studies report more robust correlations
between the levels of soluble, rather than
insoluble, A? and the extent of synaptic loss and
severity of cognitive impairment (Lue et al.,
1999; McLean et al., 1999; Wang et al., 1999).
Fig. 6. Rac1 and Cdc42 are essential for the
increased F-actin polymerization induced by
A?1-42 treatment. Primary hippocampal
cells treated with A?1-42 were transfected
with the dominant negative forms of
RacT17N-GFP and Cdc42T17N-GFP
(arrows), and then stained for F-actin using
Rhodamine-phalloidin. The untransfected
neurons are indicated by arrowheads. Scale
bar, 20 ?m. (B) Changes in F-actin levels
were determined and differences between
transfected (t) vs. untransfected (ut) control
and A?-stimulated neurons were
determined using Student’s t-test. The
values were expressed as mean ± s.e.m. for
three different experiments, with at least 12
determinations per experiment; *P<0.05.
Fig. 7. Tiam1 is activated in A?-stimulated
hippocampal cells. (A) Membrane fractions from
hippocampal cells, either untreated (control) and
stimulated for 4 and 24 hours with 10 ?M of fibrillar
A?1-42, were obtained. Samples were analyzed by
immunoblotting using Tiam1 antibody and the values
were normalized against flotillin. Tiam1 expression in
the cytoplasmic fraction was analyzed by
immunoblotting using Tiam1 antibody and actin as a
loading control. (B) Tiam1 was immunoprecipitated
from control hippocampal neurons and neurons treated
for 4 hours with A?1-42. Phosphorylation levels were
tested by immunoblotting using P-Thr monoclonal
Journal of Cell Science
polymerization observed after fibrillar A?
counteract the synaptic damage by the
soluble form of the amyloid peptide,
rescuing neurons from the loss of synaptic
increase of actin
consequence of Rho GTPases deregulation
by A? stimulation, could end in an
exacerbated accumulation of F-actin. This event would be the
trigger for the formation of F-actin aggregates, as those
described in Hirano bodies in Alzheimer pathology (Gibson
and Tomlinson, 1977).
Our findings demonstrate a direct relationship of A? with
Rac1/Cdc42 Rho GTPases deregulation and subsequent actin
cytoskeletal alterations. These results are of interest because
one of the deleterious effects in the pathway to Alzheimer’s
disease could be related to alterations in the normal actin
turnover. In summary, these results points to the actin
cytoskeleton as a target for A?-induced neurodegeneration in
the increased actin
be generated to
as dynamics, a
Materials and Methods
Rhodamine-phalloidin, pertussis toxin, protein A-agarose, monoclonal antibodies
anti-actin and Tub-Tyr were from Sigma (Sigma-Aldrich). Polyclonal antibodies for
Cdc42 (P1), NFkB (H119), LIMK1 (C18) and Tiam1 (C16) were purchased from
Santa Cruz Biotechnology Inc. (Santa Cruz, CA, USA). Polyclonal anti-P-LIMK
was from Cell Signaling (Cell Signaling Technology, Beverly, MA). Monoclonal
flotillin 1 antibody was purchased from BD Transduction Laboratories (BD
Biosciences). Beta amyloid1-42peptide was from Global Peptide Services, CO. The
Rac/Cdc42 activation assay kit and monoclonal anti-Rac1 antibody were purchased
from Upstate Biotechnology Inc. (Lake Placid, NY). Phospho-Thr antibody was
from Biodesign International. The following secondary antibodies were used: anti-
mouse (Jackson ImmunoResearch), and anti-rabbit (Pierce Biotechnology), both
horseradish peroxidase conjugated; anti-rabbit FITC-coupled (Sigma-Aldrich) and
antimouse FITC-coupled (Jackson ImmunoResearch). BAPTA-AM, Gö6976 and
roscovitine were purchased from Calbiochem. The
Western Lighting Chemiluminescence Reagent
Plus was from Perkin Elmer. The Actin
Polymerization Assay Biochem Kit was from
Cytoskeleton (Denver, CO). Neurobasal (NB), B27,
Opti-MEM and Lipofectamine-2000 were from
Invitrogen-Gibco-BRL Life Technologies. All other
chemicals were obtained from usual commercial
sources at the highest grade available.
Journal of Cell Science 120 (2)
Fig. 8. A?-mediated Tiam1 activation is affected
by Ca2+signaling. (A) Time lapse experiments
were performed to analyze the Ca2+increase in
hippocampal cells after A?1-42addition, using
the fluorescent tracer Fluo3-AM. (B) Phospho-
Thr levels from total immunoprecipitated Tiam1
were tested by immunoblot, in neurons treated
with A?1-42in the presence of BAPTA-AM;
*P<0.05. (C) Hippocampal neurons stimulated
with A?1-42were analyzed for Tiam1 association
to active Rac1 after BAPTA-AM or Gö6976
treatments. Rac1 was pulled down using PAK-
PBD agarose and the co-immunoprecipitated
Tiam1 was tested by immunoblotting using anti-
Tiam1 antibody. Values were normalized against
total Tiam1 in the crude homogenized. *P<0.05.
Fig. 9. Tiam1 mediates Rac1 activation
through Ca2+signaling in A?-stimulated
hippocampal cells. Hippocampal neurons
were incubated for 4 hours with A?1-42in
the presence of BAPTA-AM (A) or
Gö6976 (B) and then the active Rac1
fraction was pulled-down using the PAK-
PBD binding domain. The active form of
Rac1 was revealed using Rac1
monoclonal antibody. **P<0.01.
Journal of Cell Science
A? induces Rac1/Cdc42 activation in neurons
Dissociated hippocampal and cortex cultures were prepared from E19 embryos of
Sprague-Dawley rats as described previously (Caceres et al., 1986). Cells were
plated onto poly-D-lysine-coated plates and coverslips and maintained with NB
plus 10% horse serum. Three hours after plating the medium was removed and
serum-free medium B27/NB was added. Hippocampal and cortical neurons were
maintained for 4 days before treatments.
Preparation of A? aggregated peptide
A?1-42 peptide was prepared by dissolving the lyophilized synthetic peptide in
DMSO (6.67%) and phosphate buffer at pH 7.4, diluted to a concentration of 1
?g/?l and aggregated according to the ‘Aged condition’ protocol (Pike et al., 1993)
with some modifications (25°C for 7 days under vigorous agitation). Fibrillar A?1-42
was characterized by negative staining with uranyl acetate and morphologically
analyzed by electron microscopy [JEOL 100SX (100 kV)].
Hippocampal neurons exposed to A?1-42fibrils for 4 hours were: (1) fixed with 4%
paraformaldehyde plus 4% sucrose in phosphate buffer, pH 7.2 for 10 minutes and
permeabilized with 0.1% Triton X-100; or (2) extracted with detergent to prepare
‘cytoskeleton fractions’ according to the procedure of Brown (Brown et al., 1992),
then washed for 30 seconds with PHEM buffer pH 6.9 (60 mM Pipes, 25 mM
HEPES, 10 mM EGTA, 2 mM MgCl2) followed by extraction (2 minutes) with 0.2%
saponin in PHEM buffer containing 10 mM taxol. The cells were then fixed for 20
minutes with warmed 2% paraformaldehyde-0.05% glutaraldehyde in PHEM buffer.
The cells were incubated with the respective antibodies in 2.5% BSA-PBS in a
humid chamber at 4°C overnight. Rhodamine-phalloidin was included for
visualization of F-actin. The cells were examined by confocal laser scanning
microscopy (LSM 510; Carl Zeiss MicroImaging, Inc.).
The concentration of F-actin in control untreated and 4 and 24 hours A?-treated
hippocampal cells was measured with an Actin Polymerization Assay Kit, according
to the manufacturer’s recommendations. The cells were homogenized in F-actin
stabilization buffer, pH 6.9 plus protease and phosphatase inhibitors. Phalloidin at
1 ?M was used as a positive control. Equal amounts of protein were: (a) analyzed
by western blotting using actin polyclonal antibody, and (b) centrifuged, first at 600
g to separate the nuclear fraction and then at 100,000 g for 60 minutes at 30°C. The
pellets, corresponding to the F-actin fractions, were re-suspended in ice-cold H2O
plus 1 ?M cytochalasin D. They were then incubated on ice for 1 hour to dissociate
F-actin. The resuspended pellets were gently mixed every 15 minutes. SDS sample
buffer was added and the fractions were separated by 10% SDS-PAGE,
electroblotted onto nitrocellulose sheets and visualized with anti-actin antibody,
with Western Lighting Chemiluminescence Reagent Plus (Perkin Elmer). The ratio
of F-actin/total actin bands was determined by scanning and quantifying
densitometry in a Kodak Digital Science 1D 3.0.2 densitometer.
For the semi-quantitative measurement of the F-actin content, control and treated
hippocampal neurons were immunostained with phalloidin-Rhodamin. All pictures
were analyzed with the Carl Zeiss LSM 510 Image Browser program. Fluorescence
intensity was calculated by averaging the fluorescence intensity of all pixels and
subtracting the non-specific background from outside the cell. Each value is the
mean (± s.e.m.) of three different experiments, with at least 12 determinations in
each experiment. Differences were evaluated with the Student’s t-test.
Colocalization coefficients were calculated using Zeiss colocalization coefficient
function software, where all pixels above background are taken into account and
the relative number of colocalizing pixels in channels (Ch2: green and Ch3: red)
are calculated compared to total number of pixels above threshold. For a particular
color (e.g. red), the colocalization coefficient represents the ratio of red pixel
intensities showing a green component divided by the sum of all red intensities.
Data are presented as a value between 0 and 1, where 0 indicates no colocalization
and 1 indicates that all pixels colocalize (Smallcombe, 2001), and are expressed as
mean ± s.e.m. Colocalization analysis data were obtained from more than 16 double-
labeled cells in three different experiments.
Morphometric parameters were evaluated in cells stained with Rhodamine-
conjugated phalloidin. The number of filopodia and the area of the growth cone
(expressed in ?m2) of hippocampal neurons stimulated with 10 ?M A?1-42for 4
hours were analyzed, using a Zeiss LSM 510 confocal microscope with the Carl
Zeiss LSM Image Examiner program. The growth cone was defined as the distal
part of the neurite where the diameter is twice that of the neurite itself (Bradke and
Dotti, 1997). Filopodial number was determined per growth cone. Processes and
growth cones were randomly selected.
Rac1 and Cdc42 activation assay
Hippocampal neurons were treated with 0.1, 1 and 10 ?M of fibrillar A?1-42for 24
hours as well as with 10 ?M A?1-42for 30 minutes, 2, 4 and 24 hours. The GTP
loading of Rac1 and Cdc42 was measured using a Rac1/Cdc42 Activation Assay
Kit, according to manufacturer’s recommendations. Values of Rac/Cdc42 activation
were expressed as the ratio of Rac1-GTP or Cdc42-GTP/total Rac1 or Cdc42 in the
crude extract. For some experiments, cells were treated with 10 ?g/ml of roscovitin
inhibitor or pre-treated 4 hours with pertussis toxin at 500 ng/ml, plus 24 hours
stimulation with 10 ?M A?1-42. Hippocampal neurons were also stimulated for 4
hours with 10 ?M A?1-42in the presence or absence of 15 ?M BAPTA-AM or 1
?M Gö6976, both incubated for 1 hour (BAPTA-AM and Gö6976 were added 3
hours after A? stimulus). Tiam1 pulled down with active Rac1 was evaluated using
anti-Tiam1 antibody. The ratio of Tiam1 pulled down with Rac-GTP/total Tiam1 in
the crude protein extract was determined by scanning and quantifying densitometry
in a Kodak Digital Science 1D 3.0.2 densitometer.
Subcellular fractionation experiments
Hippocampal neurons untreated or stimulated for 4 and 24 hours with 10 ?M A?,
as well as 4 hours with 10 ?M A? in the presence of 15 ?M BAPTA-AM incubated
1 hour, were lysed in STM buffer and the membrane fractions were obtained
according to the protocol of Nikolic et al. (Nikolic et al., 1998). Pellets and
supernatants were re-suspended in Laemmli 4? sample buffer and analyzed by
immunoblotting using Rac1, Cdc42, Tiam1, flotillin, actin and NF?B antibodies and
a Western Lighting Chemiluminescence Reagent Plus.
Protein extracts were prepared from control and 4 hours A?-treated hippocampal
neurons in RIPA buffer plus protease and phosphatase inhibitors as we previously
described (Alvarez et al., 2001). Tiam1 polyclonal antibody (4 ?l) was incubated
at 4°C overnight. The samples were separated by SDS-PAGE, electroblotted onto
nitrocellulose sheets, and visualized with P-Thr and Tiam1 antibodies using Western
Lighting Chemiluminescence Reagent Plus. Bands were quantified in a Kodak
Digital Science 1D 3.0.2 densitometer.
DNA plasmids and transfection
Plasmids encoding dominant negative mutant (T17N) of the GTPases Rac1 and Cdc42
were sub-cloned in a plasmid encoding green fluorescence protein pEGFP-C1
(Clontech, Palo Alto, CA) by a HindIII and ApaI digestion. These vectors were
transfected in primary hippocampal neurons, using Opti-MEM buffer and
Lipofectamine 2000, according to the manufacturer’s recommendations. After 4 hours
the medium was replaced by NB/B27 and the cells were maintained in culture for a
further 24 hours. The neurons were then stimulated with 10 ?M A?1-42for 4 hours.
Transfected and non-transfected neurons were analyzed by immunofluorescence.
Measurement of Ca2+transients
An LSM 510 laser-scanning confocal microscope was used for analysis of Ca2+
transients in hippocampal neurons loaded with Fluo3-AM [prepared by mixing of 2
?M dye solution in DMSO with Pluronic acid (20% in DMSO)]. The Fluo3-AM dye
was incubated for 30 minutes at 37°C and 5% CO2at a final concentration of 5 ?M
in Krebs-Ringer/HEPES buffer pH 7.4 (KRH; 115 mM NaCl, 5 mM KCl, 1 mM
potassium phosphate, 5 mM glucose, 1.5 mM CaCl2, 1.2 mM MgSO4, 25 mM HEPES
pH 7.4). After the dye was removed, the cells were washed twice with fresh KRH
buffer. Using a time-lapse series of images, Ca2+transients were analyzed before,
during, and after stimulation of the neurons with 10 ?M A?1-42fibrils and with 15 ?M
of BAPTA-AM Ca2+chelator.
Results were analyzed using with Student’s t-test. Results were expressed as mean ±
s.d., or mean ± s.e.m. P<0.05 was considered to be significant.
This work was supported by grants from FONDECYT 1050198 to
R.B.M. and by the Millennium Institute CBB. A.M.-N. was supported
by a doctoral fellowship from the Millennium Institute for Advanced
Studies in Cell Biology and Biotechnology (CBB). We are grateful to
Alexandra Ginesta for critically reading the manuscript.
Alvarez, A., Toro, R., Caceres, A. and Maccioni, R. B. (1999). Inhibition of tau
phosphorylating protein kinase cdk5 prevents beta-amyloid-induced neuronal death. FEBS
Lett. 459, 421-426.
Alvarez, A., Munoz, J. P. and Maccioni, R. B. (2001). A Cdk5-p35 stable complex is
involved in the beta-amyloid-induced deregulation of Cdk5 activity in hippocampal
neurons. Exp. Cell Res. 264, 266-274.
Bishop, A. L. and Hall, A. (2000). Rho GTPases and their effector proteins. Biochem. J.
Bradke, F. and Dotti, C. G. (1997). Neuronal polarity: vectorial cytoplasmic flow precedes
axon formation. Neuron 19, 1175-1186.
Brown, A., Slaughter, T. and Black, M. M. (1992). Newly assembled microtubules are
Journal of Cell Science
288 Download full-text
concentrated in the proximal and distal regions of growing axons. J. Cell Biol. 119, 867-
Buchanan, F. G., Elliot, C. M., Gibbs, M. and Exton, J. H. (2000). Translocation of the
Rac1 guanine nucleotide exchange factor Tiam1 induced by platelet-derived growth factor
and lysophosphatidic acid. J. Biol. Chem. 275, 9742-9748.
Caceres, A., Banker, G. A. and Binder, L. (1986). Immunocytochemical localization of
tubulin and microtubule-associated protein 2 during the development of hippocampal
neurons in culture. J. Neurosci. 6, 714-722.
DeKosky, S. T., Scheff, S. W. and Styren, S. D. (1996). Structural correlates of cognition
in dementia: quantification and assessment of synapse change. Neurodegeneration 5, 417-
Etienne-Manneville, S. and Hall, A. (2002). Rho GTPases in cell biology. Nature 420, 629-
Fleming, I. N., Elliott, C. M., Collard, J. G. and Exton, J. H. (1997). Lysophosphatidic
acid induces threonine phosphorylation of Tiam1 in Swiss 3T3 fibroblasts via activation
of protein kinase C. J. Biol. Chem. 272, 33105-33110.
Fleming, I. N., Elliott, C. M. and Exton, J. H. (1998). Phospholipase C-gamma, protein
kinase C and Ca2+/calmodulin-dependent protein kinase II are involved in platelet-derived
growth factor-induced phosphorylation of Tiam1. FEBS Lett. 429, 229-233.
Fleming, I. N., Elliott, C. M., Buchanan, F. G., Downes, C. P. and Exton, J. H. (1999).
Ca2+/calmodulin-dependent protein kinase II regulates Tiam1 by reversible protein
phosphorylation. J. Biol. Chem. 274, 12753-12758.
Funato, H., Yoshimura, M., Kusui, K., Tamaoka, A., Ishikawa, K., Ohkoshi, N.,
Namekata, K., Okeda, R. and Ihara, Y. (1998). Quantitation of amyloid beta-protein
(A beta) in the cortex during aging and in Alzheimer’s disease. Am. J. Pathol. 152,
Galloway, P. G., Perry, G. and Gambetti, P. (1987). Hirano body filaments contain actin
and actin-associated proteins. J. Neuropathol. Exp. Neurol. 46, 185-199.
Gibson, P. H. and Tomlinson, B. E. (1977). Numbers of Hirano bodies in the hippocampus
of normal and demented people with Alzheimer’s disease. J. Neurol. Sci. 33, 199-206.
Goldman, J. E. (1983). The association of actin with Hirano bodies. J. Neuropathol. Exp.
Neurol. 42, 146-152.
Habets, G. G., Scholtes, E. H., Zuydgeest, D., van der Kammen, R. A., Stam, J. C.,
Berns, A. and Collard, J. G. (1994). Identification of an invasion-inducing gene, Tiam-
1, that encodes a protein with homology to GDP-GTP exchangers for Rho-like proteins.
Cell 77, 537-549.
Halpain, S.(2000). Actin and the agile spine: how and why do dendritic spines dance? Trends
Neurosci. 23, 141-146.
Hering, H. and Sheng, M.(2001). Dendritic spines: structure, dynamics and regulation.Nat.
Rev. Neurosci. 2, 880-888.
Hsiao, K., Chapman, P., Nilsen, S., Eckman, C., Harigaya, Y., Younkin, S., Yang, F. and
Cole, G. (1996). Correlative memory deficits, Abeta elevation, and amyloid plaques in
transgenic mice. Science 274, 99-102.
Irie, F. and Yamaguchi, Y. (2002). EphB receptors regulate dendritic spine development via
intersectin, Cdc42 and N-WASP. Nat. Neurosci. 5, 1117-1118.
Kawarabayashi, T., Younkin, L. H., Saido, T. C., Shoji, M., Ashe, K. H. and Younkin,
S. G. (2001). Age-dependent changes in brain, CSF, and plasma amyloid (beta) protein
in the Tg2576 transgenic mouse model of Alzheimer’s disease. J. Neurosci. 21, 372-
Kozma, R., Ahmed, S., Best, A. and Lim, L. (1995). The Ras-related protein Cdc42Hs and
bradykinin promote formation of peripheral actin microspikes and filopodia in Swiss 3T3
fibroblasts. Mol. Cell. Biol. 15, 1942-1952.
Kuo, Y. M., Emmerling, M. R., Lampert, H. C., Hempelman, S. R., Kokjohn, T. A.,
Woods, A. S., Cotter, R. J. and Roher, A. E. (1999). High levels of circulating
Abeta42 are sequestered by plasma proteins in Alzheimer’s disease. Biochem. Biophys.
Res. Commun. 257, 787-791.
Lee, M., You, H. J., Cho, S. H., Woo, C. H., Yoo, M. H., Joe, E. H. and Kim, J. H. (2002).
Implication of the small GTPase Rac1 in the generation of reactive oxygen species in
response to beta-amyloid in C6 astroglioma cells. Biochem. J. 366, 937-943.
Lippa, C. F., Hamos, J. E., Pulaski-Salo, D., DeGennaro, L. J. and Drachman, D. A.
(1992). Alzheimer’s disease and aging: effects on perforant pathway perikarya and
synapses. Neurobiol. Aging 13, 405-411.
Lue, L. F., Kuo, Y. M., Roher, A. E., Brachova, L., Shen, Y., Sue, L., Beach, T., Kurth,
J. H., Rydel, R. E. and Rogers, J. (1999). Soluble amyloid beta peptide concentration
as a predictor of synaptic change in Alzheimer’s disease. Am. J. Pathol. 155, 853-862.
Luo, L. (2000). Rho GTPases in neuronal morphogenesis. Nat. Rev. Neurosci. 1, 173-
Maloney, M. T., Minamide, L. S., Kinley, A. W., Boyle, J. A. and Bamburg, J. R. (2005).
Beta-secretase-cleaved amyloid precursor protein accumulates at actin inclusions induced
in neurons by stress or amyloid beta: a feedforward mechanism for Alzheimer’s disease.
J. Neurosci. 25, 11313-11321.
Masliah, E. (1995). Mechanisms of synaptic dysfunction in Alzheimer’s disease. Histol.
Histopathol. 10, 509-519.
Mattson, M. P., Barger, S. W., Cheng, B., Lieberburg, I., Smith-Swintosky, V. L. and
Rydel, R. E. (1993). beta-Amyloid precursor protein metabolites and loss of neuronal
Ca2+ homeostasis in Alzheimer’s disease. Trends Neurosci. 16, 409-414.
Matus, A. (2000). Actin-based plasticity in dendritic spines. Science 290, 754-758.
McLean, C. A., Cherny, R. A., Fraser, F. W., Fuller, S. J., Smith, M. J., Beyreuther,
K., Bush, A. I. and Masters, C. L. (1999). Soluble pool of Abeta amyloid as a
determinant of severity of neurodegeneration in Alzheimer’s disease. Ann. Neurol. 46,
Michiels, F., Stam, J. C., Hordijk, P. L., van der Kammen, R. A., Ruuls-Van Stalle, L.,
Feltkamp, C. A. and Collard, J. G. (1997). Regulated membrane localization of Tiam1,
mediated by the NH2-terminal pleckstrin homology domain, is required for Rac-
dependent membrane ruffling and C-Jun NH2-terminal kinase activation. J. Cell Biol.
Minamide, L. S., Striegl, A. M., Boyle, J. A., Meberg, P. J. and Bamburg, J. R. (2000).
Neurodegenerative stimuli induce persistent ADF/cofilin-actin rods that disrupt distal
neurite function. Nat. Cell. Biol. 2, 628-636.
Nakayama, A. Y., Harms, M. B. and Luo, L. (2000). Small GTPases Rac and Rho in
the maintenance of dendritic spines and branches in hippocampal pyramidal neurons.
J. Neurosci. 20, 5329-5338.
Nikolic, M., Chou, M. M., Lu, W., Mayer, B. J. and Tsai, L. H. (1998). The p35/Cdk5
kinase is a neuron-specific Rac effector that inhibits Pak1 activity. Nature 395, 194-198.
Nobes, C. D. and Hall, A. (1995). Rho, rac, and cdc42 GTPases regulate the assembly of
multimolecular focal complexes associated with actin stress fibers, lamellipodia, and
filopodia. Cell 81, 53-62.
Otth, C., Mendoza-Naranjo, A., Mujica, L., Zambrano, A., Concha, I. I. and Maccioni,
R. B. (2003). Modulation of the JNK and p38 pathways by cdk5 protein kinase in a
transgenic mouse model of Alzheimer’s disease. NeuroReport 14, 2403-2409.
Patrick, G. N., Zukerberg, L., Nikolic, M., de la Monte, S., Dikkes, P. and Tsai, L. H.
(1999). Conversion of p35 to p25 deregulates Cdk5 activity and promotes
neurodegeneration. Nature 402, 615-622.
Pike, C. J., Burdick, D., Walencewicz, A. J., Glabe, C. G. and Cotman, C. W. (1993).
Neurodegeneration induced by beta-amyloid peptides in vitro: the role of peptide assembly
state. J. Neurosci. 13, 1676-1687.
Price, L. S., Langeslag, M., ten Klooster, J. P., Hordijk, P. L., Jalink, K. and Collard, J.
G. (2003). Calcium signaling regulates translocation and activation of Rac. J. Biol. Chem.
Ridley, A. J., Paterson, H. F., Johnston, C. L., Diekmann, D. and Hall, A. (1992). The
small GTP-binding protein rac regulates growth factor-induced membrane ruffling. Cell
Schmitz, A. A., Govek, E. E., Bottner, B. and Van Aelst, L. (2000). Rho GTPases:
signaling, migration, and invasion. Exp. Cell Res. 261, 1-12.
Seabra, M. C. (1998). Membrane association and targeting of prenylated Ras-like GTPases.
Cell Signal. 10, 167-172.
Smallcombe, A. (2001). Multicolor imaging: the important question of co-localization.
Biotechniques 30, 1240-1242, 1244-1246.
Spires, T. L., Meyer-Luehmann, M., Stern, E. A., McLean, P. J., Skoch, J., Nguyen, P.
T., Bacskai, B. J. and Hyman, B. T. (2005). Dendritic spine abnormalities in amyloid
transgenic mice demonstrated by gene intravital multiphoton microscopy. J. Neurosci. 25,
Stern, E. A., Bacskai, B. J., Hickey, G. A., Attenello, F. J., Lombardo, J. A. and Hyman,
B. T. (2004). Cortical synaptic integration in vivo is disrupted by amyloid-plaques. J.
Neurosci. 24, 4535-4540.
Takahashi, R. H., Almeida, C. G., Kearney, P. F., Yu, F., Lin, M. T., Milner, T. A. and
Gouras, G. K.(2004). Oligomerization of Alzheimer’s beta-amyloid within processes and
synapses of cultured neurons and brain. J. Neurosci. 24, 3592-3599.
Terry, R. D., Masliah, E., Salmon, D. P., Butters, N., DeTeresa, R., Hill, R., Hansen, L.
A. and Katzman, R.(1991). Physical basis of cognitive alterations in Alzheimer’s disease:
synapse loss is the major correlate of cognitive impairment. Ann. Neurol. 30, 572-580.
Van Leeuwen, F. N., Olivo, C., Grivell, S., Giepmans, B. N., Collard, J. G. and
Moolenaar, W. H. (2003). Rac activation by lysophosphatidic acid LPA1 receptors
through the guanine nucleotide exchange factor Tiam1. J. Biol. Chem. 278, 400-406.
Wang, J., Dickson, D. W., Trojanowski, J. Q. and Lee, V. M. (1999). The levels of soluble
versus insoluble brain Abeta distinguish Alzheimer’s disease from normal and pathologic
aging. Exp. Neurol. 158, 328-337.
Zhang, H., Webb, D. J., Asmussen, H. and Horwitz, A. F. (2003). Synapse formation is
regulated by the signaling adaptor GIT1. J. Cell Biol. 161, 131-142.
Zhu, X., Raina, A. K., Boux, H., Simmons, Z. L., Takeda, A. and Smith, M. A. (2000).
Activation of oncogenic pathways in degenerating neurons in Alzheimer disease. Int. J.
Dev. Neurosci. 18, 433-437.
Journal of Cell Science 120 (2)
Journal of Cell Science