Post on 17-May-2023
ORIGINAL PAPER
Cytoskeletal Alterations and Biomechanical Properties
of parkin-Mutant Human Primary Fibroblasts
Daniele Vergara • Marzia M. Ferraro • Mariafrancesca Cascione •
Loretta L. del Mercato • Stefano Leporatti • Anna Ferretta • Paola Tanzarella •
Consiglia Pacelli • Angelo Santino • Michele Maffia • Tiziana Cocco •
Ross Rinaldi • Antonio Gaballo
� Springer Science+Business Media New York 2014
Abstract Parkinson’s disease (PD) is one of the most
common neurodegenerative diseases. Genes which have
been implicated in autosomal-recessive PD include PARK2
which codes for parkin, an E3 ubiquitin ligase that par-
ticipates in a variety of cellular activities. In this study, we
compared parkin-mutant primary fibroblasts, from a patient
with parkin compound heterozygous mutations, to healthy
control cells. Western blot analysis of proteins obtained
from patient’s fibroblasts showed quantitative differences
of many proteins involved in the cytoskeleton organization
with respect to control cells. These molecular alterations
are accompanied by changes in the organization of actin
stress fibers and biomechanical properties, as revealed by
confocal laser scanning microscopy and atomic force
microscopy. In particular, parkin deficiency is associated
with a significant increase of Young’s modulus of null-cells
in comparison to normal fibroblasts. The current study
proposes that parkin influences the spatial organization of
actin filaments, the shape of human fibroblasts, and their
elastic response to an external applied force.
Keywords Parkin � Cofilin � Atomic force microscopy �
Fibers anisotropy � Cytoskeleton � Fibroblasts
Abbreviations
a-Tub a-Tubulin
AFM Atomic force microscopy
CLSM Confocal laser scanning microscopy
ABPs Actin-binding proteins
COF Cofilin
F-actin Filamentous actin
Gsk-3 Glycogen synthase kinase-3
PD Parkinson’s disease
Daniele Vergara, Marzia M. Ferraro and Mariafrancesca Cascione
have contributed equally to this work.
Electronic supplementary material The online version of thisarticle (doi:10.1007/s12013-014-0362-1) contains supplementarymaterial, which is available to authorized users.
D. Vergara � M. M. Ferraro � M. Maffia
Department of Biological and Environmental Sciences and
Technologies, University of Salento, Lecce, Italy
D. Vergara � M. Maffia
Laboratory of Clinical Proteomic, ‘‘Giovanni Paolo II’’
Hospital, ASL-Lecce, Italy
M. M. Ferraro � M. Cascione � L. L. del Mercato �
S. Leporatti � R. Rinaldi � A. Gaballo (&)
Institute of Nanoscience-NNL, CNR, Via Arnesano, 16,
Lecce 73100, Italy
e-mail: antonio.gaballo@nano.cnr.it
M. Cascione � R. Rinaldi
Department of Mathematics and Physics ‘‘Ennio De Giorgi’’,
University of Salento, Lecce, Italy
A. Ferretta � P. Tanzarella � T. Cocco
Department of Basic Medical Sciences, Neurosciences and
Organs of Senses, University of Bari ‘A. Moro’, Bari, Italy
C. Pacelli
Department of Pharmacology, Faculty of Medicine, Universite
de Montreal, 2900 Boulevard Edouard-Montpetit, Montreal,
QC H3T1J4, Canada
A. Santino
Institute of Science of Food Production, CNR, Lecce, Italy
123
Cell Biochem Biophys
DOI 10.1007/s12013-014-0362-1
CTR Healthy control
P1 PD patient
siRNAs Interfering RNAs
LIMK LIM kinase
MLC Myosin light chain
p-COF Phosphorylated cofilin
Akt Protein kinase B
PAK Rac-Cdc42 p21-activated kinase
ROI Region of interest
ROCK Rho associated kinase
Introduction
Parkinson’s disease (PD) is one of the most common
neurodegenerative diseases characterized by the selective
loss of dopaminergic neurons in the substantia nigra.
Understanding the molecular mechanisms underlying the
pathogenesis of PD is one of the most important challenges
in the neurodegenerative disease research. Although most
cases of PD are sporadic and caused by a number of
environmental factors, there are several gene products
involved in Mendelian forms of PD. Indeed, homozygous
or compound heterozygous mutations in PARK2 (parkin),
PARK6 (pink1), and PARK7 (DJ-1) can cause autosomal
recessive forms of early-onset parkinsonism [1].
Parkin is an E3 ubiquitin ligase and participates in a
variety of cellular activities primarily through the ubiqui-
tination followed by 26S proteasome degradation of dam-
aged or misfolded proteins [2]. Parkin also plays an
important role in the autophagy of damaged mitochondria,
as well as their biogenesis, besides a well-documented role
on cellular bioenergetic deficit due to mitochondria dys-
function associated with oxidative stress [3, 4]. Recent
studies have provided new insights about the molecular
interactions between parkin and other (cytoplasmic and
mitochondrial) proteins, [5, 6] thus allowing a deeper
understanding of parkin role in the pathogenesis of PD. In
particular, it was demonstrated that parkin mutations, as
well as dysfunctions in other PD-linked proteins such as
a-synuclein and leucine-rich repeat kinase 2 (LRRK2) [7,
8] can affect cytoskeletal organization [9] and impair the
vesicular transport [10].
The actin cytoskeletal architecture affects several cel-
lular processes essential for normal development [11], as
well as many pathological conditions including cancer and
neurodegenerative diseases [12, 13]. The reorganization of
the actin cytoskeleton is controlled by actin-binding pro-
teins (ABPs) that control the tread-milling, depolymeriza-
tion, nucleation, elongation, and capping of actin filaments
[14]. Among these, cofilin (COF) is one of the major ABPs
whose main function is to sever filamentous actin (F-actin)
into short segments and create free barbed ends for actin
elongation which are critical processes for motility,
migration, morphogenesis, division, and differentiation
[15–17]. COF is inactivated by phosphorylation at the Ser3
residue by LIM Kinase 1 (LIMK1) [18]. LIMK1 is a key
downstream effector of the Rho family of small GTPase
and is activated by phosphorylation of Thr 508 by the Rho-
associated kinase (ROCK) or by Rac-Cdc42 p21-activated
kinase (PAK) [19].
In this study, we demonstrate that parkin-mutant fibro-
blasts showed alterations in the expression of cytoskeletal
proteins with respect to healthy control cells and that these
modifications affect both the cell shape and cell elasticity.
These results imply a close link between parkin, cyto-
skeletal proteins, and cell stiffness.
Materials and methods
Primary Fibroblasts and Culture Conditions
Primary fibroblasts from PD patient (P1) and from P1’s
mother, used as a healthy control (CTR), were previously
described [3]. The diagnosis of PD was made according to
the UK Brain Bank criteria: the patient underwent neuro-
logical examination including the motor part of the Unified
Parkinson’s Disease Rating Scale (UPDRS III) and Hoen–
Yahr scale (H&Y).
Primary fibroblasts from P1 and CTR were obtained by
explants from skin punch biopsy, after informed consent.
Cells were grown in high-glucose Dulbecco’s modified
Eagle’s medium (DMEM) supplemented with 10 % (v/v)
fetal bovine serum (FBS), 1 % (v/v) L-glutamine, and 1 %
(v/v) penicillin/streptomycin, at 37 �C in a humidified
atmosphere of 5 % CO2.
Western Blot Analysis
Western blot analysis was performed as previously
described [12]. In detail, cells were washed in PBS and
lysed in RIPA buffer (Cell Signaling) and insoluble
material cleared by centrifugation at 10,000 rpm for 5 min.
Proteins were quantified by the Bradford protein assay
(BIORAD). Twenty-five micrograms of cell extract were
resolved on a 10 % polyacrylamide SDS-gel, and trans-
ferred to a nitrocellulose membrane (Hybond ECL). The
membranes were blocked for at least 1 h at room temper-
ature in Blotto A (Santa Cruz), and subsequently probed by
the appropriately diluted primary antibodies for 2 h at
room temperature (see Supplementary Table 1 for the
complete list of antibodies used in western blotting). After
three washes with a solution of 0.1 % (v/v) tween 20 in
Cell Biochem Biophys
123
TBS, the blots were incubated with secondary antibodies
HRP-conjugated for 2 h at room temperature (1:2000
dilution). Immunoblots were developed using the ECL
system. Images shown in the paper are representative of at
least three replicates with similar results.
siRNA-Mediated Parkin Knockdown
CTR fibroblasts at 60–70 % confluence were transiently
transfected for 48 h with 20 nM of small interfering RNAs
(siRNAs) specific for human parkin gene (Parkin siRNA)
and scrambled siRNAs (Control siRNA) according to the
manufacturers’ instructions (SMART-pool; Dharmacon
RNA Technologies, Lafayette, CO) as described in Ref.
[20]. The sequences of siRNA for Park2 are: 50-GUAAAG
AAGCGUACCAUGA-30, 50-GAACAUCACUUGCAUUA
CG-30, 50-GAUAGUGUUUGUCAGGUUC-30, and 50-UU
AAAGAGCUCCAUCACUU-30. Non targeting sequences
are: 50-UGGUUUACAUGUCGACUAA-30, 50-UGGUUU
ACAUGUUGUGUGA-30, 50-UGGUUUACAUGUUUUC
UGA-30, 50-UGGUUUACAUGUUUUCCUA-30.
Confocal Immunofluorescence Microscopy Analysis
For confocal microscopy analysis, cells were grown onto
glass coverslips at 3 9 104 cells/ml in 6-well plates over-
night. For F-actin staining, seeded cells were fixed and
permeabilized with ice cold acetone for 15 min. Fixed cells
were washed twice with ice cold PBS and incubated with
phalloidin–tetramethylrhodamine B isothiocyanate (TRITC)
according to the manufacturer’s protocol (P1951, Sigma).
After three washes in PBS, coverslips were mounted with
Fluoroshield with DAPI (F6057, Sigma).
a-Tubulin (a-Tub) staining was performed according to
the manufacturer’s protocol (Santa Cruz). Subsequently,
samples were incubated with an anti-mouse Alexa Fluor
488 (AF488)-conjugated secondary antibody (Cell Signal-
ing). For nuclear staining, fixed cells were incubated with
1 lg ml-1 of DAPI for 5 min at room temperature.
The images of fluorescently labeled proteins were cap-
tured using a confocal laser scanning microscope (CLSM)
(TCS SP5; Leica, Microsystem GmbH, Mannheim, Ger-
many) equipped with a laser diode emitting at 405 nm, an
argon-ion laser for excitation at 488 nm (50 %), and a
helium–neon laser for excitation at 514 nm (50 %). The
fluorescence signal of DAPI (blue channel) was detected
after filtering with a band pass filter within the range of
415–500 nm, AF488-labeled secondary antibody (green
channel) was detected with a band pass filter 495–519 nm,
and TRITC-phalloidin (red channel) was detected with a
565–660 nm band pass filter. Images were taken with a
HCX PL APO lambda blue 63.0 9 1.40 oil-immersion
objective under sequential mode acquisition (scan mode:
xyz; scan speed: 200 Hz). The pinhole aperture was set to 1
Airy.
Quantification of Anisotropy of Microtubules and
F-Actin Fiber Arrays in CTR and P1 Cells
FibrilTool, an ImageJ plug-in recently described by Bou-
daoud and co-authors [21], was used for quantifying the
anisotropy of microtubules and F-actin fiber arrays in CTR
and P1 cells. FibrilTool provides the average orientation
and anisotropy of fiber arrays in a given region of interest
(ROI) that is manually designed in the fluorescent image. A
detailed description of the procedure for quantifying the
fibril anisotropy was described elsewhere [21]. Briefly,
CLSM micrographs are opened in ImageJ (1.48v). The
fluorescent channels (red or green) are first selected in the
FibrilTool dialog box and the linelength (the line segment
drawn by FibrilTool within the ROI visualizing the
anisotropy of the array) is set to 1. Then, the polygon tool is
used to manually design ROIs. At least two ROIs are
determined for each cell, avoiding regions with saturating
pixels. At the end of the analysis, the anisotropy of the fiber
arrays is quantified. With regard to the anisotropy score,
FibrilTool uses the following convention: 0 for no order
(purely isotropic arrays) and 1 for perfectly ordered, i.e.,
parallel fibrils (purely anisotropic arrays) [21]. The abso-
lute value of anisotropy is meaningful only when it is
compared with images of the same type. In our analysis,
the anisotropy scores were thus measured and compared
within the same channel (green or red).
Atomic Force Microscopy Analysis
AFM experiments were performed using a CAT (Confocal-
AFM-TIRF) microscope that combines an atomic force
microscope (AFM) (Bioscope Catalyst, Bruker Inc. USA),
a laser scanning confocal microscope (LSM 700, Zeiss,
Germany), and a total internal reflection fluorescence
microscope (Laser TIRF 3, Zeiss, Germany) in one single
compact instrument. All devices are mounted on an
inverted optical microscope (Zeiss Observer Z1, Zeiss,
Germany).
Force measurements were carried out on living fibro-
blasts seeded on Petri dishes, and maintained in Leibovitz
medium (L-15) at 37 �C in contact mode as described in
Ref. [22, 23]. V-shaped silicon nitride cantilevers (MSNL-
10 Veeco, USA) with a low nominal constant ranging from
0.01 to 0.03 nm-1 were used. Cantilever spring constant
was determined before each experiment using the thermal
noise method [24]. The mechanical properties of samples
were analyzed in force mapping mode (force volume). The
local Young’s modulus was evaluated by fitting the
Cell Biochem Biophys
123
experimental loading data with a modified Sneddon model.
Different parameters were used in order to acquire force
mapping curves: scan size 50 lm, ramp size 5 lm, ramp
rate 6 Hz, and trigger threshold 50 nm. From 20 different
cells, 25 force-distance curves were exported. Curves were
analyzed using a LabView-based home-made software.
Statistical Analysis
Data were analyzed using the statistical software GraphPad
(version 4.0). The two-tailed t test was used to test statis-
tical significance. Differences were considered statistically
significant for P-values\0.05. Densitometric analysis was
performed using the software ImageJ.
Results and Discussion
In this study, we used cultured skin fibroblasts from a
patient affected by an early-onset PD, labeled as P1, with
the PARK2 heterozygous mutations del exons 2–3/del exon
3 and from P1’s mother as parental healthy control, labeled
as CTR. The western blot analysis of parkin protein
expression showed the complete absence of the 50 kDa
full-length protein in the P1 patient cells, while parkin was
shown to be present in the parental healthy control [3]. In
our previous study, we performed a biochemical charac-
terisation of these two fibroblasts primary cultures and
demonstrated that the absence of parkin was associated
with ultrastructural abnormalities of mitochondria resulting
in an impaired energy metabolism, increase of ROS pro-
duction, and a general perturbation of mitochondrial
homeostasis [3]. Here, we provide further evidence about
the role of parkin in the regulation of specific processes of
cellular homeostasis. As shown in Fig. S1, (Supplementary
Material), the protein–protein interaction network of parkin
places this protein at the center of a network that comprises
proteins involved in several biological processes, molecu-
lar functions, and signaling pathways. Providing experi-
mental evidence for the role of parkin as mediator of these
processes may have important consequences for the diag-
nosis and treatment of PD.
By western blotting analysis, we studied the expression
of different proteins in the CTR and P1 fibroblasts (Fig. 1a,
c). These include a group of proteins involved in the reg-
ulation of cytoskeleton such as COF, Ezrin, LIMK, ROCK
and Myosin, other protein kinases, adhesion proteins, and
cell-cycle regulators. Fibroblasts did not express E-cad-
herin, a typical epithelial marker, but express N-cadherin
that showed a significant down-regulation (P\ 0.01) in P1
cells. On the contrary, cyclin-D1, a regulator of the tran-
sition G1/S, is up-regulated in P1. This result is consistent
with the modification of the Protein kinase B (Akt)/Gly-
cogen synthase kinase-3 (Gsk-3) pathway observed in P1
cells, which acts as an important regulator in the determi-
nation of cyclin D1 expression at both mRNA transcription
and protein level [25]. In fact, the phosphorylation status of
Akt (Ser473), of Gsk (Ser9), and Gsk (Tyr279/Tyr216) is
significantly reduced in P1 cells. Moreover, Gsk-3 has an
important role in regulating b-catenin levels that result
down-regulated in P1 cells (Fig. 1a, c). Rawal and col-
laborators [26] demonstrated in vivo that parkin is a reg-
ulator of Wnt/b-catenin signaling by reducing the pool of
active b-catenin upon Wnt3a stimulation. They showed
that the parkin’s effect on b-catenin is tissue type specific
and restricted to the ventral midbrain of parkin-null mice.
Here, our analysis extend this observation to human
fibroblasts and suggest a possible regulation by Gsk that
requires a future investigation.
Considering the group of cytoskeletal proteins, we find a
marked decrease of Ezrin expression in P1 cells, together
with a down-regulation of ROCK. Importantly, several
works already described a relationship between PD and
actin dynamics [27–30]. In the nervous system, actin
dynamics were shown to have a central role both in neurite
outgrowth and in the synapse formation and maintenance
[31, 32]. Therefore, a further understanding of the func-
tional link between parkin and cytoskeleton may have a
significant clinical impact. It was demonstrated that a
Parkinson’s disease-related G2019S substitution in the
kinase domain of LRRK2 enhanced the phosphorylation of
Ezrin, Radixin, and Moesin (ERM) proteins [33]. Our
findings underscore the importance of parkin in the regu-
lation of Ezrin expression.
ROCK is a kinase with a role in the regulation of the
acto-myosin contractile force. Major downstream sub-
strates include myosin light chain (MLC) and LIMK (for a
review see Ref. [19]). Activation of LIMK enhances its
ability to phosphorylate COF, a key regulator of actin fil-
ament dynamics and reorganization by stimulating depo-
lymerization and severance of actin filament [17]. This
COF effect is inhibited by phosphorylation on Ser3 that
resulted in abolishing its actin-binding activity [17]. Parkin
has been shown in the human neuroblastoma-derived
BE(2)-M17 cell line to interact to LIMK1 reducing its COF
phosphorylation activity by enhancing LIMK1-ubiquitina-
tion [30].
Here, we showed a significant upregulation of myosin
(Thr18/Ser19), LIMK 1/2 (Tyr507/Thr508), and Cof (Ser3)
in P1 compared to CTR cells. To gain deeper insight into
the involvement of parkin in the LIMK/COF pathway,
CTR cells were transfected with a parkin-targeted siRNA
to knockdown parkin expression. Parkin down-regulation
by siRNA reduced the phosphorylation status of LIMK1
and COF relative to the scramble CTR siRNA cells
Cell Biochem Biophys
123
(Fig. 1b). This result further supports a role for parkin in
the regulation of this pathway and possibly in the regula-
tion of actin filaments dynamism. For this reason, we
studied the organization of actin filaments and microtu-
bules by CLSM analysis. Figure 2a shows CLSM micro-
graphs of CTR and P1 cells stained with TRITC-phalloidin
for actin filaments (red), a specific monoclonal antibody
raised against a-Tub for microtubules (green) and DAPI for
nuclei (blue). We observed remarkable differences in
morphology (Fig. 2a) between CTR and P1 fibroblasts.
CTR cells appeared to have a typical fibroblast elongated
shape with actin filaments filling the central bodies and
radiating into peripheral lamellas. On the other hand, P1
fibroblasts displayed a completely different shapes,
appearing much wider and longer and showing a longer
doubling time with respect to the parental control
[3]. Furthermore, P1 actin filaments, as well as microtu-
bules, were neither uniformly aligned, nor evenly distrib-
uted, displaying an higher amount of stress fibers that
appeared to be more randomly oriented, in contrast to what
can be seen in CTR whose microfilaments and microtu-
bules appeared aligned along the major axis of the cell. We
then quantified the degree of alignment of fibrillar struc-
tures in a set of CLSM micrographs using the ImageJ plug-
in FibrilTool as described by Boudaoud and co-authors
[21]. In particular, we measured the anisotropy, a number
Fig. 1 a Western blot analysis of CTR and P1 samples. Whole
proteins obtained from CTR and P1 fibroblasts were probed by the
appropriately diluted primary antibodies for 2 h at room temperature.
a-Tub was used as loading control. b Western blot analysis of LIMK
1/2, LIMK 1/2 (Tyr507/Thr508), COF, and COF (Ser3) in CTR
siRNA and parkin siRNA samples. c Fold change data are presented
as mean ± standard deviation of P1/CTR ratio. Asterisks indicate
significance Student’s t test, *P\ 0.05; **P\ 0.01; ***P\ 0.001.
Images shown are representative of at least three replicates with
similar results
Cell Biochem Biophys
123
between 0 and 1 that quantifies how parallel the fibers are
in a selected ROI. In this way we could compare the
anisotropy of fibrillar structures in CTR and P1 cells
(Figures S2–4, Supplementary Material). With regard to
the anisotropy score, 0 indicates no order (purely isotropic
arrays) and 1 indicates perfectly ordered, i.e., parallel
fibrils (purely anisotropic arrays) [21]. The average
anisotropy scores of F-actin and microtubules of CTR and
Fig. 2 a CLSM micrographs of organization of F-actin (stained with
phalloidin, in red) and microtubules (stained with a-Tub, in green)
filaments in CTR and P1 cells (nuclei stained with DAPI, in blue).
The individual red and green channels are shown followed by merged
images of red, green, and blue channels (individual blue channel,
DAPI, not shown). Scale bars 50 lm. b Average of fibril arrays
anisotropy of F-actin and microtubules filaments of CTR and P1 cells.
Per each cell at least two region of interest (ROI) was determined
manually, avoiding regions with saturating pixels (see Supplementary
Figs. 3, 4 for details). The fluorescent channels (red or green) were
selected and line length was set to 1. Anisotropy scores between 0 (no
ordered fibrils, purely isotropic arrays) and 1 (perfectly ordered fibrils,
purely anisotropic arrays) were finally measured from FibrilTool.
Columns represent mean ± standard error of mean (n� cell ana-
lyzed = 10). c Average of cell area of CTR and P1 cells. Columns
represent mean ± standard error of mean (n� cell analyzed = 25).
The outlines of DIC images of single CTR and P1 cells were used for
morphometric analysis. An image software was used for entering and
storing the cell outlines and for calculating the cellular area.
d Average axes ratio of CTR and P1 cells obtained by dividing the
major axis of the cell by the minor axis of the cell. In (b) and
(c) asterisks indicate significance in two-tailed Student’s t test,
*P\ 0.05, **P\ 0.005, ***P\ 0.0005. In (d) asterisks indicate
significance in two-tailed Student’s t test, ***P\ 0.001, **P\ 0.01,
*P\ 0.05 (Color figure online)
Cell Biochem Biophys
123
P1 cells are reported in Fig. 2b. P1 cells show for both
F-actin and microtubules filaments anisotropy scores sig-
nificantly lower than CTR cells. This analysis confirmed
that P1 cells possess a lower degree of alignment of F-actin
and microtubules fibers compared to CTR cells.
As a quantitative measure of cell shape, the cell area and
the ratio between major and minor axes of CTR and P1
cells were determined (Fig. 2c, d) [9, 34]. The cell area
data showed values about 2.5 times higher in P1 with
respect to CTR cells (Fig. 2c). According to confocal
microscopy observations, P1 cells display axes ratio lower
than CTR cells, thus resembling a predominant rounded
shape of P1 cells compared to the relatively elongated
shape of CTR cells (Fig. 2d). These results denote that P1
fibroblasts undergo marked alterations in cell morphology
with an evident disorganization of the actin microfilaments
and microtubules. Moreover, these data are consistent with
the severe ultrastructural alterations revealed by electron
microscopy images presented in our previous work [3]
showing an irregular cellular shape with pseudopodia
membranes, cytoplasmic protrusions, and heterochromatic
and indented nuclei.
As alterations in cross-linking and structural protein net-
works are the major determinants of the elastic properties of
cells, we hypothesize that the observed different cytoskeletal
organizations may impact the nanomechanical properties of
CTR and P1 cells. Figure 3 shows representative AFM
height (a, top) and deflection (a, bottom) images of CTR and
P1 fibroblasts cultured in Petri dishes. Images were acquired
in contact mode at very low force, maintaining cells in L-15
culturemedium at 37 �Cduring the analysis. In CTR sample,
stress fibers (bundles of actin filaments) are clearly aligned
on a preferential direction (longitudinal) as evidenced from
deflection images. These data confirmed the structural rear-
rangements observed in fibroblasts by confocal microscopy
analysis (Fig. 2a). The different cellular architectures were
also accompanied by an increased stiffness of P1 compared
with CTR cells (Fig. 3b). Force volume images, from which
elasticity data have been derived, were obtained from single
cells indented approximately over the perinuclear region
with an indentation depth of about 50 nm. This prevents
possible effects caused by interaction with the substrate.
Histograms in Fig. 3b show that P1 cells (Young’s modulus
P1 = 4.76 ± 0.26 kPa) are significantly stiffer with respect
Fig. 3 a Representative AFM height (top) and deflection (bottom)
images of CTR and P1 fibroblasts cultured in Petri dishes. The height
of the cells is depicted by the colored bar. b Elasticity profiles of
living CTR and P1 fibroblasts. c Representative three dimensional
reconstruction of living CTR fibroblasts (scan size: 50 9 50 lm, Z-
scale 1.3 lm). Single cells were indented approximately over the
perinuclear region (indentation depth = 50 nm). The Young’s moduli
of CTR and P1 fibroblasts result from the analysis of 20 load-
ing\unloading curves per cell (n� cell analyzed = 25). Columns
represent mean ± standard error of mean
Cell Biochem Biophys
123
to CTR sample (Young’s modulus CTR = 3.72 ±
0.08 kPa). This finding correlates well with the disorgani-
zation of the actin cytoskeleton fibers observed by AFM and
confocal microscopy analysis.
We next asked whether the activation of LIMK/COF
pathway observed in P1 fibroblasts could be associated
with the nanomechanical properties of these cells. To
perform this, cells were treated with the cell permeable
compound Y-27632, a specific inhibitor of the kinase
ROCK [19, 35]. Fig. 4a shows CLSM micrographs of CTR
and P1 cells stained with TRITC–phalloidin, treated or not
with Y-27632 at the concentration of 10 lM for 24 h. We
observed that Y-27632 treatment induced changes in the
organization of actin cytoskeleton with significant effects
on patient’s cells. In particular, these cells appeared elon-
gated with narrow protrusions resembling those of CTR(-)
cells. This induced modifications of the ratio between
major and minor axes (not shown), as well as of the cell
area (Fig. 4b) whose values significantly decreased after
treatment with Y-27632. Therefore, the inhibition of
ROCK also induced modification in the cytoskeletal
architecture with the disruption of actin stress fibers.
To demonstrate that ROCK inhibition induces cellular
modifications through regulating LIMK/COF and Myosin
pathways, western blot analysis was performed on CTR
and P1 cells treated with Y-27632. As shown in Fig. 4c,
Y-27632 treatment reduced Myosin, LIMK, and COF
phosphorylation in CTR and P1 fibroblasts, suggesting that
both proteins are dependent on ROCK activity (under
ROCK control). Together, these data further support the
role of these pathways in regulating the mechanical prop-
erties of P1 cells, role also confirmed by the AFM analyses.
In fact, ROCK inhibition by Y-27632 treatment led to actin
cytoskeleton reorganization and the disappearance of stress
fibers (Fig. S5a and b, Supplementary Material). In AFM
deflection images, a retraction of the cell is also observed
(Fig. S5a and b, Supplementary Material) which can con-
tribute to a marked increase of cell stiffness (Fig. S5c,
Supplementary Material).
Conclusion
This study, using parkin-null fibroblasts derived from a
Parkinson’s patient, was aimed to investigate the role of
parkin in regulating actin filament organization and cellular
stiffness. Most notably, the integration of AFM with con-
focal microscopy and other biological techniques has
revealed that parkin-null fibroblasts showed significant
differences in elasticity, cell shape, and stress fibers orga-
nization. In particular, the absence of parkin resulted in an
increase of stiffness, in accordance with the different
expressions of actin regulators in control and parkin-null
fibroblasts. Moreover, treatment with the ROCK inhibitor
Y-27632 confirmed the contribution of LIMK/COF path-
way in the regulation of actin organization and cellular
Fig. 4 a CLSM micrographs of
CTR and P1 cells before (-)
and after (?) treatment with
10 lM Y-27632, a cell
permeable inhibitor of ROCK.
F-actin filaments stained with
phalloidin (in red). Scale bars
50 lm. Enlarged details of each
photomicrograph are also
shown. b Average of cell area of
CTR and P1 cells before (-)
and after (?) treatment with
Y-27632. Columns represent
mean ± standard error of mean
(n� cell analyzed = 25).
Asterisks indicate significance
in two-tailed Student’s t test
(**P\ 0.005). c Western blot
analysis of CTR and P1 samples
probed with specific antibodies
as indicated. a-Tub was used as
loading control (Color figure
online)
Cell Biochem Biophys
123
stiffness. Overall, these results support the hypothesis that
parkin is involved in the regulation of actin cytoskeleton by
targeting LIMK/COF pathway and represent the first direct
evidence of the role of parkin in regulating biomechanical
properties of cell. Although several studies established
quantitative biomechanical variations in human cells
exposed to several biological conditions, only few of them
correlated these differences to specific molecular altera-
tions. The feasibility of our integrated approach would
make this an attractive strategy to correlate the cellular
nanomechanical profile to the underlying molecular and
cellular mechanisms.
Acknowledgments This work was supported by the PON project
254/Ric. ‘‘Implementation of human and environment health research
center’’, Cod. PONa3_00334, REA research Grant no PITN-GA-2012-
316549 (IT LIVER) from the People Programme (Marie Curie Actions)
of the European Union’s Seventh Framework Programme (FP7/2007-
2013), PRIN 2010FPTBSH ‘‘NANO Molecular technologies for Drug
delivery - NANOMED’’, Cod. PON02_00563_3448479-F ‘‘RINO-
VATIS’’, and Cod. PONa3_00134 ‘‘Omics and nanotechnology for
early diagnosis of human diseases - ONEV’’. LLdM thanks Prof.
Arezki Boudaoud for useful discussions about FibrilTool.
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