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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