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Self-assembly of polydimethylsiloxane structures from 2D to 3D for bio-hybrid actuation
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2015 Bioinspir. Biomim. 10 056001
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Bioinspir. Biomim. 10 (2015) 056001
doi:10.1088/1748-3190/10/5/056001
PAPER
RECEIVED
24 April 2015
REVISED
8 July 2015
ACCEPTED FOR PUBLICATION
21 July 2015
PUBLISHED
20 August 2015
Self-assembly of polydimethylsiloxane structures from 2D to 3D for
bio-hybrid actuation
L Vannozzi1, L Ricotti1, M Cianchetti1, C Bearzi2,3, C Gargioli4, R Rizzi2,3, P Dario1 and A Menciassi1
1
2
3
4
The BioRobotics Institute, Scuola Superiore Sant’Anna, Pontedera (PI), Italy
Cell Biology and Neurobiology Institute, National Research Council of Italy, Rome, Italy
IRCCS MultiMedica, Milan, Italy
Department of Biology, University of Rome Tor Vergata, Rome, Italy
E-mail: [email protected]
Keywords: bio-hybrid actuation, engineered bio/non-bio interfaces, 3D self-assembled structures, living machines
Supplementary material for this article is available online
Abstract
This work aims to demonstrate the feasibility of a novel approach for the development of 3D selfassembled polydimethylsiloxane structures, to be used as engineered flexible matrices for bio-hybrid
actuation. We described the fabrication of engineered bilayers, organized in a 3D architecture by
means of a stress-induced rolling membrane technique. Such structures were provided with ad hoc
surface topographies, for both cell alignment and cell survival after membrane rolling. We reported
the results of advanced finite element model simulations, predicting the system behavior in terms of
overall contraction, induced by the contractile activity of muscle cells seeded on the membrane. Then,
we tested in vitro the structure with primary cardiomyocytes to evaluate the real bio-actuator
contraction, thus validating the simulation results. At a later stage, we provided the samples with a
stable fibronectin coating, by covalently binding the protein on the polymer surface, thus enabling
long-term cultures with C2C12 skeletal muscle cells, a more controllable cell type. These tests revealed
cell viability and alignment on the rolled structures, but also the ability of cells to differentiate and to
form multinucleated and oriented myotubes on the polymer surface, also supported by a fibroblast
feeder layer. Our results highlighted the possibility of developing 3D rolled PDMS structures,
characterized by different mechanical properties, as novel bio-hybrid actuators.
1. Introduction
One of the most important functions that characterize
a moving system is its actuation strategy. Many
technological efforts have allowed to possess nowadays
several artificial actuation solutions [1]. However,
different factors currently limit actuator performances
and their application in specific fields. Biorobotic
systems, for example, show peculiar requirements in
terms of actuation strategies. Miniature medical
robots, flexible surgical instruments, advanced and
biomimetic prosthetic limbs and other devices would
strongly benefit from advancements in the actuation
domain. Among the limitations of current actuators,
lack of stiffness control, poor scalability at small scales,
high power consumptions and no self-healing properties are the most significant ones [2]. Recently, the use
of contractile muscle cells has represented a promising
© 2015 IOP Publishing Ltd
solution to bypass these issues. In fact, the possibility
to directly use muscle cells could permit to achieve
actuation components almost matching natural muscle properties [3]. Other approaches, such as shape
memory alloys [4], electro-active polymers (EAP) [5]
or carbon-nanotube-based systems [6] have been
explored in order to emulate the functionality of an
in vivo muscle, but their properties are currently far to
well mimic muscle performances [7–9].
Muscles are linear actuators capable of large displacements and a stiffness control in a wide range [10].
A peculiar characteristic that distinguishes them
respect to other actuation means is intrinsic their scalability. Muscles allow an efficient organization of
actuation units in a range from micrometers to meters.
Muscles are long-lasting thanks to their high life cycle
and possess integrated sensors (e.g. the muscle spindles) which assure accuracy and repeatability. They
Bioinspir. Biomim. 10 (2015) 056001
L Vannozzi et al
work by consuming renewable chemical energy produced by glucose and oxygen, highly available in nature, while the wastes are biodegradable products, such
as carbon dioxide. Nowadays, the possibility to exploit
all these advantages is no longer far from reality [11–
13]. Muscle cell-based actuators could power adaptable and self-healing devices, characterized by ‘lifelike’ movements and able to overcome most of the
limitations affecting current actuation strategies. This
new actuation means could be beneficial for many
technological applications [3, 14] and it may enable in
a near future the development of more flexible and
dexterous robots, able to perform challenging
tasks [15, 16].
Recently, a significant number of studies on biohybrid actuation showed up, thus increasing the promises of achieving a usable bio-hybrid system. The
first relevant work in this field was represented by a
swimming robot, actuated by means of two explanted
frog semitendinosus muscles, able to swim for many
hours while maintaining a good efficiency [17]. Further studies followed, by exploiting the contractile
properties of different types of muscle cells: spontaneous contraction of primary cardiomyocytes
[18, 19], dorsal vessel insect tissues [20, 21] and controllable mammalian or insect skeletal muscle cells
[22–24].
Engineering challenges involve the integration of a
control system allowing cell contractile action to be
triggered and regulated. In this context, skeletal muscle cells represent an optimal solution, because they
are highly controllable, in particular by means of optical [25] or electrical stimuli [26]. A key feature of a
bio-hybrid actuation system should be the ability to
mimic the 3D fascicle structures of natural muscle tissues [27–29]. However, the majority of bio-hybrid literature examples report 2D systems, in the form of
membranes and thin films [19, 27, 30–34].
Recently, a bio-robot mimicking the structure and
the kinematics of a ‘medusoid’ [35] and a bio-hybrid
swimmer [36] were developed. Moreover, 3D printing
techniques were exploited to fabricate walking biological machines based on cardiomyocytes and C2C12
skeletal muscle cells for uni-directional movement [37, 38].
The main challenge still remains the development
of controllable 3D skeletal muscle-based in vitro constructs. This can be achieved by following two main
strategies: (i) by providing 3D scaffolds with a vasculature, in order to keep cell viability in the inner parts
of the construct; (ii) by organizing 2D materials in 3D
structures which organize and keep cells viable thanks
to their architecture, without the need of a bio-artificial vasculature. Concerning the first strategy, several
research efforts are currently ongoing to create
pseudo-vascularized biomaterials for skeletal muscle
tissue engineering [39]. However, we are quite far
from obtaining optimal results as regards the integration of a vasculature system allowing an efficient
2
transport of oxygen, glucose, nutrients and waste products [40]. Although exciting results have been
recently achieved in the field of 3D muscle tissue engineering [41], the lack of a real capillary blood flow deeply limits the long-term stability of muscle constructs.
Concerning the second strategy, some techniques permit to relatively easily move from simple 2D patterns
to complex 3D ones, through self-assembly [42]. This
kind of approach could allow to bypass the vascularization issues that affect 3D constructs. To the best of
authors’ knowledge, no bio-actuators based on 3D
self-assembled rolled structures have been proposed,
so far.
In this paper, we describe the fabrication of 3D
tubular self-assembled polymeric structures used as
matrices for the development of bio-hybrid actuators.
We report the results obtained in terms of fabrication
procedure, substrate characterization, surface functionalization and cell culture tests. Furthermore, we show
the insights achieved by performing advanced finite
element model (FEM) simulations, which are validated by experimental results and highlight the potential of these systems for bio-hybrid actuation.
2. Materials and methods
In this section we describe the materials and methodologies to fabricate and characterize the polymeric
structures, to chemically functionalize their surfaces,
to assess cell behavior by means of in vitro assays, to
predict the system performance by means of advanced
simulations and to validate the simulation results.
2.1. Membrane fabrication and assembly
The engineered structures were fabricated in polydimethylsiloxane (PDMS, Sylgard 184, Dow Corning), a
biocompatible, transparent and elastic polymer. To
obtain a relatively wide range of elastic moduli for
better matching the tissue elastic modulus, PDMS was
prepared in different monomer/curing agent ratios
(5:1, 10:1, 15:1 and 20:1 w/w). Thin films were
fabricated by spin-assisted deposition (SPIN150, SPS
Europe). All films were spinned for 60 s; different films
thicknesses were obtained by varying spin velocity
(from 500 rpm up to 5000 rpm). The strategy followed
for the assembly of engineered PDMS matrices was
based on the stress-induced rolling membrane (SIRM)
technique [42] (figures 1(a)–(d)), which permitted to
obtain tubular rolled structures from a 2D pattern,
characterized by two merged layers provided with
different surface topographies. This approach, based
on the release of internal stresses induced by the
stretching of the top layer, allowed the rolling of the
whole structure in absence of constraints at one end,
once the two layers were bonded together. Thus, while
cell orientation was controlled on the 2D surface, the
patterns were assembled in a 3D structure.
Bioinspir. Biomim. 10 (2015) 056001
L Vannozzi et al
Figure 1. Description of the SIRM technique. The top layer (a) was stretched (b) and merged with a non-stretched bottom layer (c).
The bilayer, in absence of constraints at one extremity, started to roll (d), due to the release of the previously generated internal
stresses. (e) CAD design of the rolled PDMS structure with pillars (red box, bottom layer) and grooves (green box, top layer)
evidenced. The black arrow indicates the micro-grooves direction.
We used polyvinyl alcohol (PVA, 98% hydrolized,
Mw 13 000–23 000, Sigma-Aldrich) as a sacrificial
layer. PVA is a water-soluble polymer, used to facilitate the detachment of thin membranes from Si substrates [43–45]. A PVA aqueous solution (1% wt.) was
spin-coated (4000 rpm for 20 s) on Si molds before
pouring the PDMS solution. Then, PDMS was spinned on the PVA layer and baked at 150 °C for 15 min:
this permitted to obtain a rather robust and easy-tomanipulate structure (top layer), which could be stretched 40% of its initial length (figure 1(b)). The other
PDMS membrane (bottom layer) was baked at 80 °C
for 195 s, by keeping it in a non-fully polymerized,
semi-adhesive status. Then, the top layer (stretched
and clamped) was placed on top of the bottom one,
and the whole bilayer was finally subjected to an additional thermal treatment (80 °C for 30 min) to complete the polymerization, thus stabilizing its structure.
The semicured (bottom) membrane assured a better
adhesion between the two PDMS layers during this
step. The overall thickness of the PDMS bilayer was
the sum of the thicknesses of each layer (figure 1(c)).
2.2. Fabrication of substrate microtopographies
Substrate topography represents a key feature of the
structure. In fact, matrix topography must be properly
designed to assure an anisotropic cell alignment along
one preferential direction, thus maximizing cell fusion
and force generation in the longitudinal axis direction
(black arrow in figure 1(e)) [46]. Moreover, during the
3
rolling, the different layers should be spaced in order
to avoid excessive stresses on cells; for this reason, the
bottom layer was provided with spacing pillars that
aimed at separating each concentric layer, thus maximizing cell viability (figure 1(e)).
As a first step, two photolithographic masks were
designed and subsequently printed on glass, to obtain
the desired topographies on Si wafers (400 μm thick, p
type, boron doped, 〈100〉, Silicon Materials, Kaufering, Germany). In a class 1000 clean room, a negative
and a positive photoresist were used to fabricate the
molds for PDMS bottom and top layer, respectively.
For the top layer topography (parallel microgrooves),
the molds were obtained by spinning a Microposit primer (3500 rpm for 30 s) before the positive resist
(Shipley S1813, 4500 rpm for 35 s) on Si wafers. After
pre-baking (120 °C for 1 min), the wafers were
exposed for 7 s to UV in hard-contact modality, by
means of a mask aligner (MA/BA6 Gen3, SUSS MicroTec). After the post-baking (120 °C for 1 min) the
masks were developed for 30 s in a developer solution
(MF-319 Developer, Microposit®).
For the bottom layer topography (rectangular pillars), a negative resist (SU-8 25, MicroChem) was
spinned on Si wafers by following two cycles: a spread
cycle to allow the resist to cover the entire surface
(ramp to 500 rpm at 100 rpm s−1 acceleration, then 5 s
cycle) and a spin cycle, to obtain the desired thickness
(ramp to 2000 rpm at a 300 rpm s−1 acceleration, then
30 s cycle). Then, the wafers were pre-baked (65 °C for
Bioinspir. Biomim. 10 (2015) 056001
L Vannozzi et al
3 min and 95 °C for 7 min) and exposed for 6 s in softcontact modality. A post exposure bake was performed to cross-link the exposed portions of the film
(65 °C for 1 min and 95 °C for 3 min). Finally, the
development was carried out in a SU-8 developer solution for 4 min.
PDMS top and bottom layers were obtained by
spinning the polymeric solution, as previously described, on these two Si mold types, previously provided
with a PVA sacrificial layer.
2.3. Characterization of membranes and rolled
structures
PDMS layer surfaces were imaged by means of atomic
force microscopy (AFM, Veeco) and scanning electron
microscopy (SEM, EVO MA15, Zeiss Instrument).
Before imaging, the samples were metallized by means
of Au sputtering (sputtering current: 25 mA, sputtering time: 60 s). AFM scans were carried out in tapping
mode by setting a 0.2 Hz scan frequency, 512 samples
per line and a 100 × 100 μm2 scan area. SEM scans
were performed by setting a beam voltage of 10 kV and
a current of 200 pA.
The evaluation of membrane mechanical properties was carried out by means of traction tests, using a
mechanical testing system (INSTRON 4464, equipped
with a ± 10 N load cell). The elastic moduli were
deduced from each stress/strain curve, according to a
standard procedure [47]. For each specimen, a constant traction speed of 5 mm min−1 was set until sample failure. Data were recorded with a frequency of
100 Hz. Ten independent specimens were tested, for
each sample type.
PDMS membrane thickness was assessed by
means of a profilometer (KLA Tencor). For each spin
velocity and monomer/curing agent ratio, three independent samples were tested and their thickness values
recorded.
2.4. PDMS surface functionalization
PDMS is characterized by spontaneous hydrophobic
recovery [48]. This represents a critical issue, as cells
spontaneously detach from PDMS after few days, if no
surface treatment is provided. This phenomenon can
compromise the long-term adhesion of muscle cells to
the substrate. Different methods are reported in the
literature to make PDMS surfaces hydrophilic [49].
Oxygen plasma allows to graft hydrophilic functional
groups onto PDMS. However, it is difficult to maintain
this condition for several days.
Instead, a natural cross-linker for proteins, such as
genipin [50], can prevent PDMS hydrophobic recovery, thus improving the long-term adhesion of muscle
cells on PDMS layers, as previously demonstrated
[51]. Such chemical functionalization protocol was
used to provide the PDMS matrix with a stable protein
coating on the top layer and to guarantee a long-lasting
bio-actuation system. In brief, PDMS samples were
4
incubated in a solution of 60% ethanol, 20% (3-aminopropyl) triethoxysilane and 1.2% NH3 (pH = 9) for
3 h at 60 °C. Then, they were incubated with a
5 mg ml−1 genipin (Sigma-Aldrich) solution in pure
water (Milli-Q, Millipore) for 60 min at 37 °C. We
incubated the samples immersed in a 25 μg ml−1 fibronectin (Sigma-Aldrich) solution in pure water at 37 °C
for 1 h and subsequently we kept them at 4 °C
overnight.
By using fluorescent fibronectin, it was possible to
monitor the presence of this protein on the functionalized substrates, overtime. Protein labeling was
obtained by using the Oregon Green 488 Protein
Labeling Kit (Invitrogen-Molecular Probes©). The
structures, during this experiment, were maintained
in phosphate buffered saline (PBS) solution, which
was renewed every day. Fluorescence images were
acquired at different days (day 2, 3, 6, 9 and 13) by
means of an inverted fluorescence microscope
(Eclipse Ti, FITC-TRITC filters, Nikon), equipped
with a CCD camera (DS-5MC USB2, Nikon) and with
NIS Elements imaging software. Images were then elaborated by using ImageJ, a free software available at
http://rsbweb.nih.gov/ij/.
2.5. Cell cultures and in vitro assays
Cardiomyocytes (CMs) were used to evaluate the
contraction ability of the rolled PDMS structure. CMs
were isolated from hearts of 1-day-old CD-1 neonatal
mice (Charles River Laboratories). After excision,
whole hearts were transferred into ice-cold Hanks
Balanced Salt Solution (HBBS) and the blood gently
squeezed out. The hearts were then minced into four
pieces in 0.1 mg ml−1 trypsin (Sigma-Aldrich) supplemented with 20 mg ml−1 of deoxyribonuclease I
(DNAse I, Sigma-Aldrich), and pre-digested overnight
at 4 °C. The following day, trypsin was removed and
the myocardial pieces incubated in 0.5 mg ml−1 collagenase (Worthington Biochemical) solution containing DNAse I with intermittent pipetting along
with shaking at 37 °C in a water bath for 3 min The
supernatant, containing free cells, was then collected
and kept on ice. The digestion step was repeated three
times. Cell suspensions from each digestion were
pooled, filtered through a 70 μm strainer and centrifuged at 800 rpm for 5 min at 4 °C. Cell pellet was then
resuspended in a maintenance medium composed by
high glucose Dulbecco’s modified eagle medium
(DMEM) supplemented with 10% horse serum (Invitrogen), 5% fetal bovine serum (FBS), 10 mM HEPES
(Invitrogen), 100 IU/ml penicillin, 100 μg ml−1 streptomycin and 2 mM L-glutamine. The cells were plated
and incubated at 37 °C for 3 h to allow the differential
attachment of non-myocytes, constituted mostly of
cardiac fibroblasts [52, 53]. The CMs-enriched suspension was seeded on gelatinized PDMS samples with
a density of 150 000 cells/cm2 resuspended in 100 μl,
in order to immediately reach confluence.
Bioinspir. Biomim. 10 (2015) 056001
L Vannozzi et al
The evaluation of both viability and orientation of
cells on the substrates was carried out 24 h after cell
seeding, by means of a Live/Dead® viability/cytotoxicity assay (Invitrogen). Fluorescence images were
acquired with the inverted fluorescence microscope.
Further in vitro tests were performed by using
C2C12 murine skeletal myoblasts (ATCC®, CRL1772) as muscle cell model and nHDFs (Normal
Human Dermal Fibroblast, Lonza, CC-2511). These
tests were necessary to assess the ability of the functionalized PDMS substrates to induce cell alignment, to
maintain cell viability and to permit the formation of
myotubes. The fibroblasts were used to create a feeder
layer that could facilitate the development of contractile myotubes [54], providing a suitable mechanical interface over the PDMS surface. Both cell types
were cultured in DMEM (EuroClone), with the addition of 10% FBS (EuroClone), 100 IU/ml penicillin,
100 μg ml−1 streptomycin and 2 mM L-glutamine.
Cells were seeded on micro-grooved PDMS samples at
a density of 75 000 cells/cm2 for C2C12 and 10 000
cells/cm2 for nHDFs, so that they could reach the confluence 24 h after seeding. To induce differentiation
and fusion in myotubes, the cells were kept in differentiation medium, composed of DMEM with the
addition of 1% FBS, 1% insulin–transferrin–selenium
(ITS,
Sigma-Aldrich),
100 IU/ml
penicillin,
100 μg ml−1 streptomycin and 2 mM L-glutamine.
Cell-seeded samples were maintained in an incubator with controlled conditions, namely at 37 °C in a
saturated humidity atmosphere containing 95% air
and 5% CO2. C2C12 cell viability and orientation were
assessed by means of the previously mentioned Live/
Dead® assay. To assess the differentiation and the
fusion in myotubes, samples seeded with only C2C12
cells and samples seeded with nHDFs and, 24 h later,
C2C12 cells on top of them, were kept in differentiation culture medium (renewed every day) till day 7
from the induction of differentiation. Bright field images were taken at day 7 to assess the differentiation status. In addition, at the same end-point, cells
underwent fixation (by incubating them with 4% paraformaldehyde (PFA) 30 min at room temperature)
and staining with fluorescent markers: Oregon Green®
488 phalloidin (Invitrogen), to stain F-actin, and
DAPI (Invitrogen), to stain cell nuclei. Fluorescence
images were then acquired by using a confocal microscope (Inverted Microscope TiE equipped with a
Nikon Confocal Laser System A1Rsi, Nikon). The
degree of differentiation for C2C12 multinucleated
myotubes was quantitatively evaluated by considering
the fusion index. This parameter was determined by
dividing the number of nuclei within myotubes by the
total number of nuclei in the image. A larger fusion
index indicates a higher differentiation level and thus
the formation of a more mature muscle tissue [55].
Three fluorescence images were analyzed for each
sample type.
5
The DNA content of C2C12 cells was analyzed
48 h after seeding (24 h after rolling) to assess the ability of cells to survive after the rolling the entire PDMS
structure. Two different configurations were compared: unrolled substrates seeded with 75 000 cells/
cm2 and kept in culture for 48 h, and unrolled substrates (same dimensions of the previous ones), seeded
with 75 000 cells/cm2, kept in culture for 24 h, then
rolled and kept in culture for further 24 h, in the rolled
configuration. At the end-point (48 h from seeding)
the samples were removed from the original cell culture wells and placed in new wells, which were treated
with 500 μl of d-H2O. Cell lysates were then obtained
by two freeze/thaw cycles of the samples (overnight
freezing at −20 °C and 15 min thawing at 37 °C in an
ultrasonication bath) to enable the DNA to go into the
aqueous media. The DNA content in the cell lysates
was measured by using the PicoGreen kit (Invitrogen
Co., Carlsbad, CA). The PicoGreen dye binds to DNA,
and the resulting fluorescence intensity is directly proportional to the DNA concentration. Standard solutions of DNA in d-H2O at concentrations from 0 up to
6 μg ml−1 were prepared and 50 μl of standard or sample were loaded for quantification in a 96-well black
microplate. Working buffer and PicoGreen dye solution were prepared and added according to the manufacturer’s instructions (100 and 150 μl/well,
respectively). After 10 min of incubation in the dark at
room temperature, fluorescence intensity was measured on a microplate reader (Victor X3, Perkin
Elmer), using an excitation wavelength of 485 nm and
an emission wavelength of 535 nm. Three independent samples were analyzed for each configuration,
and the volume analyzed for each sample was read in
triplicate in the microplate.
2.6. FEM simulations
To envision the contraction behavior of the bio-hybrid
samples, we carried out a series of FEM simulations.
Finite element modeling allowed to predict the overall
contraction of rolled polymeric structures along their
main axis, due to properly modeled muscle cell
contraction forces.
A specific FEM software, dedicated to the modeling of nonlinear and complex soft bodies (MARC®/
Mentat® 2010, MSC Software) was used to predict the
system behavior. The bio-hybrid structure was virtually reproduced as a rolled 15 × 10 mm2 sheet, with
bilayer thicknesses of 10, 15 and 25 μm, respectively
(figure 2). The spiral internal radius was set by using
the following formula [56]:
r=
( ( (k k
2 4
E h
+ 4kE kh3 + 6kE kh2 − 3kE kh2 ϵ0
− 2kE kh ϵ0 + 4kE kh + ϵ0 + 1) ht )/
( 6k k ϵ ( 1 + k )) ),
E h 0
h
(1)
where kE = Et/Eb (Et and Eb represent the elastic moduli
of the top and of the bottom layer, respectively),
Bioinspir. Biomim. 10 (2015) 056001
L Vannozzi et al
Figure 2. Whole spiral structure section (top images) and lateral view of a structure slice (bottom) for the rolled PDMS structures
analyzed in MARC environment, referred to 10 μm (a), 15 μm (b) and 25 (c) μm bilayer thickness.
kh = hb/ht (ht and hb represent the thickness of top and
bottom layer, respectively), and ε0 represents the
imposed stretching.
The presence of the pillars was taken into consideration by adding small parallelepipeds with size of
0.03 × 0.06 × 0.30 mm3 each, while the micro-grooves
were neglected in this analysis (their height is much
lower respect the height of pillars, as later showed in
section 3.1). The PDMS material was modeled by
using PENTA elements with full integration and the
mechanical characteristics of the material were set
resembling the real ones. In particular, the elastic
modulus was set considering the values reported in
section 3.3. The application of boundary conditions
represents a key element. Each cell was represented by
the use of a couple of forces (acting in opposite directions) and applied with a specific pattern. The necessity to place the forces at precise points along the
structure required a specific mesh subdivision. Nodes
were placed at definite coordinates and used as points
where cell forces were applied. Exploiting system symmetries, the computation burden was reduced by adding a symmetry plan at the half of the height and
considering a quarter of the structure, thus allowing
the computation of a minor amount of the elements
still maintaining unaltered the mechanical behavior.
No motion conditions at one end were imposed. The
structure height was also reduced, as it constituted a
repetition of a pattern composed of pillars and forces.
Living contractile elements were modeled as
90 × 60 μm2 (cardiomyocytes) and 250 × 50 μm2
6
(skeletal myotubes) bodies (figure S1), with their main
axis aligned with the main axis of the rolled structure
(black arrow in figure 1(e)).
For each cell, two force vectors were added, which
originated from cell extremities and pointed to the cell
nucleus. During the simulation the forces were forced
to follow the structure deformation, so that they result
to be always tangential to the membrane surface. No
spacing was provided in the Y direction, while 90 and
250 μm were kept between two consecutive cells in the
X direction respectively for cardiomyocytes and skeletal myotubes (this was needed to avoid a mutual cancellation of force components). Cell force moduli were
derived from the stress value reported by Feinberg and
colleagues for primary cardiomyocytes, which was
4 mN mm−2 [19], and the force reported by Shimizu
for skeletal myotubes [57], that was 1 μN for each
myotube.
2.7. Analysis of bio-hybrid system contraction
The overall contractile behavior of a rolled structure
seeded with CMs was evaluated by means of a motion
vector analysis program implemented in Matlab
(Matlab 2012, Math Works), using the Horn–Schunk
optical flow method [58] applied on recorded videos
of the contracting PDMS structure. This non-invasive
technique is usually used to estimate flow velocities
from a sequence of frames, showing the apparent
movement of brightness patterns in an image, and it
has been already used to quantify the contractile
behavior of CMs, as reported in the literature [59]. For
Bioinspir. Biomim. 10 (2015) 056001
L Vannozzi et al
Figure 3. Thickness of PDMS membranes measured after spinning the solution on 2 × 2 cm2 Si wafers. Three independent samples
were fabricated and measured, for each spin velocity and each monomer/curing agent ratio.
a sequence of 2D images, a pixel with coordinates (x, y)
with a brightness intensity I(x, y) will move by Δx and
Δy between two image frames; the following relationship applies:
I (x , y , t ) = I (x + Δx , y + Δy , t + Δt ) .
(2)
It follows that:
Ix *Vx + Iy *Vy = −It ,
(3)
where Vx and Vy are the components of the velocity of
I(x, y, t), while Ix, Iy and It are the derivatives of the
image at (x, y, t).
Three videos of the beating rolled CM-seeded
PDMS structure (1 min duration, each) were recorded
by using an optical microscope and digitized at 100
frames/s. Data were analyzed by considering the
movement of each pixel, frame by frame, as a vector.
The flow velocity had two components, but we were
interested in the contraction along the X direction,
which corresponded to the main axis of the rolled
structure (black arrow in figure 1(e)).
A rolled PDMS structure with a monomer/curing
agent ratio of 15:1, a thickness of 25 μm and seeded
with CMs (150 000 cells/cm2) was tested to validate
the bio-actuator performance and to compare it with
FEM simulation results.
2.8. Statistical analyses
Data analysis was performed by analysis of variance
followed by Student’s t-tests, in the case of comparison
between two groups, or by Holm-Sidak tests, in the
case of comparison between more than two groups.
The significance threshold was set at 5%.
7
3. Results and discussion
3.1. PDMS membranes: thickness and surface
topographies
The thickness of PDMS membranes with different
monomer/curing agent ratios is shown in figure 3.
Results demonstrated that the thickness of the bilayer
structure can be efficiently controlled by varying the
spin velocity. In addition, greater is the mixing ratio,
smaller is the thickness, by keeping the same velocity.
In fact, by increasing the amount of the PDMS curing
agent, the fluid results less viscous and it can spread
more easily over Si wafers.
SEM and AFM imaging showed that the top layer
was characterized by grooves with a width of
9.53 ± 0.37 μm, spaced by 10.59 ± 0.39 μm and
∼1.3 μm high (figure 4(a)). This kind of topography
has been demonstrated in the literature to improve
muscle cell alignment along the groove direction [60]
and eventually to enhance the fusion of myoblasts in
myotubes [61]. An anisotropic cell distribution is of
primary importance, since muscle cells aligned along
one direction permit to generate a force directed in
that direction, thus maximizing contraction. This is
what happens in natural skeletal muscles, which have a
structural organization based on parallel muscle
fibers.
Instead, the bottom layer was provided with rectangular pillars characterized by a height of
23.86 ± 0.67 μm, a width of 62.96 ± 1.03 μm and a
length of 322.95 ± 1.32 μm (figure 4(b)). These pillars
have the aim of separating the concentric PDMS layers, thus reducing the stress on cells during the substrate rolling phase and to assure the flow of culture
medium inside the inner channels, to keep cells alive
and to improve long-term cultures.
Bioinspir. Biomim. 10 (2015) 056001
L Vannozzi et al
Figure 4. Analysis of top and bottom layer topographies. (a) SEM (left) and AFM (right) images showing the parallel microgrooves
obtained for the top layer (profile in the red box); (b) SEM images highlighting the pillars obtained for the bottom layer.
3.2. Achievement of 3D structures
To obtain the rolled PDMS structures, a step-by-step
procedure was followed, by using a custom assembly
system fabricated by means of traditional machining
tools.
The custom extraction, stretching, supporting (in
Delrin) and clamping (in Teflon) system components
contributed to increase the efficiency of the 3D rolled
structure fabrication procedure, when relatively thin
PDMS structures were used. In fact, the detachment of
PDMS from the Si molds and its handling represented
critical issues when PDMS membrane thickness
decreased. The use of PVA as sacrificial layer in aqueous environment allowed to apply rather limited
stresses on the membranes, while the custom assembly
system permitted to efficiently carry on the procedure.
First, the PDMS top layer was detached from the Si
mold in deionized water after the dissolution of the
PVA layer, and the floating membrane was collected
by means of the extraction system (figure 5(a)). This
was then coupled with the traction system
(figure 5(b)) and the membrane was blocked at its
extremities. Then the extraction part was removed and
the layer was stretched by 40% of its initial length
8
(figure 5(c)). Top layer stretching was allowed by two
guide rails which secured the movement along one
direction, while two external clamps were screwed on
the traction system to block the stretched membrane
when desired. Subsequently, the semi-cured PDMS
bottom layer, still attached to the Si mold, was placed
below the stretched membrane by using the supporting system (figure 5(d)). After a thermal treatment
(80 °C for 30 s) to complete the PDMS polymerization, the bilayer was immersed in water, to detach the
bottom Si mold (figure 5(e)). Then, the clamping system, provided with nylon screws, allowed to block the
bilayer in its stressed status (figure 5(f)) and to introduce it in a standard 6-well plate for cell cultures
(figure 5(g)), after sterilizing it (this was achieved by
keeping the whole structure for 30 min in a PBS solution provided with a high concentration of penicillin
and streptomycin). Finally, after cell culture, the system was extracted from the well, one of its end was cut
and a rolled structure was obtained (figure 5(h),
video S1).
By means of this step-by-step procedure, we
obtained rolled PDMS structures with an Archimedean spiral-like shape, characterized by the two
Bioinspir. Biomim. 10 (2015) 056001
L Vannozzi et al
Figure 5. Different phases of the fabrication procedure, supported by a custom assembly system. An extraction system (a) allowed to
collect the PDMS membrane from water. A traction system (b) allowed to stretch it along the longitudinal axis (c). A supporting
system (d) permitted the coupling with the semi-cured bottom layer. After a thermal treatment, the PDMS bilayer was immersed in
water to detach the Si-support of the bottom part (e), blocked in its stressed status by means of a clamping system (f) and placed in
culture (g). When desired, one end was cut and a rolled structure obtained (h).
Figure 6. SEM image of a 3D rolled PDMS structure.
mentioned topographies: a micro-grooved surface on
the internal walls and rectangular pillars on the external walls (figure 6).
3.3. Chemical functionalization and mechanical
characterization
By analyzing fluorescence intensity, emitted by the
labeled fibronectin immobilized on the PDMS surface,
we demonstrated that the genipin-based procedure
9
assured the maintenance of a stable coating for at least
13 days (figure S2(a)). The green signal component
showed no statistically significant variations. This
means that genipin-based chemical functionalization
allowed to maintain a stable fibronectin coating, thus
assuring that muscle cells could remain adherent to
the PDMS substrate within this time-frame.
We also checked if such chemical functionalization procedure could influence the PDMS mechanical
Bioinspir. Biomim. 10 (2015) 056001
L Vannozzi et al
Table 1. Elastic moduli for each PDMS mixing ratio, independent
from thicknesses. The last column refers to the comparison between
bare PDMS and genipin-treated PDMS samples for each mixing
ratio: * = p < 0.05, ** = p < 0.01.
Mixing
ratio
Bare PDMS
elastic modulus (MPa)
Genipin-treated
PDMS elastic
modulus (MPa)
Statistical
comparison
5: 1
10: 1
15: 1
20: 1
2.553 ± 0.087
1.808 ± 0.073
1.013 ± 0.068
0.573 ± 0.047
2.619 ± 0.045
2.106 ± 0.095
1.308 ± 0.072
0.721 ± 0.040
**
**
**
**
properties. Stiffness is one of the cues that influence
muscle cell behavior, especially their ability to form
differentiated and functional skeletal myotubes.
Moreover, in our case this feature conditioned substrate flexibility and, consequently, the bio-actuator
performance. The aim would be, in fact, to obtain a
stiffness as low as possible for PDMS substrates, in
order to limit the mechanical opposition during the
contraction of muscle cells. We carried out tensile tests
(figure S2(b)), in which we observed that the genipinbased treatment slightly increased the stiffness of
PDMS substrates (table 1).
The treatment with genepin increased the elastic
moduli of PDMS structures for each mixing ratio.
However, for long-term cultures on PDMS substrates
we need to accept this compromise, since facing the
hydrophobic recovery represents a crucial issue.
The gap with the stiffness value reported in the literature for an optimal maturation of skeletal muscle
tissues (∼14 kPa [62]) is rather large. However, the
option to use higher monomer/curing agent ratios in
order to achieve lower elastic modulus values is hardly
feasible in our case, due essentially to manipulability
issues. In fact, the bottom PDMS layer of the structure
is semi-cured and per se almost impossible to manipulate when its elastic modulus goes below ∼500 kPa
and when its thickness goes below 10 μm. Therefore,
we considered such values, 20:1 for the monomer/curing agent ratio and 10 μm for the bilayer thickness of
the bottom layer, as our limits.
It could be argued that these mechanical characteristics would represent a strong limitation for a
correct development of muscle tissue on these substrates. Anyhow, in the case of skeletal muscle cells, we
envisioned to overcome this issue by providing the
system with a feeder layer of fibroblasts, on top of
which myoblasts can be seeded and differentiated.
This strategy already proved to be effective in providing muscle cells both with a suitable mechanical interface and with specific cytokines, produced by
fibroblasts, which help muscle tissue maturation
[31, 54]. Nonetheless, the force generation of myocytes was demonstrated also to be stiffness dependent,
as on stiffer substrates cells make use of larger forces
and spread out across the substrate, leading to a
nuclear compression and an increase of the horizontal
10
component of contraction forces respect to the vertical
one [63, 64].
3.4. FEM simulations
Simulations performed in MARC environment
revealed that, under the effect of the contraction forces
of muscle cells, each rolled PDMS structure underwent
an axial shortening. The bio-actuator overall contraction depended on the cell type, on the PDMS elastic
modulus
and
on
the
bilayer
thickness
(figures 7(a), (b)).
Obviously, higher contraction values were assured
by the most flexible substrates, characterized by lower
elastic moduli and/or thickness values. Results showed
that primary CMs would be able to contract the rolled
PDMS structure up to ∼25% of its initial length. Skeletal muscle cells would assure a much lower device contraction (up to ∼0.13%).
In silico differentiated skeletal myotubes showed
lower contraction values due to the intrinsic difficulty
to obtain mature and functional myotubes. However,
future evolutions of the system may include both technological and biological improvements. Concerning
technological ones, the current boundaries of elastic
modulus and thickness may be expanded, by changing
material or by advancing the fabrication procedure.
Concerning biological improvements, the differentiation of skeletal myotubes may be pushed, by means of
electro-mechanical stimulation [65], intracellular stimulation [31] or other cell engineering techniques,
thus achieving force values much higher than 1 μN
and more significant bio-actuator contractions.
3.5. In vitro tests
First, we evaluated cell viability on PDMS surfaces,
focusing our attention on the influence of topography
on cell behavior. Results showed that on microgrooved
PDMS samples, the majority of CMs (figure 8(a)) and
C2C12 cells (figure 8(b)) was viable (green-colored),
while only a small percentage was dead or necrotic
(red-colored). In addition, the presence of grooves
influenced the orientation of muscle cells: they aligned
along the groove axis (highlighted by white arrows),
thus exhibiting a high anisotropy.
Then, we evaluated the ability of C2C12 muscle
cells to differentiate and fuse into myotubes on the
treated substrates provided with grooved surfaces and
chemically treated with genipin, in order to achieve a
stable fibronectin coating. In parallel, we studied cocultures made of a layer of nHDFs below the C2C12
muscle cell layer. These cultures were maintained for 7
days, by replacing the differentiation medium on a
daily basis. Bright field and fluorescence (confocal)
images (figure 8(c)) show that C2C12 cells formed
highly aligned myotubes, parallel to the groove direction, at day 7. A high number of multinucleated structures can be observed, thus highlighting the myogenic
potential of these substrates. Figure 8(d) shows bright
Bioinspir. Biomim. 10 (2015) 056001
L Vannozzi et al
Figure 7. (a) FEM-based simulation results classified for cell type (primary cardiomyocytes and differentiated skeletal muscle cells),
PDMS mixing ratio (5:1, 10:1, 15:1 and 20:1) and bilayer thickness (10, 15 and 25 μm). Images depict the final contraction of each
rolled PDMS structure analyzed. (b) Graphs reporting the bio-actuator contraction (shortening percentage, respect to the initial
length) for the two cell types (primary cardiomyocytes and differentiated skeletal muscle cells), the different PDMS mixing ratios and
the different bilayer thickness values.
field and fluorescence (confocal) images of the same
PDMS substrates, previously seeded with fibroblasts,
and then with C2C12 cells.
Co-cultures with fibroblasts aimed at enhancing
the C2C12 differentiation capability on PDMS substrates. From previous literature reports, it was
demonstrated that a feeder layer of nHDFs was able to
enhance the formation of multinucleated myotubes,
both providing secretion of growth factors [54] and a
softer layer closer to the optimal substrate stiffness
value for muscle differentiation (∼14 kPa) [62]. The
quantitative assessment of the C2C12 differentiation
level was obtained by evaluating the fusion index
(figure 8(e)). A statistically significant difference was
observed after 7 days of culture in the differentiation
medium for C2C12 cells without (0.64 ± 0.04) and
with (0.76 ± 0.05) a supporting nHDFs layer. Fibroblasts were able to support an efficient cell fusion,
increase the differentiation ability of C2C12 cells and
their ability to develop anisotropically multinucleated
myotubes. The formation of more mature myotubes
will allow the development of contractile muscle capable to exert larger forces.
The rolling phase is the last step of the procedure.
To evaluate the ability of cells to survive after rolling
the entire structure, C2C12 cells were seeded on the
(stretched) top layer for 24 h, then the clamped system
11
was extracted from the 6-well plate and one extremity
was cut with a blade. When rolled up, the aligned muscle cells resulted longitudinally oriented and the pillars
separated the membrane layers, thus providing sufficient space to keep cells alive. At this time-point, we
evaluated the viability of C2C12 muscle cells inside the
tubular structure after rolling by means of the Live/
Dead® kit. Figure 9(a) shows that the rolling did not
cause evident damages on the cells, which remained
highly viable, as demonstrated by the green-colored
concentric cell layers (the image is a top view of the
cell-seeded rolled structure).
The results of a quantitative analysis of DNA content
for the unrolled and rolled structures are shown in
figure 9(b). These results permitted to further demonstrate the viability of C2C12 cells even after the rolling
phase. The DNA content measured for rolled
PDMS structures (4.42 ± 1.17 μg ml−1) was slightly
smaller than the one measured for unrolled structures
(5.61 ± 1.62 μg ml−1). However, this difference was not
statistically significant (p = 0.198). This result demonstrates that PDMS rolling does not kill a significant number of cells and that the pillars are effective in separating
each concentric layer, thus guaranteeing cell viability.
Self-assembled tubular structures have been
proved to represent a valid approach to recapitulate
milli- and micro-engineered models of blood vessels,
Bioinspir. Biomim. 10 (2015) 056001
L Vannozzi et al
Figure 8. Viability and orientation of CMs (a) and C2C12 cells (b) cultured on a microgrooved PDMS surface. Green-colored cells are
viable, red-colored cells are dead or necrotic. The white arrows indicate the groove axis in both cases. (c) Bright field image and
fluorescence (confocal) images of C2C12 cells at day 7 of differentiation, on a grooved PDMS substrate. (d) Bright field and
fluorescence (confocal) images of C2C12 + nHDF cells at day 7 of differentiation on a grooved substrate. The red boxes evidence
magnified areas of the fluorescence images. Scale bars are 50 μm. (e) Comparison between the fusion index calculated for C2C12 cells
and for the C2C12 + nHDF co-culture. ** = p < 0.01.
in previous studies [42]. Our results demonstrate that
this model can also effectively mimic the internal
structure of a muscle fiber, with the presence of
12
parallel myotubes along the longitudinal direction,
kept viable thanks to the intrinsic shape of the selfassembled structure.
Bioinspir. Biomim. 10 (2015) 056001
L Vannozzi et al
Figure 9. (a) Fluorescence image of a PDMS structure, after rolling (top view). Live cells are green-colored. Arrows evidence the cellprovided concentric layers of the structure. Scale bar is 100 μm. (b) Comparison between the DNA concentrations measured for
rolled and unrolled PDMS structures. Both structures were initially in a rolled configuration and were seeded with cells. After 24 h, the
rolled PDMS structure was self-assembled by cutting one end of the membrane, while the other sample type was kept unrolled. Both
sample types were cultured for further 24 h, before DNA quantification.
3.6. Assessment of bio-actuator contraction
The results of FEM simulations, reported in
section 3.4, were validated by analyzing the in vitro
actual behavior of a specific case, referred to a rolled
PDMS structure (15:1 mixing ratio, with a bilayer
thickness of 25 μm). Figure 10(a) shows FEM images
of the mentioned sample, characterized by a shortening of 1.32% of its length under the action of
primary CMs. Its contraction movement is showed
also in video S2.
Videos of the spontaneous contraction movements of the tested rolled PDMS structure were captured in bright field and analyzed as described in
section 2.7. The optical tracking permitted to record
the movement in a non-invasive way, without interfering on muscle cells activity. Figure 10(b) shows a
portion of the CM-seeded rolled structure. A depiction of such portion is shown by the CAD image on the
right; the white arrows indicate the movement direction of pixel areas, explained by the optical flow theory. The PDMS structure preferentially shrunk along
its longitudinal axis due to the contraction forces of
the aligned CMs (inset in figure 10(b)).
Figure 10(c) shows the structure contraction over
time. Each peak represents the contraction and relaxation phases of the device. The measured average device
contraction was 2.19 ± 0.54%. This corresponded to a
longitudinal shortening of 219 ± 54 μm, being the
actuator length equal to 1 cm. From the analysis, we
also obtained additional parameters regarding the bioactuator contraction performance, such as the duration and the beating rate, which were 0.61 ± 0.08 s and
1.14 ± 0.21 Hz, respectively.
Figure 10(d) shows the comparison between the
contraction value predicted by FEM simulation
(1.32%) and the average value measured in vitro. The
slight difference observed can be due to a series of
13
factors. Besides unavoidable distortions from the ideal
case due to variability in material properties and cell
behavior, the stress value used for CM-based simulations (4 mN mm−2, [19]) can be affected by significant
variations when different CM isolation strategies are
employed and when different surface chemistries and
matrix stiffness are considered. Moreover, the total
number of cells adhered on the substrate may be
underestimated, in the FEM environment. Overall, the
validation of FEM simulation results can be considered successful. Thus, the predicted contraction
values for different thicknesses, elastic moduli and cell
types shown in figure 7 can be considered realistic.
CMs are interesting bio-elements that have been
explored for the fabrication of self-actuated biohybrid systems, since these cells are synchronous and
do not need external power to be triggered [66]. However, as mentioned in the Introduction, skeletal muscle cells constitute an attractive alternative to CMs
in situations that require high spatial and temporal
controllability over contraction. Localized stimulation
of engineered skeletal muscle constructs has been
demonstrated [32], although the forces generated by
in vitro constructs, at present, still remain considerably
smaller in comparison with those that are generated
in vivo [12].
Our data demonstrated the feasibility of using cardiac muscle cells to actuate deformable rolled 3D
PDMS structures, as bio-hybrid devices, but also validated advanced FEM simulations, which predicted
bio-actuator contraction in the case of rolled structures coupled with mature contractile skeletal myotubes. To achieve such ambitious goal, proper
stimulation tools and additional dedicated technologies are needed [65, 67], which go beyond the scope of
this paper.
Bioinspir. Biomim. 10 (2015) 056001
L Vannozzi et al
Figure 10. (a) FEM simulations in MARC environment: lateral view of the analyzed rolled structure before muscle cells contraction
(left); bio-actuator contraction in the case of primary cardiomyocytes (center); stress analysis of the rolled PDMS structure, that
highlights the localized action of muscle cells (right). (b) Representative video frame related to the motion analysis of the rolled PDMS
structure (left) and corresponding CAD representation (right). The magnified picture shows CMs adhered to the internal walls of the
rolled structure. (c) Contraction behavior of the bio-actuator (actuator length is 10 mm) during 10 s. (d) Comparison between the
predicted (FEM simulation) and the measured (in vitro tests) contraction.
4. Conclusion
We demonstrated the feasibility of an innovative
approach for the development of 3D self-assembled
engineered substrates, to be used as enabling matrices
in the field of bio-hybrid actuation. Our results highlighted the possibility to develop rolled 3D PDMS
structures, characterized by different elastic moduli,
by exploiting a SIRM technique. These structures were
provided with defined topographical cues, able to
organize cells and influence their action. Different
surface topographies permitted to obtain the alignment of muscle cells on the top layer (thus orienting
14
their contraction along the longitudinal axis), and at
the same time to maintain a gap among concentric
layers, which guarantees cell viability. A specific surface chemical treatment allowed to obtain long-term
stable coatings over two weeks, overcoming the
problem of PDMS hydrophobic recovery. Our results
demonstrated that these structures favor the adhesion
and alignment of primary CMs and the adhesion,
proliferation and differentiation of skeletal muscle
cells, promoting the formation of myotubes, also when
supported by a fibroblast feeder layer. Advanced FEM
simulations, performed in MARC environment,
allowed to predict bio-actuator contraction for a range
Bioinspir. Biomim. 10 (2015) 056001
L Vannozzi et al
of thicknesses, elastic moduli and two cell types. By
considering a specific case (PDMS structure with a
mixing ratio of 15:1 and a bilayer thickness of 25 μm),
based on cardiomyocytes, simulations revealed an
overall system contraction of 1.32%. A bio-actuator
with these characteristics was also tested in vitro, by
seeding primary cardiomyocytes on its surface. The
system showed an overall contraction of 2.19 ± 0.54%,
comparable with the values obtained with the FEM
simulations. Since simulations were validated with the
in vitro experimental results, predictions of bioactuator contraction at lower thicknesses (e.g. 10 μm)
and elastic moduli (e.g. 0.573 MPa) can be considered
realistic, thus highlighting the potential of these
systems for reaching higher contraction values (up to
25%). Future work will be focused on the development
of the additional technologies needed to achieve
mature and functional skeletal myotubes and to trigger
their contraction, when required.
Acknowledgments
The authors would like to deeply thank Dr Gianni
Ciofani and Dr Giada Graziana Genchi, for their advices
concerning the PDMS surface functionalization procedure, Dr Tercio Terencio for the acquisition of fluorescence images with the confocal microscope, Mr
Francesco Corucci for his help in the development of the
motion vector analysis program implemented in Matlab
and Dr Maurizio Follador for the help in setting up FEM
simulations on MARC®/Mentat®. Furthermore, they
deeply acknowledge Mr Carlo Filippeschi for his support
during all the clean-room procedures, Mr Nicodemo
Funaro and Mr Andrea Melani for the fabrication of
custom elements in the BioRobotics Institute workshop.
This work was supported in part by the GeT Small
project (TarGeted Therapy at Small Scale), funded by
the Scuola Superiore di Studi Universitari e di Perfezionamento Sant’Anna (Pisa, Italy).
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