Cell-Fiber Interactions on Aligned and Suspended Nanofiber Scaffolds

Delivered by Publishing Technology to: kathryn danner
IP: 130.126.36.198 On: Tue, 27 Aug 2013 15:45:32
Copyright: American Scientific Publishers
REVIEW
Copyright © 2013 American Scientific Publishers
All rights reserved
Printed in the United States of America
Journal of
Biomaterials and Tissue Engineering
Vol. 3, 355–368, 2013
Cell-Fiber Interactions on Aligned and
Suspended Nanofiber Scaffolds
Kevin Sheets1 †
, Ji Wang2 4 †
, Sean Meehan3
, Puja Sharma1
, Colin Ng3
,
Mohammad Khan3 ‡
, Brian Koons3
, Bahareh Behkam1 3 4
,
and Amrinder S. Nain1 2 3 4 ∗
1
School of Biomedical Engineering and Sciences, Virginia Tech, Blacksburg, 24060, VA
2
Department of Engineering Science and Mechanics, Virginia Tech, Blacksburg, 24060, VA
3
Department of Mechanical Engineering, Virginia Tech, Blacksburg, 24060, VA
4
Macromolecules and Interfaces Institute, Virginia Tech, Blacksburg, 24060, VA
Cells interact with fibrous extracellular matrix (ECM) which exhibits varying degrees of alignment
throughout the body. In this review, we highlight cell-aligned fiber interactions using the recently-
developed Spinneret-based Tunable Engineered Parameters (STEP) fiber manufacturing technique
which creates fibrous scaffolds with precise control on fiber diameter, spacing, orientation, and hier-
archy. Through manipulation of each individual parameter, we show that multiple cell types (including
cancerous) display unique changes in cell shape, cytoskeletal arrangement, focal adhesion distribu-
tion, and migration speed while interacting with the suspended STEP fibers. In addition to single-cell
responses, we present our findings on higher-level monolayer formation and wound healing mod-
els, stem cell differentiation, and hepatic engineering. These single-cell and population-level studies
are conducted in the presence of aligned topographical cues that resemble native ECM. Knowl-
edge gained from such studies will help create more accurate in vitro fibrous scaffolds used for
the advancement of tissue engineering, disease treatment, and the development of diagnostic and
drug testing platforms.
Keywords: Fiber Alignment, ECM, Nanofiber, Cellular Dynamics, Mechanobiology.
CONTENTS
1. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 355
2. Step Fiber Manufacturing: Role of Scaffold
Parameters on Single Cell Behavior . . . . . . . . . . . . . . . . . . 358
2.1. Control of Fiber Diameter . . . . . . . . . . . . . . . . . . . . . 359
2.2. Inter-Fiber Spacing . . . . . . . . . . . . . . . . . . . . . . . . . 362
2.3. Fiber Orientation . . . . . . . . . . . . . . . . . . . . . . . . . . 362
2.4. Multi-Layer Assemblies and Hierarchical
Structures . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 363
2.5. Role of Structural Stiffness . . . . . . . . . . . . . . . . . . . . 364
3. Population Cell Behavior and Applications
in Tissue Engineering . . . . . . . . . . . . . . . . . . . . . . . . . . . 365
4. Concluding Remarks and Future Directions . . . . . . . . . . . . . 366
5. Experimental Details . . . . . . . . . . . . . . . . . . . . . . . . . . . 366
5.1. Fiber Manufacturing and Pillar Design . . . . . . . . . . . . . 366
5.2. Cell Culture and Imaging . . . . . . . . . . . . . . . . . . . . . 366
5.3. Statistical Analysis . . . . . . . . . . . . . . . . . . . . . . . . . 366
Acknowledgments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 366
References and Notes . . . . . . . . . . . . . . . . . . . . . . . . . . . 366
∗
Author to whom correspondence should be addressed.
†
These two authors equally contributed to this work.
‡
Mohammad Khan was a member of the STEP lab from 2010–2011.
1. INTRODUCTION
In the body, cells attach to and interact with a fibrous,
mesh-like extracellular matrix (ECM) which exhibits vary-
ing degrees of alignment throughout the body both spatially
and temporally.1
Collagen in tendons, for instance, begins
as poorly aligned fibrils in early development but becomes
highly aligned in later stages.2
The fully-developed ten-
don tissue, which is approximately 80% ECM mass by
dry weight, relies heavily on hierarchical assembly of
highly aligned ECM fibrils to provide functional muscle-
bone interfaces.3 4
ECM facilitates cell attachment, cell–
cell contacts, provides soluble growth factors and presents
gradients of elasticity to cells which directly control cell
fate.5
Cells attach to the ECM through integrin-mediated
focal adhesion complexes (FACs), which physically link
the cell to the ECM protein networks.6
Currently, it is
widely accepted that mechanical cues and forces con-
ducted through these cell-ECM junctions play a domi-
nant role in cellular and sub-cellular biological functions.7
Common examples of cellular/tissue response to such
J. Biomater. Tissue Eng. 2013, Vol. 3, No. 4 2157-9083/2013/3/355/014 doi:10.1166/jbt.2013.1105 355

IP: 130.126.36.198 On: Tue, 27 Aug 2013 15:45:32
REVIEW
Cell-Fiber Interactions on Aligned and Suspended Nanofiber Scaffolds Sheets et al.
Kevin Sheets is a Ph.D. candidate at the Virginia Tech-Wake Forest School of Biomed-
ical Engineering and Sciences. He earned his Bachelor of Science in Materials Science
and Engineering from Virginia Tech in 2009 and his Master of Science in 2010. Kevin is
working on understanding fundamental cell-fiber interactions to study cell migration, force
modulation, and disease models using STEP scaffolds.
Ji Wang is a Ph.D. candidate in the Department of Engineering Science and Mechanics at
Virginia Tech. Ji earned his Bachelor of Science degree in Polymer Science and Engineering
from the Beijing Institute of Textile Technology, China. He has worked on determining
the spinnability of various polymers using the STEP technique and also on single fiber
mechanical/material characterization.
Sean Meehan earned his Master of Science in Mechanical engineering from Virginia Tech
in 2013. He earned a Bachelor of Science in Mechanical Engineering and a minor in Physics
from Rowan University in Glassboro, NJ. Sean’s research focused on the effects of structural
stiffness (N/m) on single cell migration mechanics along a single nanofiber. He is currently
working at Duke University Medical Center in Durham, NC.
Puja Sharma is Ph.D. student at Virginia Tech-Wake Forest School of Biomedical Engi-
neering and Sciences. She received a Bachelor of Science in Biology (Honors) and minors in
Physics and Math from Hollins University, Virginia. Her research in the STEP Lab focuses
on studying single cancer cell behavior (migration, cytoskeleton and blebbing dynamics) as
a function of nanofiber structural stiffness, protein coatings and fiber curvature. She is also
studying cancer cell force modulation in a tunable mechanistic environment.
Colin Ng graduated with a Bachelor’s in Mechanical Engineering from Virginia Tech in
2012, and is currently a Masters student in the same department. His research focuses on
the cellular investigation of wound healing on nanofiber scaffolds, and single cell migratory
forces.
356 J. Biomater. Tissue Eng. 3, 355–368, 2013

IP: 130.126.36.198 On: Tue, 27 Aug 2013 15:45:32
REVIEW
Sheets et al. Cell-Fiber Interactions on Aligned and Suspended Nanofiber Scaffolds
Mohammad Khan was a member of STEP Lab from 2010–2011. He is currently attending
University of Illinois Urbana-Champaign graduate school.
Brian Koons is a graduate student studying Mechanical Engineering with an emphasis on
biomedical applications at Virginia Tech. He earned his Bachelor of Science in mechanical
engineering from Virginia Tech, 2013. Currently, his research focus is on integrating func-
tional suspended acrobatic hepatic monolayers within a bioreactor device. Brian also does
extensive work in creating customized fiber networks to determine cancer cell protrusion
dynamics.
Bahareh Behkam earned her B.Sc. degree from Sharif University of Technology (Iran),
and her M.Sc. and Ph.D. degrees from Carnegie Mellon University, all in Mechanical Engi-
neering. She has been on the faculty in the Department of Mechanical Engineering with
an affiliate appointment in School of Biomedical Engineering and Sciences and Macro-
molecules and Interfaces Institute at Virginia Tech since August 2008. Dr. Behkam is the
director of MicroN BASE Laboratory at Virginia Tech. Her research interests include bio-
hybrid micro/nano systems and living machines, biological microfluidics, wound healing
and physical chemistry of cell-surface interaction.
Amrinder S. Nain earned his B.E degree from Manipal Institute of Technology (Mechanical
Engineering, India), and his M.Sc. (Chemical Engineering) and Ph.D. (Mechanical Engi-
neering) degrees from Carnegie Mellon University. He has been on the faculty in the Depart-
ment of Mechanical Engineering with an affiliate appointment in the School of Biomedical
Engineering and Sciences, the Department of Engineering Science and Mechanics and the
Macromolecules and Interfaces Institute at Virginia Tech since August 2009. Dr. Nain is the
director of the STEP Laboratory at Virginia Tech. His research interests include advanced
materials, tissue engineering and single cell disease models.
forces include smooth muscle cell remodeling of arte-
rial walls to accommodate changes in flow rate,8
soft
tissue remodeling (collagen fibers),9
and bone loss in
microgravity.10–12
In addition, external forces carried by
the ECM guide the cell through a series of mechanical
cues, each triggering a cascade of biological responses.
For example, mesenchymal stem cells (MSCs) differenti-
ate towards osteogenic phenotypes at low strains and car-
diovascular lineages at high strains.13 14
Cell migration has
also been shown to be a product of a cell’s ability to gen-
erate and align these traction forces, and cancer cells are
thought to metastasize by separating from a primary tumor
and migrating along ECM towards blood vessels.15
Due to the varied alignments, compositions, and physical
properties ECM exhibits throughout the body, mimicking
cell-ECM interactions in vitro using state-of-the-art fiber
manufacturing platforms can be challenging.1 16
Never-
theless, a number of recent works have established key
scaffold parameters to directly influence cell behavior
including fiber diameter, spacing (or porosity), align-
ment, and hierarchy.17–20
Cellular interaction with fibers
occurs on various lengthscales affecting fundamental cell
J. Biomater. Tissue Eng. 3, 355–368, 2013 357

IP: 130.126.36.198 On: Tue, 27 Aug 2013 15:45:32
REVIEW
Fig. 1. Fibrous scaffold hierarchy and mechanobiological relevance
(shaded region depicts current manufacturing challenges).
behavior including migration, proliferation, differentiation,
and eventual tissue remodeling.21 22
The lengthscales can
be separated into four characteristic domains: macroscale
(> 100 m), microscale (1 m–100 m), sub-microscale
(100 nm–1 m), and nanoscale (< 100 nm), with each
domain detailing biological aspects critical for the scaffold
to capture (Fig. 1).23 24
At the macroscale, the scaffold should allow cell–cell
fusion and monolayer formation necessary for tissue gen-
eration. Control at the microscale (∼ cellular dimensions)
should allow individual cell dynamics and cell–cell inter-
actions to be studied where fiber spacing and orienta-
tion control single-cell decision making.18 19 23 25
While
the macro and microscales have been extensively stud-
ied in the past, the sub-micro and nanoscales are actively
being investigated for understanding fundamental single-
cell behavior (biophysical and biochemical) using a num-
ber of different fiber manufacturing techniques.26 27
These
studies aimed at elucidating single-cell response to alter-
ations in fibrous environments smaller than cellular dimen-
sions have the potential to unveil previously undescribed
modes of cell attachment, force production, and migra-
tion mechanisms, thereby providing fresh insights into tis-
sue engineering and also advancing our understanding of
mechanisms of disease onset, progression, and eventual
treatment.
Fiber fabrication technologies have advanced tremen-
dously over the past decade to the point where macroscale
and microscale resolution is rather easily achieved.28
How-
ever, studies aimed at the single cell-fiber interaction level
are still in their infancy and common fiber manufactur-
ing technologies require either new development or refine-
ment to achieve such resolution (shaded region of Fig. 1).
Processes such as electrospinning are excellent platforms
for depositing fibers of a very wide variety of mate-
rials but require specialized techniques to achieve fiber
alignment in single and double layers (Supplementary
Information).28–31
The manufacturing challenges in con-
trolling diameter, spacing, and alignment restrict the scope
to which cell-fiber interactions can be investigated using
electrospinning methods.32
In this review, we highlight
the use of the STEP manufacturing platform to create
fibrous scaffolds of well-defined diameters, spacing, ori-
entation, and hierarchical assembly and its flexibility in
investigating single-cell behavior at different length scales.
We then present our results in extending STEP technol-
ogy to develop cell population level applications in tissue
engineering.
2. STEP FIBER MANUFACTURING: ROLE
OF SCAFFOLD PARAMETERS ON
SINGLE CELL BEHAVIOR
STEP is a pseudo-dry spinning nanofiber fabrication tech-
nique which does not rely on an electric field to stretch
the solution filament. This method is fundamentally differ-
ent from electrospinning in that the absence of an electric
field eliminates random fiber deposition associated with a
charged Taylor cone and distant target,33
allowing arrays
of highly aligned fibers to be created. STEP consists of
both continuous (high density fiber array) and sequen-
tial (single fiber) fiber deposition approaches.34–36
In the
continuous approach outlined in this review, polymer solu-
tion is pumped through a nozzle and pulled into a sin-
gle filament by a rotating substrate (Fig. 2(A)). As the
rotating substrate translates at a user-defined speed ( ),
well-aligned fibers with desired scaffold porosity (inter-
fiber spacing) are obtained. In addition to depending on
polymer molecular weight and concentration, fiber diam-
eter is observed to scale with rotational speed of the sub-
strate ( ) (Fig. 2(B)).34
Multi-layer fiber networks are
obtained by depositing fiber arrays over the previous layer,
as demonstrated in Figure 2(C). By varying fiber diam-
eters and the angles between different fiber layers, fiber
assemblies with desired patterns can be fabricated in mul-
tiple layers (Figs. 2(D, E)). Fibers of different surface and
interior features are obtained by controlling fiber solidifica-
tion and phase separation processes during fabrication. As
shown in Figure 2(F), porous fiber surfaces are obtained
at high relative humidity (RH), which is attributed to a
combination of vapor induced phase separation (VIPS) and
temperature induced phase separation (TIPS) processes.37
Prolonged solidification leads to the creation of wrin-
kled fiber surfaces as a result of shrinkage mismatch
between the shell and the core (Fig. 2(G)).38 39
Further-
more, controlling the solvent volatility in the manufactur-
ing process can lead to hollow tube structures as shown
in Figure 2(H). STEP manufacturing has been extended
to deposit highly aligned fiber assemblies of different
polymers including poly(lactic-co-glycolic acid) (PLGA),
poly(methyl methacrylate)(PMMA), poly(ethylene oxide)

IP: 130.126.36.198 On: Tue, 27 Aug 2013 15:45:32
REVIEW
Fig. 2. (A) Schematic illustration of continuous STEP manufacturing process, (B) typical fiber diameter profile as a function of substrate rotational
speed ( ), in this case normalized to 400 nm for PS 1.8 M g·mol−1
at 10% by wt. (C) SEM images of a crisscross fiber network with controlled fiber
spacing, (D) fiber assembly incorporating different diameters: d1 = 800 nm, d2 = 400 nm, d3 = 80 nm, d4 = 160 nm, (E) three-dimensional scaffolds
with controlled pore sizes (1 m × 2 m) and (2 m × 2 m), with the scale bar in insert representing 1 m, (F) porous, (G) wrinkled, and (H)
hollow fibers, unidirectional and cross-hatch scaffolds of (I) PLGA, (J) PMMA, (K) PEO fiber assemblies, (L) PU, (M) collagen and (N).
(PEO), polyurethane (PU), collagen (type I), and fib-
rinogen in single and multiple layers of varying diame-
ters with control of porosity, morphology, and orientation
(Figs. 2(I)–(N)). Recently, we have also demonstrated well
aligned metal oxide such as TiO2 and BaTiO3 by combin-
ing STEP with sol gel process.40 41
The suspended nanofibers cause cells to react to surface
curvature and dimensionality that flat substrates inherently
mask, thus providing a unique platform for investigat-
ing cell-fiber interactions (Supplementary Videos 1–4).
Fig. 3. Suspended fiber design space, which accounts for the substrate’s
elastic modulus (E), structural stiffness (k), diameter, inter-fiber spacing,
orientation, and hierarchical assembly, compared to traditionally-studied
flat substrates.
As shown in the schematic of Figure 3, cells on fibers
sense and respond to changes in curvature (or diameter),
form spindle or parallel shapes due to differences in fiber
spacing, spread and attach differently on oriented fibers,
and finally form kite-like polygonal shapes on hierarchical
assemblies. We speculate that these parameters along with
the combination of material and structural stiffness in sus-
pended fiber configurations leads to differences in force
generation (F ) compared to flat substrates, which elicits
diverse behavior as measured by a large number of metrics
including cytoskeletal arrangement, nuclear shape index,
FAC cluster length and distribution, migration speed, pro-
liferation, and differentiation.
2.1. Control of Fiber Diameter
2.1.1. Molecular Chain Entanglement
Solvent-based STEP fiber spinning requires sufficient
molecular chain entanglements in order to maintain
smooth, continuous fiber formation. Taking polystyrene

IP: 130.126.36.198 On: Tue, 27 Aug 2013 15:45:32
REVIEW
Fig. 4. Isodiametric design space of PS fibers, Insert 1: beaded fibers formed below Ce, Insert 2–3: uniform fibers formed from semi-dilute to
concentrated regions, Insert 4: macro fibers formed from highly concentrated solutions, reproduced with permission.32
(PS) as a model spinning system, the solution entangle-
ment number (ne soln is defined as:42
ne soln =
pMW
M∗
e
(1)
where p is the polymer volume fraction and M∗
e is the
corrected PS entanglement molecular weight (1 25 Me =
16 600 g · mol−1
). For electrospinning, fiber initiation
(beaded fibers) starts at (ne soln ∼ 2 0 and uniform fibers
(defect free fiber) form at ne soln ∼ 3 5.42 43
In contrast,
using the STEP technique, smooth, uniform diameter fiber
with lengths of several millimeters begin forming with
(ne soln ranging from 2.0 to 3.3 for all molecular weights,
which is lower than those of electrospinning. Since the
same amount of molecular entanglements can be achieved
by either high molecular weight species in low concentra-
tions or low molecular weight species in high concentra-
tions, isodiametric lines can be generated beyond a critical
chain entanglement concentration, Ce (Fig. 4). Polymer
solutions prepared in the semi-dilute, unentangled region
lead to beaded fibers or droplets (Fig. 4, Insert 1), and con-
centrated solutions with high entanglement numbers pro-
duce macro-scale fibers (Fig. 4, Insert 4). Long, smooth,
and uniform diameter fibers are obtained in the semi-dilute
entangled and low-moderate concentrated domains (Fig. 4,
Inserts 2 and 3). Fiber diameter is found to scale with
polymer concentration at the same molecular weight, and
molecular weight at the same concentration.34 36
2.1.2. Effect of Solvent Volatility
In addition to polymer molecular weight and solution con-
centration, solvent volatility is another factor influenc-
ing fiber diameters. Solvents with low boiling points lead
to rapid solvent loss and a shortened fiber solidification
process, thus limiting the stretching of the solution fila-
ment and producing larger diameter fibers. As shown in
Figure 5, spinning PEO solutions of the same molecular
weight and concentration results in increased fiber diame-
ters with increased ethanol content of the solvent mixture.
2.1.3. Fibers of Different Polymer Systems
By adjusting these parameters, a wide range of fiber diam-
eters can be obtained for multiple polymer systems, includ-
ing PS, PU, PEO and PMMA systems (Table I). Note that
the deviation of fiber diameter is well-controlled (within
20%) due to the absence of filament branching in the
Fig. 5. PEO design space: fiber diameter change as a function of solu-
tion concentration, ethanol ratio and molecular weight.

IP: 130.126.36.198 On: Tue, 27 Aug 2013 15:45:32
REVIEW
Table I. Fiber diameter spectrum for various polymer species spun using STEP at room temperature.
Polymer Parameter Small diameter Large diameter
Polystyrene Molecular weight (g·mol−1
0.86 M 2 M
(PS) Solution concentration (%wt) 6 14
Solvent p-Xylene THF
Diameter (nm) 34±4 8067±881
Polyurethane Molecular weight (g·mol−1
0.12 M 0.12 M
(PU) Solution concentration (%wt) 20 30
Solvent THF/DMF = 3/7, V:V THF/DMF = 8/2, V:V
Diameter (nm) 382±52 2610±231
Polyethylene Molecular weight (g·mol−1
0.4 M 0.9 M
(PEO) Solution concentration (%wt) 3 9
Solvent Ethanol/Water = 2:8, V:V Ethanol/Water = 8:2, V:V
Diameter (nm) 188±35 961±180
Poly methyl Molecular weight (g·mol−1
0.075 M 1 M
methacrylate Solution concentration (%wt) 20 16
(PMMA) Solvent Chlorobenzene Nitromethane
Diameter (nm) 643±147 1921±111
Note: M = million in molecular weight description.
fiber formation process. All fibers reported in Table I have
lengths of at least several millimeters.
2.1.4. Cell Response to Fiber Diameter
Cells attach to and interact with fibers differently than
with flat substrates. When attached to single fibers,
Fig. 6. A fluorescently labeled cell on a flat substrate, (B) a fluorescently labeled cell attached to a single STEP fiber in the spindle morphology,
(C) SEM image of a cell on a thick bundle of fibers slightly elongated along the fiber axes, (D) SEM image of a spindle cell on a thin bundle of
fibers, (E) nuclear circularity measurements (circularity = 4 A/P2
, A = nucleus area and P = nucleus perimeter) show increased elongation on fibers
(N = 479), and (F) migration speed is found to be a function of fiber diameter (N = 30). Cell speeds were measured for cells migrating near the
scaffold center, and scale bars represent 10 m. ∗
indicates statistical significance (p < 0 05).
cells spread along the fiber axes and take on a highly
elongated, spindle-like morphology characterized by quan-
tifiable changes in nucleus circularity (Figs. 6(A)–(E)).
Cells attached to STEP fibers are topographically con-
fined, and the degree of this confinement and subse-
quent migratory response are dependent on fiber diameter
(Fig. 6(F)).

IP: 130.126.36.198 On: Tue, 27 Aug 2013 15:45:32
REVIEW
2.2. Inter-Fiber Spacing
STEP manufacturing provides control of spacing between
adjacent fibers by adjusting the rotational velocity of the
substrate and the translational speed of the manipulator
holding the substrate (Fig. 2(A)). Biologically, controlling
inter-fiber spacing and alignment enables cells to either
spread between neighboring parallel fibers or remain on a
single fiber if the gap is too large (Fig. 7).
On suspended fibers, FAC cluster lengths are approxi-
mately four times longer than cells on flat PS (Fig. 7(D)).
This phenomenon likely occurs due to comparably lim-
ited available cell attachment locations compared to flat
surfaces. The cell likely compensates by forming multi-
ple adhesions within the same spatial confinement, which
when fluoresced appear as a longer adhesion cluster.
With the addition of common cytoskeletal knockdown
drugs (blebbistatin-myosin II, nocodazole-microtubules,
cytochalasin-D-actin), cells still display increased FAC
cluster lengths on fibers compared to flat (with the excep-
tion of cytochalasin-D in which no appreciable adhesions
are observed for either substrate). This demonstrates that
even in the absence of several key cytoskeletal compo-
nents, surface topography still influences cell attachment.
Recognizing that substrate design influences cell spread-
ing, a direct application of aligned fiber networks is in the
design and development of monolayer-based wound heal-
ing assays. Traditionally, wound healing assays or scratch
Fig. 7. Inter-fiber spacing effects, (A) at 10 um, C2C12s spread among multiple fibers (parallel cells). They maintain this shape up to about 15–
20 um gap sizes. At higher spacing, cells are unable to spread between fibers and attach to only one instead (spindle cells) (Supplementary Video 5),
(B) schematic representation of parallel and spindle cell shapes as well as their FAC length measurements showing typical locations, (C) FAC cluster
lengths for cells treated with pharmacological agents affecting cytoskeletal components, which show ∼ 4× increase on fibers compared to flat, and
(D) differences in migration speed.44
tests probe population-based migratory potential of a cell
line under a given physiochemical state. Such tests are usu-
ally conducted by making an incision to a cell monolayer
on a flat substrate and observing the closure dynamics.45
In contrast, STEP suspended parallel fibers enable the
investigation of closure dynamics, yet cells are interact-
ing along topographic cues which they would more likely
encounter in vivo.46
Closure in our wound healing assay occurs in two prin-
cipal steps: axial migration (where NIH-3T3 fibroblasts
emerge and migrate along the fiber), and gap closure
(where cells fill in the space between the fibers). Over a
week, the cells will migrate and eventually cover the fibers
(Fig. 8). We envision that this STEP based fundamental
study of single cells spreading and migrating along aligned
and suspended fibers of different diameters deposited at
controlled spacing would aid in the development of wound
healing sutures as a means of facilitating the migration of
healthy cells into wounded damaged tissue.
2.3. Fiber Orientation
In addition to creating substrates with a single layer of
fibers parallel to one another, the angle between subse-
quent layers of fibers can be controlled. Here, we illustrate
that increments of 30 have an effect on cell spreading
behavior (Fig. 9).

IP: 130.126.36.198 On: Tue, 27 Aug 2013 15:45:32
REVIEW
Fig. 8. (A) STEP based wound healing assay which demonstrates gap closure over time, (B) cell migration along the aligned fibers after one day,
and (C) wound closure after one week. The gaps in the closure model are dependent upon inter-fiber spacing. Scale bar represents 500 m.
C2C12 cells seeded on these scaffolds typically
form stable configurations at the fiber intersections
(Supplementary Video 6). Those that do not form stable
configurations start migrating along diverging fibers. The
increasing gap size that the cell encounters as it attempts
to remain attached to both diverging fibers causes the
cell to eventually detach from one and assume a spin-
dle shape, where presumably less energy is required to
maintain this cell configuration (Supplementary Video 7).
This system enables us to visualize F-actin stress fiber
location and relate it to nucleus shape and FAC forma-
tion, potentially allowing us to determine if cells mod-
ulate their elasticity in accordance with the angle of
the fibers. Increased stress fiber presence is commonly
associated with increased cell contractility and therefore
decreased migration speed.47 48
Interestingly, we find that
Fig. 9. (A)–(D) Angular control of STEP fibers, (E)–(H) corresponding changes in cytoskeletal arrangement of C2C12 cells, and (I)–(L) Schematics
of cytoskeletal arrangements on the angled substrates highlighting orientation of stress fibers (dark red) and typical cell spread area measurements.
Scale bars represent 20 m.
cell migration speed is dependent on fiber orientation, with
spindle cells able to migrate the fastest (∼ 50 m/hr) and
polygonals migrating the slowest (∼ 20 m/hr). Spindle
cells only form two main clusters which are oriented along
the same axis, whereas parallel cells contain four along
the same axis and polygonal cells have four oppositely-
oriented adhesion clusters on two different axes. Even
though FAC cluster lengths are comparable on fibers, it is
their orientation relative to one another which influences
maximum migration speed.
2.4. Multi-Layer Assemblies and
Hierarchical Structures
Arrays of fibers arranged in hierarchical structures begin
to resemble the complex ECM fibrous arrangement. Such

IP: 130.126.36.198 On: Tue, 27 Aug 2013 15:45:32
REVIEW
hierarchical assemblies can then be employed to study and
understand how cellular migration in the presence of topo-
graphical constraints might be occurring in vivo. We have
extended the STEP platform to study migration dynam-
ics and blebbing phenomena of glioblastoma multiforme
(GBM) on aligned multi-layer fiber networks. GBM is the
most invasive form of brain cancer, causing over 15,000
annual deaths.49
It is believed that these cancerous glioma
cells prefer to migrate along aligned structures called white
matter tracts within the central nervous system, which
range from sub-500 nm to 3 m in diameter.50
Aligned
STEP fibers therefore provide a unique environment to
study glioma cell behavior. Using parallel, aligned, and sus-
pended 400 nm diameter PS STEP fibers, we observed that
the glioma cells (Denver Based Tumor Research Group,
DBTRG-05MG line) migrate almost three times faster
(range: 15–200 m/hr, average: 70 m/hr) on fibers when
compared to flat substrates (range: 5–50 m/hr, average:
25 m/hr). Furthermore, blebbing, which is associated with
cell migration and resistance to drugs/lysis, is observed
to be affected by suspended STEP fiber networks. We
observed a reversible blebbing non-blebbing phenomenon
in DBTRG cells where the bleb size and number produced
by the cells depended on the cell spread area. As the cells
acquired a spread configuration on suspended fibers, both
bleb size and number decreased. A linear regression analy-
sis showed that blebs almost completely disappeared when
cells spread beyond an area of 1400 m2
(Fig. 10).51
Given the known relationships of blebbing with amoe-
boid form of cancer migration and resistance to lysis
and drugs, our results suggest that glioma cells under
spread configurations could be relatively more vulnera-
ble to drugs, and would be migrating using non-amoeboid
migration modes. This platform may also be used to max-
imize the migration of individual cancer cells to present a
‘worst-case scenario’ that can ultimately be used as a drug
testing platform.
Fig. 10. (A) Increased cell spreading causes a decrease in bleb occurrence (count). (B) Increased cell spreading reduces bleb size, insert (i) demon-
strates a blebbing, rounded cell, and (ii) later, the same cell stretches and no longer blebs. Linear analysis of the data shows a disappearance of blebs
as the cells spread beyond an area of 1400 m2
. Scale bar represents 20 m.
2.5. Role of Structural Stiffness
It has been shown on many different substrate types
including micropillars, gels, and glass, that material-
dependent substrate stiffness (E, measured as N/m2
directly alters cellular behaviors of migration, differenti-
ation, and apoptosis.52–54
In comparison, suspended fibers
are essentially one-dimensional beams of uniform mate-
rial stiffness with varying structural stiffness (k, mea-
sured as N/m) along the length. Cells on a flat surface
of constant material stiffness and protein coating behave
similarly at all locations, but cells on a suspended fiber
are found to respond differently along the length of the
fiber due to changes in structural stiffness along its axis.
In such a system, the structural stiffness scales with fiber
diameter and length (∼ diameter4
/length3
and we specu-
late that cells attached to these fibers in spindle shapes
apply migratory forces through FACs located at the poles.
By interacting mechanically with a single fiber, the cell
probes substrate structural stiffness and FACs are found to
mature accordingly, which in turn affects behaviors such
as cell spreading, migration, and cytoskeletal arrangement
(Fig. 11).
Cells attached to STEP suspended fibers with simply
supported ends clearly respond to the mechanical gra-
dient of changes in structural stiffness by conforming
their cytoskeletons and adjusting migration speed. C2C12
mouse myoblasts tend to migrate quicker at the center of
suspended nanofibers and decrease their migration speed
as they reach the higher stiffness fiber ends. Additionally,
as cells reach areas of lower structural stiffness at the mid-
dle of the fiber span length (lowest k) compared to higher
structural stiffness at the edges (highest k), they demon-
strate shorter cell length, shorter FAC clusters, and lower
nuclear shape index (Fig. 11). The inverse relationship
between fiber diameter and fiber length corresponding to
the parameters which play dominant roles in the structural
stiffness can be extended to add to our understanding of
cancer cell metastasis mechanisms.55

IP: 130.126.36.198 On: Tue, 27 Aug 2013 15:45:32
REVIEW
Fig. 11. (A) Schematic representation of cell positioning on a single fiber suspended across two micropillars, (B) SEM image demonstrating cell
spreading’s dependence on fiber positioning, and (C) gradient structural stiffness along the length of a single 1 mm length PS nanofiber (700 nm
diameter) causes a significant difference in migration speed (normalized to 44 m/hr) (N = 293), nuclear shape index (normalized to 0.7) (N = 69),
and cell length (normalized to 100 m) (N = 62). ∗
indicates statistical significance (p < 0 05).
3. POPULATION CELL BEHAVIOR AND
APPLICATIONS IN TISSUE
ENGINEERING
In addition to demonstrating their use in single-cell stud-
ies, STEP fibers have shown promise in multiple tissue
engineering applications, highlighting the usefulness of
providing cells with physical environments that more
Fig. 12. (A) Effect of growth factor concentration on tendon differentiation,55
(B) sheet engineering (white dotted lines show underlying fiber
directions), (C) long-term culture of hepatocyte monolayers, and (D) early neural differentiation demonstrated by MAP2 fluorescence on single and
double layer scaffolds shown by dotted lines.57
closely represent the ECM (Fig. 12). For instance, with
the use of STEP scaffolds we have shown monolayers
of C2C12 cells subject to picogram/mm2
concentrations
of fibroblast growth factor 2 (FGF-2) to differentiate into
myotubes (Fig. 12 (A)).56
SEM images of monolayer
formation reveals that cell populations remain in align-
ment with their underlying fibers (Fig. 12(B)), suggesting
applications in sheet engineering.57
In addition, the fibers

IP: 130.126.36.198 On: Tue, 27 Aug 2013 15:45:32
REVIEW
have the ability to maintain long term functional behav-
ior of hepatocytes without de-differentiating (Fig. 12(C)
and unpublished data), and the same fibers have the ability
to achieve neuronal differentiation (80% neurons) from a
population of neural stem cells (NSC’s) (Fig. 12(D)).58
4. CONCLUDING REMARKS AND
FUTURE DIRECTIONS
Given the vast design space available for manufacturing
in vitro scaffolds, the ability to control key parameters in
fibrous scaffold production has opened up new possibilities
in studying cell-fiber interactions. Since they originate and
operate in different physiochemical niches, every cell type
brings with it a unique set of biophysical and biochemical
challenges which must be accounted for during substrate
design. Here, we have shown through the manipulation
of key components in fibrous substrate design that cells
exhibit diverse behavior on suspended fibers compared to
more traditionally-studied flat substrates, and that these
differences carry great potential in uncovering mechanisms
of cell adhesion and eventual force modulation. Through
the ability to control fiber diameter and length, we have
shown that cells sense and respond to changes in struc-
tural stiffness by increasing spreading on regions of higher
stiffness leading to lower migration. Fiber spacing control
allows scaffold optimization for the development of a new,
fully suspended wound healing assay. Oriented fiber arrays
in single and multiple layers at specific angles forces
cells to spread into different shapes, from highly elon-
gated spindles to more evenly-spread polygonals, which
helps us understand stress fiber organization and eventual
elasticity of cells. Fiber curvature and geometry impart
behaviors into cells which flat, two dimensional substrates
with and without topographic features find challenging to
recapitulate.
The knowledge gained from these experiments coupled
with the ability to build more accurate in vitro scaffolds
will allow the measurement of migratory cell force changes
in real-time to be related to scaffold mechanical properties.
In the future, this will allow coupling of chemical growth
factors with mechanical cues to develop a unique, dual-
cue gradient environment to advance diagnostic and drug
testing platforms and scaled up scaffold designs to accom-
modate tissue engineering on a larger scale.
5. EXPERIMENTAL DETAILS
5.1. Fiber Manufacturing and Pillar Design
Fibers were deposited according to STEP manufacturing
principles outlined above and reported previously.34
Pil-
lars for structural stiffness measurements were punched
out of a block of polydimethylsiloxane (PDMS) of 0.635
cm thickness using a 1.0 mm diameter Harris Uni-Core
Sample punch. Pairs of these pillars were then placed sev-
eral mm apart on a glass slide and fixed using an epoxy
adhesive. Fibers were spun over the pillars using the STEP
method, with a drop of uncured PDMS applied to the tip
of each pillar before spinning to ensure proper anchorage.
5.2. Cell Culture and Imaging
C2C12, NIH-3T3, and DBTRG cell lines were cultured
following suggested protocols from ATCC. SEM images
were taken with the FEI Quanta 600 FEG ESEM in high
vacuum mode. Phase contrast microscopy images were
taken with the Zeiss AxioObserver Z1 with incubation
and digital stage. Fluorescence imaging was conducted via
direct and antibody staining. To visualize FACs, cells were
fixed in 4% paraformaldehyde and permeabilized in Triton
X100. Polyclonal rabbit anti-paxillin (Invitrogen) primary
antibodies diluted 1:250 were used in conjunction with
Alexa Fluor 488 (Invitrogen) secondary antibodies. F-actin
was stained using 1:100 dilution of rhodamine phalloidin
(Santa Cruz). Nuclei were counterstained with 300 nM
4 ,6-diamidino-2-phenylindole (DAPI).
5.3. Statistical Analysis
Statistical comparisons were made using student’s t-test in
JMP-9 software. Populations were considered statistically
significant when p < 0 05 unless otherwise noted. Error
bars in figures denote one standard deviation.
Acknowledgments: The authors would like to acknow-
ledge support received from the Institute for Critical Tech-
nology and Applied Science (ICTAS) at Virginia Tech.
This research was funded in part by Jeffress Memorial
Trust Fund and the Bill and Andrea Waide Research Fund.
References and Notes
1. T. R. Cox and J. T. Erler, Remodeling and homeostasis of the extra-
cellular matrix: Implications for fibrotic diseases and cancer. Disease
Models and Mechanisms 4, 165 (2011).
2. P. P. Provenzano and R. Vanderby, Collagen fibril morphology and
organization: Implications for force transmission in ligament and ten-
don. Matrix Biology: Journal of the International Society for Matrix
Biology. 25, 71 (2006).
3. J. C. Patterson-Kane, D. L. Becker, and T. Rich, The pathogenesis
of tendon microdamage in athletes: The horse as a natural model for
basic cellular research. Journal of Comparative Pathology 147, 227
(2012).
4. S. D. Subramony, B. R. Dargis, M. Castillo, E. U. Azeloglu, M. S.
Tracey, A. Su, and H. H. Lu, The guidance of stem cell differenti-
ation by substrate alignment and mechanical stimulation. Biomater.
34, 1942 (2013).
5. T. Rozario and D. W. DeSimone, The extracellular matrix in devel-
opment and morphogenesis: A dynamic view. Developmental Biol-
ogy 341, 126 (2010).
6. B. Wehrle-Haller, Structure and function of focal adhesions. Current
Opinion in Cell Biology 24, 116 (2011).

IP: 130.126.36.198 On: Tue, 27 Aug 2013 15:45:32
REVIEW
7. K. A. Kilian, B. Bugarija, B. T. Lahn, and M. Mrksich, Geometric
cues for directing the differentiation of mesenchymal stem cells. Pro-
ceedings of the National Academy of Sciences of the United States
of America, March (2010), Vol. 107, pp. 4872–7.
8. R. S. Reneman and A. P. G. Hoeks, Wall shear stress as measured
in vivo: Consequences for the design of the arterial system. Medical
and Biological Engineering and Computing 46, 499 (2008).
9. N. J. B. Driessen, G. W. M. Peters, J. M. Huyghe, C. V. C. Bouten,
and F. P. T. Baaijens, Remodelling of continuously distributed col-
lagen fibres in soft connective tissues. Journal of Biomechanics
36, 1151 (2003).
10. A. Yegorov, L. Kakurin, and Y. Nefyodov, Effects of an 18-day flight
on the human body. Life Sciences and Space Research 57 (1972).
11. S. M. Smith, M. E. Wastney, B. V Morukov, I. M. Larina,
E. Laurence, S. A. Abrams, E. N. Taran, C. Shih, J. L. Nillen,
E. Janis, B. L. Rice, and H. W. Lane, Calcium metabolism before,
during, and after a 3-mo spaceflight: Kinetic and biochemical
changes. Am. J. Physiol. Regulatory, Integrative and Comp. Physiol.
277, R1 (1999).
12. W. Neuman, Calcium metabolism in space flight. Life Sci. Space
Res. 8, 309 (1970).
13. C. A. Simmons, S. Matlis, A. J. Thornton, S. Chen, C.-Y. Wang,
and D. J. Mooney, Cyclic strain enhances matrix mineralization by
adult human mesenchymal stem cells via the extracellular signal-
regulated kinase (ERK1/2) signaling pathway. Journal of Biome-
chanics 36, 1087 (2003).
14. M. Schmelter, B. Ateghang, S. Helmig, M. Wartenberg, and
H. Sauer, Embryonic stem cells utilize reactive oxygen species as
transducers of mechanical strain-induced cardiovascular differentia-
tion. FASEB Journal: Official Publication of the Federation of Amer-
ican Societies for Experimental Biology 20, 1182 (2006).
15. H. Yamaguchi, J. Wyckoff, and J. Condeelis, Cell migration in
tumors. Current Opinion in Cell Biology 17, 559 (2005).
16. M. J. Buehler, Nature designs tough collagen: Explaining the nano-
structure of collagen fibrils, Proceedings of the National Academy of
Sciences of the United States of America August (2006), Vol. 103,
pp. 12285–90.
17. B. M. Baker and R. L. Mauck, The effect of nanofiber align-
ment on the maturation of engineered meniscus constructs. Biomater.
28, 1967 (2007).
18. C. Erisken, X. Zhang, K. L. Moffat, W. N. Levine, and H. H. Lu,
Scaffold fiber diameter regulates human tendon fibroblast growth and
differentiation. Tissue Engineering. Part A 19, 519 (2013).
19. Y. Liu, Y. Ji, K. Ghosh, R. a F. Clark, L. Huang, and M. H.
Rafailovich, Effects of fiber orientation and diameter on the behav-
ior of human dermal fibroblasts on electrospun PMMA scaffolds.
Journal of Biomedical Materials Research. Part A 90, 1092 (2009).
20. A. M. Kloxin, K. J. R. Lewis, C. a Deforest, G. Seedorf, M. W.
Tibbitt, V. Balasubramaniam, and K. S. Anseth, Responsive culture
platform to examine the influence of microenvironmental geometry
on cell function in 3D. Integrative Biology 4, 1540 (2012).
21. M. E. Berginski, E. a Vitriol, K. M. Hahn, and S. M. Gomez, High-
resolution quantification of focal adhesion spatiotemporal dynamics
in living cells. PloS One 6, e22025 (2011).
22. R. A. Neal, A. Jean, H. Park, P. B. Wu, J. Hsiao, G. C. Engelmayr,
R. Langer, and L. E. Freed, Three-dimensional elastomeric scaffolds
designed with cardiac-mimetic structural and mechanical features.
Tissue Engineering. Part A 19, 793 (2013).
23. M. Nikkhah, F. Edalat, S. Manoucheri, and A. Khademhosseini,
Engineering microscale topographies to control the cell-substrate
interface. Biomater. 33, 5230 (2012).
24. G. V Shivashankar, Mechanosignaling to the cell nucleus and gene
regulation. Annual Review of Biophysics 40, 361 (2011).
25. Y. Liu, A. Franco, L. Huang, D. Gersappe, R. a F. Clark, and M. H.
Rafailovich, Control of cell migration in two and three dimensions
using substrate morphology. Experimental Cell Research 315, 2544
(2009).
26. W. Tutak, S. Sarkar, S. Lin-Gibson, T. M. Farooque, G. Jyotsnendu,
D. Wang, J. Kohn, D. Bolikal, and C. G. Simon, The support of
bone marrow stromal cell differentiation by airbrushed nanofiber
scaffolds. Biomater. 34, 2389 (2013).
27. G. Vidal, B. Delord, W. Neri, S. Gounel, O. Roubeau,
C. Bartholome, I. Ly, P. Poulin, C. Labrugère, E. Sellier, M.-C. Dur-
rieu, J. Amédée, and J.-P. Salvetat, The effect of surface energy,
adsorbed RGD peptides and fibronectin on the attachment and
spreading of cells on multiwalled carbon nanotube papers. Carbon
49, 2318 (2011).
28. T. J. Sill and H. a von Recum, Electrospinning: Applications in drug
delivery and tissue engineering. Biomater. 29, 1989 (2008).
29. G. Chang, G. Song, J. Yang, R. Huang, A. Kozinda, and J. Shen,
Morphology control of nanohelix by electrospinning. Appl. Phys.
Lett. 101, 263503 (2012).
30. B. Sundaray, V. Subramanian, T. S. Natarajan, R.-Z. Xiang, C.-C.
Chang, and W.-S. Fann, Electrospinning of continuous aligned poly-
mer fibers. Appl. Phys. Lett. 84, 1222 (2004).
31. W. Han Bing, E. M. Michael, M. C. Jared, H. Andres, O. Martin,
T. T. Matthew, and J. G. Ryan, Creation of highly aligned electro-
spun poly-L-lactic acid fibers for nerve regeneration applications.
Journal of Neural Engineering 6, 16001 (2009).
32. Z.-M. Huang, Y. Z. Zhang, M. Kotaki, and S. Ramakrishna, A review
on polymer nanofibers by electrospinning and their applications
in nanocomposites. Composites Science and Technology 63, 2223
(2003).
33. J. A. Matthews, G. E. Wnek, D. G. Simpson, and G. L. Bowlin,
Electrospinning of collagen nanofibers. Biomacromolecules 3, 232
(2002).
34. A. S. Nain, M. Sitti, A. Jacobson, T. Kowalewski, and C. Amon,
Dry spinning based spinneret based tunable engineered parameters
(STEP) technique for controlled and aligned deposition of poly-
meric nanofibers. Macromolecular Rapid Communications 30, 1406
(2009).
35. A. S. Nain, J. C. Wong, C. Amon, and M. Sitti, Drawing suspended
polymer micro-/nanofibers using glass micropipettes. Appl. Phys.
Lett. 89, 183105 (2006).
36. A. S. Nain and J. Wang, Polymeric nanofibers: Isodiametric design
space and methodology for depositing aligned nanofiber arrays in
single and multiple layers. Polym. J (2013).
37. F. Family and P. Meakin, Scaling of the droplet-size distribution in
vapor-deposited thin films. Phys. Rev. Lett. 61, 428 (1988).
38. L. Wang, C.-L. Pai, M. C. Boyce, and G. C. Rutledge, Wrinkled
surface topographies of electrospun polymer fibers. Appl. Phys. Lett.
94, 151913 (2009).
39. C.-L. Pai, M. C. Boyce, and G. C. Rutledge, Morphology of porous
and wrinkled fibers of polystyrene electrospun from dimethylfor-
mamide. Macromolecules 42, 2102 (2009).
40. J. Wang, J. H, M. Ellis, and A. S. Nain, Organized long titanium
dioxide nanofibers/nanotubes with controlled morphology using a
sol–gel combined STEP technique. New J. Chem. 37, 571 (2013).
41. Y. Yang, J. Wang, J. Li, D. Viehland, and A. S. Nain, Nanoparti-
cles deposition at specific sites using aligned fiber networks. OJINM
2, 55 (2012).
42. S. L. Shenoy, W. D. Bates, H. L. Frisch, and G. E. Wnek, Role
of chain entanglements on fiber formation during electrospinning
of polymer solutions: Good solvent, non-specific polymer-polymer
interaction limit. Polymer 46, 3372 (2005).
43. D. W. Mead, R. G. Larson, and M. Doi, A molecular theory for fast
flows of entangled polymers. Macromolecules 31, 7895 (1998).
44. K. Sheets, S. Wunsch, C. Ng, and A. S. Nain, Shape-dependent cell
migration and focal adhesion organization on suspended and aligned
nanofiber scaffolds. Acta Biomaterialia 9, 7169 (2013).
45. J. C. Yarrow, Z. E. Perlman, N. J. Westwood, and T. J. Mitchison,
A high-throughput cell migration assay using scratch wound healing,
a comparison of image-based readout methods. BMC Biotechnology
4, 21 (2004).

IP: 130.126.36.198 On: Tue, 27 Aug 2013 15:45:32
REVIEW
46. T. Dvir, B. P. Timko, D. S. Kohane, and R. Langer, Complex tissues.
Nature Publishing Group 6, 13 (2010).
47. D. A. Lauffenburger and a F. Horwitz, Cell migration: A physically
integrated molecular process. Cell 84, 359 (1996).
48. M. L. Gardel, I. C. Schneider, Y. Aratyn-Schaus, and C. M.
Waterman, Mechanical integration of actin and adhesion dynamics
in cell migration. Annual Review of Cell and Developmental Biology
26, 315 (2010).
49. T. A. Ulrich, E. M. de Juan Pardo, and S. Kumar, The mechanical
rigidity of the extracellular matrix regulates the structure, motility,
and proliferation of glioma cells. Cancer Research 69, 4167 (2009).
50. D. N. Louis, Molecular pathology of malignant gliomas. Annu. Rev.
Pathol. 1, 97 (2006).
51. P. Sharma, K. Sheets, E. Subbiah, and A. S. Nain, The mechanistic
influence of aligned nanofiber networks on cell shape, migration and
blebbing dynamics of glioma cells. Integr. Biol. In Press (2013).
52. A. J. Engler, S. Sen, H. L. Sweeney, and D. E. Discher, Matrix
elasticity directs stem cell lineage specification. Cell 126, 677
(2006).
53. R. H. W. Lam, S. Weng, W. Lu, and J. Fu, Live-cell subcellular
measurement of cell stiffness using a microengineered stretchable
micropost array membrane. Integrative Biology 4, 1289 (2012).
54. S. Even-Ram and K. M. Yamada, Cell migration in 3D matrix. Cur-
rent Opinion in Cell Biology 17, 524 (2005).
55. D. Wirtz, K. Konstantopoulos, and P. C. Searson, The physics of
cancer: The role of physical interactions and mechanical forces in
metastasis. Nat. Rev. Cancer 11, 512 (2011).
56. E. D. Ker, A. S. Nain, L. E. Weiss, J. Wang, J. Suhan, C. H. Amon,
and P. G. Campbell, Bioprinting of growth factors onto aligned sub-
micron fibrous scaffolds for simultaneous control of cell differentia-
tion and alignment. Biomater. 32, 8097 (2011).
57. M. Yamato, Nanotechnology-based cell sheet engineering for regen-
erative medicine, Micro-NanoMechatronics and Human Science,
2005 IEEE International Symposium (2005), pp. 251–254.
58. S. Bakhru, A. S. Nain, C. Highley, J. Wang, P. Campbell, C. Amon,
and S. Zappe, Direct and cell signaling-based, geometry-induced
neuronal differentiation of neural stem cells. Integrative Biology
3, 1207 (2011).
59. V. Chaurey, P.-C. Chiang, C. Polanco, Y.-H. Su, C.-F. Chou, and
N. S. Swami, Interplay of electrical forces for alignment of sub-
100 nm electrospun nanofibers on insulator gap collectors. Langmuir
26, 19022 (2010).
60. D. Li, Y. Wang, and Y. Xia, Electrospinning nanofibers as uniaxially
aligned arrays and layer-by-layer stacked films. Advanced Materials
16, 361 (2004).
61. Y. Liu, X. Zhang, Y. Xia, and H. Yang, Magnetic-field-assisted elec-
trospinning of aligned straight and wavy polymeric nanofibers. Adv.
Mater. 22, 2454 (2010).
62. A. Jafari, J.-H. Jeon, and I.-K. Oh, Well-aligned nano-fiberous mem-
branes based on three-pole electrospinning with channel electrode.
Macromol. Rapid Commun. 32, 921 (2011).
63. S. Sarkar, S. Deevi, and G. Tepper, Biased ac electrospinning of
aligned polymer nanofibers. Macromol. Rapid Commun. 28, 1034
(2007).
Received: 10 March 2013. Accepted: 24 April 2013.

Cell-Fiber Interactions on Aligned and Suspended Nanofiber Scaffolds

Recommended

Recommended

More Related Content

What's hot

What's hot (20)

Similar to Cell-Fiber Interactions on Aligned and Suspended Nanofiber Scaffolds

Similar to Cell-Fiber Interactions on Aligned and Suspended Nanofiber Scaffolds (20)

Recently uploaded

Recently uploaded (20)

Cell-Fiber Interactions on Aligned and Suspended Nanofiber Scaffolds