Exploring combined influences of material topography, stiffness and chemistry on cell
behavior at biointerfaces
Zhou, Qihui
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Exploring Combined Influences of
Material Topography, Stiffness and
Chemistry on Cell Behavior at
Biointerfaces
Exploring Combined Influences of Material Topography, Stiffness and Chemistry on Cell Behavior at Biointerfaces By Qihui Zhou
University Medical Center Groningen, University of Groningen Groningen, The Netherlands
Copyright © 2018 by Qihui Zhou Cover designed by Qihui Zhou
Printed by Offpage, Amsterdam, The Netherlands ISBN (printed version): 978-94-034-0780-7 ISBN (electronic version): 978-94-034-0779-1
Exploring Combined Influences of Material
Topography, Stiffness and Chemistry on Cell
Behavior at Biointerfaces
Proefschrift
ter verkrijging van de graad van doctor aan de Rijksuniversiteit Groningen
op gezag van de
rector magnificus prof. dr. E. Sterken en volgens besluit van het College voor Promoties.
De openbare verdediging zal plaatsvinden op
woensdag 11 juli 2018 om 12.45 uur
door
Qihui Zhou
geboren op 01 februari 1988 te Shandong, China
Promotor
Prof. dr. ir. H.J. Busscher
Copromoter
Dr. P. van Rijn
Beoordelingscommissie
Prof. dr. P.Y.W. Dankers Prof. dr. R.A. Bank
Table of Contents
Chapter 1 General Introduction 1
Aim of the Thesis 9
Outline of the Thesis 10
Chapter 2 Mechanical Properties of Aligned Nanotopographies for
Directing Cellular Behavior 15
Chapter 3 Directing Mesenchymal Stem Cells with Gold Nanowire Arrays 37
Chapter 4
Directional Nanotopographic Gradients: a High-throughput
Screening Platform for Cell Contact Guidance 55
Chapter 5
Screening Platform for Cell Contact Guidance Based on
Inorganic Biomaterial Micro/nanotopographical Gradients 75
Chapter 6
Orthogonal Double Gradient for Determining Combined Influences of Stiffness and Wettability on Mesenchymal Stem
Cell Behavior 101
Chapter 7 General Discussion 119
Summary 124
Samenvatting 130
Acknowledgements 136
C
HAPTER
O
NE
Faced with an ever-increasing burden of disease, congenital abnormalities and accidents each year, tissue engineering and regenerative medicine (TERM) hold great promise to repair or replace tissues or even entire organs on demand for a better quality of life [1,2].
Undoubtedly, biomaterials play an increasingly pivotal role in the development of biomedical devices and tissue engineering scaffolds [3], which seeks to unlock the
mechanism underlying cell-material interactions named as materiobiology [4]. Harnessing
the cell-material interactions to trigger the tissue regenerative potential to ensure normal tissue formation and maturation during healing is still a huge challenge. For that reason, numerous research groups have been trying to design and develop the fourth generation of biomaterials with smart or biomimetic features (Figure 1) based on recent discoveries about the complex cellular microenvironment and its interaction with cells
[5–7].
Figure 1. Evolution of biomaterials. Materials that originally were meant not to interact
with surrounding biological systems (inert) to materials that not only interact but also determine the fate of surrounding tissues and cells.
1.1.
THE
CELLULAR
MICROENVIRONMENT
In vivo, cells are known to reside in a highly dynamic and extremely complicated system surrounded by multiple biochemical and biophysical signals, defined as “cell microenvironment”, that significantly mediates cell behavior and fate [8]. Particularly,
stem cells localize within a specialized microenvironment, termed “niche” [9–11], a
concept that was originally formulated by Schofield in 1978 to describe the hematopoietic stem cell microenvironment [12]. The stem-cell niche provides the specific
physicochemical features to renew themselves and govern their survival, stemness (maintenance of their pluripotency), and the phenotype of differentiation for tissue repair and regeneration [2,10]. In general, there are the four key components within the
cell microenvironment (Figure 2), including neighboring cells, soluble factors, biophysical stimuli and the surrounding extracellular matrix (ECM), which synergistically influence cellular responses, e.g., adhesion, proliferation, migration, self-renewal, differentiation, and so on [8]. Herein, neighboring cells, soluble factors and
biophysical stimuli are briefly introduced together and then the surrounding ECM is highlighted in this section.
Figure 2. Schematic illustration of the key components of cellular microenvironment.
Adapted with permission from ref. [4]. Copyright 2017 American Chemical Society.
1.1.1. NEIGHBORING CELLS, SOLUBLE FACTORS AND BIOPHYSICAL STIMULI
Cells in vivo do not reside in isolation but rather communicate with both similar and diverse kinds of cells. Cell-cell interactions play a critical role in cell function and tissue development. Generally, cell junctions (e.g., tight junctions, anchoring junctions, and gap junctions) are induced by specific adhesion proteins, such as cadherins or related proteins (e.g., desmogleins and desmocollins) or ions (e.g., Ca2+) [8,13]. Many studies have
demonstrated the pivotal role of direct cell-cell interactions in governing cell responses and tissue development [14]. Recently, owing to integrated organ-on-a-chip or
microfluidic co-culture platforms that were developed specifically to detect these interactions, the direct cell-cell communication and interaction are relatively well understood [15,16]. The features of microfluidic systems include high-throughput,
miniaturization, high resolution, and integration, which can mimic complex microenvironment and simplify their complexity for studying different cell types, such as adult cells, stem cells, immune cells, and cancer cells [17,18].
In addition to direct cell-cell contact, indirect interaction between cells mediated by soluble factors also plays a key role in cell behavior and tissue function. In vivo, there are many soluble factors within the cellular microenvironment, including diverse nutrients (e.g., glucose, glutamine and amino acids) and signaling factors (e.g., growth factors, mitogen, hormones and cytokines) [19,20]. Among soluble factors, growth factors are the
most important and widely investigated cues for constructing the functional and biomimetic microenvironment [21]. Generally, these growth factors dissolving in aqueous
media or immobilized on the ECM are dynamic in space and time and in a concentration gradient. The specific growth factors, local concentration, and their time-dependent and spatial distribution play key roles in controlling different cell responses. Therefore, the controlled release of growth factors on demand in the cell microenvironment is an important frontier.
Besides the cell-cell interactions described above, cells in vivo are often stimulated by numerous physical parameters, such as stress, strain, and thermal fields [8]. Particularly
for the mechanical properties, cells may experience various stress and strain stimuli depending on their locations. They can sense and transduce mechanical cues into intercellular signals that modulate their cytoskeleton and respond through a process known as mechanosensing and mechanotransduction [22,23]. In general, the mechanical
stimulus needs the mediation of the ECM to work on cells.
1.1.2. THE ECM
In human tissues and organs, every cell is exposed to an intricate 3D network of fibers named the ECM which is composed of proteins (e.g., collagens, elastin, laminin, and fibronectin) and glycosaminoglycans (e.g., hyaluronic acid) [24]. These interlocked ECM
macromolecules are secreted and assembled by the resident cells of each tissue. As a result, the biochemical composition and specific physical properties of ECM are distinctive between tissues [25]. Interestingly, ECM exists in a dynamic state due to their
neighboring cells and biophysical stimulus, which can mediate cell behavior dynamically and reciprocally [26]. Interactions between cells and their ECM have been extensively
studied over the past few decades, yielding specifics on binding partners, motifs, and strengths [27]. Previously, the ECM has been considered as a passive supporting
substrate in which the resident cells were regarded as the major actors. Recently, the ECM is undoubtedly increasingly recognized as a bioactive structure, which offers structural, mechanical, and compositional signals that can direct cell activities and functions during the natural tissue regeneration process [4,8]. The ECM is well-known to
be essential for the physiology of living cells and tissue functions, while the aberrant ECM can induce unexpected cell behaviors and then result in some diseases (e.g., cancer, fibrosis) [28]. Therefore, mastering the communication between cells and ECM is
one of the most important tasks in order to mimic the structural and biologic features of the native ECM within biomimetic materials for instructing cells with specific cues. Although the physicochemical properties of the ECM can be changed over time and in space, understanding their basic features will be beneficial to the design of desired functional and biomimetic materials. The effect of biochemical cues on cells has long been recognized, but the importance of the biophysical features from the ECM were neglected. Recently, it was increasingly appreciated that the ECM offers many specific cues to cells including the structural and dimensional presentation, the mechanical stiffness, and the spatiotemporal variations. Most tissue extracellular matrixes (e.g., bone, tendon, nerve, myocardium, etc.) have hierarchically organized and anisotropic structures consisting of well-aligned micro/nano-scaled fiber [29–31]. The ECM fiber
orientation not only induces the orientation/migration of many cell types, but also directs cell function and stem cell differentiation [29,32,33]. Another important structural
porosity and pore size of the ECM mesh are crucial factors for determining the available space for cell penetration and exchange of nutrients. In addition, ultrastructural analysis of the native ECM fibers tend to have feature diameters from a few nanometers to ∼150 nm and actual fiber bundles have feature diameters from several hundred nanometers to ∼400 µm, depending on the tissue type [35–37].
The ECM biomechanical properties named as stiffness (elastic or Young's modulus) vary significantly between tissues and organs, and may play a critical role in tissue homeostasis and function [38]. It is reported that native tissues have mechanical features
spanning orders of magnitude, from static or compliant (soft) in brain or lung tissue, to hard bone tissue owing to mineralized fibers [8,39]. Cells can sense and respond to
different stiffness parameters through mechanosensing and mechanotransduction [40,41].
Importantly, changes of tissue stiffness are often considered a representative prognostic factor for diseases [42]. For instance, tumorigenesis and pathological fibrosis are closely
related with an increase in matrix stiffening, because abnormal mechanical changes in ECM enhance tumor cell invasion and myofibroblast differentiation [38,43]. Taken
together, the ECM biophysical properties can have a great impact on basic cell responses and tissue developmental processes.
1.2.
CELL
AND
BIOMATERIAL
INTERACTION
Biomaterials e.g. implants or scaffolds similar to native ECM macro-environment have been used widely to repair or engineer tissues, which are biocompatible, enable nutrient transport, and offer structural integrity [6,44]. In order to maximize the full potential of
cell-based therapies in TERM, it is vital to engineer and understand artificial ECM signals that can regulate cell behavior. Recently, inspired by the complex cellular microenvironment mentioned above, researchers found that cells are inherently sensitive to surrounding static/dynamic microenvironment of biomaterials, especially physical cues (e.g., mechanical properties, wettability, 2D topography, and 3D geometry), substrate-immobilized insoluble/soluble biochemical signals (e.g., material component, fixed proteins or diffusible factors), or other stimuli [2,4,8]. It is
well-documented that these biomaterial properties can regulate cell adhesion, morphology (e.g., cell shape, spreading, elongation, and alignment), function and stem cell differentiation [1,8].
1.2.1. CELL ADHESION AND MORPHOLOGY
Adhesion is the first step and critical requirement for anchorage-dependent cells to survive, proliferate, and consequently functionalization or differentiation on a substrate
[45]. Poor adherence of these cells to substrates causes cell quiescence or even apoptosis.
Therefore, cell adhesion is regarded as the initial indicator of cell interplay with its surrounding microenvironments, which precedes all other cellular behaviors, e.g., spreading, migration, proliferation, and differentiation.
Following cell adhesion, they start to conform to the microenvironments, which could result in changes in cell morphology, shape, spreading, elongation, alignment, and eventually cell differentiation. During embryonic development and through life, stem
cells stimulated by different cellular microenvironments are directed into specific cell types that have different morphologies, e.g., branched neurocytes, circular adipose cells, spindle shaped fibroblast, etc. Numerous researches have demonstrated that the opposite is also true: cell morphology (e.g., shape, spreading, elongation and alignment) acts as a potent regulator of cell fate [46].
For cell shape, Ding and co-workers cultured single mesenchymal stem cells (MSCs) from rats on patterned hydrogels with the same adhesive area but different shapes. It was found that the aspect ratio (AR) of the shape regulates the direction of MSC differentiation. Comparing square and rectangular cells, the optimal adipogenic differentiation appears at AR=1, but the optimal osteogenic differentiation was identified when AR≈2. Among the isotropic shapes, i.e., circular, square, triangular and star shaped cells, the optimal adipogenic and osteogenic differentiations were found in circular and star shaped cells, respectively [47]. Apart from studying shape-factors that
affect the phenotype direction of stem cells, Wang and co-workers also proved that the multipotency of MSCs (stemness) decreased with increasing aspect ratio and spreading areas [48].
For cell spreading, it was restricted by decreasing the area of substrate islands, which leads to DNA synthesis inhibition and then induce the expression of involucrin, a marker of terminal function [49]. Also, larger cell spreading on square patterned islands
caused cell survival and growth whereas smaller islands resulted in cell apoptosis [50].
Moreover, capillary endothelial cell shape was changed by ECM molecules, which can regulate cell growth and differentiation [51]. Further, it is logical to extend this
mechanism to stem cells. McBeath and colleagues reported that highly spread MSCs are more contractile, which leads to osteogenic differentiation, whereas MSCs were circularly restricted, resulted in adipocytes [52]. In addition, Wagner and co-workers
found that the elongated MSCs might drive the stem cells towards osteogenic lineage whereas the round morphology enhanced adipogenesis [53].
Cell orientation or alignment, which refers to unidirectional organization of cell body, plays a key role in various cell responses, e.g., cytoskeleton reorganization, membrane protein relocation, migration direction, proliferation, ECM remodeling, and differentiation [29,54,55]. In addition, cell alignment mimics the hierarchical structure,
provides mechanical property and special biological functions on tissue regeneration, including neuron, skeleton, cardiac muscle and tendon [29–31]. Therefore, it is necessary
to build cell alignment in vitro for exploring biomechanics and cell biology, as well as regenerating structured and functional tissue.
1.2.2. BIOMATERIAL TOPOGRAPHY AND STIFFNESS
Both biomaterial topography and stiffness serve as an important indirect signal, which can be sensed by cells through mechanosensing and mechanotransduction. Both material physical cues can be transduced into intracellular biochemical information, and vice versa the intercellular signals can be transformed back to dynamic mechanical signal (e.g., traction forces) [4]. The information communications between cell and
biointerface can activate the integrin-focal adhesion cytoskeleton actin transduction pathway, stimulating cytoskeletal tension and inducing cell morphology deformation and associated signaling cascades that thereby alter gene expression to regulate cell functions and promote tissues regeneration [56]. More interestingly, cells present directed
migration behavior toward topography and stiffness gradients, which can contribute to wound healing and tissue repair [57,58]. Cell adhesion complexes (focal adhesions) and
cytoskeletal generated forces play a significant role in transducing these signals into genetic events that eventually dictate cell fate and functions.
Generally, biomaterial topographical structures can be divided into two types, i.e., isotropic and anisotropic. As mentioned in Section 1.1.2 about the ECM of anisotropic tissues, directional topography and its interaction with cells are of particular interest. Numerous micro and nanofabrication technologies have been developed to prepare aligned topographical features, such as electrospinning [32], plasma oxidation [55],
photolithography [59], direct laser writing [60], etc. As is reported, aligned
micro/nanotopography (e.g., wrinkle, groove, grating, aligned pores, etc.), which are comparable to the natural anisotropic ECM in vivo, can guide the orientation/elongation/migration of many cell types through the contact guidance and the structure-associated organization of cell adhesion ligands [29,33,61]. In addition, the
dimension of aligned topography can significantly affect cell responses. Kim and co-workers observed the area of ventricular myocytes on hydrogel nanogratings ranging from 150 nm×50 nm×200 nm to 800 nm×800 nm×500 nm (width × gap × height) [62].
They found that the biggest features induced larger cell area and longer perimeter. Also, the cells on the hydrogel gratings were more elongated than those on the planar substrate. The majority of cell types has been demonstrated to significantly change their alignment on aligned topographies over the dimension range from nanoscale [29,62–64] to
microscale [32,55,65]. For instance, MSCs were oriented and elongated along the
nanogratings (250 nm width) whereas cells on planar substrate displayed an isotropic morphology [66]. Similarly, osteoblasts on the nanograting substrates were oriented when
the dimensions down to 75 nm in width and 33 nm in depth [67]. On the other hand,
Bashur and colleagues described that the diameter (1 µm) of electrospun PLGA microfibers also can induce the alignment of fibroblasts [68]. Even, the directional
microtopographies with the size from tens to hundreds still showed effects to cell alignment [69]. Microtopography similar to the length-scale of a mammalian cell has
effects on the whole cell body, while nanotopography mediated the signals to subcellular organelles and structures. Therefore, the optimization of topographical structures, particularly their sizes ranging from nanoscale to microscale, is critical to obtain the best cellular performance in TERM.
As mentioned in Section 1.1.2, tissues in the human body have different mechanical features ranging from soft (brain, ~0.1 kPa), to moderately stiff (skin and muscles, ~10 kPa) and stiff (precalcified bone, >1 GPa). As demonstrated in vitro, organic and inorganic biomaterials build mechanically defined microenvironments that can have a significant effect on cell adhesion, morphology, proliferation, and differentiation, resulting in tissue morphogenesis and maturation [70,71]. Yoshikawa and colleagues
reported that myoblast cells showed significant stress fiber generation and flattening with increasing the hydrogel elasticity from 1.4 kPa to 40 KPa [72]. Engler and
co-workers found that MSCs exhibited various morphologies when cultured on polyacrylamide (PAAm) substrates with different stiffness ranging from 1 to 40 kPa. MSCs grown on a soft sample (1 kPa) expressed specific neuronal cytoskeleton markers (β-3-tubulin), whereas cells on the substrates (11 kPa and 34 kPa) displayed expression of early myogenic and osteogenic transcription factors, such as MyoD and CBFα-1, respectively [73]. Importantly, cell responses on different stiffness materials depend on
cell types. Fibroblasts, epithelial cells and endothelial cells showed increased proliferation on stiffer substrates [74–76], whereas neural stem cells displayed better
proliferation on softer substrates [77], probably because of their natural ECM features
and softness of brain tissue. Interestingly, controlling the substrate stiffness could affect non-invasive gene delivery, regulating a cell's ability to uptake exogeneous signalling molecules [78]. Taken together, engineered topographcial and mechanical cues on
substrates are powerful tools for directing cell interactions with the ECM.
1.2.3. BIOMATERIAL CHEMISTRY
Biomaterials chemistry can influence significantly the mass and conformation of the adsorbing proteins and then regulate cell response, which play a critical role on the subsequent cellular behaviors [79,80]. When biomaterials are placed into a biological
environment, cells will not directly respond to the material surface but always via a protein conditioning film that originates from either the culture medium supplemented with fetal bovine serum (FBS) or proteins from biological fluids such as blood, saliva etc. This protein adsorption is generally much faster than the cell adhesion events and hence any alterations in this film will directly affect cellular behavior [81].
Generally, different materials have various chemical properties such as wettability, solubility, reactivity, charge and so on. Surface wettability, indicating interface energies of biomaterial surface (quantified as water contact angle, WCA), has previously been correlated with protein adsorption and cell behavior [79,80]. Wei and co-workers
investigated the effect of surface wettability on competitive protein adsorption (albumin: Alb; fibronectin: Fn) and found that Fn adsorbed more on hydrophilic surfaces, whereas Alb predominantly adsorbed on hydrophobic surfaces. The initial attachment of osteoblastic cells increased with increasing surface wettability, which correlated well with Fn adsorption in the competitive mode [82]. In addition, surface wettability is
critical for cell spreading and differentiation. Generally, cells have good spreading, proliferation and differentiation on hydrophilic surfaces. Mouse osteoblasts on hydrophilic surface ranging from 24 to 31° WCA, showed higher metabolic activity and expressed more osteogenic proteins (alkaline phosphatase and osteocalcin) than those on unmodified counterparts (WCA 72°) [83].
Surface wettability originating from material chemical functionalities (e.g., positive, negative or neutral) can affect protein adsorption and then mediate cell response. For instance, Lee and co-workers described how alkylsilane self-assembled monolayers with different functional groups (OH, COOH, NH2 and CH3) affected 125I-labeled
fibronectin adsorption where at pH 7, COOH is actually COO- and NH2 becomes
NH3+ while OH remains unaffected. They found the adhesion constant and binding
efficiency of the adsorbed Fn for the α5β1 integrins (CH3≈NH2<COOH ≈ OH). Fibronectin interacted more strongly with α5β1 integrins when adsorbed on COOH vs.
OH surfaces suggesting that negative charge may be a critical component of inducing efficient cellular adhesion [84]. The above studies indicate that the specific biomaterial
chemistry plays a major role in protein/cell-material interaction and this chemistry is reflected by the numerous different synthetic polymers and inorganic materials used as biomaterials. Also it illustrates that relative simple concepts such as wettability, is far more complex with large consequences for cell-material interactions.
1.3.
CHALLENGES
Although biophysical and biochemical cues located on biomaterial surfaces proved to profoundly affect (stem) cell behavior, subsequent investigations has raised more questions than they answered, especially about the complex microenvironment of the ECM and its interaction with cells. Understanding ECM complexity in regulating cellular responses is vital for optimizing biointerface design and advancing biomedical material development. How to unlock the code between biomimetic materials and cell interactions still remains relatively unknown with eminent challenges to be met such as: 1) Most physicochemical properties of biomaterials were studied individually and identifying single parameter stimulation which is not appropriate to direct cell function, since cells always interact with multiple cues simultaneously. Therefore, it is vitally important to understand and explore the combined effects of material features on the (stem) cell behavior.
2) Initially, most of these studies used independent substrates with different and randomly selected degrees of biomaterial properties, which provided interesting yet limited information. As an alternative to the traditional experimental approaches using uniform substrates, surface gradient platforms offer an ideal tool to address this challenge in vitro and in vivo, enabling the identification of optimal conditions of cell-material interactions in a high-throughput screening (HTS) strategy. The conventional approaches are incapable of screening the huge amount of potential material parameter combinations to identify the optimal cell responses, and also involves a combination of serendipity along with many series of trial-and-error experiments. For advanced TERM, high-efficient and complex bioanalysis platforms are needed to explore complex microenvironments and study healing, development, and homeostasis.
1.4.
AIM
OF
THE
THESIS
The general aim of this thesis is to develop advanced biomaterial interfaces to explore and elicit their interactions with (stem) cells for improving and accelerating the development of functional and biomimetic materials. Key features in this study are combined physicochemical stimuli and high-throughput screening platform. In order to better direct cell response, complex interfaces with different material physicochemical parameter combinations are developed and utilized for elucidating how multiple cues influence (stem) cell response. Due to the unpredictable fashion between cell and topography interaction, single surface gradients with different materials are developed to identify optimal surface characteristics for directing stem cell response in a high-throughput screening (HTS) manner. Further, surface gradients with double linear or orthogonal features are designed and used to identify optimal surface parameter combinations for directing stem cell response in a HTS fashion.
1.5.
OUTLINE
OF
THE
THESIS
Chapter 1 gives an overview of the recent great discoveries regarding the native cellular
microenvironment, particularly the ECM, and the current progress and challenges with respect to the investigation of cell-biointerface interaction for engineering the functional and biomimetic materials.
Based on the challenges and objectives mentioned above, advanced material interfaces are designed and developed using the novel preparative techniques. In Chapter 2, complex interfaces with stiffness and topography combinations are developed and applied for illustrating the different behavior of the cell-types originating from tissues with different intrinsic stiffness. In Chapter 3, Au nanowire-patterned array platforms with multi-scale design from the macroscale to the nanoscale are developed for regulating human bone marrow-derived mesenchymal stem cell (hBM-MSC) organization through the synergistic effects of surface micro/nanotopography and chemical cues.
In Chapter 4, a novel approach was developed for preparing directional nanotopographic gradients on polydimethylsiloxane (PDMS) substrates which allow us to determine optimal topographical dimension towards osteoblast adhesion and alignment more efficiently and accurately as compared to uniform wrinkle substrates.
Chapter 5 presents a novel strategy translating wrinkled topography gradients from Chapter 4 to clinically relevant inorganic biomaterials (SiO2, Ti/TiO2, Cr/CrO3, and
Al2O3) via the combination of masked plasma oxidation and metal deposition oxidation
methods. The optimal interface parameters are identified for promoting hBM-MSC alignment, elongation, filopodia development as well as cell adhesion. In Chapter 6, orthogonal stiffness and wettability gradients within a single substrate are developed for elucidating combined physical parameter influences on stem cell behavior and thereby gaining insights in desired cellular responses. hBM-MSC behavior is non-linearly regulated by surface stiffness and wettability. The optimal combined properties for promoting cell adhesion, nucleus size and vinculin expression are identified based on the orthogonal double gradients.
Finally, in Chapter 7 and Summary, the general discussion and summary of the above chapters are presented and placed into a broader perspective and provide an outlook for any potential future work.
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C
HAPTER
T
WO
M
ECHANICAL
P
ROPERTIES OF
A
LIGNED
N
ANOTOPOGRAPHIES FOR
D
IRECTING
C
ELLULAR
B
EHAVIOR
Qihui Zhou, Patrick Wünnemann, Philipp T. Kühn, Joop de Vries,
Marta Helmin, Alexander Böker, Theo van Kooten, Patrick van Rijn*
Advanced Materials Interfaces, 2016, 3, 1600275.
A
BSTRACTTailoring cell-surface interactions is important for designing medical implants as well as regenerate medicine and tissue engineering materials. Here we transcend the single parameter system via translating hard nanotopography into soft polymeric hydrogel structures via hydrogel
imprinting lithography. The response of these cells to the
nanotopography of the same dimensions but with different mechanical properties displayed unexpected behavior between “hard” tissue cells and “soft” tissue cells.
2.1. I
NTRODUCTIONCells are able to respond to various material surface properties such as stiffness, topology, chemistry and wettability [1–6]. They will adjust cellular adhesion properties,
proliferation behavior, orientation, protein expression and also differentiation [7–11].
Knowledge about the effects of these surface confined properties onto cellular behavior will have a significant impact on the design and development of medical implants as well as scaffold materials for regenerative medicine and tissue engineering.[3,12–17] Each
of the previously mentioned materials properties have been extensively investigated by systematically changing one of these properties and elucidate the response of cells to these changes [18–23]. Although, these studies have provided many new insights, they
mostly tackled a single property e.g. changing topographical features on the model polymer poly(dimethylsiloxane) (PDMS) such as wrinkle or grating structure; altering surface wettability; or using solid materials and hydrogels of different stiffness [24–28].
PDMS and hydrogels are popular materials because they are frequently used for biomedical applications when properly chosen. As all these stimuli induce cell responses, there is a need for cross-combination studies to obtain deeper insights, since a surface will always be a combination of properties such as stiffness, topography and wettability[8,29–32].
Here we demonstrate the importance of developing materials which enable investigations towards more complex interfaces, in order to elucidate cellular responses towards combined surface properties. For the system development, PDMS and pHEMA (Poly(2-hydroxyethyl methacrylate)) hydrogels were chosen as both are already used for biomedical applications such as medical implants and contact lenses, respectively. Nanostructured PDMS was translated to pHEMA hydrogels resulting in the same topography but with a different stiffness. These property combinations were expected to trigger different cell responses both macroscopically and on a molecular scale. To the best of our knowledge, for the first time nanotopographical effects are combined with mechanical material properties differing over two orders of magnitude in Young’s modulus. Results of this approach show that when cells from different tissues with an intrinsically different stiffness are targeted with the intention of inducing tissue directionality and altering protein expression behavior, the surface needs to be tailored with respect to both topography and mechanical properties depending on the cell-type. Development of materials with specific physical parameter combinations along with a diverse number of tissue specific cell-types will facilitate biomedical materials design more rapidly and accurately.
2.2.
R
ESULTS ANDD
ISCUSSIONAligned nanowrinkles on PDMS substrates were used as the hard topographical substrate. This substrate was also used as the template to prepare hydrogel wrinkles serving as the soft topological substrates with the same features and dimensions (Figure 1). The two different surfaces with similar topography but different stiffness were combined with three human cell-lines, namely osteoblast-like cells (sarcoma osteoblast-like cell-line; SaOs), fibroblasts (skin fibroblast; HSkF) and lens epithelial
cells (LEC), to illustrate the different behavior of the cell-types originating from tissues with different intrinsic stiffness. The behavior on structured surfaces was additionally compared to behavior on flat, non-structured surfaces of the same stiffness (planar PDMS and hydrogel). Combining cells from tissues of different intrinsic stiffness with substrata with similar topologies but differing in stiffness, provided insights into specific behavior towards surface topographies, and it stresses the need to combine surface parameters to derive a better understanding of cells at (bio)interfaces [16,33].
Figure 1. Schematic approach on the preparation of PDMS nanotopographical surfaces
and the translation of the nanotopography into soft polymeric hydrogel structures via “Hydrogel Nano-Imprinting Lithography”.
The wrinkle structures on the surface of PDMS substrates were induced by applying a uni-directional strain with subsequent surface oxidation via oxygen-plasma treatment[34,35]. Surface oxidation induced transformation of the PDMS into a silicon
oxide layer, which is much stiffer than the untreated PDMS (Sylgard® 184; elastomer : curing agent 10:1 (w:w)) [36]. Untreated PDMS displayed a Young’s modulus (YM) of 2.3
MPa (sd.: 0.1 MPa) which was determined by atomic force microscopy (AFM). The Young's modulus has been previously used in this context as a measure for the stiffness. Treatment with oxygen plasma for 480 seconds allowed the modulus (stiffness) to increase to 61.2 MPa (sd.: 3.4 MPa). The same oxygen plasma treatment in combination with a uni-directional strain induced by a 30% substrate extension followed by release of that strain after the oxidation, induced the formation of surface wrinkles (Figure 1,
Figure 2A-C). AFM analysis of the wrinkle dimensions obtained after the
stretch-oxidation-release procedure revealed wrinkles with 255 nm (sd.:11 nm) in amplitude (A) with a wavelength (λ) of 1032 nm (sd.: 57 nm). The oxidized, wrinkled PDMS surface displayed an inhomogeneous distribution of stiffness. On the positions with the highest slope the YM was determined to be ~45 MPa while on top of the wrinkle and on the bottom a YM of ~115 MPa was found. These values deviated from the 61 MPa found for planar oxidized PDMS and are most likely caused by differences in surface contact area of the AFM tip with the PDMS, as the wrinkled PDMS is post-treated with oxygen plasma ensuring a homogenous and full surface oxidation omitting any stiffness and
chemical inhomogeneity. The high slopes will also enable contact with the surface from the side of the AFM tip and hence increase the contact area which most likely was the cause of the lower calculated YM. Although, geometry affected the YM, the values found for structured and non-structured surfaces were in the same range and therefore considered to be comparable.
Figure 2. AFM images and analysis of the height and mechanical properties of the
PDMS (A-C) and hydrogel (D-F) nanowrinkles. Analysis was performed in PBS buffer to obtain a hydrated gel-wrinkle. The distribution of stiffness is strongly connected to the height position in the case of the PDMS wrinkles. For the gel wrinkles this connection between height and stiffness is less defined. Please note the differences in the images (B and E) and y-axes (C and F) for the Young’s moduli (C and F).
For creating soft topographies, imprinting lithography was applied. This is a technique which has been used before to create soft surface confined structures [30,37,38]. Here the
wrinkled PDMS was used as a lithographic template. Imprinting lithography method was performed under inert atmosphere (glove-box) for hydrogels to enhance the degree of polymerization by preventing oxygen-induced termination. Polymerization was performed overnight after which the PDMS substrate was removed. The surface modification of glass with acrylate moieties ensured covalent attachment of the hydrogel layers to the surface enhancing its stability. Before use as a cell substrate, PDMS surfaces were treated with air plasma to render them hydrophilic in order to match the wettability of the hydrogel. Although the hydrogel and freshly activated PDMS have similar wettability (water contact angle, WCA<10°), the presented surfaces vary in specific chemistry which could have an additional influence. The surfaces have good cytocompatibility and hence can be used for comparison, but future endeavors should also include the development of materials which are able to cover large ranges of physical parameters without affecting the specific chemistry or at least keeping the changes in chemistry to a minimum. Unfortunately, such materials are currently unknown to us.
AFM analysis of the hydrogel wrinkles in the hydrated state displayed features representing those of the PDMS. However, the features of the hydrogel were slightly shallower due to a difference in swelling between the top of the wrinkle and the bottom and the wavelength was decreased most likely due to slight contraction of the gel-layer during polymerization and drying (Figure 2D-F). The dimensions of the hydrated wrinkles are A: 168 nm (sd.:24 nm) and λ: 937 nm (sd.: 46 nm). The overall stiffness of the hydrogel was found to be more than two orders of magnitude lower than the PDMS wrinkle substrate. The stiffness of the hydrogel ranged from 300-800 kPa and also displayed an inhomogeneous distribution most likely due to combined effects of altered surface contact area of the AFM tip as well as amount of indentation into the hydrogel material (Figure 2F).
Both substrates with the same topography were combined with three human cell-lines. The cells represent a diverse range of tissues which have intrinsically different properties. Osteoblast-like cells (here represented as SaOs) are found in bone tissue which is very rigid (calcified bone tissue >>1GPa) while LEC is found in the lens of the eye, representing a much softer tissue. Fibroblasts are flexible and responsible for the structural framework of tissues and found in connective tissues that can represent quite a broad stiffness range. These three cell-types have been deposited on the structured substrata as well as on flat control substrates of the same composition. The orientation with respect to the surface structuring was determined by visualizing the surface structures underneath the cells, and relevant protein expression in relation to the used substrate was analyzed along with metabolic activity. To determine the kinetics of cellular behavior, 2 days and 5 days cell cultures were investigated. These incubation times correspond with routine culturing times towards passaging the cells.
For the SaOs cells it was initially anticipated that they would respond more to the stiffer structured substrate by adhering stronger and follow the direction of the surface pattern to a higher degree than the soft topography. The initial notion originates from the fact that conventionally they reside in calcified bone. However, before calcified bone is formed, osteoblasts adhere to a collagen matrix [39]. From microscopy analysis, two
striking behaviors were found (Figure 3). First, from the two days culture it was seen that the SaOs cells do not align on the stiff nanotopography, while the cells did align efficiently on the hydrogel wrinkles. This was not only observed from the orientation of the actin cytoskeleton but also from the focal adhesion points and even the nuclei. The two days culture on PDMS hard (H) nanowrinkle (NW) substrates (2HNW) led to a
random orientation (arrow top right of confocal laser scanning microscopy (CLSM) images indicates topography direction). Secondly, there was a significant difference between the amount of focal adhesions present between hard and soft topography. Apparently, the soft structuring provided better adhesion conditions which would indicate that indeed, as mentioned before, mimicking softer ECM components will influence osteoblasts more than mimicking the actual tissue stiffness. From the control, the 2 day culture on hard, flat PDMS (2HFlat), it became evident that PDMS was not
responsible for the lower adhesion effect since the planar substrate displays ample focal adhesion points in a random orientation (Figure 3). This implies that the presence of directional topography on a hard surface actually inhibits proper cell adhesion, which is in line with our previous studies.[22] The effects of alignment and the amount of focal
adhesion were further strengthened and more pronounced after 5 days of culturing. Even though the surface adhesion was limited for the SaOs on the HNW surfaces, it did
not inhibit proliferation as a confluent layer is reached after 5 days culturing (Figure 3). For the HFlat surface, the cell number did not increase although the amount of focal
adhesions on the flat surface increased over time indicating stronger surface interactions. Although the SaOS cells responded more strongly towards soft nanostructured surfaces, as identified by percentage of aligned cells (Figure 4), at day five there was no statistically significant difference in alignment on the hard (65.3 ± 5.6 %) and soft (57.8 ± 5.6 %) topography. At day five, both structured surfaces displayed confluent and highly aligned cells while both flat surfaces had cobblestone like morphologies and no alignment (Figure 3 and 4). Observing cellular morphologies showed that after 2 days and 5 days culturing the SaOs displayed elongated morphology on the hard and soft wrinkle surfaces (Figure 3). Furthermore, a quantitative analysis showed that, after 2 days of incubation, SaOs cultured on the soft wrinkle surface achieved a significant increase in elongation efficiencies (p< 0.01) compared to those on the soft flat and hard wrinkle and hard flat surfaces. However, at day five there was no statistically significant difference in elongation efficiencies between the hard and soft topography, similar to the alignment analysis.
Figure 3. Confocal laser scanning fluorescence microscopy images of SaOs (top two
rows), HSkF (middle two rows) and LEC (bottom two rows) cultures of two days and five days on hard surfaces (flat & wrinkled) and soft hydrogel surfaces (flat & wrinkled). Images shown are for focal adhesion (vinculin; green), actin cytoskeleton (red), extra-cellular matrix (blue, for HSkF and LEC), nucleus (blue; SaOs & HSkF for 2SFlat and 5SFlat). The arrow (top right) indicates the orientation of the nanowrinkles while the bar
indicates a flat surface. Bottom left indicates substrate used XYZ where X is the number
of days for cell culturing, Y depicts Hard (H) or Soft (S) surfaces and Z notes whether nanowrinkles (NW) were used or a flat surface (Flat).
The amount of focal adhesions was quantified in order to determine the relative degree of cell adhesion per cell for each of the surfaces. This was done by segmentation of the focal adhesion points and obtaining a binary image, which enables the determination of the effective surface coverage by focal adhesion, and by subsequent extrapolation to the amount of focal adhesion area per cell using previous reported approaches and developed software (Table 1) [40].
The quantification of the focal adhesions of the SaOs cells to the different surfaces showed the strong surface-dependent behavior not only in orientation but also in actual amount of focal adhesion area per cell based on the surface properties related to stiffness and topography. For the time point of two days, the focal adhesion area per cell was five times higher for the soft nanowrinkles (99±24 µm2/cell) than for the stiffer
nanowrinkles (21±1 µm2/cell). To a lesser degree, surface structuring also inhibited cell
adhesion on stiff surfaces demonstrated by comparing flat stiff PDMS to hard nanowrinkles, 62±8 µm2/cell versus 21±1 µm2/cell, respectively. Conversely, surface
structuring enhanced cell adhesion on soft surfaces as the focal adhesion on the flat soft was only 37±7 µm2/cell as compared to the 99±24 µm2/cell for the structured soft
surface.
Table 1. Focal adhesion area of cells on different substrates.
a) The focal adhesion area/cell was determined using the coverage value divided by the
number of cells within the covered area; b) Incubation time of 5 days could not be
determined (n.d.) due to overlapping cell layer and hence determining the proper number of cells per area was not possible.
Fibroblasts have the natural tendency to align themselves parallel to each other in their cell growth. Overall, the alignment in the entire population was random, similar to that of flat PDMS and hydrogel (Figure 3). In the case of the HSkF’s for both the hard and soft wrinkles, an alignment was observed in the direction of the nanowrinkle pattern and the alignment increased with longer culture times (Figure 3 and 4). Although both nanostructured surfaces were able to direct the cell orientation, the focal adhesion area per cell after two days (2HNW; 2SNW) was higher for the soft topography (140±68 µm2/cell) than for the stiff topography (46±2 µm2/cell). The difference between the
surface adhesions seemed to be maintained even for the five days culture at which cell populations had become confluent. However, this difference was more qualitative as there were too many cells to properly assess the focal adhesion area per cell due to proliferation beyond a single cell layer.
Figure 4. Macroscopic response of cells towards surface features comparing the
wrinkled feature with flat and hard with soft. Cells are considered aligned when cell longitude axis is within 10° of that of the direction of the surface features.
The lens epithelial cells did not display any tendency towards alignment as observed in
Figure 3 and 4. On none of the surfaces did the cells or the focal adhesion points
follow the nanotopography. Furthermore, similar morphologies were obtained on the hard nanotopography surface and on the flat surface. This is also observed from the rather low focal adhesion coverage values and the average focal adhesion area per cell (Table 1). From Figure 3 (2SNW) it could even be deduced that the LECs tend to avoid
the soft nanostructured surface and prefer the non-structured surface. There were hardly any cells present on the soft nanostructured surface except on the edges of the printed surface and on a few defective areas where printing had not structured the surface. On the stiff substrate a homogeneous distribution across the surface was observed. This cell line also supports that nanotopography has the potential to affect cell adhesion.
The comparison between the cells originating from tissues with different intrinsic stiffness in our study indicates that the “soft cell” (cell originating from soft tissue, LEC) is inhibited by the soft topography while in case of the “hard cell” (cell originating from stiff tissue, SaOs) the adhesion is inhibited in combination with the stiff nanotopography as was observed for the SaOs cells. This cannot be definitely concluded, but it does demonstrate the importance of combining different surface features such as topography and mechanics as shown here. Parameter combinations of this kind will provide deeper insights in cell behavior at interfaces, more so than working with isolated single surface parameters.
The aversion of LEC against the soft topography did not inhibit its proliferation. After five days culturing, all surfaces were well covered and with an increase in amount of focal adhesion, with the most significant increase for the soft topography (Table 1). Apparently, once the cells had sufficient support from each other, surface adhesion was also stimulated. In addition to increased surface adhesion, the morphology of the cell layer displayed a change in packing and structure. Cell layers formed by LEC have an open structure with large unoccupied areas, as was seen for samples 5H
NW and 5HFlat
(Figure 3). The LEC-layer covering the soft topography after five days culturing (5S
NW)
formed a much denser cell layer which is also generally observed for collagen coated tissue culture polystyrene (TCPS) substrates. The more homogenous packing of the LECs inside the cell layer probably has to do with the extra-cellular components secreted by these cells. In Figure 3 (5SNW) the fibronectin matrix marked in blue
to the other surfaces. More inter-cellular interactions allow for a more densely packed layer being formed, which is needed to overcome the unfavorable interactions with the hydrogel wrinkle structures.
In addition to the macroscopic responses such as adhesion and alignment, it was expected that there would also be a significant influence on the molecular level and that metabolic activity and protein expression would be affected by the physical parameters of the substratum (Figure 5 and 6). To this end, metabolic activity (XTT) was quantified and the most relevant protein expressions were qualitatively determined using fluorescence imaging. ECM upregulation was visualized by targeting collagen I, which was compared on the different surfaces for SaOs and HSkF. Alkaline phosphatase (ALP) was analyzed for the SaOs cells indicating mineralization, and alpha-smooth muscle actin (α-SMA) was analyzed for the LEC, which is a strong marker for the mesenchymal transition of epithelial cells towards myofibroblasts and considered a marker for fibrosis[41–43].
The results of the XTT assay showed that SaOs cultured on surfaces over a period of 5 days exhibited levels of metabolic activity that were statistically different from SaOs cultured on soft nanostructure. The metabolic activity of HSkF on soft nanotopography was the highest compared to on other surfaces. For LEC, the level of metabolic activity on the hard wrinkled surface was the highest compared to on other surfaces.
Figure 5. Viability assay (XTT) displaying the metabolic activity of SaOs, HSkF and
LEC on soft and hard surfaces both flat and with aligned nanotopography for 2 and 5 days culture.
Figure 6. Fluorescence imaging of expressed proteins as a consequence of the surface
parameters. For SaOs, Collagen I (green) and Alkaline Phosphatase (ALP, green) was analyzed, for HskF, Collagen I (green) was analyzed, and for LEC, α-SMA (green) was analyzed. Scale bars are 75 µm and apply to all images.
The initial hypothesis with respect to osteoblast behavior was that it would either respond to the hard substrates because of the relation with stiff calcified bone or that it would respond to soft as one of the early events is that osteoblasts adhere to collagen before producing mineralized bone tissue. It was identified that osteoblasts respond preferentially initially to soft topographies as it relates more to the initial adhesion on a collagen matrix which was also shown by the alignment efficiency in Figure 4. However, for the alignment there was no difference after 5 days culturing. Collagen production and mineralization are important functions for osteoblasts as it constitutes for the formation of mineralized bone tissue. Upon investigating the expression of collagen I and ALP some very interesting differences were detected. Both collagen I production and ALP expression were more stimulated on the hard layers after 5 days (Figure 6) whereas initially more collagen I as well as ALP was observed after 2 days on
